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Doctoral Dissertations University of Connecticut Graduate School
1-14-2019 Natural Strategies for Controlling Virulence and Antibiotic Resistance in Clostridium difficile Abraham Joseph Pellissery University of Connecticut - Storrs, [email protected]
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Recommended Citation Pellissery, Abraham Joseph, "Natural Strategies for Controlling Virulence and Antibiotic Resistance in Clostridium difficile" (2019). Doctoral Dissertations. 2036. https://opencommons.uconn.edu/dissertations/2036 Natural Strategies for Controlling Virulence and Antibiotic Resistance in Clostridium
difficile
Abraham Joseph Pellissery, PhD
University of Connecticut, 2019
Clostridium difficile is a significant enteric pathogen causing a toxin-mediated infection and diarrhea in humans. There has been an increased incidence of C. difficile infection (CDI) in the
United States with the emergence of hypervirulent strains and community associated outbreaks.
CDI is commonly observed among hospital in-patients undergoing protracted antibiotic therapy, which results in gastrointestinal dysbiosis, creating a conducive environment for spore germination, pathogen colonization in the gut, and subsequent toxin production. An ideal anti-
C.difficile therapeutic agent should inhibit critical virulence factors of the pathogen such as toxin production, sporulation and spore germination without inducing gut dysbiosis. Such agents when provided as an adjunct to C. difficile antibiotic therapy could help to improve the clinical outcome of CDI and prevent the relapse of the infection. In this Ph.D. research, the efficacy of three alternative treatment approaches as antivirulence agents was tested for potential future development as therapies against CDI. This included sodium selenite (metalloid), baicalin (flavone glycoside), and selected lactic acid bacteria. All treatment modalities were tested for inhibiting toxin production, sporulation and spore outgrowth in two hypervirulent C. difficile isolates.
Moreover, gene expression and cell culture studies were performed to elucidate the anti-toxigenic mechanism of sodium selenite and baicalin. In addition, the effect of sodium selenite in increasing pathogen sensitivity to ciprofloxacin and vancomycin, two common antibiotics used in treating C. difficile, was also tested. Furthermore, the effect of baicalin on CDI was investigated in a mouse model, with special reference to its effect on disease severity and the mouse gut microbiome. The Abraham Joseph Pellissery – University of Connecticut, 2019 results revealed that sub-minimum inhibitory concentration (sub-MIC) and sub-inhibitory concentrations (SIC) of sodium selenite and baicalin, respectively, reduced C. difficile toxin production and cytotoxicity in vitro. In addition, sodium selenite and baicalin inhibited spore outgrowth. Oral supplementation of baicalin improved the clinical outcome in challenged mice, and positively altered the gut microbiome composition. Collectively, these results indicate that the three approaches identified in this study significantly reduced C. difficile virulence, however, follow up validation in animal models for long-term safety and dose standardization, and clinical trials in human subjects are necessary.
Natural Strategies for Controlling Virulence and Antibiotic Resistance in Clostridium
difficile
Abraham Joseph Pellissery
B.V.Sc. & A.H., Kerala Agricultural University, 2006
M.V.Sc. Indian Veterinary Research Institute, 2009
A Dissertation
Submitted in Partial Fulfillment of the
Requirements for the Degree of
Doctor of Philosophy
at the
University of Connecticut
2019 Copyright by
Abraham Joseph Pellissery
2019
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APPROVAL PAGE
Doctor of Philosophy Dissertation
Natural Strategies for Controlling Virulence and Antibiotic Resistance in Clostridium
difficile
Presented by
Abraham Joseph Pellissery, B.V.Sc. & A.H., M.V.Sc.
Major Advisor……………………………………………………………….. Dr. Kumar Venkitanarayanan
Associate Advisor………………………………………………………………. Dr. Mary Anne Roshni Amalaradjou
Associate Advisor………………………………………………………………. Dr. Dennis J. D’Amico
Associate Advisor………………………………………………………………. Dr. Mazhar I. Khan
Associate Advisor………………………………………………………………. Dr. Bhushan M. Jayarao
University of Connecticut 2019
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ACKNOWLEDGEMENTS I would like to express my gratitude and immense respect to my major advisor, Dr. Kumar
Venkitanarayanan, for his continual guidance and support throughout my tenure as a graduate student. I consider myself so blessed and privileged to have got the opportunity to work under the able tutelage of such an enthusiastic and hard-working researcher. The experiences that I gained from his lab helped me to evolve as an independent researcher and a critical theorist in the microbial sciences.
I would like to express my sincere gratitude to my associate advisors, Dr. Mary Anne
Amalaradjou, Dr. Dennis D’Amico, Dr. Mazhar Khan and Dr. Bhushan Jayarao for their continuous encouragement, critical advice and suggestions throughout my doctoral research. I would also like to extend my gratitude to all the Faculty, graduate students and administrative staff of the UConn Animal Science department for all the help and support provided to me during my life here as a doctoral student.
My sincerest and profound gratitude to my dear friends: Ajith, Shafeekh, Sambhu, Shaan,
Bindu, Elza, Deepa and Poonam for your camaraderie during the sunny and rainy days of my student life at UConn.
Words are not enough to express my gratitude to my mother, Mrs. Mary Joseph and my parents-in-law, Mr. Anto E.V. and Mrs. Lissy Varghese for their unfailing support and prayers when I needed it the most. I remember and thank my father, Late Mr. P. C. Joseph for all the love, care, support and blessings that he has provided me I am sure that my father would be proud of me on my achievement up in heaven. I am grateful to my siblings James and Teresa, my in-laws Tina,
Annlyn, Manu, Sharon and Arun, and my nieces and nephew, Sarah, Rahael and Gabriel: for all their heartwarming kindness and support.
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I would like to express my deepest love and thanks to my daughters Annarita and Helenna for their laughter, naughtiness, kisses and hugs that magically made my rough days much better. I also want to thank my wife Liya, for her wonderful companionship, understanding, patience and love throughout the last eight years.
Lastly, I thank the Almighty for strengthening me to successfully complete my endeavor as a doctoral student and for all the blessings showered upon me throughout my life.
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TABLE OF CONTENTS Approval page…………………………………………………………………………... iii
Acknowledgments…………………………..………………………………………..... iv
Table of Contents……………………………………………………………………….. vi
List of figures…………………………………………………………………………… xii
List of tables……………………………………………………………………………. xv
List of abbreviations……………………………………………………………………. xvi
Chapter I: Introduction………………………………………………………………. 1
Chapter II: Review of literature……………………………………………………… 7
1. Clostridium difficile or Clostridioides difficile (CD)…………………………....……… 8
1.1.Epidemiology of CDI…………………………………...………………………………… 10
1.2.Clinical manifestations and risk factors associated with CDI in humans…..…….. 14
1.3.Pathogenesis of CDI………………………………..…………………………..………... 15
1.4.C. difficile toxins…………………………………………………………………….……. 17
1.5. Non-toxin associated virulence factors initiating host colonization..…………… 20
1.6. C. difficile sporulation ………………………………………………………………… 21
1.7. C. difficile spore germination………….……………………………………………… 23
1.8. Influence of gut microbiome on CDI susceptibility.………………………………… 24
2. Antimicrobial resistance in C. difficile…..……………………………………………… 26
3. Treatment approaches for human CDI…………………………………………………… 27
3.1. Current and novel antimicrobials for CDI…….…………………………….….. 29
3.2. Alternate and emerging treatment strategies for CDI..……………………………. 29
3.2.1. Immunization strategies…………………………………………………………... 29
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3.2.2. Fecal microbiota transplantation……………………………………………….. 30
3.2.3. Non-toxigenic strains of C. difficile……………………………………………... 30
3.2.4. Emerging strategies for CDI therapy….………………………………………… 31
4. Metals 32
5. Phytochemicals 34
6. Probiotic therapy 36
References………………………………………………………………………………………… 39
Chapter III: In vitro efficacy of sodium selenite on toxin production, spore outgrowth and antibiotic resistance in hypervirulent Clostridium difficile….…….. 60
Abstract…………………………………………………………………………………. 61
1. Introduction…………………………………………………………………………... 62
2. Materials and Methods……………………………………………………………….. 63
2.1. Bacterial isolates and culture conditions……..……………………………………... 63
2.2. Establishment of sub-minimum inhibitory concentration and minimum inhibitory concentration of sodium selenite and antibiotics ………………………………. 64
2.3. Effect of sodium selenite on C. difficile Toxin Production and Cytotoxicity…… 65
2.4. ELISA for Total Toxin A and B……………………………………………………….. 65
2.5. Cytotoxicity Assay………………………………………………………………………. 66
2.6. Real-Time Quantitative PCR (RT-qPCR)….………………………………………… 66
2.7. Effect of sodium selenite on C. difficile spore germination and outgrowth….….. 67
2.8. Effect of sodium selenite on antibiotic resistance in C. difficile….………………. 68
2.9. Statistical analysis…..………………………………………………………………….. 68
3. Results………………………………….………………………………………………….. 68
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3.1. Sub-minimum inhibitory concentrations and minimum inhibitory concentrations of sodium selenite and antibiotics…………………………...……………… 68
3.2. Effect of selenium on C. difficile toxin production…………………………………… 69
3.3. Effect of selenium on C. difficile toxin-mediated cytotoxicity on Vero cells……… 69
3.4. Effect of sodium selenite on toxin regulatory genes….……………………………… 69
3.5. Effect of sodium selenite on C. difficile spore germination and outgrowth………. 70
3.6. Effect of sodium selenite on C. difficile antibiotic resistance………………………. 70
4. Discussion……………………………………………………………………………. 71
References………………………………………………………………………………. 76
Chapter IV: Effect of Baicalin on C. difficile toxin production, sporulation and spore germination in vitro……………………………………………………………………. 88
Abstract 91
1. Introduction…………………………………………………………………………... 93
2. Materials and Methods……………………………………………………………….. 94
2.1. Bacterial strains and culture conditions……………………………………………… 94
2.2. Determination of sub-inhibitory concentration of baicalin…...…………………… 95
2.3. Effect of baicalin on C. difficile Toxin Production and Cytotoxicity……….…….. 96
2.4. ELISA for Total Toxin A and B……..…………………………………………………. 96
2.5. Cytotoxicity Assay……………………………………………………………………… 96
2.6. Effect of baicalin on C. difficile sporulation….…………………………………….. 97
2.7. Effect of baicalin on C. difficile spore germination and outgrowth.……………… 97
2.8. Real-Time Quantitative PCR (RT-qPCR)………………. …………………………… 98
2.9. Statistical analysis………………………………………….…………………………… 98
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3. Results ………………………………………………………………………………. 98
3.1. Sub-inhibitory concentration of baicalin and its effect on beneficial microbiota 98
3.2. Effect of baicalin on C. difficile toxin production..……………..………………….. 98
3.3. Effect of baicalin on C. difficile toxin-mediated cytotoxicity on Vero cells….….. 98
3.4. Effect of baicalin on C. difficile sporulation…………………………………………. 99
3.5. Effect of baicalin on C. difficile spore germination………………………………… 99
3.6. Effect of baicalin on toxin, sporulation and other virulence associated genes….. 99
4. Discussion……………………………………………………………………… 100
References………………………………………………………………………………. 103
Chapter V: Antivirulence effect of select lactic acid bacteria against Clostridium difficile ……………………………………………..………………………………….. 115
Abstract…………………………………………………………………………………. 116
1. Introduction…………………………………………………………………………... 117
2. Materials and Methods……………………………………………………………….. 119
2.1. Bacterial isolates and culture conditions…………………………………………….. 119
2.2. Effect of co-culturing of select LAB with C. difficile on toxin production..…….. 120
2.3. ELISA for Total Toxin A and B ……………………………………………………….. 121
2.4. Cytotoxicity assay………………………………………………………………….……. 121
2.5. In vitro co-culturing of selected LAB on C. difficile spore outgrowth….………… 122
2.6. Statistical analysis………………………………………………………………………. 122
3. Results………………………………………………………………………………... 122
3.1. In vitro co-culturing of selected LAB isolates with vegetative C. difficile……….. 122
3.2. Effect of LAB on C. difficile toxin production and cytotoxicity assay..………….. 123
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3.3. In vitro co-culturing of selected LAB on C. difficile spore outgrowth…………… 124
4. Discussion……………………………………………………………………………. 125
References………………………………………………………………………………. 128
Chapter VI: Effect of baicalin in reducing Clostridium difficile infection in a mouse model ………………..…………………………………………………………. 139
Abstract…………………………………………………………………………………. 140
1. Introduction…………………………………………………………………………... 141
2. Materials and Methods……………………………………………………………….. 143
2.1. Ethics statement, animals, and housing………………………………………………. 143
2.2. Prophylactic and therapeutic administration of baicalin in a mouse model of C. difficile infection……………...………………………………………………………………….. 143
2.3. DNA extraction, PCR amplification, and sequencing of taxonomic markers.….. 145
2.4. Sequence analysis……………………………………………………………………….. 145
2.5. Statistical analysis……………………….…………..………………………………… 146
3. Results ……………………………………………………………………………….. 146
3.1. Effect of baicalin supplementation on the incidence of diarrhea and severity of
C. difficile infection in mice ……..……………………………………………………………. 146
3.2. Effect of baicalin supplementation on clinical score and body weight of C. difficile infected mice ………………………………………..………………………………… 148
3.3. Effect of baicalin supplementation on the gut microbiome of C. difficile infected and non-infected mice ……………………………………………………..…………………… 149
4. Discussion……………………………………………………………………………. 151
References………………………………………………………………………………. 156
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Chapter VII: Summary………………………………………………………………. 176
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LIST OF FIGURES
Title Page
Chapter III
Figure 1 Effect of Sub-minimum inhibitory concentration of sodium selenite 77
on the growth of six beneficial bacteria
Figure 2 Effect of sodium selenite on C. difficile toxin production 78
Figure 3 Effect of sodium selenite (Se) on C. difficile induced cytotoxicity on 79
Vero cells
Figure 4 Effect of sodium selenite on C. difficile toxin regulatory genes 80
Figure 5 Effect of sodium selenite on germination and outgrowth of C. 81
difficile
Figure 6 Effect of sodium selenite on antibiotic sensitivity of C. difficile BAA 82
1870 (A) and 1803 (B) to 0.5 x MIC and MIC of ciprofloxacin
Figure 7 Effect of sodium selenite on antibiotic sensitivity of C. difficile BAA 83
1870 (A) and 1803 (B) to 0.5 x MIC and MIC of vancomycin
Chapter IV
Figure 1 Effect of sub-inhibitory concentration of baicalin on selected 103
beneficial gut bacteria growth
Figure 2 Effect of baicalin on C. difficile toxin production 104
Figure 3 Effect of baicalin (BC) on C. difficile induced cytotoxicity on Vero 105
cells
Figure 4 Effect of baicalin on C. difficile sporulation 106
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Figure 5 Effect of baicalin on germination and outgrowth of C. difficile 107
Figure 6 Effect of baicalin on C. difficile toxin regulatory genes 108
Figure 7 Effect of baicalin on C. difficile sporulation genes 109
Figure 8 Effect of baicalin on C. difficile secondary virulence associated genes 110
Chapter V
Figure 1 Effect of co-culturing vegetative CD with LAB on C. difficile growth 130
at 10, 24 and 48 h
Figure 2 LAB counts in co-culture experiment using vegetative CD at 10, 24 131
and 48 h
Figure 3 Effect of LAB on C. difficile toxin production 132
Figure 4 Effect of co-culture supernatants of C. difficile isolates BAA 1870 (A) 133
and BAA 1803 (B) on CD induced cytotoxicity on Vero cells
Figure 5 Effect of LAB isolates on spore outgrowth of C. difficile isolates 134
Chapter VI
Figure 1 Antibiotic induced murine CDI model 160
Figure 2 Effect of baicalin supplementation on the incidence of diarrhea in 161
mice after CDI
Figure 3 Effect of baicalin supplementation on the clinical severity of mice 163
after CDI
Figure 4 Effect of baicalin supplementation on relative weight loss in C. 165
difficile infected and non-infected mice
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Figure 5 Effect of baicalin supplementation on the abundance of major gut 167
microbiome taxa in the antibiotic treated and C. difficile challenged
mice
Figure 6 Effect of baicalin supplementation on the abundance of 168
Lactobacillaceae (5A), Lachnospriaceae (6A), Akkermansia (6A),
Enterobacteriaceae (6B) and Peptostreptococcaceae 65B) in the
antibiotic treated and C. difficile challenged mice
Figure 7 Effect of baicalin supplementation on the diversity of gut microbiota 170
of antibiotic treated and C. difficile challenged mice – Inverse Simpson
plot
Figure 8 Effect of baicalin supplementation on the diversity of gut microbiota 171
of antibiotic treated and C. difficile challenged mice – Bray-Curtis plot
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LIST OF TABLES Title Page
Chapter V
Table 1 pH values in co-culture experiments when vegetative CD was 129
incubated with LAB isolates in 5 log: 5log inoculation ratio
Chapter VI
Table 1 Different treatment groups used in the experiment 157
Table 2a Mouse clincal score sheet 158
Table 2b Mouse body condition chart 159
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LIST OF ABBREVIATIONS
BHIS Brain Heart Infusion supplemented with 5g/L yeast extract CCFA Cycloserine, cefoxitin and fructose agar CD Clostridium difficile CDI Clostridium difficile infection CDC Centers for Disease Control and Prevention CDMN Clostridium difficile Moxalactam Norfloxacin medium CFU Colony forming Units BC Baicalin FDA Food and Drug Administration GRAS Generally recognized as safe LAB Lactic acid bacteria MRS de Mann Rogosa Sharpe OD Optical density PBS Phosphate buffered saline PYG Peptone, yeast, glucose RT-qPCR Real-time quantitative polymerase chain reaction SAS Statistical analysis software Se Sodium selenite SIC Sub-inhibitory concentration Sub-MIC Sub-minimum inhibitory concentration USDA United States Department of Agriculture
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CHAPTER I
Introduction
1
Clostridium difficile is a Gram-positive, spore-forming, anaerobic, hospital acquired pathogen causing a toxin-mediated enteric disease in humans, which commonly arises as a result of prolonged antibiotic therapy (Bartlett, 1997; Spigaglia, 2016; Weese, 2010). More than 500,000 cases of C. difficile infection (CDI) are reported annually in the United States, resulting in approximately $6.3 billion in healthcare costs (Zhang et al., 2016). Approximately 50% of C. difficile infections occur in people younger than 65, but more than 90% of deaths occur in people
65 and older. Prolonged antibiotic therapy results in the disruption of the normal enteric microbiota, leading to C. difficile spore germination and pathogen colonization in the intestine, with subsequent production of toxins (Bartlett, 1997; Dial et al., 2005). C. difficile toxins, TcdA and TcdB, are major virulence factors that disrupt the intestinal epithelial integrity, leading to an inflammatory response, causing pseudomembrane formation in the intestine and watery diarrhea
(Hookman & Barkin, 2009; Keel & Songer, 2006; McDonald et al., 2006; Sunenshine &
McDonald, 2006; von Eichel-Streiber et al., 1999). C. difficile also produces resilient spores that promote transmission through feco-oral route in patients and cause relapse in temporarily recovered patients (Burns et al. 2010). Therefore, therapeutic agents that can reduce C. difficile virulence, especially toxin production, sporulation as well as spore germination and outgrowth in the human gut would effectively help to control CDI.
