Homologous stress adaptation, antibiotic resistance, and biofilm forming ability of Salmonella enterica serovar Heidelberg ATCC8326 on different food-contact surfaces following exposure to sublethal chlorine concentrations1

Homologous stress adaptation, antibiotic resistance, and biofilm forming ability of Salmonella... Abstract Salmonella enterica serovar Heidelberg (American Type Culture Collection; ATCC 8326) was examined for the ability to adapt to the homologous stress of chlorine through exposure to increasing chlorine concentrations (25 ppm daily increments) in tryptic soy broth (TSB). The tested strain exhibited an acquired tolerance to chlorine in TSB with the tolerant cells growing in concentrations up to 400 ppm. In addition, the chlorine stressed cells displayed rugose morphology on tryptic soy agar (TSA) plates at 37°C. The minimum inhibitory concentration (MIC) of chlorine for adapted (rugose and smooth) cells was determined to be 550 ppm and 500 ppm, respectively whereas the MIC for the control was 450 ppm. The biofilm forming ability of the adapted and control cells were examined on both plastic and stainless steel surface at room temperature and 37°C. The rugose variant, in contrast to the smooth (adapted and control) showed the ability to form strong biofilms (P ≤ 0.05) on a plastic surface at room temperature and 37°C. Rugose cells compared to smooth and control attached more (P ≤ 0.05) to steel surfaces as well. The possibility of cross-adaptation was examined by exposing the adapted and control cells to different antibiotics according to the Clinical & Laboratory Standards Institute guidelines. Adapted cells exhibited reduced susceptibility to some of the antibiotics tested as compared to control. The findings of this study suggest that exposure to sublethal chlorine concentration during the sanitization procedure can result in tolerant Salmonella cells. Chlorine may confer cross-protection that aids in the survival of the tolerant population to other environmental stresses. INTRODUCTION Salmonella enterica is well known as the leading cause of foodborne infections globally (Painter et al., 2013). The serotypes of Salmonella enterica are reported to result in over a million cases of illnesses, thousands of hospitalizations, and a few hundred deaths each year in the United States and also around the world (Mead et al., 1999; Scallan et al., 2011; CDC, 2011a). According to an estimate by the Centers for Disease Control and Prevention (CDC) in 2011, nontyphoidal Salmonella, which includes all Salmonella serotypes except Typhi and Paratyphi, was among the top five pathogens in the United States causing the most reported foodborne illnesses, hospitalizations, and deaths annually (CDC, 2011b). In addition, nontyphoidal Salmonella, which is second to norovirus, is estimated to cause 11% illnesses, 35% hospitalizations, and 28% deaths each year in the United States alone (CDC, 2011b). Moreover, of all the Salmonella enterica serotypes, the top four serotypes associated with animal products and most often reported to cause human infections are Typhimurium, Enteritidis, Newport, and Heidelberg (Hur et al., 2012). There are several food commodities implicated in foodborne illnesses, however, between 1998 and 2008, poultry accounted for 9.8% of the illnesses, 11.5% of the hospitalizations, and 19.1% of the deaths (Painter et al., 2013). Poultry is considered a major source of Salmonella contamination that results in human illness (Morris and Wells, 1970). In 2013, a multistate outbreak of multidrug-resistant Salmonella Heidelberg was linked to branded chicken. This outbreak was reported to result in 634 cases of illnesses and 38% hospitalizations (CDC, 2014). Due to the burdens attached to the control of foodborne pathogens, poultry processors employ many methods to prevent pathogens from contaminating poultry meat. One of the methods include the use of USDA approved antimicrobials for disinfecting poultry meat and sanitizing food-contact surfaces within the poultry processing facility (USDA-FSIS, 2015). The antimicrobials are capable of destroying both spoilage and pathogenic bacteria. Examples of antimicrobials used to disinfect, decontaminate, or preserve food products include peracetic acid, calcium hypochlorite, sodium nitrite, trisodium phosphate, and sodium hypochlorite (Capita and Alonso-Calleja, 2013; Alonso-Calleja et al., 2015). The misuse of these compounds has allowed pathogenic bacteria to develop adaptation (Capita et al., 2014). Previous studies have shown that using an antimicrobial at a sublethal concentration may potentially aid in the survival of foodborne pathogens by enabling them to survive challenging conditions such as antimicrobial inactivation (Sheridan et al., 2012; Capita et al., 2014). This increase in bacteria tolerance to antimicrobials is a global public health concern (Braoudaki and Hilton, 2004) Several studies have documented the ability of bacteria to adapt to stress posed by antimicrobials, which they use to induce cross-protection against other stressful conditions (Davidson and Harrison, 2002; Braoudaki and Hilton, 2005; Capita et al., 2014). The majority of these studies provides significant evidence that reduced susceptibility to one antimicrobial can cause cross-adaptation to another (Braoudaki and Hilton, 2005). Since chlorine is the antimicrobial agent that is most common in sanitizing solutions, adaptation to chlorine could constitute a potential threat to food safety by inducing cross-protection to antibiotics that are clinically important (Braoudaki and Hilton, 2004; Molina-Gonzalez et al., 2014). Antibiotic resistance has continued to be a global public health issue; research has been conducted that examines different mechanisms used by pathogens, such as Salmonella, Listeria, and E.coli, to adapt to disinfectants and induce cross-resistance to antibiotics (Buchmeier and Heffron, 1990; Lou and Yousef, 1996; Braoudaki and Hilton, 2004; Capita et al., 2014; Molina-Gonzalez et al., 2014). Some of this research demonstrates that an adapted bacteria will most likely use similar resistance mechanism to offer cross-protection to antibiotics (Braoudaki and Hilton, 2004). In addition, it was reported that a frequent strategy used by pathogenic microorganisms to adapt to antimicrobials, is the formation of biofilm (IFT, 2006). Biofilms are bacteria that cluster together on a surface and are surrounded by an exopolymeric matrix that makes the biofilm matrix impenetrable to any harmful substance (Costerton et al., 1995; Iturriaga et al., 2007; Bae et al., 2012). An important challenge faced by food processors is the control of bacteria present in the biofilm. It was documented that biofilms are more difficult to destroy because they possess tolerance to environmental stresses when compared to free cells (Frank and Koffi, 1990; Joseph et al., 2001). In poultry processing, attachment of bacteria to surfaces of equipment can be a source of product contamination (Barnes et al., 1999; Giaouris et al., 2005). Some studies have reported the incidence of cross-contamination due to bacterial attachment to equipment surfaces such as utensils used in food preparation (Scott and Bloomfield, 1990; Jiang and Doyle, 1999; Kusumaningrum et al., 2003). Other studies suggested that bacteria could attach to surfaces such as polystyrene, glass, rubber, acrylic, and cement (Czechowski, 1990; Mafu et al., 1990; Krysinski et al., 1992; Barnes et al., 1999; Nguyen and Yuk, 2013). Therefore, in this study, the homologous stress adaptation of Salmonella Heidelberg exposed to chlorine at a sublethal concentration was determined by observing changes in minimum inhibitory concentration (MIC). The biofilm formation of the adapted cells compared to non-adapted cells on different food-contact surfaces was also measured. Antibiotic susceptibility patterns of adapted cells were further investigated. MATERIALS AND METHODS Salmonella Heidelberg Strain and Culture Preparation Salmonella Heidelberg (American Type Culture Collection; ATCC 8326) was maintained on tryptic soy broth (TSB; Sigma-Aldrich Co., St. Louis, MO). Prior to experiments, frozen cells were streaked on tryptic soy agar plates (TSA; Sigma-Aldrich Co.) and incubated in a refrigerated incubator (PR505755R, Thermo Fisher Scientific, Waltham, MA) at 37°C for 24 h. A single colony was subcultured in TSB at 37°C for 20 to 24 h. Working cultures were stored at 4 ± 1°C on TSA slants and were subcultured monthly. Chlorine Source Sodium hypochlorite that contained 5% available chlorine (AC419552500, ACROS Organics, Morristown, NJ), was used as the source of chlorine in this study. The amount of free chlorine in the sodium hypochlorite was confirmed using the HACH (chlorine test kit) Pocket Colorimeter (No. 5870023, HACH Company, Loveland, CO) in accordance with the manufacturer's instructions. A sterile solution of chlorine was prepared in an appropriate media before each experiment. Determination of Minimum Inhibitory Concentration (MIC) of chlorine The MIC value was established using the broth macro and microdilution method in accordance with the Clinical & Laboratory Standards Institute guidelines (CLSI, 2008) with minimal modifications. One colony of the planktonic cells, obtained from a TSA plate was inoculated in 10 mL TSB, and incubated at 37°C for 20 to 24 h. It was determined in our laboratory that 20 to 24 h incubation of the bacterial culture contained approximate 1 × 109 CFU/mL. To start the experiment, 96-well microtiter plates (SKU 229190, Celltreat Scientific Product, Pepperell, MA) were used. A volume of 100 μL of chlorine in TSB that contained double the desired concentration was added to each well of the plate. Afterwards, 100 μL of inoculum prepared to a final concentration of ∼ 106 CFU/mL was added to make the final volume in each well 200 μL. A positive control that contained (100 μL TSB + 100 μL inoculum devoid of chlorine) and a negative control (200 μL TSB) were included in the experiment. The microwell plate was incubated at 37°C for 24 h and visible growth of bacteria in each well was determined by turbidity. The MIC was established as the lowest chlorine concentration that is necessary to inhibit the growth of Salmonella Heidelberg after 24 h of incubation. The exact MIC was determined by narrowing the range of chlorine concentrations in each well for the subsequent experiment. Stress Adaptation Study This study was performed by preparing Salmonella stock culture to a final concentration of ∼109 CFU/mL. The starting concentration of chlorine was 125 ppm, an aliquot of 100 μL of the inoculum was transferred to a polypropylene tube (Cat No. S50712, Fisher Scientific, Fair Lawn, NJ) that contained 9.9 mL (TSB + chlorine). After the transfer, the tube contained a final inoculum concentration of ∼ 107 CFU/mL. Upon incubation for 24 h at 37°C, turbidity in the tube was used to examine bacteria growth. The suspension was then diluted and plated on TSA plates to enumerate Salmonella growth. An additional aliquot of 100 μL was transferred from the same turbid tube to a sterile tube that contained 9.9 mL (TSB + higher sublethal chlorine concentration). This procedure was continued by increasing the concentration of chlorine by 25 ppm daily until no visible growth was observed in the tube. This required 12 days of incubation. The suspension in the last tube with visible bacteria growth was again diluted for Salmonella enumeration. The cells obtained on TSA plates after incubation for 24 h at 37°C were re-streaked and stored on TSA plate that did not contain chlorine. The cells were considered adapted Salmonella Heidelberg cells. The TSA plates containing the adapted cells were stored at 4 ± 1°C with the cells being transferred weekly to fresh non-chlorinated TSA plates. Adaptive Stability to Chlorine The adaptive tolerance to chlorine of the adapted Salmonella Heidelberg cells was tested by determining the MIC for the adapted cells against chlorine using the broth macro dilution method according to the CLSI guidelines (CLSI, 2012).This was done after storage on TSA plate that did not contain chlorine. A colony of the adapted cell was inoculated into a borosilicate round bottom glass culture tube (No. 1496128, Fisher Scientific) containing 10 mL TSB with chlorine at concentrations that were below, equivalent and above the MIC. Non-adapted Salmonella Heidelberg cells, which served as the control, were also tested at the same concentrations. Turbidity was used to observe bacteria growth, the glass culture tube showing no visible growth after incubation at 37°C for 24 h was considered the MIC of the adapted and control cells. Biofilm Formation on plastic surface The standard crystal violet assay as previously described by Patel and Sharma, (2010) was used to determine the biofilm forming ability of the chlorine adapted (rugose and smooth) and control cells. An overnight culture (∼ 20 h) of adapted and control Salmonella cells grown in TSB were diluted to achieve a final inoculum level of ∼ 106 CFU/mL. The treated cells (rugose and smooth) were cultured in TSB that contained 400 ppm chlorine which is the highest concentration that supports bacteria growth, whereas the control cells were cultured in chlorine-free TSB. To begin the experiment, 96-well polystyrene cell culture plates were prepared by dispensing 200 μL of the inoculum into duplicate wells for each type of adapted Salmonella Heidelberg morphotypes and the control. TSB devoid of inoculum was used as a negative control. The plates were incubated at room temperature and 37°C for 48 h. After incubation was complete, 200 μL of the inoculum was aspirated from each well of the plate, and each well was washed five times with sterile distilled water so that all loosely attached cells may be removed. The plates were later air-dried for 45 mins, and 200 μL of crystal violet solution (0.41% w/v dye, AC447570500, ACROS Organics) was dispensed into each well. The plates were incubated at room temperature for an additional 45 mins, after which the dye was aspirated from each well, and the well was washed five times using sterile distilled water. Each well was allowed another 45 mins to air dry, and then 200 μL of 95% ethanol (A406P4, Fisher Scientific) was added to each well. The content of each well was mixed to dissolve the crystal violet dye. Biofilm formation in each well was measured by an optical density (OD600) reading using a micro quant microplate spectrophotometer (Model ELx800, BioTek Instruments, Winooski, VT). The attached cells of adapted (rugose and smooth) and control on the plastic plate were enumerated. Similar to the crystal violet assay, 200 μL of the inoculum prepared to ∼ 106 CFU/mL was dispensed into each well of a 24-well tissue culture plate (SKU 229123, Celltreat Scientific Product, Pepperell, MA) in duplicate. The plate were then incubated at room temperature and 37°C for 48 h. After incubation, each well was aspirated and washed three times with sterile distilled water to remove any loosely attached bacteria. The strongly