IMPACT OF BIOACTIVE MOLECULES SECRETED BY LACTOBACILLUS ACIDOPHILUS LA-5 ON CLOSTRIDIUM DIFFICILE VIRULENCE FACTORS
by Afsaneh Najarian
A Thesis Presented to The University of Guelph
In partial fulfilment of requirements for the degree of Doctor of Philosophy in Food Science
Guelph, Ontario, Canada ©Afsaneh Najarian, November 2017 ABSTRACT
IMPACT OF BIOACTIVE MOLECULES SECRETED BY LACTOBACILLUS ACIDOPHILUS LA-5 ON CLOSTRIDIUM DIFFICILE VIRULENCE FACTORS
Afsaneh Najarian Advisors: University of Guelph, 2017 Dr. Mansel W. Griffiths Dr. Shayan Sharif
Clostridium difficile is a leading pathogen of hospital-associated diseases including infectious diarrhea and pseudomembranous colitis. Increasing frequency and severity of diseases besides inconsistent results of antibiotic therapy, demand alternative therapeutic candidates using a different strategy to control pathogenicity of C. difficile. Unlike antibiotics, antivirulence compounds control bacterial infection without killing them or developing antibiotic-resistant. Reducing essential virulence factors in C. difficile including toxin production, adherence, and biofilm formation could substantially minimize its pathogenicity and lead to a faster recovery from the disease.
This study aimed to investigate antivirulence effects of La-5 bioactive molecules produced by Lactobacillus acidophilus La-5 namely proteobiotics on the pathogenicity of C. difficile. Three clinically important strains of C. difficile (ribotypes 027, 078, and 001) were incubated with or without reconstituted La-5 cell-free supernatant (CFS). The C. difficile culture filtrates were collected for the assessment of quorum sensing, and expression of several virulence genes performing reverse transcription q-PCR. C. difficile attachment and cytotoxicity in human epithelial cells were monitored in vitro using human epithelial cells HT-29 and
Caco-2 monolayers. Additionally, L. acidophilus La-5 was incubated in a medium containing whey protein, and inhibitory effects of its CFS were tested performing quantitative biofilm assay then visualized in scanning electron microscopy (SEM).
Bioactive molecules caused a significant down-regulation in expression of some of the virulence genes mainly two essential toxin genes tcdA and tcdB and adhesion gene Cwp84 in all C. difficile strains. A substantial down-regulation in luxS expression was observed in ribotype 027 and 078, and 001 (P<0.05).
La-5 bioactive molecules also reduced cytotoxicity and cytopathic effect of C. difficile culture filtrate on
HT-29 and Caco-2 monolayers significantly (P < 0.0.5). Furthermore, pre-incubation of cell monolayers and La-5 CFS reduced attachment of the C. difficile bacterial cells on both cell monolayers. The biofilm formation and population of C. difficile cells encased in biofilms were significantly decreased within the treated cultures (P < 0.0001). SEM analysis also confirmed quantitative results showing a sufficient reduction in biofilm formation. Our study suggests that La-5 bioactive molecules provide a supportive adjunct therapy for C. difficile infection, prevent relapses, and improve disease outcome.
ACKNOWLEDGMENTS
First and foremost, I would like to thank Professor Mansel Griffiths sincerely. Thank you for providing me the opportunity to conduct this research under your supervision. Your expert, knowledge, and endless support encouraged me throughout my Ph.D. program. You let me challenge myself and taught me to think critically inside and outside the box.
My great appreciation goes to Dr. Shayan Sharif. Without your encouragement, support and guidance this thesis would not have been completed. I am extremely grateful to you for standing by me through the ups and downs I faced during my study. I am thankful to you for being supportive and for having faith in me.
Exceptional thanks to Dr. Milena Corredig. Thank you for your help and support and for being one of the great members of my advisory committee. Your expert advice and your friendship throughout my research, and trusting in my ability gave me the strength to move forward with my program.
I would like to express my sincere thanks to Dr. Mitra Amiri for being my great mentor. Your advice, vast knowledge, and friendship was great inspiration for me. Especial thanks to Dr. Alexandra (Sandy) Smith for microscopy training and assisting me with the sample preparation for scanning electron microscopy.
A great thank you to Dr. Scott Weese for his guidance and expert advice, and to Joyce Rousseau in Ontario Veterinary College for providing me the C. difficile strains. I would like to thank Dr. Rocio Morales for her supervision and guidance. Also, I acknowledge all the past and present students and researchers of Canadian Research Institute for Food Safety (CRIFS) for sharing ideas, excellent cooperation and making our laboratory a lively and friendly workplace.
I appreciate staff and students at “Dairy Lab” in Food Science Department for welcoming attitude and for supporting me during my work there and for their excellent cooperation. I am deeply thankful to the faculty and staff in the Food Science Department for their excellent support and to the University of Guelph and financial sources that supported this project to be completed.
I would like to express my most profound gratitude to my husband Mohsen and my two sons Mohammad and Matin for their unconditional love, understanding, help, and sacrifices they made for me throughout my academic journey. My sincere thanks go to my late father, and my two late brothers for inspiring me to start this path, and my huge thanks go to my beloved mother and my dear sister and brothers for sharing my problems and supporting me during our losses. Last but not least a big thanks to all of my friends for cheering me up and supporting me by having me in their thoughts and prayers.
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TABLE OF CONTENTS
ABSTRACT ...... II
ACKNOWLEDGMENTS ...... IV
TABLE OF CONTENTS ...... V
LIST OF ABREVIATIONS ...... X
LIST OF FIGURES ...... XIII
LIST OF TABLES ...... XIV
CHAPTER 1 ...... 1
GENERAL INTRODUCTION AND THESIS OBJECTIVES ...... 1
CHAPTER 2 ...... 6
LITERATURE REVIEW ...... 6
2. CLOSTRIDIA ...... 6
2.1 CLOSTRIDIUM DIFFICILE...... 7
2.1.1 Clostridium difficile Microbiology ...... 7
2.1.2 C. difficile Infection (CDI) and C. difficile associated diseases (CDAD) ...... 8
2.1.3 Epidemiology and Variation in Virulence Factors of C. difficile Strains: Hypervirulent
Strains ...... 10
2.1.4 Disease Transmission ...... 12
2.1.5 History of Clostridium difficile infectious disease ...... 13
2.2 CLOSTRIDIUM DIFFICILE VIRULENCE FACTORS ...... 14
2.2.1 Non-toxigenic Virulence Factors: Colonization and Adherence...... 14
2.2.2 Toxigenic Virulence Factors and Mechanism of Action ...... 16
2.2.3 Binary Toxins ...... 22
2.3 C. DIFFICILE OTHER VIRULENCE FACTORS ...... 23
2.3.1 C. difficile Sporulation and Germination ...... 23
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2.3.2 C. difficile Quorum Sensing...... 25
2.3.3 C. difficile Motility ...... 28
2.3.4 C. difficile Biofilm Formation ...... 29
2.4 CDI MANAGEMENT, TREATMENT AND CONTROL ...... 32
2.5 PROBIOTICS AND INHIBITION OF CDI ...... 34
2.5.1 Human Microbiota ...... 34
2.5.2 Lactic Acid Bacteria and Probiotic Strains ...... 35
2.5.3 Characteristics of probiotic Lactobacilli ...... 38
2.5.3.1 Lactobacillus Adhesion ...... 41
2.5.3.2 Tight junctions ...... 41
2.5.3.3 Cell Surface Factors ...... 42
2.5.3.4 Probiotic Proteolytic System ...... 44
2.5.4 Probiotic Safety and FDA Regulations ...... 44
2.5.5 Bioactive Peptides Mechanisms of Action ...... 45
2.5.6 Secreted Proteins and Bioactive Peptides Sources ...... 48
2.5.6.1 Secreted Proteins ...... 49
2.5.6.2 Soluble Peptides ...... 49
CHAPTER 3 ...... 50
3. BIOACTIVE MOLECULES PRODUCED BY PROBIOTIC LACTOBACILLUS ACIDOPHILUS LA-5 SHOWING INHIBITORY EFFECTS TOWARD CLOSTRIDIUM DIFFICILE VIRULENCE GENES ...... 50
ABSTRACT ...... 50
INTRODUCTION ...... 51
3.1. MATERIALS AND METHODS: ...... 54
3.1.1 Modified de Man, Rogosa, and Sharpe (m-MRS) medium ...... 54
3.1.2 Chemically Modified Medium (Whey Protein medium) ...... 55
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3.1.3 Probiotic LAB Strains ...... 55
3.1.4 Production of LAB Cell Free Supernatant ...... 57
3.2. BACTERIAL STRAINS PRE-SCANNING EXPERIMENTS ...... 57
3.2.1 Bacterial Cultures and Screening for Growth...... 58
3.3 INHIBITION OF C. DIFFICILE BIOLOGICAL ACTIVITY BY BIOACTIVE EXTRACT LA-5 CFS ...... 58
3.3.1 Bacterial Strains and Culture Conditions: ...... 58
3.3.2 Impact of La- 5 Bioactive Extract on Viability and Proliferation of C. difficile...... 60
3.3.3 LAB Bioactive Extract and Inhibition of C. difficile Autoinducer-2 ...... 60
3.3.4 Real-Time Quantitative PCR and Gene Expression Analysis ...... 61
3.3.4.1 Real-Time q- PCR Methodology ...... 61
3.3.4.2 Standard curves ...... 62
3.3.4.3 RNA extraction and Real Time q-PCR ...... 62
3.4 STATISTICAL ANALYSIS ...... 67
3.5 RESULTS ...... 67
Preliminary Results ...... 67
3.5.1 Growth Kinetics of Lactic Acid Bacteria in Different Media ...... 67
3.5.2. Effect of Bioactive Extract on C. difficile Growth Rate ...... 71
3.5.3 Quorum Sensing ...... 73
3.5.3.1 Lactic Acid Bacteria CFS Influence on C. difficile Quorum Sensing ...... 73
3.5.3.2 L. acidophilus La-5 CFS Influence on Quorum Sensing of Three Strains of C. difficile .... 73
3.5.4 Gene Expression ...... 75
3.5.4.1 Experimental Design and real-time q-PCR Data Analysis...... 75
3.5.4.2 Real-Time q-PCR and Effect of La-5 CFS on C. difficile Gene Expression ...... 77
3.6 DISCUSSION ...... 81
3.7 CONCLUSION ...... 86
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CHAPTER 4 ...... 87
4. THE IMPACT OF LA-5 PROTEOBIOTCS ON ADHERENCE AND CYTOTOXICITY OF HIGHLY INFECTIOUS C. DIFFICILE RIBOTYPE 027 ...... 87
ABSTRACT ...... 87
INTRODUCTION ...... 88
4.1 MATERIALS AND METHODS ...... 91
4.1.1 Bacterial strains and bioactive peptides preparation ...... 91
4.1.2 Cell Cultures ...... 91
4.1.2.1 Selected Cell lines ...... 91
4.1.2.2 Maintenance of Cell Culture ...... 92
4.1.2.3 SRB Assay ...... 94
4.1.3 C. difficile Cytotoxicity Assay ...... 95
4.1.3.1 Non-Toxic Concentration Assay of La-5 CFS ...... 95
4.1.3.2 Cytotoxicity of C. difficile Culture Filtrate ...... 96
4.1.3.3 La-5 CFS Influence on Cytotoxicity of C. difficile 027...... 96
4.1.4 Visualization of C. difficile Cytopathic Effect on Cell Monolayer ...... 97
4.1.5 Adhesion assay ...... 98
Fixing and Gram Staining ...... 99
4.1.6 Statistical Analysis ...... 100
4.2 RESULTS ...... 100
4.2.1 Inhibition of Cytotoxicity in C. difficile Ribotype 027 by L. acidophilus La-5 CFS ...... 100
4.2.2 Inhibition of Cytopathic effect of C. difficile Ribotype 027 by L.acidophilus La-5 CFS ..... 104
4.2.3 Inhibition of C. difficile cytotoxicity on Cell Monolayer intercellular Structure by ...... 107
L. acidophilus La-5 CFS ...... 107
4.2.4 Impact of La5 CFS on C. difficile Adhesion ...... 108
4.2.5 Visualized C. difficile Adhesion to the Cell Culture Monolayers ...... 109
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4.3. DISCUSSION ...... 111
4.4 CONCLUSION ...... 115
CHAPTER 5 ...... 116
5. IMPACT OF LA-5 PROTEOBITICS IN WHEY PROTEIN MEDIUM ON BIOFILM GROWTH AND SPORULATION OF CLOSTRIDIUM DIFFICILE ...... 116
ABSTRACT ...... 116
INTRODUCTION ...... 117
5.1 MATERIAL AND METHODS ...... 119
5. 1.1 C. difficile Biofilms ...... 119
5.1.1.1. Biofilm Formation of C. difficile with Proteobiotics ...... 119
5.1.1.2. Analysis of Biofilms and Scanning Electron Microscopy (SEM) ...... 120
5.1.2. C. difficile Motility and Effect of Proteobiotics ...... 122
5.1.3. Sporulation of C. difficile and Effect of Proteobiotics ...... 122
5.1.3.1. Preparation of C. difficile sporulation cultures in liquid media ...... 122
5.1.3.2. Heat treatment of sporulation cultures ...... 123
5.1.5. Statistical analysis ...... 123
5.2 RESULTS ...... 123
5.2.1 Biofilms formation and Effect of La-5 Proteobiotics ...... 123
5.2.2 Proteobiotics Impact on C. difficile Motility ...... 128
5.2.3 Effect of Proteobiotics on Sporulation ...... 129
5.3 DISCUSSION ...... 131
CHAPTER 6 ...... 137
6. GENERAL DISCUSSION ...... 137
CHAPTER 7 ...... 143
FUTURE DIRECTION ...... 143
REFERENCES ...... 146
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LIST OF ABREVIATIONS
× g Times gravity
AKTA AKTA protein purification system a-La a-Lactalbumin
AMP Adenosine monophosphate
ANOVA Analysis of variance
BHI Brain Heart Infusion b-Lg b-lactoglobulin
CDAD Clostridium difficile associated disease
CDC Center for disease control
CDI Clostridium difficile infection cDNA Complementary DNA
CF Culture Filtrate
CFS Cell free supernatant
CFU Colony forming unit
DNA Deoxyribonucleic acid
EDTA Ethylenediaminetetracetic acid
EHEC Enterohemorrhagic
FAO Food and Agricultural Organization
FDA Food and Drug Administration
FPLC Fast performance liquid chromatography
GMP Guanosin monophosphate
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GTP Guanosine triphosphate h Hour(s)
HCL Hydrochloric acid
HpLC High performance liquid chromatography
IgA Immunoglobulin A kbp Kilo base pair
KDa Kilo dalton
La-5 Lactobacillus acidophilus La-5
LAB Lactic acid bacteria
LCT Large clostridial toxin
LH-2 Lactobacillus helveticus LH-2
M Molar mAU Milliabsorbance unit min Minute(s) mM Millimolar
MPa Megapascal
MRS De Man, Rogosa and Sharpe
MRSA Methicillin resistant Staphylococcus aureus
NaOH Sodium hydroxide
NAP1 North America pulsed field electrophoresis oC Degree celsius
OD Optical density
PaLoc Pathogenicity locus
PBS Phosphate-buffered saline
PCR Polymerase chain reaction
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PMC Pseudomembranous colitis q-PCR Quantitative - polymerase chain reaction
RLU Relative light unit
RNA Ribonucleic acid
Rpm Round per minute
RT-PCR Reversed transcription- polymerase chain reaction
S Second(s)
SDS Sodium dodecyl sulfate
SDS-PAGE Sodium dodecyl sulfate - polyacrylamide gel electrophoresis
SEM Scanning electron microscopy
SRB Sulforodhamine B
Tris Tris (hydroxymethyl) aminomethane
U Unit v/v Volume per volume
WHO World Health Organization
WP Whey protein
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LIST OF FIGURES
Figure 2.1. Schematic of human C. difficile infection………………………………………………….10
Figure 2.2. Genetic organization of C. difficile pathogenicity locus (PaLoc) ...... 18
Figure 2.3. Toxin regulation model ……………………………………………………...……………. 21
Figure 2.4. Domain organization of C. difficile toxins……………………………...………………….22
Figure 2.5. Mechanistic view of probiotic actions by lactobacilli, illustrates adaptation factors and probiotic factors of probiotic lactobacilli ...... 40
Figure. 2.6. Cell surface composition of lactobacilli..………………………………………………….43
Figure 3.1. Optical density growth curves of selected probiotics ...... 70
Figure 3.2. Regression analysis of exponential phase of Lactobacillus acidophilus La-5 ...... 71
Figure 3.3. Growth kinetics of C. difficile strains 027, 078 and 001...... 73
Figure 3.4. Linear regression of two set of data from growth kinetic of ribotype 027 ……………...73
Figure 3.5. Production of AI-2 by C. difficile strain 027 in the presence of cell-free supernantants of lactic acid bacteria...... 74
Figure 3.6. Production of AI-2 signaling molecules by three strains of C. difficile treated with L. acidophilus La-5 CFS...... 75
Figure 3.7. Regulation of pathogenicity locus (Paloc) genes by La-5 bioactive extract influence. .... 79
Figure 3.8. Regulation of non-toxigenic virulence genes by La-5 bioactive extract influence ...... 80
Figure 4.1. Analysis of SRB assay measuring non-toxic concentration of L. acidophilus La-5 treatment on HT-29 cells...... 102
Figure 4.2. Analysis of an SRB assay measuring toxic concentration of C. difficile culture filtrate on
HT-29 cells ...... 103
Figure 4.3. C. difficile cytotoxicity changes in the presence of La-5 CFS...... 104
Figure 4.4. Morphological changes induced by C. difficile culture filtrate samples ...... 106
Figure 4.5. Confocal Image of HT-29 cell integrity pretreated with La-5 CFS...... 107
Figure 4.6. C. difficile strain 027 adherence to Caco-2 and HT-29 monolayers ...... 109
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Figure 4.7. Microscopic visualization of C. difficile adherence to HT-29 cells ...... 110
Figure 5.1. Impact of La-5 bioactive molecules on biofilm formation of three C. difficile ribotypes
027, 078, and 001 ...... 125
Figure 5.2. Scanning electron microscopy analysis of La-5 Bioactive molecules’ effect on biofilm formation by three C. difficile strains...... 126
Figure 5.3. La-5 Comparison of C. difficile cluster between La-5 treated and untreated samples . 127
Figure 5.4. Effect of La-5 bioactive molecules on motility of C. difficile strains...... 129
Figure 5.5. Impact of La-5 bioactive molecules on spore production of three C. difficile ribotypes
027, 078, and 001...... 131
LIST OF TABLES
Table 2.1. Isolation rates of toxigenic C. difficile from stool of various subject populations ...... 10
Table 3.1 Lactic acid bacteria used for this study ...... 56
Table 3.2 List of reference genes and target genes primer set sequences ……….…………………….66
Table 3. 3. Slopes and Efficiency of gene amplification performing RTq-PCR ……………………….77
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CHAPTER 1
GENERAL INTRODUCTION AND THESIS OBJECTIVES
Clostridium difficile is an intestinal opportunistic pathogen for humans and animals. This organism represents the most common cause of hospital associated diarrhea in North America, Europe and Asia and is responsible for nearly all cases of pseudomembranous colitis (Poutanen & Simor, 2004; Chang et al.,
1979). Infection by this bacterium is facilitated by disrupting intestinal microbiota after antibiotic therapy
(Bartlett, 2002) and is characterized by the complexity of its associated disease, as well as the wide range of its clinical features (Hedge et al., 2008; Mylonakis et al., 2001). C. difficile is a sporogenic microorganism that is ingested as an aerotolerant spore, which then germinates into a strictly anaerobic vegetative cell (Rupnik, 2015). The vegetative cell of C. difficile is an anaerobic bacterium that colonizes the intestinal tract and causes a range of illnesses from mild diarrhea to megacolon and may ultimately lead to death (Kuijper et al., 2007).
Recently, C. difficile has been known as a rising nosocomial and community acquired pathogen, due to several factors including 1. Extensive use of antibiotics and proton-pump inhibitors (Jump et al., 2007;
Carroll & Bartlett, 2011; Lin et al., 2015; Santos-Schaller et al., 2016), 2. An increased elderly population,
3. Longer life span that leads to longer stays in long term care facilities, and means more people are convalescing outside hospitals; and 4. C. difficile genome that is vigorously dynamic. Indeed, research shows that the number of nosocomical infections by C. difficile are higher than recorded strains of methicillin resistant Staphylococcus aureus (Voelker, 2010). The antibiotics vancomycin and metronidazole have been the most available option for treatment and more recently fidaxomicin has been used for C. difficile Infection treatment.
Although antibiotic treatment is the prominent solution for C. difficile infection (CDI) (Bartlett & Perl,
2005), unfortunately, 25% of antibiotic treatments are not successful and causes C. difficile reinfection
(Burke & Lamont, 2014; Hansen et al., 2013) and chances of reinfection increases to 50% after the second
1 infection. This is due to the emergence of a highly virulent strain (ribotype 027) of C. difficile in the recent years, which is extending its impact from hospitals into community settings, and is moving from elderly and debilitated hosts into a younger and lower-risk population. The reported case-fatality rate in North
America is 6%–30% and seems to be rising (Kelly & LaMont, 2008).
An increase in both incidence and severity of this disease, along with the inconsistent results of antibiotic therapy, demands an alternative antibacterial remedy using a different strategy to control infection and pathogenicity of C. difficile. A non-antibiotic treatment is necessary that protect the intestinal microbiota and could be used alone or collateral to the existing treatments. Currently, medical science is searching for preventive therapies or alternative antibacterial which do not increase multidrug resistance. Probiotic bioactive molecules are natural substances that are capable of prevent or treat infectious diseases (Nordeste et al., 2017), and are worthy of investigation as a natural and cost effective remedy to control Clostridium difficile reinfection. Also, newly developed experimental methods including “fecal flora transplant” and
“intravenous immunoglobulins and vaccines” are targeting C. difficile toxins (Seekatz & Young, 2014;
Shah et al., 2014; Stiles et al., 2014) but these newly developing treatments have limitations due to the immune systems of patients or CDI recurrences.
Bioactive peptides are specific amino acid sequences that may control infection and which exhibit regulatory functions in the body. There are advantages of using bioactive peptides as an alternative therapy for pathogen control, since bioactive peptides maintain the commensal microbiota without disturbing them and stimulate the immune system of the host as a result of their reaction with the body’s systems (Ruiz et al., 2014). Furthermore, using bioactive peptides may not lead to the development of antibacterial resistance because these small peptides do not kill or interfere with viability of the pathogens; rather they down- regulate their virulence genes (Sharma 2014).
Studies have shown that probiotic bioactive peptides are able to suppress biological activity of pathogenic microorganisms by down-regulating virulence gene expression in pathogens including Salmonella
Typhimurium (Brovko et al., 2003; Vinderola et al., 2007; Tellez et al., 2011). An in vitro study using a
2 bioluminescence assay confirmed that the bioactive fraction from milk fermented with probiotic
Lactobacillus helveticus down-regulated S. Typhimurium virulence gene ssrB. It also reduced clinical manifestations in mice infected with Salmonella (Tellez et al., 2011). Other studies showed that
Bifidobacterium bifidum also produced bioactive molecules that interfere with both attachment and invasion of S. Typhimurium and Escherichia coli O157:H7 (Bayoumi & Griffiths, 2012) and they also were able to down-regulate both hilA and ssrB genes carried on pathogenicity islands 1 and 2, respectively, in S.
Typhimurium (Bayoumi & Griffiths, 2010).
Moreover, bioactive molecules from L. acidophilus La-5 grown in whey protein medium exerted a protective effect against E. coli O157:H7 EHEC in a mouse model by preventing colonization and attachment of pathogen cells or due to reduction of virulence genes (Zeinhom et al., 2012). L. acidophilus
La-5 bioactive molecules also affected E. coli O157:H7 virulence by inhibiting colonization in the GI tract of mice (Medellin-Peña & Griffiths, 2009; Medellin-Peña et al., 2007). Another study by Mundi et al.,
(2013) reported that cell free medium of L. acidophilus La-5 and B. longum down-regulated Campylobacter jejuni virulence genes.
Furthermore, Yun et al., (2014) studied the effect of L. acidophilus GP1B to control C. difficile associated disease by investigating regulation of C. difficile toxin genes both in vitro and in a mouse model. They showed that a L. acidophilus cell extract decreased expression of genes luxS, tcdA, tcdB and texR genes.
All of the research mentioned above has focused mainly on extracting bioactive peptides from a modified medium containing milk or concentrated whey protein to support the growth of the probiotic L. acidophilus or other strains including Lactobacillus helveticus. These bioactive peptides have been termed
“proteobiotics” recently by Nordeste and colleagues (2017) which their research approved proteobiotics could be used as a food supplement or adjunct to reduce antibiotic use in farm animals. Proteobiotics differ from organic acids or bacteriocins in that they mainly affect quorum sensing, the cell-to-cell communication of pathogenic bacteria. Bacteria communicate through their quorum sensing system to adapt to environmental changes or to invade their host. Bioactive molecules of proteobiotics interfere quorum
3 sensing, modulate virulence of pathogens and inhibit adherence and colonization of their host cells.
According to a patent registered by Griffiths et al., (2009), novel molecules from bioactive molecules fractions produced by L. acidophilus La-5 are low molecular weight, proteinaceous, heat stable molecules.
They have amino acid sequences that include: YPVEPF, YPPGGP, YPPG and NQPY. It has been reported by the same author that these novel molecules are effective to inhibiting viral infection such as norovirus and Hepatitis A (Morales and Griffiths, unpublished results).
Hypothesis
We hypothesized that bioactive molecules of L. acidophilus La-5 produced in their culture media
(proteobiotics) could reduce the pathogenicity of C. difficile including toxin production and adherence of this pathogen. We also hypothesized that proteobiotics would affect some of the biological activities which involve in virulence of C. difficile, more specifically, biofilm-forming and motility. Instead of using live bacteria, we used cell-free supernatant (La-5 CFS) of bacterial culture to investigate the application and efficacy of L. acidophilus La-5 bioactive molecules to reduce the virulence of C. difficile.
Objectives
The objectives of this research were to:
1. Investigate the influence of La-5 CFS on quorum sensing using autoinducer bioluminescence (AI-2) assay; evaluate expression level of virulence genes of C. difficile particularly the expression of the toxin genes (tcdA, tdcB) and non-toxin genes of three C. difficile strains (Chapter 3).
2. Evaluate the effect of La-5 CFS on colonization and toxin production of the C. difficile in vitro, using human epithelial cells in adhesion and cytotoxicity assays. In addition, the impact of La-5 CFS on reducing efficacy of toxins to disrupt the cancer cell’s integrity was examined (Chapter 4).
3. Observe and compare the biological activity of C. difficile after exposing to La-5 CFS measured by biofilm formation, sporulation, and motility assays (Chapter5).
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The project was divided in four different stages. 1) At first, concentrated cell free culture medium (CFS) of different strains of probiotic lactic acid bacteria were prepared for a preliminary screening of their viability and growth pattern in two modified media 2) Appropriate concentrations of cell free cultures of the selected strain, L. acidophilus La-5, were used to evaluate its effect on quorum sensing and regulation of C. difficile virulence genes expression using real-time quantitative PCR (q-PCR). 3) The third stage of the project involved an in vitro investigation of the influence of La-5 CFS on C. difficile toxin production and proliferation of epithelial cells in the presence of C. difficile toxins. In addition, the impact of La-5 CFS on the adherence of C. difficile to Caco-2 cells and HT-29 were determined. 4) The mode of action of the bioactive molecules in La-5 CFS on the physiological behavior of C. difficile was investigated by evaluating the effect of La-5 CFS on biofilm formation, sporulation and motility.
If the bioactive compound is shown to be effective on a broader spectrum of pathogens, including Gram- positive and Gram-negative, aerobic and anaerobic bacteria, then producing these molecules on a large scale will be much more efficient and cost effective, compared to the cost of development of a new antibiotic. The outcome of this research will be the development of an alternative natural therapy, which can be a substitute for antibiotics and lead to a decrease in the problems of antibiotic-resistant bacteria. In addition, development of a bioactive molecule will benefit not only public health and health care systems, but will also help the Canadian economy by reducing costs of hospitalization and treatment. The research outcome has potential to prevent outbreaks and reduce antibiotic use both in livestock and humans. This project will also increase our understanding of the mechanism of action of probiotics.
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CHAPTER 2
LITERATURE REVIEW
2. Clostridia
Prokaryotes classified as Clostridia range from restricted anaerobes to aero tolerant bacilli. Clostridia are usually catalase negative, have a variety of colony morphologies, mainly uneven and large colonies (>2mm) and in some species, exhibit crawling behavior on the agar plate (Holt et al., 1994). Clostridia are generally in water, soil, plants and normal intestinal flora of humans (Cabral, 2010). Clostridia are well-known due to their medical significance as well as their biotechnological potential.
Clostridium, which belongs to the class of Clostridia is one of the most heterogeneous and numerous species among prokaryotic genera and is composed of about 200 species (Andreesen & Gottschalk, 1989). Some of these species are pathogenic and produce exotoxin (s) that is a key factor of their pathogenicity. Species such as Clostridium botulinum, Clostridium tetani, Clostridium perfringens and Clostridium difficile are the most important human pathogens, responsible for diseases such as botulism, tetanus, gas gangrene and colitis, respectively (Galperin et al., 2012). These species possess an outstanding ability to harm their host due to their ability to overcome harsh conditions by sporulation, growing quickly in a nutritious and oxygen- depleted environment, and producing enterotoxins and neurotoxins (Wells & Wilkins, 1996).
Some species such as Clostridium noveyi, are lethally toxigenic. However, this same species is important in clinical research when used as a bacteriolytic anti-cancer agent by destroying the tumor without harming the host (Bettegowda et al., 2006). In addition, a non-pathogenic Clostridium acetobutylicum came to the attention of researchers for its role in production of bio-butanol. Bio-butanol is a promising biofuel that is produced from renewable biomass fermentation and efficiently produces energy (Kudahettige-Nilsson et al., 2015; Lee et al., 2008).
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The Clostridium genus produce a broad range of toxins, including the most famous and the most lethal ones: BoNTs, clostridial neurotoxins (Clostridium botulinum neurotoxins) that result in muscular paralysis named botulism, and the TeNT (Clostridium tetani neurotoxin), which causes spastic paralysis (Pellizzari et al., 1999). TcdA and TcdB are C. difficile toxin A and toxin B, respectively, which are inflammatory enterotoxins (Jank & Aktories, 2008) and they are known as C. difficile main virulence factors.
2.1 CLOSTRIDIUM DIFFICILE
2.1.1 Clostridium difficile Microbiology
Clostridium difficile is a motile, gram-positive, obligate anaerobic, spore-forming bacillus that belongs to the family of Clostridiaceae, the class of Clostridia and the phylum of firmicutes (Warny et al., 2005; Carlet et al., 2011). Clostrididum difficle can colonize not only the human intestine, but also commonly inhabit the intestinal tract of animals used in food production including cattle, chickens, pigs, rabbits, and sheeps; and even in pets such as cats and dogs. Their transmission can be from animal to human and vice versa
(Gould & Limbago, 2010; Keessen et al., 2011; Weese et al., 2010). C. difficile can be found in soil and water (Finegold & Martin, 1982) and also it can be transferred through food like meat and vegetables to humans (Gould & Limbago, 2010; Harvey et al., 2011; D. Metcalf et al., 2010).
C. difficile produces two large clostridial toxins (LCT) and can exist in two forms: vegetative cells or spores.
Their vegetative cells are metabolically active in a reproductive state and very sensitive to oxygen.
However, their spores are not metabolically active. Physiologically, C. difficile can survive harsh aerobic environments; common sterilization methods, including high cooking temperatures (74oC), ultraviolet light, and harsh chemicals (Setlow, 2007; Heinlen & Ballard 2010). One specific characteristic of C. difficile that adds to its pathogenicity is that, compared to other clostridia, it is highly transmissible through their spores from person to person. The species was named “difficile” as it was difficult to grow and isolate (Kelly et al., 1994). Under laboratory conditions, C. difficile grows to large numbers at 35 - 40oC, within 24 - 48 h, in rich medium; the addition of sodium taurocholae stimulates spore germination (Wilson, 1983). C. difficile
7 has an extremely variable genome, which may explain its capacity to adapt and survive in harsh conditions and grow within the gut environment (Sebaihia et al., 2006). Their genome has a circular chromosome of over 4000-4300 kbp and a plasmid of 7-8 kbp (Sebaihia et al., 2006). C. difficile shows only 15% coding sequences in common with other four clostridial genomes; Clostridium acetobutylicum, C. botulinum, C. perfringens and C. tetani, whereas, 50% are unique to C. difficile (Sebaihia et al., 2006).
Killgore et al. in 2008 demonstrated the most commonly used molecular typing methods that classify strains include: toxinotyping (on the basis of tcdA/tcdB sequence polymorphisms); ribotyping (16s-23s rDNA intergenic spacer region patterns); pulse-field typing and restriction endonuclease analysis typing (whole genome restriction pattern polymorphisms); MLST and MLVA (multi-locus repeat- or non-repeat-based sequence variations); and SLP typing (surface-protein variations).
Tenover and colleagues (2011) reported that the most recent C. difficile-associated diseases are related to new highly infectious strains such as “ribotypes 001, 017, 027, 078 and 106 [toxinotypes 0, VIII, III, V, and pulse-field types NAP2, NAP9, NAP1, NAP7/8 and NAP1, respectively]. Among these strains, C. difficle NAP1/BI/027 is the most commonly isolated strain from outbreaks and it has caused several outbreaks in the U.S (40 states), Canada (seven provinces), England and other European countries, and Asia
(Voelker, 2010). A significant increase in severity and morbidity of C. difficile infections has made this bacterium a serious challenge for the health care system.
2.1.2 C. difficile Infection (CDI) and C. difficile associated diseases (CDAD)
C. difficile is one of the greatest concerns that healthcare systems face today. According to the CDC
(Centers for Disease Control), 5% of hospitalized patients are infected by one of the health care associated diseases such as C. difficile infection or methicillin resistant Staphylococcus aureus (MRSA). Clostridium difficile causes a range of diseases among hospitalized and out-patients. These diseases are often called C. difficile associated disease (CDAD) or C. difficile infection (CDI) (Bartlett, 2006; Smits et al., 2016;
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Heinlen & Ballard, 2010). A study show that CDAD is increasing compared to MRSA (Miller et al., 2011).