The emergence of a hypervirulent strain of C. difficile, NAP1/ribotype 027 that produces increased levels of toxins and a severe form of the disease in humans was reported in the US
(Blossom & McDonald, 2007; Hookman & Barkin, 2009; Sunenshine & McDonald, 2006).
Despite the fact that broad-spectrum antibiotics predispose patients to CDI by disrupting the normal gut microbiota (Bartlett, 1992; O'Connor et al., 2004), antibiotics are the drug of choice for treating the disease in patients. Further, the emergence of antibiotic resistance in hypervirulent
2 strains of C. difficile has been reported worldwide, which further limits the success of antibiotic treatment (Prabaker & Weinstein, 2011; Spigaglia et al., 2011).
Recently, a new approach of targeting virulence of a pathogen for controlling infectious diseases has been widely explored. Since virulence factors of pathogens contribute to the establishment of infection in a host, inhibition of these factors could prevent disease progression
(Defoirdt, 2016; Khodaverdian et al., 2013; Rasko & Sperandio, 2010). Moreover, since this approach does not target pathogen growth, it presents a lesser selective pressure on development of bacterial resistance compared to traditional antimicrobial therapy (Cegelski et al., 2008;
Clatworthy et al., 2007; Hung et al., 2005; Mellbye & Schuster, 2011; Rasko & Sperandio, 2010).
Thus, a viable strategy for controlling C. difficile infection could be the use of drugs against the virulence factors of the pathogen. Since C. difficile toxins and sporulation are critically involved its pathogenesis and transmission, respectively, identification of therapeutic agents that inhibit these virulence traits without causing gut dysbiosis would potentially constitute a viable approach for controlling CDI.
Although a variety of metals have historically been used as antimicrobial agents, their application in human medicine and agriculture began to decline in the antibiotic era. However, with the emergence of multidrug-resistant pathogens and reduced number of antibiotics being discovered, metals have received increased attention in recent years, especially as potent antimicrobial agents against antibiotic-resistant pathogens. Selenium (Se) is a naturally occurring essential microelement critical for various biological functions in the body, including enzymatic and antioxidant activities. The supplemental form of selenium, sodium selenite (Na2SeO3) has proven antifungal and antibacterial activities (Kumar et al., 2010; Soriano-Garcia, 2004).
Published research from our laboratory indicated that sodium selenite decreased verotoxin
3 production in Escherichia coli O157:H7 and cholera toxin production by Vibrio cholera
(Bhattaram et al., 2017; Surendran-Nair et al., 2016). Similarly, previous research conducted in our laboratory revealed that sodium selenite increased the sensitivity of multi-drug resistant A. baumannii to ampicillin, tetracycline, ciprofloxacin, polymixin and imipenem (Surendran-Nair et al., 2016).
Phytochemicals represent another natural group of molecules that have been used for treating various diseases in traditional medicine (Wollenweber, 1988). Plant-derived flavonoids are naturally occurring phenylchromones known to exert a wide array of biological activities, including antiallergic, antimicrobial, antimutagenic, and antioxidant activities. Some flavonoids innately contribute to the plant’s antimicrobial defense systems and possess antagonistic activities against a wide range of other pathogenic microbes (Cowan, 1999; Middleton, 1994; Tringali,
2003). Baicalin (5,6-dihydroxy-7-O-glucuronide flavone) is a flavonoid glycoside present in the roots of Scutellaria baicalensis Georgi and this herb is used for the treatment of various inflammatory diseases, hepatitis, tumors, and diarrhea in East Asian countries such as China,
Korea, Taiwan, and Japan (Chen et al., 2001; Kubo et al., 1994). The aglycone derivative of
Baicalin known as Baicalein was found to interfere directly with key bacterial virulence pathways by targeting the Salmonella Typhimurium pathogenicity island-1 (SPI-1) type III secretion system
(T3SS) effectors and translocases, and eventually prevented bacterial invasion of epithelial cells
(Tsou et al., 2016).
Probiotics are defined as live microorganisms which when consumed in appropriate amounts confer a health benefit on the host (Araya et al., 2002). Lactic acid bacteria, especially
Lactobacillus spp., and Bifidobacteria are the most commonly used probiotic bacteria, since they are considered as integral and desirable members of the intestinal microbiota (Kailasapathy &
4
Chin, 2000; Schrezenmeir & de Vrese, 2001; Soccol et al., 2010). Probiotic bacteria exert multiple health benefits to the host, including improved nutrient digestion and assimilation, potentiating host immune function, and protection against enteric pathogens (Fukuda et al., 2011; Hill et al.,
2014; Olszak et al., 2012; Soccol et al., 2010; Sonnenburg et al., 2005). In addition, prebiotics are nondigestible food ingredients that beneficially affect the host by selectively stimulating the growth and/or activity of one or a limited number of bacterial species already resident in the colon, thereby improving host health (Gibson et al. 1995). There are several reports on the correction of gut dysbiosis by administration of probiotics or prebiotics (Colombel, 1987; Kotowska et al., 2005;
Lewis et al., 2005; Plummer et al., 2004; Segarra-Newnham, 2007). Antagonistic mechanisms of probiotics include virulence interference, especially toxin inactivation, and modulation of inflammatory responses (Chen et al., 2006).
Based on published literature and preliminary research conducted in our laboratory, it is hypothesized that sodium selenite, baicalin, and probiotics reduce C. difficile virulence. Further, it is hypothesized that sodium selenite decreases C. difficile resistance to ciprofloxacin and vancomycin. Although vancomycin is one of the antibiotics of choice for treating CDI, an increased prevalence of vancomycin resistant C. difficile strains has been recently reported (Peng et al., 2017). Moreover, fluoroquinolones, which are used for treating bacterial infections in clinical settings have been linked to promoting recurrent CDI in susceptible individuals (Peng et al., 2017). Therefore, identifying synergistic non-antibiotic agents that can enhance the therapeutic efficacy of these antibiotics against C. difficile is critical for controlling the infection in humans.
The overall goal of this dissertation is to identify adjunct/alternative approaches to antibiotics that have the potential to be developed as therapies for controlling CDI. The specific objectives are
5
1. To investigate the effect of sodium selenite on C. difficile toxin production, spore germination, and resistance to ciprofloxacin and vancomycin in vitro.
2. To investigate the effect of baicalin on C. difficile toxin production, sporulation and spore germination in vitro.
3. To study the effect of selected lactic acid bacteria on C. difficile toxin production and spore germination in vitro.
4. To determine the effect of baicalin on C. difficile pathogenicity in a mouse model.
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CHAPTER II
Review of Literature
7
C. difficile is a Gram-positive, spore forming obligate anaerobic bacillus, which causes a serious toxin-mediated enteritis in humans (Hookman and Barkin 2009). More than 500,000 cases of C. difficile infection (CDI) are reported annually in the US, resulting in ~ $6.3 billion as health- care costs (Zhang et al. 2016). The bacterium colonizes the intestine and produces a toxin-mediated enteric disease characterized by abdominal pain, fever, fulminant colitis, toxic megacolon, sepsis and shock (Rupnik et al. 2009). Asymptomatic carrier status of C. difficile with or without mild diarrhea has also been reported in some patients (Hensgens et al. 2012). The infection is generally associated with long-term use of antibiotics, proton inhibitors and anti-inflammatory agents, which can result in gut dysbiosis, predisposing C. difficile spore germination and colonization in the gastrointestinal tract (Bartlett 1992; Dial et al. 2006; Kelly and LaMont 1998). The bacterium has also emerged as a community-associated pathogen (Beaugerie et al. 2003; Hensgens et al. 2012), with increased reports of community-associated Clostridium difficile infection (CA-CDI) among young, healthy individuals, who were not previously exposed to antibiotics. However, the role of various community sources such as soil, food, water, and animals in CDI epidemiology is not well understood (Chitnis et al. 2013; Matamouros et al. 2007). In addition, emergence and spread of multi-drug resistant C. difficile isolates has been reported globally, raising potential concerns on the efficacy of antibiotic-based current treatment strategies against CDI.
1. Clostridium difficile or Clostridioides difficile (CD)
The morphological and biochemical characteristics mentioned below have been adapted from
Hall and O’Toole (1935), Prevot 1938, 84AL, with minor additions (Lawson et al. 2016). C. difficile vegetative cells are Gram-positive, usually motile in broth cultures and peritrichous, with dimensions of 0.5-1.9 x 3.0-16.9 µm. Spores are oval and subterminal, which appear swollen and
8 cell walls contain meso-Diaminopimelic acid. C. difficile sporulation can be induced when grown on Brucella blood agar for two days, whereas, spore germination can be enhanced on solid agar media containing 0.1% sodium taurocholate. Agar surface colonies appear circular, occasionally rhizoid, flat or low convex, opaque, grayish or whitish, and have a matte to glossy surface. All strains produce pale green fluorescence under long wavelength of UV light after 48 h of incubation on Brucella blood agar supplemented with hemin and vitamin K. Optimal growth temperature ranges between 30-37°C, although, growth also occurs at 25°C and 45°C. With regards to carbohydrate utilization, C. difficile strains can utilize fructose, but not amygdalin, arabinose, galactose, glycogen, inositol, inulin, lactose, maltose, melezitose, raffinose, rhamnose, ribose, starch, and sucrose; weak reactions are obtained for the utilization of cellobiose, salicin, sorbitol, trehalose, and xylose. Amino acids such as proline, aspartic acid, serine, leucine, alanine, threonine, valine, phenylalanine, methionine, and isoleucine are utilized for growth with metabolites such as d-aminovalerate and a-aminobutyrate being produced (Lawson et al. 2016). A selective minimal medium containing selected amino acids as a source of carbon and energy, only a trace of fructose (0.1%), 2% bile as a growth stimulant, with 16 pg/ml cefoxitin, and 500 mg/ml streptomycin for reduction of associated microbiota, has been found useful for isolation of these organisms from feces (Hubert et al. 1981). In addition, another frequently used agar consists of egg yolk agar base with cycloserine (500 mg/ml), cefoxitin (16 mg/ml) and fructose, (CCFA)
(George et al. 1979). C. difficile strains are abundant gas producers in peptone, yeast, glucose
(PYG) deep agar cultures (Lawson et al. 2016)
Recently, C. difficile has been reclassified from the Clostridium sensu stricto group
(Lawson et al. 2016). This reclassification was proposed since C. difficile was shown to be phylogenetically distant from the rRNA clostridial cluster I and located in cluster XI, which has
9 been moved to the family Peptostreptococcaceae. Based on the phenotypic, chemotaxonomic and phylogenetic analysis, C. difficile was proposed to be renamed as Clostridioides difficile (Lawson et al. 2016). Currently, both Clostridium difficile and Clostridioides difficile are validly used under the provisions of the Prokaryotic Code (Oren and Rupnik 2018).
1.1. Epidemiology of CDI
Nearly three decades ago, the importance of CDI was minimal due to its reduced incidence rate and the high recovery rate since patients responded well to either metronidazole or vancomycin administration. Although, recurrent CDI was documented in the past, the condition was easily manageable with infrequent incidences of severe CDI (George 1988). However, in the past twenty years, the emergence of hypervirulent C. difficile with severe pathological implications and increased antibiotic resistance has become a significant burden to the health-care systems worldwide (Spigaglia 2016).
The annual incidence of C. difficile infection (CDI) in the United States is estimated to be
453,000 cases, with 29,000 deaths, resulting in a significant financial burden ranging from $1.7-
7.0 billion to the US healthcare systems (Peng et al. 2017; Zhang et al. 2016). Recently, a 10-year review on the CDI trends in incidence, mortality and hospital charges during 2005-2014 was analyzed (Luo and Barlam 2018). The incidences of general CDI, hospital onset and non-hospital onset CDI increased by 3.3%, 1.4% and 2.0% respectively. However, the overall rate of mortality due to CDI was 8.5%, with a decline from 9.7% in 2005 to 6.8% in 2014. The reduced mortality may be attributed to the adoption of antibiotic stewardship programs in hospitals (McDonald et al.
2018), however, there still remains the underlying risk of the emergening antibiotic resistant C. difficile isolates (Centers for Disease Control and Prevention, 2013). In addition, the individual median CDI hospitalization charge increased from $41,974 in 2005 to $46,663 in 2009, which
10 subsequently declined from $45,725 in 2010 to $41,875 in 2014 (Luo and Barlam 2018). Although considered as a nosocomial pathogen predominantly affecting the elderly, immunocompromised and long-term hospital inpatients, there has been an increased incidence of community-acquired
CDI (CA-CDI), especially in low-risk populations such as individuals who have neither been recently hospitalized nor have been exposed to antibiotics (Beaugerie et al. 2003; Chitnis et al.
2013; Gupta and Khanna 2014; Khanna et al. 2012; Lessa et al. 2015). Community-associated CDI has been documented in individuals with no hospitalization within the past 12 weeks or infections that occur within 48 h of hospital admission (Lessa, 2013). It is estimated that CA-CDI accounts for 32% of all CDI cases, implicating the possibility of a supplementary C. difficile source causing the disease in non-hospitalized patients (Hensgens et al. 2012; Lessa 2013; McDonald et al., 2006;
Rupnik et al. 2009). Deshpande and coworkers (2013) reported antibiotics, especially clindamycin, fluoroquinolones and cephalosporins, enhance the risk of CA-CDI (Deshpande et al. 2013). In another other study, Khanna et al. (2012) reported that CA-CDI affected younger individuals more than hospital-acquired CDI, with a median age of 50 and 72, respectively. The incidence of CA-
CDI in Monroe County, New York was studied by Dumyati et al. (2012), who found it responsible for 18% of total CDI cases over one year. These researchers also found that 21% of these patients reported a past visit to a health-care facility with a family member within 12 weeks before CDI onset (Dumyati et al. 2012), concluding that minor exposure to health-care settings or contact with individuals under long-term hospital care could increase the risk of CA-CDI.
In addition, CDI epidemiology has dramatically changed over the past two decades with etiological implications for foodborne or zoonotic sources (Knight et al. 2015). Studies conducted by several investigators have indicated C. difficile occurrence in a variety of food animals. With increased isolation of the pathogen in animal reservoirs, it is considered as one of the reasons for
11 the increased incidence of human CDI (Indra et al. 2009; Rodriguez-Palacios et al. 2013; Rupnik et al. 2008). Food animals such as calves, pigs, sheep and chicken are known to harbor C. difficile in the gastrointestinal tract and reports of CDI have been observed among these animal species
(Alvarez-Perez et al. 2009; Knight and Riley, 2013; Rodriguez-Palacios et al. 2006; Weese et al.
2010). Similarly, companion animals, such as dogs, cats and horses, can also be colonized with the bacterium, thus becoming susceptibile to CDI (Riley et al. 1991; Songer et al. 2009).
C. difficile NAP7 (ribotype 078) and NAP8 have been isolated from food animals in the
United States and were found to be closely related or indistinguishable from human cases of ribotype 078 by pulse field gel electrophoresis (PFGE). This ribotype has been reported to be more commonly associated with CA-CDI than nosocomial CDI (Jhung et al. 2008). In the Netherlands, ribotype 078 has been found to predominate in pigs and calves with an increased incidence of human CDI particularly in rural geographical regions among younger patients (Goorhuis et al.
2008). The hypervirulent ribotype 027 has also been isolated from both food and companion animals. In Canada, a survey identified ribotype 027 to be the third most common type isolated from calves from widely dispersed geographical locations (Rodriguez-Palacios et al. 2006). In
Germany, Rabold and coworkers (2018) conducted a large-scale survey on the fecal occurrences of C. difficile among companion animals (dogs and cats) and their owners to assess the potential epidemiological links within the community. Human infection related ribotypes 014, 027 and 078 were isolated from both pets and humans. Although the isolation rate of C. difficile was very low, the study highlights the potential zoonotic risk for CA-CDI via companion animals (Rabold et al.
2018).