attached cells were scraped into 0.1% peptone water, vortexed, and serially diluted. After dilution, an aliquot of 100 μL was plated on TSA plates, incubated at 37°C for 24 h in order to count the number of attached cells. Biofilm Formation on stainless steel (SS) surface To examine the influence of chlorine stress on the ability of Salmonella Heidelberg to attach to stainless steel (SS), a previously described method with minimal modifications was used (Hood and Zottola, 1997). Stainless steel scoops were purchased from Spring Chef (www.springchef.com, Dallas, TX). The SS scoops were washed with detergent upon arrival, sanitized in 70% ethanol and autoclaved at 121°C for 15 mins. A total of 2 mL of previously prepared inoculum (∼ 106 CFU/mL) of adapted and control Salmonella Heidelberg were dispensed into each 5 mL scoop. The scoops were placed in sterile Nalgene pans (Cat No. 1336110, Fisher Scientific) that were placed in a covered storage box (No. 552834742, Walmart Stores, Inc. Bentonville, AR) and incubated at both room temperature and 37°C for 48 h. After incubation, each scoop was aspirated and washed three times with sterile distilled water to remove any loosely attached bacteria cell. The strongly attached cells on the scoop were scraped into a solution of 0.1% peptone water and vortexed for 2 minutes before being subjected to a 10-fold dilution in TSB. An aliquot of 100 μL from each dilution was plated on TSA plates, incubated at 37°C for 24 h in order to quantify the number of attached cells. Determination of Antibiotic Susceptibility Chlorine-adapted Salmonella Heidelberg and control cells were screened against different antibiotics on Mueller-Hinton broth and Mueller-Hinton agar (MHB, MHA; Oxoid Co., Nepean, ON, Canada) to determine susceptibility. Adapted cells were grown in TSB containing chlorine (400 ppm) and the control cells were grown in TSB devoid of chlorine. The inoculum was prepared in MHB to a final concentration of ∼ 106 CFU/mL. MIC broth microdilution and disk diffusion method as described by the CLSI (2008) guideline with slight modifications were used in the study. The following antibiotic disks (Fisher Scientific) were used: sulphamethoxazole/trimethoprim (SXT, 25 μg), gentamicin (GN, 10 μg), streptomycin (S, 10 μg), amoxicillin/clavulanic acid (AMC, 30 μg), nalidixic acid (NA, 30 μg), ciprofloxacin (CIP, 5 μg), ceftriaxone (CTX, 30 μg), ampicillin (AMP, 10 μg). The inhibition zones were measured and recorded as susceptible, intermediate, or resistant according to the guidelines by CLSI (2008). Cultures of Escherichia coli 25922 with known antibiotic resistance patterns were used as control reference strain. The MIC for the antibiotics was determined using a 96-well cell culture plate. Each panel in the plate contained 5 dilutions using the MIC breakpoints provided by the CLSI (2008) guidelines for each antibiotic tested in this study. Positive control (MHB + inoculum) and negative control (MHB only) were maintained throughout the experiment. The plate was incubated at 37°C for 24 h, and the results were recorded as the least concentration of antibiotic that prevent the growth of bacteria either as susceptible, intermediate, or resistance as stated in the CLSI (2008) guidelines. Statistical Analysis Each Experiment was Evaluated Sequentially on Different Days All data analysis in this study was carried out using analysis of variance (ANOVA) in the General Linear Model (GLM) of SAS v. 9.4 (SAS Institute, Cary, NC; Steel and Torrie, 1980). Means separations were acquired using Fisher's least significant difference test. The treatments and controls were determined to be significant at the 5% (P ≤ 0.05) level. RESULTS Homologous Stress Adaptation to Chorine It was observed in previous research (Obe et al., 2016) that Salmonella Typhimurium undergoes a morphological change upon a prolong exposure to sublethal chlorine concentration. In the present study, Salmonella Heidelberg changes it morphology to the rugose variant after 4 days exposure to chlorine stress at 37°C. Rugose and smooth Salmonella Heidelberg continued to grow together on TSA plate until the chlorine concentration in TSB reached 400 ppm. That was after 12 days of incubation. This concentration was recorded as the maximum chlorine concentration that allowed the growth of stressed Salmonella Heidelberg and the cells recovered at this concentration was referred to as the chlorine-adapted Salmonella cells. The MIC of chlorine for Salmonella Heidelberg before sublethal chlorine exposure was determined to be approximately 400 ppm (Table 1). The adaptive tolerance to chlorine was later measured by sub-culturing the stressed cells in a nonselective broth (TSB) after storage on TSA plates devoid of chlorine. The stressed cells (rugose and smooth) were able to grow above the MIC values up to 500 ppm and 450 ppm respectively, compared to the non-exposed cell that could not grow above 400 ppm (Figure 1). This suggests that Salmonella Heidelberg ATCC 8326 possesses a stable homologous adaptation to chlorine. Figure 1. View largeDownload slide Minimum inhibitory concentration (MIC) for chlorine-adapted and control Salmonella Heidelberg after stress adaptation. Data represent the average of 2 replicates. The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Figure 1. View largeDownload slide Minimum inhibitory concentration (MIC) for chlorine-adapted and control Salmonella Heidelberg after stress adaptation. Data represent the average of 2 replicates. The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Table 1. Minimum inhibitory concentrations (MICs) and minimum bactericidal concentrations (MBCs) of chlorine (ppm) for Salmonella Heidelberga (ATCC8326) before adaptation. Dilution method  MIC (ppm)  MBC (ppm)  Micro  400  500  Macro  500  500  Dilution method  MIC (ppm)  MBC (ppm)  Micro  400  500  Macro  500  500  aSalmonella Heidelberg culture prior to chlorine exposure. Data represent the average of 3 replicates. View Large Biofilm Formation on Plastic Surface To determine whether adaptation to chlorine would influence the strength of biofilms formed, Salmonella Heidelberg was cultured in the presence of chlorine (400 ppm) for the adapted cells and in the absence of chlorine for the non-adapted (control) cells. The optical density reading (OD600) of the adapted and control cells were observed and used to determine biofilm forming ability of each of the Salmonella variants. In the present study, Salmonella Heidelberg formed biofilms on polystyrene plastic surface both at room temperature and at 37°C. At room temperature, there was a significant difference (P < 0.05) in the biofilms formed by the adapted cells versus control. Adapted rugose formed the strongest biofilm with OD600 value of 3.4, followed by adapted smooth with OD600 value 1.13 representing a moderate biofilm former, compared to control that formed the weakest biofilm with OD600 value of 0.68 (Figure 2A). At 37°C, similar to room temperature, the morphotypes tested were significantly different (P < 0.05) in their ability to form a biofilm. Adapted rugose showed a strong biofilm forming ability with an OD600 value of 3.4. There were no differences observed for smooth morphologies (both adapted and control) which showed OD600 values of 0.7 and 0.57, respectively (Figure 2B). Biofilm formation on the plastic surface was also determined by enumerating the strongly attached cells on TSA plates. The difference in log values was used to determine the difference in biofilm forming ability of chlorine-adapted cells against control. The result were reported in Log CFU/mL. At room temperature, there was no significant difference (P > 0.05) observed for adapted cells (rugose compared to smooth), and adapted smooth compared to control. However, adapted rugose has a significant higher attached cell concentration of 5.05 log cfu/mL when compared to control which is 4.85 log cfu/mL but the difference was not enough to establish a biological significance (Figure 3A). A significant difference (P < 0.05) was determined for the attachment of chlorine-adapted cells (rugose and smooth) having 5.25 and 4.9 log cfu/mL respectively compared to control, which has a cell concentration of 4.7 log cfu/mL at 37°C (Figure 3B). In addition, at 37°C, rugose appears to have more CFU/mL of attached cells on plastic surface compared to smooth (adapted and non-adapted) cells. Figure 2. View largeDownload slide (A) Biofilm formation by Salmonella Heidelberg after 48 h on plastic surface using 96-well polystyrene microtiter plate at room temperature. Means with different superscripts indicate significant differences in the biofilm forming ability of chlorine-adapted and control Salmonella Heidelberg morphotype. (P = 0.0001; SEM = 0.109; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, dark gray bar represents non-exposed positive control, and light gray bar represent (broth only) negative control. (B) Biofilm formation by Salmonella Heidelberg after 48 h on plastic surface using 96-well polystyrene microtiter plate at 37°C. Means with different superscripts indicate significant differences in the biofilm forming ability of chlorine-adapted and control Salmonella Heidelberg morphotype. (P < 0.0001; SEM = 0.058; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, dark gray bar represents non-exposed positive control, and light gray bar represent (broth only) negative control. Figure 2. View largeDownload slide (A) Biofilm formation by Salmonella Heidelberg after 48 h on plastic surface using 96-well polystyrene microtiter plate at room temperature. Means with different superscripts indicate significant differences in the biofilm forming ability of chlorine-adapted and control Salmonella Heidelberg morphotype. (P = 0.0001; SEM = 0.109; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, dark gray bar represents non-exposed positive control, and light gray bar represent (broth only) negative control. (B) Biofilm formation by Salmonella Heidelberg after 48 h on plastic surface using 96-well polystyrene microtiter plate at 37°C. Means with different superscripts indicate significant differences in the biofilm forming ability of chlorine-adapted and control Salmonella Heidelberg morphotype. (P < 0.0001; SEM = 0.058; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, dark gray bar represents non-exposed positive control, and light gray bar represent (broth only) negative control. Figure 3. View largeDownload slide (A) Quantification of Salmonella Heidelberg after 48 h attached to 24-well polystyrene plate at room temperature. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the polystyrene plastic plate. (P = 0.081; SEM = 0.041; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. (B) Quantification of Salmonella Heidelberg after 48 h attached to 24-well polystyrene plate at 37°C. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the polystyrene plastic plate. (P = 0.002; SEM = 0.029; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Figure 3. View largeDownload slide (A) Quantification of Salmonella Heidelberg after 48 h attached to 24-well polystyrene plate at room temperature. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the polystyrene plastic plate. (P = 0.081; SEM = 0.041; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. (B) Quantification of Salmonella Heidelberg after 48 h attached to 24-well polystyrene plate at 37°C. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the polystyrene plastic plate. (P = 0.002; SEM = 0.029; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Biofilm Formation on SS Surface To determine the biofilm formation of chlorine adapted and control Salmonella Heidelberg on a different food-contact surface other than plastic, the strongly attached cells on stainless steel scoops were measured on TSA plates. Similar to plastic quantification, the results were reported in log CFU/mL and the difference in log values were used to determine the difference in biofilm formation between adapted and control cells. There was a significant difference (P > 0.05) observed in the biofilm formation of the adapted rugose cell compared to adapted smooth and control at room temperature. Adapted rugose has a higher attached cell concentration of 5.25 log cfu/mL when compared to the smooth (adapted and control), which has cell concentrations of 4.9 and 4.75 log cfu/mL respectively (Figure 4A). Whereas at 37°C, there was no difference in the concentration of adapted cells (rugose and smooth) that has 5.3 and 5.25 log cfu/mL respectively. The attached cells for chlorine adapted rugose and smooth were higher when compared to the concentration of the control cells, which is 5.02 log cfu/mL (Figure 4B). Salmonella Heidelberg cells attached more to the SS scoop surface at 37°C than that observed at room temperature. Figure 4. View largeDownload slide (A) Quantification of Salmonella Heidelberg after 48 h attached to stainless steel scoop at room temperature. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the stainless steel scoop. (P = 0.033; SEM = 0.071; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. (B) Quantification of Salmonella Heidelberg after 48 h attached to stainless steel scoop at 37°C. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the stainless steel scoop. (P = 0.015; SEM = 0.031; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Figure 4. View largeDownload slide (A) Quantification of Salmonella Heidelberg after 48 h attached to stainless steel scoop at room temperature. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the stainless steel scoop. (P = 0.033; SEM = 0.071; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. (B) Quantification of Salmonella Heidelberg after 48 h attached to stainless steel scoop at 37°C. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the stainless steel scoop. (P = 0.015; SEM = 0.031; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Antibiotic Cross-Resistance Sensitivity to antibiotics was determined for Salmonella Heidelberg following adaptation to chlorine through exposure to sublethal chlorine concentrations. Zones of inhibition were measured (in millimeters) around an impregnated antibiotic disk. A broth microdilution assay was also performed to determine if there was a change in MIC between the non-exposed and the chlorine tolerant cells. The results for the antibiotic susceptibility patterns of adapted (rugose and smooth) and control cells is shown in Table 2 and 3. The results showed no cross-resistance to antibiotics for the majority of the antibiotics tested when adapted cells were compared to control. However, a reduction in susceptibility was observed in some of the antibiotics. A ≤ 2-mm inhibition zone difference was observed for adapted rugose to GN, S, AMC, and CIP when compared to smooth (adapted and control) (Table 2). The same difference was observed for adapted smooth against SXT and S when compared to control (Table 2). Similarly, a slight increase in MIC was recorded for adapted rugose against S, NA, T, and AMX, which moved from the susceptible to the intermediate category (Table 3). In addition, the adapted (rugose and smooth) cells moved from the susceptible to the intermediate category for T (Table 3). Table 2. Disk diffusion assay of antibiotic cross-resistance patterns of adapted and control S. Heidelberg. Salmonella Heidelberg cultures previously adapted to chlorineb  Susceptibility to indicated antibiotica (μg) SXT GN S AMC NA CIP CTX AMP  Adapted rugose  21/S 15/S 8/R 19/S 18/I 26/S 23/S 20/S  Adapted smooth  19/S 16/S 8/R 21/S 18/I 28/S 23/S 20/S  Control  21/S 16/S 10/R 21/S 18/I 28/S 23/S 20/S  Salmonella Heidelberg cultures previously adapted to chlorineb  Susceptibility to indicated antibiotica (μg) SXT GN S AMC NA CIP CTX AMP  Adapted rugose  21/S 15/S 8/R 19/S 18/I 26/S 23/S 20/S  Adapted smooth  19/S 16/S 8/R 21/S 18/I 28/S 23/S 20/S  Control  21/S 16/S 10/R 21/S 18/I 28/S 23/S 20/S  aSXT, sulphamethoxazole/trimethoprim (25 μg), GN, gentamicin (10 μg), S, streptomycin (10 μg), AMC, amoxicillin/clavulanic acid (30 μg), NA, nalidixic acid (30 μg), CIP, ciprofloxacin (5 μg), CTX, ceftriaxone (30 μg), AMP, ampicillin (10 μg). Susceptibility zones of inhibition (millimeters) are reported as S, susceptible strains; I, intermediate susceptible strains; R, resistance strains. Boldfaced data indicate reduced susceptibility relative to unexposed (control) strains; data not bolded indicate exposed strains with no difference in susceptibility patterns relative to unexposed (control) strains. An increase in resistance was defined as a change in S (before chlorine exposure) to R (after chlorine exposure). bFor adaptation, cultures were previously exposed to increasing sublethal chlorine concentrations, control represent unexposed Salmonella culture. Data represent the average of 3 replicates. View Large Table 3. Broth microdilution assay of antibiotic cross-resistance patterns of adapted and control S. Heidelberg. Salmonella Heidelberg cultures previously adapted to chlorineb  Antibiotica (μg/ml) AMP GN S NA T CIP AMX  Adapted rugose  4/S 4/S 64 16/S 8/I 1/S 16/I  Adapted smooth  4/S 4/S 32 8/S 8/I 1/S 8/S  Control  4/S 4/S 32 8/S 4/S 1/S 8/S  Salmonella Heidelberg cultures previously adapted to chlorineb  Antibiotica (μg/ml) AMP GN S NA T CIP AMX  Adapted rugose  4/S 4/S 64 16/S 8/I 1/S 16/I  Adapted smooth  4/S 4/S 32 8/S 8/I 1/S 8/S  Control  4/S 4/S 32 8/S 4/S 1/S 8/S  aAMP, ampicillin; GN, gentamicin; S, streptomycin; NA, nalidixic acid; T, tetracycline; CIP; ciprofloxacin; AMX, amoxicillin. Data are reported as S, susceptible strains; I, intermediate susceptible strains; R, resistance strains. Boldfaced data indicate reduced susceptibility relative to unexposed (control) strains; data not bolded indicate exposed strains with no difference in susceptibility patterns relative to unexposed (control) strains. An increase in resistance was defined as a change in S (before chlorine exposure) to R (after chlorine exposure). b For adaptation, cultures were previously exposed to increasing sublethal chlorine concentrations, control represent unexposed Salmonella culture. Data represent the average of 2 replicates. View Large DISCUSSION After exposure to increasing sublethal concentrations of chlorine, Salmonella Heidelberg demonstrated the ability to acquire resistance. The ability of the adapted cells to tolerate higher chlorine concentrations continued even after repeated storage on chlorine-free TSA plates, which suggests the stability of adapted cells to the homologous stress of chlorine. In the current study, the MIC for the chlorine adapted (rugose and smooth) cells was 1.2 and 1.1 times higher than non-adapted control. This increase in MIC observed for chlorine-stressed Salmonella Heidelberg was in line with a previous study using Salmonella Typhimurium (Obe et al., 2016) and other studies on Salmonella enterica strains that exhibited adaptation to sublethal stress posed by different antimicrobials (Braoudaki and Hilton, 2004; Kim and Day, 2007; Alonso-Hernando et al., 2009; Stanojevic et al., 2010; Molina-Gonzalez et al., 2014). The inappropriate use of antimicrobials either directly or indirectly in the food processing establishment can expose any bacteria present to sublethal doses of disinfectants and sanitizers that are supposed to reduce the growth of bacteria (Capita et al., 2014). Another important finding from this study is the ability of the chlorine-adapted Salmonella Heidelberg to grow in the presence of high chlorine concentrations even above the concentration approved by the USDA for sanitation purposes on food-contact surfaces (200 ppm). The increase in bacteria tolerance following sublethal antimicrobial concentrations observed in this study have been well reported by others (Braoudaki and Hilton, 2004; Sheridan et al., 2012). Additionally, sublethal doses of antimicrobials have been observed to induce resistance in foodborne pathogens (Braoudaki and Hilton, 2005; Alonso-Calleja et al., 2015). The bacterial cells respond to stressful conditions either by adaptation or elimination, a common means that bacteria use to adjust to strenuous environmental conditions is by changing their morphology (Foster, 2005; Young, 2007). Other authors have reported these changes as a survival mechanism for bacteria (Justice et al., 2004; Young, 2007; Capital et al., 2014). In this study, Salmonella Heidelberg undergoes a morphological change to the rugose morphotype in order to cope with the stress posed by increasing sublethal chlorine concentrations. The change in morphology of the smooth Salmonella Heidelberg to the rugose variant in TSB was observed through the formation of a pellicle that is composed of cell aggregates, thereby allowing the rugose cell to remain as an aggregate in solution. Similar changes have been reported for Vibrio cholerae O1 E1 Tor and Salmonella Typhimurium DT104 (Morris et al., 1996; Anriany et al., 2001). The rugose variant was reported to be more virulent and not easy to kill compared to the smooth variant (Rice et al., 1993; Morris et al., 1996). In addition, studies have reported similar findings on the ability of some microorganisms including E. coli, Listeria monocytogenes, and Pseudomonas aeruginosa to tolerate different environmental stress. These microorganisms are reported to survive by changing their morphology through degradation of the cell wall, elongation of cells, aggregation of damaged bacteria cells and disturbances during cell division (To et al., 2002; Shalamanov, 2005; Giotis et al., 2009; Capita et al., 2014). The conditions at which rugose developed in this study is by daily transfer of an aliquot of 100 μL of culture containing sublethal chlorine concentration. This is different from the observations of other authors on the formation of the rugose variant in both Vibrio and Salmonella. In Salmonella Typhimurium DT104, the rugose variant was observed after 4 days of storage on TSA at room temperature and in Vibrio cholerae O1 strain TSI-4; the rugose variant was observed 2 months post inoculation when the smooth variant was re-cultured under a starved conditions at 16°C (Wai et al., 1998; Anriany et al., 2001). Whereas in the current study, the rugose variant was observed after exposure to increasing sublethal chlorine concentration (200 ppm) but failed to develop when cultured at room temperature with or without exposure to chlorine under similar experimental conditions. Another important finding was the ability of the rugose variant in Salmonella Typhimurium ATCC14028 (Obe et al., 2016) and currently in Salmonella Heidelberg ATCC8326 to retain their morphology even after storage on chlorine-free TSA. The adhesion and subsequent attachment of foodborne pathogens to food processing equipment and environment can result in a major food safety challenge when the pathogen contaminate food products thus causing a foodborne outbreak (Dourou et al., 2011). The attachment of Salmonella Heidelberg to different food-contact surfaces was examined in this study. The ability of the chlorine-adapted and control cells to adhere to both plastic and stainless steel surface was assessed at room temperature and 37°C. The results demonstrates that Salmonella Heidelberg possess the ability to attach to both surfaces tested. It was previously reported that Salmonella has the ability to colonize and attach to different surfaces including plastic, rubber, stainless steel, and glass (Helke et al., 1993; Sinde and Carballo, 2000; Nguyen et al., 2014; Yang et al., 2016). The rugose cells attached and formed stronger biofilms at both temperatures tested and showed no preference to a particular surface. The expression of rugose in Salmonella was previously reported to be due to aggregation of cells and formation of exopolysaccharides (EPS), which might aid in the strong attachment to food-contact surfaces (Marshall, 1992; Morris et al., 1996; Wai et al., 1998; Yildiz and Schoolnik, 1998). The smooth cells both adapted and control seems to attach better on the steel surface. The properties of the surface that Salmonella cells attached to, helps with their survival on such surface (O’Leary et al., 2013). Although Salmonella was reported to attach strongly to a hydrophobic surface, the ability to attach to a particular surface has been reported to be dependent on the strain of Salmonella (Sinde and Carballo. 2000; Chia et al., 2009). Nguyen et al. (2014), reported that Salmonella Typhimurium possess the ability to attach to both stainless steel and acrylic surface but showed greater attachment on stainless steel surfaces. This is because stainless steel is hydrophilic and bacteria attach more to such surfaces as compared to hydrophobic surface (Mafu et al., 1990; Sinde and Carballo. 2000). The results from this study shows that chlorine-adapted Salmonella Heidelberg exhibited no preference to a particular temperature. Salmonella generally grow well and has been observed to form biofilm at 37°C (Nguyen and Yuk, 2013). Room temperature was used in this study to examine the ability of Salmonella to form biofilm when exposed to an unfavorable condition such as temperature abuse. In addition, the adapted cells attached well on both surfaces tested in this study. Plastic and stainless steel surface was used because they are the most common surfaces encountered in food processing (Chmielewski and Frank, 2003; Ismail et al., 2013). Residues of food processing left on these surfaces can contribute to the formation of film if they are not promptly removed during cleaning (Joseph et al., 2001; Chmielewski and Frank, 2003). The application of effective sanitizers on food-contact surfaces following cleaning is important to inactivate and prevent the development of resistance in any pathogenic bacteria present. This will help prevent the acquisition of cross-protection to any other stress conditions encountered in processing The chlorine-adapted cells were examined for their ability to exhibit cross-adaptation to antibiotics. From the results of this study, it appears there were some interactions between adaptations to chlorine and cross-adaption to antibiotics. Salmonella Heidelberg cells that were adapted to sublethal concentrations of chlorine exhibited a certain degree of reduced susceptibility to some of the antibiotics tested. For the adapted cells, a reduced zone of inhibition was observed when compared to non-adapted (control), but the reduction was not significant enough to move above the limit set by the CLSI (2012) guidelines on antibiotic susceptibility testing for the “susceptible” category. Other authors have reported similar observations, Molina-Gonzalez et al. (2014), observed some Salmonella enterica strains that were previously exposed to various antimicrobials including sodium hypochlorite exhibited a lower zone of inhibition when compared to non-exposed strains, but are still susceptible to the antibiotics tested. In another study on the cross-adaption patterns of Salmonella enterica, the authors suggested that a slight reduction in susceptibility is noteworthy; this is because the pathogen may not be inhibited in the presence of the antibiotic over time (Braoudaki and Hilton, 2004). Chlorine-adapted rugose showed a slight reduction in the zone of inhibition to fluoroquinolones (CIP), quinolones (NA), aminoglycosides (S, GN), penicillin (AMX/AMC), and tetracycline (T). In addition, the MIC for the rugose variants increased and moved from the “susceptible” (i.e., bacterial infection will most likely respond to antibiotic treatment) category to the “intermediate” (bacterial infection may or may not respond to antibiotic treatment) category when tested against amoxicillin. A similar trend has been observed in Salmonella Typhimurium (Obe et al., 2016). Other studies have reported similar changes in antibiotic susceptibility patterns of previously adapted foodborne pathogens to different classes of antibiotics (Suller and Russell, 2000; Braoudaki and Hilton, 2004; Capita et al., 2014). Some of these studies suggested that the adaptive nature of foodborne pathogens to antimicrobials like chlorine is directly associated with a broad spectrum mechanism of resistance which includes the presence of an active efflux and alterations to cell permeability. This mechanism makes it difficult for different chemical molecules to enter the adapted cells (Tattawasart et al., 1999; Suller and Russell, 2000; Braoudaki and Hilton, 2004; Capita and Alonso-Calleja, 2013; Molina-Gonzalez et al., 2014). Even though, some authors have reported that sublethal dosage of antimicrobials could select for resistance to antibiotics in Salmonella, some studies do not observe similar findings. The results of this study in part agree with those authors that do not observe a change in susceptibility to antibiotics after exposure to sublethal concentrations of antimicrobials (Thomas et al., 2000; Ledder et al., 2006). In summary, the findings of the current study supports the hypothesis that Salmonella Heidelberg would adapt to sublethal concentrations of chlorine. However, the adaptive tolerance to chlorine observed in this study resulted in the formation of a more virulent Salmonella variant. The adapted cells were better biofilm formers on both food-contact surfaces tested and exhibited a slight reduction in zones of inhibition to different classes of antibiotics. This shows that chlorine stressed Salmonella may not be easily inactivated with high concentrations of chlorine. From this study, it is speculated that the possession of similar broad mechanisms of adaptation may eventually enable the adapted cells to become resistant to certain antibiotic treatment. The findings in this study signify a possible challenge to food safety and suggest that the misuse of the antimicrobial agent at a sublethal concentration could represent a potential public health risk. REFERENCES Alonso-Calleja C., Guerrero-Ramos E., Alonso-Hernando A., Capita R.. 2015. Adaptation and cross-adaptation of Escherichia coli ATCC 12806 to several food-grade biocides. Food Control . 