This increase has occurred in North America and Europe since 2000, and now is becoming a prominent concern in Asian countries (Burke & Lamont, 2014; Gerding & Lessa, 2015). Kelly and Lamont in 1998 reported that 3% of healthy adults and 20% of treated patients carry C. difficile (Table 2.1) and asymptomatic individuals as well as infected people are also sources of C. difficile infection (Gould &
McDonald, 2008). Crogan & Evans, (2007) state that CDAD is responsible for 25% of deaths in vulnerable elderly people. An increasing number of fatal cases of CDAD during the last two decades has made C. difficile among the most important pathogens in the United States (Ananthakrishnan et al., 2008; McDonald et al,. 2005; O’Brien et al., 2007).
Clinical expression of CDAD depends on the virulence of the strains and reactions of the host’s immune system. C. difficile is part of the commensal flora of 1-3% of adults, and their spores are also present in healthcare settings. Therefore, CDI occurs when spores are ingested by a susceptible host through contact with a contaminated surface or eating contaminated food or water. Under favorable conditions, spores germinate in the intestine and vegetative cells colonize and produce toxins in the colon (Figure 2.1). CDI usually begins with symptoms of mild to severe diarrhea, and progresses with fever and bloating. As soon as bacteria colonize and produce toxins, they continue to harm the intestinal cells. Toxins result in inflammation forming pseudomembranous due to accumulation of dead epithelial and leukocyte cells, fibrin and mucous layer. When these layers are taken off, it results in pseudomembranous colitis and toxic mega colon, a life threatening disease (Knoop et al., 1993).
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Table 2.1. Isolation rates of toxigenic C. difficile from stool of various subject populations
Subject population C. difficile positive Pseudomembranous colitis 95-100% Hospital in-patients 20% Healthy adults 0-3% Healthy neonates and infants 25-80%
Table has been modified from Kelly and LaMont, (1998): Clostridium difficile Infection
Figure 2. 1. Schematic of human C. difficile infection. Spores, vegetative cells, bacterial factors and host factors that impact/modulate disease are depicted. Adapted from Vedantam et al., 2012.
2.1.3 Epidemiology and Variation in Virulence Factors of C. difficile Strains: Hypervirulent Strains
As stated earlier, the recent rise of C. difficile infection in hospitals and in communities can be related to several factors: inadequate hygiene in hospital environments and increase of antibiotic prescription as well as hospital design, highly dynamic bacterial genome, long-term care residency, longer lifespan (older than
10
65 years, resulting in decreased immunity) and increased visits to healthcare facilities (Jump et al., 2007;
Stiles et al., 2014; Borgmann et al., 2008; Cartman et al., 2010; Hedge et al., 2008). Another reason for the increasing number of C-difficile outbreaks is the use of gastric acid suppressants, including proton pump inhibitors and histamine type 2 blockers (Khanna & Pardi, 2012). Immunocompromised patients, such as people with HIV, cancer, or those who have had organ transplants (Maschmeyer & Haas, 2008) are at risk of CDI as well. A recent study shows that CDI has risen in surgical patients (Sartelli et al., 2015) and in children that have cancer or undergone cell transplants (Nicholson et al., 2015).
Antibiotic medication, especially antibiotics which are anti-aerobes, are associated with CDI (McFarland,
2006; Pepin et al., 2005). As a result, a highly resistant virulent strain of C. difficile (027/ribotype NAP1) triggered after wide use of fluoroquinolone in hospitals and communities (Barbut et al., 2011). The emerging clone of C. difficile, named as ribotype 027 or hypervirulent strain, is a toxinotype III, North
America PFGE pulsotype 1 (NAP1) and restriction endonuclease analysis type BI. The strain 027/NAP1/BI became epidemic and caused more complex disease and higher fatality as well as more resistance to metronidazole (Pépin et al., 2004). Consequently, CDI has increased specifically three to eight times in the
USA (Gerding & Lessa, 2015) and severe clinical types of infection have increased; resulting in a 50% mortality rate (Venugopal et al., 2013). In Europe, increasing hypervirulent strains of C. difficile
(027/NAP1) are responsible for more severe and complicated clinical forms and recurrences (Davies et al.,
2014; Loo et al., 2005; Pépin et al., 2004).
Incidence of this dominant virulent strain varies with region, which is partly due to differences in toxin testing and reporting methods. The most popular toxin test methods have been based on ELISA tests, which have disputed sensitivity and reliability. Mitchel and colleagues (2012) argued for standard testing in hospitals including testing methods, study duration and testing effort. In their review article, they demonstrated the need for standard testing as well as incident reporting methods nationally and internationally. Currently the approved diagnosis test is detecting the C. difficile toxin genes directly from the stool specimen using PCR (nucleic acid amplification technic) (Mitchell et al., 2012).
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Strains such as ribotype 027 and ribotype 078 are the two most common strains in patients with CDI in
North America and Europe (Bauer et al., 2011; Hensgens et al., 2011). Community-acquired C. difficile infections have also increased with ribotype strain 001 causing 60% of cases (Wilcox et al., 2008).
Additionally, incidence of C. difficile has risen as a pathogen or commensal in different animals such as pigs, calves and chickens (Wilcox et al., 2008) and PCR ribotype 078 is the most often detected strain in cattle and swine (Janezic et al., 2012). This change in epidemiology is the consequence of emerging “hyper virulent” variants of C. difficile, which results in a greater incidence, and severity of illness and more fatalities (Denève et al., 2009). Warny et al., in 2005 demonstrated that hypervirulent strains are able to produce more toxins (resulting in a 16-fold increase in production of TcdA and a 23-fold increase in production of TcdB) or spores in both exponential and stationary phases of their growth duration.
2.1.4 Disease Transmission
C. difficile spores are abundant in hospital facilities and also exist in young children, some healthy or asymptomatic populations, the environment, and food supply, that transmit infections nosocomially and into the community (Barbut et al., 2011; Rupnik et al., 2009). Although it is known that C. difficile is transferred through the oral-fecal-route (Kelly et al., 1994) , another study by Otten et al., in 2010 described a model that shows the many ways by which this bacterium may be transmitted. These methods of transmission are: consumption of contaminated food /water, person-to-person contact among both diseased and colonized individuals working in hospitals, long-term care centers and veterinary clinics, and animal- to-person transmission from pets and domestic animals (Buggy et al., 1985; Otten et al., 2010; Rodriguez-
Palacios et al., 2006).
Recent studies have isolated C. difficile from retail meats, pet food, and livestock and there is a concern that handling and consumption of food may play a role in transmission of community-acquired infection.
With respect to food safety, a study in 2005 in two Canadian provinces reported 20% of retail meat products
12 were contaminated with C. difficile (Rodriguez-Palacios et al., 2007) including 21% of ground beef and
14% of ground veal. A similar study in the United States by Songer et al., in 2009 demonstrated 42% contamination in a variety of raw and ready-to-eat products. Geographical or seasonal differences may cause variation in the prevalence of C. difficile in food and food products. Contamination during processing and final preparation of food by an infected family member or food handler could also result in transmission of C. difficile. Rodriguez-Palacios (2007) reported that spores of this bacterium can survive in ground beef cooked at 71oC for at least 120 minutes. These findings suggest that C. difficile can be commonly found in retail food and it is possible that meat contamination originated from animal hides contaminated with the spores or by internal contamination of the skeletal muscles (Vengust et al., 2003).
2.1.5 History of Clostridium difficile infectious disease
C. difficile and its toxigenic potential has been known since 1935 when Hall and O’Toole first found it during research on normal microflora of new borne infants. Because they had difficulty cultivating this organism, they called it Bacillus difficilis. They reported that a strong and lethal neurotoxin was produced from this bacterium, but there was no clinical reference that could project pathogenicity of this anaerobic bacterium (Hall & O’Toole, 1935). Although there was evidence of a link between antibiotic intake and developing infectious diarrhea, the relationship between those diseases to this bacterium was unknown until the 1970s.
Many years before, in 1893 Finney reported a Pseudomembranous colitis (PMC) ulcer of the intestine in a
22 year-old female patient who died after post-operative diarrhea (Finney, 1893). After the introduction of antibiotics reports of PMC became more common. In that era, Staphylococcus aureus was the only bacterium known as a pathogen of stool microbiology. At that time PMC was called “pseudomembranous enterocolitis” or “antibiotic-associated enterocolitis” (Penner & Bernheim, 1939). In the early 1970s
Tedesco and coleagues (1974) reported several cases of severe diarrhea attributed to clindamycin, an antibiotic that was preferred for anaerobic bacterial infection. Upjohn Company, the producer of clindamycin, sponsored research on patients who had diarrhea during clindamycin intake. This study 13 showed that 21% of patients given clindamycin had diarrhea and 10% had PMC at endoscopy that was distinguished by FDA as a “potential life threatening complication with the use of this drug” (Bartlett,
2009).
In 1974, Hafiz was studying C. difficile and its glutamine dehydrogenase antigen, when he reported that C. difficile is widely distributed in the environment and in the stool of animals including donkeys, horses and camels. While Hafiz and Tedesco were conducting their research, Green et al., (1974), studied stools of animals with cecum inflammation (cecitis). They found that penicillin use induced cecitis associated with a cytopathic toxin. Therefore, 1974 was a milestone year in the history of research on C. difficile infectious disease. Three different studies were relevant to CDI but the link between the toxins that Green detected
(Green, 1974), the bacterium that Hafiz was working on (Hafiz, 1974) and the complex disease that Tedesco described (Tedesco et al., 1974) was not established.
A few years later, C. difficile was recognized as a bacterium that produces toxin and causes pseudomembranous colitis in patients treated with clindamycin (Bartlett et al., 1978; Tedesco & Alpers,
1974). In 1978, a tissue culture test led to the establishment of the cytotoxin assay as a test method to demonstrate the cytopathic effect of toxin that caused actinomorphic changes in tissue culture fibroblast cells (Chang et al., 1979). After that several other groups began CDI research to detect the cause of antibiotic associated PMC including Bartlett, Burton, Keighley, Fekety and Silva (Bartlett, 2009).
2.2 CLOSTRIDIUM DIFFICILE VIRULENCE FACTORS
2.2.1 Non-toxigenic Virulence Factors: Colonization and Adherence
Clostridium difficile infection (CDI) has two main phases that progress from colonization to toxin production. Colonization is a necessary introductory stage of pathogenesis that happens when a range of adhesion factors are activated. After disruption of the normal intestinal microbiota of a host as a result of antibiotic chemotherapy, C. difficile spores germinate and multiply. Bacteria adhere to intestinal mucus and cover the epithelial surface of the gastrointestinal (GI) tract. Then, they penetrate the mucus layer using
14 their flagella and proteases (Tasteyre et al., 2000) resulting in colonization; this is the first phase of pathogenesis (Shim et al., 1998).
Gram-positive cells have a different surface structure and diverse proteins that affect their pathogenic mechanism in their host such as disrupting tissue, adhesion or other factors that control bacterial growth and metabolism. C. difficile is one of the important Gram-positive species that possesses “proteinaceous” surface layer proteins (S-layer) for their adhesion to host cells. The S-layer is comprised of various proteins arranged in a crystalline lattice (Sara & Sleytr, 2000), with surface layer protein A (SlpA).
Vedantam et al., (2012) established that SlpA is a major contributor to C. difficile attachment in vitro by exposing the C. difficile to cell cultures pretreated with anti-SlpA antibodies and measuring adherence of the bacterium to the cell culture in comparison with the unexposed cells with antibodies. Two major proteins are encoded by slpA gene: proteins P47 (high molecular weight) and P36 (low molecular weight). Cell wall proteins encoding genes are located in the same 37-kbp DNA cluster in the C. difficile genome (Karjalainen et al., 2001).
Studies showed slpA genes encode cell adhesion factors including surface-associated proteins Cwp66,
Fbp68, GroEL and the S-layer protein P47. These proteins moderate attachment of bacteria to the epithelial cells (Calabi, 2002; Denève et al., 2008; Hennequin et al., 2001; Hennequin et al., 2003; Waligora et al.,
2001). Several studies concluded that surface associated proteins always express during CDI to induce immune responses in CDAD patients (Cerquetti et al., 1992; Drudy et al., 2004; Pechine, 2005; Pechine et al., 2005).
In addition, Cwp84, a cysteine protease that is able to degrade proteins of the extracellular matrix of host intestinal tissue, disturbs cell integrity in the early stages of CDI (Denève et al., 2008; Janoir et al., 2007).
Cwp84 is one of two cysteine proteases that forms the S-layer proteins of C. difficile. It is a “dynamic” molecule with the ability to assemble proteins to the cell surface for merging into the cell walls and also facilitate pathogenicity by its enzymatic nature against the host’s proteins (Vedantam et al., 2012). SlpA and other surface layer proteins (SLPs) are under a larger category of proteins named cell wall proteins
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(CWPs). Studying CWPs may result in the discovery of a way to prevent their action; thereby inhibiting gastrointestinal colonization of C. difficile.
2.2.2 Toxigenic Virulence Factors and Mechanism of Action
Once C. difficile is colonised and established in the GI tract, it begins its second phase: toxin production.
Toxins A and B (TcdA and TcdB) are potent cytotoxic enzymes that damage the human colonic mucosa and their production leads to disorganization of the cell cytoskeleton (Denève et al., 2009). Toxin A
(enterotoxin) attracts and stimulates neutrophils to move toward the ileum with the release of cytokines. It also disturbs cell-to-cell tight junctions, enhances flow of liquids through the gut wall, and begins diarrhea.
Beside, toxin B (cytotoxin), provokes the breakdown of the actin and destroys cellular cytoskeleton both in vivo an in vitro. This causes the loosening of the intestinal epithelial junction and, subsequently, inflammation and diarrhoea (Dingle et al., 2010). Both toxins A and B are homologous glucosyltransferases that are considered to be the primary C. difficile virulence factors (Voth & Ballard, 2005). These two toxins,
A and B, are large clostridial toxins (LCTs) and have a molecular mass of 308 kDa and 269.6 kDa, respectively (Kuehne et al., 2010).
All of the Clostridium spp. belonging to this LCT family affect their target cells by modifying GTPases
(enzymes that can bind and hydrolyze guanosine triphosphate). These high-molecular-weight proteins are transported to the cytoplasm where they link to specific receptors on the luminal side of the colonic epithelium (Pothoulakis et al., 1996). In the LCT category, there is also a family of large clostridial glucosylating toxins (LCGT). They suggested that toxins are single stranded proteins that have four structural domains and specifically include toxins from Clostridium difficile (TcdA and TcdB), lethal toxins from Clostridium sordellii (TcsL) and α-toxins from Clostridium novyi (Genth et al., 2014; Guttenberg et al., 2011; Just & Gerhard, 2004; Popoff & Bouvet, 2009; Varela Chavez et al., 2015).
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Studies show that although there are more colonized C. difficile in the large intestine of human infants, they are more resistant to CDAD than adults and it is hypothesized that lack of toxin receptors in infants prevents
CDAD at an early stage of a mammal’s’ life (Hall & O’Toole, 1935; Pothoulakis et al., 1996). While human infants are resistant to C. difficile infection, newborn pigs are infected by C. difficile. The reason for this could be the α–galactosyl antigen, which is more abundant in pigs than in other species and is absent in humans. If this antigen that is made in the early stages of a pig’s life is related to their disposition to CDI, this may explain the difference between human and animal infants (Keel & Songer, 2007).
TcdA protein, has short and long multiple repeated sequences at the end of C-terminal receptor binding sites, and are responsible for finding surface receptors. The TcdA protein and its identified receptors including Gala1-3b1-4GlcNAc-R (α -galactosyl or the α-Gal epitope) have been the focus of many well- documented studies (Keel & Songer, 2007; Lowe et al., 1980; Tucker & Wilkins, 1991). In depth studies have been conducted to investigate toxin structure in order to gain more insight into their molecular mechanism and their intoxication actions. Negative stain electron microscopy showed that toxins A and B possess a similar structure and sequencing showed that they share 68% sequence similarity and 47% sequence identity (Pruitt et al., 2010). Albesa-Jové and his colleagues (2010) studied C. difficile toxin structure by X-ray crystallography and small angle X-ray scattering models (SAXS) of TcdB. The two toxins contain a biologically active N-terminal domain (glucosyl-transferase domain); a cysteine protease domain; a middle translocation (transmembrane domain) section; and a C-terminal receptor-binding domain (Davies et al., 2011; Vedantam et al., 2012; Yang et al., 2008) (Figure 2.4) but the precise functions of all the toxin domains are, as yet, unknown.
Using compounds which are able to interact with the C. difficile receptor-binding domain or inhibit their glucosyltransferase activity are feasible approaches toward controlling infection (Jank & Aktories, 2008).
Heerze et al., (1994) proposed a strategy that manipulates host cell receptor analogs to inhibit TcdA and
TcdB binding to the epithelial cells in the intestine. They applied direct electrospray ionization mass
17 spectrometry (ES-MS) assay and cytotoxicity neutralization assay to check if human milk oligosaccharides
(HMOs) are able to bind to toxins and to protect Vero cells in vitro. However, their data showed that HMOs do not significantly inhibit the cytotoxic effects of TcdA or TcdB. The absence of protection was attributed to the very weak intrinsic affinities that the toxins exhibited toward the HMOs (El-Hawiet et al., 2011).
The genes encoding toxin A and toxin B are tcdA and tcdB, respectively, which together with three other accessory genes tcdR, tcdC, and tcdE (Figure 2.2) are located within the 19.6 kb pathogenicity locus
(PaLoc) ( Dupuy et al., 2008; Dingle et al., 2011). While tcdR and tcdC, are negative and positive regulators of toxin production, respectively, tcdE encodes a putative holin and its role in pathogenicity of C. difficile is not known yet. Transcription of these genes depends on the growth phase: tcdC transcription mostly occurs during the logarithmic phase whereas the four other genes are transcribed during the stationary phase
(Borriello, 1998).
Figure 2.2 Genetic organization of C. difficile pathogenicity locus (PaLoc). Adapted from (Dupuy et al., 2008).
Two transcription regulators TcdR and TcdC control the expression of the two toxin promoters in the
PaLoc. TcdR is a transcriptional activator that positively regulates toxin genes (Mani et al., 2006; Mani &
Dupuy 2001; Moncrief et al., 1997). The tcdR gene acts as an RNA polymerase sigma factor that activates tcdA and tcdB transcription (Mani & Dupuy 2001) and also activates expression of itself too (Figure 2.3)
(Mani et al., 2002). TcdR is found in other Clostridium species like C. botulinum (BotR) and C. tetani
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(TetR). Although it has a different structure in every species, TcdR is still a positive regulator of neurotoxins in those bacteria (Dupuy et al., 2005; Raffestin et al., 2005; Mani & Dupuy, 2001; Mani et al., 2002). On the contrary, TcdC acts as a negative regulator of toxin synthesis (Carter et al., 2011; Hundsberger et al.,
1997; Matamouros, 2007).
Multiple studies show that deletion of tcdC increases toxin production in epidemic C. difficile (NAP1/ 027) strains (Carter et al., 2011; Hundsberger et al., 1997; McDonald et al., 2005; Warny et al., 2005).
Additionally, there is no evidence of any change in toxin synthesis due to tcdC mutation (Bakker et al.,
2012), which indicates that there might be other elements interfering with the toxin production system
(Darkoh et al., 2015). This recent study (Darkoh et al., 2015) also, suggests a new ‘thiolactone quorum- signaling peptide’ that is associated with toxin production in C. difficile strains and is not controlled by
‘tcdC-mediated regulation.’ This finding can be used in developing future antibiotics to target the newly found regulatory peptide to block the quorum signaling pathway of C. difficile effectively. The role of TcdE has yet to be confirmed but it is similar to phage holin proteins, and it can help the toxin to be released into the environment and penetrate into the host cells (Tan et al., 2001).
There has been a dispute over the importance of toxin A or toxin B in C. difficile infection. Lyras et al.,
(2009) determined that strains expressing toxin B are more virulent and the B toxin is more important than toxin A. Kuehne and colleagues (2010), by using a Clos-Tron system, were able to re-establish that both toxins are vital in infecting the host and both toxins must be measured in diagnostic tests for effective results. The most recent study by Chumbler et al., (2016) demonstrated the difference between the toxicity of TcdA and TcdB. They examined different concentrations of each toxin at different time points by using mouse colonic epithelial cells and measuring lactate dehydrogenase (LDH) release. They concluded that both toxins can damage cell membrane integrity but TcdB causes more rapid harm to cells than TcdA. Also they reported that TcdB kills the cells by a different mechanism when their concentration is higher than 100 picomolar. TcdA toxicity causes “an apoptotic cell death” and depends on glucosyltransferase activity of
19 the toxin. TcdB applies different mechanisms to intoxicate the cells depending on its concentration. Its
“bimodal mechanism” is glucosyltransferase-dependent when the concentration is low and is glucosyltransferase-independent when its concentration is high and consequently causes necrotic death in cells (Chumbler et al., 2016).
According to the literature, there is a significant difference between the Gram-negative pathogen
Escherichia coli and Gram-positive C. difficile pathogenesis regarding the mechanism of action of their toxins. Unlike Escherichia coli heat-stable toxin, and cholera toxin, that influence intracellular levels of cyclic AMP or GMP, C. difficile toxins break down their targeted cells in two ways: direct cytotoxication and indirect cytotoxication. The mechanism of direct intoxication is due to the toxin A and toxin B glucosyl- transferase domains (Figure 2.3). CD Toxins inactivate small regulatory proteins [Rho-GTPase (Rac, Rho,
Cdc42)] of eukaryotic cells after quick addition of a glucose moiety to a specific threonine on them. Since
Rho proteins are regulators for the actin cytoskeleton, when they are deactivated by glucosylation, this mechanism leads to cytoskeletal breakdown in the target cells (Aktories & Just, 1995) and the disruption of some other processes that are controlled by GTPases (Schmidt et al., 1996). In summary, these toxins inactivate the host GTPases, which leads to alteration in the actin cytoskeleton and disruption of barrier function and finally cell death (Just et al., 1995a; Just et al., 1995b).
Toxin A and Toxin B also induce inflammatory responses that cause tissue damage and sometimes leads to pseudomembranous colitis (PMC). Their action provokes inflammatory cytokine interleukin IL-1β production (IL-1β is a group of cytokines that controls immune responses to infections) in a tissue culture model (Vedantam et al., 2012). However, studies show that the mechanism of action of C. difficile toxins in general, has some similarity with some other bacterial toxins that target Rho proteins; including toxins from Clostridium sordellii and Clostridium novyi (cyto-toxins), which add a glucose to Rho, and toxins from Bacillus cereus and Staphylococcus aureus that also alter the Rho family proteins (Kelly & LaMont,
1998).
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The reason that these two toxins have different mechanisms is unknown but one assumption is that they might have different receptors to engage. The human receptors for TcdA are not known yet, but there are two receptors that have been described for TcdB: poliovirus receptor-like protein 3 (PVRL3) and chondroitin sulfate proteoglycan 4 (CSPG4) (LaFrance et al., 2015; Yuan et al., 2015). It has been shown that PVRL3 can mediate both necrotic and apoptotic mechanisms. Therefore, lack of binding of such a receptor for TcdA in the hosts is the reason that toxin A performs differently from TcdB for killing the cells
(Chumbler et al., 2016).
Figure 2.3. Toxin regulation model (txeR: tcdR; PtxeR: tcdR promoter PtoxA: toxA promoter; PtoxB: toxB promoter; RNAP: RNA polymerase) Picture was taken from Mani and Dupuy, 2001. This model illustrates the role of txeR in response to an extracellular signal from stress or lack of nutrients. Increasing text synthesis direct “RNA polymerase core enzyme” and connect it to the tox (toxin) promoter, then activates transcription of toxin genes tcdA and tcdB.
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Figure 2.4. Domain organization of C. difficile toxins. (A) The LCTs are composed of four domains, the glucosylating enzymatic domain ‘A’ (dark blue), the autocatalytic processing domain ‘C’ (green), the translocating domain ‘D’ (magenta) and the binding domain ‘B’ (purple). Source: Super toxins from a super bug: structure and function of Clostridium difficile Toxins (Davies et al., 2011).
2.2.3 Binary Toxins
In addition to toxin A and toxin B, the genome of some C. difficile virulent strains encodes a binary toxin called C. difficile transferase (CDT) that is composed of an enzymatic component encoded by cdtA, and cdtB genes. The binary toxin is an ADP-ribosylating toxin, made up of two separately fabricated components: an enzymatic component (CDTa) and the transport component (CDTb); the latter component’s role is to translocate CDTa into target cells (Davies et al., 2011). Binary toxins in Clostridium and Bacillus are encoded by chromosome or plasmid-based genes and have 27% - 31% G+C nucleotides (Popoff, 2003).
C. difficile strains, which are more virulent and more dominant, such as C. difficile 027, 078 and CD196, also produce binary toxins, in addition to their two main exotoxins (Perelle et al., 1997). Nevertheless, it is not known whether CDT has any relation to other toxins nor if it contributes to C. difficile pathogenicity
(Metcalf & Weese, 2011).
One study showed that the role of binary toxins is unclear and it is not necessary for C. difficile infection
(Metcalf & Weese, 2011). On the contrary, Bacci et al compared the 30-day case-fatality rate of 212 CDI cases in which strains were toxin A and B-positive, and 265 CDI cases in which the aetiological agent was
Toxin A and B positive as well as CDT positive. One hundred and ninety-three cases were from 027 strain
22 and 72 cases were from non-027 strains. Their study showed that infection with binary toxin-positive strains had a higher case-fatality rate than the other strains. Similar observations were made with strain 027 and non-027 strains. Their study concluded that “CDT is a marker for more virulent strains or directly contributes to strain virulence” (Bacci et al., 2011). One study in 2009 by Schwan et al. suggested that binary toxins may increase adherence of bacterial cells to Caco-2 cells (Schwan et al., 2009).
2.3 C. DIFFICILE OTHER VIRULENCE FACTORS
2.3.1 C. difficile Sporulation and Germination
C. difficile, like any other bacteria of the firmicutes phylum can be present as vegetative cells and endospores. Bacillus spp. and Clostridium spp. are two Gram-positive bacilli of spore forming bacteria that are clinically important. The C. difficile spore morphotype is not as well-known as other spore-forming bacteria like Bacillus subtilis or Clostridium perfringens, but research shows that there are considerable differences between the former and latter bacteria in terms of their outer layer (exosporium and spore coat) and genes encoding spore formation.
Vegetative cells shift to a dormant stage when there are unfavorable conditions, such as lack of nutrient sources including carbon and nitrogen, exposure to oxygen, very acidic conditions, and high temperatures
(Nicholson et al., 2000; Setlow B. & Setlow P., 1980; Setlow, 2006). This state (sporulation) is not related to their reproduction process but to their survival, and they do not exhibit any metabolic activity in this state. Their endospores are very resistant to harsh conditions such as desiccation, high temperatures up to
100 o C, chemicals, oxygen, radiation, and antibiotics. The C. difficile endospore contains the whole genetic material of the bacterium (DNA), a thick keratin like coat, peptidoglycan, cell membrane, very little water, and nutrients from cytoplasm. Spore surface structure includes their exosporium, but their associated surface proteins are not yet completely known and they are potential research subjects for future studies.
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The survival of C. difficile in the environment is due to the persistency of their spores on surfaces in the presence of oxygen. C. difficile spores can survive in the environment for years; when the desirable growth condition is provided, they are able to germinate and return to their active state (Bruns et al., 2010). Spores can germinate when their receptors link to a specific germinant, then monovalent cations (H+, Na+ and K+) and a large amount of calcium dipicolinic acid will be released followed by spore peptidoglycan cortex hydrolysis. In this last stage, the water content of the spore increases to the same level as vegetative cells.
The core expands and activation of enzyme production and metabolism occurs (Setlow, 2003).
Understanding the germination and the mechanism of this transformation helps researchers who are working on therapeutic products to inhibit CDI in humans or animals. C. difficile resistant spores are present on the surfaces of different environments and, to date, it is unknown which exact situation causes germination of spores. Studies that have been conducted by Wilson (1983) showed that the presence of bile acids like taurocholate are key factors to resuscitate the dormant spores. Then in 2008, Sorg and Sonenshein determined that bile acids and glycine are co-germinant factors. They showed that the secondary bile acid, deoxycholate, prevents germination. They theorized that one of the reasons that a healthy host is not infected is because the concentration of deoxycholate is high in their intestinal tract.
Among the normal bacterial flora of the gut, there are some species, including Clostridium scindens that transforms primary bile acids to secondary bile acids and protects against infection (Rupnik, 2015). On the contrary, in a disturbed microbiota secondary bile salt is not produced to compete with bile salts and, therefore, germination is not inhibited. (Sorg & Sonenshein, 2008) also found that, in addition to high absorption of primary bile salts by the colon, there is a higher concentration of cholate derivatives in the large intestine despite the anaerobic condition of the colon, which makes the condition suitable for outgrowth of C. difficile vegetative cells and colonization of the large intestine.
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In a review, Paredes et al., (2005) compared the genome sequence of Bacillus spp. and clostridia and their analysis revealed that Spo0A, a master regulator, its downstream regulators, and signaling molecules are conserved in both genera. Upstream signaling molecules of Spo0A that contributed to phosphorylation and activation is absent or different amongst Bacillus and clostridia genomes. In conclusion, studies have found that germination and colonization of C. difficile happens by synergy of bile acids, antibiotics, and a disturbed intestinal microbiota (dysbiosis) (Burns et al., 2010).
2.3.2 C. difficile Quorum Sensing
Bacteria organize their activity in their environment through their population, and by the information that they receive through cell-to cell communication accordingly. Bacterial communication is often referred to as Quorum Sensing is mediated through the production and detection of secreted signaling molecules, auto- inducers (Ng & Bassler, 2009).This quorum signaling system regulates expression of different genes and allows the population of pathogens to control their energy, to influence their environment and their host
(Reading & Sperandio, 2006). Signaling molecules build up in their surroundings as bacterial cell numbers increase. They then are able to control the environment as a group (Miller & Bassler, 2002; Schauder &
Bassler, 2002). Many pathogens are virulent when they are a group of cells and they control their activity by quorum sensing (Sperandio et al., 1999). Quorum sensing was first reported in regulation of bioluminescence in Vibrio fischeri and Vibrio harveyi (Federle, 2009), and after that it was recognized as a common mechanism in gene regulation in other bacteria.
Gram-positive and Gram-negative bacteria use different auto inducer molecules. Gram-positives use processed oligo-peptides as signaling molecules, produced from auto-inducing peptides (AIPs) and secreted from their cells. When AIP quantity reaches to its high level, it binds to a two-component histidine kinase receptor in the membrane and then after further mechanism, it activates gene transcription of the QS regulon
(Rutherford & Bassler, 2012). Gram-negatives use either acylated homoserine lactones (AHLs) or other molecules whose production depends on S-adenosyl methionine (SAM) as a substrate (Wei, 2011) to
25 communicate and change their physiological activities, which includes: symbiosis, virulence, competence, conjugation, motility, sporulation (Miller & Bassler, 2001).
Quorum sensing participates in a variety of mechanisms such as bioluminescence, sporulation, antibiotic production, virulence factor regulation, and biofilm formation (Dapa et al., 2013; Falcão et al., 2004;
Hammer & Bassler, 2003; Kendall & Sperandio, 2016; Rutherford & Bassler, 2012). Although bacteria are different in their quorum sensing mechanisms, chemical molecules, and related target genes; all of them are able to communicate within their community and change their gene expression and their behavior within the resident community (Miller and Bassler, 2001).
Microorganisms as a population use various pheromones for their activity coordination. Among them, autoinducer-2 (AI-2) is a unique molecule for interspecies communication. AI-2 and its encoding gene luxS have been found in many Gram-negative and Gram-positive bacteria (Surette & Bassler, 1998; Surette et al., 1999). It is interesting that quorum sensing in Vibrio harveyi has features of both Gram-negative and
Gram-positive quorum sensing systems. It uses acylated homoserine lactones as Gram-negatives, but its quorum signaling occurs through a two-component circuit similar to Gram-positives (Bassler, 1999). In addition to that, V. harveyi uses AI-2 in their QS circuit that is used for interspecies cell-to-cell communication (Defoirdt et al., 2008; Waters & Bassler, 2006; Mok et al 2003; Ng & Bassler, 2009). The structure of AI-2 molecules is a furanosyl borate diester, and it has been identified that luxS gene is the responsible gene for producing AI-2, and LuxS protein is one of the major players in enzymatic synthesis of AI-2. Also luxS gene deactivation cancels the AI-2 secretion (Lyon et al., 2001; Bassler 1999; Surette et al., 1999).
AI-2 can play different roles. One of the roles that has been reported in different studies is AI-2’s role in the ‘regulation of pathogenicity in E. coli O157:H7, Vibrio cholerae, C. perfringens, Streptococcus pyogenes, and Shigella flexneri (Day & Maurelli, 2001; Lyon et al., 2001; Ohtani et al., 2002; Sperandio et al., 1999). More than 50 bacteria have homologues of V. harveyi’s luxS in their genome (Miller et al.,
2004). Some species of many bacteria that carry luxS genes are: E. coli, S. Typhimurium, Salmonella Typhi,
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Haemophilus influenzae, Helicobacter pylori, B. subtilis, Neisseria meningitidis, Campylobacter jejuni,
Vibrio cholerae, Mycobacterium tuberculosis, S. aureus, Clostridium perfringens and Clostridium difficile
(Bassler, 1999; Surette et al., 1999).
Later, V. harveyi was used as a reporter of AI-2 quorum sensing bacteria to study other bacterial quorum sensing systems such as E. coli O157:H7 (Delcenserie et al., 2012) , Listeria monocytogenes (Turovskiy &
Chikindas, 2006), Campylobacter jejuni (Mundi et al., 2013) and Clostridium difficile (Carter et al., 2005;
Lee & Song, 2005;Yun et al., 2014). A commonly called auto inducer bioassay is used to detect AI-2 molecules and the results of such a bioassay depends on Vibrio harveyi and its ability to produce bioluminescence (Federle, 2009).
Increasing concern regarding C. difficile associated disease in elderly populations and its occurrence in communities emphasized the need for additional remedies and in this regard some studies have focussed on drugs that can target luxS genes and manage gene expression through cell-to-cell communication (Carter et al., 2005; Lee & Song, 2005). Vibrio harveyi is a bioluminescent reporter strain that can be used to study and detect AI-2 production of C. difficile (Carter et al., 2005; Michael et al., 1999). They showed that V. harveyi BB170 has the ability to bioluminesce in response to AI-2 and at low population (106 – 107CFU/ml).
Addition of AI-2 from other bacteria to V. harveyi BB170, induces bioluminescence in V. harveyi and can be detected at specific wavelengths (ex. 600nm).