Besides food animal sources, C. difficile has also been isolated from soil, water, raw vegetables samples and milk (Hengsgens et al. 2012; Janezic et al. 2012; Jobstl et al. 2010; Kotila
12 et al. 2013; Metcalf et al. 2010; Rodriguez-Palacios et al. 2013). Prevalence studies conducted in the United States and Canada revealed an isolation rate ranging from 20-36% from retail meat samples such as ground beef, ready-to-eat beef, ground pork, ground turkey, pork sausage, summer sausage, pork chorizo and pork braunschweiger (Rodriguez-Palacios et al. 2007; Loo et al. 2005; Rodriguez-Palacios et al. 2009; Songer et al. 2009). Nearly two decades ago, a prevalence study to assess C. difficile spores from environmental sources was conducted within and outside the hospital milieu such as surface samples from health care facilities, water, soil and raw vegetables (Al Saif and Brazier, 1996). An overall 7% prevalence was observed across all environmental sources, with water samples yielding the highest culture positivity, followed by soil and healthcare environs (Al Saif and Brazier, 1996). In addition, a recent study revealed a higher prevalence of toxigenic C. difficile in public space lawns in western Australia, wherein the toxigenic ribotypes of 014 and 020 were predominant (Moono et al. 2017). Also, a study in Canada demonstrated a 39% positivity for ribotype 078 from sediments of rivers receiving discharge effluent pipes from wastewater treatment plants (Xu et al. 2014). Although the exact routes of pathogen dissemination are not completely delineated, all the aforementioned findings suggest the likelihood of food and other environmental sources as plausible transmission routes of human CDI especially CA-CDI.
The increased rate of recurrent CDI reported in the United States has been chiefly attributed to the emergence and dissemination of the hypervirulent North American Pulsotype 1 (NAP 1), classified as toxinotype III and ribotype 027 (Arroyo et al. 2005; Loo et al. 2005). Concurrently, many investigators have documented the emergence of antibiotic resistance in C. difficile, especially against fluoroquinolones, clindamycin, erythromycin, metronidazole and vancomycin
(Drudy et al. 2007; Kelly and LaMont 2008; Musher et al. 2005; Pepin et al. 2005; Spigaglia et al.
13
2008; Spigaglia 2016b). Consequently, the Centers for Disease Control and Prevention (CDC) in its report on emerging pathogens with antibiotic resistance, categorized C. difficile as one of the three urgent threats to public health (Centers for Disease Control and Prevention, 2013).
1.2. Clinical manifestations and risk factors associated with CDI in humans
CDI is manifested as mild, moderate or severe forms of disease in humans (Hookman and
Barkin, 2009; Weese et al. 2010). Mild form is usually asymptomatic or clinically presented with mild fever and abdominal cramps. The primary symptom of moderate to severe CDI is watery diarrhea in affected patients (Kelly & LaMont, 1998). Apart from watery and rarely bloody diarrhea manifested in severe CDI, abdominal pain due to colonic distension and toxic megacolon are also observed, which lead to complications and potentially death (Hookman & Barkin, 2009;
Kelly & LaMont, 1998; Knight & Surawicz, 2013). Pseudomembranous colitis is a classical sign of complicated CDI formed by necrotic epithelial debris, proteinaceous exudates and inflammatory cells in the colonic lumen, which are endoscopically visualized as yellow membranous plaques in the colonic lumen (Kelly & LaMont, 1998; Knight & Surawicz, 2013). The complications associated with pseudomembranous colitis include colonic perforation, and in advanced stages, can lead to sepsis and death in some patients, especially in the elderly population (Knight &
Surawicz, 2013).
Antibiotic exposure, advanced age, duration of hospital stay and severity of underlying disease conditions are the major risk factors associated with CDI (McDonald et al. 2018). For many years, almost all known antibiotics have been associated with CDI, but in particular, 3rd and
4th generation cephalosporins, carbapenems (Hensgens et al. 2011), fluoroquinolones (Loo et al.
2005b; Muto et al. 2005; Pepin et al. 2005) and clindamycin (Johnson et al. 1999; Thibault et al.
1991) have been found to be the high-risk categories. Antibiotic administration increases the risk
14 of CDI as it suppresses the normal gastrointestinal microbiota, thereby creating an accommodating niche for C. difficile to colonize and cause infection (Dethlefsen et al. 2008). The relative risk for
CDI with a particular antibiotic agent depends mainly on the local prevalence of C. difficile strains that are specifically resistant to the antibiotic being administered (Johnson et al. 1999). The residual gut dysbiosis due to antibiotic administration could be long-lasting, implicating a risk for
CDI during the therapeutic period, with a 7-10-fold higher risk during the first month of post- antibiotic exposure, and the risk protracting for up to three months following the termination of antibiotics. It is also known that exposure to multiple antibiotics increases the risk for CDI
(Hensgens et al. 2011).
1.3. Pathogenesis of CDI
Feco-oral route is the primary mode of C. difficile transmission in humans (Hookman and
Barkin 2009). C. difficile spores are resistant structures shed by affected patients that contaminate and survive the hospital environment such as surfaces and equipment for months, and are extremely resistant to physical and chemical sanitizing agents (Bettin et al. 1994b; Jabbar et al.
2010; Kim et al. 1981; Siani, Cooper, Maillard 2011). Spores ingested by susceptible patients can survive the low pH of the gastric environment and eventually reach the intestine. With the absence of normal gut microbiota, spores germinate to vegetative cells in the presence of primary bile salts present in the small intestine. In healthy individuals, the actively secreted primary bile salts from gall bladder, especially sodium taurocholate (Railbaud et al. 1974) are reabsorbed in the distal ileum (Kelly and LaMont 1998; Knight and Surawicz 2013). Taurocholate residues seen in the distal part of the intestine are readily transformed to secondary bile salt metabolites by the normal benign microbiota. This reduces the availability of primary bile salts to induce C. difficile spore germination, thereby minimizing initiation of C. difficile pathogenesis in healthy individuals
15
(Kelly and LaMont 1998b; Knight and Surawicz 2013). Also, secondary bile salts such as chenodeoxycholate, are known to suppress C. difficile spore germination and vegetative growth
(Sorg and Sonenshein 2010). Hence, a healthy microbiota modulates resistance against CDI by influencing the metabolism of primary bile salts, which is critically required for C. difficile spore germination (Giel et al. 2010; Theriot et al. 2016).
The vegetative C. difficile colonize and multiply in the intestinal crypts to produce major exotoxins, namely toxin A and B, which are critical virulence factors for CDI (Kuehne et al. 2011).
These exotoxins possess glucosyltransferase domain (GTD) that glycosylate Rho and Rac
GTPases in the colonic epithelial cells and destabilize critical functions such as cytoskeletal disruption eventually leading to tight junction dissociation between colonic epithelial cells and the loss of epithelial integrity (Hunt and Ballard 2013). This results in an increased intestinal permeability and translocation of bacteria from the gut lumen into deeper tissues (Naaber et al.
1998). Damaged epithelial cells release cytokines and chemokines such as IL-1β, IL-8, CXCL-1 and CXCL-2, which initiate neutrophil recruitment and activation of resident dendritic cells and macrophage, favoring the release of additional proinflammatory cytokines, including IL-1β, IL-12 and IL-23. This process stimulates innate lymphoid cells to release IL-22 and IFN-γ to increase macrophage and neutrophil phagocytic activity, production of antimicrobial peptides, reactive oxygen and nitrogen species (RNS and ROS), which in turn further limit the translocation of other intestinal bacteria. Although inflammatory responses are essential for host survival subsequent to
CDI, an overactivation of inflammatory responses proceeds to a condition called pseudomembranous colitis, which in advanced cases, can be detrimental to the host. C. difficile toxins in damaged epithelia further promote the release of cytokines such as IL-1, IL-8 and leukotriene-B, which further recruit more neutrophils to the affected region causing additional
16 mucosal injury and focal micro-abscesses and pseudomembrane formation. In adverse conditions, an exaggerated immune response and release of systemically active cytokines, complicated by fluid loss from the resultant severe diarrhea may lead to systemic shock and death (Knight and
Surawicz 2013).
1.4. C. difficile toxins
C. difficile toxins are high molecular weight proteins classified as members of the family of large clostrial toxins (LCTs) (Voth and Ballard, 2005; Popoff and Bouvet, 2009). C. difficile produces two potent exotoxins: the enterotoxin, toxin A (TcdA) and the cytotoxin, toxin B (TcdB), which are the major virulence factors of the pathogen responsible for CDI in humans (Popoff and
Bouvet; Kelly & LaMont, 1998; Stanley et al. 2013). Apart from these toxins, hypervirulent C. difficile strains are known to produce binary toxins (CdtA and CdtB) which can also contribute to the severity of CDI in affected patients. (Block 2004; Bondo et al. 2015; Gerding et al. 2014). The toxins, TcdA and TcdB, have molecular weights of 308 and 270 kDa respectively, with 49% identity and 63% similarity in their amino acid sequence (Popoff and Bouvet, 2009). Both TcdA and TcdB holotoxins possess the same ABCD domain structure: the “A” domain possesses the N- terminal glucosyltransferase activity; the “B” domain located at the C-terminal part of the holotoxin harbors combined repetitive oligopeptides (CROPs); the “C” domain consists of a cysteine protease domain that aids to cleave the holotoxin to release the glucosyltransferase domain, and the “D” domain corresponding to the internal hydrophobic region. The holotoxin activity involves four major steps: (a) endocytosis mediated by the B domain; (b) D domain- mediated intrusion of host membranes for the translocation of a catalytic domain into the cytosol;
(c) release of the A domain from host lysosomes subsequent to autoproteolytic processing by the
C domain and (d) inactivation of the host Rho family GTPases protein by glycosylation (Janoir
17
2015; Oezguen et al. 2012; Pruitt and Lacy 2012). This process induces cytoskeleton destruction and cell death as a result of apoptosis and necrosis (Kuehne et al. 2011). Although the toxins have comparable structure and mechanism of action, TcdB is 100–10,000 times more potent than TcdA in its cytopathic effect on various cell types (Voth and Ballard 2005). Earlier studies have proposed
TcdA as a key virulence factor in CDI that aids to facilitate TcdB entry and potentiate its activity
(Gerhard et al. 2008; Kuehne et al. 2011; Sutton et al. 2008). However, follow up investigations have demonstrated that TcdB is the major virulence factor and TcdA is not critically essential for
CDI in humans (Lyras et al. 2009; McDonald et al. 2005). In addition, a similar degree of damage is evinced due to infection with either TcdA-TcdB+ or TcdA+TcdB+ strains of C. difficile, and no incidences of pathogenic TcdA+TcdB- strains have been reported from human CDI cases (Sambol et al. 2000). Binary toxins, CdtA and CdtB (or C. difficile transferase; CDT) of C. difficile possess similar mechanistic activities as that of TcdA and TcdB. CdtB contributes to virulence by augmenting C. difficile adherence to gut epithelial cells, whereas CdtA serves as the catalytic component having ADP-ribosyltransferase activity, which disrupts actin filament polymerization and cause cell rounding (Barth and Stiles 2008; Gerding et al. 2014; Hemmasi et al. 2015; Schwan et al. 2009). Interestingly, it was observed that naturally occurring TcdA-TcdB-CDT+ strains caused enterocyte damage in an ex situ rabbit ileal loop assay (Geric et al. 2006), however, this strain was not virulent in a hamster model (Eckert et al. 2015).
The operon that expresses and regulates toxin production is encoded by a 19.6 kb region called the Pathogenicity locus (PaLoc) in the C. difficile genome (Dingle et al. 2013; Neyrolles et al. 2011). This locus comprises of tcdA and tcdB genes encoding the holotoxins; tcdC and tcdR genes encoding factors that regulate transcription of the PaLoc region; and tcdE gene that encodes a holin-like protein. TcdA and TcdB gene expression occurs during late log phase and early
18 stationary phases of bacterial growth (Voth and Ballard 2005). TcdR is an alternative sigma factor that positively regulates the expression of toxin genes (Dupuy and Matamouros 2006; Popoff and
Bouvet 2009). TcdC is the suppressor of toxin genes, which acts by inhibiting TcdR complex required for toxin gene transcription (Dupuy et al. 2008; Matamouros et al. 2007). In addition,
TcdE is a porin protein that helps to shuttle C. difficile toxins to the exterior of the bacterial cell
(Govind and Dupuy 2012; Govind et al. 2015).
Regulatory gene elements located outside the PaLoc region such as codY and ccpA are also known to influence C. difficile toxin gene regulation (Antunes et al. 2011; Dineen et al. 2010).
CodY is a global gene regulator that strongly represses the expression of virulence associated genes in numerous Gram-positive bacteria, including Clostridia (Dineen et al. 2007; Fisher et al.
1996; Guedon et al. 2001; Lobel et al. 2012). CodY is expressed in response to branched chain amino acids, which binds to the tcdR promoter and in turn suppresses toxin production (Dineen et al. 2007). CcpA is a member of the LacI/GalR family of transcriptional regulators commonly seen in low G+C Gram-positive bacteria that rapidly recognizes easily metabolizable carbon sources such as glucose and influences genes responsible for carbon and nitrogen metabolism (Antunes et al. 2011; Antunes et al. 2012; Li et al. 2015). In addition to its influence on carbon-nitrogen metabolism, it negatively regulates virulence-associated genes in pathogenic Gram-positive bacteria classified under the Clostridium and Bacillus genus. Likewise, in C. difficile, CcpA aids to sense the nutrient rich environment in culture media and eventually suppress the toxin production (Antunes et al. 2011). Moreover, it has also been observed that the sporulation gene, spo0A regulates both sporulation and toxin production in C. difficile (Pettit et al. 2014), although its mechanistic pathways are yet to be delineated. Recently, researchers have identified a gene,
CD3668 that is responsible for reducing sporulation and increasing toxin production and motility.
19
This locus was renamed as rstA as an acronym for regulator of sporulation and toxins. rstA expresses a bifunctional protein that upregulates sporulation through an unknown pathway, and simultaneously represses motility and toxin production by influencing sigD transcription (Edwards et al. 2016).
1.5. Non-toxin associated virulence factors initiating host colonization
The initial interaction of C. difficile with the host requires numerous surface bacterial components. Most of the surface proteins identified are known to be involved in mucus and cell adhesion, and some of these factors are involved in triggering the host immune response. The main cell wall proteins (Cwp) associated with C. difficile surface include the S-layer proteins (SLPs), a cysteine protease (Cwp84), and an adhesin, Cwp66 (Calabi et al. 2001; Fagan et al. 2011; Janoir et al. 2007; Karjalainen et al. 2001; Waligora et al. 2001). The S-layer proteins (SLPs) are highly immunogenic proteins that form a crystalline array over the entire bacterial cell surface. This comprises two major components, namely the low molecular weight (LMW) SLP and high molecular weight (HMW) SLP, which are derived by the enzymatic cleavage of the precursor SlpA protein by Cwp84. The low molecular weight SLP is the surface exposed component required for adhesion, and is known to be highly variable among different strains (Eidhin et al. 2006). On the other hand, the high molecular weight SLP segment is anchored to the cell wall and is required for adhering to intestinal tissues and extracellular proteins such as collagen, thrombospondine and vitronectin (Calabi et al. 2001; Karjalainen et al. 2001). Cwp84 is another highly immunogenic, surface exposed cysteine protease that is responsible for cleaving the SlpA precursor protein into their respective LMW- and HMW- SLPs (Janoir et al. 2007; Montes et al. 2013; Pechine et al.
2005). This enzyme possesses an N-terminal proteolytic site and C-terminal domain that aids in cell wall anchoring (Dang et al. 2010; Kirby et al. 2009). Moreover, it also known to degrade the
20 host’s extracellular matrix components such as fibronectin, laminin and vitronectin and facilitate bacterial spread (Janoir et al. 2007). Cwp66 is another highly immunogenic, two-domain adhesin protein involved in C. difficile adherence to intestinal epithelial cells. The C-terminal region is a highly immunogenic, surface exposed domain and observed to possess a high degree of variability among C. difficile isolates (Waligora et al. 2001). Fibronectin binding protein, Fbp, is a highly conserved, surface exposed protein responsible for fibronectin binding and adherence to intestinal epithelial cells. Fbp is also known to induce immunogenicity in C. difficile patients (Barketi-Klai et al. 2011; Hennequin et al. 2003). Other miscellaneous proteins required for C. difficile colonization include, surface-localized, collagen binding protein CbpA (Tulli et al. 2013), a zinc- metalloprotease capable of cleaving host IgA2, fibronectin or fibrinogen (Cafardi et al. 2013;
Hensbergen et al. 2014), and flagellar proteins, FliC and FlicD that play an important role in intestinal adherence and colonization (Dingle et al. 2011; Stevenson et al. 2015).
1.6. C. difficile sporulation
Sporulation or otherwise spore formation, is a virulence factor that plays a critical role in
C. difficile infection in humans. Spores are dormant survival structures that withstand unfavorable physical, chemical and metabolic conditions, that aid C. difficile for its environmental persistence, transmission to susceptible individuals, and to induce relapse in temporarily recovered patients
(Paredes-Sabja et al. 2014). The sporulation process in C. difficile is a very complex and less understood process. However, various environmental stimuli such as nutrient deprivation, quorum sensing, physical/chemical stress and oxidative stress are considered to trigger the process (Darkoh et al. 2016; Higgins and Dworkin 2012; Paredes-Sabja et al. 2014). Spore formation in C. difficile comprises of four morphogenetic phases: (a) the asymmetric septation of vegetative cells into a smaller compartment and larger mother cell; (b) the mother cell engulfing the smaller compartment
21 in a phagocytic-like event resulting in a forespore within the cytoplasm of the mother cell; (c) assembly of the spore cortex and coat layers; (d) lysis of the mother cell and release of mature spore into the environment. The core region of the spore contains the genomic DNA, mRNA, ribosomes, protein, and is very rich in the calcium salt of pyridine-2,6-dicarboxylic acid (DPA).
The spore core is centripetally overlaid by five other membranes such as the inner membrane, a peptidoglycan-containing germ cell wall, a specialized peptidoglycan-containing cortex, an outer membrane and multiple layers of coat protein (Edwards and McBride 2014; Gil et al. 2017).
