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Investigating factors contributing to the survival of Salmonella enterica on mini cucumbers Chen, Huihui 2017

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	   1	  INVESTIGATING FACTORS CONTRIBUTING TO THE SURVIVAL OF SALMONELLA ENTERICA ON MINI CUCUMBERS By Huihui Chen BSc., Zhejiang University, 2015 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE   in  THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Food Science) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) August 2017 © Huihui Chen, 2017     	   ii	  Abstract Cucumbers have been associated with recent Salmonella enterica (S.enterica) outbreaks. The ability of S. enterica to attach or internalize into produce may be a factor that make these produce items more likely to be sources of S. enterica contamination. The purpose of this study was to evaluate the survival capability of S. enterica on mini cucumbers and explore the factors contributing to the survival of this foodborne pathogen on the surface of cucumbers. Five strains of S. enterica representing different serotypes were individually inoculated onto mini cucumbers and subsequently incubated at 22 ± 2 °C for 8 days or at 4 ± 2 °C for 19 days respectively. Crystal violet assay was performed to quantify the biofilm formation and attachment capability based on the value of optical density at 595 nm of the destaining crystal violet at the specific interval time (0, 3, 6, 9, 12, 24, 48, 72, 96 hours). The phenotypic evaluation of red dry and rough (rdar) morphotype formation of S. enterica were conducted on Luria-Bertani (LB) agar complemented with Congo red (40 µg/mL) and Coomassie brilliant blue (20 µg/mL). The results suggested different S. enterica strains showed differential survival rates at both temperatures. S. Poona exhibited the strongest survival ability at 22 ± 2 °C with the highest Δlog CFU and maximum achieved density (Nmax) of 0.84 ± 0.01 and 6.72 ± 0.05, respectively. However, at 4 ± 2 °C, S. Enteritidis survived better compared with S. Poona due to the least cell density decrease of -0.91 ± 0.01 Δlog CFU and maximum achieved density of 6.04 ± 0.09. Besides, survival behaviors of S. enterica were found to be associated with biofilm formation ability and the biofilm ability differed among different strains. This means that biofilm formation contributes to the survival ability of S. enterica on mini cucumbers. 	   iii	  Lastly, different strains exhibited specific morphotypes on Congo red agar, indicating that both curli and cellulose contribute to biofilm formation of S. enterica. Unique survival characteristics among S. enterica reveal that corresponding interventions need to be applied to eliminate contamination of produce with specific S. enterica strains.              	   iv	  Lay summary The purpose of this study was to evaluate the survival capability of S. enterica on mini cucumbers and explore the factors contributing to the survival of this foodborne pathogen on the surface of cucumbers. Unique survival characteristics among S. enterica strains at different temperatures increase our understanding of the factors influencing microbial safety of fresh produce and reveal that corresponding interventions need to be applied to eliminate contamination of produce with specific S. enterica strains.           	   v	  Preface This report is original, unpublished, independent work by the author, Huihui Chen.                  	   vi	  Table of Contents   Abstract………………………………………………………………………................. ii Lay summary…………………………………………………………………...………. iv Preface………………………………………………………………………………....... v Table of Contents………………………….…………………………………................ vi List of Tables………………….…………………………………................................... ix List of Figures………………….………………………………….................................. x List of Abbreviations………………….…………………………………...................... xi Acknowledgements…………………….…………………………………................... xiii Chapter 1: Literature review and research purpose…………………………………. 1     1.1 Salmonella………………………………………………………………………..... 1         1.1.1 Microbial characteristics of Salmonella…………………………………...….. 1         1.1.2 Classification of Salmonella……………………………………………….…. 2         1.1.3 Foodborne disease burden associated with S. enterica ………………….…… 4         1.1.4 Food products associated with S. enterica …………………………………… 5             1.1.4.1 Meat products……………………………………………………….……. 5             1.1.4.2 Egg and dairy products…………………………………………….…….. 7             1.1.4.3 Vegetable and fruit produce ……………………………………………... 7         1.1.5 Factors contributing to the survival of S. enterica on fresh produce…………10     1.2 Microbial biofilms……………………………………………………………..… 11         1.2.1 Characteristics of biofilms…………………………………………………... 11         1.2.2 Mechanism of biofilm formation………………………………………….… 13         1.2.3 Impact of biofilms on public health……………………………………….… 14         1.2.4 Factors affecting biofilm formation…………………………………………. 16 	   vii	      1.3 Major biofilm components involved in rdar morphotype……………………...… 17         1.3.1 The role of curli in biofilm formation……………………………………..… 17         1.3.2 The role of cellulose in biofilm formation………………………………...… 19         1.3.3 Congo red (CR) binding assays……………………………………………... 20     1.4 Research purpose……………………………………………………………….... 22 Chapter 2: Survival of S. enterica on mini cucumbers…………………………...…. 23     2.1 Introduction ……………………………………………………………………… 23     2.2 Materials and methods………………………………………...……………….… 24         2.2.1 Origin and maintenance of S. enterica strains………………………………. 24         2.2.2 Preparation of strains and mini cucumbers……………………………..…… 25         2.2.3 Mini cucumber inoculation and recovery of S. enterica ………………….… 26         2.2.4 Colony enumeration using Thin Agar Layer (TAL) method……………..…. 27         2.2.5 Data and statistical analysis…………………………………………….…… 28     2.3 Results and discussion…………………………………………………………… 28         2.3.1 Growth (Δlog CFU) of S. enterica on mini cucumbers…………………..…. 29         2.3.2 Maximum achieved density (Nmax) of S. enterica on mini cucumbers…….... 32     2.4 Conclusions…………………………………………………………………….… 33     2.5 Potential interventions for specific S. enterica strains……………...…….……… 35 Chapter 3: Examination of biofilm formation and attachment ability of S. enterica……………………………................................................................................. 37     3.1 Introduction………………………………………………………………………. 37     3.2 Materials and methods...…………………………………………………….…… 39         3.2.1 Quantification of biofilm formation of S. enterica ……………………...….. 39         3.2.2 Quantification of cell attachment of S. enterica ………………………….… 40         3.2.3 Data and statistical analysis……………………………………………….… 41     3.3 Results and discussion………………………………………………………..….. 43 	   viii	          3.3.1 Quantification of S. enterica biofilm formation…………………………..… 43         3.3.2 Quantification of cell attachment………………………………………….… 47             3.3.2.1 S. enterica attachment ability………………………………………...…. 47             3.3.2.2 Relationship between attachment ability and survival capability…….… 49     3.4 Conclusions…………………………………………………………………….… 52 Chapter 4: Characterization of the S. enterica rdar morphotype ……………….… 53     4.1 Introduction…………………………………………………………………….… 53     4.2 Materials and methods………………………………………………………...…. 54         4.2.1 Bacterial strains……………………………………………………………… 54         4.2.2 Phenotypic evaluation of rdar morphotype………………………………..… 55     4.3 Results and discussion…………………………………………………………… 56     4.4 Conclusions…………………………………………………………………….… 59 Chapter 5: Conclusion and future direction……………………………………...…. 61     5.1 Conclusion……………………………………………………………………….. 61     5.2 Future direction………………………………………………………………...… 62 References…………………………………………………………………………….... 64          	   ix	  List of Tables  Table 1.1 Selected outbreaks of salmonellosis associated with the consumption of fresh produce…………………………………………………………………….…….. 9 Table 2.1 S. enterica strains used in this study.…………………………………...…… 25 Table 2.2 Δlog CFU and maximum achieved density (Nmax) of S. enterica on mini cucumbers at room temperature (22 ± 2 °C) and at refrigeration temperature (4 ± 2 °C)………………………………………………………………………….…. 32 Table 3.1 Area under the curve (AUC) and optical density at 595 nm (OD595) and attachment ability of Salmonella serotypes incubated at room temperature (22 ± 2 °C) and at refrigeration temperature (4 ± 2 °C)………………………………… 48 Table 4.1 Morphotypes of S. enterica, curli fimbriae and cellulose production on Congo red plates.……………………………………………………………..………… 57            	   x	  List of Figures  Fig. 2.1 Schematic illustration of the modified one-step Thin Agar Layer method ……………………………………………………………………....….. 27 Fig. 2.2 Survival of Salmonella strains on mini cucumbers stored at room temperature (22 ± 2 °C, a) and at refrigeration temperature (4 ± 2 °C, b). Bacterial cells were recovered on BHI-XLD agar. Error bars indicate the standard errors of the means of three replicates.………………………………………………...…………….. 30 Fig. 3.1 Biofilm formation by S. enterica strains represented by OD595 at 22 ± 2 °C. The data are presented as the mean of three replicates and the error bars indicate the standard errors of the means……………………………………………………. 43 Fig. 3.2 Mean values of attachment ability of S. enterica strains represented by OD595 at room temperature (22 ± 2 °C) (a) and refrigeration temperature (4 ± 2 °C) (b). The data are presented as the mean of triplicate replications and the error bars indicate the standard errors of the means……………………………………………...… 50 Fig. 4.1 Rdar morphotypes of S. enterica on Congo red plates. (+) Pseudomonas aeruginosa PA14, positive control; (-) L. monocytogenes HPB642, negative control……………………………………………………………………...…… 59      	   xi	  List of Abbreviations  ANOVA  AUC  bdar  BHI  CDC  CFU  CR  CV  D/E  ECM  EPS  IS  LB  Nmax  OD595  OD600  PAI  PBS  PCR  QMRA  rdar    Salmonella enterica sar Analysis of variance Area under the curve Brown, dry, and rough Brain-Heart-Infusion Centers for Disease Control  Colony forming units Congo red Crystal violet Dey and Engley Extracellular matrix Extracellular polymeric substances Insertion sequence Luria-Bertani Maximum achieved density Optical density at 595 nm Optical density at 600 nm Pathogenicity islands Phosphate buffered saline Polymerase chain reaction Quantitative microbial risk assessment Red, dry, and rough S. enterica Smooth and red	   xii	  saw  SPI 1  SPI 2  TAL  TTS   T3SS  WHO  XLD     Smooth and white Salmonella pathogenicity island 1 Salmonella pathogenicity island 2 Thin Agar Layer Type III secretion Type III secretion systems World Health Organization Xylose-lysine-deoxycholate         	  xiii Acknowledgements First and foremost, I extend my sincere gratitude to my supervisor, Dr. Siyun Wang for being an excellent mentor and role model throughout this project. Your continued support, optimism and wealth of knowledge are especially appreciated. Thank you for your guidance and faith in my abilities. Secondly, I gratefully acknowledge my committee members, Dr. Vivien Measday and Dr. Susan Bach, for their innovative ideas and probing questions and Dr. Zhaoming Xu, Dr. Kim Cheng for their unfailing support. Your considerations have been ever so helpful during the progression of my project. A very special gratitude goes out to all down at Research Fund and also Rich Foundation for helping and providing the funding for the work. Thank you to the University of British Columbia for funding that made this project possible. Technical support from Aljosa Trmcic, Karen Fong, Patricia Hingston, Justin Falardeau and Yue Dai are gratefully acknowledged. Thank you for your assistance and fun laboratory discussions! To all members of the Wang lab, thank you for undertaking this fun roller coaster ride of a MSc. with me, and keeping me sane. Meeting such like-minded people has definitely made this journey exciting. And last but not least, I am so thankful for my family and friends. I appreciate all the support and stability you have given me. Thank you for being my biggest cheerleaders.	   1 Chapter 1: Literature review and research purpose  1.1 Salmonella       1.1.1 Microbial characteristics of Salmonella Salmonella is a genus of rod-shaped or bacillus gram-negative bacteria (Madigan, Martinko and Parker, 1997) and are divided into Salmonella bongori and Salmonella enterica. Salmonella can be found in poultry, pigs, cows, pets and many wild animals. Salmonella bacteria were discovered by Theobald Smith, but named in honor of his boss Daniel Elmer Salmon (Jay, 2012; Lee et al., 2015). Salmonella bacteria are between 2 and 5 micrometers long and 0.7 to 1.5 micrometers in diameter (Wray, 2000). They have flagella, which are tail-like projections made of proteins that help the bacteria to move (Prendergast et al., 2009). Salmonella typically live in the intestines of both cold and warm blooded animals and can also be found throughout the natural environment, including soil and water that have been contaminated with animal excrement (Rittmann and McCarty, 2012; Cabral, 2010). Although the bacteria cannot multiply outside of the host digestive tract, they can live for several weeks in water and several years in soil when conditions such as humidity, pH and temperature are favorable (Podolak et al., 2010). S. enterica is a common contaminant of food and can be hazardous to consumers if the food is not thoroughly cooked, representing a public health risk (Prendergast et al., 2009).   More than 2,500 serotypes of S. enterica have already been recognized (Hidekazu et al., 2009). S. enterica are zoonotic, which means that they can be transmitted from animals to humans or from humans to animals. Fecal-oral route is the dominant way of transmission 	   2 (Fedorka-Cray et al., 1995). Human usually get infected by the consumption of foods or water contaminated with fecal matter that contains S. enterica. Proper food thawing and handling techniques along with proper sanitation and hand-washing are effective ways to prevent S. enterica infections. S. enterica, through some of its serotypes such as Typhimurium and Enteriditis, shows signs of the ability to infect several different mammalian host species, while other serotypes such as Typhi seem to be restricted to only a few hosts (Thomson et al., 2008). Loss of genetic elements and mutation is one of the ways that S. enterica serotypes adapt to their hosts (Tsolis et al., 1999; Uzzau et al., 2000). In more complex mammalian species, immune systems, which are responsible for pathogen specific immune responses, target serotypes of S. enterica through binding of antibodies to structures like flagella (Fierer and Guiney, 2001). S. enterica can evade a host's immune system by losing the genetic elements that code for a flagellum. Kisela et al. found that more pathogenic serotypes of S. enterica have certain adhesins in common that have developed out of convergent evolution (Kisiela et al., 2012), which means that, as these strains of S. enterica have been exposed to similar conditions such as immune systems, similar structures evolved separately to negate these similar, more advanced defenses in hosts.       