56: 86– 94. Google Scholar CrossRef Search ADS   Alonso-Hernando A., Capita R., Prieto M., Alonso-Calleja C.. 2009. Comparison of antibiotic resistance patterns in Listeria monocytogenes and Salmonella enterica strains pre-exposed and exposed to poultry decontaminants. Food Control . 20: 1108– 1111. Google Scholar CrossRef Search ADS   Anriany Y. A., Weiner R. M., Johnson J. A., De Rezende C. E., Joseph S. W.. 2001. Salmonella enterica serovar Typhimurium DT104 displays a rugose phenotype. Appl. Environ. Microbiol . 67: 4048– 4056. Google Scholar CrossRef Search ADS PubMed  Bae Y. S., Baek, Lee S.. 2012. Resistance of pathogenic bacteria on the surface of stainless steel depending on attachment form and efficacy of chemical sanitizers. Int. J. Food Microbol.  153: 465– 473. Google Scholar CrossRef Search ADS   Barnes L. M., Lo M. F., Adams M. R., Chamberlain A. H. L. 1999. Effect of milk proteins on adhesion of bacteria to stainless steel surfaces. Appl. Env. Microbiol.  65: 4543– 4548. Braoudaki M., Hilton A. C.. 2004. Adaptive resistance to biocides in Salmonella enterica and Escherichia coli O157 and cross-resistance to antimicrobial agents. J. Clin. Microbiol.  42: 73– 78. Google Scholar CrossRef Search ADS PubMed  Braoudaki M., Hilton A. C.. 2005. Mechanism of resistance in Salmonella enterica adapted to erythromycin, benzalkonium chloride and triclosan. Int. J. Anti. Agents.  25: 31– 37. Google Scholar CrossRef Search ADS   Buchmeier N. A., Heffron F.. 1990. Induction of Salmonella stress proteins upon infection of macrophages. Science . 248: 730– 732. Google Scholar CrossRef Search ADS PubMed  Capita R., Alonso-Calleja C.. 2013. Antibiotic-resistant bacteria: a challenge for the food industry. Crit. Rev. Food Sci. Nutr.  53: 11– 48. Google Scholar CrossRef Search ADS PubMed  Capita R., Riesco-Pelaez F., Alonso-Hernando A., Alonso-Calleja C.. 2014. Exposure of Escherichia coli ATCC 12806 to sublethal concentrations of food-grade biocides influences its ability to form biofilm, resistance to antimicrobials, and ultrastructure. Appl. Environ. Microbiol.  80: 1268– 1280. Google Scholar CrossRef Search ADS PubMed  CDC. (Center for Disease Control and Prevention). 2011a. Estimates findings: estimates of foodborne illnesses in the United States [online]. Accessed Aug. 10, 2016. http://www.cdc.gov/foodborneburden/2011-foodborne-estimates.html. CDC. (Center for Disease Control and Prevention). 2011b. Burdens of Foodborne Illnesses: Findings [online]. Accessed Jan. 14, 2017. https://www.cdc.gov/foodborneburden/2011-foodborne-estimates.html. CDC. (Center for Disease Control and Prevention). 2014. Reports of selected Salmonella outbreak investigations [online]. Accessed Jan. 14, 2017. https://www.cdc.gov/salmonella/outbreaks.html. Chia T., Goulter R., McMeekin T., Dykes G., Fegan N.. 2009. Attachment of different Salmonella serovars to materials commonly used in a poultry processing plant. Food Microbiol . 26: 853– 859. Google Scholar CrossRef Search ADS PubMed  Chmielewski R. A. N., Frank J. F.. 2003. Biofilm formation and control in food processing facilities. Compr. Rev. Food Sci. Food Saf.  2: 22– 32. Google Scholar CrossRef Search ADS   CLSI. (Pennsylvania: National Committee for Clinical Laboratory Standards). 2008. Reference method for broth dilution antifungal susceptibility testing of filamentous fungi. Approved Standard – Second Edition. M38-A2. CLSI. (Pennsylvania: National Committee for Clinical Laboratory Standards). 2012. Performance standards for antimicrobial disk susceptibility test. Approved Standard – Twelfth Edition. M02-A12. Costerton J. W., Lewandowski Z., Caldwell D. E., Kober D. R., Lappin-Scott H. M.. 1995. Microbial biofilms. Ann. Rev. Microbiol.  49: 711– 745. Google Scholar CrossRef Search ADS   Czechowski M. H. 1990. Bacterial attachment to Buna-N gaskets in milk processing equipment. Australian J. Dairy Tech.  45: 113– 114. Davidson P. M., Harrison M. A.. 2002. Resistance and adaptation to food antimicrobials, sanitizers, and other process controls. Food Tech . 56: 69– 78. Dourou D., Beauchamp C. S., Yoon Y., Geornaras I., Belk K. E., Smith G. C., Nychas G. J., Sofos J. N.. 2011. Attachment and biofilm formation by Escherichia coli O157:H7 at different temperatures, on various food contact surfaces encountered in beef processing. Int. J. Food Microbiol.  149: 262– 268. Google Scholar CrossRef Search ADS PubMed  Foster P. L. 2005. Stress responses and genetic variation in bacteria. Mut. Res. Rev.  569: 3– 11. Google Scholar CrossRef Search ADS   Frank J. K., Koffi R. A.. 1990. Surface-adherent growth of Listeria monocytogenes is associated with increased resistance to surfactant sanitizers and heat. J. Food Protect.  53: 550– 554. Google Scholar CrossRef Search ADS   Giaouris E., Chorianopoulos N., Nychas G. J. E.. 2005. Effect of temperature, pH, and water activity on biofilm formation Salmonella Enteritidis PT4 on stainless steel surfaces as indicated by the bead vortexing method and conductance measurements. J. Food Protect.  68: 2149– 2154. Google Scholar CrossRef Search ADS   Giotis E., Blair I. S., McDowell D. A.. 2009. Effects of short-term alkaline adaptation on surface properties of Listeria monocytogenes 10403S. The Open Food Sci. J.  3: 62– 65. Google Scholar CrossRef Search ADS   Helke D., Somers E., Wong A.. 1993. Attachment of Listeria monocytogenes and Salmonella Typhimurium to stainless steel and buna-N rubber in the presence of milk and individual milk components. J. Food Protect.  56: 479– 484. Google Scholar CrossRef Search ADS   Hood S. K., Zottola E. A.. 1997. Adherence to stainless steel by foodborne microorganisms during growth in model food systems. Int. J. Food Microbiol.  37: 145– 153. Google Scholar CrossRef Search ADS PubMed  Hur J. C., Jawale, Lee J. H.. 2012. Antimicrobial resistance of Salmonella isolated from food animals: review. Food Res. Int.  45: 819– 830. Google Scholar CrossRef Search ADS   IFT (Institute of Food Technologists). 2006. Antimicrobial resistance: Implications for the food systems. Compr. Rev. in Food Sci. Food Saf.  5: 71– 137. CrossRef Search ADS   Ismaïl R., Aviat F., Michel V., Le Bayon I., Gay-Perret P., Kutnik M., Fédérighi M.. 2013. Methods for recovering microorganisms from solid surfaces used in the food industry: a review of the literature. Int. J. Environ. Res. Pub. Health.  10: 6169– 6183. Google Scholar CrossRef Search ADS   Iturriaga M. H., Tramplin M. L., Escartin E. F.. 2007. Colonization of tomatoes by Salmonella Montevideo is affected by relative and storage temperature. J. Food Protect.  70: 30– 34. Google Scholar CrossRef Search ADS   Jiang X., Doyle M. P.. 1999. Fate of Escherichia coli O157:H7 and Salmonella Enteritidis on currency. J. Food Protect.  62: 805– 807. Google Scholar CrossRef Search ADS   Joseph B., Otta S., Karunasagar I., Karunasagar I.. 2001. Biofilm formation by Salmonella spp. on food contact surfaces and their sensitivity to sanitizers. Int. J. Food Microbiol.  64: 367– 372. Justice S. S., Hung C., Theriot J. A., Fletcher D. A., Anderson G. G., Footer M. J., Hultgren S. J.. 2004. Differentiation and developmental pathways of uropathogenic Escherichia coli in urinary tract pathogenesis. Proc. Natl. Acad. Sci.  101: 1333– 1338. Google Scholar CrossRef Search ADS   Kim D., Day D. F.. 2007. A biocidal combination capable of sanitizing raw chicken skin. Food Control . 18: 1272– 1276. Google Scholar CrossRef Search ADS   Krysinski E. P., Brown L. J., Marchisello T. J.. 1992. Effect of cleaners and sanitizers on Listeria monocytogenes attached to product contact surfaces. J. Food Protect.  55: 246– 251. Google Scholar CrossRef Search ADS   Kusumaningrum H. D., Riboldi G., Hazeleger W. C., Beumer R. R.. 2003. Survival of foodborne pathogens on stainless steel surfaces and cross-contamination to foods. Int. J. Food Microbiol . 85: 227– 236. Google Scholar CrossRef Search ADS PubMed  Ledder R. G., Gilbert P., Wilis C., McBain A. J.. 2006. Effects of chronic triclosan exposure upon the antimicrobial susceptibility of 40 ex-situ environmental and human isolates. J. Appl. Microbiol.  100: 1132– 1140. Google Scholar CrossRef Search ADS PubMed  Lou Y., Yousef A. E.. 1996. Resistance of Listeria monocytogenes to heat after adaptation to environmental stresses. J. Food Protect.  5: 465– 471. Google Scholar CrossRef Search ADS   Mafu A. A., Roy D., Goulet J., Magny P.. 1990. Attachment of Listeria monocytogenes to stainless steel, glass, polypropylene and rubber surfaces after short contact times. J. Food Protect . 53: 742– 746. Google Scholar CrossRef Search ADS   Marshall K. C. 1992. Biofilms: an overview of bacterial adhesion, activity, and control at surfaces. ASM News . 58: 202– 207. Mead P. S., Slutsker L., Dietz V., McCaig L. F., Bresee J. S., Shapiro C., Griffin P. M., Tauxe R. V.. 1999. Food-related illness and death in the United States. Emerg. Infect. Dis . 5: 607– 625. Google Scholar CrossRef Search ADS PubMed  Molina-Gonzalez D., Alonso-Calleja C., Alonso-Hernando A.. 2014. Effect of sublethal concentrations of biocides on the susceptibility to antibiotics of multi-drug resistance Salmonella enterica strains. Food Control . 40: 329– 334. Google Scholar CrossRef Search ADS   Morris G. K., Wells J. G.. 1970. Salmonella contamination in a poultry-processing plant. Appl. Microbiol . 38: 2465– 2467. Morris J. G. Jr, Sztein M. B., Rice E. W., Nataro J. P., Losonsky G. L., Panigrahi P., Tacket C. O., Johnson J. A.. 1996. Vibrio cholerae O1 can assume a chlorine-resistant rugose survival form that is virulent for humans. The J. Infect. Dis . 174: 1364– 1368. Google Scholar CrossRef Search ADS   Nguyen H. D. N., Yuk H. G.. 2013. Changes in resistance of Salmonella Typhimurium biofilms formed under various conditions to industrial sanitizers. Food Control . 29: 236– 240. Google Scholar CrossRef Search ADS   Nguyen H. D. N., Yang Y. S., Yuk H. G.. 2014. Biofilm formation of Salmonella Typhimurium on stainless steel and acrylic surfaces as affected by temperature and pH level. LWT – Food Sci Technol . 55: 283– 288. Google Scholar CrossRef Search ADS   O’Leary D., Mc Cabe E. M., McCusker M. P., Martin M., Fanning S., Duffy G.. 2013. Microbiological study of biofilm formation in isolates of Salmonella enterica Typhimurium DT104 and DT104b cultured from the modern pork chain. Int. J. Food Microbiol.  161: 36– 43. Google Scholar CrossRef Search ADS PubMed  Obe T., Nannapaneni R., Sharma C. S.. 2016. Development of rugose morphotype of Salmonella Typhimurium following exposure to sub-inhibitory chlorine concentrations that exhibit chlorine resistance and strong biofilm forming ability. Poult. Sci.  95( E – Suppl. 1): 35. Painter J. A., Hoekstra R. M., Ayers T., Tauxe R. V., Braden C. R., Angulo F. J., Griffin P. M.. 2013. Attribution of foodborne illnesses, hospitalizations, and deaths to food commodities by using outbreak data, United States, 1998–2008. Emerg. Infect. Dis . 19: 407– 415. Google Scholar CrossRef Search ADS PubMed  Patel J., Sharma M.. 2010. Differences in attachment of Salmonella enterica serovars to cabbage and lettuce leaves. Int. J. Food Microbiol.  139: 41– 47. Google Scholar CrossRef Search ADS PubMed  Rice E. W., Johnson C. H., Clark R. M., Fox K. R., Reasoner D. J., Dunnigan M. E.. 1993. Vibrio cholerae O1 can assume a ‘rugose’ survival form that resists killing by chlorine, yet retains virulence. Int. J. Env. Health Res . 3: 89– 98. Google Scholar CrossRef Search ADS   Scallan E., Hoekstra R. M., Angulo F. J., Tauxe R. V., Widdowson M. A., Jones S. L., Griffin P. M.. 2011. Foodborne illnesses acquired in the United States- major pathogen. Emerg. Infect. Dis.  17: 7– 15. Google Scholar CrossRef Search ADS PubMed  Scott E., Bloomfield S. F.. 1990. The survival and transfer of microbial contamination via cloths, hands and utensils. The J. Appl. Bacteriol.  68: 271– 278. Google Scholar CrossRef Search ADS   Shalamanov D. S. 2005. Chlorhexidine gluconate-induced morphological changes in gram negative microorganisms. Biotechnol. Biotechnol. Eq.  19: 121– 124. Google Scholar CrossRef Search ADS   Sheridan A., Lenahan M., Duffy G., Fanning S., Burgess C.. 2012. The potential for biocide tolerance in Escherichia coli and its impact on the response to food processing stresses. Food Control . 26: 98– 106. Google Scholar CrossRef Search ADS   Sinde E., Carballo J.. 2000. Attachment of Salmonella spp. and Listeria monocytogenes to stainless steel, rubber and polytetrafluoroethylene: the influence of free energy and the effect of commercial sanitizers. Food Microbiol.  17: 439– 447. Google Scholar CrossRef Search ADS   Stanojevic D., Comic L., Stefanovic O., Solujic-Sudolak S.. 2010. In vitro synergistic antibacterial activity of Salvia officinalis L. and some preservatives. Arch. Biol. Sci.  62: 175– 183. Google Scholar CrossRef Search ADS   Suller M. T. E., Russell A. D.. 2000. Triclosan and antibiotic resistance in Staphylococcus aureus. J. Antimicrob. Chemother.  46: 11– 18. Google Scholar CrossRef Search ADS PubMed  Tattawasart U., Maillard J. Y., Furr J. R., Russell A. D.. 1999. Development of resistance to chlorhexidine diacetate and cetylpyridinium chloride in Pseudomonas stuzeri and changes in antibiotic susceptibility. J. Hos. Infect.  42: 219– 229. Google Scholar CrossRef Search ADS   Thomas L., Maillard J. Y., Lambert R. J. W., Russell A. D.. 2000. Development of resistance to chlorhexidine diacetate in Pseudomonas aeruginosa and the effect of a “residual” concentration. J. Hos. Infect.  46: 297– 303. Google Scholar CrossRef Search ADS   To M. S., Favrin S., Romanova N., Griffiths M. W.. 2002. Post-adaptational resistance to benzalkonium chloride and subsequent physicochemical modifications of Listeria Monocytogenes. Appl. Environ. Microbiol.  68: 5258– 5264. Google Scholar CrossRef Search ADS PubMed  USDA-FSIS. (United States Department of Agriculture, Food Safety Inspection Service). 2015. Safe and suitable ingredients in the production of meat, poultry, and egg products. FSIS Directive 7120.1 Revision 36 [online]. Accessed Aug. 2, 2016. http://www.fsis.usda.gov/wps/wcm/connect/bab10e09-aefa-483b-8be8809a1f051d4c/7120.1.pdf?MOD=AJPERES.. Wai S. N., Mizunoe Y., Takade A., Kawabata S. I., Yoshida S. I.. 1998. Vibrio cholerae O1 strain TSI-4 produces the exopolysaccharide materials that determine colony morphology, stress resistance, and biofilm formation. Appl. Environ. Microbiol . 64: 3648– 3655. Google Scholar PubMed  Yang Y., Miks-Krajnik M., Zheng Q., Lee S. B., Lee S. C., Yuk H. G.. 2016. Biofilm formation of Salmonella Enteritidis under food-related environmental stress conditions and its subsequent resistance to chlorine treatment. Food Microbiol . 54: 98– 105. Google Scholar CrossRef Search ADS   Yildiz F. H., Schoolnick G. K.. 1998. Role of rpoS in stress survival and virulence of Vibrio cholerae. J. Bacteriol.  180: 773– 784. Google Scholar PubMed  Young D. 2007. Bacterial morphology: why have different shapes? Curr. Opin. Microbiol.  10: 596– 600. Google Scholar CrossRef Search ADS PubMed  © 2018 Poultry Science Association Inc. http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Poultry Science Oxford University Press