In 2005, a study demonstrated that AI-2 molecules produced by C. difficile 630, induced bioluminescence in a V. harveyi AI-2 reporter strain (Carter et al., 2005). They reported that AI-2 was at its maximal concentration at late-log phase and it reduced quickly when bacteria entered stationary phase. Other studies presented similar results from Salmonella Typhimurium (Surette et al., 1999) and Porphyromonas gingivalis (Chung et al., 2001; Scheres et al., 2015). Another study reported that AI-2 molecules in C. difficile induces tcdA, tcdB and tcdE transcription in the early log phase, although C. diffcile toxins (TcdA and TcdB) are produced in stationary phase (Lee and Song 2005). They suggested that C. difficile can have a ‘LuxS/AI-2 dependent signaling mechanism’ which perhaps contributes to the pathogenicity of this
27 bacterium. Molecular analysis of more than 100 clinical isolates showed that luxS gene and LuxS protein in all strains of C. difficile are conserved. Environmental factors such as pH, temperature and osmolarity can affect AI-2 production in C. difficile (Lee and Song 2005). In conclusion, to inactivate the virulence of bacteria, a potential therapeutic target is to block bacterial cell-to cell communication (Hirakawa & Tomita,
2013). A study on its quorum signalling system shows that toxin production of C. difficile is regulated through a quorum signalling system, a gene regulator smaller than 1000 Da that is detectable in stool samples of CDI patients (Darkoh et al., 2015). These researchers identified a new quorum-signaling system termed Agr system that its Agr-like signaling peptide (TI signal) regulates C. difficile toxin gene transcription and toxin production.
2.3.3 C. difficile Motility
Motility of microorganisms is one of the significant physiological factors in their viability and survival.
Bacterial flagella are filamentous organelles that drive cell movement. However, in some bacterial species, it can be an invading factor or a means to colonize at the right sites in the gastrointestinal tract. For instance,
Campylobacter jejuni flagella are responsible for internalization of the organism (Grant et al., 1993; Guerry,
2007), but Helicobacter pylori flagella are in charge of adherence and colonization (Bergonzelli et al.,
2006; Dunne et al., 2014; Eaton et al., 1996).
As for many other pathogens, adherence of C. difficile to the colonic mucus is the first stage for invading large intestine cells. It is necessary for C. difficile to find a way through the layer of mucus to reach the epithelial cells. C. difficile bypass the defensive barrier to reach the mucosal layer by means of proteases and flagella and then binds to the enterocytes by its adhesion molecules (Karjalainen et al., 1994; Tasteyre et al., 2001). Therefore, motility and chemotaxis are involved in this process. A study conducted by
Tasteyre and colleagues (2000) has shown that not all pathogenic strains of C. difficile show a flagellated phenotype by electron microscopy but the flagellin gene could be found in the genome of both flagellated and non-flagellated strains of C. difficile. They also concluded that fliC and fliD genes encode the respective
28 flagellin monomers and these genes can be used as a basis for rapid detection of pathogenic strains of C. difficile (Tasteyre et al., 2000). Non-flagellated but toxigenic C. difficile strains showed 10 times less adherence to the mouse cecum compared to the flagellated strains of the same serogroup; indicating the importance of flagella for adherence (Tasteyre, et al., 2001a; Tasteyre et al., 2001b).
It is noteworthy that there are several factors including carbon source, temperature and viscosity of the medium, salt, and sugar can induce motility of the pathogens (Girón, 1995; Holt et al., 1994;Lemoine et al., 2013). Some facts that could explain interspecies -diversity of C. difficile strains include: 1) the presence of a capsule, an anti-pathogenic factor (Davies & Borriello, 1990), 2) proteolytic enzymes that produce suitable protein from available sources for them to metabolize nutrients and to infiltrate the mucus (Poilane et al., 1998; Seddon & Borriello, 1992), 3) the ability to adhere to mucus and subsequently invade epithelial cells (Eveillard et al., 1993; Karjalainen et al., 1994), 4) the presence of fimbriae whose role is unknown
(Borriello, 1998a), 5) the presence of flagella, which have a role in colonizing ceccal tissue (Taystere et al.
2001).
There are some controversies in relationship to flagella and pathogenicity of C. difficile. For instance, an in vitro study conducted by Tasteyre and others (2000) reported that flagella of C. difficile are not responsible for adherence of bacteria because the presence of antibodies raised against the flagellin protein could not inhibit adherence of bacteria to epithelial cells (Tasteyre et al., 2000). However, the same authors in another study in 2001 conducted an in vivo study and reported that FliC and FliD proteins were involved in the attachment of C. difficile cells to the mucus layer in the intestine so they concluded that flagellated strains are more capable to attach to the cecal wall in vivo (Tasteyre et al., 2001).
2.3.4 C. difficile Biofilm Formation
Biofilm formation has been an essential part of the life-cycle in different organisms from prokaryotes to
Archaea for billions of years. The evidence proving the existence of biofilms has been found in fossils
29 found in deep sea rocks in Australia dating from 3.2 billion years ago. Biofilm formation of pathogenic bacteria are linked to the escalation of antibiotic resistance and frequent chronic infection.
Consequently, biofilms can have a detrimental impact on both public health and economics. Many pathogenic bacteria form biofilms to provide a physical barrier to external stresses, such as desiccation, antimicrobials and biocides, as well as to evade the host immune response, preserve nutrients and retain mechanical integrity (Costerton & Lewandowski, 1995; Flemming & Wingender, 2010; O’Toole et al.,
2000). During biofilm formation, excessive biotic mechanisms involve cell motility (Klausen et al., 2003),
EPS production (Flemming & Wingender, 2010; Flemming et al., 2007), metabolic and other phenotypic differentiation (Stoodley et al., 2002), and cell-to-cell interactions such as quorum sensing (Cady et al.,
2012; Diggle et al., 2007; Kong et al., 2006).
Biofilm is a self-produced extracellular matrix made of protein, DNA and polysaccharide. Biofilms are a common form of bacterial existence in many environments and are a community of aggregated micro- colonies attached together and adhered to a natural or artificial surface (Costerton & Lewandowski, 1995) through a matrix of EPS (extracellular polymeric substance). In other words, biofilms are poly-microbial aggregates linked to each other and to surfaces (Costerton, 1999). It requires a complicated multifactorial process; and protects bacteria from stresses like antibacterial agents and host immune responses. EPS is a complex mixture of extracellular polymer macromolecules originating from bacterial cells in the biofilm
(Flemming & Wingender, 2010) and it is the slime matrix that maintains stability of biofilm and their adherence to surfaces (Hall-Stoodley et al., 2012). When single bacteria are deprived of nutrients
(starvation), they turn from planktonic single cells into aggregates (Reichenbach, 1985). There is evidence that many bacteria are disposed to aggregate in different environments (Burmølle et al., 2010; Monier &
Lindow, 2003; Stoodley et al., 2002) and it is understood that aggregation is a preliminary step before bacteria form biofilm (Melaugh et al., 2015).
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Important pathogens such as Staphylococcus aureus and Pseudomonas aeruginosa are able to produce biofilm in order to colonize their particular niche to protect themselves from environmental stresses.
Researchers in recent years showed the ability of C. difficile to form biofilm in vitro, an important ability that needs to be studied and is essential in their colonization and persistence in the host (Dapa et al., 2013;
Dawson et al., 2012).
There is insufficient information about biofilm formation by anaerobic gut pathogens but some studies show that biofilms enhance persistence and virulence in C. difficile. Di and In (2014) demonstrated that C. difficile R20291, an outbreak isolated strain (027 ribotype), accumulates more biofilm in comparison with
C. difficile 630, a clinical strain. Their study also shows that Cwp84 and LuxS are compulsory factors, called virulence associated proteins for more biofilm formation by C. difficile. This same study also showed that biofilm associated cells of C. difficile are more resistant to vancomycin, a commonly used antibiotic to treat CDI and they suggested that biofilm formation by C. difficile is a complicated process and can be the pivotal for persistence of clostridia in the host. In addition, biofilm formation can play a vital role in recurrent infection in individuals after antibiotic treatment, because of their ability to resist the antibacterial effect and, hence, recolonize their host (Dawson et al., 2012).
Cwp84, flagella and LuxS are some of the factors that affect biofilm formation in C. difficile and LuxS protein, which is a putative quorum sensing regulator, is required for maximal biofilm formation by C. difficile (Dapa & Unnikrishnan, 2013). Therefore, quorum sensing plays an important role in C. difficile biofilm formation in early and late stationary phases when bacteria start to produce biofilm components, which is modulated by luxS gene. C. difficile flagella also are key elements during biofilm formation and biofilm maturation by enabling the bacteria to move toward the place that is right for attachment (Dapa and
Unnikrishnan, 2013).
Dawson and others (2012) demonstrated a new characteristic of C. difficile. They found that C. difficile form aggregates prior to biofilm formation on abiotic surfaces. They monitored development of biofilm in
C. difficile cultures for 6 days after bacterial cells sediment (aggregate) in the first 16-24 hours. SEM
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(scanning electron microscopy) pictures showed that C. difficile make their biofilms after they attach to a surface and link to each other within the EPS matrix. On the other hand, C. difficile, known as the most important cause of nosocomial infections, has shown biofilm formation during long-term colonization in mice (Dawson et al., 2012). C. difficile makes aggregates after cell surface protein CwpV is continuously expressed (Reynolds et al., 2011) or when C-di-GMP, a biofilm inducer, is present (Purcell et al., 2012).
2.4 CDI Management, Treatment and Control
Clostridium difficile infection is a burden on the health care system because it increases costs for extra care for at risk patients, recurrent infections are common, and requires longer hospital stays and increased medication. According to Darkoh and colleagues (2015), the cost of C. difficile infection is estimated at between 1 to 4.8 billion U.S. dollars annually, and the total cost for the United States, Canada and United
Kingdom has been estimated at about 7 billion US dollars each year (Dubberke & Olsen, 2012; Kyne et al.,
2001; O’Brien et al., 2007; Wilkins & Lyerly, 2003). To date, the cost of CDI has not been precisely reported because most of the studies are from acute care hospitals. In the event that number of cases has risen in long term care facilities (LTCF), they have underestimated the real cost of CDI including hospitals and communities. A CDI surveillance in Ohio, USA, reported that the number of CDI cases diagnosed in
LTCF are more than hospitals and this may be because most of the individuals with recurrent CDI are cured outside of hospitals (Dubberke & Olsen, 2012; European CDC, 2013).
Several environmental factors impacting expression of colonization genes, such as presence of different antibiotics, have been investigated in vitro. Studies showed that clindamycin and ampicillin increased expression of the genes encoding adhesion and also increased adherence of C. difficile bacteria to mammalian cells. Among the adherence genes, cwp84 was the most upregulated gene (Denève et al.,
2008b). They also investigated the influence of ampicillin, clindamycin, ofloxacin, moxifloxacin and kanamycin on the expression of adherence genes and the protease gene cwp84 by real-time PCR, and they confirmed that exposing C. difficile to ampicillin and clindamycin increased expression of genes
32 responsible for colonization of bacterium to the Caco-2 cells. A research group at Sanofi Pasteur are developing a new vaccine called “C. diffense” and it is in phase III in its clinical study stage to investigate how their vaccine can help to protect at risk patients for CDI (http://www.sanofipasteur.ca/node/40701).
Three essential factors for C. difficile infection to occur are: 1. antimicrobial intake that ruins normal and protective microbiota in the colon, 2. ingestion of a toxigenic strain of C. difficile or its presence in the colon 3. The presence of host factors including immune system response to toxins (serum IgG antibody).
When a watery stool test is positive or pseudomembranes are present in the stool, that patient is CDI positive. To manage the infection effectively, the next necessary step is to detect the infection very quickly before any further medication. Production of toxins generates inflammation by attracting more white cells from the blood to the mucosal lining. Cells start to die and intestine movement and permeability increase.
These factors will lead to diarrhea and colitis (Durai, 2007). Clinical features of C. difficile infection in symptomatic patients are: diarrhea 100%, fever 28%, abdominal pain in 22%, leukocytosis (> 10,000/mm3) in 50% (Gerding et al., 1986), and thickened colonic mucosa in more severe situations. Toxic megacolon and colonic perforations are life threatening consequences of infection (Pépin et al., 2004; Stieglbauer et al., 1995).
Even though this life threatening pathogen causes immense problems for the health care system, there still is not enough information about its toxin production system and factors that regulate its virulence genes to onset the disease. Another study conducted by Darkoh and colleagues (2015) revealed new information about a mechanism to regulate C. difficile toxin production, which can be used for the rapid detection of
CDI by assay of a small (<1000kDa) thiolactone molecule that mediates quorum sensing signal regulators and is detectable in the patient’s stool. Previously Bartlett (2008) showed that the main factor for CDI is associated with antibiotic use, but another study showed that using antibiotics is not the primary risk factor since 61% of patients diagnosed with CDI did not have any antibiotic intake at least 90 days before the infection occurred (Kutty et al., 2010).
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Among the wide range of antibiotics, some of them are more problematic in causing CDI, including fluoroquinolones, clindamycin and beta-lactam agents (Hedge et al., 2008; Kuijper et al., 2007).
Other risk factors are internal tube feeding, anti-ulcer medication, and proton pump inhibitors (PPI) (Dial et al., 2005). PPI is a popularly prescribed medicine in the USA for long term treatment of acid-related disorder (Fashner & Gitu, 2013) and it is overused in Europe (Ramirez et al., 2010). A study in 2008 did not confirm any risk by PPI in patients (Gerding et al., 2008b) but other researchers reported that PPI is a risk factor for CDI (Kwok et al., 2012) and another recent study concluded that PPI therapy may increase the chance of C. difficile re-infection or death, especially in male patients (Santos-Schaller et al., 2016).
Debilitated individuals with lower immunity and fewer antibodies to neutralize C. difficile toxins are at higher risk for CDI (Durai, 2007). Different studies show that females are more susceptible to C. difficile infection than males particularly between the age of 21 to 50 years and over the age of 70 years in the community, although the ratio of female to male patients are lower in hospitals (Aronsson et al., 1985;
Smith et al., 2015).
Sodium hypochlorite (bleach) diluted 1:10 is effective to deactivate the C. difficile spores. Vaporized hydrogen peroxide also is an effective way to disinfect surfaces from the spores. On the contrary, ammonium based products and alcohol should not be used for disinfection against C. difficile because they may stimulate germination. Use of gloves, aprons, snd hand washing with soap are effective methods to reduce the spread of spores in the environment and lower the risk of infection (Hedge et al., 2008).
2.5 Probiotics and Inhibition of CDI
2.5.1 Human Microbiota
It is known that the human body and microbes form a complex ecosystem. The gastrointestinal tract is the biggest portion of the body that interfaces with the environment. Microorganisms that exist in the gastrointestinal tract of humans and animals are called intestinal microbiota. This community of live microorganisms consists of hundreds of bacterial species as well as viruses, fungi and parasites (Guo et al.,
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2015; Howarth & Wang, 2013). These microorganisms belong to four phyla: Actinobacteria, Bacteroidetes,
Firmicutes and Verrucomicrobia and many different types of bacteria exist within each phyla, however, bacteria are the dominant ecosystem and they exist in every healthy individual (Rajilić-Stojanović et al.,
2007).
The human microbiota varies between individuals and changes during a person’s life depending on their age and age related events, including different diets, lifestyles and their immune system. Although the mechanism of action for gut microbiota is not yet completely known or understood, some of their impacts on human health include: mediating immune responses like allergies and autoimmune diseases, digesting foods, absorbing nutrients, preventing inflammation, and guarding the host from pathogen colonization by competing for food and epithelial surfaces (Backhed, 2005; Oozeer et al., 2005). Some members of the gut microbiota, such as lactic acid bacteria especially probiotics, have attracted a great deal of attention as beneficial microorganisms that can be important in medicine, nutrition, and in the market. Their importance is because they can compete for nutrients, produce bacteriocins or neutralize toxins, and inhibit pathogens like C. difficile and many more that will be explained later in this chapter (Rupnik, 2015).
2.5.2 Lactic Acid Bacteria and Probiotic Strains
Lactic acid bacteria are a group of bacteria that have diverse phylogenetic classifications but similar metabolic activities. They are a family of microorganisms that are wide-spread in the environment, in humans and animals, and are traditionally defined as heterogeneous bacteria. They are capable of fermenting various foods, and producing lactic acid as a main by-product. Lactic acid bacteria (LAB) are
Gram-positive, non-sporulating rods and cocci, microaerophilic, acid-tolerant, usually catalase negative that exist in diverse environments. They live in different parts of the human body such as the gastrointestinal tract, mouth, respiratory tract and vaginal cavity. In addition to human and animal bodies, they exist in numerous niches in the environment including plants, dairy products, vegetables, and meat products
(Klaenhammer et al., 2002; Klaenhammer et al., 2005).
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Taxonomically, LAB are categorized in two phyla; Firmicutes and Actinobacteria. Some of the important genera of LAB in the Firmicutes phylum, are Enterococcus, Lactobacillus, Lactococcus, Leuconostoc,
Pediococcus, Streptococcus and Weissella, that are all belong to the Lactobacillale order and are low-GC content organisms (31–49 %). These bacteria have a high demand for nutrients such as amino acids, peptides, carbohydrates, vitamins, fatty acids, salts and nucleic acid derivatives (Kandler & Weiss, 1986).
They cannot get energy from respiration, therefore amino acids are essential factors for them to grow, and they have developed a sophisticated proteolytic system to break down protein sources to their required nutrients (Pessione, 2012). The Bifidobacterium genus is a member of the Actinobacteria phylum, with a high-GC content (58–61 %) and is also a lactic acid bacterium (Klaenhammer, 2005; Pfeiler &
Klaenhammer, 2007; Schleifer & Ludwig, 1995).
Metabolically, LAB are categorized in two groups according to their metabolic pathways: homofermentative and heterofermentative. Homofermentative species of LAB produce about 85% lactic acid as an end product, whereas heterofermentative species produce CO2 and ethanol or acetate in addition to lactic acid. Approximately 50% of their final product carbon is in the form of lactate. LAB have significant industrial applications in fermentation processes and in the human diet. Their existence in the gastrointestinal tract has brought some of them recognition as probiotics due to their human origin and their beneficial impact on human health (Klaenhammer et al., 2008; Makarova et al., 2006).
The term “probiotic” is derived from the combination of two words in Latin (pro = in favor of) and Greek
(bios = life). Initially, this term was used to describe the microorganism’s growth supporting materials
(Lilley & Stillwell, 1965) but later researchers refined the meaning of the term to identify “a live microbial feed supplement which beneficially affects the host animal by improving its intestinal balance” (Fuller,
1989). Since the 1990s considerable research has been done on probiotic bacteria and the mechanisms responsible for their beneficial impact, such as their effect on the gut and the immune system. At the same
36 time, industry and academia supported many projects devoted to probiotics and functional foods. On the other hand, there were no specific definitions and regulations for probiotics regarding consumer safety. In
2001 and 2002, an evaluation process was undertaken by FAO and WHO to provide international guidelines on the practical and safety aspects of probiotics. Results of meetings of international experts were documented and the term “probiotic” was defined as “a live microorganism that, when administered in adequate amounts, confers a health benefit on the host” (FAO/WHO 2001).
Probiotics are often bacteria and also yeasts, isolated from human microbiota depending on their special characteristics (Perricone et al., 2015). Some of the important criteria for selecting probiotics include phenotype and genotype stability (plasmid stability); metabolism of proteins and carbohydrates; bile uptake; adhesion to the intestinal epithelial cells; secretion of antimicrobial substances; antibiotic resistance patterns; ability to inhibit spoilage organisms and pathogens; their immunogenicity; and their survival in different food products or in the host’s gastrointestinal tract (Tuomola et al., 2001).
Probiotics and pathogens both need to survive in the harsh conditions of the stomach, and tolerate acid and bile. The difference is the way they establish themselves in their host’s intestine. Some pathogens like E. coli and Salmonella invade the epithelial cells or some like C. difficile toxico-infect their host and cause infection. But probiotics have a mutual interaction with their host, which is a health-promoting mechanism benefitting both the host and the probiotic (Medzhitov, 2007). Mechanisms of action of probiotics are complex and different between strains, they cannot be generalized for all of them. Some of their known benefits include: healing intestinal bowel disease (IBD), curing infectious diarrhea, (Hedin et al., 2007), preventing colorectal cancer (Rafter et al., 2007) and healing irritable bowel syndrome (Camilleri, 2006).
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2.5.3 Characteristics of probiotic Lactobacilli
The most common probiotic bacteria are Bifidobacteria and Lactobacilli. Bifidobacterium belongs to the family Bifidobacteriaceae, order Bifidobacteriales, phylum Actinobacteria (Biavati & Mattarelli, 2006).
Bifidobacteria mostly are obligate anaerobes but some of their species such as B. psychraerophilum, B. scardovii and B. tsurumiense, can tolerate O2 and grow under aerobic conditions (Okamoto et al., 2008).
Lactobacilli have a long history of use in fermentation of foods from animal sources (milk and meat) or plants (vegetables and olives). Lactobacilli are lactic acid bacteria and mainly produce lactic acid from carbohydrate metabolism. They live throughout the GIT, mostly in the proximal small intestine, which is a niche full of nutrients for them to grow and survive (Ng et al., 2009).
Members of the genus Lactobacillus are large heterogeneous Gram-positive, non-sporulating and anaerobic bacteria (Claesson et al., 2007). Taxonomically they belong to phylum Firmicutes, class Bacilli, order
Lactobacillales, family Lactobacillaceae. They are factidious bacteria which need a rich medium with
“carbohydrate, amino acids, peptides, fatty acids esters, salts, nucleic acid derivatives and vitamins” for growth (Kandler and Weiss, 1986). Among the human fecal microbiota, Lactobacilli comprise only 0.01% to 0.06% of total microbial flora and are found mostly in the small intestine and colon (Maukonen et al.,
2008; Neville et al., 2012). Some of the species are permanently inhabit the gastrointestinal tract (GIT) including L. gasseri, L. reuteri, L. crispatus, L. salivarius, and L. ruminis species (Walter 2005). There are other species of Lactobacilli such as L. acidophilus, L. fermentum, L. casei, L. rhamnosus, L. johnsonii, L. plantarum, L. brevis, L. delbrueckii, L. curvatus, and L. sakei that can also be found in the GIT but they are not at a stable level (Heilig et al., 2002; Walter, 2005).
Various studies have been conducted, especially in the last two decades, on many different Lactobacillus strains to investigate their benefit to human well-being, namely probiotic effects. There are controversial results from negative results (Claesson et al., 2007; Lebeer, 2008; Ligaarden et al., 2010) and positive and
38 supportive findings that show some Lactobacillus strains actually improve human health (Brouwer et al.,
2006; Folster-Holst et al., 2006; Isolauri et al., 2000; Kalliomäki et al., 2001; Kalliomäki et al., 2003;
Viljanen et al., 2005; Wells et al., 2011).
Clinical effectiveness of probiotics have been supported by a number of meta-analyses and it is generally agreed that probiotics have promising health benefits in humans. Some specific species of Lactobacillus are effective in prevention of acute infectious diarrhea and antibiotic infectious diarrhea (Sazawal et al.,
2006). Some studies also revealed that certain Lactobacilli effectively reduce the recurrence of C. difficile associated diarrhea (Pillai & Nelson, 2008). Other research demonstrated that specific Lactobacilli can exert health benefits for treating and preventing inflammatory bowel disease (IBD) (Prantera & Scribano,
2002), colorectal cancer (Rafter et al., 2007), and irritable bowel syndrome (Camilleri, 2006). Other research groups suggested that some Lactobacillus strains can prevent and treat urogenital disease and bacterial vaginosis in women (Falagas et al., 2007), atopic disease and food hypersensitivity (Boyle &
Tang, 2006), and dental caries (Meurman & Stamatova, 2007).
The link between probiotic application and health benefits is based on their mechanism of action categorized in two main factors (Figure 2.5). One is called “adaptation factor”, which is related to their adaptation to their new environment including: stress resistance, adapted metabolism and adherence.
Another contribution to their mode of action, termed “probiotic factors”, is directly related to one or all of the following three categories: (i) anti-pathogenic activity and maintaining a stable and constant internal environment for the microbial flora, (ii) improving epithelial barrier function, (iii) adjusting immune responses.
It is noteworthy to consider that the mode of action of probiotics is strain-specific and depends on the host response (Lebeer et al., 2008) and that not all the strains have a beneficial effect on their host (Sengupta et al., 2013). Strains of the same species of lactobacilli can vary in their impact on the host; this variation can
39 be related to their cell surface structure and their response to their environment or the production of different compounds, such as adhesion factors, or the health of their host and its interaction with those beneficial bacteria.
Figure 2.5. Mechanistic view of probiotic actions by lactobacilli, illustrates adaptation factors and probiotic factors of probiotic lactobacilli. Adaptation factors that outlines the elements of surviving these bacteria in their human host whereas probiotic factors pointed to the health promoting factors including pathogen inhibition, rebuilding of the microbial balance, enhancement of epithelial barrier function, and immunomodulatory effects via interactions with immune cells such as dendric cells (DCs). Adapted from Lebeer et al., 2008.
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2.5.3.1 Lactobacillus Adhesion
Adhesion of probiotic bacteria to the mucus and epithelial cells is one of the main reasons why
Lactobacillus compete with pathogens for receptor binding, and colonization ( Howarth & Wang, 2013).
Adherence of probiotic strains of Lactobacillus depends on their host’s large cell surface proteins and mucus-binding proteins that have regions compatible with binding domains of proteins secreted from those
Lactobacillus. Adhesion also increases interaction between the probiotic and the host intestine (Van Tassell
& Miller, 2011). Different surface binding proteins have been identified in different probiotic strains that are important factors for colonization of the gut (Lukic et al., 2012; González-Rodríguez et al., 2012; Van
Tassell & Miller, 2011). In addition, the impact of probiotic adhesion to the epithelial cells is important for the balance of cell death and cell proliferation and this feature of probiotics has potential for further research for cancer treatment (Howarth & Wang, 2013).
2.5.3.2 Tight junctions
The intestinal barrier is controlled by various tight junction proteins, which are located at the “sub-apical aspect of the lateral membrane” and which physically connect the cells responsible for intestinal permeability (Zhou et al., 2010; Putaala et al., 2012; Rao & Samak, 2013). Barrier dysfunction occurs when diseases such as inflammatory bowel disease increase permeability due to interruption of certain tight junction proteins in the gut cells (Corridoni et al., 2012; Zhou et al., 2010). Studies showed that probiotic bacteria are able to suppress expression of proteins that increase intestinal cell permeability by modifying regulation of the tight junction proteins (Corridoni et al., 2012). Probiotics also have the potential to maintain cell integrity through maintaining the balance between epithelial cell death and reproduction
(Corridoni et al., 2012; Tsirtsikos et al., 2012; Zhou et al., 2010).
Probiotics may improve cell proliferation and they are able to control cell apoptosis by their cytoprotective activities (Howarth & Wang, 2013). According to the literature, more than two thirds of the immune system
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“as gut-associated lymphoid tissue” is located within the intestinal tract (Pessione, 2012). Therefore, the association between intestinal microbiota and host immunity plays a very important role in protecting the host during stressful times or against pathogenic invasion (Honda & Littman, 2012; Manichanh et al., 2012;
Pessione, 2012). Probiotics can help control the antigen-specific antibodies to enhance the immune responses of the host (Yang et al., 2008).
2.5.3.3 Cell Surface Factors
The cell wall of Lactobacillus species is composed of several macromolecules that vary between species and strains and that determines specific properties of every strain or species such as adaptation to different conditions or interactions with host immune receptors of the epithelial cells. Gram-positives such as lactobacilli have a thick layer of peptidoglycan. As Figure 2.6 illustrates, they possess a thick multilayer of peptidoglycan that is furnished with teichoic acids, exopolysaccharides and cell surface proteins (Delcour et al., 1999). Lactobacillus acidophilus and some other lactobacilli cell walls are covered by a superimposed surface layer (S-layer) protein or other secreted proteins (Avall-Jaaskelainen & Palva, 2005).
There are slight differences between glycan chains in different bacteria, but there are remarkable differences between proteins and peptides that manifest different influences of LAB with their environment.
Exopolysaccharides (EPS) can be attached to the cell wall or released to their environments. Lactobacilli cell walls have different surface proteins, usually large proteins that can be linked to the cell wall by different mechanisms (Navarre & Schneewind, 1999). Proteins that are released from the cell wall will interact with their surroundings or can reach the cell wall by electrostatic interaction. S-layer proteins are small and very basic proteins of 40 to 60 kDa with a stable tertiary structure and are noncovalently attached to the cell wall (Lebeer et al., 2008).
S-layer proteins and sortase-dependent proteins (SDPs) which are well studied in lactobacilli cell wall structure play a crucial role for the organism’s interaction with host cells (Lebeer, 2008). SDPs adhere to
42 the host cells through different structures such as N-terminal signal peptide and C-terminal motif (LPxTG)
(Navarre and Schneewind, 1994). Other surface proteins also can play a role in their adhesion. Complete genome analysis of L. acidophilus for presence of adhesion encoding protein showed that a fibronectin- binding protein (FbpA), a mucin-binding protein (Mub), and a surface layer protein (slpA) all are involved in adherence of L. acidophilus to Caco-2 cells in vitro with the adhesion being regulated by different factors
(Buck et al., 2005; Lebeer et al., 2008).
Figure. 2.6. Cell surface composition of lactobacilli. Lactobacilli cell walls are a complex of different macromolecules that creates specific properties in each strain including adapting to different environments and interacting with epithelial cells and host immune receptors. Cell wall structure is composed of a thick multilayered peptidogly which is decorated with WTA, LTA, EPS, SDPs, S-layer proteins and other secreted proteins. WTA (Wall teichoic acids), LTA (lipoteichoic acids), EPS (exopolysaccharide), PG (peptidoglycan), SDPs (Sortase-dependent proteins). Picture has been taken from Lebeer et al 2008, “Genes and Molecules of Lactobacilli Supporting Probiotic Action”.
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2.5.3.4 Probiotic Proteolytic System
Widespread studies have been conducted on the proteolytic systems of lactic acid bacteria. Studies on
Lactobacillus helveticus and Leoconostoc lactis show that LAB can produce external proteases, which play an important role in their proteolytic system (Christensen et al., 1999). This significant feature of LAB provides essential amino acids for bacterial growth. Proteinases, along with peptidases and their specific transportation systems, break down the proteins and enable optimum growth conditions for the bacteria in protein rich media such as milk (Fira et al., 2001; Gobbetti et al., 2002).
As stated earlier, LAB use amino acids as their nutritional source and for their energy metabolism.
Therefore, they have a complex proteolytic and peptidolytic system to break down proteins and peptides to smaller peptides and amino acids. In their proteolytic system, they possess three components: 1) extracellular proteases to produce oligopeptides, 2) membrane transport proteins, catalyzing oligopeptides and tri-peptides and 3) intracellular peptidases to produce amino acids from peptides. Autolysis of probiotic cells releases these components into their environment. These proteolytic enzymes are able to produce bioactive peptides from milk proteins or other protein sources (Pessione, 2012).
2.5.4 Probiotic Safety and FDA Regulations
According to FAO/WHO documents, the assessment of “human safety” is based on the available information from the long history of using lactobacilli as probiotics (‘long history of safe use in food’) without having any established risk to healthy people. There is no record of pathogenicity or virulence of lactobacilli, Bifidobacterium or lactococci. However, use of bacteria in pathological conditions as ‘bio therapeutics’ have to be assessed for their new condition of application (FAO/WHO 2001). Probiotics are
“generally-regarded-as-safe” (GRAS), however there have been rare cases where probiotics have caused infection in immunocompromised patients or in patients with severe health conditions (Besselink et al.,
2008; Liong, 2008). For instance, the probiotic Saccharomyces boulardii which has been used to treat
44 patients with severe CDI or recurrent CDI in hospital, sometimes causes undesirable results in immunocompromised patients (Venugopalan et al., 2010).
2.5.5 Bioactive Peptides Mechanisms of Action
Recent research in food science and nutrition has shown that food can provide special functions and properties in addition to being a source of energy for the body. Food can benefit us through improving our health as it contains bioactive molecules such as phenolic acids, flavonoids, vitamins, omega-3 fatty acids, proteins and peptides (Carrasco-Castilla et al., 2012). Peptides are an important group of bioactive components, defined as specific protein fragments that supply amino acids and nitrogen. Depending on their amino acid sequences, peptides can improve health of the cardiovascular, digestive, immune, and nervous systems (Kitts & Weiler, 2003; Korhonen & Pihlanto, 2006). Sequences of bioactive peptides can be between 2- 20 amino acids but some peptides can have more than 20 amino acids (Saito et al., 2000).
Therefore, bioactive peptides are classified in three groups: small peptides with less than 7 amino acids, medium peptides with 7-25 amino acids, and large peptides that contain more than 25 amino acids. These biomolecules are accessible from food protein sources and there can be many of them in a single food.
These peptides are available in some foods called Functional Foods or Nutraceuticals. Dairy products, especially milk, are the most investigated sources of bioactive peptides. Bioactive peptides are novel and potential dietary ingredients capable of promoting health but which may also be sometimes detrimental for human health. Some of the bioactivities linked to bioactive peptides include anti-carcinogenic, antioxidant, antihypertensive, mineral-binding and stress-relieving properties (Korhonen & Pihlanto, 2006). Bioactive peptides can be produced through fermentation, hydrolysis of proteins by selected microorganisms mainly bacteria, or by proteolytic enzymes or a combination of bacteria and enzymes (Korhonen & Pihlanto, 2006).
Microbial hydrolysis of proteins that are degraded to small size peptides are interesting field of study.
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Emerging infections and increasing of drug-resistant bacterial pathogens universally, requires new strategies and novel approaches to overcome this important problem (National Research Council (US)
Committee on New Directions in the Study of Antimicrobial Therapeutics, 2006; Alirol et al., 2011). To date, the scientific community and the food industry is very interested in isolating and identifying bioactive peptides that have anti-virulent characteristics. Anti-virulence refers to a mechanism that disarms pathogens by preventing production of virulence factors and its strategy in vivo would be similar to vaccines. There are advantages of using bioactive peptides as an alternative therapy for pathogen control, since bioactive peptides maintain the commensal microbiota without disturbing them and stimulate the immune system of the host as a result of their reaction with the body’s systems. Furthermore, using bioactive peptides will not lead to the development of antibacterial resistance because these small peptides do not kill or interfere with viability of the pathogens but rather down-regulate their virulence genes (Sharma, 2014).