1.1.2 Classification of Salmonella S. bongori is predominantly associated with cold-blooded animals and rarely causes disease in humans, while S. enterica is responsible for the majority of Salmonella infections in warm-blooded animals (Porwollik et al., 2004) and the majority of human foodborne illnesses (Malorny et al., 2011). Biochemical assays of the somatic (O) and 	   3 flagellar (H) antigens are used to characterize the bacteria into the different serotypes. S. enterica can be further divided into six sub-species (enterica, salamae, arizonae, diarizonae, indica and houtenae, or I, II, IIIa, IIIb, IV, and VI, respectively), each with hundreds of different serotypes (Porwollik et al., 2004; Malorny et al., 2011). Among these six subspecies, only subspecies I is associated with disease in warm-blooded animals (Porwollik et al., 2004) with over 2,300 serotypes identified within subspecies I. Three serotypes of S. enterica subspecies (Typhimurium, Enteritidis and Newport) are responsible for over 50 percent of all human infections	   in the U.S, as reported by the Journal of Bacteriology (Porwollik et al., 2004). The most common serotypes of S. enterica are Enteritidis and Typhimurium, which account for half of all human infections (Saphra and Winter, 1957). S. enterica serotype Enteritidis are considered the most important group of foodborne S. enterica causing gastrointestinal illness of varying severity in humans (Forshell and Wierup, 2006). S. Typhimurium is the predominant serotype isolated from humans in Europe and pigs are an important reservoir of this particular serotype (Boyen et al., 2008). According to the host preference and disease manifestations in humans, Salmonella serotypes can also be clinically classified into typhoidal or nontyphoidal (Li et al., 2017). Nontyphoidal Salmonella serotypes are more common, and usually cause self-limiting gastrointestinal disease, infecting a range of animals. Besides, nontyphoidal Salmonella are zoonotic, which means that they can spread between humans and other animals. Typhoidal serotypes including Salmonella Typhi and Salmonella Paratyphi are adapted to humans or higher primates and do not cause systemic infections in other animals. 	   4       1.1.3 Foodborne disease burden associated with S. enterica Disease caused by S. enterica is one of the most frequently reported causes of foodborne gastroenteritis. The Public Health Agency of Canada estimates that every year about 1 in 8 Canadians (4 million people) get sick from the food they eat (Thomas et al., 2006). A study (Thomas et al., 2013) in 2013 reported that non-typhoidal Salmonella causes approximately 88,000 cases per year in Canada. In the United States, Salmonella was estimated to cause 1.0 million cases each year (Scallan et al., 2011) and to be the leading cause of hospitalizations and deaths from foodborne disease.  Salmonella serotypes differ in their natural reservoirs and ability to cause human infections. Only a small proportion of >2,500 serotypes cause most human infections (Jackson 2013), particularly Salmonella Enteritidis and Salmonella Typhimurium which caused more than 50 percent of gastrointestinal illness of varying severity in humans (Forshell and Wierup, 2006). World Health Organization (WHO) reported that these two serotypes were considered the most frequently isolated serotypes in human outbreaks of salmonellosis in Europe from 1993 to 1998, leading to 77.1% of the reportable outbreaks (Tirado, Schmidt, 2001).  Similar data from other countries almost certainly underestimate the magnitude of the problem, as many cases of salmonellosis are not reported. Because usually, people that get infected with S. enterica do not receive therapy in time and many laboratory findings are not reported. Apart from the human health implications, S. enterica has been a threat to worldwide food production and many strains are antibiotic-resistant, principally due to the therapeutic use of antimicrobials in animals. Therefore, increased attention should be 	   5 focused on the prevention and control of Salmonella in food production since the work is an urgent challenge.       1.1.4 Food products associated with Salmonella Various food vehicles are implicated in salmonellosis outbreaks, including meat, poultry, eggs, produce, salad, butter, nuts and many other foods. The most commonly implicated food commodity differs by Salmonella serotype. Overall, eggs are considered as the most commonly implicated food commodity, followed by chicken, pork, beef, fruit and turkey (Jackson 2013). Notable relationships between Salmonella serotypes and food commodities that point to major food reservoirs for different serotypes were found by Jackson (Jackson, 2013). The contamination of food commodities with S. enterica can occur at multiple stages throughout the food chain and there are many factors contributing to the outbreaks, including inadequate temperature control, infected food handlers, contaminated raw ingredients, cross-contamination, and inadequate heat treatment. From a risk assessment perspective, the prevalence and numbers of this pathogen on food commodities is a clear reflection of consumer exposure to the pathogen.       1.1.4.1 Meat products S. enterica infections are highly related to poultry and poultry products and these animals are the primary source of salmonellosis. A wide variety of animals, particularly food animals, have been identified as reservoirs for nontyphoidal Salmonella spp. (Aarestrup, 2007). S. enterica is a common pathogen contaminating raw meat and is hazardous to 	   6 human if the meat is not thoroughly cooked (Prendergast et al., 2009). S. enterica infections are often spread through international trade in animal feed, live animals and non-heated animal food products (Forshell and Wierup, 2006). Raw meat or uncooked poultry meat products possibly pose a huge hazard to public health, reported by World Health Organization (WHO) (WHO, 1995), although situations are likely to vary between countries, according to different levels of S. enterica contamination and patterns of consumption (Forshell and Wierup, 2006). Contamination of poultry or poultry meat may occur throughout the whole production chain and important risk factors for contamination at each stage of this process has been identified (Heyndrickx et al., 2002; Mead, 2004). The most common serotypes for pork are Uganda and Infantis while chicken is the most common food commodity for serotype Typhimurium (Jackson 2013). The prevalence of S. enterica on retail pork has been evaluated in many laboratory studies by the Republic of Ireland (Prendergast et al., 2009). Salmonella spp. was discovered from raw pork sausages (pre-packed and loose) between 2001 and 2002 at prevalence’s of 2.9% (27/921) (Boughton et al., 2004). Jordan et al. recovered Salmonella spp. in raw pork samples in 2002, 2003 and 2004 at prevalence's of 2.3% (160/6,823), 2.0% (136/6,638) and 2.1% (158/7,683), respectively (Jordan et al., 2006). It is increasingly accepted that the prevalence of Salmonella in animal production must be decreased and, in the European Union, plans to achieve this are currently being implemented. However, more works on Salmonella spp. on animal meat at retail are still required due to the limited quantitative data, which is crucial for quantitative microbial risk assessment (QMRA) models and the development of strategies to reduce risk from this pathogen (Malorny et al., 2008; Prendergast et al., 2009; Leuschner et al., 2010). 	   7       1.1.4.2 Egg and dairy products Besides poultry and meat products, egg and dairy products (e.g., shell egg, raw milk, and unpasteurized cheese) are also regarded as common implicated food vehicles for S. enterica. One hundred and twelve (28%) outbreaks were reported to be associated with eggs in United States in 2013 (Jackson et al., 2013). Enteritidis and Heidelberg are two major serotypes causing the outbreaks associated with eggs. For example, a high (83%) proportion of egg-associated outbreaks were caused by serotype Enteritidis in United States from 1998 to 2008 (Jackson et al., 2013). Serotype Heidelberg also contributes a high percentage of egg-associated outbreaks supported by findings from case-control studies and previous reviews (Hennessy et al., 2004; Chittick et al., 2006). S. enterica was also associated with several raw milk and cheese outbreaks during 2004 ~ 2011 (Ebel et al., 2016). About sixteen dairy-associated outbreaks were reported in United States during 1998 ~ 2008, of which most were caused by Salmonella Typhimurium (56%) and Newport (25%) (Jackson et al., 2013). S. enterica, particularly Salmonella Typhimurium and Dublin, are commonly found in cattle and are excreted in the feces (McClelland and Pinder, 1994). This provides an easy route of contamination during milking and milk processing. The consumption of contaminated, unpasteurized milk or products in which unpasteurized milk is used as an ingredient may cause lethal Salmonella infections in human.       1.1.4.3 Vegetable and fruit produce It is generally perceived that consumption of raw fresh produce products is safe and that most foodborne disease outbreaks are primarily caused by foods of animal origin 	   8 (Sivapalasingam et al., 2004). However, in the recent years several studies have assessed S. enterica prevalence on produce (Table 1.1). Many S. enterica serotypes were implicated in salmonellosis outbreaks associated with raw and minimally processed fruit and vegetables, mainly including Litchfield, Poona, Oranienburg and Javiana depending on the category of fruit or vegetable items (Prendergast et al., 2009). Produce-related salmonellosis outbreaks will probably increase in the future with the increase consumption of various forms of fresh vegetable and fruits around the world (Davis, Elaine et al., 2009). A large number of salmonellosis outbreaks are found to be associated with tomatoes, seed sprouts, serrano and jalapeno peppers, and cucumbers (see Table 1.1). These outbreaks have usually been linked to fecal contamination of fresh produce during postharvest handling, shipping, or processing in circumstances that permitted bacterial multiplication (Edelstein, 2009). Guo Xuan et al. recovered that most common serotypes causing tomato-associated infections are Newport (~32%), followed by Typhimurium (~16%), Braenderup (~11%), Enteritidis (~11%), and Javiana (~11%) (Guo Xuan et al., 2001). Seed sprouts are reported to be the most common food commodity for serotype Mbandaka, Typhimurium and Agona (Mahon et al., 1997; Kocharunchitt, Ross and McNeil, 2009). A study was conducted in 2006 to determine the prevalence of S. enterica on a farm that cultivates chili peppers in Mexico and both Salmonella Typhimurium and Enteritidis serotypes were found associated with the chili pepper production system (Gallegos-Robles et al., 2008). However, limited previous studies were conducted on fresh cucumbers, which caused multistate outbreaks from 2013 to 2015. Therefore, this study provided an insight on the characteristics and potent factors contributing to the survival of S. enterica serotypes. 	   9  Table 1.1 Selected outbreaks of salmonellosis associated with the consumption of fresh produce. Year Salmonella  enterica serotype Produce commodity Reference 1990 Javiana Round tomatoes Sivapalasingam et al., 2004 1991 Poona Cantaloupe CDC, 1991 1993 Montevideo Round tomatoes CDC, 1993 1995 Stanley Alfalfa sprouts Mahon et al., 1997 1996 Montevideo Alfalfa sprouts Taormina et al., 1999 1995-1996 Newport Alfalfa sprouts Van Beneden et al., 1999 1998 Havana Alfalfa sprouts Taormina et al., 1999 1999 Baildon Round tomatoes Cummings et al., 2001 1999 Newport Mango Sivapalasingam et al., 2003 2000-2001 Enteritidis Mung sprouts Mohle-Boetani et al., 1009 2001 Saintpaul Mango Beatty et al., 2004 2001 Poona Cantaloupe CDC, 2002 2002 Newport Round tomatoes Greene et al., 2008 2004 Braenderup Roma tomatoes Gupta et al., 2007 2005 Newport Round tomatoes Greene et al., 2008 2006 Typhimurium Round tomatoes CDC, 2007 2008 Saintpaul Serrano and jalapeño peppers CDC, 2008; Barton et al., 2011 2009 Saintpaul Alfalfa sprouts CDC, 2013 2010 Unknown Alfalfa sprouts CDC, 2011 2011 Agona Papayas CDC, 2011 2012 Typhimurium, Newport Cantaloupe CDC, 2012 2013 Saintpaul Cucumbers CDC, 2013 2014 Newport, Enteritidis Cucumbers, Bean Sprouts Angelo et al., 2015; CDC, 2015 2015 Poona Cucumbers CDC, 2016 2016 Muenchen Alfalfa Sprouts CDC, 2016  	   10        1.1.5 Factors contributing to the survival of S. enterica on fresh produce Understanding the factors contributing to the survival of S. enterica and their ability to cause disease would enable researchers to prevent suffering and economic losses caused by this pathogen. Environmental factors including contaminated water sources used to irrigate and wash produce crops have been associated with a large number of outbreaks (Steele and Odumeru, 2004). S. enterica is able to attach to the surface as well as internalize into fruits and vegetables, which is probably the reason that a lot of preventative efforts are focusing on limiting the initial contamination (Fatica et al., 2011). S. enterica is adapted to food-related environments and can survive under a wide range of pH and temperatures encountered in various food products (Alford and Palumbo, 1969). Most S. enterica serotypes can grow at temperatures ranging from 7 °C to 48 °C (Pui et al., 2011). There are also some rare serotypes that can grow at temperatures as low as 4 °C (Helmuth et al., 1985; Pui et al., 2011), although the general understanding is that the growth of S. enterica is substantially reduced bellow 10 °C. (Sreedharan et al., 2014). The optimum pH range for the growth of S. enterica is between 6.5 and 7.5, whereas some serotypes can also survive at pH values as low as 3.7 or as high as 9.5 (Podolak et al., 2010). Adaptability of S. enterica to the temperature, pH or moisture on the produce surface as well as interactions with produce (e.g. attachment and persistence ability) are two key factors contributing to produce-linked outbreaks in plant environment (Bartz et al., 1981; Sivapalasingam et al., 2003).  	   