Homologous stress adaptation, antibiotic resistance, and biofilm forming ability of Salmonella enterica serovar Heidelberg ATCC8326 on different food-contact surfaces following exposure to sublethal chlorine concentrations1

Loading next page...
 
/lp/ou_press/homologous-stress-adaptation-antibiotic-resistance-and-biofilm-forming-JqCZ0Pa0br
Publisher
Oxford University Press
Copyright
© 2018 Poultry Science Association Inc.
ISSN
0032-5791
eISSN
1525-3171
D.O.I.
10.3382/ps/pex346
Publisher site
See Article on Publisher Site

Abstract

Abstract Salmonella enterica serovar Heidelberg (American Type Culture Collection; ATCC 8326) was examined for the ability to adapt to the homologous stress of chlorine through exposure to increasing chlorine concentrations (25 ppm daily increments) in tryptic soy broth (TSB). The tested strain exhibited an acquired tolerance to chlorine in TSB with the tolerant cells growing in concentrations up to 400 ppm. In addition, the chlorine stressed cells displayed rugose morphology on tryptic soy agar (TSA) plates at 37°C. The minimum inhibitory concentration (MIC) of chlorine for adapted (rugose and smooth) cells was determined to be 550 ppm and 500 ppm, respectively whereas the MIC for the control was 450 ppm. The biofilm forming ability of the adapted and control cells were examined on both plastic and stainless steel surface at room temperature and 37°C. The rugose variant, in contrast to the smooth (adapted and control) showed the ability to form strong biofilms (P ≤ 0.05) on a plastic surface at room temperature and 37°C. Rugose cells compared to smooth and control attached more (P ≤ 0.05) to steel surfaces as well. The possibility of cross-adaptation was examined by exposing the adapted and control cells to different antibiotics according to the Clinical & Laboratory Standards Institute guidelines. Adapted cells exhibited reduced susceptibility to some of the antibiotics tested as compared to control. The findings of this study suggest that exposure to sublethal chlorine concentration during the sanitization procedure can result in tolerant Salmonella cells. Chlorine may confer cross-protection that aids in the survival of the tolerant population to other environmental stresses. INTRODUCTION Salmonella enterica is well known as the leading cause of foodborne infections globally (Painter et al., 2013). The serotypes of Salmonella enterica are reported to result in over a million cases of illnesses, thousands of hospitalizations, and a few hundred deaths each year in the United States and also around the world (Mead et al., 1999; Scallan et al., 2011; CDC, 2011a). According to an estimate by the Centers for Disease Control and Prevention (CDC) in 2011, nontyphoidal Salmonella, which includes all Salmonella serotypes except Typhi and Paratyphi, was among the top five pathogens in the United States causing the most reported foodborne illnesses, hospitalizations, and deaths annually (CDC, 2011b). In addition, nontyphoidal Salmonella, which is second to norovirus, is estimated to cause 11% illnesses, 35% hospitalizations, and 28% deaths each year in the United States alone (CDC, 2011b). Moreover, of all the Salmonella enterica serotypes, the top four serotypes associated with animal products and most often reported to cause human infections are Typhimurium, Enteritidis, Newport, and Heidelberg (Hur et al., 2012). There are several food commodities implicated in foodborne illnesses, however, between 1998 and 2008, poultry accounted for 9.8% of the illnesses, 11.5% of the hospitalizations, and 19.1% of the deaths (Painter et al., 2013). Poultry is considered a major source of Salmonella contamination that results in human illness (Morris and Wells, 1970). In 2013, a multistate outbreak of multidrug-resistant Salmonella Heidelberg was linked to branded chicken. This outbreak was reported to result in 634 cases of illnesses and 38% hospitalizations (CDC, 2014). Due to the burdens attached to the control of foodborne pathogens, poultry processors employ many methods to prevent pathogens from contaminating poultry meat. One of the methods include the use of USDA approved antimicrobials for disinfecting poultry meat and sanitizing food-contact surfaces within the poultry processing facility (USDA-FSIS, 2015). The antimicrobials are capable of destroying both spoilage and pathogenic bacteria. Examples of antimicrobials used to disinfect, decontaminate, or preserve food products include peracetic acid, calcium hypochlorite, sodium nitrite, trisodium phosphate, and sodium hypochlorite (Capita and Alonso-Calleja, 2013; Alonso-Calleja et al., 2015). The misuse of these compounds has allowed pathogenic bacteria to develop adaptation (Capita et al., 2014). Previous studies have shown that using an antimicrobial at a sublethal concentration may potentially aid in the survival of foodborne pathogens by enabling them to survive challenging conditions such as antimicrobial inactivation (Sheridan et al., 2012; Capita et al., 2014). This increase in bacteria tolerance to antimicrobials is a global public health concern (Braoudaki and Hilton, 2004) Several studies have documented the ability of bacteria to adapt to stress posed by antimicrobials, which they use to induce cross-protection against other stressful conditions (Davidson and Harrison, 2002; Braoudaki and Hilton, 2005; Capita et al., 2014). The majority of these studies provides significant evidence that reduced susceptibility to one antimicrobial can cause cross-adaptation to another (Braoudaki and Hilton, 2005). Since chlorine is the antimicrobial agent that is most common in sanitizing solutions, adaptation to chlorine could constitute a potential threat to food safety by inducing cross-protection to antibiotics that are clinically important (Braoudaki and Hilton, 2004; Molina-Gonzalez et al., 2014). Antibiotic resistance has continued to be a global public health issue; research has been conducted that examines different mechanisms used by pathogens, such as Salmonella, Listeria, and E.coli, to adapt to disinfectants and induce cross-resistance to antibiotics (Buchmeier and Heffron, 1990; Lou and Yousef, 1996; Braoudaki and Hilton, 2004; Capita et al., 2014; Molina-Gonzalez et al., 2014). Some of this research demonstrates that an adapted bacteria will most likely use similar resistance mechanism to offer cross-protection to antibiotics (Braoudaki and Hilton, 2004). In addition, it was reported that a frequent strategy used by pathogenic microorganisms to adapt to antimicrobials, is the formation of biofilm (IFT, 2006). Biofilms are bacteria that cluster together on a surface and are surrounded by an exopolymeric matrix that makes the biofilm matrix impenetrable to any harmful substance (Costerton et al., 1995; Iturriaga et al., 2007; Bae et al., 2012). An important challenge faced by food processors is the control of bacteria present in the biofilm. It was documented that biofilms are more difficult to destroy because they possess tolerance to environmental stresses when compared to free cells (Frank and Koffi, 1990; Joseph et al., 2001). In poultry processing, attachment of bacteria to surfaces of equipment can be a source of product contamination (Barnes et al., 1999; Giaouris et al., 2005). Some studies have reported the incidence of cross-contamination due to bacterial attachment to equipment surfaces such as utensils used in food preparation (Scott and Bloomfield, 1990; Jiang and Doyle, 1999; Kusumaningrum et al., 2003). Other studies suggested that bacteria could attach to surfaces such as polystyrene, glass, rubber, acrylic, and cement (Czechowski, 1990; Mafu et al., 1990; Krysinski et al., 1992; Barnes et al., 1999; Nguyen and Yuk, 2013). Therefore, in this study, the homologous stress adaptation of Salmonella Heidelberg exposed to chlorine at a sublethal concentration was determined by observing changes in minimum inhibitory concentration (MIC). The biofilm formation of the adapted cells compared to non-adapted cells on different food-contact surfaces was also measured. Antibiotic susceptibility patterns of adapted cells were further investigated. MATERIALS AND METHODS Salmonella Heidelberg Strain and Culture Preparation Salmonella Heidelberg (American Type Culture Collection; ATCC 8326) was maintained on tryptic soy broth (TSB; Sigma-Aldrich Co., St. Louis, MO). Prior to experiments, frozen cells were streaked on tryptic soy agar plates (TSA; Sigma-Aldrich Co.) and incubated in a refrigerated incubator (PR505755R, Thermo Fisher Scientific, Waltham, MA) at 37°C for 24 h. A single colony was subcultured in TSB at 37°C for 20 to 24 h. Working cultures were stored at 4 ± 1°C on TSA slants and were subcultured monthly. Chlorine Source Sodium hypochlorite that contained 5% available chlorine (AC419552500, ACROS Organics, Morristown, NJ), was used as the source of chlorine in this study. The amount of free chlorine in the sodium hypochlorite was confirmed using the HACH (chlorine test kit) Pocket Colorimeter (No. 5870023, HACH Company, Loveland, CO) in accordance with the manufacturer's instructions. A sterile solution of chlorine was prepared in an appropriate media before each experiment. Determination of Minimum Inhibitory Concentration (MIC) of chlorine The MIC value was established using the broth macro and microdilution method in accordance with the Clinical & Laboratory Standards Institute guidelines (CLSI, 2008) with minimal modifications. One colony of the planktonic cells, obtained from a TSA plate was inoculated in 10 mL TSB, and incubated at 37°C for 20 to 24 h. It was determined in our laboratory that 20 to 24 h incubation of the bacterial culture contained approximate 1 × 109 CFU/mL. To start the experiment, 96-well microtiter plates (SKU 229190, Celltreat Scientific Product, Pepperell, MA) were used. A volume of 100 μL of chlorine in TSB that contained double the desired concentration was added to each well of the plate. Afterwards, 100 μL of inoculum prepared to a final concentration of ∼ 106 CFU/mL was added to make the final volume in each well 200 μL. A positive control that contained (100 μL TSB + 100 μL inoculum devoid of chlorine) and a negative control (200 μL TSB) were included in the experiment. The microwell plate was incubated at 37°C for 24 h and visible growth of bacteria in each well was determined by turbidity. The MIC was established as the lowest chlorine concentration that is necessary to inhibit the growth of Salmonella Heidelberg after 24 h of incubation. The exact MIC was determined by narrowing the range of chlorine concentrations in each well for the subsequent experiment. Stress Adaptation Study This study was performed by preparing Salmonella stock culture to a final concentration of ∼109 CFU/mL. The starting concentration of chlorine was 125 ppm, an aliquot of 100 μL of the inoculum was transferred to a polypropylene tube (Cat No. S50712, Fisher Scientific, Fair Lawn, NJ) that contained 9.9 mL (TSB + chlorine). After the transfer, the tube contained a final inoculum concentration of ∼ 107 CFU/mL. Upon incubation for 24 h at 37°C, turbidity in the tube was used to examine bacteria growth. The suspension was then diluted and plated on TSA plates to enumerate Salmonella growth. An additional aliquot of 100 μL was transferred from the same turbid tube to a sterile tube that contained 9.9 mL (TSB + higher sublethal chlorine concentration). This procedure was continued by increasing the concentration of chlorine by 25 ppm daily until no visible growth was observed in the tube. This required 12 days of incubation. The suspension in the last tube with visible bacteria growth was again diluted for Salmonella enumeration. The cells obtained on TSA plates after incubation for 24 h at 37°C were re-streaked and stored on TSA plate that did not contain chlorine. The cells were considered adapted Salmonella Heidelberg cells. The TSA plates containing the adapted cells were stored at 4 ± 1°C with the cells being transferred weekly to fresh non-chlorinated TSA plates. Adaptive Stability to Chlorine The adaptive tolerance to chlorine of the adapted Salmonella Heidelberg cells was tested by determining the MIC for the adapted cells against chlorine using the broth macro dilution method according to the CLSI guidelines (CLSI, 2012).This was done after storage on TSA plate that did not contain chlorine. A colony of the adapted cell was inoculated into a borosilicate round bottom glass culture tube (No. 1496128, Fisher Scientific) containing 10 mL TSB with chlorine at concentrations that were below, equivalent and above the MIC. Non-adapted Salmonella Heidelberg cells, which served as the control, were also tested at the same concentrations. Turbidity was used to observe bacteria growth, the glass culture tube showing no visible growth after incubation at 37°C for 24 h was considered the MIC of the adapted and control cells. Biofilm Formation on plastic surface The standard crystal violet assay as previously described by Patel and Sharma, (2010) was used to determine the biofilm forming ability of the chlorine adapted (rugose and smooth) and control cells. An overnight culture (∼ 20 h) of adapted and control Salmonella cells grown in TSB were diluted to achieve a final inoculum level of ∼ 106 CFU/mL. The treated cells (rugose and smooth) were cultured in TSB that contained 400 ppm chlorine which is the highest concentration that supports bacteria growth, whereas the control cells were cultured in chlorine-free TSB. To begin the experiment, 96-well polystyrene cell culture plates were prepared by dispensing 200 μL of the inoculum into duplicate wells for each type of adapted Salmonella Heidelberg morphotypes and the control. TSB devoid of inoculum was used as a negative control. The plates were incubated at room temperature and 37°C for 48 h. After incubation was complete, 200 μL of the inoculum was aspirated from each well of the plate, and each well was washed five times with sterile distilled water so that all loosely attached cells may be removed. The plates were later air-dried for 45 mins, and 200 μL of crystal violet solution (0.41% w/v dye, AC447570500, ACROS Organics) was dispensed into each well. The plates were incubated at room temperature for an additional 45 mins, after which the dye was aspirated from each well, and the well was washed five times using sterile distilled water. Each well was allowed another 45 mins to air dry, and then 200 μL of 95% ethanol (A406P4, Fisher Scientific) was added to each well. The content of each well was mixed to dissolve the crystal violet dye. Biofilm formation in each well was measured by an optical density (OD600) reading using a micro quant microplate spectrophotometer (Model ELx800, BioTek Instruments, Winooski, VT). The attached cells of adapted (rugose and smooth) and control on the plastic plate were enumerated. Similar to the crystal violet assay, 200 μL of the inoculum prepared to ∼ 106 CFU/mL was dispensed into each well of a 24-well tissue culture plate (SKU 229123, Celltreat Scientific Product, Pepperell, MA) in duplicate. The plate were then incubated at room temperature and 37°C for 48 h. After incubation, each well was aspirated and washed three times with sterile distilled water to remove any loosely attached bacteria. The strongly attached cells were scraped into 0.1% peptone water, vortexed, and serially diluted. After dilution, an aliquot of 100 μL was plated on TSA plates, incubated at 37°C for 24 h in order to count the number of attached cells. Biofilm