L. acidophilus is a homofermentative Lactobacillus that produces lactic acid from lactose or any fermentable sugar. L. acidophilus La-5 is a well-known probiotic bacterium and is commercially available from Chr Hansen Company. It has been approved as a food supplement because of its acknowledged health benefits, and it is well documented for its potential to balance the normal microbiota of the intestine. It has been used for more than 30 years without any identified side effects (Bunterngsook et al., 2012). Its impact on Clostridium difficile colitis, antibiotic-associated diarrhea, inflammatory bowel disease in combination with Bifidobacteria animalis and Lactococcus lactis has been studied (Aragon et al., 2010).
Most of the research has investigated the role of probiotic bacteria on human health in the last decade, focusing on clinical applications of probiotics to treat intestinal disorders (Demirel et al., 2013; Hunter et al., 2012; Yan et al., 2011). In contrast, the effectiveness of the bioactive molecules secreted by probiotics is not well documented (Paszti-Gere et al., 2012; Yan et al., 2007). Advanced research in this area will discover therapeutic benefits of these bioactive molecules. Administration of live bacteria has some risks for immunocompromised patients with a weaker immune system or patients that have gastrointestinal issues. Research on bioactive molecules will also provide a better understanding of the mechanisms of
46 probiotic actions (Howarth, 2010; Howarth & Wang 2013). In conclusion, finding the composition of the bioactive molecules produced by probiotics is challenging and there are many difficulties such as dependency of these derivatives upon several factors including bacterial species, bacterial strains and their media components. The effectiveness of bioactive peptides depends on dose and frequency of administration among microbiota architecture.
Studies on probiotic L. acidophilus La-5 have demonstrated that the bioactive fractions derived from L. acidophilus La-5 down-regulate virulence genes in Gram-negative bacteria. For instance, secreted peptides of L. acidophilus were investigated for their impact on virulence genes of enterohemorrhagic Escherichia coli O157:H7 (EHEC O157). They showed that cell free media that had supported the growth of L. acidophilus reduced pathogenicity of E. coli EHEC, by interrupting quorum sensing; leading to significant down-regulation of a number of virulence genes of EHEC O157:H7 (Medellin-Peña & Griffiths, 2009;
Medellin-Peña et al., 2007; Zeinhom et al., 2012). Tissue culture assays showed that these bioactive molecules decreased EHEC O157 attachment and a mouse model revealed that L. acidophilus La-5 produces bioactive molecules that suppress severe infection (Medellin-Pena et al., 2009). It was reported that EHEC O157 infection decreased significantly after mice were fed yogurt containing La-5 cell free fractions containing bioactive peptides produced by L. acidophilus, which led to reduced colonization and attachment of EHEC on the epithelial cells of mice (Zeinhom et al., 2012).
An in vitro study by Peng, (2014) showed the effects of bioactive components secreted by probiotics such as L. acidophilus La-5 and L. helveticus LH-2 on improving the integrity of epithelial cells against S.
Typhimurium infection. Another study by Sharma, (2014), using the same strains of probiotic against S.
Typhimurium showed that these bioactive molecules were able to down-regulate expression of genes affiliated to AI-2 activity and invasion of this pathogen, two important factors for their interaction with their host. Additionally, Bayoumi and Griffiths (2010) reported that L. acidophilus bioactive peptides down regulated virulence genes located in pathogenicity islands 1 and 2 of S. Typhimurium.
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Furthermore, Yun and colleagues (2014) investigated the impact of L. acidophilus GP1B on quorum sensing and virulence activity of C. difficile in vitro and in vivo. Their research revealed that components of the probiotic culture reduced quorum sensing of C. difficile and inhibited its growth in the GIT of mice.
The study also showed that toxin-related virulence genes in C. difficile were down regulated as a result of those bioactive molecules. Also, (Chapman et al., 2012) conducted research to investigate the growth inhibition of C. difficile, Escherichia coli and S. Typhimurium using a variety of probiotic strains and they concluded that inhibition of pathogen growth by probiotics was strain dependent and that among different strains of lactobacilli, L. acidophilus showed a significant inhibition zone in agar spot tests.
Tessema (2015) screened several LAB strains for production of bioactive peptides against S. Typhimurium.
He also examined different culture media including whey protein, de Man, Rogosa, and Sharpe (MRS), brain heart infusion (BHI), and showed that L. acidophilus La-5 grown in the presence of whey protein concentrate produced the bioactive agent leading to the highest suppression of virulence genes of
Salmonella. Results have shown that inhibition of infection is mediated through down-regulation of genes associated with the Type 3 secretion system or homologous genes in Gram-negative bacteria using whey protein concentrate.
2.5.6 Secreted Proteins and Bioactive Peptides Sources
The beneficial effect of probiotic bacteria is directly related to what they produce in their environment.
Therefore, isolation and characterizing of those secreted molecules is a focus of current and future research.
Bioactive molecules can be produced by probiotic bacteria in their culture media or they can be derived from their culture fraction. They are proteinaceous isolates with low molecular weight and very few amino acids (Griffiths et al., 2009).
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2.5.6.1 Secreted Proteins
Studies have shown that some of the advantages of probiotics are gained through their direct contact with the intestinal epithelial cells; they act as an anti-inflammatory agent, breakdown fatty acids, or aid in vitamin
B synthesis. On the other hand, there have been studies that showed some other advantages that do not need direct cell contact. For instance, recently, proteins were identified which are secreted by L. rhamnosus GG
(p40 and p75) from their culture medium and it has been claimed that they affect signaling pathways involved in the promotion of intestinal epithelial homeostasis (Yan et al., 2002). These proteins stimulated cell growth, and repressed apoptosis of epithelial cells (Yan & Polk, 2002), supported epithelium tight junctions, and protected them as a barrier against cytokines (Martínez-Augustin, , 2014).
Other studies reported that cell free culture of other probiotics such as L. acidophilus and L. plantarum contained unidentified and large proteinaceous soluble factors that stabilized tight junctions and induced mucins in intestinal epithelial cells (Otte & Podolsky, 2004). Further research on cell free culture medium of various probiotic lactobacilli will help us learn more about the unknown factors that may induce defense mechanisms of the intestine epithelial cells.
2.5.6.2 Soluble Peptides
Additionally, studies reported that soluble, heat and acid stable small peptides have been identified with low-molecular-weight in the cell free culture of probiotics. These small peptides are able to induce heat shock proteins to protect epithelial cells (Tao, 2006) and to help the cells to stay intact and keep the epithelial barrier undisturbed (Fujiya et al., 2007). Membrane separation and ultra-filtration techniques were used to enrich and separate peptides with a specific molecular weight range (Korhonen and Pihlanto,
2006). Other molecules such as probiotic DNA, cell surface proteins, cell wall peptidoglycan, and exopolysaccharide (EPS) that can be found in the growth media of lactobacilli, all these secreted molecules from lactobacilli perform as protective factors for the intestinal epithelial cells.
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CHAPTER 3
3. BIOACTIVE MOLECULES PRODUCED BY PROBIOTIC LACTOBACILLUS
ACIDOPHILUS LA-5 SHOWING INHIBITORY EFFECTS TOWARD CLOSTRIDIUM
DIFFICILE VIRULENCE GENES
Abstract
Lactobacillus acidophilus has unique qualities which differentiate it from other probiotic bacteria. L. acidophilus produces antivirulence substances named bioactive molecules, compounds inhibitory to the pathogenicity of various pathogens including Clostridium difficile. In the present study, after comparing the effects of L. acidophilus La-5 with some other probiotic strains on quorum sensing of a hypervirulent strain of C. difficile ribotype 027 in vitro, we aimed to assess its impact on several virulence genes of three highly infectious strains of C. difficile (ribotype 027, 078 and 001). Quorum sensing changes were determined by quantitative measurement of light emission using a bioluminescent reporter strain, Vibrio harveyi BB170, and bioactive molecules produced a significant reduction (P<0.05) in autoinducer-2 in all of the C. difficile strains. The effect of bioactive molecules on the expression of several virulence genes were assessed with reverse transcription real-time quantitative PCR (qPCR) and it was shown that bioactive molecules from L. acidophilus La-5 were able to down-regulate some virulence genes of C. difficile strains.
Key words: Probiotic, Lactobacillus acidophilus La-5, antivirulence, bioactive molecules, proteobiotics,
Clostridium difficile, gene expression, quorum sensing, LuxS.
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Introduction
C. difficile is a leading pathogen in hospital associated diseases and, unfortunately, C. difficile infections have risen since 2000 due to changes in physiological and biomolecular characteristics of C. difficile as a result of extensive use of broad-spectrum antibiotics (Borriello, 1998). C. difficile associated diseases range from mild to severe diarrhea to life-threatening megacolon disease associated with high mortality and morbidity (Rupnik et al., 2009). Symptoms of C. difficile infection (CDI) are mostly caused by its two exotoxins: TcdA and TcdB. Toxin genes tcdA and tcdB are located in a pathogenicity locus (PaLoc) that is a genetic element in toxigenic strains. Both toxins are glucosyltransferases that inactivate Rho, Rac and
Cdc42 proteins in the host epithelial cells. This mechanism results in disrupting cytoskeleton and disregulating cell function that leads to apoptosis (Jochim et al., 2011; Just & Gerhard, 2004). PaLoc also contains genes encoding an anti-sigma factor (tcdC) and a sigma factor (tcdR). TcdC is suggested to be an inhibitor of toxin expression (Carter et al., 2011; Dupuy et al., 2008); whereas TcdR is a positive regulator of toxin expression (Bakker et al., 2012; Mani & Dupuy, 2001; Merrigan et al., 2010; Paredes et al., 2005).
Studies indicated that non-toxigenic virulence factors including surface layer proteins (SLPs), and cell wall proteins (CWPs), their encoding genes slpA (encoding a surface layer protein) and cwp84 (encoding an 84 kDa cell wall protein – protease) are involved in the attachment and colonization process (Denève et al.,
2008). SLP proteins function as a mechanical barrier and as a virulence facilitator for harming the host. It also, along with CWPs, is involved in biofilm formation (Gerbino et al., 2015). CWP also acts as an enzyme against host proteins to facilitate pathogenicity of the bacterium (Vedantam et al., 2012). Furthermore, motility and chemotaxis are potential virulence factors. Before the colonization of the gastrointestinal tract can occur, C. difficile must penetrate the mucus layer by the help of their flagella and protease enzymes.
FliC, main subunit in flagellar structure and FliD, the flagellar cap protein are of the flagellar proteins that is heavily involved in adhesion to the epithelial cells and they play important role in C. difficile virulence mechanism (Tasteyre et al., 2001).
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Studies confirmed that bacterial virulence factors are more effective when their population is higher in their community due to quorum signalling (Chung et al., 2001; Fong et al., 2001; Ohtani et al., 2002; Zhu et al.,
2002). Bacterial communication is mediated through the production and detection of secreted signalling molecules and is often referred to as Quorum Sensing (Bassler, 2010). Quorum sensing is the mechanism by which bacterial populations coordinately control gene expression; often in response to cell population density (Fuqua et al., 1994).
Research also showed that changes in biological activities of pathogenic C. difficile may be due to modulation of its quorum signaling system (Ary, 2011; Darkoh et al., 2015; Wilson et al., 2012). One encoding gene involved in quorum sensing of Gram-positive bacteria including C. difficile is luxS gene and it is produced at its highest level when C. difficile strains are at mid-exponential phase in their growth cycle.
The LuxS enzyme which is part of the methyl cycle pathway, takes part in the production of AI-2 from S- ribosylhomocysteine (Pereira et al., 2013) and also regulates some of PaLoc genes (Lee & Song, 2005).
Inhibiting virulence gene transcription and their regulators is a key factor to prevent infection. Bioactive molecules are anti-infective agents that are able to reduce the harmful effects of enteric pathogens in mammals (Yun et al., 2014; Zeinhom et al., 2012). Bioactive molecules are peptide sequences that are released during metabolic activities of probiotic lactic acid bacteria, and hydrolysis of nitrogen sources present in the gut (Pessione and Cirrincione, 2016). These are small oligopeptides (Pessione, 2012) extracted from culture media and may account for some of the health benefits of probiotics. In this study, probiotic cell free supernatant (CFS) containing bioactive molecules (recently termed proteobiotics)
(Nordeste et al., 2017) was used to evaluate changes in expression of the C. difficile virulence genes of three toxin positive C. difficile isolates (ribotype 027, 001 and 078) when exposed to cell free supernatant obtained from L. acidophilus La-5 culture.
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Toxin production differs between strains and is influenced by the available nutrients and stresses in the cell’s environment. In this study, we have chosen three strains of toxin positive C. difficile including ribotype 027, 078, and 001. Ribotype 027 is a hypervirulent strain involved in several outbreaks (Kuijper et al., 2006; Warny et al., 2005), ribotype 078 is an animal origin toxigenic strain (Weese et al., 2010) and strain PCR-ribotype 001 was identified as a high prevalence toxigenic strain in European countries found in communities (Arvand et al., 2009; Borgmann et al., 2008; Nyc et al., 2015; Zaiß et al., 2010). We investigated he inhibitory effect of bioactive molecules from La-5 cell free supernatant against growth pattern, viability and quorum sensing signaling molecules (autoinducer-2) production of three toxigenic strains of C. difficile. Also, the regulatory activity of La-5 and fold-changes in gene expression were monitored in all three C. difficile strains ribotype 027, 001, and 078 by using real-time q-PCR.
Additionally, we investigated the probiotic characteristics of some lactic acid bacteria: growth pattern in different protein rich media as well as the growth inhibition of C. difficile by limiting their community communication. Selected lactic acid bacteria (LAB) were all of human origin or known for human use.
Then, one probiotic strain was chosen for an investigation to measure other important impacts of bioactive molecules on C. difficile virulence factors. The rationale behind this selection was based on the probiotic’s ability to 1) grow well in both selected media (modified MRS and whey protein concentrate), 2) show more inhibition of AI-2 production on C. difficile, 3 on previous research results (Tessema 2015; Sharma, 2014;
Medellin-Peña et al., 2007). The de Man, Rogosa, and Sharpe medium (MRS) ingredients were modified in order to reduce the content of sugar and L-cysteine to limit the impact of both on autoinducer signal production and toxin gene expression of C. difficile (Karlsson et al., 1999). A high protein content medium
(whey protein concentrate) with low sucrose content was prepared to test the growth performance of probiotics in comparison to the control medium, modified MRS.
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Our hypothesis was that bioactive extracts would decrease the expression level of virulence genes in all three C. difficile strains. Also, it was hypothesized that bioactive extract is effective inhibitor of mechanisms responsible for growth performance and quorum sensing of C. difficile cells.
The contribution of this research is to understand the mechanism of action of bioactive molecules that are secreted by lactobacillus acidophilus strain, on pathogenesis of C. difficile.
The main objectives of this chapter were to:
1. Measure C. difficile quorum sensing in the presence of LAB bioactive extract;
2. Investigate how La-5 bioactive extract influence C. difficile viability and growth pattern;
3. Observe the quantitative changes in expression of C. difficile virulence genes in addition of La-5 bioactive extract.
3.1. Materials and Methods:
3.1.1 Modified de Man, Rogosa, and Sharpe (m-MRS) medium
MRS (de Man, Rogosa, and Sharpe) medium is a commercial medium that is a suitable source of different proteins, sugar (glucose) and essential minerals for bacterial metabolism. The source of proteins in MRS medium are hydrolyzed proteins such as peptone from pancreatic digest of casein, beef extract and yeast extract. In the current study, we used modified MRS broth (Delcenserie et al., 2011) which has 75% less sugar (sucrose) than the original formula of MRS (20g/L). Modified MRS contains 10 g peptone from pancreatic digest of casein, 8 g beef extract, 4 g yeast extract (BD Diagnostic Systems, Sparks, MD), 5 g sucrose, 1mL Tween 80, 2 g dipotassium hydrogen phosphate, 2 g di-ammonium hydrogen citrate, 5 g sodium acetate, 0.2 g magnesium sulfate, 0.2 g L-cysteine, and 0.04 g manganese sulfate in 1 liter of distilled water. The prepared m-MRS was filter-sterilized using a 0.22 μm-pore-size filter (EMD Millipore,
Billerica, MA, USA) and PVDF sterile syringe filters (Fisher Scientific, Canada). Sterile medium was kept in a sealed container at 4oC and was used within 24 h of preparation.
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3.1.2 Chemically Modified Medium (Whey Protein medium)
Another interesting source of valuable nutrition to grow lactic acid bacteria is bovine whey (Pescuma et al.,
2011). The major constituents of bovine whey proteins include β-lactoglobulin (β-lg) and α-lactalbumin, serum albumin, lactoferrin (LF), immunoglobulin (IgG), glycomacropeptides (GMP), lactoperoxidase and other immunoglobulins, growth factors and a number of other proteins of lower concentrations (Korhonen
& Pihlanto, 2007; Korhonen & Pihlanto, 2006). Whey protein is also an interesting substrate to use as a nitrogen source for lactic acid bacteria. In this study, the whey protein isolate that was used has high amounts of protein but no sugar or fat content. To prepare the optimal medium from whey protein for probiotics, sucrose was added to the reconstituted powder before sterilization.
Reconstituted whey protein powder was prepared by adding 28 g whey protein [Whey Protein (GRAS,
Ergogenics Nutrition Ltd., Vancouver)] (5.6 g/100 mL CMM product) and 10 mL sterile sucrose solution
(0.25 g/mL) [Sucrose (S) – (Sigma Aldrich Mississauga, ON Canada)] to 500 mL sterile distilled water.
The chemically modified medium or CMM was sterilized through two step filtration. First it was filtered using Millipore 0.7 µm APFF047 (Millipore) using a vacuum. A second filtration was accomplished using
Fisher 0.45 µm low protein binding Millipore-size filter (09-719D, Fisher Scientific Canada) using a 30 mL syringe. After sterilization of CMM, to make sure that there is no contamination during the process of sterilization, each time, the sterile media was tested by spreading 100 μL of it on the MRS agar and BHI agar and incubating in an anaerobic jar for 48 h at 37oC.
3.1.3 Probiotic LAB Strains
Seven lactic acid bacteria strains were obtained from the culture collection of Canadian Research Institute for Food Safety (CRIFS) at the University of Guelph. These strains are listed in Table 3.1 and belong to four genera including Lactobacillus; Bifidobacterium; Pedoiococcus; and Lactococcus. To activate the strains, one loop full of frozen stock cultures was streaked on Lactobacillus MRS agar (BD Difco™,
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Mississauga, ON, Canada) and incubated under anaerobic conditions with a BBL GasPak system (BD
Bioscience, Mississauga, ON, Canada) for 48h at 37 °C. Then the purity of the cultures was tested by checking morphology of the single colonies, gram staining, and looking at cells under bright field microscopy (Omegon Binocular Microscope, 100× objective lens). Viability of the bacterial cells was tested by plating on lactobacillus MRS agar plates that were incubated at 37 °C for 48 h and CFU/mL (colony- forming units per mililiter) were determined and expressed as log CFU/mL. For future work, pure stock cultures of each strain were prepared by growing the strains to the late exponential phase with a colony count of 1 × 108 CFU /mL in m-MRS (Medellin-Pena et al., 2007). The stock cultures were stored at -80
°C in 30% v/v sterile glycerol (Fisher Scientific, Nepean, Ontario, Canada). The pH of the probiotic cultures was measured at the end of incubation and then CFU/mL and pH values were compared between the probiotic strains. The strains were cultured in triplicate and analyses were performed in duplicate.
Table 3.1 Lactic acid bacteria used for this study LAB Genera LAB Strain Source
Lactobacillus L. acidophilus La-5 Canadian Research Institute for Food Safety Lactobacillus L. rhamnosus GG Canadian Research & Development Centre for Probiotics, London, ON, Canada
Lactococcus Lc. lactis DSM 4366 Canadian Research Institute for Food Safety Bifidobacterium B. longum NCC2705 Canadian Research Institute for Food Safety
Bifidobacterium B. infantis ATCC 15697 Canadian Research Institute for Food Safety
Pediococcus P. pentosaceus C1337 Donated by Chr. Hansen, Denmark Pediococcus P. acidilactici C1336 Donated by Chr. Hansen, Denmark
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3.1.4 Production of LAB Cell Free Supernatant
As described in section 3.1.3, selected probiotic bacteria were activated from a frozen cryogenic vial. One colony per 10 mL of a freshly prepared m-MRS or WP was inoculated and each time one un-inoculated medium was used as a control and treated the same way as inoculated samples.
After 48 hours incubation time, the probiotic cells were separated from the growth culture by centrifugation at 15,000 ×g for 20 min at 4°C (Avanti J-20 XPI, Beckman Coulter, Canada). Remaining cells were removed by filtration through a 0.45 µm low protein-binding Millipore-size filter (EMD Millipore,
Billerica, MA, USA). To avoid the influence of pH on the results of other experiments, before filtration the supernatant of the probiotic culture was neutralized with 5N NaOH. Uninoculated m-MRS medium was treated with the same conditions as La-5 culture as an experimental control in all the experiments. Finally, cell free culture supernatants from probiotic cultures and m-MRS were lyophilized using a freeze-drying system (Unitop 600 SL, VirTis Co., Inc. Gardiner, NY, USA). Lyophilized CFS was labeled and sealed properly and kept at -80oC for further use. Each time, an appropriate amount of lyophilized powder was reconstituted with sterile deionized water to make a soloution 10× more concentrated than the original CFS.
Protein concentration of La5 CFS was measured using an ND-1000 Spectrophotometer (Thermoscientific,
Wilmington, DE) in protein A280 section according to the manufacturer’s instruction. Normally, the protein concentration of CFS was adjusted to achieve a protein content of 100 mg/mL. In order to prevent changes in the protein and peptide structure, reconstituted CF was always handled in a cooler and, after reconstitution, it was used only for a single experiment. Reconstituted powders will be termed La-5 CFS throughout this study
3.2. Bacterial Strains Pre-scanning Experiments
In a preliminary study, we focused on screening of some of the probiotic strains from the culture collection to make sure that the bacterium that we selected down-regulated C. difficile virulence genes.
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3.2.1 Bacterial Cultures and Screening for Growth
A bio-screening experiment was conducted to investigate growth pattern of selected lactic acid bacteria
(LAB) known to produce bioactive peptides, in two different media: 1. m-MRS and 2. Whey protein medium, and then bacterial growth curves were compared between the two media. Both media were prepared as described in the previous sections (3.1.1 and 3.1.2).
In this study, we screened the bacterial’s growth pattern by incubating inoculated cultures in both media under the same conditions. LAB strains were inoculated from one overnight culture (2 × 10 8 CFU/mL) into both media with a dilution ratio of 5:100. Each probiotic culture was dispensed in a volume of 100µl into
100-wells plates (Honeycomb, Fisher Scientific, USA), inside an anaerobic chamber (80% nitrogen, 10% hydrogen, 10% carbon dioxide) and then plates were sealed with clear adhesive films (Micro Amp™
Obtical Adhesive Film Cat. 4311971) to make sure that oxygen did not penetrate into the wells during incubation. Plates were transferred to the BioScreen C (Automated MicrobiologyGrowth Curve Analysis
System, MTX Lab Systems) and incubation temperature was maintained at 37oC. Optical density
(OD600nm) was measured at 60 min intervals for 48 hours. Bioscreening of probiotics was performed with two independent experiments and three technical replicates. Growth curves were obtained by plotting the
OD against the exposure time and the slope of the growth curves at their period of maximum growth velocity were determined graphically by the GraphPad Prism Software 7.0.
3.3 Inhibition of C. difficile Biological Activity by Bioactive Extract La-5 CFS
3.3.1 Bacterial Strains and Culture Conditions:
In the present study, we observed the influence of bioactive molecules secreted by the chosen probiotic strains of LAB on C. difficile viability and proliferation through monitoring growth pattern; quorum sensing through AI-2 bioassay; and expression of virulence genes through real-time q-PCR.
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C. difficile Strains. C. difficile strains selected for this study were a NAP1/ribotype 027 human clinical strain, a NAP7/ ribotype 078 porcine strain, and a ribotype 001 human clinical isolate. These three epidemic strains of C. difficile are among the most commonly reported ribotypes associated with outbreaks over the last decade. They were provided kindly by Dr. Scott Weese, Pathobiology Department, University of
Guelph, Canada. It is noticeable that C. difficile is a biosafety level 2 (BSL-2) pathogen. All the guidelines and regulations for handling BSL-2 plus were followed throughout the study. In each independent experiment, C. difficile strains were activated from frozen stock (-80oC) by streaking a loop-full stock culture on a fresh BHIS agar plate (Brain Heart Infusion agar, 0.1% sodium taurocholate) (Smith et al.,
1981) and incubated in an anaerobic chamber (80% nitrogen, 10% hydrogen, 10% carbon dioxide) at 37°C,
48 hours. Then, one colony of each strain was inoculated in BHIS broth and incubated in an anaerobic chamber at 37°C overnight. Sodium taurocholate stimulates spore germination in C. difficile selective medium (Buggy et al., 1985).
Treatment Procedure. 1mL of reconstituted La-5 CFS was added to BHIS broth in a ratio of 1:10 and pre- incubated for 4 hours to reduce the oxygen content of the medium. Then, 100 µl of overnight culture were added to treated BHIS broth in a ratio of 1:100 and stirred by pipetting gently, and samples and controls were incubated at 37 oC anaerobically for about 16-20 hours until the OD 600nm reached 0.6 - 0.7 (late exponential phase). Cells were harvested by centrifugation (9000 ×g, 10 min at 4°C).
C. difficile Culture Filtrate. Cells were harvested by centrifugation at 9000 ×g and 4oC for 10 min, and the supernatants were filtered using 0.22-mm syringe filters (EMD Millipore, Billerica, MA, USA). C. difficile filtrates were kept at -80oC for future use. Toxins are delicate proteins that can be degraded or deformed at room temperature.
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3.3.2 Impact of La- 5 Bioactive Extract on Viability and Proliferation of C. difficile
In order to see the impact of La-5 CFS treatment on viability and growth of C. difficile, optical density
(OD), or total count were assessed at different time points in the presence and absence of the CFS. Also,
AI-2 production, gene expression and toxin production in C. difficile is phase-dependent, therefore, knowing the growth pattern and population of three strains at certain time points was advantageous for the design of future experiments.
C. difficile stock cultures were activated and liquid starter cultures of C. difficile C027, C078 and C001 strains were prepared as in section 3.3.1. Next, 25 mL cultures were prepared with a dilution of 1:100 of
C. difficile culture into pre-incubated BHIS. At the same time, cultures were treated with La-5 CFS
(10mg/mL). Untreated cultures were used as a control to compare the treatment between strains and within each treatment and its control. All the cultures were incubated in an anaerobic chamber (80% nitrogen, 10% hydrogen and 10% carbon dioxide) for 48 hours. Every 3 h, 1 mL of each sample was taken from the anaerobic chamber and OD600nm was recorded manually using 1.5 mL disposable cuvettes (UV-Cuvettes,
Semi-micro, Cole-Parmer). Each sample was duplicated and the experiment was repeated three times.
3.3.3 LAB Bioactive Extract and Inhibition of C. difficile Autoinducer-2
The AI-2 bioluminescence assay was performed as described by Carter et al., (2005) and Yun et al., (2014).
Briefly, C. difficile culture filtrate of strains 027, 078 and 001 were examined for presence of the AI-2 signaling molecule using a V. harveyi reporter strain BB170. C. difficile cultures treated with CFS of six probiotic strains were prepared as explained in section 3.3.1. Briefly 100 µl of C. difficile cultures were inoculated into 10 mL of BHIS broth and incubated anaerobically at 37oC for 18-24 hours to reach an OD
600 nm of 0.5. According to other studies, C. difficile releases AI-2, mostly during exponential phase
(Darkoh et al., 2015). Therefore, cell free cultures of C. difficile were collected at the exponential phase and after removing the cells by centrifuging at 9000 ×g, 10 min, at 4 oC, the supernatant was passed through a 0.22 µm-pore size membrane filter (EMD Millipore, Billerica, MA, USA).
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The reporter strain V. harveyi BB170, was grown for 16 h at 30°C in a shaking incubator (175rpm) in AB medium (autoinducer bioassay medium) consisting of main ingredients 0.3 M NaCl, 0.05 M MgSO4, 0.2% vitamin-free casamino acids in 1 L distilled water adjusted pH to 7.5 with KOH and autoclaved before adding filter sterilized supplement components of 1M potassium phosphate (pH 7.0), 0.1 M L-arginine
Hydrochloride, 50% glycerol, 10 mg/mL riboflavin, and 1 mg/mL thiamine. Then V. harveyi culture was diluted 1:1,000 into fresh AB medium. C. difficile culture filtrate was added to a final concentration of 10%
(v/v) before dispensing 100 µl of the diluted V. harveyi BB170 to 96-well microtiter plates (Corning No.
3610, Fisher Scientific, Ottawa, Ontario, Canada). Positive control consisted of AB medium containing
10% (v/v) cell free supernatant collected from V. harveyi BB152, and blank samples included sterile AB or
BHIS media added to the base culture to a concentration of 10% (v/v). The microtiter plate was shaken at
175 rpm at 30°C in the VictorTM Multilabel counter (Wallac, PerkinElmer Life Sciences Canada,
Woodbridge, Ontario, Canada). Luminescence production was recorded every hour for 24 hours at 30 oC.
The V. harveyi cell density was measured by recording the absorbance at 600 nm at the same time as light measurement. All experiments were performed at least three times and all the samples were tested in triplicate. Maximal bioluminescence was measured between 7-9 hours after adding cell free supernatant.
The data were reported as relative light unit (RLU) per unit of absorbance (cell density; OD 600nm).
3.3.4 Real-Time Quantitative PCR and Gene Expression Analysis
3.3.4.1 Real-Time q- PCR Methodology
One of the best ways to illustrate the functional status of a specific cell is to observe its gene expression pattern. Bacterial cells express genes differently under different conditions or metabolic stages. Real-time quantitative PCR (q-PCR) presents a sensitive and powerful method to quantify nucleic acids and analysis of gene expression in bacterial pathogens. Two-step real-time q-PCR is one of the techniques that is usually performed. In this technique, the first step involves RNA extraction and reverse transcription to obtain
61 complementary DNA (cDNA) followed by the second step, PCR amplification of specific genes.
Fluorescent quantitative real-time PCR is one of the most precise techniques. SYBR Green dye generates fluorescent signals by intercalating into double stranded DNA. Normally, the expression of a target gene together with a reference gene are analyzed for normalization of the amount of the PCR template and for the calculation of the relative gene expression level of the target gene (Muller et al., 2002).
3.3.4.2 Standard curves
Standard curves are prepared by plotting the regression line of the “threshold cycle” (Ct) values against cDNA concentration. Standard curves were generated for all the genes using duplicate of all the dilutions and three biological replicates. The AB system Vii 7 PCR system analyzed Ct values and automatically generated standard curve plots, (1) slope (A slope close to − 3.3 indicates optimal, 100% PCR amplification efficiency), (2) R2 values (correlation coefficient) “a measure of the closeness of fit between the regression line and the individual CT data points of the standard reactions” (AB system manual). A value of 0.99 to
1.00 shows the best fit between the regression line and the data points. (3) The threshold cycle (CT) is the
PCR cycle number at which the fluorescence level meets the threshold. A CT value >35 indicates a low amount of target in the reaction; for CT values >35, and a higher standard deviation is expected.
3.3.4.3 RNA extraction and Real Time q-PCR
Three strains of C. difficile treated with or without La-5 CFS or alone were prepared in BHIS medium, as explained in section 3.3.1. Briefly, cultures were incubated until the OD 600nm reached between 0.5 and
0.7. Some experiments needed samples from the exponential phase (tcdC) and some samples needed stationary phase cultures (tcdA, tcdB and tcdR). Pellets were precipitated by centrifugation of 1-2 mL of culture for 3 min at 9000×g, and room temperature. The supernatant was discarded and any residual medium was removed. Pellets were re-suspended in 2 mL of RNA protect bacteria reagent (Qiagen Inc.,
Mississauga, Ontario, Canada) and incubated for 5min at room temperature (25oC). Centrifugation was
62 repeated for 3 min at 13,000 ×g (Spectrafuge-16M, Labnet International, Edison, USA). The supernatant was removed and the pellet was resuspended in 200 µl Tris-EDTA buffer (Bio-ultra, PH 8.0, Fluka, Sigma-
Aldrich Co. St. Louis, USA) containing 60 µl freshly made lysozyme (50 mg/mL; Sigma-Aldrich Co., St.
Louis, USA) and 20 µl proteinase K (Qiagen) to lyse the cells according to the procedure for Gram-positive rods. The mixture was incubated for 1 h at 37 oC with shaking at 450 rpm (Thermomixer, Eppendorf,
Humburg, Germany). RNA extraction followed with the RNA extraction procedure (Qiagen’s RNeasy mini kit, 74104) as described in the manufacturer’s instruction manual.
DNAse treatment was performed for each sample to eliminate residual DNA. RNase-free DNase enzyme
(Turbo™ DNase, 2 U/ µl, Ambion) was added to the RNA samples. The mixture was incubated for 15 min at 37oC and treated with 0.5 M molecular grade EDTA at 70 oC, 10 min, to deactivate the DNase. After that,
RNA was purified using RNeasy Mini Elute (Clean up Kit, Qiagen 74204) following the manufacturer’s instructions. Purified RNA was solubilized in 20 µl RNase free water.
The quality and concentration of extracted RNA (ng/ µl) were determined spectrophotometrically (Fleige
& Pfaffl, 2006; Sambrook, Fritsch, & Maniatis, 1989) using the ND-1000 spectrophotometer
(ThermoScientific, Wilmington, DE). A spectrophotometer measures RNA concentration and quality using the ratio of the absorbance at 260/280 nm.
In addition, agarose gel electrophoresis (Bio-Rad, Mini-Sub® Cell GT System, Bio-Rad, Mississauga, ON) was performed to confirm the quality of the total RNA. Briefly, 1 % agarose gel was prepared containing
0.5 g agarose gel (Bio-Rad, Mississauga, ON), in 50 mL TAE buffer (40mM Tris (pH 7.6), 20 mM acetic acid, 1mM EDTA). Agarose was dissolved by boiling, and 0.5 µl ethidium bromide were added after cooling down to 60 oC. The gel was cast and a comb was inserted. After 30 minutes, the gel became solid.
It was transferred to electrophoresis tank containing TAE buffer. A ratio of 6:1 of total RNA samples and
6× loading dye (Fermentas, Fisher Scientific, Ottawa, ON) was prepared, and five µl of total RNA was
63 loaded into the wells. Two µ1 of 100 bp DNA ladder were added to a well as a reference (Fermentas, Fisher
Scientific, Ottawa, ON). Electrophoresis was performed with the same buffer at 70 volts for 90 minutes at room temperature. The gel was transferred to the Gel Doc System (Bio-Rad, Mississauga, ON) and RNA bands were visualized using Quantity One software (version 4.4.0, Bio-Rad, Mississauga, ON).