11  1.2 Microbial biofilms        1.2.1 Characteristics of biofilms A biofilm is a consortium of microorganisms in which cells associate with each other and adhere onto a surface (Ghannoum and O’Toole, 2004). Microorganisms develop biofilms comprising microbial cells and extracellular polymeric substances (EPS) after attaching to surfaces, which is a complex process and regulated by diverse characteristics of the growth medium, substratum, and cell surface (Donlan, 2002; Pang et al., 2005). A prominent function of biofilms is to provide an optimal environment for the exchange of genetic material between cells (Donlan, 2002). Van Leeuwenhoek revealed that surface-associated microorganisms (biofilms) that attach to and grow universally on exposed surfaces demonstrated a distinct phenotype with respect to gene transcription and growth rate (Donlan, 2002; Donlan and Costerton, 2002; Hall-Stoodley, Costerton and Stoodley, 2004). The mechanisms for initial attachment of these microorganisms to a surface, detachment, development of a community structure and ecosystem are specific, constituting a protected mode of growth that allows survival in hostile environment (Donlan, 2002; Donlan and Costerton, 2002). Biofilm-associated microorganisms differ from their planktonic (freely suspended) counterparts in their growth rate, bacteria composition and increased resistance to biocides, antibiotics and antibodies with respect to the transcribed genes (Donlan and Costerton, 2002; Jain et al., 2007). Biofilms can form on various living or non-living surfaces (Hall-Stoodley, Costerton and Stoodley, 2004), like living tissues, industrial or 	   12 potable water system piping, indwelling medical devices, or natural aquatic systems (Parsek and Singh, 2003; Percival et al., 2011). Components such as water, polysaccharides and other macromolecules contribute not only for the heterogeneity of the matrix but also for its multicellular function (Prakash, Veeregowda and Krishnappa, 2003). There are two major components of a biofilm, namely the sessile bacterial cells and a matrix of self-produced EPS (Li et al., 2017). EPS is considered the primary matrix material of the biofilm, accounting for 50% to 90% of the total organic carbon of biofilms (Flemming et al., 1999). Although EPS may vary in chemical and physical properties, its primary composition is polysaccharides (Donlan, 2002; Vu et al., 2009). EPS are anionic in some gram-negative bacteria due to the presence of uronic acids (such as D-glucuronic, D-galacturonic, and mannuronic acids) or ketal-linked pryruvates, allowing the association of divalent cations such as calcium and magnesium, which have been shown to cross-link with the polymer strands and provide greater binding force in a developed biofilm (Kokare et al., 2009). However, the chemical composition of EPS is primarily cationic, which may be quite different in some gram-positive bacteria (Hussain, Wilcox and White, 1993; Altaf et al., 2017; Ahmad et al., 2017). The readily observable characteristics of biofilm include that cells irreversibly attached to a surface or interface, embedded in a matrix of EPS produced by these cells and exhibited an altered phenotype with respect to growth rate and gene transcription (Donlan and Costerton, 2002; Thomas, Ramage and Lopez-Ribot, 2004). In nature, bacterial biofilms allow the bacteria to survive in hostile environments via constituting a protected growth modality and colonizing any humid surface (Prakash, Veeregowda and Krishnappa, 2003). Occasionally, these bacterial aggregates release individual cells that disperse and 	   13 rapidly multiply, thereby colonizing other places (Prakash, Veeregowda and Krishnappa, 2003; Mampel et al., 2006).       1.2.2 Mechanism of biofilm formation The process of biofilm development is complex, including specific steps (e.g., initial adhesion, microcolony production, three-dimensional community structure development and maturation as well as detachment (Donlan, 2002; Prakash, Veeregowda and Krishnappa, 2003). Detailed examination of the mechanisms of biofilm formation have already been done by the scientific and engineering community (Pedersen, 1990; Donlan and Costerton, 2002; Lappin-Scott et al., 2003). Interestingly, bacteria form biofilms preferentially in very high shear environments (i.e., rapidly flowing milieus) (Donlan and Costerton, 2002). The motility of bacteria is slowed via approaching the surface very closely and bacteria form a transient association with the surface and/or other microbes previously attached to the surface (O'Toole, Kaplan and Kolter, 2000; Watnick and Kolter, 2000). Smooth surfaces might escape bacterial colonization (Donlan and Costerton, 2002; Shunmugaperumal, 2010). In other words, surfaces that are rougher, more hydrophobic provide a more ideal environment for the attachment and growth of microorganisms (Knobloch et al., 2001). Besides, attachment will occur most readily on the solid-liquid interface between a surface and an aqueous medium (e.g., water and blood) (Costerton, Stewart and Greenberg, 1999; Prakash, Veeregowda and Krishnappa, 2003). And the attachment flow may be affected by velocity, water temperature and nutrient concentration once these factors do not exceed critical levels (Melo and Bott, 1997; Donlan, 2002).  	   14 Microcolonies begin to form after the bacteria attach to the inert surface/living tissue. The bacteria multiply within the embedded exopolysaccharide matrix and emit chemical signals that intercommunicate among the bacterial cells (Prakash, Veeregowda and Krishnappa, 2003; Jamal et al., 2015), which activate the genetic mechanisms underlying EPS production and give rise to the formation of a microcolony (Landini et al., 2010). Subsequently, the transcription of specific genes required for the synthesis of EPS occurs and 3-dimensional structures are formed, resulting in a complex biofilm (Davey and O'toole, 2000). Occasionally, some bacteria are shed from the colony or stop producing EPS for purely mechanical reasons (Prakash et al., 2005; Saikia et al., 2005), leading to dispersal and detachment (or shedding) of bacterial cells and shearing of biofilm aggregates (continuous removal of small portions of the biofilm) because of flow (Donlan, 2002; Malic, 2008). And the phenotypic characteristics of the organisms are greatly affected by the mode of dispersal. In another words, biofilm development is a dynamic process of growth and detachment (or shedding) of bacterial cells and aggregates, which can lead to the ingestion or inhalation of a condensed infective dose (Stoodley et al., 2001; Cogan et al., 2016).       1.2.3 Impact of biofilms on public health Biofilms can form on various surfaces, including biotic surface and abiotic surface. Their high prevalence and resistance to antimicrobial treatments lead to significant public health concerns, reported by Canadian Antimicrobial Resistance Surveillance System in 2016. A large number of studies of infectious disease processed from a biofilm perspective have been conducted based on the recognition that microbial biofilms are ubiquitous in nature (Davey and O'toole, 2000; Rinaudi and Giordano, 2010; Faria, Joao 	   15 and Jordao, 2015). Bacterial biofilms colonize on any humid surface, causing cystic fibrosis, native valve endocarditis, otitis media, periodontitis or chronic prostatitis, and are prevalent on most wet surfaces in nature, resulting in environmental problems (Costerton, Stewart and Greenberg, 1999; Cortés, Bonilla and Sinisterra, 2011). Nevertheless, Nickel and Costerton concluded that there was not definitive evidence that the infection resulted from these organisms (Nickel and Costerton, 1992; Donlan and Costerton, 2002). All that could be stated was that there was an association between the presence of the organisms and the disease (Donlan and Costerton, 2002). For several of the diseases, such as periodontitis, native valve endocarditis, and cystic fibrosis, that association is stronger. For others, such as otitis media, the association is less well established (Martínez et al., 2006). Epidemiologic evidence illustrates that biofilms play a role in infectious diseases, both for specific conditions such as cystic fibrosis and periodontitis and in bloodstream and urinary tract infections as a result of indwelling medical devices (Parsek and Singh, 2003). The resistance of bacteria is increased with the growth of biofilm, which may make organisms less conspicuous to the immune system (Lewis, 2001). However, in many human infections, the bacteria are difficult to access and biofilm and planktonic growth may coexist (Parsek and Singh, 2003; Hall-Stoodley et al., 2012). The characteristics of biofilms play an important role in infectious disease processes (Donlan, 2002; Aguilar-Romero et al., 2010). Additionally, the role of biofilm in antimicrobial drug is also of importance from a public health perspective (Donlan, 2001). Preventing biofilm formation rather than applying treatment to	  eliminate is generally recommended because the resistance of microbes residing in the biofilms towards different types of 	   16 antimicrobial agents poses a serious threat to the pharmaceutical industries (Ryder, 2005;	  Kokare et al., 2009; Lebeaux et al., 2013).       1.2.4 Factors affecting biofilm formation Biofilm may be formed on a wide range of surfaces (Morton and Surman, 1994). The biofilm system is highly complex and affected by many factors.  Biofilm formation ability is related to bacterial attachment ability, which could be influenced by a number of factors, including the surrounding environment and cell surface properties. For instance, as the surface roughness increases, the extent of microbial colonization tends to increase, which contributes to the maximum attachment (O'toole and Kolter, 1998; Krivorot et al., 2011; Yu et al., 2016). The degree of attachment was found to depend on a number of factors, including the nature of the substrate, pH, electrolyte concentration, ionic charge of the polymer, and bacterial strain tested (Douglas, 2003; Kokare et al., 2009; Shunmugaperumal, 2010). The attached bacterial cells are enclosed in a self-produced polymeric matrix, which makes biofilm a very dangerous biological structure in attachment because biofilm improves their ability to colonize and survive in a harsh condition (Monier and Lindow, 2003) and can become a persistence source of contamination (Costerton et al., 1999; Houdt and Michiels, 2010). Chemical characteristics of aqueous medium such as nutrient level, ionic strength, temperature, etc. could possibly provide competitive advantages for the rate of microbial attachment of one organism (Kokare et al., 2009; Van Houdt and Michiels, 2010). Gram-negative bacteria, such as Salmonella, respond to nutrient limitation and other environmental stresses by synthesizing sigma factors (Watson, Clements and Foster, 	   17 1998). Furthermore, hydrodynamic factors, like flow rate, presence of shear, batch versus open system, retention time can also affect the ability of bacteria to form biofilms and are considered to be important in the development of a model biofilm system (Shunmugaperumal, 2010). It is found that bacteria have greater adherence to hydrophobic surfaces and lenses composed of nonionic polymers (Dutta, Cole and Willcox, 2012) as bacteria with increased hydrophobicity possess reduced repulsion between the extracellular matrix and the bacterium (Donlan, 2002). Lastly, properties of the cell surface, specifically the presence of fimbriae, flagella and surface-associated polysaccharides or proteins are conducive to the bacterial communities where bacterial biofilms are produced (Donlan, 2002;	  Bogino et al., 2013).  1.3 Major biofilm components involved in rdar morphotype        1.3.1 The role of curli in biofilm formation Biofilm formation by Salmonella has only recently been investigated. The production of thin aggregative curli fimbriae (in another words, curli) by virulent strains was previously documented in some early work but a conclusive role for these fibers was not elucidated (Römling et al., 1998; Solomon et al., 2005). The production of curli was later found to be an important component in the formation of an extracellular matrix by cells of S. enterica (Römling et al., 2000; Cookson, Cooley and Woodward, 2002; Solano et al., 2002). Contamination of surfaces due to microbial attachment occurs in many environments and may create serious economic and health problems associated with food spoilage and disease transmission (Costerton, Stewart and Greenberg, 1999). Initial bacterial colonization of surfaces is reversible and may be mediated by surface-expressed 	   18 appendages such as curli (Pratt and Kolter, 1998) and flagella (Pratt and Kolter, 1998; O'Toole and Kolter, 1998). Curli, the major proteinaceous component of a complex extracellular matrix produced by many Enterobacteriaceae were first discovered in the late 1980s on Escherichia coli (E. coli) strains that caused bovine mastitis, and have been implicated in many physiological and pathogenic processes of E. coli and S. enterica (Barnhart and Chapman, 2006). Curli fibers play a role in adhesion to surfaces, cell aggregation, biofilm formation and mediate host cell adhesion and invasion as they are potent inducers of the host inflammatory response (Barnhart and Chapman, 2006; Kline et al., 2009). Curli showed a unique structure and biogenesis among bacterial fibers (Jonson, Normark and Rhen, 2005; Barnhart and Chapman, 2006). Curli was structurally and biochemically categorized under a growing class of fibers known as amyloids, which is involved in several human diseases (e.g., Alzheimer’s, Huntington’s, and prion diseases) (Barnhart and Chapman, 2006; Tian et al., 2014). Different morphotypes were expressed by different enteric bacteria, which correspond to differences in the extracellular matrix that they produce (Bokranz et al., 2005; Barnhart and Chapman, 2006) and can be visualized by growing bacteria on CR-indicator plates. Curli play an important role in the initial stages of biofilm development, which is a multi-step developmental process in both E. coli and S. enterica (Blanco et al., 2012; Chambers and Sauer, 2013). In the food industry, foods get infected after contacting with contaminated Teflon and stainless steel where S. enterica adhere to and form biofilms with the help of curli (Austin et al., 1998; Barnhart and Chapman, 2006; Van and Michiels, 2010; Giaouris et al., 2012). Interestingly, cellulose-deficient strains were 	   19 discovered only among produce isolates (Solomon et al., 2005). Barak et al. found that S. enterica adhered more strongly to alfalfa sprouts than did E. coli O157:H7 and postulated that this resulted from the lack of curli production by E. coli O157: H7 (Barak, Whitehand and Charkowski, 2002). Besides, they reported that eighty percent of the produce-related isolates produced curli and all of the produce-related strains that did not produce curli were from produce-related outbreaks rather than collected food samples, which may further underscore the importance of curli in attachment to produce surfaces (Barak, Whitehand and Charkowski, 2002). Another study suggested that biofilms formed by curli-proficient strains and curli-deficient strains are different in their morphology (Barnhart and Chapman, 2006). Curli-deficient strains form flat biofilms on polyurethane sheets, while curli-expressing strains produce mature biofilms (Kikuchi et al., 2005).       