The master transcriptional regulator, Spo0A plays a critical role during C. difficile sporulation. Activation of Spo0A requires phosphorylation, which is potentially mediated by a histidine kinase that is encoded from five putative orphan histidine kinase genes: CD1352,
CD1492, CD1579, CD1949 and CD2492. The histidine kinase, CD 1579, is known to autophosphorylate and transfer a phosphate group to Spo0A (Underwood et al. 2009). In addition, studies have shown that deletion mutants of CD 2492 (Underwood et al. 2009) have been shown to possess reduced sporulation in C. difficile. Contrastingly, researchers have identified that CD
1492 functions to repress sporulation in C. difficile (Childress et al. 2016). However, the role of
CD1352 and CD1949 histidine kinases in Spo0A phosphorylation remains uncertain (Zhu, Sorg,
Sun 2018). CodY and CcpA are known to negatively influence sporulation in C. difficile. Although the mechanistic pathways have not been completely delineated, CodY is shown to negatively affect spo0A expression and positively regulates sporulation inhibitor genes such as sinI and sinR
(Edwards and McBride 2014). CcpA represses critical factors such as spo0A and sigF, which are required in the early phase of sporulation (Antunes et al. 2012). In addition, researchers have identified a novel regulator, RstA that positively regulates C. difficile sporulation and simultaneously represses toxin production and motility by indirectly affecting the expression of
22 flagellar specific sigma factor, SigD (Edwards et al. 2016). Moreover, RNA polymerase sigma factors, σF ,σE ,σG and σK are involved in the complex regulatory process of sporulation. Forespore formation is initiated by sigma factors σF and σG, whereas σE and σK are required for mother cell formation (Fimlaid et al. 2013). Spo0A activates the forespore protein, SpoIIR via σF, which in turn is required for the activation of mother cell sigma factor, σE (Saujet et al. 2013). The downstream activation of SpoIIID mediated by σE is eventually required for σK production and activation in the mother cell (Paredes-Sabja et al. 2014). The synthesis of spore coat surrounding the forespore is mediated by SpoIVA (Putnam et al. 2013).
1.7. C. difficile spore germination
Transmission of C. difficile in humans occurs as a result of ingestion of spores. The critical step involved in the establishment of CDI involves the germination of spores in the small intestine.
Spore germination in C. difficile occurs in response to the presence of bile salt germinants
(taurocholic acid or cholic acid derivative) and amino acids (e.g., glycine or alanine) (Sorg and
Sonenshein 2008). Compounds structurally similar to cholic acid, such as chenodeoxycholic acid derivatives are known to be competitive inhibitors of cholic acid mediated spore germination
(Francis et al. 2013). In spore forming bacteria, three subtilisin-like serine proteases namely, CspA,
CspB and CspC, present in spore assist in the initiation of spore germination (Adams et al. 2013;
Kevorkian et al. 2016). However, C. difficile is known to encode a pseudoprotease from the CspC gene, which acts as a bile acid germinant receptor. The CspB is a catalytically active protease that is required for the cleavage of a pro-SleC protein into its active, spore cortex degrading form, known as the cortex hydrolase protein or SleC. The activated cortex hydrolase protein degrades the cortex causing to an osmotic swelling of the inner spore membrane and leads to the release of the calcium salt of dipicolinic acid (CaDPA) from the spore core to the exterior. This process
23 results in the hydration of the spore core and reactivation of the spore core metabolism (Paredes-
Sabja et al. 2011).
1.8. Influence of gut microbiome on CDI susceptibility
A normal and healthy gut microbiome composition provides colonization resistance to most enteric pathogens, including C. difficile (Britton and Young 2014). Hence, disruption of the normal gut microbiota is the most important predisposing factor for CDI (Hookman and Barkin
2009). Antibiotic therapy affects microbial diversity and the induced changes tend to persist even after the withdrawal of antibiotics (Antonopoulos et al. 2009). This leads to a loss of colonization resistance and increases the susceptibility to CDI. Some of the major antimicrobial classes or antibiotics that predispose humans to CDI are fluoroquinolones, cephalosporins, penicillins and clindamycin (Blossom and McDonald 2007; McFarland 2008; Spigaglia 2016).
Continuous shifts and alterations in the gut microbiota occur throughout the human lifespan
(Hopkins et al. 2001). In healthy adults, the gut microbiome is almost stable, however, the microbial composition undergoes significant alterations and becomes less diverse as age advances
(Biagi et al. 2010; Claesson et al. 2011; Hopkins et al. 2001). In elderly individuals, there is a considerable drop in the protective populations such as Bifidobacterium and other members associated with Firmicutes, along with an increase in undesirable Proteobacteria in the gut (Biagi et al. 2010; Claesson et al. 2011; Hopkins et al. 2001). In addition, immunosenescence along with frequent hospital stays among the elderly may contribute to a negative alteration in the microbiome that favors C. difficile colonization (Seekatz and Young 2014).
Chronic gastrointestinal diseases and the use of proton pump inhibitors are other important factors that detrimentally affect the gut microbiota and predispose an individual to CDI (Berg et
24 al. 2013; Dial et al. 2005; Vesper et al. 2009). The use of proton pump inhibitors alters the pH of the gut, thereby affecting microbial populations, especially beneficial bacteria such as
Lactobacillus spp. Moreover, inflammatory bowel disease also contributes to gut dysbiosis typically showing reduced diversity of Firmicutes and Bacteroides populations, along with an increased Proteobacteria in the gut of affected patients (Manichanh et al. 2006; Nagalingam and
Lynch 2012).
C. difficile infected patients possess a less diverse gut microbiome compared to healthy, non-infected adults (Rea et al. 2012). With the advancement of age, the ratio of gut Firmicutes to
Bacteroidetes increases, which is indicative of gut dysbiosis (Ling et al. 2014; Mariat et al. 2009).
Previous researchers have identified this specific gut microbiome pattern also predisposes individuals to CDI. These specific patterns are indicative of a reduced colonization resistance, which would directly favor the initiation and persistence of C. difficile (Rea et al. 2012). CDI patients generally have a reduced abundance of Ruminococcaceae and Lachnospiraceae, as well as butyrate-producing bacteria compared to healthy subjects, whereas
Enterococcus and Lactobacillus are more abundant in CDI patients (Antharam et al. 2013).
Several others have indicated that a decrease in Enterococcaceae along with a concurrent increase in Lactobacillaceae and Enterobacteriaceae is commonly observed in C. difficile positive patients
(Buffie et al. 2012; Crobach et al. 2018; Perez-Cobas et al. 2014; Rea et al. 2012; Schubert et al.
2014; Skraban et al. 2013; Zhang et al. 2015). Colonization resistance to C. difficile in humans is mainly contributed by abundance of Lachnospiraceae, Ruminococcaceae, and Bacteroidaceae families (Antharam et al. 2013; Schubert et al. 2014; Schubert et al. 2015). Surprisingly, an appreciable majority of infants is colonized with C. difficile, but they do not develop CDI
(Rousseau et al. 2011). This is attributed to a comparatively higher population of Bifidobacteria
25 in the gut that may exert a protective effect against C. difficile in infants (Rousseau et al. 2011,
Rea et al. 2012).
2. Antimicrobial resistance in C. difficile
The prolonged use of antibiotics is considered the most important risk factor to CDI in humans (Leffler and Lamont 2015). Currently, C. difficile is known to be resistant to multiple antibiotics such as fluoroquinolones, cephalosporins, penicillins, aminoglycosides, lincomycin, tetracyclines, erythromycin and clindamycin, which are commonly used to treat bacterial infections in a clinical setting (Johanesen et al. 2015; Spigaglia 2016). Generally, clindamycin, cephalosporins and fluoroquinolones are known to promote CDI (Johanesen et al. 2015; Slimings and Riley 2013; Spigaglia 2016). Based on CLSI breakpoints for susceptibility testing in anaerobic bacteria, approximately 30% of ribotype 027 strains in North America were resistant to multiple drugs, including clindamycin, moxifloxacin and rifampin (Tenover et al. 2012). Although, metronidazole and vancomycin remain to be the first line of antibiotics for CDI treatment, C. difficile isolates with significantly reduced susceptibility to these drugs have been reported (Adler et al. 2015; Goudarzi et al. 2013). In addition, resistance to vancomycin, rifamycins, fidaxomicin, tetracylines and chloramphenicol has also been documented (Freeman et al. 2015; Goudarzi et al.
2013; Leeds et al. 2013).
The factors contributing to the development of antibiotic resistance in C. difficile include the presence of resistance associated genes in the chromosome, mobile genetic elements (MGEs), alterations in the antibiotic targets and associated metabolic pathways (Peng et al. 2017). Genes encoding penicillin binding proteins and β-lactamase-like proteins confer resistance to penicillins and cephalosporins, respectively (Spigaglia 2016). The conjugation, transduction and transformation of mobile genetic elements such as transposons among C. difficile strains and/or
26 between C. difficile and other bacterial species are important mechanisms of antibiotic resistance acquisition in the pathogen (Spigaglia 2016). The transposons, Tn5398, Tn5398-like derivative,
Tn6194 and Tn6215 mediate resistance to antibiotics of the MLSB (macrolide-lincosamide- streptogramin B) family (Tsutsumi et al. 2014). Tetracycline resistance is considered to be associated with the Tn5397, Tn916 and Tn6164 transposons, which harbor the tet class of genes comprising of tet(M), tet(44), and tet(W) (Tsutsumi et al. 2014). Mutations in the peptidoglycan biosynthesis-associated proteins such as MurG is considered as the possible mechanism for conferral of vancomycin resistance in C. difficile (Leeds et al. 2013). Likewise, alterations in the rpoB gene, that encodes the bacterial RNA polymerase, are attributed for resistance to rifamycin class of antibiotics (O'Connor et al. 2008). In addition, the plausible alteration in the quinolone- resistance determining region of both GyrA and GyrB is believed to mediate resistance to fluoroquinolones (Spigaglia 2016).
3. Treatment approaches for human CDI
The current challenge with CDI is the medical management of the condition in hospital settings. Relapse in CDI treatment is known to range from 20% after the first episode, to 60% after multiple occurrences of relapse (Ianiro et al. 2018). Treatment and management of CDI require the use of antibiotics as well as other adjunct treatment strategies that target C. difficle colonization, toxin activity, relapse and most importantly, the disruption of the gut microbiota.
3.1. Current and novel antimicrobials for CDI
The choice of antibiotic therapy is based on CDI infection severity and recurrence rate. The currently recommended antibiotic therapy for CDI includes drugs under the class of imidazoles
(e.g., metronidazole), glycopeptides (e.g., vancomycin) and macrolides (e.g., fidaxomicin) (Bauer
27 et al. 2009; Cohen et al. 2010; Debast et al. 2014). Of these antibiotics, metronidazole and vancomycin have been used for the past thirty years (Peng et al. 2018). Treatment with metronidazole or vancomycin for 10-14 days has been found to be effective in 50% of the patients after the first episode of recurrent CDI (Longo et al. 2015). In addition, treatment for subsequent episodes of recurrent CDI is difficult as a result of persistence of spores in the gut along with an inadequate immunological response against C. difficile toxins. In such instances, tapered and pulsed regimen of vancomycin administration combined with fecal microbial transplantation, or fidaxomicin for 10 days is recommended (Leffler and Lamont 2015; Surawicz et al. 2013). In addition to the aforementioned treatment options, other antibiotics such as nitazoxanide, rifamixin, ramoplanin, tigecycline and teicoplanin have been used in cases where severe and adverse effects have been observed with standard therapy. These antibiotics are usually considered for salvage therapy in cases of fulminant CDI, multiple recurrences, and where surgical interventions are impossible. However, these antibiotics are not considered as a primary choice for treatment because of the limited clinical data, high cost, along with an unfavorable adverse-event profile.
Moreover, they are also implicated for the development of resistance in C. difficile (Leffler and
Lamont 2015).
Novel antibiotics are currently being evaluated for the therapy against CDI, which are currently in different phases of clinical trials. Cadazolid (developed by Actelion) and Surtomycin
(developed by Merck) have completed phase III clinical studies. In addition, LFF571 (Novartis) and Ridinilazole/SMT19969 (Summit Pharmaceuticals) have completed phase II studies.
Moreover, CRS3123 developed by the National Institute of Allergy and Infectious Diseases, is currently being evaluated in phase I studies (Fehér et al. 2017).
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3.2. Alternate and emerging treatment strategies for CDI
3.2.1. Immunization strategies
Bezlotoxumab (ZINPLAVATM, Merck) is the first US-FDA (United States Food and Drug
Administration) approved human monoclonal antibody therapy aimed at preventing the cytotoxic effects of TcdB (Peng et al. 2018). In addition, Merck also developed a human monoclonal antibody, actoxumab that is capable of neutralizing TcdA. A phase 3 clinical trial found that a significantly lower recurrence of CDI was observed when actoxumab and bezlotoxumab were provided in combination (Bartlett 2017; Wilcox et al. 2015). Moreover, in a mouse CDI model, administration of actoxumab-bezlotoxumab combination facilitated normalization of the gut microbiota in CDI mice (Džunková et al. 2016).
There are several promising vaccine targets that target TcdA and TcdB, which are currently being studied and at different phases of clinical trials. Cdiffense is a vaccine that contains toxoids of TcdA and TcdB developed by Sanofi Pasteur that is currently in phase III trials (Peng et al.
2018). Two other vaccines are currently in phase II clinical trials: IC84 is a recombinant truncated version of the TcdA and TcdB from Valneva, and Bivalent toxin vaccine from Pfizer. Other vaccine candidates developed include recombinant vaccines based on polysaccharide glycans, glycoconjugate vaccines and DNA-based vaccines, which have displayed good efficacy against
CDI under laboratory conditions or clinical trials (Peng et al. 2018). However, it should be noted that active immunization does not prevent C. difficile colonization in the gut, but it can ameliorate the severity of infection by blocking the toxin-mediated pathology (Longo et al. 2015).
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3.2.2. Fecal microbiota transplantation
Fecal microbiota transplantation (FMT) is intended to restore the normal gut microbiota in
CDI patients. It is considered as a safe, valid and effective method for treating multiple recurrent
CDI since 2010 (Liubakka and Vaughn 2016), with a success rate of >90% in patients with recurrent CDI (Kassam et al. 2013). In particular, FMT is recommended as a therapeutic option if there is a third recurrence after a pulsed vancomycin regimen (Surawicz et al. 2013). However, the exact role of FMT in treating primary and recurrent CDI is still not well-understood (Peng et al.
2018). The general routes of FMT administration includes (a) the upper gastrointestinal tract by means of endoscopy, pill ingestion or nasointestinal tubes; (b) the proximal part of colon by colonoscopy, and (c) the distal part of the colon by enema, rectal tube or sigmoidoscopy (Liubakka and Vaughn 2016). Recently, a study observed that delivery of fecal sample through the lower gastrointestinal tract is a safe and effective treatment for recurrent CDI, and is known to yield quicker results than delivering through the upper gastrointestinal tract (Cohen et al. 2016).
3.2.3. Non-toxigenic strains of C. difficile
The use of non-toxigenic C. difficile helps to facilitate competitive exclusion of pathogenic
C. difficile in the gut. The C. difficile strain, VP20621 (NTCD-M3) was shown to colonize the gastrointestinal tract of CDI patients (Villano et al. 2012). In addition, this strain was capable of reducing pathogenic C. difficile colonization when used in conjunction with antibiotics such as vancomycin and metronidazole (Gerding et al. 2015). Although the approach has been demonstrated to exert a positive impact, there are concerns of whether the non-toxigenic C. difficile strains might acquire the PaLoc region of the chromosome that encodes C. difficile toxins via horizontal gene transfer (Ivarsson et al. 2015).
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3.2.4. Emerging strategies for CDI therapy
As most of the current emerging approaches are still in the laboratory phase, they might turn out to be future therapies for the treatment of CDI. The use of bacteriophage lysin protein and its catalytic domain (PlyCD1-174) cloned from the prophage sequence of the C. difficile CD630 genome has proven C. difficile lytic activity, with the catalytic domain possessing a broader lytic spectrum against the pathogen (Wang et al. 2015). Moreover, in vitro studies have shown that sub-inhibitory doses of vancomycin combined with the catalytic domain were significantly more bactericidal against C. difficile compared to vancomycin alone treatments (Wang et al. 2015).
Antimicrobial peptides such as human alpha-defensins: HNP-1, HNP-3 and HD-5 have the potential in preventing TcdB induced cytotoxicity in intestinal epithelial cells. In addition, HNP-1 and HD-5 also displayed good killing effects against C. difficile, with HD-5 having an improved bactericidal activity against the pathogen (Giesemann et al. 2008). Moreover, it is suggested that combination of HD-5 with FMT therapies can be useful for treating recurrent CDI because of their inherent properties (Furci et al. 2015).
A low molecular weight organoselenium compound, ebselen, elicits its anti-toxin activity by directly targeting the glucosyltransfease domain of TcdA and TcdB, thereby neutralizing C. difficile toxin mediated pathology in mice (Bender et al. 2015). Non-absorbable anionic polymers such as Tolevamer, has the potential to absorb TcdA and TcdB, and clinical trials on Tolevamer therapy against CDI have been initiated. Although, Tolevamer did achieve a lower CDI recurrence rate, the results were not promising in terms of the duration of resolution of diarrhea and with the lower rate of clinical success (Johnson et al. 2014). However, such anionic polymers with toxin binding properties could be considered as an adjunct treatment option with antibiotic therapy against CDI.
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4. Metals
Metals that form a major part of essential minerals have historically been used as antimicrobial agents. However, with the advent of antibiotics, the application of metals as antimicrobial agents in human medicine and agriculture began to decline. In light of emergence of multidrug-resistant pathogens and reduced number of antibiotics currently being discovered, metals received increased attention in recent years, especially as potent antimicrobial agents against antibiotic-resistant pathogens. Recently, researchers have identified a reciprocal relationship for serum trace metal concentrations, especially iron, zinc and selenium, with C. difficile disease severity. The researchers suggest that interventions improving the bioavailability of trace metals may have a substantial impact on the disease outcome in CDI. However, further mechanistic studies are required to determine its effects on host-pathogen interaction and pathogen virulence (Monaghan et al. 2018).
Selenium (Se) is a metalloid, essentially consumed as a dietary antioxidant and its related compounds are commonly used in nutrition and as a chemopreventive agent (Estevam et al. 2015).