Formation on stainless steel (SS) surface To examine the influence of chlorine stress on the ability of Salmonella Heidelberg to attach to stainless steel (SS), a previously described method with minimal modifications was used (Hood and Zottola, 1997). Stainless steel scoops were purchased from Spring Chef (www.springchef.com, Dallas, TX). The SS scoops were washed with detergent upon arrival, sanitized in 70% ethanol and autoclaved at 121°C for 15 mins. A total of 2 mL of previously prepared inoculum (∼ 106 CFU/mL) of adapted and control Salmonella Heidelberg were dispensed into each 5 mL scoop. The scoops were placed in sterile Nalgene pans (Cat No. 1336110, Fisher Scientific) that were placed in a covered storage box (No. 552834742, Walmart Stores, Inc. Bentonville, AR) and incubated at both room temperature and 37°C for 48 h. After incubation, each scoop was aspirated and washed three times with sterile distilled water to remove any loosely attached bacteria cell. The strongly attached cells on the scoop were scraped into a solution of 0.1% peptone water and vortexed for 2 minutes before being subjected to a 10-fold dilution in TSB. An aliquot of 100 μL from each dilution was plated on TSA plates, incubated at 37°C for 24 h in order to quantify the number of attached cells. Determination of Antibiotic Susceptibility Chlorine-adapted Salmonella Heidelberg and control cells were screened against different antibiotics on Mueller-Hinton broth and Mueller-Hinton agar (MHB, MHA; Oxoid Co., Nepean, ON, Canada) to determine susceptibility. Adapted cells were grown in TSB containing chlorine (400 ppm) and the control cells were grown in TSB devoid of chlorine. The inoculum was prepared in MHB to a final concentration of ∼ 106 CFU/mL. MIC broth microdilution and disk diffusion method as described by the CLSI (2008) guideline with slight modifications were used in the study. The following antibiotic disks (Fisher Scientific) were used: sulphamethoxazole/trimethoprim (SXT, 25 μg), gentamicin (GN, 10 μg), streptomycin (S, 10 μg), amoxicillin/clavulanic acid (AMC, 30 μg), nalidixic acid (NA, 30 μg), ciprofloxacin (CIP, 5 μg), ceftriaxone (CTX, 30 μg), ampicillin (AMP, 10 μg). The inhibition zones were measured and recorded as susceptible, intermediate, or resistant according to the guidelines by CLSI (2008). Cultures of Escherichia coli 25922 with known antibiotic resistance patterns were used as control reference strain. The MIC for the antibiotics was determined using a 96-well cell culture plate. Each panel in the plate contained 5 dilutions using the MIC breakpoints provided by the CLSI (2008) guidelines for each antibiotic tested in this study. Positive control (MHB + inoculum) and negative control (MHB only) were maintained throughout the experiment. The plate was incubated at 37°C for 24 h, and the results were recorded as the least concentration of antibiotic that prevent the growth of bacteria either as susceptible, intermediate, or resistance as stated in the CLSI (2008) guidelines. Statistical Analysis Each Experiment was Evaluated Sequentially on Different Days All data analysis in this study was carried out using analysis of variance (ANOVA) in the General Linear Model (GLM) of SAS v. 9.4 (SAS Institute, Cary, NC; Steel and Torrie, 1980). Means separations were acquired using Fisher's least significant difference test. The treatments and controls were determined to be significant at the 5% (P ≤ 0.05) level. RESULTS Homologous Stress Adaptation to Chorine It was observed in previous research (Obe et al., 2016) that Salmonella Typhimurium undergoes a morphological change upon a prolong exposure to sublethal chlorine concentration. In the present study, Salmonella Heidelberg changes it morphology to the rugose variant after 4 days exposure to chlorine stress at 37°C. Rugose and smooth Salmonella Heidelberg continued to grow together on TSA plate until the chlorine concentration in TSB reached 400 ppm. That was after 12 days of incubation. This concentration was recorded as the maximum chlorine concentration that allowed the growth of stressed Salmonella Heidelberg and the cells recovered at this concentration was referred to as the chlorine-adapted Salmonella cells. The MIC of chlorine for Salmonella Heidelberg before sublethal chlorine exposure was determined to be approximately 400 ppm (Table 1). The adaptive tolerance to chlorine was later measured by sub-culturing the stressed cells in a nonselective broth (TSB) after storage on TSA plates devoid of chlorine. The stressed cells (rugose and smooth) were able to grow above the MIC values up to 500 ppm and 450 ppm respectively, compared to the non-exposed cell that could not grow above 400 ppm (Figure 1). This suggests that Salmonella Heidelberg ATCC 8326 possesses a stable homologous adaptation to chlorine. Figure 1. View largeDownload slide Minimum inhibitory concentration (MIC) for chlorine-adapted and control Salmonella Heidelberg after stress adaptation. Data represent the average of 2 replicates. The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Figure 1. View largeDownload slide Minimum inhibitory concentration (MIC) for chlorine-adapted and control Salmonella Heidelberg after stress adaptation. Data represent the average of 2 replicates. The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Table 1. Minimum inhibitory concentrations (MICs) and minimum bactericidal concentrations (MBCs) of chlorine (ppm) for Salmonella Heidelberga (ATCC8326) before adaptation. Dilution method  MIC (ppm)  MBC (ppm)  Micro  400  500  Macro  500  500  Dilution method  MIC (ppm)  MBC (ppm)  Micro  400  500  Macro  500  500  aSalmonella Heidelberg culture prior to chlorine exposure. Data represent the average of 3 replicates. View Large Biofilm Formation on Plastic Surface To determine whether adaptation to chlorine would influence the strength of biofilms formed, Salmonella Heidelberg was cultured in the presence of chlorine (400 ppm) for the adapted cells and in the absence of chlorine for the non-adapted (control) cells. The optical density reading (OD600) of the adapted and control cells were observed and used to determine biofilm forming ability of each of the Salmonella variants. In the present study, Salmonella Heidelberg formed biofilms on polystyrene plastic surface both at room temperature and at 37°C. At room temperature, there was a significant difference (P < 0.05) in the biofilms formed by the adapted cells versus control. Adapted rugose formed the strongest biofilm with OD600 value of 3.4, followed by adapted smooth with OD600 value 1.13 representing a moderate biofilm former, compared to control that formed the weakest biofilm with OD600 value of 0.68 (Figure 2A). At 37°C, similar to room temperature, the morphotypes tested were significantly different (P < 0.05) in their ability to form a biofilm. Adapted rugose showed a strong biofilm forming ability with an OD600 value of 3.4. There were no differences observed for smooth morphologies (both adapted and control) which showed OD600 values of 0.7 and 0.57, respectively (Figure 2B). Biofilm formation on the plastic surface was also determined by enumerating the strongly attached cells on TSA plates. The difference in log values was used to determine the difference in biofilm forming ability of chlorine-adapted cells against control. The result were reported in Log CFU/mL. At room temperature, there was no significant difference (P > 0.05) observed for adapted cells (rugose compared to smooth), and adapted smooth compared to control. However, adapted rugose has a significant higher attached cell concentration of 5.05 log cfu/mL when compared to control which is 4.85 log cfu/mL but the difference was not enough to establish a biological significance (Figure 3A). A significant difference (P < 0.05) was determined for the attachment of chlorine-adapted cells (rugose and smooth) having 5.25 and 4.9 log cfu/mL respectively compared to control, which has a cell concentration of 4.7 log cfu/mL at 37°C (Figure 3B). In addition, at 37°C, rugose appears to have more CFU/mL of attached cells on plastic surface compared to smooth (adapted and non-adapted) cells. Figure 2. View largeDownload slide (A) Biofilm formation by Salmonella Heidelberg after 48 h on plastic surface using 96-well polystyrene microtiter plate at room temperature. Means with different superscripts indicate significant differences in the biofilm forming ability of chlorine-adapted and control Salmonella Heidelberg morphotype. (P = 0.0001; SEM = 0.109; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, dark gray bar represents non-exposed positive control, and light gray bar represent (broth only) negative control. (B) Biofilm formation by Salmonella Heidelberg after 48 h on plastic surface using 96-well polystyrene microtiter plate at 37°C. Means with different superscripts indicate significant differences in the biofilm forming ability of chlorine-adapted and control Salmonella Heidelberg morphotype. (P < 0.0001; SEM = 0.058; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, dark gray bar represents non-exposed positive control, and light gray bar represent (broth only) negative control. Figure 2. View largeDownload slide (A) Biofilm formation by Salmonella Heidelberg after 48 h on plastic surface using 96-well polystyrene microtiter plate at room temperature. Means with different superscripts indicate significant differences in the biofilm forming ability of chlorine-adapted and control Salmonella Heidelberg morphotype. (P = 0.0001; SEM = 0.109; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, dark gray bar represents non-exposed positive control, and light gray bar represent (broth only) negative control. (B) Biofilm formation by Salmonella Heidelberg after 48 h on plastic surface using 96-well polystyrene microtiter plate at 37°C. Means with different superscripts indicate significant differences in the biofilm forming ability of chlorine-adapted and control Salmonella Heidelberg morphotype. (P < 0.0001; SEM = 0.058; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, dark gray bar represents non-exposed positive control, and light gray bar represent (broth only) negative control. Figure 3. View largeDownload slide (A) Quantification of Salmonella Heidelberg after 48 h attached to 24-well polystyrene plate at room temperature. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the polystyrene plastic plate. (P = 0.081; SEM = 0.041; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. (B) Quantification of Salmonella Heidelberg after 48 h attached to 24-well polystyrene plate at 37°C. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the polystyrene plastic plate. (P = 0.002; SEM = 0.029; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Figure 3. View largeDownload slide (A) Quantification of Salmonella Heidelberg after 48 h attached to 24-well polystyrene plate at room temperature. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the polystyrene plastic plate. (P = 0.081; SEM = 0.041; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. (B) Quantification of Salmonella Heidelberg after 48 h attached to 24-well polystyrene plate at 37°C. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the polystyrene plastic plate. (P = 0.002; SEM = 0.029; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Biofilm Formation on SS Surface To determine the biofilm formation of chlorine adapted and control Salmonella Heidelberg on a different food-contact surface other than plastic, the strongly attached cells on stainless steel scoops were measured on TSA plates. Similar to plastic quantification, the results were reported in log CFU/mL and the difference in log values were used to determine the difference in biofilm formation between adapted and control cells. There was a significant difference (P > 0.05) observed in the biofilm formation of the adapted rugose cell compared to adapted smooth and control at room temperature. Adapted rugose has a higher attached cell concentration of 5.25 log cfu/mL when compared to the smooth (adapted and control), which has cell concentrations of 4.9 and 4.75 log cfu/mL respectively (Figure 4A). Whereas at 37°C, there was no difference in the concentration of adapted cells (rugose and smooth) that has 5.3 and 5.25 log cfu/mL respectively. The attached cells for chlorine adapted rugose and smooth were higher when compared to the concentration of the control cells, which is 5.02 log cfu/mL (Figure 4B). Salmonella Heidelberg cells attached more to the SS scoop surface at 37°C than that observed at room temperature. Figure 4. View largeDownload slide (A) Quantification of Salmonella Heidelberg after 48 h attached to stainless steel scoop at room temperature. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the stainless steel scoop. (P = 0.033; SEM = 0.071; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. (B) Quantification of Salmonella Heidelberg after 48 h attached to stainless steel scoop at 37°C. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the stainless steel scoop. (P = 0.015; SEM = 0.031; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Figure 4. View largeDownload slide (A) Quantification of Salmonella Heidelberg after 48 h attached to stainless steel scoop at room temperature. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the stainless steel scoop. (P = 0.033; SEM = 0.071; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. (B) Quantification of Salmonella Heidelberg after 48 h attached to stainless steel scoop at 37°C. Means with different superscripts indicate significant differences in the quantification of the attached chlorine-adapted and control Salmonella Heidelberg morphotype on the stainless steel scoop. (P = 0.015; SEM = 0.031; N = 2). The black bar represents chlorine-adapted rugose morphotype, white bar represents chlorine-adapted smooth morphotype, and dark gray bar represents non-exposed control. Antibiotic Cross-Resistance Sensitivity to antibiotics was determined for Salmonella Heidelberg following adaptation to chlorine through exposure to sublethal chlorine concentrations. Zones of inhibition were measured (in millimeters) around an impregnated antibiotic disk. A broth microdilution assay was also performed to determine if there was a change in MIC between the non-exposed and the chlorine tolerant cells. The results for the antibiotic susceptibility patterns of adapted (rugose and smooth) and control cells is shown in Table 2 and 3. The results showed no cross-resistance to antibiotics for the majority of the antibiotics tested when adapted cells were compared to control. However, a reduction in susceptibility was observed in some of the antibiotics. A ≤ 2-mm inhibition zone difference was observed for adapted rugose to GN, S, AMC, and CIP when compared to smooth (adapted and control) (Table 2). The same difference was observed for adapted smooth against SXT and S when compared to control (Table 2). Similarly, a slight increase in MIC was recorded for adapted rugose against S, NA, T, and AMX, which moved from the susceptible to the intermediate category (Table 3). In addition, the adapted (rugose and smooth) cells moved from the susceptible to the intermediate category for T (Table 3). Table 2. Disk diffusion assay of antibiotic cross-resistance patterns of adapted and control S. Heidelberg. Salmonella Heidelberg cultures previously adapted to chlorineb  Susceptibility to indicated antibiotica (μg) SXT GN S AMC NA CIP CTX AMP  Adapted rugose  21/S 15/S 8/R 19/S 18/I 26/S 23/S 20/S  Adapted smooth  19/S 16/S 8/R 21/S 18/I 28/S 23/S 20/S  Control  21/S 16/S 10/R 21/S 18/I 28/S 23/S 20/S  Salmonella Heidelberg cultures previously adapted to chlorineb  Susceptibility to indicated antibiotica (μg) SXT GN S AMC NA CIP CTX AMP  Adapted rugose  21/S 15/S 8/R 19/S 18/I 26/S 23/S 20/S  Adapted smooth  19/S 16/S 8/R 21/S 18/I 28/S 23/S 20/S  Control  21/S 16/S 10/R 21/S 18/I 28/S 23/S 20/S  aSXT, sulphamethoxazole/trimethoprim (25 μg), GN, gentamicin (10 μg), S, streptomycin (10 μg), AMC, amoxicillin/clavulanic acid (30 μg), NA, nalidixic acid (30 μg), CIP, ciprofloxacin (5 μg), CTX, ceftriaxone (30 μg), AMP, ampicillin (10 μg). Susceptibility zones of inhibition (millimeters) are reported as S, susceptible strains; I, intermediate susceptible strains; R, resistance strains. Boldfaced data indicate reduced susceptibility relative to unexposed (control) strains; data not bolded indicate exposed strains with no difference in susceptibility patterns relative to unexposed (control) strains. An increase in resistance was defined as a change in S (before chlorine exposure) to R (after chlorine exposure). bFor adaptation, cultures were previously exposed to increasing sublethal chlorine concentrations, control represent unexposed Salmonella culture. Data represent the average of 3 replicates. View Large Table 3. Broth microdilution assay of antibiotic cross-resistance patterns of adapted and control S. Heidelberg. Salmonella Heidelberg cultures previously adapted to chlorineb  Antibiotica (μg/ml) AMP GN S NA T CIP AMX  Adapted rugose  4/S 4/S 64 16/S 8/I 1/S 16/I  Adapted smooth  4/S 4/S 32 8/S 8/I 1/S 8/S  Control  4/S 4/S 32 8/S 4/S 1/S 8/S  Salmonella Heidelberg cultures previously adapted to chlorineb  Antibiotica (μg/ml) AMP GN S NA T CIP AMX  Adapted rugose  4/S 4/S 64 16/S 8/I 1/S 16/I  Adapted smooth  4/S 4/S 32 8/S 8/I 1/S 8/S  Control  4/S 4/S 32 8/S 4/S 1/S 8/S  aAMP, ampicillin; GN, gentamicin; S, streptomycin; NA, nalidixic acid; T, tetracycline; CIP; ciprofloxacin; AMX, amoxicillin. Data are reported as S, susceptible strains; I, intermediate susceptible strains; R, resistance strains. Boldfaced data indicate reduced susceptibility relative to unexposed (control) strains; data not bolded indicate exposed strains with no difference in susceptibility patterns relative to unexposed (control) strains. An increase in resistance was defined as a change in S (before chlorine exposure) to R (after chlorine exposure). b For adaptation, cultures were previously exposed to increasing sublethal chlorine concentrations, control represent unexposed Salmonella culture. Data represent the average of 2 replicates. View Large DISCUSSION After exposure to increasing sublethal concentrations of chlorine, Salmonella Heidelberg demonstrated the ability to acquire resistance. The ability of the adapted cells to tolerate higher chlorine concentrations continued even after repeated storage on chlorine-free TSA plates, which suggests the stability of adapted cells to the homologous stress of chlorine. In the current study, the MIC for the chlorine adapted (rugose and smooth) cells was 1.2 and 1.1 times higher than non-adapted control. This increase in MIC observed for chlorine-stressed Salmonella Heidelberg was in line with a previous study using Salmonella Typhimurium (Obe et al., 2016) and other studies on Salmonella enterica strains that exhibited adaptation to sublethal stress posed by different antimicrobials (Braoudaki and Hilton, 2004; Kim and Day, 2007; Alonso-Hernando et al., 2009; Stanojevic et al., 2010; Molina-Gonzalez et al., 2014). The inappropriate use of antimicrobials either directly or indirectly in the food processing establishment can expose any bacteria present to sublethal doses of disinfectants and sanitizers that are supposed to reduce the growth of bacteria (Capita et al., 2014). Another important finding from this study is the ability of the chlorine-adapted Salmonella Heidelberg to grow in the presence of high chlorine concentrations even above the concentration approved by the USDA for sanitation purposes on food-contact surfaces (200 ppm). The increase in bacteria tolerance following sublethal antimicrobial concentrations observed in this study have been well reported by others (Braoudaki and Hilton, 2004; Sheridan et al., 2012). Additionally, sublethal doses of antimicrobials have been observed to induce resistance in foodborne pathogens (Braoudaki and Hilton, 2005; Alonso-Calleja et al., 2015). The bacterial cells respond to stressful conditions either by adaptation or elimination, a common means that bacteria use to adjust to strenuous environmental conditions is by changing their morphology (Foster, 2005; Young, 2007). Other authors have reported these changes as a survival mechanism for bacteria (Justice et al., 2004; Young, 2007; Capital et al., 2014). In this study, Salmonella Heidelberg undergoes a morphological change to the rugose morphotype in order to cope with the stress posed by increasing sublethal chlorine concentrations. The change in morphology of the smooth Salmonella Heidelberg to the rugose variant in TSB was observed through the formation of a pellicle that is composed of cell aggregates, thereby allowing the rugose cell to remain as an aggregate in solution. Similar changes have been reported for Vibrio cholerae O1 E1 Tor and Salmonella Typhimurium DT104 (Morris et al., 1996; Anriany et al., 2001). The rugose variant was reported to be more virulent and not easy to kill compared to the smooth variant (Rice et al., 1993; Morris et al., 1996). In addition, studies have reported similar findings on the ability of some microorganisms including E. coli, Listeria monocytogenes, and Pseudomonas aeruginosa to tolerate different environmental stress. These microorganisms are reported to survive by changing their morphology through degradation of the cell wall, elongation of cells, aggregation of damaged bacteria cells and disturbances during cell division (To et al., 2002; Shalamanov, 2005; Giotis et al., 2009; Capita et al., 2014). The conditions at which rugose developed in this study is by daily transfer of an aliquot of 100 μL of culture containing sublethal chlorine concentration. This is different from the observations of other authors on the formation of the rugose variant in both Vibrio and Salmonella. In Salmonella Typhimurium DT104, the rugose variant was observed after 4 days of storage on TSA at room temperature and in Vibrio cholerae O1 strain TSI-4; the rugose variant was observed 2 months post inoculation when the smooth variant was re-cultured under a starved conditions at 16°C (Wai et al., 1998; Anriany et al., 2001). Whereas in the current study, the rugose variant was observed after exposure to increasing sublethal chlorine concentration (200 ppm) but failed to develop when cultured at room temperature with or without exposure to chlorine under similar experimental conditions. Another important finding was the ability of the rugose variant in Salmonella Typhimurium ATCC14028 (Obe et al., 2016) and currently in Salmonella Heidelberg ATCC8326 to retain their morphology even after storage on chlorine-free TSA. The adhesion and subsequent attachment of foodborne pathogens to food processing equipment and environment can result in a major food safety challenge when the pathogen contaminate food products thus causing a foodborne outbreak (Dourou et al., 2011). The attachment of Salmonella Heidelberg to different food-contact surfaces was examined in this study. The ability of the chlorine-adapted and control cells to adhere to both plastic and stainless steel surface was assessed at room temperature and 37°C. The results demonstrates that Salmonella Heidelberg possess the ability to attach to both surfaces tested. It was previously reported that Salmonella has the ability to colonize and attach to different surfaces including plastic, rubber, stainless steel, and glass (Helke et al., 1993; Sinde and Carballo, 2000; Nguyen et al., 2014; Yang et al., 2016). The rugose cells attached and formed stronger biofilms at both temperatures tested and showed no preference to a particular surface. The expression of rugose in Salmonella was previously reported to be due to aggregation of cells and formation of exopolysaccharides (EPS), which might aid in the strong attachment to food-contact surfaces (Marshall, 1992; Morris et al., 1996; Wai et al., 1998; Yildiz and Schoolnik, 1998). The smooth cells both adapted and control seems to attach better on the steel surface. The properties of the surface that Salmonella cells attached to, helps with their survival on such surface (O’Leary et al., 2013). Although Salmonella was reported to attach strongly to a hydrophobic surface, the ability to attach to a particular surface has been reported to be dependent on the strain of Salmonella (Sinde and Carballo. 2000; Chia et al., 2009). Nguyen et al. (2014), reported that Salmonella Typhimurium possess the ability to attach to both stainless steel and acrylic surface but showed greater attachment on stainless steel surfaces. This is because stainless steel is hydrophilic and bacteria attach more to such surfaces as compared to hydrophobic surface (Mafu et al., 1990; Sinde and Carballo. 2000). The results from this study shows that chlorine-adapted Salmonella Heidelberg exhibited no preference to a particular temperature. Salmonella generally grow well and has been observed to form biofilm at 37°C (Nguyen and Yuk, 2013). Room temperature was used in this study to examine the ability of Salmonella to form biofilm when exposed to an unfavorable condition such as temperature abuse. In addition, the adapted cells attached well on both surfaces tested in this study. Plastic and stainless steel surface was used because they are the most common surfaces encountered in food processing (Chmielewski and Frank, 2003; Ismail et al., 2013). Residues of food processing left on these surfaces can contribute to the formation of film if they are not promptly removed during cleaning (Joseph et al., 2001; Chmielewski and Frank, 2003). The application of effective sanitizers on food-contact surfaces following cleaning is important to inactivate and prevent the development of resistance in any pathogenic bacteria present. This will help prevent the acquisition of cross-protection to any other stress conditions encountered in processing The chlorine-adapted cells were examined for their ability to exhibit cross-adaptation to antibiotics. From the results of this study, it appears there were some interactions between adaptations to chlorine and cross-adaption to antibiotics. Salmonella Heidelberg cells that were adapted to sublethal concentrations of chlorine exhibited a certain degree of reduced susceptibility to some of the antibiotics tested. For the adapted cells, a reduced zone of inhibition was observed when compared to non-adapted (control), but the reduction was not significant enough to move above the limit set by the CLSI (2012) guidelines on antibiotic susceptibility testing for the “susceptible” category. Other authors have reported similar observations, Molina-Gonzalez et al. (2014), observed some Salmonella enterica strains that were previously exposed to various antimicrobials including sodium hypochlorite exhibited a lower zone of inhibition when compared to non-exposed strains, but are still susceptible to the antibiotics tested. In another study on the cross-adaption patterns of Salmonella enterica, the authors suggested that a slight reduction in susceptibility is noteworthy; this is because the pathogen may not be inhibited in the presence of the antibiotic over time (Braoudaki and Hilton, 2004). Chlorine-adapted rugose showed a slight reduction in the zone of inhibition to fluoroquinolones (CIP), quinolones (NA), aminoglycosides (S, GN), penicillin (AMX/AMC), and tetracycline (T). In addition, the MIC for the rugose variants increased and moved from the “susceptible” (i.e., bacterial infection will most likely respond to antibiotic treatment) category to the “intermediate” (bacterial infection may or may not respond to antibiotic treatment) category when tested against amoxicillin. A similar trend has been observed in Salmonella Typhimurium (Obe et al., 2016). Other studies have reported similar changes in antibiotic susceptibility patterns of previously adapted foodborne pathogens to different classes of antibiotics (Suller and Russell, 2000; Braoudaki and Hilton, 2004; Capita et al., 2014). Some of these studies suggested that the adaptive nature of foodborne pathogens to antimicrobials like chlorine is directly associated with a broad spectrum mechanism of resistance which includes the presence of an active efflux and alterations to cell permeability. This mechanism makes it difficult for different chemical molecules to enter the adapted cells (Tattawasart et al., 1999; Suller and Russell, 2000; Braoudaki and Hilton, 2004; Capita and Alonso-Calleja, 2013; Molina-Gonzalez et al., 2014). Even though, some authors have reported that sublethal dosage of antimicrobials could select for resistance to antibiotics in Salmonella, some studies do not observe similar findings. The results of this study in part agree with those authors that do not observe a change in susceptibility to antibiotics after exposure to sublethal concentrations of antimicrobials (Thomas et al., 2000; Ledder et al., 2006). In summary, the findings of the current study supports the hypothesis that Salmonella Heidelberg would adapt to sublethal concentrations of chlorine. However, the adaptive tolerance to chlorine observed in this study resulted in the formation of a more virulent Salmonella variant. The adapted cells were better biofilm formers on both food-contact surfaces tested and exhibited a slight reduction in zones of inhibition to different classes of antibiotics. This shows that chlorine stressed Salmonella may not be easily inactivated with high concentrations of chlorine. From this study, it is speculated that the possession of similar broad mechanisms of adaptation may eventually enable the adapted cells to become resistant to certain antibiotic treatment. The findings in this study signify a possible challenge to food safety and suggest that the misuse of the antimicrobial agent at a sublethal concentration could represent a potential public health risk. REFERENCES Alonso-Calleja C., Guerrero-Ramos E., Alonso-Hernando A., Capita R.. 2015. Adaptation and cross-adaptation of Escherichia coli ATCC 12806 to several food-grade biocides. Food Control . 56: 86– 94. Google Scholar CrossRef Search ADS   Alonso-Hernando A., Capita R., Prieto M., Alonso-Calleja C.. 2009. Comparison of antibiotic resistance patterns in Listeria monocytogenes and Salmonella enterica strains pre-exposed and exposed to poultry decontaminants. Food Control . 20: 1108– 1111. Google Scholar CrossRef Search ADS   Anriany Y. A., Weiner R. M., Johnson J. A., De Rezende C. E., Joseph S. W.. 2001. Salmonella enterica serovar Typhimurium DT104 displays a rugose phenotype. Appl. Environ. Microbiol . 67: 4048– 4056. Google Scholar CrossRef Search ADS PubMed  Bae Y. S., Baek, Lee S.. 2012. Resistance of pathogenic bacteria on the surface of stainless steel depending on attachment form and efficacy of chemical sanitizers. Int. J. Food Microbol.  153: 465– 473. Google Scholar CrossRef Search ADS   Barnes L. M., Lo M. F., Adams M. R., Chamberlain A. H. L. 1999. Effect of milk proteins on adhesion of bacteria to stainless steel surfaces. Appl. Env. Microbiol.  