1.3.2 The role of cellulose in biofilm formation Cellulose is a second component of the S. enterica extracellular matrix (ECM) involved in bacterial biofilm formation and attachment to surfaces and responsible for the rdar phenotype (Uhlich, Cooke and Solomon, 2006). Although the most abundant source of cellulose (Augimeri, Varley and Strap, 2015) is plant cellulose, which is the main constituent in the cell wall of vascular plants (Somerville, 2006), cellulose synthesis has been observed in numerous microorganisms, such as green algae and oomycetes that use cellulose in their cell walls, as well as various bacterial species (Richmond, 1991; Fugelstad et al., 2009). The most thoroughly characterized organism with respect to cellulose synthesis is A. xylinum, which include Pseudomonas, Achromobacter, Alcaligenes, Aerobacter, Azotobacter, Agrobacterium, Rhizobium, and Sarcina species 	   20 (Ross, Mayer and Benziman, 1991). The primary function of bacteria cellulose-containing biofilms is to establish close contact with a preferred host to facilitate efficient host-bacteria interactions (Augimeri, Varley and Strap, 2015). Bacteria cellulose producers display a pathogenic or symbiotic relationship with animal, plant or fungal hosts according to the environment (Douglas, 2003; Augimeri, Varley and Strap, 2015). It is surprising that although bacteria cellulose is not essential for survival, it does provide a survival advantage for bacteria (Augimeri, Varley and Strap, 2015). Biofilms containing bacteria cellulose can facilitate host-bacteria interactions (Augimeri, Varley and Strap, 2015).  Due to the development of advanced technologies and the increased availability of genetic tools (e.g., next-generation sequencing technology), research regarding cellulose synthesis among a wide variety of microorganisms has been increased steadily during the last decades (Augimeri, Varley and Strap, 2015; Römling and Galperin, 2015). However, there is still much to learn regarding the environmental interactions mediated by bacteria cellulose. The ecological diversity of bacteria cellulose-producing organisms reveals the importance of bacteria cellulose as a mediator of environmental interaction (Römling and Galperin, 2015). Future studies should be geared toward further elucidating bacteria cellulose-mediated environmental interactions to provide novel insights into the mechanisms that control bacteria cellulose biosynthesis.       1.3.3 Congo red (CR) binding assays Bacteria can escape from deleterious conditions when surviving in biofilms (Davey and O'toole, 2000). Therefore, the formation of biofilms tends to be a crucial factor in the 	   21 disease cycle of bacterial pathogens in both animals and plants (Bogino et al., 2013). In recent years, many studies have been conducted on the role of bacterial surface components in combination with bacterial functional signals in the process of biofilm formation (Bogino et al., 2013). Investigation of bacteria biofilm can be carried out using various phenotypic methods. One method is based on culture in brain heart infusion (BHI) agar containing sucrose and red Congo dye (original Congo red agar) developed by Freeman et al. in 1989 (Freeman, Falkiner and Keane, 1989). Due to the visual analysis of the color of the colonies that grow on the agar, this method is considered to have low accuracy, but it is really cheap and easy to perform. This method can be modified with the addition or substitution of some substances or the modification of some parameters (Mariana et al., 2009; Los et al., 2010), which improves the accuracy of detecting biofilm produced by various S. enterica strains. Curli fimbriae and cellulose are known to be important components of the biofilm extracellular matrix. The co-expression of thin aggregative fimbriae and cellulose leads to an aggregative colony phenotype (red, dry, and rough [rdar]), which is a multicellular behaviour in bacteria when grown on medium containing the dye Congo red (Solano et al., 2002). The rdar morphotype is the best-studied form of S. enterica multicellular behavior with respect to regulation and exopolysaccharide (EPS) composition (Romling, 2005). Curli fimbriae and cellulose play a synergetic role in biofilm formation (Saldana et al., 2009). The production of cellulose and curli by S. enterica leads to a matrix of tightly packed cells covered in a hydrophobic network, which is important in biofilm formation as well as in its persistence on various surfaces (Solomon et al., 2005). Curli production and cellulose production can be monitored by assessing morphotypes on Luria-Bertani 	   22 agar without salt containing Congo red and by assessing fluorescence on Luria-Bertani agar containing calcofluor, respectively (Solomon et al., 2005). The Congo red dye directly interacts with certain polysaccharides, forming colored complexes. Isolates were grouped into five distinct morphotypes (Solomon et al., 2005): (a) red, dry, and rough (rdar), indicating curli and cellulose production; (b) brown, dry, and rough (bdar), indicating curli production but a lack of cellulose synthesis; (c) smooth and red (sar), indicating cellulose production but a lack of curli synthesis (Castelijn et al., 2012); and (d) smooth and white (saw), indicating a lack of both curli and cellulose production. The incidence of cellulose production and curli biosynthesis is different for isolates from a variety of sources (Bokranz et al., 2005). The production of both cellulose and curli is important for the survival and persistence of S. enterica on surface environments (Römling, 2002; Römling et al., 2003). The role of biofilm formation, curli, and cellulose in establishing bacteria on the surface of fruits and vegetables must be characterized in order to put forth more effective postharvest intervention treatments. 1.4 Research purpose The purpose of this study was to evaluate the survival characteristics of S. enterica strains at different temperatures and analyze the factors contributing to their survival. This project comprises three objectives: (i) to examine the survival and growth of five S. enterica strains on mini cucumbers at 22 ± 2 °C and 4 ± 2 °C; (ii) to quantify the biofilm formation and attachment ability of the five S. enterica strains; and (iii) to examine the S. enterica cell morphology by Congo Red Agar (CRA) method.  	   23  Chapter 2: Survival of S. enterica on mini cucumbers 2.1 Introduction Although eggs and poultry are still two leading food products that contribute to salmonellosis outbreaks, accounting for nearly half of the cases, a growing range of fresh produce have been implicated in S. enterica infection (Guo et al., 2016). Fresh produce may be exposed to soil, insects, animals, water, and/or humans during growing or harvesting and in processing plants (Ukuku, 2004). Additionally, crops may be fertilized with the manure from domestic animals. Consequently, produce intended for raw consumption may be exposed to pathogenic bacteria, parasites, and viruses from animal excreta (birds, insects, rodents, and reptiles), from water (irrigation and rain), or from infected workers, manure, and food-processing facilities with poor sanitation (Wei et al., 1995). In order to start addressing these issues, the survival of S. enterica on fresh produce needs to be determined. Cucumbers have been identified as the cause for recent salmonellosis outbreaks with the increasing consumption of fresh produce. One S. enterica outbreak was traced each year to the consumption of contaminated cucumbers in USA since 2013, viz., Salmonella Saintpaul infections in 2013 (CDC, 2013), Salmonella Newport infections in 2014 (Angelo et al., 2015) and Salmonella Poona infections in 2015 (CDC, 2015), which caused nearly 2000 human infections and four deaths. The fourth S. enterica outbreak linked to fresh cucumbers since 2013 sickened people in at least eight states. In this study Salmonella Newport and Poona were two of the strains examined for both their survival 	   24 capability and characteristics. Previous studies mainly focused on the survival of S. enterica on tomatoes, seed sprouts, serrano and jalapeno peppers but little on cucumbers, which also implicated in numerous outbreaks of S. enterica in recent years. Many environmental factors contribute to the survival of S. enterica on fresh produce surface, including contaminated water sources used to irrigate and wash produce crops, pH, temperature, nutrition, oxygen etc. In addition, the ability of S. enterica to attach or internalize into produce may also be a factor that make these produce items more likely to be sources of S. enterica contamination. We hypothesized that S. enterica is able to survive for extended days on cucumbers. In addition, S. enterica strains will exhibit differential survival capacities on mini cucumbers during storage at these two temperatures. Therefore, the research objective was to examine the survival and growth of five S. enterica strains on mini cucumbers at 22 ± 2 °C and 4 ± 2 °C, which are two commonly used storage temperature conditions in practice.  2.2 Materials and methods        2.2.1 Origin and maintenance of S. enterica strains Five S. enterica strains were used in this project (Newport, Typhimurium, Enteritidis, Daytona, Poona). The stock cultures were maintained in Brain-Heart-Infusion (BHI) broth (BD, Difco, East Rutherford, NJ) supplemented with 15% glycerol and held at -80 °C. For use, strains were aseptically streaked onto Luria-Bertani (LB) agar and incubated at 37 °C for 24 ± 2 hours. Cultures were stored on LB agar plates at 4 °C up to one month, using individual colonies in subsequent experiments.  	   25 Table 2.1 S. enterica strains used in this study. Serotype Isolate Origin Strain ID Newport Human FSL S5-639 Typhimurium Irrigation water LMFS-S-JF-001 Enteritidis Irrigation water  LMFS-S-JF-005 Daytona Irrigation water LMFS-S-JF-009 Poona Beef roast S306        2.2.2 Preparation of strains and mini cucumbers A single colony from LB agar plates, stored at 4 °C, was aseptically inoculated into 10 ml of Brain-Heart-Infusion (BHI) broth (Becton, Dickinson And Co.) using a bacteriological loop. Each strain was inoculated into three separate BHI broth tubes to represent three biological replicates, as well as one un-inoculated broth tube as a negative control. Inoculated cultures were incubated in a shaking incubator (Thermo Fisher Scientific, Waltham, MA, USA) at 37 °C and 170 rpm until reaching stationary phase (18h). The optical density of overnight cultures was measured at 600 nm using a spectrophotometer (Shimadzu Corp., Kyoto, Japan) and used to adjust the cell concentration for subsequent experiments (Sun et al., 2001). Intact mini cucumbers without any physical processing were used to inoculate S. enterica strains because people usually eat cucumber slices in ready-to-eat cold dishes without peeling and the fruit peel is where most easily to be contaminated by S. enterica. Fresh mini cucumbers used in this assay were purchased from a local supermarket in Vancouver, BC. The whole mini cucumbers were weighed individually and washed in running water for 5 s each to remove any observable dirt. The cucumbers were then 	   26 placed in sterile Whirl-Pak® bags (Nasco, Fort Atkinson, Wisconsin, USA) and dried in a type A2 biological safety cabinet (Esco, Portland, OR, USA) overnight with the bags open.       2.2.3 Mini cucumber inoculation and recovery of S. enterica One mL of overnight culture with adjusted cell concentration (~1011 CFU/mL) was transferred from each tube into each centrifuge tube. The cells were centrifuged at room temperature for 10 min at 5000 x g (Model 5424R, Thermo Fisher, Waltham, MA, USA) and the supernatant was subsequently decanted. The pellet was washed twice with phosphate buffered saline (PBS) (Amresco) and suspended in 1 mL of sterile PBS. Inoculum was diluted 100:1 fold in PBS and 100 µL of the diluted bacterial culture was inoculated on cucumbers by pipetting small droplets on the surface. Then all cucumbers were air-dried for an hour in a type A2 biological safety cabinet (Esco, Portland, OR, USA) in order to allow the cells to adhere to the surface. Samples were subsequently stored at 22 ± 2 °C or 4 ± 2 °C in an incubator (Thermo Fisher, Waltham, MA, USA) for 8 and 19 days, to mimic the maximum shelf life of cucumbers at room temperature and refrigeration temperature, respectively (Khattak et al., 2005). To obtain initial cell counts, suspensions were serially diluted with PBS, spread plated in duplicate onto BHI agar, and colonies were counted after 24 ± 2 hours incubation at 37 °C. Cell densities were assessed at day 0, 2, 4, 6, 8 for 22 ± 2 °C and 0, 3, 5, 9, 12, 15, 17, 19 for 4 ± 2 °C. At each sampling time, 25 mL of PBS was transferred into each sampling bag to wash cells from cucumber surfaces and the suspensions in the sampling bags were serially diluted in PBS. Subsequently, 100 µL of the diluted suspension was spread onto BHI-XLD agar (Difco, East Rutherford, New Jersey, USA) in duplicates using Thin Agar Layer (TAL) 	   27 method. Plates were incubated at 37 °C for 24 ± 2 hours.       