Reverse Transcription PCR. After quality and quantity of total RNA was measured as it was mentioned above, the purified RNA was transformed to cDNA immediately using a cDNA reverse transcription kit
(Applied Biosystems, Burlington, Ontario, Canada). cDNA synthesis was completed according to the manufacturer’s instructions using 50 ng total RNA and 0.5 µM of each primer in a 20 µl reaction volume.
The negative control with no reverse transcriptase enzyme for each RNA sample was prepared to make sure there was no DNA contamination. Reverse transcription was performed in a Mastercycler Gradient
Thermocycler (Eppendrof, Mississauga, Ontario, Canada) using cycles of 25 oC for 10 min, 37 oC for 120 min, 85 oC for 5 min, and then a hold step at 4 oC. cDNA can be stored at 4 oC for up to 1 week or at -20 oC for a longer time.
Quantitative Real-time qPCR. A real-time q-PCR assay was carried out in order to determine gene transcription analysis. Real-time qPCRs were performed using a ViiA™ 7 Real-Time PCR System
(Applied Biosystems, Burlington, Ontario, Canada) and Power SYBR Green PCR master mix following the manufacturer’s instruction. Aliquots of cDNAs from three strains and from both treated and untreated samples resulting from a reverse transcription assay were used as templates for real-time qPCR amplification in triplicate for validation of reference genes and target genes. A non-template control and a calibrator (a relevant dilution series of standard sample) were used in every run.
The three reference genes used for normalizing C. difficile 027 and 078 target genes were rrs (16s), rpsJ and adk; whereas reference genes rrs, adk and gluD showed more stable results for C. difficile 001. All the
64 primers used in qPCR assays were synthesized by Laboratory Services, University of Guelph, using previously published sequences (Table 3.2). Since gene stability is strain dependent (Metcalf et al., 2010), selection of reference genes for different C. difficile strains needed to be assessed and the most stable reference genes between the strains were selected. Optimized thermal cycling conditions were used for real- time qPCR and were as follows: initial denaturation at 95 oC for 10 minutes, followed by 40 cycles of denaturation at 95 oC for 15 s; hybridization at 56 oC for 1 minute and melting curve analysis at 95 oC for
15 s, 60 oC for 15s and 95 oC for 15s.
The ViiA 7 analyzed the data and showed the slope and also standard curve of each gene. To make sure that the amplification product was of the expected size for each gene, electrophoresis was performed with
1% agarose gel using ethidium bromide. The analysis of gene expression data from quantitative real-time q-PCR experiments was performed using REST 2009 Software (www, Qiagen.com/REST). This software is ideal for analysis of relative gene expression, normalizes expression levels of target genes when there are more than one reference gene involve. It improves reliability of the results by using expression of multiple reference genes in different samples or groups.
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Table 3.2. List of reference genes and target genes primer set sequences
Genes Function of encoding Primers Amplicon Source protein size (bp)
rrs (16S) RNA ribosomal GGGAGACTTGAGTGCAGGAG 120 Metcalf et al., subunit GGGAGACTTGAGTGCAGGAG 2010
rpsJ 30S Ribosomal protein GATCACAAGTTTCAGGACCTG 151 Metcalf et al., S10 GTCTTAGGTGGATTAGC 2010
adk adenylate kinase GTGTATGTGATGTATGCCAAG 195 Metcalf, 2012 CCTAAGGCTGCGACAATAC gluD glutamate ATGCAGTAGGGCCAACAAA 135 Denève et al., TTCCACCTTTACCTCCACCA 2008 tcdA Toxin A production AATTCCAATACAAGCCCTGTAG 91 Bakker et al., TATCAGCCCATTGTTTTATGTATTC 2012 tcdB Toxin B production GTTCTATTTTGGTGAAGATGGTGTC 89 Bakker et al., CCTCACCATCAATAATAACTGAACC 2012
tcdC Negative regulator of GGTTCAAAATGAAAGACGACG 98 Bakker et al., toxin gene expression GCACCTCATCACCATCTTCA 2012
tcdR Sigma factor for toxin TCAAAGTAAGTCTGTTTTTGAGGAA 92 Bakker et al., gene transcription TGCTCTATTTTTAGCCTTATTAACAGC 2012
luxS Quorum sensing GTGTACTTGATGGAGTAAAGGGAGA 180 Carten et al., TTCTACATCCCATTGGAGATAAGTC 2005
fli C Flagellin ATGAGAGTTAATACAAATGTAAGTGC 130 Dingle et al., CTATCCTAATAATTGTAAAACTCC 2011 cwp84 Cell wall protein – TGGGCAACTGGTGGAAAATA 151 Denève et al., protease TAGTTGCACCTTGTGCCTCA 2008
slpA Surface layer protein- AATGATAAAGCATTTGTAGTTGGTG 126 Denève et al., adhesion TATTGGAGTAGCATCTCCATC 2008
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3.4 Statistical Analysis
Statistical analysis was performed using two way ANOVA with Tukey’s multiple comparison test for significant differences between means of different groups in GraphPad Prism Software (version 7.0, La
Jolla, CA). Differences between the means reported with 95% confidence intervals (Tukey’s multiple comparison test) were considered significant at P < 0.05.
3.5 RESULTS
Preliminary Results
3.5.1 Growth Kinetics of Lactic Acid Bacteria in Different Media
Growth curves of seven probiotic strains were obtained with the average OD 600nm value at 60 minutes intervals at 37 oC in m-MRS and WP media. Analyzed data showed slightly a higher OD 600nm for strains
L. acidophilus La-5 and L. rhamnosus compared to other strains grown in m-MRS. L. aciduphilus La-5 also had a higher OD 600nm in WP medium. The slope of all the growth curves in one medium was parallel in the exponential phase, and growth rates were similar between the bacteria. However, a large difference (P
< 0.05) in growth patterns was observed corresponding to different media. All the strains grew faster in m-
MRS medium (Figure 3.1A) with a shorter lag phase. Strains started their logarithmic growth after 2-3 h incubation and then reached stationary phase after 10-12 h. On the other hand Figure 3.1B, illustrated strains needed longer duration between 8 to 10 h to adapt to the WP medium and no bacterial growth was detected during this time. After adaptation period, unlike the growth pattern in m-MRS medium that resulted a very steep pattern for their exponential phase, the same strains in WP medium consumed longer time to grow exponentially. As a result, their growth speed was slower in their exponential phase than the m-MRS medium. Their OD 600 nm reached to 1.6 in some strain but after 48 h incubation whereas two of the strain stayed in plateau pattern from very low OD600nm (L. rhamnosus and P. acidilactici). The slope of exponential phase of L. acidophilus La-5 growth curve (based on OD 600 nm) was statistically compared between two media using Linear Regression features of Microsoft excel (Figure 3.2). The slopes of 67 exponential phase of L. acidophilus la-5 grown on two media (0.19 in m-MRS and 0.007 in WP) were significantly different (P=0.004).
In the present study, growth kinetics were monitored by the assessment of optical density, viable cell count, and pH changes in both culture media. After 48 hours of incubation, the pH range in selected probiotic cultures grown in m-MRS was 4.5 to 4.9, while these same probiotic cultures grown in WP medium exhibited a pH above 5.5 for all the strains except L. acidophilus. La-5 for which the pH was 5.2 (data not shown here). The decline in pH and acidity of the culture is a good indication of an active culture, but for our purpose, to eliminate the possibility that low pH could affect virulence gene expression, the pH was adjusted to 6.5 in all CFSs.
Analysis of growth performance, pH, and viability data confirm that growth kinetics is strain dependent
(Elfahri et al., 2014; Matar et al., 1996). The difference between the growth pattern of probiotics in each medium can be related to the medium contents and in contrast to the m-MRS that is a complex synthetic media, the whey medium is not very complex and it may not have all the essential growth factors, therefore it is time consuming for the bacteria to adapt to such an environment and start exponentially proliferating, which explains the increase in the lag phase in WP medium.
Lactic acid bacteria (LAB) and probiotics are fastidious organisms and they need plentiful growth elements in their medium. Although whey is a relatively cheap, protein rich medium to be used in mass production, it does not have so many free amino acid and small peptides available for supporting bacterial growth
(Donkor et al., 2007; Shihata & Shah, 2000). Probiotic bacteria essentially use their own proteinase and peptidase enzymes to breakdown polypeptides and large proteins to maintain their source of nitrogen (Smid et al., 1991). They need highly advanced proteolytic and glycolytic systems to provide essential growth factors (amino acids and glucose) for their growth (Kunji et al., 1996). Some probiotics are more proteolytic compared to others. In addition, the proteolytic system of probiotics has become more important since it
68 was realized that bioactive peptides are released during proteolysis by LAB (Donkor et al., 2007; Gobbetti et al., 2002; Nakamura et al., 1995). Furthermore, these bacteria must have peptidolytic activity to produce different oligopeptides with 4-18 amino acids and mostly with 4-8 amino acids which are claimed to be bioactive peptides (Kunji et al., 1996). LAB cell walls are able to transport short peptides to the cell cytoplasm for further degradation; releasing essential amino acids for growth activities. Those longer peptides that stay outside of the bacterial cells are those peptides that can be biologically active peptides
(Ramchandran & Shah, 2008).
Modification of media in this study was important due to the studies that have demonstrated 1) glucose can interfere with the LuxS system in different bacteria by catabolite repression that leads to different outcomes of quorum sensing and increasing virulence expression of the bacteria (Delcenserie et al., 2011; Bassler et al., 1993), and 2) cysteine, a sulfur containing amino acid that can be precursor of enzyme activities and is able to repress toxin production in clinical isolates of C. difficile (Karlsson et al., 2008; Karlsson et al.,
2000). Therefore, m-MRS medium has been modified by reducing L-cysteine supplementation and substituting glucose with sucrose in MRS medium. In conclusion, the composition of growth media had a great impact on growth pattern and biological activity of our selected lactic acid bacteria.
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A
2 .5
M R S c o n t r o l 2 .0 P .a c id ila c tic i
P . p e n to s a c e u s
m 1 .5 n
0 B . in fa n tis
0
6
D B .lo n g u m
1 .0 O L c .la c tis
0 .5 L . r h a m n o s u s
L .a c id o p h ilu s L a - 5
0 .0 0 1 0 2 0 3 0 4 0 5 0
In c u b a tio n tim e (h )
B 2 .0
W P C o n tro l
1 .5 P .a c id ila c tic i
P . p e n to s a c e u s
m n
0 B . in fa n tis
0 1 .0 6
B .lo n g u m
D O L c .la c tis
0 .5 L . rh a m n o s u s L .a c id o p h ilu s L a -5
0 .0 0 1 0 2 0 3 0 4 0 5 0
In c u b a tio n tim e (h )
Figure 3.1. Optical density growth curves. Seven probiotic strains in two protein rich but low sugar media grew anaerobically in the same culture condition: modified MRS at 37 oC for 48 hours (two replicate), compared to a sample from uninoculated medium as a control; (A) Modified MRS (m-MRS) and (B) whey protein media (WP) inoculated in a ratio of 2:100. Symbols, Mean; error bars, mean ± SD.
70
A 1.4
1.2 y = 0.2438x - 1.0073 R² = 0.9915 1
0.8
0.6 OD 600nm OD 0.4
0.2
0 0 2 4 6 8 10 hour(s)
0.3 B y = 0.0067x + 0.0989 0.25 R² = 0.9778
0.2
0.15 OD 600nm OD 0.1
0.05
0 0 5 10 15 20 25 30 hour(s)
Figure 3.2. Regression analysis of exponential phase of Lactobacillus acidophilus La-5. Slope of exponential phase was compared in m-MRS (A) vs. WP (B) using linear regression comparison of means.
3.5.2. Effect of Bioactive Extract on C. difficile Growth Rate
The influence (antibacterial effect) of the probiotic-extracted supernatant (La-5 CFS) on growth behaviour of three toxigenic strains of C. difficile was assessed by measuring the optical density at 600nm, at 37oC, for 24 hours at 3 hour intervals, starting at OD 0.01. Clostridium difficile treated cultures were compared with C. difficile samples without treatment (Figure 3.3). The growth pattern illustrated no significant
71 difference (P > 0.05) within the group of untreated cultures and they had parallel slopes in their exponential phase. The lag phase for control samples was about 6 h, whereas the treated group had a longer lag phase about 12-18 h (strains dependent). The linear regression analysis accomplished using excel software, showed that the slopes of three strains’ growth curve are not substantially different (P > 0.05). Since the slopes are not significantly different between strains, it is possible to calculate one slope for all the data.
Linear regression analysis for the ribotype 027 and its treated culture (Figure 3.4, A and B) revealed that the differences between the slopes are not significant (P = 0.0879) but intercepts (where regression line intersect an axis) analysis showed that there is a significant difference (P = 0.044) between the exponential phase onset of treated sample with La-5 CFS (-0.812; R2 =0.99) compare to the control 027 sample (-0.414;
R2 = 0.95).
1 . 2
C 0 2 7 0 . 9
C 0 7 8
m n
0 C 0 0 1
0 0 . 6
6
D C 0 2 7 # L a - 5 O
0 . 3 C 0 7 8 # L a - 5
C 0 0 1 # L a - 5
0 . 0 0 6 1 2 1 8 2 4 3 0 3 6 4 2 4 8
In c u b a tio n tim e (h )
Figure 3.3. Growth kinetics of C. difficile strains 027, 078 and 001. Optical density growth curves. Growth curve of C. difficile strain 027, 078 and 001 used in this study, and their treated cultures with L. acidophilus La-5 CFS. Means and standard errors from three biological replicates (N=3 repeats)
72
1.2 0.9 A 1 B y = 0.0563x - 0.8119 0.8 y = 0.0699x - 0.414 R² = 0.99 0.7 R² = 0.97 0.8 0.6 0.5 0.6 0.4 OD 600 nm 600 OD 0.4
OD 600nm OD 0.3 0.2 0.2 0.1 0 0 0 5 10 15 20 0 10 20 30 40 hour(s) hour(s)
Figure 3.4. Linear regression of two set of data from growth kinetic of ribotype 027 (A) and its treated samples (B) as monitored by measuring OD600nm.
3.5.3 Quorum Sensing
3.5.4.1 Lactic Acid Bacteria CFS Influence on C. difficile Quorum Sensing
The cell free supernatant of the six probiotic strains were tested against C. difficile ribotype 027, and the luminescence was measured. Results showed that some of the probiotic CFSs suppressed AI-2 induction of the C. difficile ribotype 027 culture when it was added at 10% v/v (Figure 3.5). The results showed different effects between the tested CFSs. C. difficile grown in the presence of 10% L. acidophilus La-5, Lc. lactis,
P. pentosaceus and L. rhamnosus CFSs showed significant suppression of AI-2 bioactivity, while B. infantis and B. longum did not cause subatantial reduction in luminescence for C. difficile ribotype 027. Moreover, m-MRS did not show any large effect on results.
3.5.4.2 L. acidophilus La-5 CFS Influence on Quorum Sensing of Three Strains of C. difficile
To support our hypothesis that bioactive molecules from La-5 impacts quorum sensing of the three strains of C. difficile, an AI-2 bioluminescence assay was performed using the three strains of C. difficile (ribotypes
027, 078 and 001). Based on the results discussed in previous sections, the probiotic strain of choice (L.
73 acidophilus La-5) was used to test the differences of autoinducer activity between these strains. Strains of
C. difficile were treated with La-5 CFS and their luminescence measured and compared with their control groups (without treatment). C. difficile culture filtrates were prepared and added to the V. harveyi reporter strain (BB170) to a concentration of 10% v/v. Interestingly, the results showed that La-5 CFS exhibited the greatest suppression of autoinducer activity in all three strains. Differences between La-5 treated CFSs and their control cultures were statistically significant (P < 0.0001) (Figure 3.6). These results will be assessed by analysis of real-time q-PCR (fold change in luxS gene expression) in the next section.
2 5 0 0 0 0 0
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0
l 7 5 s o 2 - is s m is a r 5 2 t u u t S i t 1 0 a c e s u n R d n h L a c o g a e C .l n n f M o . s a o n M u c s m l i C V il L o . . h t u B n h B B A p e r o p . id . L c a P . L
L a c tic a c id b a c te ria C F S M s
Figure 3.5. Production of AI-2 by C. difficile strain 027 in the presence of cell-free supernantants of lactic acid bacteria. C. difficile 027 grown for 24 h in BHI broth supplemented with CFSs from different probiotic strains at a concentration of 10%. C027: C. difficile culture filtrate without treatment; AB medium: Autoinducer bioassay (AB) media was used for V. harveyi BB170; results were expressed as relative light units (RLU), defined as counts per minute and adjusted to OD600 (RLU/OD600). The error bars represent standard deviation (Bar marked with * is statistically significant P < 0.05). Bars with **** means differences between this group of bars is statistically extremely significant from other bars (P < 0.0001). All determinations were made from three biological replicates of samples, each tested in triplicate.
74
0 2 7 1 5 0 0 0 0 0 C 0 7 8 * * * * C 0 0 1 * * * *
1 0 0 0 0 0 0 * * * *
0
0
6
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U L
R 5 0 0 0 0 0
0
ff ff ff i -5 i -5 i -5 .d a .d a .d a L L L C + C + C + d d d C C C
S t r a in s
Figure 3.6. Production of AI-2 signaling molecules by three strains of C. difficile treated with La- 5 CFS. C. difficile 027, 078, 001 grown for 24 h in BHI broth supplemented with La-5 CFS at a percentage of 10% and the controls (cultures without La-5 CFS). Results were expressed as relative light units (RLU), defined as counts per minute and adjusted to (RLU/OD600). The data are the means ± standard deviation. Bars with **** means differences between this group of bars is statistically extremely significant (P < 0.0001). All determinations were made from three biological replicates of samples, each tested in triplicate.
3.5.4 Gene Expression
3.5.5.1 Experimental Design and real-time q-PCR Data Analysis
Relative gene expression analysis was conducted after expression levels of reference genes were validated for the C. difficile selected strains. Primer pairs were tested for the lowest Ct values and stability. Fold changes was determined using comparative quantification algorithms. Standard curve method and fold
∆Ct target ∆Ct reference changes were determined using: Fold difference = (E target) / (E reference) , efficiency from
(-1/slope) standard curve E= 10 , ∆Ct target = Ct control - Ct sample, and ∆Ct reference = Ct control- Ct sample.
This equation has been taken from Pfaffl (2001) and any changes of 2-fold or more are considered significant.
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The aim of the real-time q-PCR and gene expression assay was to determine efficacy of La- 5 bioactive molecules to manipulate regulation, and transcription of potent C. difficile virulence genes in vitro. It was predicted that bioactive molecules will impact virulence factors of C. difficile strains. For extremely thorough investigation, different sets of experiments were designed to explore this possibility. Firstly, we investigated changes in the pathogenicity locus (PaLoc) genes in human and highly infectious animal strains
C. difficile ribotypes 027, 078 and 001, after treating them with La-5 CFS. Therefore, expression of toxin genes (tcdA and tcdB) and their regulators (tcdR and tcdC) from PaLoc were examined by reverse transcriptase q-PCR. As other studies reported, although toxin genes and their positive regulators are active in exponential phase of all the three strains, their quantity is not robust. To detect those genes and their expression changes, samples were collected from the stationary phase (Merrigan et al., 2010). Additionally, to examine the putative anti-sigma factor tcdC expression, samples were collected from the exponential phase.
RNA was extracted and templates of cDNA were prepared using the reverse transcription method. Standard curves were prepared by plotting the regression line of the Ct values against cDNA concentration and efficiency of each gene amplification was calculated (Table 3.3). Genes were normalized to three reference genes depending on the strains. Down regulation of tcdA and tcdB was measured relative to transcription of reference genes. Based on the work of Metcalf et al. (2010) and our experiments, rrs, rpsJ and adk were the most stable genes for strains 027 and 078, whereas rrs, adk and gluD were the most stable reference genes for strain 001. Statistical analysis of the fold changes was conducted using the REST© program
(Pfaffl, 2001). The software is based on the efficiency-calibrated model that uses randomization tests to find the significance level.
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Table 3.3. Slopes and efficiency of gene amplification performing RTq-PCR of three C. difficile strains.
Cd027 Cd078 Cd001
Genes Slope Efficiency Slope Efficiency Slope Efficiency 1 rrs(16s) -3.34 1.99 -3.40 1.96 -3.34 1.99 2 rpsJ -3.31 2.00 -3.17 2.07 -3.20 2.05 3 adk -3.52 1.92 -3.36 1.98 -3.03 2.13 4 gluD -3.56 1.90 -3.28 1.99 -3.45 1.99 5 slp -3.18 2.06 -3.30 2.0 -3.56 1.9 6 cwp84 -3.27 1.99 -3.12 2.09 -3.27 2.00 7 fliC -3.17 2.07 -3.55 1.91 -3.29 2.01 8 LuxS -3.25 2.03 -3.51 1.92 -3.61 1.9 9 TcdA -3.31 1.97 -3.11 2.1 -3.26 2.02 10 TcdB -3.34 1.99 -3.51 1.92 -3.24 2.02 11 TcdC -3.49 1.93 -3.56 1.9 -3.37 1.98 12 TcdR -3.45 1.99 -3.55 1.9 -3.21 2.04 Note: Standard curves were generated for all the genes using duplicate of all the dilutions and three biological replicates. Each slope represents average of three measured slopes by real Time q-PCR.
3.5.5.2 Real-Time q-PCR and Effect of La-5 CFS on C. difficile Gene Expression
L. acidophilus La-5 was selected as a potential probiotic bacterium in the current study as previous studies confirmed that L. acidophilus La-5 produces anti-virulent bioactive peptides against Salmonella, E. coli
O157:H7, and Campylobacter jejuni (Bayoumi and Griffiths 2010; Mundi et al. 2013; Tellez et al. 2011;
Tessema 2015.; Sharma 2014; Zeinhom et al. 2012). Also, a study showed that bioactive molecules produced by L. acidophilus GP1B probiotics reduced C. difficile pathogenicity (Yun et al., 2014). They showed that L. acidophilus GP1B cell extract inhibited C. difficile growth when the pH was less than 4.
They also reported that GP1B cell extract down-regulated the level of gene expression in luxS, texR, tcdA, and tcdB using real-time q-PCR. Their result was supported in a previous study by Lee & Song (2005) in which it was stated that luxS regulates toxin production in C. difficile. Also, some of our selected probiotic strains were examined in another study which showed that probiotic cell free culture media impacted specific virulence genes of Salmonella Typhimurium (Bayoumi & Griffiths 2012; Peng 2014; Sharmna
77
2014). We monitored antivirulence activity of La-5 bioactive molecules against three strains of C. difficile and expression of several important virulence genes (toxigenic and non-toxigenic genes) was assessed.
Pathogenicity Locus genes. REST© analysis showed that exposure to La-5 bioactive molecules modified the expression of tcdA and tcdB with significant down-regulation in all three strains (Figure 3.7). Expression ratio of PaLoc genes in ribotype 027 showed significant down regulation in tcdA and tcdB (-5.9 ± 0.87, and
-3.40 ± 0.41fold-changes respectively, P < 0.05) and in tcdR (-3.74 ± 0.41fold-changes, P < 0.05). In ribotype 078, tcdA and tcdB expression was down regulated substantially (-9.71 ± 0.37 and -7.58 ± 0.19 fold-changes respectively, P < 0.001), as was expression of tcdR (-3.63± 0.31 fold-changes, P< 0.05).
Additionally, ribotype 001 exhibited statistically significant down regulation of tcdA and tcdB gene expression (-5.15 ±0.32, and -2.71± 0.28 fold-changes, respectively (P < 0.05). Results also show that there is a significant up regulation of tcdC expression in ribotype 001 (6.59 ± 1.92, P < 0.05). However, La-5 bioactive molecules did not modify tcdR in ribotype 001. It also did not change the expression of the negative regulator tcdC of ribotype 027 and 078 significantly (P > 0.05).
Adhesin gene expression. Additionally, regulation of genes encoding cell wall proteins and flagella were investigated, including Cwp84, slpA and fliC and fold-change calculations showed significant down regulation of Cwp84 of all three ribotypes 027, 078, 001 (-12.15 ± 1.30, -14.48 ± 1.32, 12.34 ± 0.86 fold- changes, P < 0.05), whereas, significant up regulation of the fliC gene was observed only in ribotype 001
(23 ± 3.9 fold-changes, (P < 0.05) (Figure 3.8 A).The expression of slpA was also decreased in three strains but to a lesser extent (-1.4 ± 0.75, -2.87 ± 0.24 and -2 ± 0.7 fold-changes in 027, 078 and 001, respectively).
LuxS gene Expression. Relative expression of gene luxS of three treated C. difficile ribotypes with La-5
CFS was compared (Figure 3.8 B). Exposure of C. difficile bacteria to La-5 bioactive molecules resulted in
78 a down regulation of expression of luxS, resulting in -2.30 ± 0.25, -20.83±1.32, -5.28 ± 1.7 fold changes in ribotypes 027, 078, and 001 respectively (P< 0.05).
1 0
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-1 0
-1 5 t c d A t c d B t c d R t c d C
P a lo c g e n e s
Figure 3.7. Regulation of pathogenicity locus (Paloc) genes by bioactive extract influence. Real- time q-PCR analysis represented as fold-changes of tcdA, tcdB, tcdR, and tcdC genes expression of C. difficile ribotype 027, 078, and, 001 toxin genes and toxin regulators over treatment of La-5 CFS with respect of control samples. Data were normalized to rrs, rspJ, adk and gluD expression (reference genes). Error bars represent standard deviation of triplicate measurement of three biological samples.
79
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0 2 7 0 7 8 0 0 1
Figure 3.8. Regulation of non-toxigenic genes by La-5 bioactive extract influence. A) Adhesin genes: Cwp84, slpA, and fliC genes, B) luxS genes. Real-time q-PCR analysis represented as fold- changes of gene expression of C. difficile ribotype 027,078, and 001 virulence genes over treatment of La-5 CFS with respect of control samples. Data were normalized to rrs, rspJ, adk and gluD expression (reference genes). Error bars represent standard deviation of triplicate measurement of three biological samples.
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3.6 DISCUSSION
In the present study, growth kinetics was monitored by the assessment of optical density, viable cell count, and pH changes in both culture media. After 48 hours of incubation, the pH range in selected probiotic cultures grown in m-MRS was 4.5 to 4.9, while these same probiotic cultures grown in WP medium exhibited a pH above 5.5 for all the strains except L. acidophilus La-5 for which the pH was 5.2 (data not shown here). The decline in pH and acidity of the culture is a good indication of an active culture, but for our purpose, to eliminate the possibility that low pH could affect virulence gene expression, the pH was adjusted to 6.5 in all CFSs.
Analysis of growth performance, pH, and viability data confirm that growth kinetics is strain dependent
(Elfahri et al., 2014; Matar et al., 1996). The difference between the growth patterns of probiotics in each medium can be related to the medium contents and in contrast to the m-MRS that is a complex synthetic media, the whey medium (WP) is not very complex and it may not have all the essential growth factors.
Therefore it is time consuming for the bacteria to adapt to such an environment and start exponentially proliferating, which explains the increase in the lag phase in WP medium.
Furthermore, Lactic acid bacteria (LAB) and probiotics are fastidious organisms and they need plentiful growth elements in their medium. Although whey is a relatively cheap, protein rich medium to be used in mass production, it does not have so many free amino acid and small peptides available for supporting bacterial growth (Donkor et al., 2007; Shihata & Shah, 2000). Probiotic bacteria essentially use their own proteinase and peptidase enzymes to breakdown polypeptides and large proteins to maintain their source of nitrogen (Smid et al., 1991). They need highly advanced proteolytic and glycolytic systems to provide essential growth factors (amino acids and glucose) for their growth (Kunji et al., 1996). Some probiotics are more proteolytic compared to others and proteolytic system components that were recognized in L. acidophilus group including L. acidophilus, L. johnsonii, L. gasseri, L. bulgaricus and L. helveticus are relatively higher in quantity and variety in comparison to other LAB. For instance, cell-wall-bound
81 proteinase was only identified in few lactobacillus spp. including L. acidophilus strains (Liu et al., 2010).
In addition, the proteolytic system of probiotics has become more important since it was realized that bioactive peptides are released during proteolysis by LAB (Donkor et al. 2007; Gobbetti et al. 2002;
Nakamura et al., 1995). Also, these bacteria must have peptidolytic activity to produce different oligopeptides with 4-18 amino acids and mostly with 4-8 amino acids which are claimed to be bioactive peptides (Kunji et al., 1996). LAB cell walls are able to transport short peptides to the cell cytoplasm for further degradation; releasing essential amino acids for growth activities. Those longer peptides that stay outside of the bacterial cells are those peptides that can be biologically active peptides (Ramchandran &
Shah, 2008).
Modification of media in this study was important due to the studies that have demonstrated 1) glucose can interfere with the LuxS system in different bacteria by catabolite repression that leads to different outcomes of quorum sensing and increasing virulence expression of the bacteria (Delcenserie et al., 2011; Bassler et al., 1993), and 2) cysteine, a sulfur containing amino acid that can be precursor of enzyme activities and is able to repress toxin production in clinical isolates of C. difficile (Karlsson et al., 2000; Kalsson et al.,2008).
Therefore, m-MRS medium has been modified by reducing L-cysteine supplementation and substituting glucose with sucrose in MRS medium. In conclusion, the composition of growth media had a great impact on growth pattern and biological activity of our selected Lactic acid bacteria (LAB).
C. difficile growth curve coincubated with La-5 CFS analysis supports the hypothesis that treatment has an effect on the growth pattern of C. difficile. However, La-5 CFS does not inhibit their growth completely nor does it kill the bacteria according to the plate count results but it causes an increase in lag phase.
Influence of new condition of their growth media because of introducing them to La-5 CFS resulted to a more extended lag time compared to their growth pattern in their original growth media (BHIS without treatment which is called adaptation time with at least 6 to 12 hours delay. Observation of growth curve pattern and changes occurs during coincubation with La-5 CFS was necessary to be used in further assessments. We monitored the antivirulence activity of bioactive molecules La-5, using growth curve and
82 viability data from these preliminary experiments to assess three strains of C. difficile quorum sensing and gene expression changes.
Quorum sensing changes were determined by quantitative measurement of light emission using a bioluminescent reporter strain, Vibrio harveyi BB170, and bioactive molecules produced a significant reduction (P<0.05) in autoinducer-2 in all of the C. difficile strains. These result later will be confirmed by testing luxS gene’s transcription level in the treated C. difficile samples with La-5 CFS performing real- time q-PCR. When bacteria are under any stress such as starvation, exposure to chemical and antibacterial agents, heat, different oxygen levels, and presence of other bacteria, they change their catabolic activity through communication with other cells, which results in either greater virulence or more repression of their activities. When the density of the bacterial population reaches a certain point, they produce a sufficient concentration of signaling molecules to modify expression of other genes. Therefore, a possible mechanism of bioactive molecules is that they can prevent infection through their effect on quorum sensing or cell-to- cell signaling of C. difficile bacteria.
For three strains of C. difficile studied, the expression of toxin genes tcdA and tcdB was affected by La-5 bioactive molecules to some extent. Generally, the range of changes in expression was diverse among the three strains. Strain 078 showed more down regulation (p < 0.001) in tcdA and tcdB expression and less up regulation in tcdC expression than the two other strains (Figure 3.7). This trend was similar in fold changes of PaLoc genes in two other strains too (ribotypes 027 and 001). Transcription of tcdR showed substantial down regulation in two ribotypes 027 and 078 in the presence of bioactive molecules, while they did not down regulate this gene in ribotype 001. That could be related to the up regulation of tcdC gene in ribotype
001 as study showed that tcdC expression inhibits tcdR transcription (Dupuy et al., 2008).
Transcription of tcdC genes between treated samples and controls showed only minor up regulation in ribotypes 027 and 078, whereas, bioactive molecules produced significant up regulation in ribotype 001.
This suggests that tcdC was not affected by bioactive molecules in ribotypes 027 and 078, and it may be due to the fact that genes and regulators respond to bioactive molecules in a strain dependent manner or 83 their RNA have been extracted from different growth phase between the samples, as PaLoc genes are growth phase dependant. Interestingly, up-regulation of tcdC in ribotype 001 supressed expression of the positive regulator tcdR of the same strain that led to its significant, though slight, down regulation. In two other ribotypes 027 and 078 for which tcdR was down regulated more than 2 fold, tcdR had an opposite relationship with negative sigma factor tcdC and consequently, tcdC up regulation was insignificant.
The negative regulator TcdC is a small acidic protein which does not possess any conserved DNA-binding motif, but it is able to form dimers and in its N-terminal region has a putative transmembrane domain
(Dupuy et al., 2008). Although it is not known how tcdC works, it is suggested that it uses mechanisms similar to any anti-sigma factor to interact with sigma factors and modulate the activity of the sigma factor
(Bakker et al., 2012; Carter et al., 2011; Dupuy et al., 2008). It prevents RNA polymerase in TcdR from recognizing the tcdA and tcdB promoters and it might be the way that it represses toxin gene expression
(Dupuy et al., 2008, and Dupuy, 2007). According to the literature (Bakker et al., 2012), there are controversies over the major function of tcdC in Paloc. Some research claims tcdC to be a repressor of toxin gene regulation (Carter et al., 2011; B. Dupuy et al., 2008) while other studies (Merrigan et al. 2010; Bakker et al. 2012) suggest that tcdC might not be a repressor but a modulator of the regulation of the toxin genes.
Data analysis revealed that La-5 bioactive extract influenced transcription of quorum sensing gene luxS.
The luxS result was correlated with the AI-2 production which showed that La-5 and other probiotic strains reduced AI-2 production in C. difficile and also with the previous study (Yun et al., 2014). Another study supported direct impact of luxS in controlling the pathogenesis of C. difficile (Lee & Song 2005). Analysis of relative expression of non-toxin genes in the presence of bioactive molecules showed that ribotype 027 exhibited the lowest down regulation in its non-toxin genes, Cwp84, slpA and luxS, while ribotype 078 had the greatest down regulation of those three genes. On the contrary, toxin genes tcdA and tcdB and their positive regulator tcdR presented higher level of down regulation in ribotype 027 compared to the two other ribotypes.