65: 4543– 4548. Braoudaki M., Hilton A. C.. 2004. Adaptive resistance to biocides in Salmonella enterica and Escherichia coli O157 and cross-resistance to antimicrobial agents. J. Clin. Microbiol.  42: 73– 78. Google Scholar CrossRef Search ADS PubMed  Braoudaki M., Hilton A. C.. 2005. Mechanism of resistance in Salmonella enterica adapted to erythromycin, benzalkonium chloride and triclosan. Int. J. Anti. Agents.  25: 31– 37. Google Scholar CrossRef Search ADS   Buchmeier N. A., Heffron F.. 1990. Induction of Salmonella stress proteins upon infection of macrophages. Science . 248: 730– 732. Google Scholar CrossRef Search ADS PubMed  Capita R., Alonso-Calleja C.. 2013. Antibiotic-resistant bacteria: a challenge for the food industry. Crit. Rev. Food Sci. Nutr.  53: 11– 48. Google Scholar CrossRef Search ADS PubMed  Capita R., Riesco-Pelaez F., Alonso-Hernando A., Alonso-Calleja C.. 2014. Exposure of Escherichia coli ATCC 12806 to sublethal concentrations of food-grade biocides influences its ability to form biofilm, resistance to antimicrobials, and ultrastructure. Appl. Environ. Microbiol.  80: 1268– 1280. Google Scholar CrossRef Search ADS PubMed  CDC. (Center for Disease Control and Prevention). 2011a. Estimates findings: estimates of foodborne illnesses in the United States [online]. Accessed Aug. 10, 2016. http://www.cdc.gov/foodborneburden/2011-foodborne-estimates.html. CDC. (Center for Disease Control and Prevention). 2011b. Burdens of Foodborne Illnesses: Findings [online]. Accessed Jan. 14, 2017. https://www.cdc.gov/foodborneburden/2011-foodborne-estimates.html. CDC. (Center for Disease Control and Prevention). 2014. Reports of selected Salmonella outbreak investigations [online]. Accessed Jan. 14, 2017. https://www.cdc.gov/salmonella/outbreaks.html. Chia T., Goulter R., McMeekin T., Dykes G., Fegan N.. 2009. Attachment of different Salmonella serovars to materials commonly used in a poultry processing plant. Food Microbiol . 26: 853– 859. Google Scholar CrossRef Search ADS PubMed  Chmielewski R. A. N., Frank J. F.. 2003. Biofilm formation and control in food processing facilities. Compr. Rev. Food Sci. Food Saf.  2: 22– 32. Google Scholar CrossRef Search ADS   CLSI. (Pennsylvania: National Committee for Clinical Laboratory Standards). 2008. Reference method for broth dilution antifungal susceptibility testing of filamentous fungi. Approved Standard – Second Edition. M38-A2. CLSI. (Pennsylvania: National Committee for Clinical Laboratory Standards). 2012. Performance standards for antimicrobial disk susceptibility test. Approved Standard – Twelfth Edition. M02-A12. Costerton J. W., Lewandowski Z., Caldwell D. E., Kober D. R., Lappin-Scott H. M.. 1995. Microbial biofilms. Ann. Rev. Microbiol.  49: 711– 745. Google Scholar CrossRef Search ADS   Czechowski M. H. 1990. Bacterial attachment to Buna-N gaskets in milk processing equipment. Australian J. Dairy Tech.  45: 113– 114. Davidson P. M., Harrison M. A.. 2002. Resistance and adaptation to food antimicrobials, sanitizers, and other process controls. Food Tech . 56: 69– 78. Dourou D., Beauchamp C. S., Yoon Y., Geornaras I., Belk K. E., Smith G. C., Nychas G. J., Sofos J. N.. 2011. Attachment and biofilm formation by Escherichia coli O157:H7 at different temperatures, on various food contact surfaces encountered in beef processing. Int. J. Food Microbiol.  149: 262– 268. Google Scholar CrossRef Search ADS PubMed  Foster P. L. 2005. Stress responses and genetic variation in bacteria. Mut. Res. Rev.  569: 3– 11. Google Scholar CrossRef Search ADS   Frank J. K., Koffi R. A.. 1990. Surface-adherent growth of Listeria monocytogenes is associated with increased resistance to surfactant sanitizers and heat. J. Food Protect.  53: 550– 554. Google Scholar CrossRef Search ADS   Giaouris E., Chorianopoulos N., Nychas G. J. E.. 2005. Effect of temperature, pH, and water activity on biofilm formation Salmonella Enteritidis PT4 on stainless steel surfaces as indicated by the bead vortexing method and conductance measurements. J. Food Protect.  68: 2149– 2154. Google Scholar CrossRef Search ADS   Giotis E., Blair I. S., McDowell D. A.. 2009. Effects of short-term alkaline adaptation on surface properties of Listeria monocytogenes 10403S. The Open Food Sci. J.  3: 62– 65. Google Scholar CrossRef Search ADS   Helke D., Somers E., Wong A.. 1993. Attachment of Listeria monocytogenes and Salmonella Typhimurium to stainless steel and buna-N rubber in the presence of milk and individual milk components. J. Food Protect.  56: 479– 484. Google Scholar CrossRef Search ADS   Hood S. K., Zottola E. A.. 1997. Adherence to stainless steel by foodborne microorganisms during growth in model food systems. Int. J. Food Microbiol.  37: 145– 153. Google Scholar CrossRef Search ADS PubMed  Hur J. C., Jawale, Lee J. H.. 2012. Antimicrobial resistance of Salmonella isolated from food animals: review. Food Res. Int.  45: 819– 830. Google Scholar CrossRef Search ADS   IFT (Institute of Food Technologists). 2006. Antimicrobial resistance: Implications for the food systems. Compr. Rev. in Food Sci. Food Saf.  5: 71– 137. CrossRef Search ADS   Ismaïl R., Aviat F., Michel V., Le Bayon I., Gay-Perret P., Kutnik M., Fédérighi M.. 2013. Methods for recovering microorganisms from solid surfaces used in the food industry: a review of the literature. Int. J. Environ. Res. Pub. Health.  10: 6169– 6183. Google Scholar CrossRef Search ADS   Iturriaga M. H., Tramplin M. L., Escartin E. F.. 2007. Colonization of tomatoes by Salmonella Montevideo is affected by relative and storage temperature. J. Food Protect.  70: 30– 34. Google Scholar CrossRef Search ADS   Jiang X., Doyle M. P.. 1999. Fate of Escherichia coli O157:H7 and Salmonella Enteritidis on currency. J. Food Protect.  62: 805– 807. Google Scholar CrossRef Search ADS   Joseph B., Otta S., Karunasagar I., Karunasagar I.. 2001. Biofilm formation by Salmonella spp. on food contact surfaces and their sensitivity to sanitizers. Int. J. Food Microbiol.  64: 367– 372. Justice S. S., Hung C., Theriot J. A., Fletcher D. A., Anderson G. G., Footer M. J., Hultgren S. J.. 2004. Differentiation and developmental pathways of uropathogenic Escherichia coli in urinary tract pathogenesis. Proc. Natl. Acad. Sci.  101: 1333– 1338. Google Scholar CrossRef Search ADS   Kim D., Day D. F.. 2007. A biocidal combination capable of sanitizing raw chicken skin. Food Control . 18: 1272– 1276. Google Scholar CrossRef Search ADS   Krysinski E. P., Brown L. J., Marchisello T. J.. 1992. Effect of cleaners and sanitizers on Listeria monocytogenes attached to product contact surfaces. J. Food Protect.  55: 246– 251. Google Scholar CrossRef Search ADS   Kusumaningrum H. D., Riboldi G., Hazeleger W. C., Beumer R. R.. 2003. Survival of foodborne pathogens on stainless steel surfaces and cross-contamination to foods. Int. J. Food Microbiol . 85: 227– 236. Google Scholar CrossRef Search ADS PubMed  Ledder R. G., Gilbert P., Wilis C., McBain A. J.. 2006. Effects of chronic triclosan exposure upon the antimicrobial susceptibility of 40 ex-situ environmental and human isolates. J. Appl. Microbiol.  100: 1132– 1140. Google Scholar CrossRef Search ADS PubMed  Lou Y., Yousef A. E.. 1996. Resistance of Listeria monocytogenes to heat after adaptation to environmental stresses. J. Food Protect.  5: 465– 471. Google Scholar CrossRef Search ADS   Mafu A. A., Roy D., Goulet J., Magny P.. 1990. Attachment of Listeria monocytogenes to stainless steel, glass, polypropylene and rubber surfaces after short contact times. J. Food Protect . 53: 742– 746. Google Scholar CrossRef Search ADS   Marshall K. C. 1992. Biofilms: an overview of bacterial adhesion, activity, and control at surfaces. ASM News . 58: 202– 207. Mead P. S., Slutsker L., Dietz V., McCaig L. F., Bresee J. S., Shapiro C., Griffin P. M., Tauxe R. V.. 1999. Food-related illness and death in the United States. Emerg. Infect. Dis . 5: 607– 625. Google Scholar CrossRef Search ADS PubMed  Molina-Gonzalez D., Alonso-Calleja C., Alonso-Hernando A.. 2014. Effect of sublethal concentrations of biocides on the susceptibility to antibiotics of multi-drug resistance Salmonella enterica strains. Food Control . 40: 329– 334. Google Scholar CrossRef Search ADS   Morris G. K., Wells J. G.. 1970. Salmonella contamination in a poultry-processing plant. Appl. Microbiol . 38: 2465– 2467. Morris J. G. Jr, Sztein M. B., Rice E. W., Nataro J. P., Losonsky G. L., Panigrahi P., Tacket C. O., Johnson J. A.. 1996. Vibrio cholerae O1 can assume a chlorine-resistant rugose survival form that is virulent for humans. The J. Infect. Dis . 174: 1364– 1368. Google Scholar CrossRef Search ADS   Nguyen H. D. N., Yuk H. G.. 2013. Changes in resistance of Salmonella Typhimurium biofilms formed under various conditions to industrial sanitizers. Food Control . 29: 236– 240. Google Scholar CrossRef Search ADS   Nguyen H. D. N., Yang Y. S., Yuk H. G.. 2014. Biofilm formation of Salmonella Typhimurium on stainless steel and acrylic surfaces as affected by temperature and pH level. LWT – Food Sci Technol . 55: 283– 288. Google Scholar CrossRef Search ADS   O’Leary D., Mc Cabe E. M., McCusker M. P., Martin M., Fanning S., Duffy G.. 2013. Microbiological study of biofilm formation in isolates of Salmonella enterica Typhimurium DT104 and DT104b cultured from the modern pork chain. Int. J. Food Microbiol.  161: 36– 43. Google Scholar CrossRef Search ADS PubMed  Obe T., Nannapaneni R., Sharma C. S.. 2016. Development of rugose morphotype of Salmonella Typhimurium following exposure to sub-inhibitory chlorine concentrations that exhibit chlorine resistance and strong biofilm forming ability. Poult. Sci.  95( E – Suppl. 1): 35. Painter J. A., Hoekstra R. M., Ayers T., Tauxe R. V., Braden C. R., Angulo F. J., Griffin P. M.. 2013. Attribution of foodborne illnesses, hospitalizations, and deaths to food commodities by using outbreak data, United States, 1998–2008. Emerg. Infect. Dis . 19: 407– 415. Google Scholar CrossRef Search ADS PubMed  Patel J., Sharma M.. 2010. Differences in attachment of Salmonella enterica serovars to cabbage and lettuce leaves. Int. J. Food Microbiol.  139: 41– 47. Google Scholar CrossRef Search ADS PubMed  Rice E. W., Johnson C. H., Clark R. M., Fox K. R., Reasoner D. J., Dunnigan M. E.. 1993. Vibrio cholerae O1 can assume a ‘rugose’ survival form that resists killing by chlorine, yet retains virulence. Int. J. Env. Health Res . 3: 89– 98. Google Scholar CrossRef Search ADS   Scallan E., Hoekstra R. M., Angulo F. J., Tauxe R. V., Widdowson M. A., Jones S. L., Griffin P. M.. 2011. Foodborne illnesses acquired in the United States- major pathogen. Emerg. Infect. Dis.  17: 7– 15. Google Scholar CrossRef Search ADS PubMed  Scott E., Bloomfield S. F.. 1990. The survival and transfer of microbial contamination via cloths, hands and utensils. The J. Appl. Bacteriol.  68: 271– 278. Google Scholar CrossRef Search ADS   Shalamanov D. S. 2005. Chlorhexidine gluconate-induced morphological changes in gram negative microorganisms. Biotechnol. Biotechnol. Eq.  19: 121– 124. Google Scholar CrossRef Search ADS   Sheridan A., Lenahan M., Duffy G., Fanning S., Burgess C.. 2012. The potential for biocide tolerance in Escherichia coli and its impact on the response to food processing stresses. Food Control . 26: 98– 106. Google Scholar CrossRef Search ADS   Sinde E., Carballo J.. 2000. Attachment of Salmonella spp. and Listeria monocytogenes to stainless steel, rubber and polytetrafluoroethylene: the influence of free energy and the effect of commercial sanitizers. Food Microbiol.  17: 439– 447. Google Scholar CrossRef Search ADS   Stanojevic D., Comic L., Stefanovic O., Solujic-Sudolak S.. 2010. In vitro synergistic antibacterial activity of Salvia officinalis L. and some preservatives. Arch. Biol. Sci.  62: 175– 183. Google Scholar CrossRef Search ADS   Suller M. T. E., Russell A. D.. 2000. Triclosan and antibiotic resistance in Staphylococcus aureus. J. Antimicrob. Chemother.  46: 11– 18. Google Scholar CrossRef Search ADS PubMed  Tattawasart U., Maillard J. Y., Furr J. R., Russell A. D.. 1999. Development of resistance to chlorhexidine diacetate and cetylpyridinium chloride in Pseudomonas stuzeri and changes in antibiotic susceptibility. J. Hos. Infect.  42: 219– 229. Google Scholar CrossRef Search ADS   Thomas L., Maillard J. Y., Lambert R. J. W., Russell A. D.. 2000. Development of resistance to chlorhexidine diacetate in Pseudomonas aeruginosa and the effect of a “residual” concentration. J. Hos. Infect.  46: 297– 303. Google Scholar CrossRef Search ADS   To M. S., Favrin S., Romanova N., Griffiths M. W.. 2002. Post-adaptational resistance to benzalkonium chloride and subsequent physicochemical modifications of Listeria Monocytogenes. Appl. Environ. Microbiol.  68: 5258– 5264. Google Scholar CrossRef Search ADS PubMed  USDA-FSIS. (United States Department of Agriculture, Food Safety Inspection Service). 2015. Safe and suitable ingredients in the production of meat, poultry, and egg products. FSIS Directive 7120.1 Revision 36 [online]. Accessed Aug. 2, 2016. http://www.fsis.usda.gov/wps/wcm/connect/bab10e09-aefa-483b-8be8809a1f051d4c/7120.1.pdf?MOD=AJPERES.. Wai S. N., Mizunoe Y., Takade A., Kawabata S. I., Yoshida S. I.. 1998. Vibrio cholerae O1 strain TSI-4 produces the exopolysaccharide materials that determine colony morphology, stress resistance, and biofilm formation. Appl. Environ. Microbiol . 64: 3648– 3655. Google Scholar PubMed  Yang Y., Miks-Krajnik M., Zheng Q., Lee S. B., Lee S. C., Yuk H. G.. 2016. Biofilm formation of Salmonella Enteritidis under food-related environmental stress conditions and its subsequent resistance to chlorine treatment. Food Microbiol . 54: 98– 105. Google Scholar CrossRef Search ADS   Yildiz F. H., Schoolnick G. K.. 1998. Role of rpoS in stress survival and virulence of Vibrio cholerae. J. Bacteriol.  180: 773– 784. Google Scholar PubMed  Young D. 2007. Bacterial morphology: why have different shapes? Curr. Opin. Microbiol.  10: 596– 600. Google Scholar CrossRef Search ADS PubMed  © 2018 Poultry Science Association Inc.

Journal

Poultry ScienceOxford University Press

Published: Mar 1, 2018

There are no references for this article.

You’re reading a free preview. Subscribe to read the entire article.


DeepDyve is your
personal research library

It’s your single place to instantly
discover and read the research
that matters to you.

Enjoy affordable access to
over 12 million articles from more than
10,000 peer-reviewed journals.

All for just $49/month

Explore the DeepDyve Library

Unlimited reading

Read as many articles as you need. Full articles with original layout, charts and figures. Read online, from anywhere.

Stay up to date

Keep up with your field with Personalized Recommendations and Follow Journals to get automatic updates.

Organize your research

It’s easy to organize your research with our built-in tools.

Your journals are on DeepDyve

Read from thousands of the leading scholarly journals from SpringerNature, Elsevier, Wiley-Blackwell, Oxford University Press and more.

All the latest content is available, no embargo periods.

See the journals in your area

DeepDyve Freelancer

DeepDyve Pro

Price
FREE
$49/month

$360/year
Save searches from Google Scholar, PubMed
Create lists to organize your research
Export lists, citations
Access to DeepDyve database
Abstract access only
Unlimited access to over
18 million full-text articles
Print
20 pages/month
PDF Discount
20% off