2.2.4 Colony enumeration using Thin Agar Layer (TAL) method The survival of individual S. enterica strains on mini cucumbers was monitored using the one-step Thin Agar Layer method, which was further improved by Kang and Fung et al. to recover injured pathogens (Kang and Fung, 2000; Wu and Fung, 2001; Wu et al., 2001a, b). This modified method was performed by pre-pouring selective medium (XLD, ca. 25 mL) and overlaying it with 14 mL of non-selective medium (BHI); see Figure 1. The top layer (BHI) provides a favorable environment for injured cells to resuscitate and become functionally normal in the first few hours of incubation, while the bottom layer (XLD) selectively facilitates or inhibits typical reactions of certain microorganisms. Besides, the non-selective medium (BHI) does not hinder typical color of colonies produced by target microorganisms, making this novel one-step procedure a significant improvement in comparison to the traditional cumbersome two-step overlay (OV) methods (Kang and Fung, 1999, 2000; Wu and Fung, 2001; Wu et al., 2001a, b).      Fig. 2.1 Schematic illustration of the modified one-step Thin Agar Layer method (Wu and Fung, 2001; Wu et al., 2001a, b).        	  Inoculation of injured microorganisms directly on non-selective thin agar layer plate	   	   	   	   	  	  	  25 mL of selective medium (XLD)	  14 mL of non-selective medium (BHI)	  	  	  	  	   28       2.2.5 Data and statistical analysis Surviving populations of S. enterica were reported as the mean CFU on individual cucumber using three replicates. Two parameters were used to assess survival of the S. enterica strains: (i) the difference in log CFU between first day and last day of sampling (Δlog CFU), which evaluates the cell proliferation at the end of the inoculation period; (ii) the maximum achieved density following initial inoculation (Nmax), which indicates the relative growth and death fluctuation rates. The one-way analysis of variance models, using each of the two parameters as the response, were carried out to evaluate the contributions of the strain and food matrix. A P value of < 0.05 was considered statistically significant. Statistical analyses were performed using JMP version 11.1.1 (SAS Institute, Inc., Cary, NC). 2.3 Results and discussion To evaluate the survival characteristics of five S. enterica strains on mini cucumbers, ~ 107 CFU of each strain was spotted onto cucumbers in triplicate and incubated either at 22 ± 2 °C or 4 ± 2 °C. The inoculated cucumbers were incubated for up to 8 days at 22 ± 2 °C and every 2 days cucumbers were sampled for S. enterica by washing the surface with PBS and plating onto BHI-XLD agar. The cucumbers held at 4 ± 2 °C were sampled every 3 days for up to 19 days.  Five different S. enterica strains (Newport, Typhimurium, Enteritidis, Daytona, Poona) were used in the study. The initial cells inoculated on cucumber surfaces are 7.83 ± 0.42 log CFU. The large number of cells applied to the mini cucumbers may not reflect 	   29 realistic terms levels of contamination that may occur in the environment. Nevertheless, contamination caused by contact with fecal material containing large populations of S. enterica could occur (Guo, Xuan, et al., 2001).       2.3.1 Suvival or growth (Δlog CFU) of S. enterica on mini cucumbers Δlog CFU is the difference of log CFU between the last day and the first day of sampling, which illustrates the variance of cell population at the end of the period. Specifically, positive Δlog CFU means the cell number of S. enterica increased at the end, while negative Δlog CFU means the cell number of S. enetrica decreased at the end. At 22 ± 2 °C, only Salmonella Poona (S. Poona) and Salmonella Daytona (S. Daytona) increased in populations at last day. S. Poona increased most obviously at the last day with the highest Δlog CFU of 0.84 ± 0.01 (Table 2.2), followed by S. Daytona (Δlog CFU ~ 0.10 ± 0.01). Salmonella Typhimurium (S. Typhimurium), Salmonella Newport (S. Newport) and Salmonella Enteritidis (S. Enteritidis) all experienced a decline in cell numbers. However, these decrease was minimal with the Δlog CFU of -0.23 ± 0.01, -0.20 ± 0.01 and -0.17 ± 0.01, respectively.    	   30  Fig. 2.2 Survival of Salmonella strains on mini cucumbers stored at room temperature (22 ± 2 °C, a) and at refrigeration temperature (4 ± 2 °C, b). Bacterial cells were recovered on BHI-XLD agar. Error bars indicate the standard errors of the means of three replicates.  However, at 4 ± 2 °C, the Δlog CFU of these five S. enterica strains are all negative, indicating that the cell numbers of these five strains all decreased at the last day. At refrigeration temperature, S. Poona exhibited the largest decrease in cell population (~ -1.82 ± 0.01 of Δlog CFU). This was followed by S. Typhimurium, S. Newport and S. Daytona. S. Enteritidis showed the smallest decrease in cell population with the decrease of 0.91 ± 0.01 log CFU, highlighting the diversity of strain-specific responses to cold stress. S. Enteritidis was the most resistant to cold stress encountered on cucumber surface with the smallest cell reduction of -1.64 ± 0.01 log CFU. The cold resistance of S. enterica has not been extensively studied and these results revealed the potential risk of (a) 22 ± 2 °C 	   (b) 4 ± 2 °C 	  Newport  Typhimurium Enteritidis  Daytona  Poona  	   31 these particular strains to food industry. S. Poona, which survived the best at room temperature in this study was also one of the serotypes implicated in a 2015 cucumber outbreak in United States (CDC, 2015). The less resistant strain, S. Newport, has also been reported in literature to be capable of causing cucumber contamination. A multistate cluster of S. Newport infections was reported in August 2014 causing 275 cases reported from 29 states (Angelo et al., 2015). In one previous study, S. Newport was inoculated on cucumber surface and stored at 10 or 22 °C. It was found that between day 0 and 1, populations on cucumbers without treatment stored at 10 and 22 °C declined by 2.47 and 1.07 log CFU per cucumber, respectively (Sharma et al., 2017). These findings all show that S. Newport was able to survive on cucumbers and temperature can greatly influence their survival capabilities.          	   32 Table 2.2 Δlog CFU and maximum achieved density (Nmax) of Salmonella on mini cucumbers at room temperature (22 ± 2 °C) and at refrigeration temperature (4 ± 2 °C). Serotype Δlog CFUa Nmaxb(log CFU) 22 ± 2 °C 4 ± 2 °C 22 ± 2 °C 4 ± 2 °C Newport -0.20 ± 0.01C -1.39 ± 0.01B 6.09 ± 0.49B 5.94 ± 0.11A Typhimurium -0.23 ± 0.01C -1.64 ± 0.01C 5.73 ± 0.12C 5.85 ± 0.11A Enteritidis -0.17 ± 0.01C -0.91 ± 0.01A 6.14 ± 0.16B 6.04 ± 0.09A Daytona 0.10 ± 0.01B -1.33 ± 0.01B 6.20 ± 0.07B 5.99 ± 0.08A Poona 0.84 ± 0.01A -1.82 ± 0.01C 6.72 ± 0.05A 6.02 ± 0.03A aΔlog CFU, log CFU (Last Day) – log CFU (Day 0). bNmax, maximum achieved density. Results are summarized by means ± standard deviations for three biological replicates plated in duplicate. Means within a given column with the same letter are not significantly different from each other (overall α = 0.05, Tukey’s correction).         2.3.2 Maximum achieved density (Nmax) of S. enterica on mini cucumbers In addition to possessing the highest Δlog CFU on mini cucumbers at 22 ± 2 °C, S. Poona also had the highest Nmax of 6.72 ± 0.05 log CUF at room temperature (Table 2.2), which was significantly higher compared to the other serotypes (P < 0.05) and was able to continue growing for few days. This was followed by S. Daytona, Enteritidis, Newport, demonstrating a lower Nmax but there was no significant difference between them (P > 0.05). S. Typhimurium decreased the most in cell numbers. Besides, it possessed the lowest Nmax, demonstrating a weak survival on mini cucumbers at room temperature. S. Daytona and S. Typhimurium achieved the maximum cell density at day 2, while S. 	   33 Newport and S. Enteritidis achieved the maximum cell density at day 4. Since the population of S. Poona kept growing during the whole storage, its cell number achieved to the highest at last day. At 4 ± 2 °C, the maximum achieved density (Nmax) of five S. enterica strains was equal to the cell population at day 0. The cell number at day 0 was not significantly different due to the limitation of contact time. S. Newport, S. Typhimurium and S. Daytona was not able to maintain > log 5.0 CFU per cucumber on the first 6 days, suggesting that these strains are less adapted to survival on cucumber on first few days (Figure 2.2). The ability of S. enterica to maintain high cell densities may exacerbate the risks associated with consumption of contaminated cucumbers, although high concentrations are not necessarily to cause disease when contaminated produce is consumed. 2.4 Conclusions These results demonstrate that S. enterica was able to survive on mini cucumbers for extended days at both 4 ± 2 °C and 22 ± 2 °C. Different S. enterica strains showed differential survival capability at both temperatures. At 22 ± 2 °C, S. Poona S306 showed a steady growth on cucumbers during the 8 day storage at 22 ± 2 °C while all other tested strains showed at least some level of die-off on cucumbers under the same conditions. S. Poona was identified to be the most adapted to mini cucumber environment with the highest Δlog CFU of 0.84 ± 0.01 and Nmax of 6.72 ± 0.05 (Table 2.2), followed by S. Daytona, then S. Enteritidis, S. Newport, and lastly S. Typhimurium. Strong survival ability of S. Poona on cucumbers at 22 ± 2 °C substantially increases the risk of infection. At 4 ± 2 °C, the survival curves of all S. enterica strains indicated notable decrease in cell 	   34 density which highlights a poor response to cold stress. S. Enteritidis exhibited the strongest survival capability with the smallest log CFU reduction of 0.908 (P < 0.05) compared with other strains. This was followed by S. Daytona, S. Typhimurium and S. Newport, and lastly, S. Poona. Interestingly, S. Poona, which survived the best at 22 ± 2 °C, survived the worst at 4 ± 2 °C. Furthermore, there was an evident recovery of growth from the initial sharp decrease for S. Typhimurium, S. Daytona, S. Newport (Day 12) and S. Poona (Day 15). This is most likely to be caused by adaption to cold stress and recovery of survival capabilities after incubated at refrigeration temperature for a few days. The temperature fluctuation of the incubator, which was monitored by the incubator indicator and probably caused by frequently opening and closing the fridge door, could also be the reason.  Overall, these results indicate that these five S. enterica strains can survive on mini cucumbers at both temperatures. As expected all S. enterica strains survived better at 22 ± 2 °C than at 4 ± 2 °C because room temperature is closer to the optimum survival condition of S. enterica, which is around 37 °C. S. Poona showed the largest difference in survival between the two temperatures as it was able to grow on mini cucumbers at 22 ± 2 °C and died off at the highest rate at 4 ± 2 °C. Temperature is an important environmental factor associated with survival and storing cucumbers at refrigeration temperatures can lower the risks of S. enterica infections through consumption of raw cucumbers.   	   35 2.5 Potential interventions for specific S. enterica strains These results suggest that S. enterica can survive on mini cucumbers at room temperature and refrigeration and provided insight on external factors contributing to survival (i.e., temperature, the ability to internalize or attach to the surface). Unique survival characteristics among S. enterica strains at different temperatures increase our understanding of the factors influencing microbial safety of fresh produce and reveal that corresponding interventions need to be applied to eliminate contamination of produce with specific S. enterica strains. Refrigeration is an effective and economical way to kill S. enterica on fresh produce, which is supported by the results of this study. However, due to some specific factors related to produce, there are unique challenges in produce industry for eliminating S. enterica contamination compared to other food products. For instance, produce (e.g. cucumbers) is typically physically processed and consumed without cooking. Moreover, produce is usually not packaged both at markets and home. Also, the ability of pathogens to internalize into produce exists, some of which could even last for few days. However, even with the proper plan and systems in place, it may still be possible for some microbial contamination to occur and it is at that point that corrective actions need to be taken. Effective actions include determining and fixing the point in the production chain at which contamination was introduced and determining what to do with the contaminated produce. The magnitude of contamination and the risk it presents has to be determined before considering treatments that can render the produce safe or if the produce will have to be discarded. Other studies demonstrated some specific strategies are applied for specific S. enterica strains. Sharma et al. found that phage application to whole cucumbers before slicing did 	   36 not reduce the transfer of S. Newport to fresh-cut slices. However, lytic phage application could be a potential intervention to reduce S. enterica populations on whole cucumbers (Sharma et al., 2017) as a study found a significant decline in S. Newport populations on phage-treated whole cucumbers compared to control-treated cucumbers. Treatment of ultrasound and organic acids is also an effective way to reduce S. enterica. A previous study found that a combined treatment of ultrasound and organic acids resulted in additional 0.8 to 1.0 log CFU reduction in cell numbers compared to individual treatments, without causing huge quality changes (color and texture) on lettuce during 7 day storage (Sagong et al., 2015). S. Enteritidis is the second most commonly isolated serotype after S. Typhimurium in Canada, making it an important foodborne pathogen. A variety of potentially adherent fimbrial types produced by S. Enteritidis can enhance the ability of S. Enteritidis to internalize and adhered to produce surface. At room temperature, S. Enteritidis did not survive well as its cell number decreased 0.17 ± 0.01 log CFU. However, at refrigeration temperature S. Enteritidis was able to survive well as its population only decreased 0.91 ± 0.01 log CFU. Besides, sanitizers are commonly used in produce industry to eliminate the contamination of foodborne pathogens like S. enterica on fresh produce surface. There are numerous studies on the efficacy of a variety of sanitizers in killing S. enterica on fresh produce. Linda et al. compared the efficacy of sterile USP water, Dey and Engley (D/E) neutralizer broth, and alkaline solution comprised of ingredients generally recognized as safe to remove S. enterica from tomato surface and found that the alkaline solution treatment resulted in the most effective reduction in the number of S. enterica compared to the sterile water or D/E neutralizer broth controls (Linda et al., 2001). Usage of ionizing radiation was investigated as a 	   37 means to reduce or to totally inactivate S. enterica, if present, on the produce (Kathleen et al., 2000). But its effects on the quality (e.g. shelf-life and taste) of produce need to be further explored before widely applied in food industry. Moreover, a Sparta polyester brush was identified to be less effective than a scouring pad for removing S. enterica from carrots (Erickson, et al., 2015). Peeling was found not able to eliminate the pathogens from the produce items, which may due to contamination of the utensil during use. A considerable challenge remains to ensure safe produce due to a lack of packaging and the possibility of the produce becoming recontaminated after any post-harvest treatment. Multiple combined strategies are most promising approach when applied to eliminate S. enterica from produce. As people currently have a better understanding of the molecular basis of the internalization and interactions with produce, this may provide valuable references to develop more effective intervention strategies in the future (Irene et al., 2009).  