It is an essential constituent of several redox enzymes in the biological system, such as glutathione reductase. The US-FDA recommends dietary intake of selenium, with an upper tolerable intake of
400-800 µg/day (Ross et al. 2014). There are several selenium-based formulations such as selenomethionine and sodium selenite, which are commercially available as food supplements, anticancer agents and immune stimulators. The antimicrobial properties of selenium were studied in detail during the early 20th century (Kumar et al. 2010; Soriano-Garcia 2004). Previous researchers have identified that sodium selenite affected the growth and protein synthesis in various pathogens such as Bacillus subtilis, B. mycoides, Escherichia coli, and Pseudomonas. In addition, the researchers also identified that sodium selenite was capable of increasing the
32 pathogen’s sensitivity to antibiotics (Vasić et al. 2011). Several researchers have also shown that selenium was capable of strongly inhibiting spore germination, germ tube elongation and mycelial spread in various spoilage fungi associated with fresh produce such as Penicillium expansum and
Botrytis cinerea (Wu et al. 2016). Recent reports have revealed in vitro anti-toxigenic effect of selenium against V. cholerae (Bhattaram et al. 2017) and E. coli O157:H7 (Surendran Nair, 2016).
Moreover, the biological activity and role of different selenium compounds were studied against antibiotic resistant S. aureus. In this study, it was shown that selenium was by itself toxic to the pathogen and it increased bacterial antibiotic sensitivity. In addition, biologically produced and synthetic nanoparticles of selenium were demonstrated to possess anti-biofilm activity against several foodborne pathogens, including B. cereus, Enterococcus faecalis, S. Typhimurium, and S.
Enteritidis as well as drug resistant nosocomial pathogens such as methicillin-resistant S. aureus
(MRSA) (Khiralla and El-Deeb 2015; Tran and Webster 2011).
The suggested mechanistic activity of selenium is indicated as interference with the bacterial proteasome machinery and by the bioreductive formation of insoluble elemental deposits
(Estevam et al. 2015). Likewise, previous studies have also mentioned that sodium selenite exert their antimicrobial activity by increasing oxidative stress, damaging the DNA and by depleting the cellular thiostat (Jacob 2011; Jacob et al. 2011). A role for thiol depletion has also been demonstrated in the antimicrobial action mediated by Se nanoparticles. Thiol groups undergo reduction in the presence of selenium and release S-, which in turn favors the generation of reactive oxygen species such as superoxide. This process subsequently increases the expression of genes involved in ROS elimination (Spallholz et al. 2001; Wang and Webster 2012) and also induce
DNA damage, thereby causing bacterial cell toxicity (Jacob 2011).
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5. Phytochemicals
Phytochemicals represent a natural group of molecules that have been used for treating various diseases in traditional medicine (Wollenweber, 1988). Previous research conducted in our laboratory revealed that phytochemicals such as trans-cinnamaldehyde and carvacrol increased the sensitivity of multi-drug resistant Salmonella Typhimurium DT 104 to antibiotics by down- regulating antibiotic resistance genes and the efflux pump, tolC (Kollanoor Johny et al., 2010).
Moreover, in the context of CDI, our laboratory previously observed carvacrol and trans- cinnamaldehyde significantly reduced C. difficile toxin production by downregulating toxin production genes (Mooyottu et al., 2014). Carvacrol also inhibited in vitro sporulation and spore outgrowth in C. difficile (Mooyottu et al., 2017a). Follow up studies in a mouse model showed that carvacrol supplementation significantly reduced diarrhea, mitigated clinical symptoms of CDI and promoted a favorable gut microbiota shift without detrimentally affecting the gut microbiome diversity (Mooyottu et al., 2017b).
Plant-derived flavonoids are naturally occurring phenylchromones known to exert a wide array of biological activities, including antiallergic, antimicrobial, antimutagenic and antioxidant activities. Some flavonoids innately contribute to the plant’s antimicrobial defense systems and possess antagonistic activities against a wide range of other pathogenic microbes (Cowan, 1999;
Middleton, 1994; Tringali, 2003). Baicalin (5,6,7-trihydroxyflavone) is a major flavone glycoside purified from the roots of Scutellaria baicalensis, and has been described as an herb in the Chinese
Pharmacopoeia (Liu et al. 2000). Baicalin is a component in numerous traditional Chinese medicine (TCM) formulae, and has been widely used clinically to treat fever, bronchitis and upper respiratory tract infection (Havsteen 2002; Zhang et al. 2016; Zhang et al. 2013). Previous researchers observed that oral administration of baicalin to rats at a dose of 20 mg/kg indicated the
34 presence of baicalein 6-O-β-d-glucopyranuronoside in the plasma suggesting that baicalin is directly absorbed from the gastrointestinal tract (Akao et al. 2013). However, previous research conducted by the same group on germ-free rats revealed that only a small amount of the metabolite was detected in plasma, indicating the influence of microbiome on the biotransformation and absorption of baicalin (Akao et al. 2000). In addition, researchers have also identified that subsequent to oral baicalin administration, glucuronides and sulfates of baicalein were also observed in plasma. Moreover, when comparing the rates of relative of absorption of baicalin and baicalein (aglycone form), baicalin had 65% relative absorption rate, whereas the relative absorption rate of baicalein was negligible. However, intravenous administration of baicalin had an improved level of baicalein metabolites, indicating the hepatic contribution to their presence
(Lai et al. 2003).
Baicalin has been found to possess significant antibacterial activity against methicillin- resistant Staphylococcus aureus, Helicobacteri pylori, and Escherichia coli (Huang et al. 2015;
Novy et al. 2011; Zhou et al. 2016). In a mouse model of H. pylori, researchers have observed that baicalin and baicalein were capable of significantly inhibiting the pathogen in the murine stomach
(Chen et al. 2014; Chen et al. 2018). Previously, researchers have identified that baicalin improved renal function and significantly reduced Stx2-induced lethality in mice. Further, structural and biophysical analyses revealed that baicalin directly binds Stx to inactivate the toxin and favor the formation of toxin oligomers (Dong et al, 2015). Moreover, the same research group challenged mice with Enterohemorrhagic E. coli O157:H7 and showed that baicalin was capable of reducing lethality in mice, however, the exact mechanism behind the reduced pathogenesis is unclear
(Zhang et al. 2017).
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Considering the mechanism of antimicrobial activity, previous researchers have documented that flavonoid compounds exhibit antibacterial activity by inhibiting nucleic acid synthesis (Ohemeng et al. 1993), cytoplasmic membrane integrity (Mori et al. 1987) as well as energy metabolism (Haraguchi et al. 1998). In addition, Yun and coworkers (2012) identified that the aglycone form of baicalin (baicalein) inhibited Staphylococcus aureus by compromising membrane permeability, cell respiration, protein synthesis and DNA topoisomerase activity to exert its antibacterial function (Yun et al. 2012).
6. Probiotic therapy
Probiotics are defined as live microorganisms, which when consumed in appropriate amounts confer a health benefit on the host (Araya et al. 2002; Hill et al. 2014). The US Food and
Drug Administration has classified probiotic microorganisms as generally recognized as safe
(GRAS) (Hotel and Cordoba 2001). Probiotics are generally prescribed as an adjunct therapy with antibiotics to maintain gut microbiota, which could be disrupted as a result of antibiotic administration. Previous research has shown that probiotic administration significantly reduced the recurrence of CDI in infected patients (Crow et al. 2015). However, several theories have been hypothesized to explain their role in protecting against various enteric pathogens including C. difficile. The protective action of probiotics is mainly attributed to their role in inhibiting or modulating pro- or anti-inflammatory signaling pathways in the gut epithelium (Kumari et al.
2011; Patel and Lin 2010).
Several clinical trials have identified that the administration of multi-strain probiotics to
CDI patients have proven to be safe and effective (Goldenberg et al. 2017; Mills et al. 2018). In a decade long, observational study conducted in Quebec, hospital inpatients were supplemented with a probiotic mixture of L. acidophilus CL1285, L. casei LBC80R, and Lactobacillus rhamnosus
36
CLR2 (Bio-K+) within 12 hours of receiving an antibiotic dosage. The results revealed that the incidence of CDI reduced from 18.0 cases per 10,000 patient days to an average of 2.3 cases per
10,000 patients (Maziade et al. 2015). Likewise, in another controlled trial, the incidence of CDI was studied in hospital patients undergoing antibiotic therapy with a cocktail of Lactobacillus casei, L. bulgaricus and Streptococcus thermophilus or a placebo control. There were no reports of CDI in the probiotic cocktail group as opposed to 17% incidence in the placebo control group
(Hickson et al. 2007). Based on the recent Cochrane systematic review on the preventive efficacy of probiotics against CDI in adults and children, short-term use of probiotics appeared to be safe and effective when used along with antibiotics in patients, who are not immunocompromised or severely debilitated (Goldenberg et al. 2017).
In summary, C. difficile is a major nosocomial pathogen causing toxin-mediated enteritis and severe diarrhea in humans. C. difficile predominantly affects hospital inpatients undergoing protracted antibiotic treatment, which results in gastrointestinal dysbiosis, leading to C. difficile spore germination, pathogen colonization in the intestine and subsequent toxin production. Most of the antibiotics used for treating various diseases, including anti- C. difficile antibiotics are found to predispose patients to CDI and its relapse by inducing gastrointestinal dysbiosis. Moreover, the global emergence of antibiotic-resistant strains of hypervirulent C. difficile warrants the identification alternative therapeutic strategies against CDI.
The goal of this Ph.D. dissertation is to identify novel and safe strategies that have the potential to be developed as an alternate/adjunct therapy to antibiotics for controlling CDI without adversely affecting the gut microbiome. Based on published literature and preliminary research conducted in our laboratory, it is hypothesized that sodium selenite, baicalin and select lactic acid bacteria attenuate major C. difficile virulence determinants, including toxin production and
37 sporulation. Further, it is hypothesized that sodium selenite decreases C. difficile resistance to ciprofloxacin and vancomycin. The specific objectives of this dissertation include
1. To investigate the effect of sodium selenite on C. difficile toxin production, spore germination, and resistance to ciprofloxacin and vancomycin in vitro.
2. To investigate the effect of baicalin on C. difficile toxin production, sporulation and spore germination in vitro.
3. To study the effect of selected lactic acid bacteria on C. difficile toxin production and spore germination in vitro.
4. To determine the effect of baicalin on C. difficile pathogenicity in a mouse model.
38
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59
CHAPTER III
In vitro efficacy of sodium selenite on toxin production, spore outgrowth and antibiotic
resistance in hypervirulent Clostridium difficile
60
Abstract
This study investigated the efficacy of essential mineral, selenium (sodium selenite) in reducing toxin production, spore outgrowth and antibiotic resistance of Clostridium difficile in vitro. Two hypervirulent C. difficile strains were cultured in brain heart infusion broth with and without the sub-minimum inhibitory concentration of sodium selelnite, and the supernatant and bacterial pellet were harvested for total toxin quantitation and RT-qPCR analysis of toxin- encoding genes, respectively. Additionally, C. difficile strains were cultured in brain heart infusion broth containing 0.5 or 1 x minimum inhibitory concentration (MIC) of either ciprofloxacin or vancomycin with or without the sub-MICs of sodium selenite (0.14 mM, 2.1 mM or 3.6 mM).
Sodium selenite significantly reduced C. difficile toxin production, cytotoxicity and spore outgrowth (p<0.05). Also, the expression of toxin production genes, tcdA and tcdB were downregulated in the presence of sodium selenite (P<0.05). Further, sodium selenite significantly increased C. difficile sensitivity to ciprofloxacin (P<0.05), but not vancomycin, as revealed by a decreased bacterial growth in samples containing the ciprofloxacin + selenium compared to antibiotic control. These results suggest that sodium selenite could potentially be used to control
C. difficile, and warrant future studies in vivo.
61
1. Introduction
Clostridium difficile is an anaerobic, spore-forming, bacterial pathogen causing a toxin- mediated enteric disease in humans [1-3]. More than 500,000 cases of C. difficile infection (CDI) are reported annually in the United States, resulting in more than $6.3 billion as healthcare costs
[4]. C. difficile mostly affects hospitalized individuals particularly the elderly undergoing prolonged antibiotic therapy [5]. Prolonged antibiotic therapy disrupts the normal gut microbiota, leading to C. difficile spore germination, pathogen colonization and subsequent toxin production and release [1,6]. The toxins, TcdA and TcdB, are the major virulence factors of C. difficile that disrupt intestinal epithelial integrity and elicit strong inflammatory response leading to severe colonic inflammation eventually resulting in pseudomembranous colitis [7, 8,9]. The emergence of a hypervirulent C. difficile strain, NAP1/ribotype 027 that is capable of producing increased levels of toxins has been associated with CDI outbreaks worldwide in hospital and community settings [7,8,10]. The Centers for Disease Control and Prevention (CDC) in its recent report on emerging pathogens with antibiotic resistance listed C. difficile as one of the three urgent threats to human health [11].
Although broad-spectrum antibiotics predispose patients to CDI by disrupting the normal gut microbiota [12,13], antibiotics continue to be the drug of choice for treating the disease in patients.
Additionally, the worldwide emergence of antibiotic resistance in hypervirulent C. difficile strains further limits the continued use of antibiotics for treating CDI [14,15]. Since the toxins are the major virulence factors responsible for the pathogenesis of CDI, the use of antivirulence agents that inhibit CD toxin production could be a potential strategy for controlling CDI in patients.
Essential minerals are nutrients that are required by living organisms for performing physiological functions. Metals that form a significant part of essential minerals have historically
62 been used as antimicrobial agents [16]. However, with the advent of antibiotics, the application of metals as antimicrobial agents in human medicine began to decline. With the emergence of multidrug-resistant pathogens and reduced number of antibiotics being discovered, metals have received increased attention in recent years, especially as potent antimicrobial agents against antibiotic-resistant pathogens [17,18]. Numerous transition metals and metalloids, such as silver, copper, selenium, and tellurium have been identified as effective antimicrobial and antibiofilm agents against bacterial pathogens such as Staphylococcus aureus, Escherichia coli, Salmonella and Pseudomonas aeruginosa [18,19,20,21].
Selenium is a naturally occurring essential microelement critical for various biological functions in the body, including enzymatic and antioxidant activities [17]. The commonly used dietary supplement, sodium selenite (Na2SeO3), is proven to possess antifungal and antibacterial properties [22,23]. Moreover, sodium selenite has been reported to increase the sensitivity of methicillin-resistant Staphylococcus aureus (MRSA) to antibiotics such as oxacillin, cloxacillin, ampicillin/sulbactam and neomycin [17]. The objective of the current study was to determine the in vitro efficacy of selenium in reducing toxin production, spore outgrowth and increasing antibiotic susceptibility in C. difficile.
2. Materials and Methods
2.1. Bacterial Strains and Culture Conditions
Two hypervirulent C. difficile cultures (ATCC BAA 1870 strain and 1803 isolate) were grown in brain heart infusion broth (Difco, Sparks, MD) supplemented with 5g/L yeast extract
(BHIS) in a Don Whitley A35 anaerobic workstation (Don Whitley Scientific, West Yorkshire,
63
UK) with atmospheric conditions of 80% nitrogen, 10% hydrogen and 10% carbon dioxide at 37°C for 24 h [24]. The bacterial population was determined by plating 100 µL volumes of suitable dilutions on BHIS agar, and Clostridium difficile moxalactam norfloxacin (CDMN) agar (Oxoid,
Hampshire, UK) supplemented with 5% horse blood, under strict anaerobic conditions at 37 °C for 24 h. Moreover, six selected beneficial bacteria obtained from the USDA-ARS culture collection – Lactococcus lactis ssp. lactis, Lactobacillus rhamnosus, Lactobacillus delbrueckii bulgaricus, Lactobacillus reuteri, Lactobacillus brevis and Lactobacillus plantarum were separately grown in de Man, Rogosa and Sharpe (MRS) broth (Remel Inc., Lenexa, KS) under anaerobic conditions at 37°C for 24 h. The growth of each isolate was determined by measuring the optical density using Synergy plate reader (Biotek, Winooski, VT) at 600 nm during 0 h, 6 h,
12 h and 24 h time points [24].
2.2. Establishment of sub-minimum inhibitory concentration and minimum inhibitory concentration of sodium selenite and antibiotics
The highest concentration of sodium selenite that did not significantly affect bacterial growth was considered as the sub-minimum inhibitory concentration (sub-MIC). The minimum inhibitory concentration (MIC) was the lowest concentration that inhibits bacterial growth after 24 h of incubation. The sub-MIC of sodium selenite and MIC of antibiotics (ciprofloxacin and vancomycin) against C. difficile were determined by tube broth dilution assay, as previously reported [25]. Sodium selenite (Sigma Aldrich, St. Louis, MO, USA), Ciprofloxacin hydrochloride
(Alfa Aesar, Haverhill, MA, USA) and Vancomycin hydrochloride (Sigma Aldrich, St. Louis, MO,
USA) stock solutions were prepared in sterile deionized water. The samples groups for different concentrations of sodium selenite and antibiotics were incubated in an anaerobic workstation at
37°C for 24 h, and bacterial growth was enumerated by serial dilution and plating. Duplicate
64 samples were included, and the experiment was replicated three times with each strain. For the selenium-antibiotic combination experiments, we used 0.5x MIC and MICs of antibiotics, and the
1x sub-MIC, 15x and 25x sub-MIC of sodium selenite. The concentrations of sodium selenite used in the current study were based on preliminary experiments conducted in our laboratory. Similarly, the effect of sub-MIC of sodium selenite on the growth of the aforementioned beneficial gut bacteria was determined by culturing them separately in 10 mL of MRS broth under anaerobic conditions at 37 °C with or without the sodium selenite for 24 h. The growth of each culture was determined by measuring optical density at 600 nm.