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These three strains are three important nosocomial and community acquired strains and all are from the most common ribotypes isolated from clinical patients in North America and Europe. Although, ribotype
078 has been the most commonly isolated strain from swine (83% of 119) and from calves (94% of 33) in the United States (Keel et al., 2007), few strains of this ribotype have been isolated from hospitalized patients. However, in recent years, this ribotype has been found in more CDI patients possibly due to its transmission from meat to humans (Songer et al., 2007). However, according to the literature, ribotype 027 is the most frequently isolated and is the most antibiotic resistant strain (Pépin et al., 2004; Denève et al.,
2009; Tschudin-Sutter et al., 2016). Differences in gene expression observed in these three ribotypes also might be due to the nature of these bacteria and also because C. difficile is a heterogeneous bacterium (He et al., 2010).
In the process of CDI, colonization, adherence of the C. difficile cells to the intestinal mucus, and adhesion proteins which are involved in this process are important (Kyne et al., 2001; Vedantam et al., 2012). Various
C. difficile adhesins have been reported such as FliC, the flagellin protein (Tasteyre et al., 2001); SlpA, the surface layer protein (Calabi et al., 2002) and Cwp84, the cysteine protease (Bradshaw et al., 2015; Janoir
& Collignon, 2013). C. difficile possesses proteolytic activity that destroys its host intestinal tissue due to the action of its cysteine protease Cwp84 (Janoir & Collignon, 2013). Gene expression of the analysed adhesin genes revealed that Cwp84, a gene encoding a cysteine protease active in infection, was significantly down regulated in all three ribotypes in the presence of bioactive molecules.
The assessment of objective of this study, growth performance of three strains of C. difficile, autoinducer-
2 bioassay and data analysis of their toxin genes, tcdA and tcdB, in treated samples established that La-5 bioactive molecules only affected C. difficile growth slightly, without any bactericidal impact. Also, it suppressed quorum signalling of all three strains. On the other hand, it influenced their molecular behavior and resulted in a moderate to large reduction of virulence gene expression in a way that could be considered to influence virulence in vivo.
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3.7 CONCLUSION
Our study focused on the application of bioactive molecules produced by L. acidophilus La-5 and analysis confirmed it has an antivirulence impact against three highly infectious strains of C. difficile. Consequently, several pathogenicity factors, most importantly quorum sensing and toxin gene expression, were repressed in the presence of La-5 bioactive molecules. We can conclude that protein fragments that have been produced in the L. acidophilus La-5 culture showed an inhibitory effect on the pathological activity of C. difficile. Given its proven use for animal supplements, and our findings, we believe that L. acidophilus is deserving of more research to be applied in human food supplements to improve human health and more importantly, for preventing C. difficile re-infection in patients treated with antibiotics.
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CHAPTER 4
4. THE IMPACT OF La-5 PROTEOBIOTCS ON ADHERENCE AND CYTOTOXICITY OF
HIGHLY INFECTIOUS C. DIFFICILE RIBOTYPE 027
Abstract
Clostridium difficile infection is a range of toxin - mediated intestinal diseases that is often acquired in hospitals and small communities in developed countries. The main virulence factors of C. difficile are two exotoxins, toxin A and toxin B, which damage epithelial cells and manifest as colonic inflammation and mild to severe diarrhea. Inhibiting C. difficile adherence, colonization, and reducing its toxin production could substantially minimize its pathogenicity and lead to faster recovery from the disease. This study investigated the efficacy of probiotic secreted bioactive molecules from Lactobacillus acidophilus La-5, termed bioactive molecules, in decreasing C. difficile attachment and cytotoxicity in human epithelial cells in vitro. L. acidophilus La-5 cell free supernatant (CFS) was used to treat the hypervirulent C. difficile strain
027 culture with subsequent monitoring of cytotoxicity and adhesion. In addition, the effect of pretreating cell lines with La-5 CFS in protecting cells from the cytotoxicity of C. difficile culture filtrate or bacterial cell attachment was examined. La-5 CFS substantially reduced the cytotoxicity and cytopathic effect of C. difficile culture filtrate on HT-29 and Caco-2 cells. Furthermore, La-5 CFS significantly reduced attachment of the C. difficile bacterial cells on both cell lines. It was also found that pretreatment of cell lines with La-
5 CFS effectively protected cell lines from cytotoxicity and adherence of C. difficile. Our study suggests that La-5 CFS could potentially be used to prevent and cure C. difficile infection and relapses.
Keywords: Clostridium difficile, Lactobacillus acidophilus La-5, bioactive molecules, cytotoxicity, cytopathic, adhesion
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Introduction
Clostridium difficile is a Gram-positive anaerobic bacterium known to be the main cause of antibiotic associated diseases and hospital-acquired diarrhea, which mostly targets older adults and immunocompromised individuals in hospitals and communities (Leffler & Lamont, 2015). C. difficile infectious diseases are an economic burden on the healthcare system: in 2002 the United States estimated its impact at $1 billion annually (Kyne et al., 2002) with an increasing trend to $6 billion in 2016. Similarly, this bacterium has an impact on hospitalized patients, resulting in 240,000 patients with C. difficile infection
(CDI) and 46,000 deaths nationwide annually in United State (Shorr et al., 2016). C. difficile causes a range of symptoms from mild diarrhea to severe pseudomembranous colitis. Although C. difficile is present in the intestine of 70% of infants and 17% of adults, it is not biologically active when microbial flora of the host is intact and acts normally within a stable immune system (Jangi & Lamont, 2010; Ozaki et al.,
2004). Following antibiotic therapy or weakening of the immune system and disturbing normal bacterial flora of the colon, C. difficile becomes opportunistic and colonizes the large intestine followed by production of toxins and infection (Bien et al., 2013; Carding et al., 2015).
The increasing trend in CDI since the 1970s can be explained by several parameters, including development of detection methods, more frequent use of antibiotics and chemotherapies, followed by further spread of
C. difficile spores as a result of contamination in hospitals and long-term care facilities due to more patients/residents with CDI (Voth & Ballard, 2005). Another reason for the increasing rate of CDI after
2000 was related to the emerging clone of C. difficile named as ribotype 027, Toxinotype III, North America
PFGE pulsotype 1 (NAP1) and restriction endonuclease analysis BI. The strain 027/NAP1/BI became epidemic and caused more complex disease and higher fatality as well as gaining greater resistance to metronidazole (Pépin et al., 2004; Denève et al., 2009; Tschudin-Sutter et al., 2016).
CDI begins with the transformation of C. difficile spores to vegetative cells in the desirable condition presented by the large intestine of a mammalian host with a weakened immune system or after antibiotic therapy. Vegetative cells adhere to the epithelial mucosa and start colonizing this comfortable niche in the
88 absence of normal microflora. After adherence to the mucus layer, C. difficile penetrates the mucus and reaches the epithelial cells to bind to specific receptors before transfer of its toxin to the cells. Several studies on the physiopathological mechanism of C. difficile virulence factors have been done in recent years that show adhesion of C. difficile to the human or animal intestinal mucosa is an important step in its pathogenicity and could be a therapeutic target for developing alternative treatments other than antibiotic therapy to cure CDI and recurrent CDI (Eveillard et al., 1993; Naaber et al. 1996; Tasteyre et al., 2001;
Janoir, 2016).
Following C. difficile adhesion and colonization of the intestine, it starts to produce toxins A and toxin B.
The C. difficile toxins TcdA and TcdB are the main virulence factors of CDI that have various effects on mammalian cells, such as cytopathic and cytotoxic effects. Receptor binding is essential for toxins to be carried into the cytoplasm. In the cell cytoplasm, toxins act as glucosyltransferases, which disrupt cell-cell junctions and inactivate Rho, Rac and Cdc-42 GTPase, which are associated with F-actin regulation, and consequently leads to increased epithelial permeability and luminal fluid accumulation associated with C. difficile associated disease (CDAD) (Lyras et al., 2009; Nusrat et al., 2001). Cytopathic effects of the toxins on the cell lines result in drastic morphological changes, including shrinking and rounding of cells caused by damage to the tight junction structures and functions (Chen et al., 2002; McCormack et al., 2013). In addition to cytopathic effects, both toxins have cytotoxic effects on the intoxicated cells that cause cell death. Studies showed that cultured epithelial cells exposed to both C. difficile toxins exhibited all the features of apoptosis (Fiorentini et al., 1998; Mahida et al., 1996) or necrosis (Voth & Ballard, 2005).
Due to the existence of highly infectious strains of C. difficile (hypervirulent strains) (Yakob et al., 2015;
Pfaffl 2001; Rupnik. et al., 2009), C. difficile infection is occurring in communities which are traditionally considered low-risk (Carter et al., 2012). Although CDI can be treated by antibiotics, there are still high rates of recurrence. In this regard, researchers are looking for therapeutic alternatives that prevent or treat
CDI recurrences. Various systematic reviews suggest that probiotic bacteria show promise to solve the issue
89 of antibiotic associated disease, especially recurrences (Rex, 2017; Johnson et al., 2012; Hickson, 2011;
Kathleen, 2010; Isakow et al., 2007).
The bioactive molecules produced by probiotic lactic acid bacteria (LAB) are believed to be able to attach to the mucosal surface and act as a barrier to prevent internalization of pathogens, including C. difficile
(Sánchez, et al., 2010). In addition, it is known that proteins released by probiotics in the gut lumen may inhibit the adhesion or colonization of enteropathogens (Sánchez et al., 2008). Therefore the current study will determine whether bioactive molecules secreted by L. acidophilus La-5 in their culture supernatant are able to mitigate the effect of C. difficile toxins in cell culture. For in vitro quantification, we used cytotoxicity and adhesion assays. Two cell lines were used in this study: HT-29 and Caco-2 cell lines. Both are mammalian epithelial cells and they are characterized by their growth rate, junction strength and mucus production. HT-29 cells are more sensitive to the C. difficile toxins particularly to the TcdA. Caco-2 cells have tighter junctions, slower proliferation speed, and less mucus production than HT-29 cells.
Previous studies on probiotics revealed that bioactive molecules produced by probiotics such as L. acidophilus and Lactobacillus rhamnosus impacted the virulence of pathogenic bacteria including
Salmonella, E. coli O157:H7, C. difficile and Campylobacter jejuni (Bayoumi & Griffiths 2010; Mundi et al., 2013; Tellez et al., 2011; Tessema, 2015.; Sharma, 2014; Peng, 2014; Yun et al., 2014; Zeinhom et al.,
2012). Following those studies, our study seeks to contribute more knowledge about the efficacy of these bioactive molecules (proteobiotics) on virulence factors of a hypervirulent strain of C. difficile, including toxin production and adhesion. From our previous study (Chapter 3) we concluded that bioactive molecules have the potential to decrease C. difficile pathogenicity by down-regulating virulence gene expression and interfering with quorum sensing. In the current study our main objective was to investigate how proteobiotic molecules produced by L. acidophilus La-5 impact toxin production and adhesion of the C. difficile bacteria on human epithelial cells. We also investigated whether the presence of La-5 with C. difficile in epithelial cell culture changes the level of damage that is caused by C. difficile.
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4.1 Materials and Methods
4.1.1 Bacterial strains and bioactive peptides preparation
Lactobacillus acidophilus La-5 culture was prepared by inoculating freshly activated bacteria in modified de Mann Rogosa Sharp (m-MRS) and whey protein (WP) media, at 37oC for 48 hours in an anaerobic environment. Cell free supernatant (La-5 CFS) was prepared by centrifugation, filtration, and freeze drying, as described in Sections 3.1.1 to 3.1.4 of this thesis. An appropriate quantity (100mg/mL) of lyophilized
La-5 CFS was reconstituted and kept on ice for instant use.
C. difficile Ribotype 027 culture was prepared in BHIS, (Brain Heart Infusion supplemented with 0.1% sodium taurocholate) as previously described in section 3.3.1. In summary, for the toxicity assay, bacterial strains were inoculated into 10 mL of reduced BHIS broth and incubated anaerobically at 37oC for 18-24 h. Cell density was standardized for all the samples including treated culture with La-5 CFS and C. difficile culture without any treatment. Throughout the assay, one treated sample containing MRS CFS as a negative control has been used. C. difficile culture filtrate was prepared as described in section 3.3.1.
4.1.2 Cell Cultures
4.1.2.1 Selected Cell lines
The carcinogenic human epithelial cell lines, HT-29 and Caco-2 cells, were obtained from the culture collection of the Canadian Research Institute for Food Safety (CRIFS, University of Guelph, ON, Canada).
HT-29 Cells (18-30 passages) were cultured in Dulbecco’s modified Eagle medium (DMEM, High glucose,
Life Technologies, Burlington, ON, Canada) supplemented with 1% (v/v) 10,000 U/mL penicillin– streptomycin (Life Technologies, Burlington, ON, Canada), 10% heat-inactivated fetal bovine serum (HI
FBS, Life Technologies, Burlington, ON, Canada) and 1% L-glutamine (Life Technologies, Burlington,
ON, Canada).
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Caco-2 cells (22-34 passages) were cultured in DMEM supplemented with 1% (v/v) 10,000 U/mL penicillin–streptomycin, 10% heat-inactivated fetal bovine serum, and 1% L-glutamine (1%) HEPES (Life
Technologies, Burlington, ON, Canada) and 1mM non-essential amino acid (Life Technologies,
Burlington, ON, Canada). Cells were kept in a humid atmosphere of 5% CO2 at 37°C in an incubator
(Forma™ Series II3110 Water-Jacketed CO2 Incubators, Thermo Fisher Scientific, Mississauga, ON,
Canada), and grown as a monolayer in 25 or 75 cm2 flasks (Corning, NY, USA). The medium was changed every two days and cells were passed (90% confluence) using 0.25% trypsin-EDTA reagent (Life
Technologies, Burlington, ON, Canada).
4.1.2.2 Maintenance of Cell Culture a. Thawing HT-29 cells. One vial of each cell line (HT-29 and Caco-2) was taken from a -80℃ freezer and put on ice immediately. Both cell lines were grown the same way using different growth media and supplements. Caco-2 cells took longer to differentiate and become confluent than HT-29 cells. Frozen cells in vials were thawed in a 37℃ water bath by gently shaking the vial until only a piece of ice was left in the vial. Cells were quickly added to a centrifuge tube with 9 mL of complete DMEM (supplemented with 1%
(v/v) 10,000 U/mL penicillin–streptomycin, 10% heat-inactivated fetal bovine serum and 1% L-glutamine) and centrifuged (AllegraTM 21R Centrifuge, Beckman CoulterTM, Mississauga, ON, Canada) at 700× g for 7 mins at room temperature. All the medium was removed and 1 mL of fresh growth medium was added to suspend the pellet and mixed thoroughly by pipetting. One mL of the cell suspension was added to a T-
25 cm2 flask, with 7~10 mL of growth medium. If a lot of cells were attached to the bottom of the flask, the medium needed to be changed after 24 hours. Cells were checked under the microscope every day to make sure they were growing well. The medium was changed every two days using a 10 mL pipette; 7 mL of complete DMEM were added to the flask and cells were checked again under the microscope.
92 b. Passing the cells. Cells were taken from the incubator and checked under the microscope for contamination and the cell number. In order to pass the cells, they must be in their exponential growth phase; otherwise, it needs to be diluted. The required cell concentration for the experiment was 1 × 10 6 cells. The overflow was removed using the 10 mL pipette and cells were washed with 7 mL phosphate buffer saline (PBS) by gently moving the flask to wash everything inside. The PBS was removed from the flask. One mL of trypsin-EDTA 0.25% reagent (Life Technologies, Burlington, ON, Canada) was added to the cell culture in the flask and was left in the incubator for 5 mins. Cells were checked under the microscope to see if they were floating, otherwise the incubation period was extended. Three mL of a complete medium
(DMEM containing all the required supplements) were added and mixed by pipetting up & down at least 6 times to maintain cell homogeneity. A cell suspension (100 ~ 400 μl) was added to 7 mL complete DMEM and incubated at 37 °C in an atmosphere of 5 % CO2. c. Cryogenic Preservation of Cells. In order to preserve the younger passages of the cells for the culture collection, cryogenic preservation of HT-29 and Caco-2 cells was carried out. Cells were harvested in the mid-exponential phase of growth with viability higher than 90%, and were then centrifuged and re- suspended in a freezing medium to a concentration of approximately 1×106 cells per mL. The freezing medium contained 80% of the fresh complete DMEM; 10% of the phosphate buffer saline (PBS) in order to improve post-freezing survival and recovery of the cells; and 10% filtered dimethyl sulphoxide (DMSO)
- a cryo-protective agent. Aliquots of the prepared cell suspension were poured into labeled 1mL cryovials, immediately placed in a -20˚C freezer overnight, and then transferred to -80 ˚C for 24 h subsequent to which they were transferred to -150 ˚C for long term storage.
Live cell counting was carried out by adding trypan blue to the 10% v/v diluted cells (taking 50 μl of the cells and 450 μl of the trypan blue for each tube) and checking the cells under the light microscope using a hemocytometer. The trypan blue exclusion test is a rapid method to assess cell viability in a cell suspension
(Strober 2001). It is simple and inexpensive. The dye exclusion test is based on the ability of viable cells to be impermeable to trypan blue, naphthalene black, erythrosine, and other dyes. When membrane integrity
93 of the cells is compromised, there is uptake of the dye into the cells so that viable cells, which are unstained, appear clear with a refractile ring around them and non-viable cells appear dark blue with no refractile ring around them.
4.1.2.3 SRB Assay
The sulforhodamine B (SRB) assay is applied widely in research and in large scale screening of inhibitory compounds and drug toxicity on cell lines. This is a colorimetric assay that indirectly measures cell viability by assessing the ability of SRB dye to bind to protein in trichloroacetic acid (TCA) fixed cells (Vichai &
Kirtikara, 2006). SRB is a negatively charged aminoxanthene dye with a bright pink color that electrostatistically binds to basic amino acid residues in medium acidic conditions and disassociates in weak acidic conditions (Skehan et al., 1990). There is a correlation between cell mass and the amount of the dye taken up by cells. The greater the number of cells, the more intense the color will be and the more absorption there will be at the end. Therefore, the rate of proliferation is measured indirectly by the intensity of the color of the SRB dye (pink) under mild acidic conditions. SRB has a colorimetric end point, and it is a stable, cost effective and nondestructive assay with sensitivity to measure cytotoxicity of drugs. In the SRB assay, mammalian cell lines are grown in 96 well culture plates and fixed by trichloroacetic acid (TCA).
To measure the intensity of the color in each well, which is related to the amount of protein content of the cells, 10 mM Tris buffer (pH 10.5) solubilizes the protein-bound dye and the OD of the plate is measured at 570 nm in a microplate reader. SRB assay will be explained with more details in the next sections 4.1.3.1 and 4.1.3.2.
푀푒푎푛 푂퐷 푠푎푚푝푙푒 − 푀푒푎푛 푂퐷 푐표푛푡푟표푙 % 표푓 푐표푛푡푟표푙 푐푒푙푙 푔푟표푤푡ℎ = × 10 푀푒푎푛 푂퐷 푛푒푔푎푡𝑖푣푒 푐표푛푡푟표푙 − 푀푒푎푛 푂퐷 푑푎푦 0
94
4.1.3 C. difficile Cytotoxicity Assay
The cytotoxicity assay determines how actively one or more mammalian cells grow in the presence or absence of different test substances (Houghton et al., 2007). It is known that both C. difficile toxins, TcdA and TcdB, are cytocidal for epithelial cells (Kelly et al., 1994; Voth and Ballard, 2005). A cytotoxicity assay determines if the compound is cytocidal (kills the cells) or is cytostatic (stops cells growing or dividing). To determine the protective effect of L. acidophilus La-5 extracted bioactive peptides
(proteobiotics) against C. difficile toxicity, a number of cell culture treatments were designed using the SRB assay. First the non-toxic dose of La-5 CFS and its control media were determined. Then to select the last dilution of C. difficile culture filtrate that yielded complete cytopathic effect on HT-29 cells, cytotoxicity of different dilutions of C. difficile culture filtrate needed to be measured.
4.1.3.1 Non-Toxic Concentration Assay of La-5 CFS
First, the non-toxic concentration of CFSs of La-5 and control was determined using HT-29 cells. Cell cultures were co-incubated with different concentrations (1-6 % v/v) of reconstituted La-5 CFS and m-
MRS (modified de Man, Rogosa, and Sharpe) CFS, then the viability was measured by SRB assay. Three
4 96-well plates were seeded by 2 × 10 HT-29 cells per well and incubated for 24h at 37°C in a 5% CO2 incubator. One plate was fixed with 50 μl of 50% (w/v) cold Trichloroacetic Acid (TCA, Sigma,
Markham, ON, Canada) after cells attached to the well and used as a day 0 sample in our calculation. In one plate, cells were treated with different concentrations of La-5 or MRS (1-6%) CFS. On another plate with the same number of HT-29 cells, the medium was replaced with the same fresh medium to use as a control. Plates were incubated for another 24 h at 37°C. After removing the La-5 CFS, cells were fixed with
50 μl of 50% (w/v) cold trichloroacetic Acid. The TCA was removed and 0.4% (w/v) SRB (Sigma,
Markham, ON, Canada) in 1% acetic acid (Glacial Acetic Acid, Fisher Scientific, Canada) was used to stain the cells for 30 min. A hundred µl per well of acetic acid (1%) was used to remove the unbound dye then
100 µl per well of 10 mM Tris-base (Tris-hydroxymethyl-aminomethane, Sigma, Markham, ON, Canada)
95 was added to dissolve the protein-bound dye. The developed color was quantified by spectrophotometry at
570 nm using a microplate reader (Synergy H5, BioTek, Winooski, VT, USA) and results calculated as percentage using control wells to represent 100% growth under constant condition without any treatment.
4.1.3.2 Cytotoxicity of C. difficile Culture Filtrate
The in vitro cytotoxicity of hypervirulent C. difficile Ribotype 027 toxin was assayed using a monolayer culture of HT-29 cells maintained in DMEM supplemented with 10% fetal bovine serum in 96 well microtitration plates. The C. difficile culture filtrate was harvested at the late stationary phase (48 h) when both toxins have been already produced in the medium (Dupuy et al., 2008). C. difficile cells were separated from supernatant by centrifugation at 12000 ×g, 10 min, at 4oC. The supernatant was filtered through a
0.22-µm syringe filters (EMD Millipore, Billerica, MA, USA) and kept at -20oC until needed. The HT-29 cells (grown in DMEM containing supplements) were seeded in two 96 well culture plates (2×104 cells/
o well). Cells were grown at 37 C in a 5% CO2 incubator for 24 h to form a monolayer. After attachment of the cells and formation of a monolayer, one plate was treated with different concentrations of C. difficile culture filtrate (CF) using serial two-fold dilutions of C. difficile CF. Treated cell culture with C. difficile
o CF and control were incubated for 24 h at 37 C and 5% CO2. After incubation, a SRB assay was performed as described in previously (section 4.1.3.1), to quantify the cytotoxicity of the C. difficile CF.
4.1.3.3 La-5 CFS Influence on Cytotoxicity of C. difficile 027
Untreated and treated C. difficile cultures both with final concentrations of 10 mg/mL of reconstituted La-
5 CFS were incubated anaerobically for 48h at 37 oC. Culture filtrates were harvested after 48 h as described in section 4.1.3.2. Two-fold dilution of C. difficile CF was prepared for cytotoxicity assay. The SRB assay was performed on La-5 CFS treated and control sample of C. difficile 027 CF.
96
Alternatively, HT-29 cell monolayer pretreated with reconstituted La-5 CFS to a final concentration of 3% of treatment in 100 µl complete DMEM and it was used for cytotoxicity assay. 100 µl of C. difficile CF was added to the pretreated cell monolayers to have a two-fold diluted CF, and plates were incubated for
24 h, at 37oC, 5% CO2 and results were compared with another cytotoxicity assay which used cocultured
C. difficile with La-5 adding to the non-treated cell monolayers.
Neutralization Assay. A neutralization assay was carried out to ensure that cytotoxic activity on cell line was the outcome of the activity of toxin A and toxin B, using neutralizing antibodies. Specific antibodies for toxin A and toxin B (monoclonal VHHs antibodies) were kindly provided by Jamshid Tanha and Greg
Hussack from NRC-CNRC (National Research Council Canada; Ottawa, Canada). VHH antibodies from llama are the variable domains found in heavy-chain IgGs and only 15 kDa in size. The antibodies for TcdA were designated A20.1 and A5.1 and the antibody for TcdB was B39. Culture filtrates of different C. difficile samples were incubated with toxin A and toxin B specific antibodies one hour before inoculation onto HT-29 monolayers in 96 well plates that contained assay reagents. The final concentration of VHHs in each assay was 1,000 nM (Hussack et al., 2011). A negative control was included to compare HT-29 cells without C. difficile CF. A positive control consisted of HT-29 cells exposed to C. difficile CF.
Morphological changes of monolayers were observed using an inverted light microscope and images were taken by the CCD camera (Leica, Q Imaging Retica 1300i, Burnaby, BC, Canada). Undisturbed monolayer cells were observed after the neutralization of C. difficile toxins by their antibodies.
4.1.4 Visualization of C. difficile Cytopathic Effect on Cell Monolayer
To investigate the inhibitory potential of La-5 active compounds on C. difficile toxins, the cytopathic effects of C. difficile CFS containing toxins was carried out by observing rounded cells under the microscope
(Yang et al., 2008). HT-29 and Caco-2 cells were seeded in glass bottom plates (1×10 4 cell/mL) and
o incubated at 37 C, 5% CO2 for 48 h to become more than 50% confluent. C. difficile CF were derived from co-cultured cells with 10 % v/v La-5 CFS and WP La-5 CFS and their control media, to late stationary
97 phase as described before (section 4.1.3.2). These were diluted by adding DMEM in 50/50 ratio to have a
2 fold diluted C. difficile CF. This was added to the HT-29 and Caco-2 cell monolayer and incubated for
o 24 h at 37 C, 5% CO2. After 24 h incubation, the cell monolayers were observed under an inverted microscope and images were taken by the CCD camera (Q Imaging Retica 1300i, Burnaby, BC, Canada).
Experiments were conducted in duplicate and repeated three times.
The effectiveness of presence of La-5 CFS in the cell culture before intoxication with C. difficile CF was also tested using confocal scanning laser microscopy to compare the cell integrity changes with and without
4 o La-5 treatment. Caco-2 cells seeded in glass bottom plates (1×10 cell/mL) and incubated at 37 C, 5% CO2 for 48 h to become 50% confluent to be used for the cell monolayer were pretreated with 3% v/v La-5 CFS one hour before exposing to C. difficile CF 2 fold diluted culture filtrate. Plates were incubated for 24 h at
o 37 C, 5% CO2. After 24 h, the cell monolayer was washed with PBS and fixed with 3 mL cold paraformaldehyde (4%) for 15 min and 3 mL Triton-X-100 for 15min followed by adding 1 mL 3% BSA for 1 h. Finally, the fixing solution was removed and fixed Caco-2 cells were stained with a 1mL 1:1000 diluted TRITC (Tetramethyl Rhodamine Iso-Thiocyanate) for 1 h and 1 mL 1:100 DAPI (4',6-diamidino-
2-phenylindole is a fluorescent stain) 4 mins. Experiments were conducted in duplicate and repeated two times. Confocal scanning laser microscopy (DMIRB Inverted Fluorescence Microscope, Leica
Microsystem, Concord, ON, Canada) was used to visualize the intercellular integrity of the epithelial monolayer when they were exposed to La5-CFS.
4.1.5 Adhesion assay
In this study, the impact of L. acidophilus La-5 culture supernatant on the adherence of C. difficile to Caco-
2 and HT-29 cells was investigated in vitro. The C. difficile adhesion assay was carried out by pre- incubating both bacterial cells and cell culture monolayers with optimal concentrations of La-5 CFS before infecting the cells with the bacterium. Caco-2 and HT-29 cells were grown in DMEM supplemented with
10% FBS for both cell lines and 1% non-essential amino acids for Caco-2 cells. The cell monolayers were prepared in 24 well tissue culture plates and were used when late confluence was achieved (15 days after 98 seeding for Caco-2 and 5-7 days for HT-29 cell line to have fully differentiated confluent cell monolayers).
One day prior to adhesion assays, cells were washed with PBS and then DMEM without antibiotic supplement was added to each well. Cell culture monolayers were pre-incubated with 3% La-5 CFS 2 h prior to the infection experiment in an anaerobic condition, to minimize the air in the medium before adding
C. difficile bacterial cells. C. difficile strain 027 was grown overnight until late exponential phase, OD600 nm= 0.6, was reached.
For infecting the cell cultures, in both sets of experiments, bacteria and tissue culture cells were incubated together for 1.5 h at 37oC under anaerobic conditions, and then plates were washed three times with preincubated PBS in order to discard non-adherent bacteria. In each plate, 2 wells were used to enumerate cell-adherent bacteria after trypsinization and spreading appropriate dilutions on BHI (Brain Heart Infusion) agar. Results were reported as CFU/mL of cell-adherent bacteria. The assays were repeated on three biological replicates as recommended by Denève et al., (2008). For each treatment, one well was used for fixation and staining (Gram stain) that had circular cover slips embedded under the cell culture monolayer before being seeded.
Fixing and Gram Staining
The cell culture medium was removed and gently washed three times with the PBS buffer. PBS buffer was removed and 1-2 mL of 4% paraformaldehyde were added to the cell culture medium. Plates were left at 4 oC for 1 h. For gram staining the cells in the adhesion experiment, the fixing solution was removed and five drops of crystal violet dye solution were added to the fixed culture and left with the stain for 60 seconds. In order not to disturb the cells, adding the stains and washing were performed from the sides of the well. The stain was gently withdrawn with a disposable pipette and the excess stain rinsed with water. A few drops of the iodine solution were added to the culture, enough to cover the fixed culture and was left to stand for
50 seconds. The iodine solution was poured off and the well contents were rinsed with water and the excess water was removed. A few drops of ethanol (95%) were added and subsequently rinsed off with water after
20 seconds. The exact time to rinse the ethanol is when the solvent is no longer colored as it flows over the
99 culture. Further delay will cause excess decolourization in the gram-positive cells, and the purpose of staining will be defeated. In the final steps, cells were counterstained with 5 drops of the red safranin solution for 50 seconds, which was then washed off with water. The excess water was removed by pipetting.
The stained slide was examined under a microscope with a 40× objective lens. Each image is representative of images from at least in 10 random microscopic areas.
4.1.6 Statistical Analysis
All experiments were performed in three independent biological samples and each sample was duplicated unless otherwise stated. Quantitative data were analyzed using Graph Pad Prism software version 7.3.
Results were expressed as mean ± standard deviation. Comparisons were conducted between groups using one-way and two way analysis of variance ANOVA, and differences when P < 0.05 were considered statistically significant.
4.2 RESULTS
4.2.1 Inhibition of Cytotoxicity in C. difficile Ribotype 027 by L. acidophilus La-5 CFS
Before we assessed the inhibitory effect of La-5 CFS on toxin production and potential protective effect of
La-5 CFS on the cell lines, it was necessary to know the maximum concentration of La-5 CFS that produces no adverse effect on the cells. A SRB assay was performed to determine the non-toxic concentration of the
La-5 CFS. In vitro studies suggested bioactive molecules and peptides can show cytotoxicity when their quantity exceeds a certain concentration (Saar et al., 2005; Pushpanathan et al., 2013). Furthermore, an in vivo study by Tellez et al., (2011) showed that mice fed with partially purified LH-2 CFSM fractions with a high dose (0.08 µg/day) resulted in lower tolerance and survival rates after Salmonella infection compared to non-stimulated mice. The HT-29 monolayer was co-incubated with different concentrations
(1-6 % v/v) of reconstituted La-5 CFS and m-MRS CFS, then the viability was measured by the SRB assay as described by Dingle et al., (2011).
100
The SRB assay results showed little effect from CFS La-5 concentrations of 1% and 2%. Indeed, these concentrations seemed to increase cell growth. However, some cell damage occurred when La-5 CFS concentrations reached 4% and higher (P < 0.05) (Figure 4.1). A concentration of 3% was the optimum concentration with no positive or negative effect. The m-MRS medium alone also showed some influence on the cell viability at a concentration of 6%. Both 1- and 2-fold dilutions of C. difficile culture filtrate killed HT-29 cells at rates of 100% and 90 %, respectively (Figure 4.2 A) Thus, a 2× dilution of culture filtrate was used in subsequent experiments. BHI medium did not show any significant impact on toxicity of the cell monolayer (Figure 4.2 B).
The effect of La-5 CFS on C. difficile toxin production was determined by SRB assay as described previously. La-5 CFS was introduced to the HT-29 monolayer in two ways: 1) pretreating cells with La-5
CFS directly and 2) coincubating C. difficile with La-5 CFS. In the second treatment, C. difficile culture filtrate was prepared the same way that as for the gene expression assays (Section 3.3) by adding 10% v/v
La-5 CFS to the C. difficile culture in BHIS (brain heart infusion, supplemented with 0.1% sodium taurocholate) medium and incubated in an anaerobic incubator for 48 h. Results showed that pretreated cell monolayers containing La-5 CFS significantly reduced toxicity effects on HT-29 cells by protecting 41.1
% ± 1.98 (P = 0.012) viable cells compared to HT-29 cells infected with the pathogen and even 8.1 % more than when C. difficile was co-cultured with La-5 CFS and the culture filtrate was added to HT-29 (33% ±
2.54 (P < 0.05) protection of viable cells). m-MRS did not show significant inhibition of toxin production in treatment 2 when coincubated with C. difficile. (Figure 4.3 A & B).
C. difficile neutralization assay using toxins A and B VHH antibodies was conducted on the HT-29 and
Caco-2 cells. The cytopathic effect of C. difficile culture filtrate co-incubated with antibodies was observed under the inverted light microscopy. Percentage of inhibition of cell rounding was witnessed showed more than 75% of cells stayed intact without shrinking or rounding in the presence of TcdA antibodies (A20.1
101 and A5.1). This confirmed that there is toxin A produced in the culture filtrate by C. difficile strain 027.
This experiment was performed in over two successive passages of epithelial cells.
1 5 0
n s
)
% (
1 0 0 y
t * *
i
l i
b *** a
i ****
V
l
l 5 0
e C
0
d e % % % % % % t 1 2 3 4 5 6 a e r s t ll n e U C L a -5 C F S C o n c e n tra tio n s
Figure 4.1. Analysis of SRB assay measuring non-toxic concentration of L. acidophilus La-5 treatment on HT-29 cells. HT-29 cells seeded in 96 well plate were treated with a series of concentrations of La-5 CFS and incubated for 24h. Percentage of cell viability was calculated over untreated cells for each treatment. Results represent the means ± SD of tested samples in triplicate from three independent experiments. Bars identified with ns are not significantly different. Bars identified with **** are extremely different (P<0.0001), bars identified with *** are very significantly different (P < 0.001), bars identified with ** are significantly different (P < 0.05), and ns represent not-significant differences.