Chapter 3: Examination of biofilm formation and attachment ability of S. enterica 3.1 Introduction Biofilm is a community of microbes embedded in an organic polymer matrix, adhering to a surface (Carpentier et al., 1993). Microbial biofilms develop when microorganisms attach to living and nonliving surfaces, where EPS were generated to facilitate adhesion and provide a structural matrix. Biofilms are dynamic and responsive to the environment, 	   38 helping bacteria adapt to changes in the environment (Sutherland, Ian, 2001). Bacteria can detach from surface individually or in clumps (Stoodley et al., 2002). When they detach in clumps, they retain the reduced susceptibility to antimicrobials characteristic of biofilms. The clumps that are pieces of the biofilms do not attach to the surface but maintain the protective properties of the original biofilm and are thus much more difficult to be killed (Stoodley et al., 2002). Biofilm can show in the aspects of both solids and liquids, which is similar to slug slime (Khan and Butt, 2015). The solid-liquid interface between a surface and an aqueous medium (e.g., water, blood) provides an ideal environment for the attachment and growth of microorganisms (Donlan, Rodney, 2002). It was found that different chemical substances or physical parameters could affect the biofilm expression, such as NaCl concentration, temperature and presence or absence of oxygen (Kaiser et al., 2013). Many factors are considered when exploring the framework of attachment, e.g. the effects of the substratum, conditioning films formed on the substratum, hydrodynamics of the aqueous medium, characteristics of the medium, and various properties of the cell surface (Donlan, 2002). Bacterial attachment, colonization, and biofilm formation on fresh produce surfaces can serve as a source of cross-contamination of produce, leading to health issues and increased produce spoilage (Carpentier et al., 1993; Kumar et al., 1998). Crystal violet is a common dye used to test the formation of biofilm. Gram-negative bacteria do not retain the crystal violet dye used in the gram staining method due to the fact that they possess a lipid-rich outer membrane (as well as a plasma membrane) and a thin peptidoglycan layer (Beveridge, Terry, 1999). Gram-negative bacteria are often harmful to a host, which is the case for many of the S. enterica bacteria. The primary stain (crystal violet) is washed from the cells in the 	   39 alcohol decolorizing step of Gram staining. Biofilm formation by S. enterica has only recently been investigated (Annous et al., 2005). Therefore, the second research objective was to evaluate the biofilm formation and attachment ability of the five S. enterica strains by microplate assay and explore the relationship between survival capabilities and biofilm formation ablilities. We hypothesized that biofilm formation and attachment ability will differ across the five strains of S. enterica, and this will correlate with their capacities to survive on the surface of mini cucumbers. 3.2 Materials and methods        3.2.1 Quantification of biofilm formation of Salmonella Pure overnight culture of five S. enterica strains was diluted in 105 folds (i.e., from ~109 CFU/mL to ~104 CFU/mL) in fresh BHI broth. Specifically, 10 µL of overnight culture was added to 990 µL fresh BHI broth and the tubes were briefly vortexed. Then, 10 µL of this 100X diluted culture was added to 990 µL fresh BHI broth with a fresh tip to obtain 10,000X diluted culture. The tubes were also briefly vortex. Finally, 1 mL of this 10,000X dilution was added into 9 mL of BHI and used as the final 100,000X diluted culture used in the biofilm formation assay. Wells of a 96-well black microplate with a clear lid (sterile, nunc, ThermoFisher) were filled with 200 µL of the 100,000X diluted culture (BHI). Each strain was monitored at four time points (24, 48, 72, 96 h) with each containing 4 technical replicate wells. The edge of the plate was wrapped in Parafilm to prevent evaporation and incubated at 22 ± 2 °C for 24 hours as the same conditions used in survival assay. After incubation, the microplate was washed three times with room temperature Tris buffer (pH 7.4, 0.05M) to remove loosely adhered cells. One set of wells 	   40 (4 per each strain) was left for biofilm quantification and the remaining 3 sets were filled with 200 µL of fresh (sterile) BHI broth and the plate was again incubated at 22 ± 2 °C, for additional 24, 48 and 72h. After each time point, one set of wells were washed three times with Tris buffer (pH 7.4, 0.05 M) and left for biofilm quantification using crystal violet. Biofilms were stained by adding 200 µL of 0.1% crystal violet solution to each well and incubating for 1h. After incubation, the crystal violet was removed and wells were washed three times with Tris buffer (pH 7.4, 0.05 M) by pipetting up and down several times. After washing, the entire plate was inverted on clean paper towels and dried overnight. After drying, the attached crystal violet was solubilized with 150 µL of 100% ethanol for 30 min (the plate was gently shaken on a vortex platform for 60 s every 10 min). The ethanol solution from plate was transferred into a new 96-well clear microplate (sterile, nunc, Corning). Solubilized crystal violet in the ethanol solution was quantified by measuring absorbance at 595 nm using a Tecan GENios fluorescent-plate reader (Phoenix Research Products, Hayward, CA). The assay was repeated in three biological replicates.       3.2.2 Quantification of cell attachment of S. enterica Crystal violet detection of attached cells was examined at 0, 3, 6, 9, 12, 24, 48, 72, 96 hours at 22 ± 2 °C and 4 ± 2 °C. The temperatures were consistent with the temperatures used in survival assay. The procedure was similar to the quantification of biofilm formation described in 3.2.1. The only difference was that the cell culture was re-suspended in PBS when examining the attachment ability while in biofilm formation assay the overnight culture was diluted in BHI broth. To be specific, overnight culture 	   41 (10 mL) was centrifuged (Model 5424R, Thermo Fisher, Waltham, MA, USA) at 5000 x g for 10 min (room temperature) and the supernatant was subsequently decanted. The cell pellet was washed twice with PBS and re-suspend in 10 mL of sterile PBS. 200 µL of the re-suspended culture was transferred to designated wells of a 96-well black microplate with a clear lid. Each S. enterica strain was represented by 4 sets of wells one each of 2 plates (8 time points), each set contained 4 technical replicate wells. The edge of the plate was wrapped in Parafilm to prevent evaporation and incubated at 22 ± 2 °C and 4 ± 2 °C for 72 and 96h, respectively. At each sampling time point, the same washing procedure was performed as described in the biofilm formation assay. The sampling time points were 0, 3, 6, 9, 12, 24, 48, 72 h for 22 ± 2 °C and 3, 6, 9, 12, 24, 48, 72, 96 h for 4 ± 2 °C. The final quantification of crystal violet was performed as described in the biofilm formation assay (Section 3.2.1).       3.2.3 Data and statistical analysis The total area under the survival curve (AUC), which combines both the inoculation period and Nmax parameters to yield an overall estimate of biofilm formation was used to assess biofilm-producing abilities of the S. enterica strains. A high AUC value indicates a high ability in the biofilm formation, whereas a low AUC value indicates a lower ability. The AUC was calculated according to the trapezoidal rule, in which the area is divided into trapezoids and summed to yield the AUC (Fong et al., 2016): 𝐴𝑈𝐶 = (𝑡! − 𝑡!)𝑂𝐷!"!(𝑡!)+ 𝑂𝐷!"!(𝑡!)2  where 𝑂𝐷!"! (t) is equal to the OD595 at a given time, t. In the current study, t refers to 	   42 the storage time in days. The variables t1 and t2 correspond to the first and second time points, respectively, because each successive trapezoid is summated to give the total AUC. The one-way analysis of variance models, using each of the parameter as the response, were carried out to evaluate the contributions of the strain and food matrix. A P value of < 0.05 was considered statistically significant. Statistical analyses were performed using JMP version 11.1.1 (SAS Institute, Inc., Cary, NC).            	   43 3.3 Results and discussion        3.3.1 Quantification of Salmonella biofilm formation   Fig. 3.1 Biofilm formation by Salmonella strains represented by OD595 at 22 ± 2 °C. The data are presented as the mean of three replicates and the error bars indicate the standard errors of the means.  The biofilm formation ability of the five S. enterica strains was tested in 96-well microplates with three replicates. Strains were incubated in a black 96-well microplate and stained by crystal violet. Cells that did not attach to the surface were washed by Tris buffer at each time point. Residual crystal violet binding with cells was eluted by 100% ethanol and transferred to a clear 96-well microplate. Solubilized crystal violet in the ethanol solution was quantified by measuring absorbance at 595 nm using fluorescent-plate reader. The higher OD595 values indicate that more biofilms are produced by S. 	   44 enterica. The biofilm forming capacity was dependent on medium and temperature, but also on the origin of the strain. Plastic and polystyrene were selected as substrate materials due to their extensive use in produce processing industry (Arora et al., 1998; Chen, 2009). There were previous studies applying crystal violet directly on cucumber, mango and guava surface to determine the biofilm formation ability by S. enterica (Tang et al., 2012), but these produce were cut into uniform sizes and placed onto petri dish prior to use. In this case, the intact produce surface was destroyed and the water or nutrition were released to the surface, providing an unlimited environment for S. enterica to contaminate with their inner fruit part. Most importantly, conducting this assay on fresh cucumber could cause large variation due to the tough fruit peel and the abundant fruit juice generated by fresh cucumber. After the 24-hour incubation, the biofilm formation by S. enterica showed no significant increase with the average OD595 of ~ 0.16 ± 0.05 compared with the negative control (P > 0.05). In 24 hours, S. enterica had time to contact with the substrate but biofilm did not start forming due to the low OD595 tested at 24 h. Biofilm was formed when bacterial cells attach to one another and/or adhere to a living or inert contact surface. At 24 hours, the old BHI broth was replaced by a fresh one, after which the biofilm formation by S. enterica began to show different levels of increase. For the biofilm formation by S. Poona, the OD595 value increased significantly (P < 0.05) during this 96-hour period. From 24 h to 48 h, the OD595 of S. Poona increase slowly from 0.15 ± 0.04 to 0.33 ± 0.03 but increased sharply to 0.70 ± 0.07 at 72 h. However, during the last 24 hours, the biofilm formation of S. Poona increased slowly again, with the OD595 eventually reaching 0.82 ± 	   45 0.08. These results illustrated that the biofilm formation ability of S. Poona had a quick enhance during 48 h to 72 h at 22 ± 2 °C. In addition, too short or too long incubation time will both result in the slowing of biofilm formation. S. Daytona produced almost no biofilm at the first 48 hours, but its biofilm formation increased gradually after 48 hours, with the OD595 reaching 0.32 ± 0.06 and 0.42 ± 0.07 at 72 h and 96 h, respectively. The biofilm formation of S. Newport, S. Typhimurium and S. Enteritidis experienced a similar trend. Their OD595 values both increased steadily over the whole period. For all the tested S. enterica strains, the OD595 values that represented the quantity of biofilm formation were quite low (~ 0.17 ± 0.03) at time 24 h. This indicated that there was little biofilm formation by S. enterica at 22 ± 2 °C. The reason behind is that the cells need time to adapt to the new environment conditions when the bacterial cells are newly transferred into the substrate. Besides, the inoculated bacterial cells needed time to migrate on the surface to seek for suitable secure sites for adhesion (Pui et al., 2011a). The inoculated cucumbers stored at 4 ± 2 °C for up to 19 days showed a > 1.5 log CFU reduction