2.3. Effect of sodium selenite on C. difficile Toxin Production and Cytotoxicity
Brain Heart Infusion broth supplemented with 5g/L of yeast extract (BHIS) with or without the sub-MIC of sodium selenite was inoculated (~105 CFU/ml) separately with each C. difficile isolate and incubated at 37°C for 48 h anaerobically, as previously mentioned. The culture supernatants were collected at 24 h and 48 h of incubation for total toxin A and B quantitation by
ELISA [26] and for determining cytotoxicity on Vero cells [24,27]. The bacterial pellets were harvested at 12 h by centrifugation (14,000 x g for 10 min) for RNA isolation for RT-qPCR analysis of C. difficile genes associated with toxin production [24].
2.4. ELISA for Total Toxin A and B
The amount of toxin in the culture supernatants at 24 h and 48 h was quantitated using the
Wampole ToxA/B II kit (TechLabs, Inc., VA, USA), as described previously [26]. Purified toxin
B (Sigma Aldrich) was used to plot a standard curve. The culture supernatants were diluted, and
ELISA was performed according to the manufacturer’s instructions. The optical densities at 450
65 nm were compared with the linear range of the standard curve, and total toxin concentration was estimated [24].
2.5. Cytotoxicity Assay
The effect of sodium selenite on the cytotoxicity of C. difficile culture supernatant was determined by Vero cell cytotoxicity assay, as previously described [24,28]. C. difficile culture supernatant from the 24 h and 48 h incubation were serially diluted (1:2) and added onto confluent
Vero cell monolayers in 96-well microtiter plates. Subsequently, the plates were incubated in a 5%
CO2 incubator at 37°C for 24 h and observed for cytopathic changes under an inverted microscope.
Positive reactions were indicated by the characteristic Vero cell rounding accompanied by parallel neutralization of cytotoxicity with Clostridium sordellii antitoxin (TechLabs, Inc., VA, USA). The cytotoxicity titer was considered as the highest well dilution showing 80% cell rounding, and titer values were expressed as the reciprocal of the identified dilution.
2.6. Real-Time Quantitative PCR (RT-qPCR)
To analyze the effect of sodium selenite on C. difficile genes involved in toxin production, total RNA was isolated from early stationary phase (12 h) cultures. The cultures from each treatment group were collected by centrifugation at 14,000 × g for 10 min at 4 °C. The bacterial pellet was resuspended in RNAwiz solution (Ambion, Austin, TX), flash frozen in liquid nitrogen, and stored at −80°C. Total RNA was extracted using the Ambion RiboPure Bacteria RNA kit, followed by DNase I digestion using Turbo DNase I (Ambion). The RNA derived from each
DNase I digestion was purified further using the Qiagen RNeasy RNA column purification kit.
The cDNA was produced using the Bio-Rad iScript cDNA synthesis kit (Bio-Rad, Hercules, CA).
RT-qPCR analysis of the toxin production associated genes was performed using published
66 primers [29] normalized against 16S rRNA gene expression. Twenty microliter reactions were performed in triplicate using Bio-Rad iTaq Universal SYBR green supermix (Bio-Rad, Hercules,
CA). The relative fold change in gene expression was calculated using the 2−ΔΔCt method [30].
2.7. Effect of sodium selenite on C. difficile spore germination and outgrowth
Freshly grown ATCC BAA 1870 and 1803 single colonies were separately inoculated into
BHIS broth and cultured overnight at 37 °C under anaerobic conditions. Anaerobically pre- reduced BHIS agar prepared in six-well plates were inoculated with 150 µL aliquots of the overnight culture, spread evenly and incubated anaerobically at 37 °C for 10 days in an anaerobic workstation (Don Whitley Scientific, West Yorkshire, UK) to allow sporulation. Subsequently, spores were collected from the six-well plates by flooding with 2 ml ice-cold sterile water. The spore suspension was subjected to heat-shock at 60 °C for 20 min to kill any vegetative cells and washed six times in distilled water by centrifuging at 14, 000 x g for 5 min. Spore suspensions were examined for purity by phase-contrast microscopy before storage at −20 °C before use [27,31].
To assess the germination of C. difficile isolates, 100 µL suspension containing 105 spores/mL was added to the wells of a 12-well plate containing 1.9 mL of pre-warmed, pre-reduced
BHIS supplemented with 0.1% sodium taurocholate (Sigma Aldrich, St. Louis, MO), separately incorporated with the sub-MIC of sodium selenite inside an anaerobic workstation. The plates were closed inside the anaerobic workstation with lids and sealed with a sealant. BHIS media without taurocholate, and spore suspensions or culture media replaced with dH2O were included as controls. A well with resazurin (0.1 mg/mL) was included for examining anaerobiasis in 12 well plates during reading. The optical density of the spore-medium mixture in the wells at 600 nm was recorded using Synergy plate reader (Biotek, Winooski, VT) at 37°C, over a 24 h period with readings taken at 10 min intervals and expressed as a percentage of the initial OD600 (t/t0). The
67 spore germination was measured as the initial loss of OD600 and spore outgrowth was measured by recording the increase in OD600 followed by spore germination, as described previously [32,33].
2.8. Effect of sodium selenite on antibiotic resistance in C. difficile
To determine whether sodium selenite increased the antibiotic sensitivity of C. difficile, sub-MIC, 15 x sub-MIC and 25 x sub-MIC of sodium selenite were added separately to duplicate tubes containing 10 mL of BHIS inoculated with ~ 5.0 to 5.5 log10 CFU of C. difficile and supplemented with each antibiotic separately at 0.5 x MIC or MIC. Suitable controls were included such as C. difficile (positive control), C. difficile with 0.5 x MIC of antibiotics, C. difficile with
MIC of antibiotics and C. difficile with one of the three sub-MICs of sodium selenite. The culture tubes were incubated at 37°C for 24 h in an anaerobic workstation. Bacterial growth was monitored by serial dilution and plating; duplicate samples were included, and the experiment was replicated three times.
2.9. Statistical analysis
All experiments were performed as duplicates and repeated three times. The data were analyzed using PROC MIXED of SAS v 9.3 and differences between means were considered significantly different at p<0.05.
3. Results
3.1. Sub-minimum inhibitory concentrations and Minimum inhibitory concentration of sodium selenite and antibiotics
The sub-MIC of sodium selenite against both C. difficile strains was 0.14 mM (25 μg/mL).
The 15x and 25x sub-MIC of sodium selenite were 2.1 mM (375 μg/mL) and 3.6 mM (625 μg/mL),
68 respectively. The MIC for Ciprofloxacin against C. difficile was 100 μg/mL and 90 μg/ mL for
BAA 1870 and BAA 1803, respectively. Also, the MIC of Vancomycin was 2.5 μg/mL and 2
μg/mL against BAA 1870 and BAA 1803, respectively. The OD600 values of the six selected beneficial bacterial isolates cultured in the presence of sub-MIC of sodium selenite were not significantly different from their respective controls (Figure 1), indicating that the sub-MIC of sodium selenite was non-inhibitory to the growth of selected beneficial bacteria (p>0.05).
3.2. Effect of Selenium on C. difficile toxin production
The effect of sodium selenite on C. difficile toxin production was determined by ELISA, as described previously [24,26]. Sodium selenite significantly reduced toxin production in both C. difficile isolates at 24 h and 48 h compared to controls (p < 0.05). There was a reduction of approximately 85% in toxin production in both isolates, and the results are shown in Figure 2 (A
& B).
3.3. Effect of Selenium on C. difficile toxin-mediated cytotoxicity on Vero cells
The efficacy of sodium selenite in reducing C. difficile induced cytopathic effects was evaluated on Vero cells based on a previously published protocol [24,27]. Sodium selenite treated
C. difficile culture supernatants had reduced cytotoxicity on Vero cells compared to the culture supernatants of untreated cells. (p < 0.05). Cytotoxicity was reduced by ~ 90% at 48 h compared to the control (Figure 3 A & B).
3.4. Effect of sodium selenite on toxin regulatory genes
In order to delineate the effect of sodium selenite on the expression of C. difficile toxin coding genes, transcriptional analysis by qPCR was performed. Sodium selenite significantly reduced the expression of genes critical for toxin production (tcdA and tcdB) in both C. difficile
69 isolates (p < 0.05). A significant downregulation was also observed in the expression of tcdR, the positive regulator of toxin production in both isolates treated with sodium selenite. Additionally, sodium selenite significantly upregulated the expression of tcdC, which is a negative regulator of toxin production that translates the TcdR antagonist (p <0.05) (Figure 4 A & B).
3.5. Effect of sodium selenite on C. difficile spore germination and outgrowth
The effect of sub-MIC sodium selenite on C. difficile spore germination and outgrowth over a 24 h period was determined. For both isolates, spore outgrowth was observed in untreated wells, as indicated by an increase in OD 600 beginning at hour 11 of incubation, with a continued increase over the 24 h duration (Figure 5 A & B). However, the sub-MIC of sodium selenite completely inhibited spore outgrowth in both isolates, as shown by a lack of increase in absorbance at 600 nm (p<0.05). Also, there was no significant difference in the initial dip in OD600 during spore germination for control, and sodium selenite treated groups, suggesting that sodium selenite did not affect C. difficile spore germination process [34] (Figure 5 A & B).
3.6. Effect of sodium selenite on C. difficile antibiotic resistance
To determine the effect of sodium selenite on C. difficile sensitivity to two selected antibiotics, three sub-MIC levels of sodium selenite were separately combined with 0.5 x MIC or the MIC of either ciprofloxacin or vancomycin. Bacterial growth in selenium-antibiotic treatment groups was compared with that in antibiotic control for the respective experiment (i.e., with either
0.5 or 1x MICs of ciprofloxacin or vancomycin alone treatments). When C. difficile isolates were cultured in the presence of ciprofloxacin and sodium selenite, bacterial growth after 24 h was significantly decreased for all the treatment combinations compared to that in antibiotic control (p
<0.05) (Figure 6A & B). Additionally, when C. difficile isolates were cultured in the presence of
70 vancomycin and sodium selenite, bacterial growth after 24 h was significantly reduced in cultures treated with 0.5 x MIC of vancomycin and any of the three sodium selenite treatments (p <0.05).
However, there was no appreciable reduction in bacterial growth when the MIC of vancomycin was combined with the different levels of sodium selenite (Figure 7A & B).
4. Discussion
Prolonged antibiotic therapy and the resultant gut dysbiosis are critical factors that increase susceptibility to CDI. Such a milieu favors C. difficile colonization, production and release of exotoxins, TcdA and TcdB in the colon, which lead to increased gut permeability, cytokine and chemokine release, leukocyte infiltration and release of reactive oxygen intermediates, consequently damaging the intestinal mucosa [35,36,37]. Therefore, reducing C. difficile toxin production is critical in controlling CDI pathogenesis in affected patients.
Targeting critical virulence mechanisms in pathogens is a relatively new strategy that aims at reducing microbial pathogenesis rather than their growth. Such molecules that target pathogen virulence mechanisms are neither bacteriostatic nor bactericidal, and the likelihood of developing bacterial resistance against these molecules is minimal since they exert a lesser selection pressure
[38]. Moreover, these targets would be marginally deleterious to the host gut microbiota [38-40].
Therefore, we investigated the efficacy of sodium selenite as an alternative therapeutic agent that could reduce the virulence of C. difficile by reducing exotoxin production. For these experiments, we used the sub-minimum inhibitory concentrations (sub-MIC) of sodium selenite, which is defined by the National Committee for Clinical Laboratory Standards, as the concentration of an antimicrobial agent that is not inhibitory to the bacterial growth, but is still effective in altering bacterial physiology and shape in vitro and in vivo, and thus reducing bacterial virulence [41,42].
The antivirulence compounds at sub-MIC could potentially be used as an adjunct therapy in
71 combination with currently practiced antibiotic regimens to reduce disease progression and improve the prognosis in patients.
In our study, we observed that the sub-MIC of sodium selenite against C. difficile growth did not inhibit the growth of six different species of endogenous bacteria (p >0.05) commonly found in the human gastrointestinal tract, which included the genera Lactococcus and
Lactobacillus, that play a vital role in gut health maintenance [43]. Research by Kasaikina and coworkers suggested that dietary selenium supplementation in conventionalized germ-free mice could increase the overall gut microbial diversity with differential effects on specific phylotype abundance [44], indicating that Se supplementation may not adversely affect beneficial bacterial populations and cause gut dysbiosis.
Our results indicate that sub-MIC of sodium selenite significantly reduced total toxin production in both C. difficile isolates tested, compared to untreated controls (p <0.05) (Figure 2).
Moreover, the culture supernatants from sodium selenite-treated C. difficile cultures significantly reduced the toxin-mediated cytopathic effects on Vero cells (p <0.05) (Figure 3), which effectively translates our C. difficile toxin ELISA result at cellular level. Previous research conducted in our lab revealed that sodium selenite reduced extracellular and intracellular verotoxin concentration in
E. coli O157:H7 [45]. Additionally, published research from our laboratory demonstrated that sodium selenite could significantly reduce Vibrio cholerae motility, cell-adhesion, and cholera toxin production in vitro, and ex vivo [46]. Since the sub-MICs of sodium selenite were used in our experiments, the attenuation of virulence factors observed was not due to C. difficile growth inhibition, but it could be attributed to its potential ability to reduce the transcription of virulence genes associated with toxin production. Therefore, we performed an RT-qPCR to determine the effect of sodium selenite on the transcription of genes involved in toxin synthesis, regulation and
72 secretion. Our data revealed a significant downregulation in C. difficile toxin-encoding genes (tcdA and tcdB), and positive regulator gene of toxin production (tcdR) in sodium selenite treated cultures. Interestingly, tcdC, the negative regulator of toxin production in C. difficile pathogenicity locus, and codY, a global regulator of virulence and a potent repressor of tcdR, was found to be significantly upregulated in sodium selenite treated cultures (p <0.05). These results suggest that sodium selenite inhibits C. difficile toxin production by downregulating toxin-encoding genes and modulating toxin regulatory genes. Previous research suggests that sodium selenite could promote oxidative stress and DNA damage [17] by various mechanisms. Moreover, previous studies have demonstrated that sodium selenite gets reduced spontaneously in the presence of thiol group- containing proteins. These chemical reactions could induce oxidative posttranslational modifications that could significantly impact the function of critical C. difficile proteins [47,48].
Spore germination and outgrowth of newly germinated C. difficile spores in the gut are crucial for initiation of CDI pathogenesis [49,50]. Therefore inhibiting C. difficile germination and spore outgrowth is critical in controlling CDI in humans. Results demonstrated that sodium selenite had no effect on C. difficile spore germination (p >0.05). However, the sub-MIC of sodium selenite significantly inhibited C. difficile spore outgrowth compared to controls (p <0.05). These data suggest that vegetative cells from newly germinated spores are apparently more sensitive to sodium selenite compared to vegetative C. difficile cells grown in broth (Figure 5). Previous reports suggest that generation of reactive oxygen species by metal compounds inhibits spore germination in agriculturally important fungal organisms [51,52]. Likewise, accumulation of reactive oxygen species in newly germinated nascent vegetative C. difficile could be a plausible mechanism for inhibiting C. difficile spore outgrowth.
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The effect of sodium selenite on C. difficile antibiotic resistance against two commonly used antibiotics that are clinically important in the therapy against CDI was also investigated.
Fluoroquinolones are broad-spectrum antibacterial agents extensively used against various infectious diseases and have high bioavailability and safety profile [53]. However, the recent emergence of C. difficile ribotypes resistant to fluoroquinolones has limited their use in the treatment against CDI [54-56]. Further, researchers have identified that hypervirulent, fluoroquinolone-resistant C. difficile overexpress toxin production genes in the presence of ciprofloxacin [57]. Similarly, although vancomycin is the first-line antibiotic used to treat moderate to severe CDI, C. difficile clinical isolates with reduced susceptibility to vancomycin have recently emerged [2]. Therefore, in this study, the efficacy of sodium selenite in increasing
C. difficile sensitivity to ciprofloxacin and vancomycin was investigated in vitro.
We observed that sodium selenite significantly increased the sensitivity of C. difficile BAA
1870 and BAA 1803 to ciprofloxacin (p <0.05). However, the effect of sodium selenite on C. difficile vancomycin sensitivity was minimal. Previously Vasić et al. (2011) reported that sodium selenite enhanced antibiotic susceptibility (ampicillin, streptomycin and sulfamethoxazole– trimethoprim) of Bacillus subtilis, B. mycoides, E. coli and Pseudomonas spp. in vitro [58].
Likewise, previous research conducted in our laboratory found that SIC of sodium selenite increased the sensitivity of multi-drug resistant A. baumannii to multiple antibiotics, including ciprofloxacin [59]. Although our results reveal a statistically significant effect of sodium selenite in increasing C. difficile sensitivity to ciprofloxacin (p < 0.05), the reduction in bacterial counts was marginal (growth reduction by 1.1-2.2 log10 CFU/mL). Therefore, the practical significance of the combination therapy with ciprofloxacin and sodium selenite to treat human CDI needs to be investigated in vivo.
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To conclude, our aforementioned results indicate that sodium selenite is effective in reducing C. difficile toxin production. Sodium selenite mediated reduction in toxin production also translated to cellular level on Vero cells. Additionally, sodium selenite significantly reduced C. difficile sporulation and spore outgrowth. Furthermore, sodium selenite increased the sensitivity of C. difficile to ciprofloxacin. These findings suggest the potential use of sodium selenite as an adjunct therapy for CDI in humans, however, in vivo studies are required to validate these findings.
The upper tolerable level and the no-observed-adverse-effect level (NOAEL) of Se are 400 µg and
800 µg, respectively [60]. The highest antivirulence concentration of sodium selenite used in this study falls below the aforementioned safety levels. Further, the intended application of sodium selenite is limited only for the treatment duration of CDI. However, long-term studies on the safety and efficacy of sodium selenite for the treatment of CDI are warranted.