102
1 5 0
n s
A ) %
( 1 0 0
y
t
i
l
i
b
a
i
V
l
l 5 0
e C
**** **** 0
d X X X X X X e 1 2 4 8 6 2 t 1 3 a e r s t ll n e U C C . d if f ic ile C F S s e r ia l D ilu t io n
1 5 0
n s
)
% (
1 0 0
y
t i
B l
i
b
a
i
V
l
l 5 0
e C
0
x x x x d 2 4 8 6 te 1 a e r s t ll n e U C B H I S e r ia l D ilu t io n
Figure 4.2. Analysis of SRB assay measuring toxic concentration of C. difficile culture filtrate on HT-29 cells. (A) toxic concentration of C. difficile culture filtrates and (B) toxic concentration of BHI medium as control. HT-29 cells seeded in 96 well plate were exposed to C. difficile 027 culture filtrate (grown in BHI medium), in 2-fold series of 1, 2, 4, 8, 16 and 32 times dilution. The SRB assay was performed and cell viability was expressed as the percentage of survival HT-29 cells that were exposed to C. difficile culture filtrate. Percentage of cell viability was calculated over untreated HT-29 cells for each treatment. Results represent the means ± SD of tested samples in triplicate from three independent experiments. ns = not-significantly different. Bars identified with**** represents extremely significant difference (P < 0.0001).
103
1 2 0 )
9 0
%
(
y
t
i l
i * * * *
b 6 0 a
i *
V
s
l l
e *
3 0 C
0
9 7 S S 2 2 F - 0 C R T C 5 M H - 7 a 2 L 0 d C
P r e tr e a te d C . d iffic ile t P r e tr e a te d H T 2 9
Figure 4.3. C. difficile cytotoxicity changes in the presence of La-5 CFS. Black bars indicate pretreated HT-29 cells with La-5 CFS and m-MRS, were exposed to 2 fold diluted C. difficile 027 culture filtrate, and incubated 24h at 37oC. The SRB assay was performed and percentage of cell viability was calculated over untreated HT-29 cells negative control. Grey bars indicate HT-29 cell cultures were exposed to C. difficile 027 co-cultured with 10% La-5 CFS for 24h at 37oC. Results represent the means ± SD of tested samples in triplicate from three independent experiments. Bars identified with *, **** are significant, and extremely significant different (P < 0.05; P < 0.001; P < 0.0001 respectively).
4.2.2 Inhibition of Cytopathic effect of C. difficile Ribotype 027 by L.acidophilus La-5 CFS
To determine the efficacy of La-5 CFS in reducing C. difficile cytopathic effects, cell rounding was observed under the inverted microscope as described in section 4.1.4. Exposure of HT-29 and Caco-2 monolayer cells to the C. difficile culture filtrate co-cultured with La-5 CFS and m-MRS, reduced cytopathic effects and the results are shown in Figure 4.4. In the presence of untreated C. difficile CF,
(images A and F) morphology of the HT-29 and Caco-2 cells was changed and almost all the cells became rounded in appearance, while Image B and G illustrate that C. difficile co-cultured with m-MRS triggered
104 a reduced amount of rounded cells, but it damaged the cell-to-cell borders and caused detachment of both monolayers from the well’s bottom.
On the contrary, C. difficile CF that was co-cultured with La-5 CFS did not alter cell morphology achieving results similar to the untreated controls, and as images C and H indicate, very few cells in image C were shrink but not rounded and very few cells in image H were partially rounded, which confirms that the effects were specifically related to the activity of La-5 bioactive molecules in that treatment compared to the m-
MRS treatment. Furthermore, C. difficile CF from the co-cultured C. difficile with WPLa-5 CFS or WP did not cause any cytopathic or cell rounding effects on monolayers of HT-29 (D and E) or Caco-2 (I, J).
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HT-29 Caco-2
A F
50 µl
B G
C H
D I
E J
Figure 4.4. Morphological changes induced by C. difficile culture filtrate samples. Images A-E represent HT-29 cells and F-J represent Caco-2 cells. A and F: cell lines were exposed to the C. difficile culture filtrate. B and G: C. difficile co-cultured with m-MRS; C and H: C. difficile co- cultured with La-5 CFS; D and I: C. difficile co-cultured with La5 CFS WP (whey protein medium); E and J C. difficile co-cultured with WP.
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4.2.3 Inhibition of C. difficile cytotoxicity on Cell Monolayer intercellular Structure by
L. acidophilus La-5 CFS
The effectiveness of La-5 CFS treatment in protecting cell monolayer structure from C. difficile toxins
(culture filtrate) was examined using confocal scanning microscopy and HT-29 monolayer with more than
50% confluency. Phase contrast images are presented here (Figure 4.5). From the images, we noticed significant differences between two samples. In image A, we visualized a cluster of rounded or dead HT-
29 cells; in image B we observed that HT-29 monolayer pretreated with 3% La-5 CFS and then exposed to
C. difficile CF, was free of any rounded or dead cells. The cytoplasm and nucleus structures were intact, with the nucleus located in the middle of the cell without marginalization which is a sign of cytopathic impact on the cells.
A. B.
Figure 4.5. Confocal Image of HT-29 cell integrity pretreated with La-5 CFS. 1 ×10 4 cell/mL was seeded in glass bottom plates and cell monolayers were more than 50% confluent, pretreated with La-5 CFS and exposed to C. difficile CF. image A is the control sample HT-29 and C. difficile CF; Image B is pretreated HT-29 cells with La-5 CFS and exposed to C. difficile.
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4.2.4 Impact of La5 CFS on C. difficile Adhesion
The aim of this study was to understand the impact of La-5 CFS on non- toxigenic virulence factors of C. difficile 027. To achieve this goal, an anaerobic adhesion assay was developed and adherence of C. difficile bacterial cells to Caco-2 and HT-29 cells was tested by comparing two different treatments with the controls. First, adhesion assay results of treatment 1 (i.e. both cell cultures and C. difficile culture were pre- incubated with La-5 CFS before an adhesion test) were compared with a negative control (C. difficile cells without pre-incubation with La-5 CFS) infected untreated cell cultures. The Sidak’s comparison tests in one-way ANOVA analysis of adhesion assay results showed that treatment 1 significantly inhibited highly infectious C. difficile strain 027/ NAP1 bacterial cells from association with the cell culture monolayers.
Figure 4.6 illustrates that the count of adherent C. difficile cells was decreased by 1.52 log CFU/mL for the
Caco-2 cells and by 2.2 log CFU/mL for the HT-29 cells compared to the negative control (P < 0.001).
Additionally, Figure 4.6 shows adhesion assay results of treatment 2 (i.e. untreated C. difficile infected pretreated cell cultures containing 3% La-5 CFS), when compared to the negative control, and revealed that
C. difficile strain 027 adherence on HT-29 was significantly reduced by the presence of La-5 CFS in the cell culture before infection (P =0.0001; 1.76 log CFU/mL reduction). Although La-5 CFS reduced adhesion of C. difficile to Caco-2 cells, the effect was not statistically significant (P=0.051). Therefore, bacterial cells that were exposed to the La-5 CFS before the adhesion assay had lower attachment to the cell monolayers than those C. difficile cells without treatment.
Furthermore, comparing the adhesion of C. difficile bacteria on the two cell lines used in our study showed that although they exhibited similar outcomes in adherence to the negative controls, there was a statistically significant variation of adhesion between them in cells treated with La-5 CFS. The count of C. difficile attached to the HT-29 in treatment 1 was 0.39 log less than Caco-2 cells (P < 0.002) and the count of C. difficile attached to the treated HT-29 in treatment 2 was 0.5 log less than treated Caco-2 cells (P < 0.0005)
(Figure 4.6).
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8 C a c o - 2 C e lls **** H T - 2 9 c e lls 6
**** L
m n s
/ ** U
F 4 *** C
**
g **
o L 2
0
7 7 7 7 2 2 2 2 0 m 0 0 u 0 s d l d D d ll C C s C u r l ls C e c l r l r C o T e n T e T r In C n c T U U r & T &
T re a tm e n ts
Figure 4.6. C. difficile strain 027 adherence to Caco-2 and HT-29 monolayers. One strain of C. difficile and two types of cell culture were tested in an adhesion assay mostly in anaerobic conditions. Cd027 is the total live bacterial count that was inoculated in the cell culture monolayers. UT is for untreated samples with La-5 CFS and T is for treated samples with La-5 CFS Adherent C. difficile cells were counted by plate counting and log 10 CFU/mL was calculated. All experiments were performed in duplicate and repeated three times. Error bars are standard deviation of the means. Bars identified with **, ***, **** are significantly, very significantly, and extremely significant different (P < 0.05; P < 0.001; P < 0.0001 respectively) and ns means not significant differences.
4.2.5 Visualized C. difficile Adhesion to the Cell Culture Monolayers
Adhesion of C. difficile to the treated HT-29 cells containing 3% La-5 CFS was observed using light microscopy (Figure 4.7). Images showed that C. difficile bacterial cells were evenly attached to the HT-29 cell monolayers. Interestingly C. difficile bacterial cells were associated with the cell-cell borders, but they did not show any clustering effect in any specific field visualized by microscopy. We also noticed that HT-
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29 monolayer stayed intact under anaerobic pre-incubation and incubation and its morphology was consistent in all the samples, except in treated samples with modified MRS when they detached from the slides embedded under the monolayers; this happened in all replicates. Therefore, cells and bacteria were washed off completely or partially after washing three times with PBS. Image B as negative control shows that there are much higher numbers of C. difficile bacteria attached to the untreated HT-29 monolayer than in image C where C. difficile was added to the HT-29 monolayer pretreated with La-5 CFS (Treatment 1).
5CFS -
B:Cd027
C: Cd027 La C:Cd027
29
- A: A: HT
MRS D:Cd027
Figure 4.7. Microscopic visualization of C. difficile adherence to HT-29 cells. Untreated live C. difficile 027 bacteria were inoculated onto pretreated HT-29 cell monolayers and incubated for 1.5 h in anaerobic conditions at 37 oC. Unattached bacteria were washed and the attached bacteria were gram stained with the standard gram staining method. Images were captured at 40× magnification with light microscopy and are representative of multiple images observed from the whole culture and two replications. Black spots indicate bacterial cells attached to monolayers. Image A represents HT-29 cells as a control. Image B represents C. difficile 027 adherence on HT-29 monolayer without any treatment. Image C represent C. difficile adherence on the pretreated HT-29 monolayer.
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4.3. DISCUSSION
C. difficile is a non-invasive toxin producer pathogen which does not need to enter host cells to cause infection; instead it produces exotoxins A and B to disturb the actin arrangement of the cytoskeleton. One of the potential therapeutic targets for treating CDI is the reduction of C. difficile toxin production, and in our study, we investigated the impact of La-5 CFS on inhibition or control of toxin production. Our earlier study demonstrated that L. acidophilus La-5 bioactive molecules influenced the C. difficile PaLoc gene and other virulence genes. Down-regulation of sigma factors and toxin genes was detected after C. difficile was grown in a medium treated with La-5 CFS and results showed that tcdA and tcdB toxin genes were significantly suppressed following treatment with La-5 CFS. In this study, we examined the influence of
La-5 CFS on C. difficile cytotoxicity to a human epithelial cell line, HT-29, to determine the correlation between cytotoxicity and reduction of toxin gene expression. It was confirmed that L. acidophilus bioactive molecules have the ability to reduce toxin production, neutralize toxicity of the C. difficile culture filtrate, and reduce association of C. difficile bacterial cells to host cells in vitro. Cytotoxicity assays proved that
La-5 CFS reduced toxin production or interfered with the toxin’s mechanism to act normally, since our results showed that the presence of La-5 CFS in the C. difficile culture or in the cell monolayers, had significantly higher percentages of viable cells after infection.
Inhibition of C. difficile pathogenicity has been under investigation in different fields of research, including probiotics and its production of bioactive peptides (Rex, 2017; Banerjee et al., 2009; Allen, 2015; Plummer et al., 2004; Yun et al., 2014). An in vivo study by Yun et al. (2014) revealed that L. acidophilus strain
GP1B protected the intestinal epithelial cells of mice against C. difficile infection. L. acidophilus inhibited inflammation of the intestinal epithelial cells and C. difficile growth in the CDI mouse model. Additionally, other studies documented that bioactive peptides derived from lactic acid bacteria (proteobiotics) modulated the virulence of other pathogens such as E. coli, Salmonella and Campylobacter jejuni (Bayoumi &
Griffiths 2010; Mundi et al., 2013; Tellez et al, 2011; Sharma, 2014; Zeinhom et al., 2012). Another study also revealed that probiotics produce bioactive molecules which affect protein expression, which are in
111 charge of pathogenicity of the bacteria (Kos et al., 2003). Studies also showed that a recombinant probiotic
Lactobacillus paracasei inhibited adhesion invasion and translocation of Listeria monocytogenes in Caco-
2 cells by expressing its specific adhesion protein. It also reduced cytotoxicity and protected tight junctions from L. monocytogenes infection (Koo et al., 2012).
Our cytopathic assay visualized the effect of La-5 CFS on protecting epithelial cells in vitro. Interestingly, it proved that L. acidophilus La-5 CFS extracted from two media (m-MRS La-5 CFS and WP La-5 CFS) efficiently inhibited cell cytotoxicity of C. difficile CF when tested in cell lines of HT-29 and Caco-2 (Figure
4.4). A significant increase in efficacy of La-5 CFS was seen when WP La-5 CFS was used as treatment compare to m-MRS La-5 CFS. Moreover, bioactivity of L. acidophilus La-5 CFS was very noticeable in both cytotoxicity and adhesion assays, compared to the results obtained from control medium m-MRS. WP
La-5 CFS also inhibited cytotoxicity of C. difficile CF on monolayers when it was co-cultured with C. difficile, but also it seems that whey protein itself also has inhibitory influence against C. difficile toxicity.
Changes in cell morphology are the first signs of toxin impacts that accompany loss of cell integrity. Toxins cause changes through suppression of F-actin, alteration in cell ultrastructure, and rearrangement of microvilli (Chang et al., 1979). Studies proved that morphological changes are similar between TcdA and
TcdB toxicity. These changes always predicted when cells were exposed to both toxins because actin polymerization determines regulation of structural processes (Voth & Ballard, 2005). Marginalization of the nucleus results because of changes to the cytoskeleton (Fiorentini et al., 1990) and is another sign of alteration in cell morphology (Voth & Ballard, 2005).
The confocal microscopic images of HT-29 cells illustrated the impact of La-5 CFS and showed that monolayer cells containing La-5 CFS that have been exposed to C. difficile culture filtrate were not affected by toxicity and the major damage that toxins cause, including rounding and barrier destruction. As these images show, the integrity of the cells has not been damaged, the cell nucleus is intact, and cells are still
112 alive. In contrast, control cells lost their morphology and cells were rounded without showing any signs of viability.
Confocal microscopy that captures 3D images is able to show images from different layers of a sample, but it is assumed that because the normal functions of dead or rounded cells have been impaired, they were not able to uptake stains from the staining solution. Therefore, fluorescence was emitted only from the outer layer of the cells, which could be the reason that the inner structure was not visible under the microscope
(Dailey et al., 2006). The same results have been reported by Gratz et al., 2007 using L. rhamnosus GG on
Caco-2 cells against aflatoxin B1. They also suggested that using probiotics could be an alternative to inhibit toxin absorption in the gastrointestinal tract.
As several studies show, adhesion of C. difficile and other pathogenic bacteria to the epithelial cell surface is a prerequisite for colonization, virulence manifestation and development of infection, and is required to modulate the host immune system (Borriello et al., 1988; Naaber et al., 1996). Therefore, colonization and attachment of the pathogen to host cells is a critical event to be studied in C. difficile pathogenesis. Adhesion of bacteria is based firstly on non-specific interaction between two surfaces, and secondly through specific interaction between adhesins (proteins) and complementary receptors (Pérez et al., 1998; Rojas & Conway,
1996). When live probiotic bacteria are present in the gut, they prevent pathogenic colonization by auto- aggregation and co-aggregation. Auto-aggregation acts as a barrier to protect the epithelial cells and co- aggregation functions to slow down or inhibit pathogens from attachment to the mucus layer and subsequent colonization (Boris et al., 1997; Reid et al., 1988). L. acidophilus is one of the most commonly found species in the intestine that has a very high aggregation ability and its co-aggregation is considered as a potential antagonistic action against pathogenic bacteria to defend their host from infection (Kos et al.,
2003).
As adherence is the vital stage for pathogenic bacteria in order to colonize and establish in their niche
(Finlay et al., 1989), they possess intricate adhesive surface components to bind to the cells surface
113 components to be able to carry their toxins to their desirable targets. Adherence is an active process that provokes some outcome, including response from their target cells. One of the parameters that affects the adherence outcome is whether the receptors have been up or down regulated when the ligation occurs. Other parameters that influence adherence include adhesion alone or with a group of other products, and the receptor domain involved in binding such as carbohydrate antigens, protein antigens, and fimbriae
(Hoepelman & Tuomanen, 1992).
Studies indicated that non-viable secreted components from probiotic bacteria interfere with the infection process of enteropathogens. In this study, instead of live probiotics, we used L. acidophilus cell free supernatant which contained bioactive molecules produced by the bacteria. We added this supernatant to epithelial cells (Caco-2 and HT-29 cells) which resulted in the significant inhibition of adhesion, cytopathic effects and cytotoxicity of C. difficile. Quantitative results were in agreement with the images from adhesion and cytopathic assay. The gene expression results did not show any significant changes to slpA a gene encoding surface layer proteins but another study (Johnson-Henry et al., (2007) suggested that probiotic surface layer proteins could potentially outcompete intestinal pathogen adhesins. They indicated that
Lactobacillus helveticus extracted S-layer proteins prevent adherence of E. coli O157:H7 to intestinal epithelial cells and they also suggested that probiotic bacterial adherence depends on cell surface hydrophobicity. Their investigation on the effectiveness of S-layer extract on protection of epithelial cell integrity concluded that probiotic surface proteins maintain epithelial cell barrier function and their integrity.
Downregulation of quorum sensing gene luxS and reduction of AI-2 production could be a factor to inhibit adhesion of bacterial cells to the epithelial cells. There may be other factors that can interfere with the binding of C. difficile surface proteins or glycoproteins to the receptors in the cell culture, such as (glycol-
) proteinaceous material that has been separated from the L. acidophilus cells in their cultures as well as other molecules such as SLPs, and adhesins. A surface associated protein GroE, a highly conserved protein and essential for all living organisms, is one of the proteins that has been found not only in probiotic strains
114 including L. johnsonii that mediate aggregation of Helicobater pylori (Bergonzelli et al., 2006), but also in some pathogens like C. difficile (Hennequin et al., 2001).
Our study showed that pre-incubation of mammalian cell lines with La-5 CFS significantly reduced C. difficile adhesion to the Caco-2 and HT-29 monolayer, although it is more effective when La-5 CFS is present in cell monolayers and C. difficile CF at the same time. The comparison of two cell lines showed that the two cell lines exhibit different levels of adhesion inhibition that could be due to the types of cell lines. HT-29 cell lines are mucus secreting cell lines (Martínez-Maqueda et al., 2015) and there are contradictions as to whether the mucus itself can increase attachment of bacterial cells.
Furthermore, C. difficile usually binds to the mucus layer as a cluster or as isolated cells. The presence of bioactive peptides can alter the surface and prevent diffusion of pathogens like C. difficile into the dense mucosal layer. This may be mediated by blocking the binding sites through formation of protein-peptide bonds or inhibit the auto-aggregation of bacteria, and prevent aggregation and biofilm formation. Studies suggested that components of C. difficile culture media (treatments) can associate with their bacterial cells during the co-incubation, which has been shown by the SDS-PAGE profile of C. difficile (Eveillard et al.,
1993). Their results showed that small proteins and peptides bind to the bacterium during growth and this could prevent bacterial adhesion to surfaces or inhibit auto-aggregation. Surface associated proteins are also important elements for the adhesion of C. difficile (Eveillard et al., 1993). These results suggest that bioactive molecules interfere with the cell receptors and adhesion factors of C. difficile or block the binding site of their cells to prevent cell-to-surface attachment.
4.4 CONCLUSION
L. acidophilus secreted bioactive molecules effectively reduced the cytostatic and cytocidal effects of C. difficile toxins. The protective effect of L. acidophilus bioactive molecules in our cell-base in vitro study suggests that bioactive molecules could be considered as a supplement to the therapies used for CDI patients but further work using animal models is needed to confirm this effect.
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CHAPTER 5
5. IMPACT OF LA-5 PROTEOBITICS IN WHEY PROTEIN MEDIUM ON BIOFILM
GROWTH AND SPORULATION OF CLOSTRIDIUM DIFFICILE
Abstract
Bioactive molecules are health-enhancing compounds produced by probiotics. The aim of this work was to determine the effect of bioactive molecules produced by Lactobacillus acidophilus La-5, namely proteobiotics or WpLa-5 CFS, on the biological activity of C. difficile; including biofilm formation, spore production and motility. C. difficile is a major cause of antibiotic - associated diarrhea as a result of antibiotic therapy and disruption of normal microbiota of the intestine. In addition to the main virulence factors (toxin A and B) of C. difficile, biofilm forming ability of this pathogen increases antibiotic resistance and recurrent infections. Motility also plays a role in the establishment of infection while spores are the vehicle through which the pathogen is transmitted and spread through the environment to human and animal populations. We investigated whether these three biological activities of three toxigenic strains of C. difficile (ribotype 027, 078, and 001) differed due to the presence of active molecules of bioactive molecules. L. acidophilus La-5 was grown in a chemically modified medium containing whey protein concentrate for 48 hours and the cell free supernatant obtained was concentrated by freeze drying. Three C. difficile strains were grown with or without WpLa-5 CFS, and then cultures were used for a quantitative biofilm assay, as well as sporulation and motility assays after 3, 5, and 2 day incubation periods, respectively. Scanning electron microscopy (SEM) was used to visualize the impact of treatments on biofilm formation. WPLa-5 CFS reduced attachment and population of C. difficile cells encased in the biofilm and increased non-aggregated planktonic cells in the media significantly (P < 0.0001). SEM analysis was consistent with the quantitative data. Motility was not affected by the treatment and varied between strains. Sporulation decreased in ribotypes 078 and 001 but in ribotype 027 spore numbers
116 increased. Our results suggest that WpLa-5 CFS could potentially be used to inhibit biofilm formation of
C. difficile and prevent recurrent C. difficile infection (CDI) in patients.
Keywords: Clostridium difficile, Probiotics, Biofilm, Sporulation, La-5 bioactive molecules, proteobiotics, whey protein.
Introduction
Clostridium difficile, one of the greatest concerns that healthcare systems face today, is an intestinal opportunistic pathogen for humans and animals and is the main cause of antibiotic-associated disease
(Runpik et al., 2009). Most research has focused on C. difficile toxins and mechanisms of pathogenesis.
However, it is noticeable that non-toxigenic virulence factors also play an important role for the bacteria to establish in the host intestine. Since 2000, the rate and severity of CDI has increased, particularly in North
America and Europe (Redelings et al., 200; Bauer et al., 2011). CDI mostly targets patients on antibiotic therapy, immunocompromised and the elderly (Barbut et al., 2000). Recently emerged C. difficile strains are more resistant to multiple antibiotics. This relates to an increasing number of recurrent infection in CDI patients (Viswanathan et al., 2010). Reinfection is common in 15-35 % of susceptible individuals within two months of CDI treatment (Vedantam et al., 2012; Janoir, 2016). The cause of reinfection and persistence of C. difficile is not completely known. Biofilm formation by C. difficile could be one factor leading to reinfection.
Bacterial biofilms play a role in the protection of the bacterial community from external damage and increased persistent infection. Additionally, biofilm formation by bacteria confers on them greater antibiotic-resistance and protects them from the host immune responses (Davies, 2003). Up to 80% of bacterial infection is related to biofilms (ref). Biofilm formation also increases chronic recurrent infection by pathogenic bacteria such as Clostridium difficile (Dapa et al., 2013; Dawson et al., 2012). Biofilm formation is a potential threat in food processing and nosocomial settings as biofilms are observed in the
117 environment and on surfaces like plastics, metals, glass, and more. The bacterial world is dominated by two modes of growth: planktonic and biofilm. Individual organisms that move freely are planktonic cells, whereas biofilms are communities of cells encased in a gel like matrix and characterized as multifaceted, heterogeneous, and structurally composed of single or multispecies entities (O’Toole et al., 2000). Although bacteria living in a biofilm grow naturally, most antimicrobial research has focused on bacteria during planktonic growth, (Hernández-Jiménez et al., 2013).
Gene expression and physiological behavior of bacteria differ between the two modes of growth (planktonic and biofilm). Bacteria in their biofilm mode are less active than bacteria in the planktonic growth mode; in the biofilm mode they exhibit a slower metabolism (Bester et al., 2011) and decreased motility (Guttenplan
& Kearns, 2013). In many bacteria, biofilm formation is linked to inter-species communication known as quorum sensing. Bacterial transition from planktonic to multicellular biofilm is completed using quorum sensing and their auto inducers (signaling molecules) (Kaufmann et al., 2008).
As C. difficile is a spore-forming, Gram-positive, anaerobic bacterium, spore formation is a natural survival mechanism of the bacterium, and an important trait that allows it to spread in the environment. As we focused on the influence of proteobiotics on the biofilm forming activity of C. difficile, we also tested whether these compounds affect spore production during biofilm growth of the bacterium by counting heat resistant spores produced by C. difficile in different cultures.
C. difficile secretes two major toxins, toxin A and B. Before two exotoxins start damaging the mucosa and intestinal inflammation occurs, C. difficile must find a way to reach the epithelial cells by penetrating through a thick layer of dense mucous (Karjalainen et al., 1994), using their flagella or proteases (Poilane et al., 1998). Karjalainen et al. (1994) also showed that the flagellin gene is present in all flagellated and non-flagellated C. difficile strains. Tasteyre and colleagues (2000) suggested that although the exact role of flagella is as yet unknown, chemotaxis and the flagellar system may be involved in all the steps of infection from adapting to the microenvironment to adherence and colonization.
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According to studies, motility of bacteria give them the ability to adapt to divergent environments including water, soil and microenvironments in their host. Motility is executed mostly by the bacterial flagellum.
Flagellum-driven cell movement in some bacterial species acts as an invasion factor or aids colonization of the gastrointestinal tract. Motility assists pathogens to penetrate through the mucus layer to reach their targeted epithelial cells before colonization and biofilm formation occur. A study by Tasteyre et al. (2001) confirmed that motility has a role in adherence and colonization. McKee et al. (2013) reported that the same signaling molecule that regulates TcdR (sigma factor for toxin A and B) also regulates motility and they concluded that motility has a role in virulence of C. difficile. Therefore we tested if our proteobiotic compound influenced motility of C. difficile by performing a motility assay.
Our approach toward the issue of rising CDI relapses and C. difficile associated disease (CDAD), was to investigate if proteobiotics limit biological activity of C. difficile and reduce its pathogenicity through inhibiting biofilm formation. The objective of this study was to investigate whether proteobiotics from L. acidophilus La-5, grown in whey protein medium, control biofilm formation and at the same time impact spore production and motility of the three strains of C. difficile, viz. ribotypes 027, 078 (Metcalf et al 2010) and 001 (Vohra and Poxton, 2011).
5.1 Material and Methods
5. 1.1 C. difficile Biofilms
5.1.1.1. Biofilm Formation of C. difficile with Proteobiotics
The effect of bioactive molecules (proteobiotics) on biofilm forming characteristics of C. difficile strain
C027 was determined. Bioactive molecules can act as a stressor for bacterial cells, sometimes causing biofilms to form differently. To investigate this phenomenon, pre-equilibrated BHIS medium (Brain Heart
Infusion agar, 0.1% % w/v sodium taurocholate) (C. J. Smith et al., 1981) was inoculated with 5 colonies of C. difficile per 10 mL BHIS cultures of three strains of C. difficile 027, 078, and 001. All the cultures
119 were incubated at 37oC in an anaerobic chamber (80% nitrogen, 10% hydrogen, 10% carbon dioxide).
Whey protein medium (WP) was prepared as described in section 3.1.2 and whey protein L. acidophilus
La-5 cell free supernatant (WpLa-5 CFS) was prepared as described in section 3.1.4.
A dilution of 1/100 of the overnight cultures of each C. difficile strains was treated with both 10% WpLa-
5 CFS and 10% WP and a control sample was taken that was BHIS containing C. difficile culture without any treatment. Lyophilized powders of WpLa-5 CFS and WP were reconstituted in 2 times more distilled water to have 100mg/ml protein per treatment. Treated and untreated cultures were grown in 2 ml of pre- equilibrated BHIS in two 24-well plates (Corning Costar TC-treated Multiple Well Plates, Sigma Aldrich) for 3 days at 37oC in an anaerobic chamber. One plate was used for total count (planktonic and attached as biofilm), the other plate was used for counting encased bacterial cells within biofilms attached to the well surfaces after 3 days’ incubation. Every sample was analysed in triplicate, and two media blank controls were included in each plate. Plates were sealed with parafilm to prevent evaporation.
After three days, in order to total count the planktonic and attached bacteria in the culture, excess BHIS medium was removed by pipetting, each well was washed three times with 1 ml phosphate buffer saline
(PBS), followed by adding 1 ml PBS again to transfer the disrupted biofilm to the test tube. Viability counting was performed by plating 1:10 dilution in triplicate onto BHIS plates supplemented with 0.1% taurocholate. Assays were performed with three technical replicates and two biological replicates.
5.1.1.2. Analysis of Biofilms and Scanning Electron Microscopy (SEM)
C. difficile liquid cultures were prepared in BHIS treated with 10% L. acidophilus WpLa-5CFS as described previously. Treated cultures in a volume of 1 mL were dispensed into 24 well tissue culture plates equipped with a low evaporation lid (Corning Costar TC-treated Multiple Well Plates, Sigma Aldrich) containing pre-incubated BHIS medium. A glass cover slip (NeuVitro, Round German Glass, 10mm) was embedded
120 at the bottom of each well to obtain SEM imaging samples. Plates were incubated for three days at 37oC anaerobically. Each analysis was performed in duplicate and the experiment was carried out twice in order to include non-treated cultures and media blank control. The samples were fixed and removed from the anaerobic chamber for Scanning Electron Microscopy (SEM) with 10 kilovoltage, 3.0 um=10000× magnification and 30um= 5000× magnification.
C. difficile biofilms preparation for Scanning Electron Microscopy (SEM)
Prior to SEM, each biofilm sample attached to the glass cover slip in each well was fixed immediately.
Wells were washed once with SEM buffer [0.07 M sodium phosphate dibasic (Na2HPO4) and 0.07 M potassium phosphate (KH2PO4), in a 50: 50 ratio)]. Biofilms were fixed on the glass slides using 500 µl of
2% glutaraldehyde in SEM buffer and left overnight. The biofilms were rinsed with SEM buffer and washed with the same buffer three times. The cover slips were post-fixed with 1% osmium tetroxide (OsO4) in SEM buffer at room temperature for 30 mins, rinsed off with SEM buffer once, and then a dehydration process was carried out using sequential concentrations of 50, 70, 80, 90 and 100 % ethanol with each being applied for 10 mins.
The final step involved dehydrating the cells completely using 100 % ethanol added 3 times within 15 minutes to dry the bacterial cells without introducing surface tension artifacts. Samples were dried at their critical control point using pressure and dry ice in a Critical Pivot Dryer. Each glass cover slip was located
o in the dryer baskets. The ethanol content of the bacterial cells was replaced by CO2 at 32.7 C and not more than 1350 psi vapor pressure was used to replace ethanol with CO2 gradually. After drying, samples were carefully mounted on an aluminum stub using double stick carbon tape. Samples were coated with gold in an Emitech K550 sputter coater for 2 min to reach a thickness of 10 nm. Each sample was observed with a
Hitachi S-570 electron microscope (Hitachi High Technologies Inc., Tokyo, Japan) at accelerating voltage of 10-20 kV and a working distance of 5-10 mm. Digital Images were obtained using Quartz PCI software
121 version 9 (Quartz Imaging, Vancouver, British Columbia, Canada) and each sample was prepared and examined by SEM in duplicate.
5.1.2. C. difficile Motility and Effect of Proteobiotics
The effect of La-5 proteobiotics on motility of three strains of C. difficile was determined by using traditional techniques as explained by Dingle et al., (2011) and Tastayre (2000). In order to visualize motility of these bacteria, a motility assay was performed using brain heart infusion (BHI) broth containing
0.175% agar (semi-solid agar). Semi-solid BHI agar, in glass tubes, was placed in an anaerobic incubator at 37oC overnight to be pre-equilibrated. An overnight culture of each of the three strains of C. difficile
(ribotype 027, 078 and 001) were prepared containing a final concentration of 10mg/mL WpLa-5 CFS or
10mg/mL WP, and cultures were incubated overnight to reach an OD of 0.7 at 600nm wavelength.
Untreated samples of each strain were prepared as controls. The next day, one loop-full of every culture was stabbed into the semi-solid BHI agar. Inoculated tubes were incubated again for 48 h anaerobically at
37°C. This experiment was repeated twice.
5.1.3. Sporulation of C. difficile and Effect of Proteobiotics
5.1.3.1. Preparation of C. difficile sporulation cultures in liquid media
The effect of bioactive compound in WpLa-5 CFS on accumulation of spores and germination of C. difficile strains was tested using five day old cultures grown in BHIS broth. Liquid cultures of C. difficile strains
C027 and C078 and 001 were prepared as described previously. Ten millitres of liquid overnight cultures were treated with WpLa-5 CFS or WP to a final concentration of 10g/ml in the cultures. Each treatment was performed in triplicate and the experiment was repeated twice. To make sure that there were only vegetative cells in the inoculum, a stock culture was prepared with BHIS broth that was inoculated in ratio of 1:100 and incubated until an OD600 between 0.2 and 0.5 was reached. The sporulation medium was then inoculated 1:100 with this stock culture (Patel, 2011).
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5.1.3.2. Heat treatment of sporulation cultures
In order to enumerate the number of bacteria (spores and vegetative cells) existing in the sporulation medium, 1 mL of each culture was collected from the anaerobic chamber after 5 days, centrifuged at 5000×g for 10 minutes at which time bacterial pellets were washed in PBS and heat-shocked at 65oC for 15 minutes to kill all the vegetative cells (Sambol et al., 2002). In order to obtain a total count of spores and vegetative cells together, a 1 ml non-heated sample of all the cultures was also collected. To prevent the impact of oxygen exposure, both series of samples (heated and non-heated) were taken in the chamber, serially diluted in PBS, and plated onto BHIS agar supplemented with 0.1 % taurocholate (Sigma, United Kingdom). After a 48 hour incubation, bacterial colonies (CFU/mL) were counted and the number of spores was subtracted from total counts and vegetative cell numbers calculated (Patel, 2011). All experiments were performed in triplicate with two biological replicates. Results were plotted graphically in Prism 7.0.