in S. Poona populations (Figure 2.2). These results indicated that refrigeration can suppress the growth of this microorganism. This prompted us to investigate the effect of room temperature on biofilm formation by S. enterica on cucumbers. The quantity of biofilm formation represented by the OD595 values can be affected by many factors. For instance, variable hydrodynamic speed that normally occurs when manually washing loose cells from the surface by pipetting several times will change the flow rate leading to different levels of detachment (Jyoti et al., 2001). The variation of the incubation temperature will also affect the production of biofilms as the reproduction and metabolism of S. enterica are dependent on temperature. Besides, the drawback of the 	   46 crystal violet assay is that crystal violet does not differentiate between live and dead cells or between cells and extracellular polymers, and different strains may produce varying levels of extracellular polysaccharides, which may result in an overestimate of the biofilm formation by S. enterica. Overall, the biofilm formation on plastic surfaces by S. enterica presented by OD595 values increased with contact time, as shown in Figure 3.1. The background signal was defined as the mean OD595 of the negative control (~ 0.20 ± 0.05). S. Poona possessed greatly higher ability of biofilm formation with the OD595 increasing notably from 0.15 to 0.82 compared to the other four strains, which generally showed less biofilm formation at the tested culture conditions. S. Daytona was identified to possess the second strongest ability of biofilm formation as the OD595 increased particularly after 48 hours. There was no significant difference among other three S. enterica strains of OD595 values (P > 0.05), which showed little to no biofilm formation. These data indicate that biofilm forming behavior is associated with strain origins. As previous survival assay showed, in addition to possessing the strongest biofilm formation ability, S. Poona also survived the best at 22 ± 2 °C, followed by S. Daytona, then S. Newport, S. Typhimurium, S. Enteritidis. These results are in line with the results of survival assay on mini cucumbers. S. Poona is both capable of growing on the surface of mini cucumbers and forms the strongest biofilm during incubation at 22 ± 2 °C, which means biofilm formation could be a potent factor that contributes to the survival of S. enterica and cause severe cross-contamination. Higher growth in combination with biofilm formation represents a higher food safety risk and should be addressed in future studies on produce safety.  	   47       3.3.2 Quantification of cell attachment       3.3.2.1 Salmonella attachment ability This experiment was conducted under a simulative condition of produce surface due to the difficult operation on fresh cucumber surface. S. enterica was evaluated for attachment at 22 ± 2 °C and 4 ± 2 °C during 72 to 96 hours using crystal violet. Since their initial OD595 values are no significantly different (~ 0.24 ± 0.13, P > 0.05), AUC (OD595) values can be used to calculated to assess the attachment abilities of S. enterica at both temperatures. The AUCs were tested from 0 h at 22 ± 2 °C and from 3 h at 4 ± 2 °C. That is because the whole experiment was conducted at room temperature in the cabinet. It is impractical to obtain the AUC of attachment at 4 ± 2 °C. The data demonstrated that S. Daytona possesses the strongest attachment ability with the highest AUC (OD595) value of 22.82 ± 0.76 at 22 ± 2 °C (Table 3.1). This is followed by S. Poona (19.88 ± 1.23), and subsequently S. Typhimurium, S. Enteritidis and S. Newport (17.35 ± 0.98, 16.67 ± 1.91 and 16.58 ± 1.12, respectively). The AUCs obtained for S. Daytona and S. Poona were significantly greater (P < 0.05) compared to the AUC of S. Enteritidis, S. Newport and S. Typhimurium, indicating a stronger adherence ability at RT. Interestingly, we previously reported that Salmonella was able to survive better at 22 ± 2 °C compared to 4 ± 2 °C but not all Salmonella processed higher attachment abilities at 22 ± 2 °C as shown in Table 3.1. S. Daytona, Poona and Enteritidis demonstrated greater AUCs at 22 ± 2 °C compared to 4 ± 2 °C. The AUC obtained for S. Daytona decreased most notably when incubated at refrigeration temperature, followed by Poona and lastly, Enteritidis. Salmonella Daytona and Poona no longer attached the best as at 22 ± 2 °C. In 	   48 contrast, they had the lowest AUCs at 4 ± 2 °C. For Salmonella Newport, there was no significant difference between the AUCs obtained at 22 ± 2 °C and 4 ± 2 °C (P > 0.05), with the AUC of 16.58 ± 1.12 and 17.84 ± 0.95, respectively, which means the variation of temperature only exerted little influence on the attachment ability of Salmonella Newport. Interestingly, the AUC for S. Typhimurium increased a little when incubated at 4 ± 2 °C than at 22 ± 2 °C, indicating stronger attachment strength at lower temperature. Table 3.1 Area under the curve (AUC), optical density at 595 nm (OD595) and attachment ability of Salmonella serotypes incubated at room temperature (22 ± 2 °C) and at refrigeration temperature (4 ± 2 °C). Serotype Initial OD595a AUC (OD595)b Attachment abilityc 22 °C (0 h) 4 °C (3 h) 22 °C 4 °C 22 °C 4 °C Newport 0.21 ± 0.01A 0.21 ± 0.01A 16.58 ± 1.1C 17.84 ± 0.95A ++ ++ Typhimurium 0.23 ± 0.02A 0.22 ± 0.04A 17.35 ± 1.0BC 20.30 ± 0.84A ++ +++ Enteritidis 0.23 ± 0.01A 0.24 ± 0.01A 16.67 ± 1.91C 13.84 ± 1.01B ++ + Daytona 0.25 ± 0.01A 0.20 ± 0.01A 22.82 ± 0.76A 12.45 ± 0.98B ++++ + Poona 0.26 ± 0.01A 0.23 ± 0.01A 19.88 ± 1.23B 10.96 ± 0.89C +++ + aInitial OD595, OD values tested at 0 h (22 °C) or 3 h (4 °C). bAUC, area under the curve. cAttachment ability, strains were tentatively differentiated as strongly (++++) versus weakly (+) adherent based on the maximum achieved OD595 and AUC values. Results are summarized by means ± standard deviations for three biological replicates plated in duplicate. Means within a given column with the same letter are not significantly different from each other (P > 0.05).      	   49       3.3.2.2 Relationship between attachment ability and survival capability  The ability of S. enterica to attach to plastic was highly variable during initial stages of the incubation time as can be seen by the large fluctuation in OD595 at both 22 ± 2 ºC and 4 ± 2 ºC (Fig. 3.2a, b). Incubation time here can be known as contact time which is the time for the bacterial cells to contact with the surface.  There was a significant increase in OD595 during first 6 hours at 22 ± 2 ºC (Figure 3.2a), after which their OD595 decreased gradually and	   reached a stable level. The maximum OD595 was obtained at 6 h for S. Newport, S. Typhimurium, S. Poona and at 3 h for S. Daytona, S. Enteritidis at 22 ± 2 °C, with all OD595 decreasing significantly after achieving maximum ODs (Fig. 3.2a). These variations indicated that bacterial cells are going through cycles of attachment and detachment from the surface during the initial incubation. S. Poona and S. Daytona had higher survival capabilities compared to S. Typhimurium, S. Enteritidis and S. Newport at 22 ± 2 ºC. Similarly, S. Poona and S. Daytona have greater attachment ability at room temperature, which means that the attachment abilities of S. enterica may contribute to the survival rate when incubated at room temperature but the influence was tiny. However, this was only a preliminary conjecture because no excellent correlation and linearity could be provided.     	   50           Fig. 3.2 Mean values of attachment ability of Salmonella strains represented by OD595 at room temperature (22 ± 2 ºC) (a) and at refrigeration temperature (4 ± 2 ºC) (b). The data are presented as the mean of three replicates and the error bars indicate the standard errors of the means.  The OD595 values increased evidently during preliminary period. That may be due to the incremental adhered cells with the increasing incubation time, which is the time for the bacterial cells to contact with the surface. When the contact time increases, more bacterial cells have enough time to attach to the surface. The subsequent decreasing OD595 values may likewise be due to metabolic quenching and maximum life expectancy suspended in phosphate buffered saline (PBS) (Amresco), since the reproduction capability of Salmonella was restricted if provided with no external nutrition and the dead cells would be removed by washing immediately after the incubation period.  (a) 22 ± 2 ºC (b) 4 ± 2 ºC 	   51 However, at refrigeration temperature (4 ± 2 °C), the attachment abilities of S. enterica were fluctuating during the entire incubation period, with various increase and decrease. S. Daytona, S. Poona and S. Enteritidis demonstrated higher attachment abilities at 22 ± 2 °C than at refrigeration temperature, especially at preliminary incubation period. This may be caused by the less physical strength and endurance when stressed with cold. When more attached bacterial cells are found on the fresh produce surface, this indicates that more interaction forces can be formed between the biofilm and the fresh produce surface. The properties of the attachment surface contribute a lot to the potential of biofilm formation (Tang et al., 2012). They include the surface roughness, wetability (determined by hydrophobicity), cleanability, disinfectability and vulnerability to wear influence the ability of bacterial cells to adhere to a particular surface (Van and Michiels, 2010; Sarjit and Dykes, 2017). In this study, the cells were incubated in a sterile 96-well flat-bottomed polystyrene microplate, which makes it easier to control the consistency of the attachment surface properties. Overall, S. enterica strains used in this study exhibited different capabilities of attachment at both temperatures. The attachment ability may contribute weakly to the survival capability, especially at room temperature. However, the direct correlation between the attachment ability and survival rate was not proven. Furthermore, since crystal violet does not differentiate between live and dead cells or between cells and extracellular polymers, and different strains may produce varying levels of extracellular polysaccharides, the strength of adherence should be further confirmed by more quantitative means in the future. 	   52  3.4 Conclusions Evaluation of biofilm formation by S. enterica in this study revealed that S. Poona possessed a high potential for biofilm formation on plastic surfaces while S. Newport, S. Typhimurium, S. Enteritidis and S. Daytona possessed weaker capacities to produce biofilm at room temperature. These results confirm some previous findings, which showed that S. enterica are able to form biofilm on plastic surfaces (Römling and Rohde, 1999; Joseph et al. 2001; Mireles, Toguchi and Harshey, 2001; Djordjevic et al. 2002; Stepanovic et al. 2003). Generally, glass and stainless steel are assumed to be hydrophilic materials while rubber and plastic are hydrophobic materials (Sinde and Carballo 2000; Donlan 2002). Previous studies showed that S. enterica strains adhere in higher numbers to more hydrophobic materials (Cunliffe et al. 1999; Sinde and Carballo 2000; Donlan 2002). Therefore, this could be one possible explanation for the ability of these bacteria to produce biofilm in high numbers on plastic surface. On the other hand, it is well known that many factors influence attachment ability of S. enterica, like temperature. Therefore, two temperatures were selected in this study. Room temperature is better condition for the survival of S. enterica compared to refrigeration temperature. This study demonstrated that S. enterica, as foodborne pathogens, readily form biofilm on plastic surfaces, which are nowadays frequently used in food-processing environments. Biofilm formation and attachment ability by these bacteria are affected by temperature and S. enterica serotypes. In general, biofilm formation behavior is associated with S. enterica serotypes. S. Poona was able to grow on the surface of mini cucumbers and possessed the strongest biofilm formation ability during incubation at 22 ± 2 °C, compared to the other 	   53 four strains. S. Daytona was identified to possess the second strongest ability of biofilm formation. There was no significant difference among other three S. enterica strains in OD595 values (P > 0.05), which showed little to no biofilm formation. Besides, S. enterica exhibited different capabilities of attachment and the adhesive ability may contribute weakly to the survival capability, especially at room temperature. However, the direct relationship between the attachment ability and survival rate was not significantly correlated as the results showed.  Chapter 4: Characterization of the S. enterica rdar morphotype 4.1 Introduction Curli fimbriae is an important component of the biofilm extracellular matrix. In S. enterica, biofilm formation is associated with the production of curli and exopolysaccharide, such as cellulose and colanic acid (Austin et al., 1998; Solano et al., 2002; Danese et al., 2000; Vidal et al., 1998). Certain S. enterica are capable of producing thin aggregative fimbriae that bind the dye Congo red and mediate attachment to both inert surfaces and biological proteins. In S. Typhimurium and S. Enteritidis, the co-expression of thin aggregative fimbriae and cellulose leads to an aggregative colony morphotype (red, dry, and rough [rdar]) when grown on medium containing the dye Congo red and Coomassie brilliant blue G-250 (Solano et al., 2002). The rdar morphotype was a multicellular behaviour of S. enterica and characterized by the expression of the adhesive extracellular matrix components cellulose and curli fimbriae (Römling, 2005; White and Surette, 2006).  	   54 The rdar (red, dry and rough) morphotype, a multicellular behavior of S. enterica was characterized by the expression of the adhesive extracellular matrix components cellulose and curli fimbriae, which are predominant matrix-compounds in S. enterica (Jonas et al., 2007) and play a synergistic role in biofilm formation (Pamp et al., 2007; Steenackers et al., 2012). The disruption of both or either of these components will lead to distinct changes in colony morphology on Congo red agar plates (Friedman et al., 2004; Jonas et al., 2007). The addition of the planar hydrophobic diazo dye Congo red to the agar medium can enhance visualization of the colony morphology (Etienne et al., 2002; Sondén et al., 2005).  