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45. Surendran-Nair, M.; Kollanoor-Johny, A.; Ananda-Baskaran, S.; Norris, C.; Lee, J.; Venkitanarayanan, K. Selenium reduces enterohemorrhagic Escherichia coli O157: H7 verotoxin production and globotriaosylceramide receptor expression on host cells. Future microbiology 2016, 11, 745-756. 46. Bhattaram, V.; Upadhyay, A.; Yin, H.; Mooyottu, S.; Venkitanarayanan, K. Effect of dietary minerals on virulence attributes of Vibrio cholerae. Frontiers in microbiology 2017, 8. 47. Jacob, C. Redox signalling via the cellular thiolstat. Biochem Soc Trans 2011, 39, 1247-1253. 48. Jacob, C.; Jamier, V.; Ba, L.A. Redox active secondary metabolites. Curr Opin Chem Biol 2011, 15, 149-155. 49. Riggs, M.M.; Sethi, A.K.; Zabarsky, T.F.; Eckstein, E.C.; Jump, R.L.; Donskey, C.J. Asymptomatic carriers are a potential source for transmission of epidemic and nonepidemic Clostridium difficile strains among long-term care facility residents. Clin Infect Dis 2007, 45, 992- 998. 50. Sun, X.; Wang, H.; Zhang, Y.; Chen, K.; Davis, B.; Feng, H. Mouse relapse model of Clostridium difficile infection. Infect Immun 2011, 79, 2856-2864. 51. Shi, X.; Li, B.; Qin, G.; Tian, S. Mechanism of antifungal action of borate against Colletotrichum gloeosporioides related to mitochondrial degradation in spores. Postharvest Biol Technol 2012, 67, 138-143. 52. Wu, Z.; Yin, X.; Lin, Z.; Bañuelos, G.S.; Yuan, L.; Liu, Y.; Li, M. Inhibitory effect of selenium against Penicillium expansum and its possible mechanisms of action. Curr Microbiol 2014, 69, 192-201. 53. Tomé, A.M.; Filipe, A. Quinolones. Drug safety 2011, 34, 465-488. 54. Saxton, K.; Baines, S.D.; Freeman, J.; O'Connor, R.; Wilcox, M.H. Effects of exposure of Clostridium difficile PCR ribotypes 027 and 001 to fluoroquinolones in a human gut model. Antimicrob Agents Chemother 2009, 53, 412-420. 55. McDonald, L.C.; Killgore, G.E.; Thompson, A.; Owens, R.C.,Jr; Kazakova, S.V.; Sambol, S.P.; Johnson, S.; Gerding, D.N. An epidemic, toxin gene-variant strain of Clostridium difficile. N Engl J Med 2005, 353, 2433-2441. 56. Pépin, J.; Saheb, N.; Coulombe, M.; Alary, M.; Corriveau, M.; Authier, S.; Leblanc, M.; Rivard, G.; Bettez, M.; Primeau, V. Emergence of fluoroquinolones as the predominant risk factor for Clostridium difficile–associated diarrhea: a cohort study during an epidemic in Quebec. Clinical Infectious Diseases 2005, 41, 1254-1260. 57. Aldape, M.J.; Packham, A.E.; Nute, D.W.; Bryant, A.E.; Stevens, D.L. Effects of ciprofloxacin on the expression and production of exotoxins by Clostridium difficile. J Med Microbiol 2013, 62, 741-747. 58. Vasić, S.; Radojević, I.; Pešić, N.; Čomić, L. Influence of sodium selenite on the growth of selected bacteria species and their sensitivity to antibiotics. Kragujevac Journal of Science 2011, 33, 55-61.
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Figure 1. Effect of Sub-minimum inhibitory concentration of sodium selenite (0.14 mM) on the growth of six beneficial bacteria: (A) Lactococcus lactis lactis; (B) Lactobacillus rhamnosus; (C) Lactobacillus delbrueckii bulgaricus; (D) Lactobacillus reuteri; (E) Lactobacillus brevis; (F) Lactobacillus plantarum . The dotted line indicates the control group and the dashed line indicates sodium selenite treated group. Sodium selenite treated beneficial bacterial populations did not change significantly from the controls (P> 0.05).
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Figure 2. Effect of sodium selenite on C. difficile toxin production* p < 0.05. Brain heart infusion- yeast extact with or without the Sub-minimum inhibitory concentration (sub-MIC) of sodium selenite (0.14mM) was inoculated (105 CFU/mL) separately with two hypervirulent C. difficile isolates ATCC BAA 1870 (A) or ATCC BAA 1803 (B) and incubated anaerobically at 37 °C for 48 h. The culture supernatant from groups Control (black bar graph) and Sodium selenite treated (white bar graph) were collected at 24, and 48 h of incubation for assessing total toxin quantity by ELISA. * Treatments significantly differed from the control (p < 0.05).
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Figure 3. Effect of sodium selenite (Se) on C. difficile induced cytotoxicity on Vero cells. Brain heart infusion with or without the Sub-minimum inhibitory concentration (sub-MIC) of sodium selenite (0.14mM) was inoculated (105 CFU/mL) separately with two hypervirulent C. difficile isolates, ATCC BAA 1870 (A) or ATCC BAA 1803 (B), and incubated anaerobically at 37 °C for 48 h. The culture supernatant from groups sodium selenite and control were harvested at 24, and 48 h of and the cytotoxicity titer on Vero cells in 96-well plates were determined. Serially diluted (1:2) C. difficile culture supernatants were added to the 96-well plates and incubated at 37 °C under 5% CO2 for 24 h. Vero cell cytotoxicity was detected by the presence of cell rounding under an inverted microscope.
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Figure 4. Effect of sodium selenite on C. difficile toxin regulatory genes. BHIS broth with or without the sub-minimum inhibitory concentration (sub-MIC) of sodium selenite (0.14 mM) was inoculated separately with two hypervirulent C. difficile isolates, ATCC BAA 1870 (A) or ATCC BAA 1803 (B) and incubated anaerobically at 37 °C for 12 h. Bacterial pellets were harvested at early stationary phase for gene expression analysis. *Treatments significantly differed from the control group (p < 0.05).
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Figure 5. Effect of sodium selenite on germination and outgrowth of C. difficile BAA 1870 (A) and 1803 (B) spores. A spore suspension containing 105 spores/ml was added to pre-reduced BHI supplemented with 0.1 % sodium taurocholate and incubated with 0 mM or 0.14 mM sodium selenite inside an anaerobic workstation. Optical density at 600 nm (OD 600) was recorded and is expressed as a percentage of the initial OD 600 (t/t0). Germination was measured as the initial loss of OD 600 and spore outgrowth was measured by recording the increase in OD 600 following spore germination.
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Fig. 6. Effect of sodium selenite on antibiotic sensitivity of C. difficile BAA 1870 (A) and 1803 (B) to 0.5 x MIC and MIC of ciprofloxacin. Approximately 105 CFU/ml of vegetative C. difficile was added to pre-reduced BHI containing 0.5 x MIC or 1 x MIC of ciprofloxacin with any of the three concentrations of sodium selenite: sub-MIC (0.14 mM), 15x sub-MIC (2.1 mM) and 25x sub-MIC (3.6 mM). * Treatments significantly differed from the antibiotic controls (0.5x or 1x MIC of ciprofloxacin) (p < 0.05).
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Fig. 7. Effect of sodium selenite on antibiotic sensitivity of C. difficile BAA 1870 (A) and 1803 (B) to 0.5 x MIC and MIC of vancomycin. Approximately 105 CFU/ml of vegetative C. difficile was added to pre-reduced BHI containing 0.5 x MIC or 1 x MIC of vancomycin with any of the three concentrations of sodium selenite: sub-MIC (0.14 mM), 15x sub-MIC (2.1 mM) and 25x sub-MIC (3.6 mM). * Treatments significantly differed from the antibiotic controls (0.5x or 1x MIC of vancomycin) (p < 0.05).
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CHAPTER IV
Effect of Baicalin on C. difficile toxin production, sporulation and spore germination in
vitro
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Abstract
Clostridium difficile is a nosocomial, spore-forming pathogen that causes a serious toxin- mediated enteric disease in humans. The major virulence factors, toxin A (TcdA) and toxin B
(TcdB), promote the disruption of cytoskeleton and intestinal epithelial tight junctions, thereby leading to severe morbidity and mortality. Moreover, sporulation and spore germination are virulence components critical for transmission and relapse of the disease. Therefore, reducing the aforementioned virulence traits could significantly minimize C. difficile pathogenicity and disease outcome in affected individuals.
This study investigated the efficacy of a natural flavone glycoside, baicalin in reducing toxin production, sporulation and spore germination in C. difficile in vitro. Additionally, the effect of baicalin on the growth of six selected endogenous gut bacteria was determined. Hypervirulent
C. difficile strains BAA 1870 or BAA 1803 (5 log CFU/ml) were cultured in Brain Heart Infusion
(BHI) broth with yeast extract (5g/L) with or without the sub-inhibitory concentration (SIC) of baicalin (0.16 mM), and incubated at 37°C for 24 h under strict anaerobic conditions. The supernatant was harvested after 24 h for determining C. difficile toxin production using C. difficile
ToxA/B ELISA kit. In addition, a similar experiment was performed wherein samples were harvested for assessing total viable counts and heat-resistant spore counts (survivors of incubation at 60°C for 20 min) by serial dilution and plating on BHI agar plates at 72 h. Moreover, to determine the effect of baicalin on C. difficile germination and spore outgrowth, C. difficile spores were seeded in germination medium with or without the SIC of baicalin, and spore germination and spore outgrowth were measured by recording optical density at 600 nm. Additionally, the effect of baicalin on C. difficile toxin, sporulation and virulence associated genes was investigated using real-time qPCR.
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The SIC of baicalin did not inhibit C. difficile growth after 24 h of incubation at 37°C, however significantly reduced toxin production, where total toxin production was reduced by ~
85% when compared to control (P < 0.05). Also, a significant decrease in spore counts was observed at 72 h of incubation in baicalin-treated samples compared to control (P<0.05), wherein the SIC of baicalin reduced spore counts by ~1.2 log c.f.u.ml−1 in both isolates. Baicalin did not inhibit C. difficile spore germination; however, completely inhibited spore outgrowth. In addition,
C. difficile genes critical for pathogenesis were significantly down-regulated (P < 0.05) in the presence of baicalin. Notably, the antivirulence concentration of baicalin exerted no inhibitory effect on the growth of tested beneficial bacteria. Collectively, the aforementioned findings suggest the potential efficacy of baicalin for controlling C. difficile; however, follow up in vivo studies are warranted to validate the results.
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1. Introduction
Clostridium difficile is an anaerobic, spore former that produces a toxin-mediated enteritis in humans (Spigaglia 2016; Weese 2010). Gut dysbiosis as a result of protracted antibiotic therapy in hospitalized patients is a significant factor that promotes the development of C. difficile infection
(CDI) (Hookman and Barkin 2009). Although extended duration of antibiotic therapy induces gut dysbiosis, thereby predisposing patients to CDI, antibiotics still remain as the treatment of choice against the disease (Bartlett 1992; O'Connor et al. 2004). Moreover, incidences of emerging hypervirulent, antibiotic resistant C. difficile strains are increasing worldwide, and this further highlights the need to devise novel therapies against CDI (Centres for Disease Control and
Prevention (US) 2013; Spigaglia 2016).
The initial phase of C. difficile colonization requires non-toxin related factors such as flagellar proteins, adhesion factors, hydrolytic and proteolytic enzymes that aid the bacterium to adhere to the predilection site, colonize and initiate pathogenesis (Hennequin et al. 2003; Janoir et al. 2007; Janoir 2015; Kato et al., Arakawa 2005; Tasteyre et al. 2001). As a result of gut dysbiosis,
C. difficile is capable of colonizing and producing major virulence factors, Toxin A (TcdA) and
Toxin B (TcdB) that induce severe gut inflammation in affected individuals. These toxins exhibit glucosyl transferase activity capable of inactivating the host cell Rho family GTPases involved in cytoplasmic F-actin regulation, consequently promoting the disruption of cytoskeleton and gut epithelial tight junctions (Keel and Songer 2006; von Eichel-Streiber et al. 1999). This process triggers an inflammatory response favoring the release of cytokines and leukotrienes, which when prolonged ensues into pseudomembranous colitis and watery diarrhea (Dial et al. 2005; Hookman and Barkin 2009; McDonald et al. 2006; Sunenshine and McDonald 2006). In addition, C. difficile spores are highly resistant structures that promote transmission through feco-oral route in patients
91 or cause relapse in temporarily recovered patients (Burns et al. 2010). Therefore, therapeutic agents that can reduce C. difficile virulence, especially toxin production, sporulation as well as spore germination and outgrowth in the human gut would effectively help to control CDI.
Antivirulence therapy is an approach currently gaining interest for controlling antibiotic resistant infectious diseases. This treatment modality primarily attenuates bacterial virulence rather than growth, which is critical for promoting infection or disease in the host (Khodaverdian et al. 2013;
Rasko and Sperandio 2010). Since antivirulence agents are neither bacteriostatic nor bactericidal, there is a reduced tendency for pathogen resistance development (Cegelski et al. 2008; Clatworthy et al. 2007; Hung et al. 2005; Mellbye and Schuster 2011), with the plausibility of minimally affecting the host beneficial microbiota.
Plant derived compounds have been historically used in traditional medicine to treat various diseases (Kollanoor Johny et al. 2010; Upadhyay et al. 2014). Baicalin (5,6-dihydroxy-7-
O-glucuronide flavone) is a flavone glycoside present in Scutellaria baicalensis Georgi, which is commonly used in traditional Chinese medicine to treat a wide range of infectious diseases such as respiratory tract infections, pneumonia, scarlet fever, jaundice, hepatitis, and dysentery (Liu et al. 2000). The objective of this study was to investigate the potential of baicalin to reduce toxin production, sporulation, spore germination and outgrowth, which are significant in the pathogenesis and relapse of C. difficile infection.
2. Materials and methods
2.1. Bacterial Strains and Culture Conditions
Two hypervirulent C. difficile isolates (ATCC BAA 1870 and 1803) were grown in brain heart infusion broth (Difco, Sparks, MD) supplemented with 5g/L yeast extract (BHIS) in a Don Whitley
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A35 anaerobic workstation (Don Whitley Scientific, West Yorkshire, UK) under atmospheric conditions of 80% nitrogen, 10% hydrogen and 10% carbon dioxide at 37°C for 24 h (Mooyottu et al. 2014). The bacterial population in the cultures was determined by plating 100 µL volumes of suitable dilutions on BHIS agar, and Clostrodium difficile moxalactam norfloxacin (CDMN) agar (Oxoid, Hampshire, UK) supplemented with 5% horse blood, under strict anaerobic conditions at 37 °C for 24 h. Moreover, six beneficial lactic acid bacteria (LAB) obtained from the
USDA-ARS culture collection – Lactococcus lactis ssp. lactis, Lactobacillus rhamnosus,
Lactobacillus delbrueckii bulgaricus, Lactobacillus reuteri, Lactobacillus brevis and
Lactobacillus plantarum were separately grown in de Man, Rogosa and Sharpe (MRS) broth
(Remel Inc., Lenexa, KS) under anaerobic conditions at 37°C for 24 h. The growth of each LAB isolate with or without the sub-inhibitory concentration (SIC) of baicalin was determined by measuring the optical density at 600 nm during 0 h, 6 h, 12 h and 24 h of incubation (Mooyottu et al. 2014).
2.2. Determination of sub-inhibitory concentration of baicalin
The highest concentration of baicalin that does not affect bacterial growth was considered as its sub-inhibitory concentration (SIC). The SIC of baicalin against C. difficile was determined by broth dilution assay, as previously reported (Kollanoor Johny et al. 2010). Baicalin (Indofine
Chemicals, Hillsborough, NJ) was dissolved in BHIS broth to a final concentration of 0.2% and filter sterilized before use for each experiment. BHIS broth containing various concentrations of baicalin were inoculated with C. difficile (~105 CFU/ml) and incubated in an anaerobic workstation at 37°C for 24 h, and bacterial growth was enumerated by serial dilution and plating. Duplicate samples were included, and the experiment was replicated three times with each strain.
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2.3. Effect of baicalin on C. difficile Toxin Production and Cytotoxicity
BHIS broth with or without the SIC of baicalin was inoculated (~105 CFU/ml) separately with each C. difficile isolate and incubated at 37°C for 48 h anaerobically, as previously described
(Merrigan et al. 2010; Mooyottu et al. 2014). The culture supernatants were collected at 24 h and
48 h of incubation for total toxin A and B quantitation by ELISA and for determining cytotoxicity on Vero cells. In addition, bacterial pellets were also harvested at 12 h for RNA isolation for RT- qPCR analysis of C. difficile genes associated with toxin production, sporulation and critical virulence factors.
2.4. ELISA for Total Toxin A and B
The amount of toxin in culture supernatants at 24 h and 48 h was quantitated using the
Wampole ToxA/B II kit (TechLabs, Inc.,VA, USA), as described previously (Merrigan et al. 2010;
Mooyottu et al. 2014). Purified toxin B (Sigma Aldrich) was used to plot a standard curve. The culture supernatants were diluted and ELISA was performed according to the manufacturer’s instructions. The optical density measured at 450 nm was compared with the linear range of the standard curve, and total toxin concentration was estimated (Mooyottu et al. 2014).
2.5. Cytotoxicity Assay
The effect of baicalin on the cytotoxicity of C. difficile culture supernatant was determined by
Vero cell cytotoxicity assay (Baines et al. 2005). C. difficile culture supernatant from 24 h and 48 h was serially diluted (1:2) and added onto confluent Vero cell monolayers in 96-well microtiter plates. Subsequently, the plates were incubated in a 5% CO2 incubator at 37°C for 24 h and observed for cytopathic changes under an inverted microscope. Positive reactions were indicated by the characteristic Vero cell rounding accompanied by parallel neutralization of cytotoxicity with Clostridium sordellii antitoxin (TechLabs, Inc., VA, USA). The cytotoxicity titer was
94 considered as highest well dilution showing 80% cell rounding, and titer values expressed as the reciprocal of the identified dilution.