5.1.5. Statistical analysis
Statistical analysis was performed using graph pad prism 7.03 software (Graph Pad Software Inc., La Jolla,
CA, USA). Two way ANOVA was performed to determine if there were significant differences in biofilm formation (P < 0.05) indicates treatment specific differences). Tukey’s multiple comparison test was performed to determine whether there were any remarkable differences in biofilm formation of treated samples with La-5 CFS and WP CFS and both treatment samples compared to a control culture without any treatment (P < 0.05). Additionally, collected data from sporulation was analyzed using one-way ANOVA and Tukey’s multiple comparison test to determine statistically significant variance (P < 0.05).
5.2 RESULTS
5.2.1 Biofilms formation and Effect of La-5 Proteobiotics
The hypothesis of our study was that bioactive molecules produced by L. acidophilus La-5 affect aggregation of C. difficile bacterial cells and therefore biofilm formation will be inhibited. In order to
123 support this hypothesis two experiments were conducted. The first experiment conducted measured total count vs. encased bacteria in the attached biofilm. The second experiment observed the differences between biofilms of treated and control samples in each C. difficile strain using scanning electron microscopy
(SEM).
Biofilms formed in all three strains were tested and data analysis was performed using two-way ANOVA.
Data analysis revealed that the total count (CFU/mL) of bacteria (Figure 5.1), including planktonic and attached cells of the three selected strains, were similar when strain 001 exhibiting a slightly lower count.
Interestingly, treated cultures of all strains showed an increase in the number of colonies; WP treatment showed the highest growth among the treatments but it was not statistically different. On the contrary, after we washed the wells to collect only attached cells, the counts obtained from control samples (only C. difficile culture) were significantly higher than those treated with WPLa-5 CFS (more than three log10 reduction of CFU/mL in treated cultures). Similar counts were obtained from the samples treated with WP.
There was between two to more than three log10 CFU/mL reduction of attached bacterial cells depending on strains, compare to the untreated controls. Since the difference between control and treated samples was statistically significant, therefore, biofilm formation was affected by WpLa-5 CFS and its proteobiotic components and the effect is considered extremely significant (P <0.0001) for all three C. difficile strains.
It is notable that differences between two treatments WP and WpLa-5 were slightly different but not statistically different from each other. That means whey protein also has some effect on attachment, possibly due to protein blocking of the well surface.
Clostridium difficile biofilms were visualized after three days’ incubation by SEM as shown in Figure 5.2.
There was a full layer of biofilm attached to the cover slips at the bottom of each well in untreated cultures, which was visible with the naked eye. However, there was only a residue of biofilm or only a few cells attached to the cover slips in both treated cultures containing WpLa-5 CFS or WP, which was not visible without a microscope. SEM images with magnification of × 3000 and ×5000 were taken from three biological replicates and from three C. difficile strains (C. difficile ribotype 027, 078, 001). From the SEM
124 images of the control cultures, we observed a community of interconnected cells and a network of dead cells, spores, and extracellular material connected to cells (Figure 5.2). Large rod shape cells were attached together with the EPS matrix (encapsulated) and to the cover slips. Conversely, there were few spots on the cover slips from treated samples with or without EPS matrix. In addition, it was apparent that both treatments resulted in a preventative or reducing effect on aggregation and production of biofilm in the cultures. Although bacterial cells did not aggregate and did not attach to the abiotic surfaces of the plates, it was interesting that the total number of bacterial cells in treated samples increased significantly compared to the non-treated cultures of three strains as we reported earlier (P<0.05) (see Figure 5.1). Analysis of
SEM images (Figure 5.3) also clearly shows that population of encased bacterial cells in the biofilm is dramatically different from the images of the treated samples when observed with smaller magnification
(larger scale= 30µM).
1 8 * * * * * * * * * * * *
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T o ta l c o u n t E n c a s e d c e lls in B io film
Figure 5.1. Impact of La-5 proteobiotics on biofilm formation of three C. difficile ribotypes 027, 078, and 001. The error bars represent standard deviations of the means of triplicate determinations. Quantity of C. difficile ribotype 027 strain cells encased inside biofilm was compared by one way ANOVA and Tukey’s multiple comparison test. Results are presented in log scale and error bars represent standard deviation (P< 0.05). * represent significant differences, ** represent very significant differences and **** represent extremely significant differences between bars.
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A B C
D E F
G H I
Figure 5.2. Scanning electron microscopy analysis of La-5 proteobiotics effect on biofilm formation of three C. difficile ribotypes. A) Cd* 027; B) WpLa-5 Cd027; C) WP Cd 027; D) Cd 078; E) WpLa-5 Cd078; F) WP Cd078; G) Cd001; H) WpLa-5 Cd001; I) WP Cd001; * Cd: C. difficile ribotype. Images are representative of three independent experiments. Images are representative of at least twenty viewed field.
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A: C027 B: WPLa-5027
C: C078 D: C078+WPN LA5
E: C001 F: C001+WPN LA5
100µm
Figure 5.3. Comparison of C. difficile cluster between La-5 treated and untreated samples. Scanning electron microscopy analysis of A: ribotype 027, B: treated sample from ribotype 027 co-cultured with WpLa-5 CFS. C: ribotype 078, D: treated sample from ribotype 078 co-cultured with WpLa-5 CFS. E: ribotype 001, F: treated sample from ribotype 001 co-cultured with WpLa- 5 CFS. Biofilm formation assay performed for SEM in duplicated samples and images are representative of three independent experiments. Images are representative of at least twenty viewed field.
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5.2.2 Proteobiotics Impact on C. difficile Motility
Motility is an important tool for gastrointestinal pathogens including C. difficile. In order to determine whether exposure to proteobiotics changes the motility of ribotype 027, 078, and 001, a motility assay was performed to observe differences between the three strains Figure 5.4. When samples of treated and untreated C. difficile strains were cultured by stabbing in motility agar and visually assessed, the appearance and movement of the groups of treated samples were not different from their controls. However, ribotypes
078 and its treated samples were quite different from the two other ribotypes. The cultures with ribotype
078 and its co-cultured samples had very little movement around the stabbing zone and appeared to remain in the original place of inoculation (Figure 5.4, lane 2, 3 and 4). In the control culture of ribotype 027 and its two treated samples both with WpLa-5 CFS and WP, we observed that bacterial cells occupied a wider zone and they had moved to the bottom of the motility agar (Figure 5.4, lane 5-7). This suggests that the cells from the two treated samples of ribotype 027 were more motile than cells in the control. Also, cells of ribotype 001 showed a spreading pattern around the original zone but its two treated samples showed slightly less motility compared with treated cultures of ribotype 027.
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1 2 3 4 5 6 7 8 9 10
Motility
C. difficile Control WpLa-5 Wp Strains
Ribotype 027 Positive with significant Positive with significant Positive with significant movement to the bottom movement to the bottom movement to the bottom
Ribotype 078 Negative remained in the Negative remained in the Negative remained in the stabbed zone stabbed zone stabbed zone
Ribotype 001 Positive with significant positive Negative movement to the bottom Insignificant movement Insignificant movement
Figure 5.4. Effect of La-5 CFS on motility of C. difficile strains. Stabs of C. difficile strains were performed in BHI containing 0.175% agar. Motility of the strains was visible in agar stabs. 1) BHI agar; 2) Ribotype 078; 3) co-cultured 078 and La-5 CFS; 4) co-cultured 078 and Wp; 5) Ribotype 027, 6) co- cultured 027 and WpLa-5 CFS; 7) co-cultured 027 and WP; 8) Ribotype 001; 9) co-cultured 001 with WpLa-5 CFS; 10) co-cultured 001 and WP. Black arrows indicates diffusion of C. difficile cells to the bottom of their stab zone.
5.2.3 Effect of Proteobiotics on Sporulation
Sporulation plays a vital role for C. difficile to survive and spread in the environment and spores are produced in nutrient limited conditions. To determine whether proteobiotics inhibit spore production in C. difficile a sporulation assay using heat treatment was performed. We sought to add WpLa-5 CFS or WP to
C. difficile culture and let the culture grow and become mature for five days (Semenyuk et al., 2014) in order to examine sporulation rate. Three strains of C. difficile ribotype 027, 078, and 001 were tested.
129
Results revealed that all three strains produced spores in the presence and absence of treatments, but their sporulation rates were different (Figure 5.5). First a one-way ANOVA analysis using Sidak’s multiple comparison test revealed that WpLa-5 and WP increased total count of ribotype 027 (0.95 and 1.03 log respectively, P < 0.0001) compared to their control. Also, there was a one log10 cycle increase on count in the treated sample 001 La-5, P < 0.0001 and 2.1 log10 cycle increase (P < 0.0001) in total count of
001Wp compared to their control ribotype 001 total count. In contrast, for the 078 La-5 sample, the total count was decreased by about 0.93 log10 CFU/mL (P < 0.0001), but WP 078 did not show any significant difference from its control culture.
One-way ANOVA, Sidak analysis also compared each ribotype control culture and its two treated samples for number of spores remaining in the culture after heat treatment. For ribotype 027 both treatments resulted in an increase in total count before heat treatment, their spore count also was significantly higher than for the control culture (by 1.6 log10 CFU/mL for each treatment, P < 0.0001). The number of spores of ribotype
001 was not significantly different between treatments, but spore numbers of ribotype 078were significantly decreased in treated samples, 0.8 log10 cycle decrease in 078 La-5, and 0.6 log10 cycle decrease for 078 WP
(P < 0.0001). On further analysis, it was determined that numbers of vegetative cells that did not sporulate was significantly higher for ribotype 001 and 078 treated with proteobiotics (P < 0.0001) when compared to ribotype 027 treatments, for which all of the bacteria transformed to spores.
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T o ta l C o u n t S p o r e
Figure 5.5. Impact of La-5 proteobiotics on spore production of three C. difficile ribotypes 027, 078, and 001. Black bars represent C. difficile total count CFU/ mL of culture. Grey bars represent heat-resistant C. difficile spores. La-5 represent WpLa-5 CFS treatment and WP used as control treatment. Bars represent means and standard deviation of three biological replicates. Statistically significant differences (P < 0.0001 and P < 0.05) were measured by one-way ANOVA, Tukey’s multiple comparison test. ** and *** represent very significant differences and **** represent extremely significant differences between bars. ns represents not significant differences between bars.
5.3 DISCUSSION
The opportunistic pathogenic bacteria possess biological behaviors that help them to stablish in their host’s gastrointestinal tract, or to make them resistant to any external stress and aid them survive in nutrient deprived environment. Bacterial cells aggregate and adhere to living cells, including human epithelium, or non-living surfaces and form biofilms. Cells in biofilms are more resistant to antibiotics and biofilm
131 formation is one of the reasons for the persistence of chronic infections (Whiteley et al., 2001). Our current study examines the different biological behaviors of three highly infectious strains of C. difficile, including biofilm formation, motility, and spore production in the presence of bioactive compound extracted from L. acidophilus La-5 culture (WpLa-5 CFS). First, we show that C. difficile cultures that grow in the presence of WpLa-5 CFS failed to produce a mature biofilm. These C. difficile cultures were more visible as planktonic cells and exhibited less aggregative behavior. This study also showed that bacteria that are in biofilm growth proliferate less than when they are in planktonic growth; this finding is supported by Bester et al. (2011) who showed that encapsulated bacterial cells in biofilm have a slower metabolism than free floating bacteria.
As our earlier study revealed, L. acidophilus La-5 produces quorum sensing inhibitors. Bacteria cooperate with each other to build biofilms through cell-to-cell communication or quorum sensing. In Gram-positive bacteria, quorum sensing regulates dispersal and maintenance of biofilms, instead of regulating biofilm formation (Pratten et al., 2001; Yarwood et al., 2004). Therefore, quorum sensing and regulatory factors involved in biofilm-induced infections are of great interest for finding antimicrobial compounds against pathogenic bacteria (del Pozo & Patel 2007; Davies 2003; Dapa et al., 2013; Recsei et al., 1986; Janzon &
Arvidson, 1990; Abdelnour et al., 1993; Kong et al., 2006).
Another study showed that heptapeptide RNA III inhibiting peptide (RIP) inhibits pathogenesis of drug- resistant Staphylococcus aureus by interfering with its quorum sensing system; RIP also disrupts biofilm formation of S. aureus in a rat model and therefore, it was concluded that RIP with a synergistic activity with antibiotic inhibits biofilm formation and infection caused by drug-resistant Staphylococcus aureus
(Kong et al., 2006). However, it is assumed that RIP does not target the quorum sensing system directly and its underlying mechanism remains still unknown (Giacometti et al., 2003).
Other anti-biofilm agents are nucleotide biosysnthesis inhibitors. For instance, biosynthesis of cyclic di- guanosyl-5’ monophosphate (C-di-GMP) nucleotides are pivotal signaling molecules for biofilm regulation
(Landini et al., 2010) and a commercially available antibiotic named sulfathiazole, inhibits C-di-GMP
132 nucleotide biosynthesis and prevents biofilm formation. Thus, C-di-GMP is a signaling molecule that controls bacterial lifestyle choices. This molecule also controls the transition between planktonic and biofilm growth. One study showed that high level of C-di-GMP increases biofilm formation, and low level of C-di-GMP has a relationship with motility (Antoniani et al., 2010; Simm et al., 2004). C-di-GMP elevation strongly represses toxin gene expression by C. difficile by inhibiting tcdR (toxin gene sigma factor) expression and it also is able to inhibit motility of the bacterium by inhibiting sigD (flagella regulator) (McKee et al., 2013).
Conventional antibiotics prevent bacterial cell division or kill the bacterium, but evidence shows that they damage normal microbiota, which facilitates reinfection and increase numbers of antibiotic-resistant bacteria (Maria et al., 2014). Therefore strategies that can prevent biofilm formation without killing bacteria and that target different stages of developing biofilms are important (Maria et al., 2014). Anti-adhesion agents are also another strategy that could be a potential field of research as they prevent the main stage of biofilm formation, bacterial adherence. Studies revealed that using anti-adhesion compounds such as anti- pili that interfere with pilin subunits released from the bacterial cells inhibited biofilm formation in vitro
(Kline et al., 2009).
Motility of pathogens including C. difficile is a potential factor for colonization and adherence to mucus and epithelial cells in the gastrointestinal tract (Janoir, 2016; Baban et al., 2013;Tasteyre et al., 2000; Shrout et al., 2006; Logan, 2006; Borriello, 1998a). Our study, using motility agar to determine motility of C. difficile strains resulted in different outcomes. In contrast to ribotype 078 controls and its two treated cultures that failed to exhibit a spreading pattern around the stabbing location, two other ribotypes (027 and
001) and their treated cultures produced noticeable diffuse growth away from the inoculum stab into their cultures. Looking at results of the motility agars suggests that WP or WpLa-5 CFS does not change C. difficile from non-motile to motile phenotype (ribotype 078), but if the bacterial strains are motile, including ribotype 027 and 001, then their treated cultures also show motility. However we can see a slight decrease in motility of the ribotype 001 co-cultured with WP and a slight increase in motility of ribotype 027 treated
133 with WpLa-5 CFS and we can conclude that effectiveness of bioactive compound on C. difficile can be strain dependent.
As studies revealed, C. difficile is able to manifest peritrichous flagella with a higher level of adherence to the mouse cecum; being 10 times higher than non-flagellated strains (Tasteyre., 2001). Two genomic microarray studies proposed that virulence of the pathogen may not require motility and that is because flagellar coding content among the strains are diverse (Janvilisri et al., 2009; Stabler et al., 2006). However, motility is still considered a key player in colonization during infection due to the relationship between the product of motility locus and other surface associated proteins during infection (Twine et al., 2009).
Baban et al., (2013) showed that the importance of motility and flagellum to adherence and colonization, varies between different strains of C. difficile, for instance, their study indicated that a flagellum is not necessary for clinical strain 630, since 630∆erm exhibited successful adherence and colonization with paralyzed flagellum and mutation of fliC, fliD and flgE in vivo and in vitro without any flagella. Contrary to that, they demonstrated that in a hypervirulent strain R20291, flagella play a role in colonization and adherence. They strongly claimed that the importance of motility and flagellum is different amongst different strains and characterizing more than one strain is necessary before making general conclusions about specific pathogenesis mechanism.
Additionally, C. difficile lives on surfaces in the environment or in the mammalian GIT in the form of resistant spores and what makes them one of the risk factors of CDI is when contamination occurs and spores transmitted through the oral-fecal route germinate inside their host. In this study we showed that C. difficile sporulation is affected by proteobiotics for ribotype 001 and 078, but not much change was observed in ribotype 027; meaning that either the effect of proteobiotics is strain dependent or the drug resistant strain, ribotype 027 is more resistant to biological perturbation than the other two strains. It can also be concluded that their response to environmental signals might be different between the strains.
Ribotype 027 total counts before and after heat shock were not significantly different which it indicated
134 high sporulation of this hypervirulence strain compared to other two strains tested in this study and it supported other studies which have claimed this phenomenon before (Vohra & Poxton, 2010).
Once C. difficile is established in its suitable location with disrupted normal microbiota, it colonizes and produces toxin A and B. There are controversial debates that sporulation and toxin production are related to each-other for survival of this organism. Kamiya et al., (1992) suggest that these two mechanisms are not alternate mechanisms to aid C. difficile survival. Other studies indicated that the master regulator,
Spo0A, negatively regulates toxin A and B production in epidemic strain ribotype 027 only (Mackin et al.,
2013; Rosenbusch et al., 2012; Pettit et al., 2014), while Vohra and Poxton (2010) reported that 12 different isolates of ribotype 027 produce high levels of both toxins and spores. Vohra and Poxton (2010) also suggested that if the spo0A gene expresses during the early growth phase, it increases expression of other genes encoding spores and therefore, a higher number of spores will be produced later by the bacterium in their environment.
Conclusion
Bioactive molecules secreted by L. acidophilus culture in concentrated whey powder, named proteobiotics, had an effective inhibitory impact on C. difficile biofilm formation. Proteobiotics prevented bacterial cell- to-surface and cell-to-cell attachment significantly in all three strains. Moreover, results of the sporulation assay showed that the impact of proteobiotics differed between the C. difficile ribotype 027 and two other ribotypes. In addition, motility of the C. difficile is strain dependent. Proteobiotics did not change the natural behavior of C. difficile from non-motile to motile bacteria or vice versa, but if they are motile, it might increase or decrease their motility depending on strain. Therefore, L. acidophilus La-5 culture produces anti-adhesion and anti-quorum sensing compounds which can be biofilm inhibitors. Further research is
135 required to use purified proteobiotics and determine if the effect is exclusively because of the bioactive peptide produced or released by L. acidophilus La-5, or whether medium components are also involved.
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CHAPTER 6
6. GENERAL DISCUSSION
The aim of this study was to investigate and identify antivirulence impact of L. acidophilus La-5 biologically active substances on virulence factors of three clinically important strains of C. difficile, ribotypes 027, 078, and 001. Also, we tried to improve our understanding of proteobiotics and their inhibitory effects on biological activities that are important for pathogenicity of C. difficile. The objectives were to 1) investigate the influence of lactic acid bactria (LAB) cell free supernatant (CFS) on C. difficile growth and viability; 2) evaluate the influence of CFS of selected LAB strain L. acidophilus La-5 on quorum sensing of C. difficile using autoinducer bioluminescence (AI-2) assay; 3) comparison of expression of virulence genes of C. difficile particularly the expression of the toxin genes (tcdA, tcdB) and non-toxin genes of three C. difficile strains with and without La-5 CFS treatment; 4) evaluate the effect of La-5 CFS on colonization and toxin production of C. difficile in vitro, by performing adhesion and cytotoxicity assays; 5) visualize the impact of La-5 CFS on reducing efficacy of toxins to disrupt mammalian cell integrity; 6) observe and compare the effectiveness of La-5 CFS on biofilm formation and sporulation of
C. difficile after exposing to La-5 CFS.
The preliminary study showed that our selected probiotic strain L. acidophilus La-5 was well adapted to both media among several tested probiotics, although different growth patterns were observed in each medium. The differences between growth was due to the composition of the two media, and bacteria required different lag times to adjust their metabolism to each medium (Griffiths &Tellez, 2013; Karlsson et al. 1999). The analysis of C. difficile growth confirmed that La-5 CFS was not inhibitory for all three strains of C. difficile after 48 h co-incubation, but it postponed onset of exponential phase in treated cultures due to the adaptation time that bacteria needed when they were exposed to La-5 CFS in their growth media.
Therefore, this preliminary finding leads us to conclude that instead of an antibacterial effect, there would be antivirulence impact of the La-5 CFS, that is a strategy postulated by Rasko & Sperandio (2010).
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In Chapter three of this thesis the effect of La-5 bioactive extract was studied on quorum sensing, which is known to play a role in virulence of several pathogens, including C. difficile (Nealson & Hastings, 1979;
Taga & Bassler, 2003). First, we examined different LAB strains which are well documented for their antivirulence activities for their effect on autoinducer 2 (AI-2) production using Vibrio harveyi BB170 as a bioluminescent reporter for the presence of AI-2. Data showed that bioluminescence of V. harveyi was induced by C. difficile culture filtrate; indicating the production of AI-2. However, autoinducer signaling was decreased in most treated samples and particularly following La-5 CFS treatment, due to its inhibitory impact on cell-to-cell communication of C. difficile bacterial cells and the inhibitory impact was clearly effective for all three ribotypes. Then, to support other studies and to investigate if there is a link between quorum sensing and C. difficile gene expression, and also to examine the antivirulence impact of La-5 CFS by reducing transcription of toxin and luxS genes, we performed real-time q-PCR using three ribotypes 027,
078, and 001.
Gene expression analysis indicated that transcription levels of luxS gene that is involved in quorum sensing showed significant down regulation, and these results were in agreement with previous studies conducted by others (Yun et al., 2014). At the same time, toxin genes tcdA and tcdB and their positive regulator, tcdR were affected by La-5 CFS with statistically significant down regulation in all three strains except for tcdR in ribotype 001. Another notable finding of our study was that gene Cwp84 (one of the adhesion genes important for colonization and attachment of pathogens to their targeted cells), was significantly down regulated in all three strains. These findings lead us to conclude that there is a relationship between transcription of different virulence genes in the presence of La-5 bioactive extract and it prompted us to investigate changes of adherence and cytotoxicity of C. difficile on human epithelial cells in the presence of La-5 CFS containing bioactive molecules.
In Chapter four, cytotoxicity assays indicated that coincubating C. difficile and La-5 CFS substantially reduced toxin damage to HT-29 and Caco-2 cell lines (human epithelial monolayer). Cytotoxicity results
138 showed significant reduction in dead cells; and cytopathic assay using light microscopic images, confirmed inhibition of cell rounding damage from C. difficile culture filtrate pretreated with La-5 CFS. This impact could have been by reducing toxin production through repressing toxin regulators, or interfering with toxin function and toxin delivery of C. difficile (Kaufmann et al., 2008; Rasko & Sperandio, 2010).
Downregulation of Cwp84 gene encoding cysteine protease enzyme might be another determinant for reducing cytotoxicity in epithelial cells. We also, compared whey protein treatments (WP and WpLa-5
CFS) with La-5 CFS prepared from modified MRS. From the analysis of images, we concluded that WpLa-
5 CFS compared to La-5 CFS was more protective against cytotoxicity caused by C. difficile culture filtrate.
Reducing cytotoxicity could be related to quorum sensing. Quorum sensing could be inhibited by interfering with recognition and then transcription response to the signaling molecules, LuxR homologues (Hentzer et al., 2002; Hentzer et al., 2003). Therefore, by disruption of quorum sensing, virulence genes expression which is activated by accumulation of quorum signaling molecules, and consequently, toxin production is prevented. Zhang and Dong, 2003 indicated mechanisms are interfering Gram-positive quorum sensing system. The two-component sensor and response regulator pair, autoinducing peptides (AIP) and AgrCA can be interrupted by receptor competition for signaling molecules or by inhibition of sensor kinase. Also, interfering with quorum sensing could be through inhibiting the production of signaling molecules.
Inactivation of quorum sensing locus including agrA (response gene) (Darkoh et al., 2015) or quorum sensing encoding genes including luxS, can decrease signaling molecules and toxins production in C. difficile ribotype 027 (Martin-Verstraete et al., 2016).
Traditional antibacterial compounds inhibit DNA or RNA synthesis, protein synthesis, cell wall synthesis, folate synthesis or depolarizes membrane potential. On the contrary, antivirulence or virulence inhibitors mostly target virulence gene regulation through quorum sensing, and/or toxin gene regulation. In addition, toxin delivery can be targeted such as achieved by inhibiting bacterial systems including type II and type
III secretion systems, which consequently prevents translocation of effector molecules (Kauppi et al.,
139
2003). Therefore, interference of toxin delivery of toxins to their site of action when small molecules bind to C. difficile toxins or to the receptors in their target cells inhibits their delivery to intestinal epithelial cells and decreases the effect of toxins (King and Barriere, 1981).
We also studied adhesion of C. difficile, which is an important virulence factor and is the prerequisite of colonization and commencement of infection. It was found that pretreatment of HT-29 cell monolayer with
La-5 CFS was prevented C. difficile adherence to the cell line efficiently. Chapter four findings were consistent with those described in Chapter three as both chapters documented studies on important virulence factors of toxigenic strain of C. difficile ribotype 027. The results indicated that when C. difficile preincubated with La-5 proteobiotics in their cultures or when they were exposed to proteobiotics in the target cell culture for infection challenges, their adherence is affected in both situation, and that confirms an antivirulence impact of proteobiotics. According to other studies, a pathogen’s ability to adhere and form biofilm make it able to colonize and establish an infection (Nobbs et al., 2009; Allsopp et al., 2010). This led us to further study biofilm formation by C. difficile when the pathogen was coincubated with La-5 proteobiotics.
In Chapter five, we assessed biofilm growth of three ribotypes of C. difficile 027, 078, and 001 with and without treatment with La-5 proteobiotics presented as WpLa-5 CFS. Scanning electron microscopy (SEM) was used to visualize the impact of treatments on biofilm formation. WpLa-5 CFS reduced attachment and population of C. difficile cells encased in the biofilm but increased non-aggregated planktonic cells in the medium significantly (P < 0.001). SEM analysis was consistent with the quantitative data. From the total count data and sporulation assay, we found that nutritional composition of whey protein triggered growth of C. difficile. Their population increased in co-cultured samples and the SEM images showed that biofilm formation of treated cultured was remarkably reduced in treated samples. That could be due to availability of nutrients and consequently higher metabolic activity of the pathogen, which did not allow them to transform into sessile aggregated cells and subsequently biofilm.
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Lactic acid is one of the antibacterial factors produced when probiotics are present in the gut, but in our study we used cell free culture supernatant with a neutral pH, to prevent antibacterial effect of lactic acid from causing misleading results. We excluded glucose from the m-MRS medium and replaced it with sucrose to eliminate the effect of rapidly metabolizable sugar or pH changes on C. difficile tcdR gene transcription or toxin production (Nagraj Mani et al., 2002). Likewise, studies proved that nutritional signals, including carbon sources or specific amino acids, are very important environmental factors that influence toxin production (Dupuy & Sonenshein, 1998; Yamakawa et al., 1996). That is because rapidly metabolizable carbon sources mostly enter bacterial cells, which invokes the carbon catabolite repression
(CCR) mechanism. The CcpA regulon (a global regulator of carbon metabolism in Gram-positive bacteria)
(Antunes et al., 2012) is a link between the carbon and nitrogen pathway and also virulence gene expression in C. difficile (Dupuy & Sonenshein, 1998). CcpA directly interacts with the promoter region upstream of the tcdR regulator and affects several PaLoc genes. This leads us to conclude that C. difficile toxin production in response to accessibility of glucose and nitrogen involves a complicated regulatory network with the CcpA and codY as its key regulators (Martin-Verstraete et al., 2016).
According to literature, other studies also reported that protein fraction and some amino acids such as proline, glycine, hydroxyproline, leucine, isoleucine, and alanine have an inhibiting impact on toxin yield by C. difficile (Bouillaut et al., 2013; Karlsson et al., 1999; Yamakawa et al., 1998). Studies by Dupuy et al., (1998) and Hundsberger et al., (1997) established that various environmental changes affect the level of toxin production in C. difficile, with nutritional signals among the most influential impacts from the environment.
Cytotoxicity and biofilm results confirmed that whey protein treatments (WpLa-5 and WP) resulted in effective protection for epithelial cell monolayer against C. difficile infection. One of the limitations of our study was that unfortunately, the effect of WpLa-5 CFS on the regulation of virulence genes by real-time
PCR was not examined due to the challenges that we faced for RNA extraction from the treated samples.
Seemingly, whole culture were sporulating, which may have frustrated attempts to extract RNA. It is
141 predicted that high protein content of whey protein would affect toxin gene expression due to evidence from other studies.
Conclusion
In conclusion, our findings suggest that Lactobacillus acidophilus La-5 bioactive extract from whey protein concentrate (proteobiotics) is relatively crude biological mixture produced from a food grade bacterium that showed some favorable in vitro characteristics. It is deserving of more research to show that they are likely could be applied in human food supplements for better health and to be used in healthcare to prevent and reduce reinfection in CDI patients.
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CHAPTER 7
FUTURE DIRECTION
1) Application of current research for C. difficile reinfection
Development of highly virulent strains of C. difficile caused an increasing trend in recurrent Clostridium difficile infection (RCDI) (Warny et al., 2005) with more than 20% chance of developing reinfection after the first episode of CDI and 40-65% chance of recurrence after the first recurrence (McFarland et al., 1999).
Treating the recurrences is more challenging than the first infection and clinical manifestation are more severe. Current treatments for RCDI is metronidazole for moderate infection and vancomycin for severe cases of CDI. Metronidazole has a neurotoxicity and hepatic toxicity effects and is the cause of 14%-27% recurrences. Therefore, subsequent recurrences need to be treated with a different antibiotic such as vancomycin administered orally and for a more extended period which is the cause of 7% to 23% recurrences. Some of the alternatives are Fidaxomycin, fecal bacteriotherapy, and probiotics which there are limits in their application too. Probiotic strains including Sacharomyces bulardii and Lactobacillus helveticus GG have been used as an adjunct therapy with antibiotic to treat recurrences but yet there are no effective alternative to prevent RCDI (McFarland et al., 2005). Restoration of intestinal microbiota is a critical approach for preventing reinfection in CDI patients, and to speed the healing process.
Proteobiotics showed effective anti-biofilm impact against C. difficile as we described in Chapter five of this study. We also observed the significant protective effect of the bioactive on human epithelial cell monolayer in Chapter 4. They reduced cytotoxicity of C. difficile culture filtrate very efficiently. Also, a very recent research by Nordeste and colleague (2017), confirmed the anti-infective effect of proteobiotics to prevent the enterogenic E. coli (ETEC) infection in pigs. They fed the piglets with different concentration of bioactive extract from L. acidophilus La-5 before the infection challenge with E. coli K88. Feeding with a low and high dose of proteobiotics significantly decreased infection symptoms. These proteobiotics also improved gut health by reducing colonization of E. coli K88 in piglets. Their research suggested using this bioactive will reduce antibiotic usage on the farm animals. Also, evidence shows bioactive molecules are
143 known to maintain mucus of epithelial cells and also can protect epithelial cells and their tight junctions, and they have the advantage of not being live microorganism to cause any problem for immunocompromised patients (to cause the blood stream infections). Therefore, application of proteobiotics in human possess multi-function for improving the health of gut in healthy or low-risk individuals and an potentially effective therapy to reduce or prevent RCDI. Bioactive prepared from metabolic activity of a well-documented Lactobacillus acidophilus strains in a whey protein-based media
(Yun et al., 2014; Nordeste et al., 2017), deserve to be considered to be studied in CDI recurrences in hospitals. It could be prepared as a dietary supplement or a yogurt-like product containing different concentration of proteobiotics. A pilot study is suggested to administer WpLa-5 CFS (in a liquid form), to the CDI treated patients orally during their antibiotic therapy or after that for 2 to 8 weeks every day, 100 mL from an optimum concentration. A concentration of 1.5- 3 mg/mL was the optimum level to be used in in vitro study (higher dose about 10-20 mg/mL might be more effective inside the gut because of presence of other substances, and liquids inside the GI tract that dilute the extract). Population of fecal C. difficile concentration as well as other gut microbiota needs to be monitored. The study must focus on high-risk individuals such as hospitalized patients with advanced age. Also it is suggested for both male and female hospitalized patient with surgery, longer stay in intensive care unit which generally they become carrier of
C. difficile after 4 week of hospitalization even without antibiotic therapy. Safety, effectiveness and tolerance of the supplement must be recorded as a self-diary by the patient every day in outpatients and by the nurses for the inpatients. Frequency of the stool and form of the stool need to be reported too.
The aim of study also could be the investigation of the influence of proteobiotics to recover and maintain the disturbed microbiota of the gut after C. difficile infection. Probiotic bacteria and their secreted factors are not only effective to change the microbial flora of the gastrointestinal tract, but also they can be beneficial to maintain the gut homeostasis. To identify recovery of bacteria from disturbed microbiota of
C. difficile infected hosts RNA of fecal samples will be examined using next-generation based 16S ribosomal RNA (rRNA) sequencing. It will identify different strains of divergent samples. Data will be interpreted using 16S metagenomics an Illumina-curated taxonomic database. 144
2) Mechanism of Action of Proteobiotics: Another future direction that is suggested is further study on mechanism of action of characterized sequences of proteobiotics on toxin production and biofilm inhibition.
It has been well documented that probiotic LAB improve health and help prevent disease, and they can be a substitute for antibiotics but there is limited knowledge about their mechanism of action and how they exert their beneficial effects. The fact that probiotics are able to interact with the intestinal mucus, thereby improving the immune system while acting as a barrier along with production of antibacterial and antivirulence compounds are among the most interesting fields of research. An in vivo study measuring the
IgA levels in the intestine of an animal model to investigate anti-toxin secretory after their proteobiotics intake and infection with C. difficile. Examining proteases that possibly produced by the L. acidophilus in their supernatant and identifying their effect on degrading C. difficile toxins A and B, or degrading toxin receptors in the colon also will be worth trying.
3) Bioactives and Regulatory Network of pathogens: Another future research idea could be investigation of possible link between proteobiotics and the regulatory network of C. difficile that controls virulence of the bacterium including cyclic di-guanosyl-5’monophosphate (C-di-GMP), a second messenger that controls transition of bacterial life style from planktonic to biofilm growth.
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