The extracellular matrix components are able to interact with Congo red and Coomassie brilliant blue G-250, resulting in typical color of the colony. Therefore, the last research objective was to examine the S. enterica cell morphology characterized by the expression of the adhesive extracellular matrix components cellulose and curli fimbriae on Congo red agar (CRA) plates and evaluate how these two components contribute to the biofilm formation in S. enterica. We hypothesized that S. enterica cell morphologies will differ across strains with different biofilm formation and attachment abilities. 4.2 Materials and methods       4.2.1 Bacterial strains In addition to the five S. enterica strains used in Objectives I & II, reference strains including a positive biofilm producer Pseudomonas aeruginosa PA14 (Friedman et al., 2004) and a negative biofilm producer Listeria monocytogenes HPB642 (Uhlich et al., 2006) were used as control. All isolates were obtained and maintained in Brain-Heart- 	   55 Infusion (BHI) broth with 15% glycerol and stored at -80 °C.       4.2.2 Phenotypic evaluation of rdar morphotype  To detect curli fimbriae and cellulose production, the colony morphology phenotype was assessed on Congo red agar plates using the spot test and streak test. Strains were precultured on BHI agar plates at 37 °C for approximately 18 h. Cells were suspended in 10 mL of BHI broth to obtain pure culture. Pure overnight culture was diluted for 100 folds (i.e., from ~109 CFU/mL to ~107 CFU/mL) in fresh BHI broth. Specifically, 10 µL of overnight culture was added to 990 µL fresh BHI broth and the tubes were briefly vortexed. S. enterica strains were cultivated on LB without salt agar plates supplemented with Congo red (40 µg/mL) (Sigma-Amresco) and Coomassie brilliant blue G-250 (20 µg/mL) (Amresco) (Monteiro et al., 2011). Congo red stain and Coomassie brilliant blue were prepared as a concentrated aqueous solution and added to LB agar after each was autoclaved (121 °C for 15 minutes). The strains were streaked onto Congo red plates and incubated at 22 ± 2 °C under aerobic conditions for 48 hours before determining morphotypes by comparing them to control strains. Pseudomonas aeruginosa PA14 was used as a positive control and Listeria monocytogenes HPB642 was used as negative control for polysaccharide formation. The Congo red dye directly interacts with certain polysaccharides, forming colored complexes. Isolates were grouped into five distinct morphotypes (Solomon et al., 2005): (i) red, dry, and rough (rdar), indicating curli and cellulose production; (ii) brown, dry, and rough (bdar), indicating curli production but a lack of cellulose synthesis; (iii) smooth and red (sar), indicating cellulose production but a lack of curli synthesis (Castelijn et al., 2012); 	   56 and (iv) smooth and white (saw), indicating a lack of both curli and cellulose production. Besides the inoculation of strains in streaks on the modified CRA plates, spot inoculation was also evaluated. A 2 µL-aliquot of 102-fold diluted overnight culture with ~107 CFU/mL was inoculated in a spot and incubated at 22 ± 2 °C under aerobic condition for 48 hours. Seven strains were inoculated per plate and the experiment was performed in triplicates. For colony size studies, three independent cultures of each strain were used to generate 21 replicate spots/plate, which was monitored daily for 48 h. The experiments were performed at least twice to ensure the consistency of the results. 4.3 Results and discussion Curli fimbriae and cellulose production was analyzed on agar plates containing Congo red and Coomassie brilliant blue G-250. Morphotypes were judged by comparing test strains to control strains as follows. S. Poona expressed the rdar morphotype (Table 4.1 & Fig. 4.1), which correlates with the production of both curli fimbriae and cellulose (Solomon et al., 2005). S. Daytona showed a bdar morphotype, which indicates the production of curli but the deficiency in cellulose biosynthesis (Table 4.1 & Fig. 4.1). For S. Newport, S. Typhimurium and S. Enteritidis, a sar morphotype was observed, which indicates the production of cellulose but the deficiency in curli biosynthesis (Table 4.1 & Fig. 4.1). The positive control Pseudomonas aeruginosa PA14 also expressed a red, dry, and rough morphotype, while a saw morphotype was observed in the negative control Listeria monocytogenes HPB642, which means no biosynthesis of both curli fimbriae and cellulose (Table 4.1 & Fig. 4.1). 	   57  Table 4.1 Morphotypes of S. enterica and the curli fimbriae and cellulose production on Congo red plates. Serotype Morphotype Control Morphotype Newport sar Positive control: Pseudomonas  aeruginosa PA14 rdar Typhimurium sar Enteritidis sar Negative control:  Daytona bdar Listeria monocytogenes  HPB642  saw Poona rdar Note: Morphotypes expressed on Congo red agar plates: rdar (red, dry and rough), bdar (brown, dry and rough), saw (smooth and white), and sar (smooth and red). The morphotypes observed are comparable to those found in other studies (Barnhart and Chapman 2006; Romling et al. 1998; Jones et al. 2007).  In this study, when S. enterica were grown on Congo red agar at RT for up to 48 h, four distinct phenotypic classes were observed (Table 4.1): S. Poona and the positive control Pseudomonas aeruginosa PA14 formed rdar colonies with complete surface patterns that could be lifted off the agar surface intact, S. Daytona formed bdar colonies with complete patterns, S. Newport, S. Typhimurium and S. Enteritidis formed smooth, non-aggregative red colonies without surface patterns, and the negative control Listeria monocytogenes HPB642 formed smooth and white colonies without surface patterns. S. Poona appeared the same multicellular and aggregative behavior [rdar (red, dry and rough) morphotype] as the positive control Pseudomonas aeruginosa PA14, suggesting that S. Poona produced both curli fimbriae and cellulose. Previous results showed that S. Poona produced significant amounts of biofilm when cultivated in the appropriate medium at room temperature. In addition, S. Poona possessed the best survival capacity at room 	   58 temperature compared to the other S. enterica isolates. These results indicate that cellulose and curli may play a role in the survival and resistance of S. enterica in the food environment. Interestingly, the biofilm formation ability of S. Poona was significantly higher than S. Newport, S. Typhimurium and S. Enteritidis, which produced the least amount of biofilm and expressed a smooth and red (sar) morphotype without surface patterns on Congo red agar indicating the production of cellulose but the deficiency in curli biosynthesis. These results gives rise to the hypothesis that although curli fimbriae and cellulose work synergetically to promote biofilm formation in S. enterica, the effects of cellulose on biofilm capacity are less important compared to curli fimbriae (Saldaña et al., 2009). Besides, bdar (brown, dry and rough) colonies with surface patterns were only observed in S. Daytona. The bdar morphotype was found to be associated with the production of curli fimbriae but the deficiency in cellulose biosynthesis. In previous assays, S. Daytona ranked the second in biofilm formation abilities as well as survival capacities, which confirmed the hypothesis that curli fimbriae and cellulose play a synergetic role in biofilm formation by S. enterica.       	   59                                                      Fig. 4.1 Rdar morphotypes of S. enterica on Congo red plates. (+) Pseudomonas aeruginosa PA14, positive control; (-) Listeria monocytogenes HPB642, negative control.  4.4  Conclusions Pseudomonas aeruginosa PA14 was proved to be a strong biofilm producer in previous study (Friedman et al., 2004) and showed a rdar morphotype on Congo red plates. In contrast, the negative control Listeria monocytogenes HPB642 that produced no biofilms (Uhlich et al., 2006) had no biosynthesis of both curli fimbriae and cellulose. Based on the results from biofilm assay, S. Poona was identified to possess the strongest biofilm capacity. The rdar morphotype illustrated that S. Poona can produce both the curli fimbriae and cellulose, which are considered as two predominant compounds contributing to biofilm development. There was production of curli but deficiency in cellulose in S. Daytona with the morphotype of bdar on Congo red plates, which possessed the second highest biofilm capacity in the biofilm assay. For S. Newport, S. Typhimurium and S. Enteritidis, they all expressed a smooth and red (sar) morphotype, S. Newport (sar) S. Typhimurium (sar) S. Enteritidis (sar)S. Daytona (bdar)  Pseudomonas aeruginosa PA14 (rdar)  S. Poona (rdar) Listeria monocytogenes HPB642 (saw) 	   60 which means only cellulose biosynthesis occurred during the biofilm development. However, the biofilm formation abilities of S. Newport, S. Typhimurium and S. Enteritidis were negligibly weak, which may due to the lack of curli fimbriae that are primary components beneficial to biofilm formation in S. enterica. According to these data, it is reasonable to propose that the strong biofilm formation ability of S. enterica was the result of both curli and cellulose, which act synergistically to promote early cell adherence. However, although both curli fimbriae and cellulose benefit the biofilm development in S. enterica, the influence of curli fimbriae is of greater importance compared to cellulose. Early hypotheses that fimbriae were involved in adherence to host cells suggested that numerous fimbrial types would contribute to the host specificities and tissue tropisms of different S. enterica (Patti et al., 1994; Forest et al., 2007). However, despite much research, the role of fimbriae in the pathogenesis of S. enterica is still not well understood (White and Surette, 2006; Wagner and Hensel, 2011). In addition, no clear links have been made connecting one fimbrial type to a particular animal host or disease process (Parsek and Singh, 2003). Therefore, the ability of more strains of S. enterica originating from produce, meat, or clinical sources to form biofilms as well as their phenotypic styles should be further investigated based on the recent increase in produce-related outbreaks of salmonellosis, along with the indication that S. enterica are able to form biofilms on both biotic (host) and abiotic surfaces.   	   61 Chapter 5: Conclusion and future direction  5.1 Conclusion The present study characterized the survival of S. Newport, S. Enteritidis, S. Daytona, S. Typhimurium and S. Poona on fresh cucumbers. In objective I, all five strains were assessed for survival on cucumber surfaces at different temperatures. All strains were capable of strong survival at room temperature but exhibited weak survival at refrigeration temperature. S. Poona was identified as being the most adapted to mini cucumber environment, followed by S. Daytona, then S. Enteritidis, S. Newport, and lastly S. Typhimurium at room temperature. However, at 4 ± 2 °C, all S. enterica strains exhibited weak resistance to cold stress based on the notable decrease in cell density. These results indicate that S. enterica can survive on mini cucumbers but their survival behaviours were affected by temperature and varied among different S. enterica serotypes. In objective II, the biofilm formation ability and attachment ability of five S. enterica strains were assessed by crystal violate method. Evaluation of biofilm formation by S. enterica in this study revealed that S. Poona possessed a high potential for biofilm formation on plastic surfaces while S. Newport, S. Typhimurium, S. Enteritidis and S. Daytona possessed weaker capacities to produce biofilm at room temperature. We also found that biofilm formation and attachment ability by these bacteria were affected by temperature and S. enterica serotypes. Lastly, in objective III, the rdar morphotype was characterized by the expression of the adhesive extracellular matrix components cellulose and curli fimbriae. The role of curli and cellulose were evaluated on Congo red agar. Different strains exhibited different 	   62 morphotypes and curli and cellulose act synergistically to promote biofilm formation. In conclusions, the clod stress response of five S. enterica serotypes on fresh cucumber surfaces was examined in this work. Survival behaviours varied with temperature and serotypes. The findings emphasize the adaptable nature of this pathogen and highlight the importance in S. enterica control on fresh produce. 5.2 Future direction In this study, the molecular mechanisms behind the effects of different temperatures on S. enterica survival behaviours were not identified. The gene expression should be examined in future studies to gain better insight into this poorly-understood phenomenon. Doing so would undoubtedly promote the development of innovative strategies to mitigate the implied risk in the produce industry. Further, fresh cucumbers were the only produce used in this study. However, there were many salmonellosis outbreaks associated with other produce items in recent decades. A recommendation for future studies would be to explore the survival of S. enterica on different produce surfaces (e.g., seed sprouts, peppers).  Besides, it is meaningful to study the survival of S. enterica at transit temperature that is around 10 °C to 12 °C. In reality, the temperature during the transportation of fresh produce from farms to markets differs with the storage temperature used in this study and the transportation could be a potential risk for cross-contamination. Therefore, it is necessary to evaluate the survival of S. enterica at transit temperature in the future. Also, although using crystal violet to examine the attachment ability of S. enterica is 	   63 common and straightforward, there is limitation of this method. Crystal violet does not differentiate between live and dead cells or between cells and extracellular polymers, and different strains may produce varying levels of extracellular polysaccharides. Therefore, the strength of adherence should be further confirmed by more quantitative means (e.g., fluorescence method).  Lastly, five strians of S. enterica were used in this study, and their relative abilities to survive at different temperatures were compared. 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