2.6. Effect of baicalin on C. difficile sporulation
The effect of baicalin on C. difficile sporulation was determined by a previously published protocol (Babakhani et al. 2012; Mooyottu et al. 2017). Briefly, BHIS broth with or without the
SIC of baicalin was inoculated separately with each C. difficile isolate (105 CFU/ml) and incubated at 37°C for 72 h anaerobically. Samples were harvested at 72 h for quantitation of heat-resistant spores (survivors of heating at 60°C for 20 minutes) and total viable count (TVC) by serially diluting each sample in PBS and plating each on duplicate on BHIS agar supplemented with 0.1% taurocholate.
2.7.Effect of baicalin on C. difficile spore germination and outgrowth
Freshly grown ATCC BAA 1870 and 1803 single colonies were separately inoculated into
BHIS broth and cultured overnight at 37°C under anaerobic conditions. Anaerobically pre-reduced
BHIS agar prepared in six-well plates were inoculated with 150 µL aliquot of the overnight culture was spread evenly and incubated anaerobically at 37 °C for 10 days in an anaerobic workstation
(Don Whitley Scientific) to induce sporulation. Subsequently, spores were collected from the six- well plates by flooding with 2 ml ice-cold sterile water. The spore suspension was subjected to heat-shock at 60°C for 20 min to kill any vegetative cells and washed six times in sterile distilled water by centrifuging at 14, 000 g for 5 min. Spore suspensions were examined for purity by phase- contrast microscopy before storage at −20 °C (Baines et al. 2009; Mooyottu et al. 2017).
To assess the germination of C. difficile isolates, 100 µL suspension containing 105 spores/mL was added to the wells of a 12-well plate containing 1.9 mL of pre-warmed, pre-reduced
BHIS supplemented with 0.1% sodium taurocholate (Sigma Aldrich, St. Louis, MO), and
95 separately added with the SIC of baicalin inside an anaerobic workstation. The plates were closed inside the anaerobic workstation with lids and sealed with a sealant. BHIS media without taurocholate, and spore suspensions or culture media replaced with dH2O were included as controls. A well with resazurin (0.1 mg/mL) was included for examining anaerobiasis in 12 well plates during reading. The optical density of the spore-medium mixture in the wells at 600 nm was recorded using Synergy plate reader (Biotek, Winooski, VT) at 37°C, over a 24 h period with readings taken at 10 min intervals, and expressed as a percentage of the initial OD600 (t/t0). The spore germination was measured as the initial loss of OD600 and spore outgrowth was measured by recording the increase in OD600 followed by spore germination, as described previously (Allen et al. 2013; Paredes-Sabja et al. 2008).
2.8.Real-Time Quantitative PCR (RT-qPCR)
To analyze the effect of baicalin on C. difficile genes involved in toxin production, sporulation and other secondary virulence factors, total RNA was isolated from early stationary phase (12 h) cultures. The cultures from each treatment group were collected by centrifugation at 14,000 × g for 10 min at 4 °C. The bacterial pellet was resuspended in RNAwiz solution (Ambion, Austin,
TX), flash frozen in liquid nitrogen, and stored at −80°C. Total RNA was extracted using the
Ambion RiboPure Bacteria RNA kit, followed by DNase I digestion using Turbo DNase I
(Ambion). The RNA derived from each DNase I digestion was purified further using the Qiagen
RNeasy RNA column purification kit. The cDNA was produced using the Bio-Rad iScript cDNA synthesis kit (Bio-Rad, Hercules, CA). RT-qPCR analysis of the toxin genes, sporulation genes and secondary virulence associated genes was performed using published primers (Babakhani et al. 2012; Deneve et al. 2009; Saujet et al. 2011; Soutourina et al. 2013; Vohra and Poxton 2011) and normalized against 16S rRNA gene expression. Twenty microliter reactions were performed
96 in triplicate using iTaq SYBR (Bio-Rad, Hercules, CA). The relative fold change in gene expression was calculated using the 2−ΔΔCt method (Livak and Schmittgen 2001).
2.9. Statistical analysis
All experiments were performed as duplicates and repeated at least three times. The data were analyzed using PROC MIXED of SAS v 9.3 and differences between means were considered significantly different at p<0.05.
3. Results
3.1. Sub-inhibitory concentration of baicalin and its effect on beneficial bacteria
The sub-inhibitory concentration of baicalin against both C. difficile strains was 1.6 mM
(70 μg/mL). The counts of six selected beneficial bacterial isolates cultured in the presence of SIC of baicalin were not different from their respective controls (Figure 1), indicating that the SIC of baicalin was also non-inhibitory to the growth of beneficial bacteria.
3.2. Effect of baicalin on C. difficile toxin production
Baicalin significantly reduced toxin production in both hypervirulent isolates of C. difficile at
24 h and 48 h (p < 0.05). In comparison to control C. difficile cultures grown without baicalin, a reduction of ~ 70-85% in toxin concentration was observed in the presence of baicalin (Figure 2
A & B).
3.3. Effect of baicalin on C. difficile toxin-mediated cytotoxicity on Vero cells
The efficacy of baicalin in reducing C. difficile induced cytopathic effects was conducted on Vero cells, where a reduced ability of C. difficile culture supernatants to produce Vero cell cytotoxicity compared to culture supernatants of untreated cells was observed (p < 0.05). Results
97 revealed that cytotoxicity in baicalin-treated Vero cells was decreased by approximately 85% compared to control (Figure 3 A & B).
3.4. Effect of baicalin on C. difficile sporulation
The effect of SIC of baicalin on C. difficile spore production is shown in Figure 4 (A & B).
In both C. difficile isolates, a significant decrease in spore counts was observed after 72 h of incubation in baicalin treated samples compared to control (p<0.05). For both isolates, baicalin at
1.6 mM resulted in approximately 1.2 log10 CFU/mL reduction in heat shock spore counts compared to control (p<0.05) (Figure 4 A & B). However, the differences in total viable counts
(plated without heat treatment) between baicalin treated and untreated control samples were negligible (p>0.05).
3.5. Effect of baicalin on C. difficile spore germination
The effect of SIC of baicalin on C. difficile spore germination and outgrowth over a 24 h period was determined. For both isolates, spore outgrowth was observed in untreated wells, as indicated by an increase in OD600 starting around 11 hours of incubation, with a continued increase over the 24 h duration (Figure 8 A & B). However, the SIC of baicalin completely inhibited spore outgrowth in both isolates, as shown by a lack of increase in absorbance at 600 nm (p>0.05).
Moreover, there was no significant difference in the initial dip of OD600 during spore germination for control and baicalin treated groups, indicating that baicalin did not affect C. difficile spore germination process (p>0.05).
3.6. Effect of baicalin on toxin, sporulation and other virulence associated genes
In order to delineate the effect of baicalin on genes associated with toxin production, sporulation and other virulence factors of C. difficile, transcriptional analysis by qPCR was performed. The transcriptional analysis was performed on samples that were harvested at the early
98 stationary phase (12 h) of growth. Baicalin reduced the transcription of genes critical for toxin production (tcdA and tcdB) in both C. difficile isolates (p< 0.05). A significant downregulation was also observed in the expression of the positive regulator tcdR in both isolates treated with baicalin (Figure 5 A & B). In addition, a down-regulation of tcdC (TcdR antagonist) and codY was observed in the presence of baicalin. A similar trend of down-regulation was also noted in the transcription of all sporulation associated genes (Figure 6 A & B) and four other secondary virulence genes critical for host colonization of C. difficile (Figure 7 A & B).
4. Discussion
Mounting evidence during the past two decades indicates that the stability of human gut microbiome is critical for maintaining optimum health, especially gastrointestinal health. Since disturbances in the gut microbiome following long-term antimicrobial therapy predisposes to C. difficile proliferation and subsequent disease onset, novel strategies against CDI should target the pathogen without adversely affecting the normal gastrointestinal microbiota. Therefore, this study utilized antivirulence strategy against C. difficile, where the SIC of baicalin was evaluated to inhibit the virulence determinants in the pathogen without affecting bacterial growth. As observed, the SIC of baicalin did not inhibit the growth of six different species of endogenous bacteria commonly found in the human gastrointestinal tract (p>0.05) (Figure 1), which included the genera
Lactococcus and Lactobacillus that play a vital role in gut health maintenance (Rupa and Mine
2012). Previous research by Zhang and coworkers reported that intragastric administration of baicalin in mice favoured the growth of Lactobacilli and enhanced the formation of more stable intestinal microbial communities (Zhang et al. 2010), underscoring that baicalin supplementation may not adversely influence beneficial bacterial populations and promote gut dysbiosis.
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TcdA and TcdB, being the major C. difficile exotoxins that significantly contribute to disease pathogenesis (Burns et al. 2010; Dial et al. 2005; Hookman and Barkin 2009; McDonald et al. 2006; Sunenshine and McDonald 2006), drug targets that inhibit synthesis or disrupt the function of C. difficile exotoxins would represent a potential avenue for the development of antivirulence agents against CDI. As observed in Figure 2, toxin production was significantly reduced in both C. difficile isolates when treated with baicalin in comparison to untreated control
(p<0.05). Further, C. difficile culture supernatants from baicalin treated samples significantly decreased cytopathic effects mediated by the exotoxins on Vero cells (p<0.05) (Figure 3), underscoring the potential use of baicalin for controlling CDI. Besides reducing toxin production, results indicated that baicalin exerted a significant inhibition on sporulation and spore outgrowth in both C. difficile isolates (Figure 4 A & B; Figure 5 A & B). This is notable since a decreased spore production in the gut could minimize the transmission and relapse of CDI in patients (Barbut et al. 2009; Burns et al. 2010; Hookman and Barkin 2009).
Since the SIC of baicalin was used to target C. difficile virulence in this study, the observed reduction in exotoxin production and spore formation was not due to growth inhibition, but could be attributed to baicalin’s effect in modulating the expression of genes associated with these virulence traits. Therefore, we performed RT-qPCR to determine the effect of baicalin on transcription of C. difficile genes associated with toxin production regulation, spore formation as well as critical colonization factors in the host. Toxin gene regulation in C. difficile occurs in a gene cluster identified as the ‘pathogenicity locus’ or PaLoc region. The transcription of tcdA and tcdB, which encode the exotoxins is positively regulated by TcdR, an RNA polymerase sigma factor encoded by tcdR. Additionally, tcdE which encodes a holin-like protein called TcdE is required for toxin secretion outside the bacterial cell. The PaLoc region is negatively regulated by
100 an antagonist of TcdR known as TcdC, which is coded by tcdC (McDonald et al. 2006). The codY gene is a factor present outside the PaLoc region that aids to repress toxin gene transcription
(Dineen et al. 2007). The RT-qPCR results revealed a significant downregulation of tcdA and tcdB genes in baicalin-treated C. difficile, which concurred with the ELISA that showed substantial reduction in toxin levels in the presence of baicalin (Figure 6 A & B). However, there was downregulation in TcdC, the negative regulator of toxin genes, as well as CodY, which is a potent repressor of tcdR (p<0.05). This suggest that the anti-toxin mechanism of baicalin did not possibly involve CodY, and follow up research using global transcriptomic approaches would be critical to delineate pathways modulated by the plant compound.
In addition to the down-regulation of critical toxin genes, baicalin was effective in decreasing the transcription of several genes involved in C. difficile sporulation and host colonization. The gene regulation of spore production in C. difficile involves Spo0A, the master regulator, along with SigH and CD2492, an RNA polymerase factor and a histidine kinase, that is involved in the initiation of sporulation. Other auxiliary proteins that are known to exert a regulatory function on sporulation include SpoIIA, SpoIIR and SpoIIID, which are involved in the various stages of sporulation and spore maturation (Babakhani et al. 2012; Dembek et al. 2015;
Saujet et al. 2011). As observed in Figure 7, baicalin was found to repress all sporulation associated genes (p<0.05), thereby corroborating with the sporulation data (Figure 4 and 7).
As observed for most enteric pathogens, C. difficile colonization also requires adhesion factors and various hydrolytic or proteolytic enzymes. slpA gene codes for the S-layer proteins on the bacterium’s outer surface that is required for strong binding to host intestinal tissues and extracellular matrix proteins. Likewise, other cell wall associated factors that augment C. difficile colonization include cwp84 (cell wall protease), fbp68 (fibronectin binding protein) and fliC
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(flagellar protein). These proteins help the pathogen to move to the colonization site and subsequently attach and destroy host tissue extracellular matrix, and favor dissemination of infection (Hennequin et al. 2003; Janoir et al. 2007; Janoir 2015; Kato et al. 2005; Tasteyre et al.
2001). The RT-qPCR data indicated a significant downregulation of colonization associated virulence factors (p<0.05) that are critical for C. difficile motility, adhesion and breakdown of tissue extracellular matrices (Figure 8 A & B). Previous researchers have documented that flavonoid compounds exhibit antibacterial activity by inhibiting nucleic acid synthesis (Ohemeng et al. 1993), cytoplasmic membrane integrity (Mori et al. 1987) as well as energy metabolism
(Haraguchi et al. 1998). In addition, Yun and coworkers identified that the aglycone form of baicalin (baicalein) inhibited Staphylococcus aureus by compromising membrane permeability, cell respiration, protein synthesis and DNA topoisomerase activity to exert its antibacterial function (Yun et al. 2012).
To conclude, this study indicated that baicalin was effective in significantly reducing C. difficile toxin production, sporulation and spore outgrowth without adversely affecting the growth of selected gut beneficial bacteria in vitro. These results suggest the potential use of baicalin in controlling CDI, however, in vivo studies using a valid animal model are warranted before moving to clinical trials.
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Figure 1. Effect of sub-inhibitory concentration of baicalin on selected beneficial gut bacteria growth. Six selected beneficial gut bacteria ( (A) Lactococcus lactis lactis; (B) Lactobacillus rhamnosus; (C) Lactobacillus delbrueckii bulgaricus; (D) Lactobacillus reuteri; (E) Lactobacillus brevis; (F) Lactobacillus plantarum) were grown in de Man, Rogosa and Sharpe broth in anaerobic condition at 37°C with and without SIC of baicalin (1.6 mM or 0.07% w/v) The bacterial growth was monitored by measuring optical density at 600 nm measured at 6, 12 and 24 h. Baicalin-treated gut bacterial populations did not change significantly from the controls (p> 0.05).
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Figure 2. Effect of baicalin on C. difficile toxin production, * P < 0.05. Brain heart infusion-yeast extract broth with or without the SIC of baicalin (1.6 mM or 0.07% w/v) was inoculated (105 CFU/mL) separately with two hypervirulent C. difficile isolates ATCC BAA 1870 (A) or ATCC BAA 1803 (B), and incubated anaerobically at 37 °C for 48 h. The culture supernatant from groups Control (black bar graph) and Baicalin treated (BC; white bar graph) was collected at 24, and 48 h of incubation for total toxin A and B quantitation by ELISA. * Treatments significantly differed from the control (p < 0.05).
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Figure 3. Effect of baicalin (BC) on C. difficile induced cytotoxicity on Vero cells. BHIS with or without the SIC of baicalin (1.6 mM) was inoculated (105 CFU/mL) separately with two hypervirulent C. difficile isolates, ATCC BAA 1870 (A) or ATCC BAA 1803 (B), and incubated anaerobically at 37°C for 48 h. The culture supernatant from baicalin treated and control samples were collected at 24, and 48 h and cytotoxicity titer on Vero cells was determined. Serially diluted (1:2) C. difficile culture supernatants were added to the monolayers in 96-well plates and incubated at 37°C under 5% CO2 for 24 h. The cell morphology was examined under an inverted microscope for characteristic rounding as an indication of cytotoxicity.
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Figure 4. Effect of baicalin on C. difficile sporulation. Brain heart infusion with or without the Sub-inhibitory concentration (SIC) of baicalin (1.6mM) was inoculated (105 CFU/mL) separately with two hypervirulent C. difficile isolates, ATCC BAA 1870 (A) or ATCC BAA 1803 (B), and incubated anaerobically at 37 °C for 72 h. Samples were withdrawn at different times for quantitation of heat-resistant spores (survivors of incubation at 60°C for 20 minutes) and total viable count (TVC) by serially diluting each sample in PBS and plating each in duplicate on BHIS agar supplemented with 0.1% taurocholate. * Treatments significantly differed from the respective controls (p<0.05)
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Figure 5. Effect of baicalin on germination and outgrowth of C. difficile BAA 1870 (A) and 1803 (B) spores. A spore suspension containing 105 spores/ml was added to pre-reduced BHI supplemented with 0.1 % sodium taurocholate and incubated with 0 mM or 1.6 mM baicalin inside an anaerobic workstation. Optical density at 600 nm (OD 600) was recorded and is expressed as a percentage of the initial OD 600 (t/t0). Germination was measured as the initial loss of OD 600 and spore outgrowth was measured by recording the increase in OD 600 following spore germination.
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Figure 6. Effect of baicalin on C. difficile toxin regulatory genes. BHIS broth with or without the sub-inhibitory concentration (SIC) of baicalin (1.6 mM) was inoculated separately with two hypervirulent C. difficile isolates, ATCC BAA 1870 (A) or ATCC BAA 1803 (B) and incubated anaerobically at 37 °C for 12 h. Bacterial pellets were harvested at early stationary phase for gene expression analysis. *Treatments significantly differed from the control group (p < 0.05).
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Figure 7. Effect of baicalin on C. difficile sporulation genes. BHIS broth with or without the sub- inhibitory concentration (SIC) of baicalin (1.6 mM) was inoculated separately with two hypervirulent C. difficile isolates, ATCC BAA 1870 (A) or ATCC BAA 1803 (B) and incubated anaerobically at 37 °C for 12 h. Bacterial pellets were harvested at early stationary phase for gene expression analysis. *Treatments significantly differed from the control group (p < 0.05).
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Figure 8. Effect of baicalin on C. difficile secondary virulence associated genes. BHIS broth with or without the sub-inhibitory concentration (SIC) of baicalin (1.6 mM) was inoculated separately with two hypervirulent C. difficile isolates, ATCC BAA 1870 (A) or ATCC BAA 1803 (B) and incubated anaerobically at 37 °C for 12 h. Bacterial pellets were harvested at early stationary phase for gene expression analysis. *Treatments significantly differed from the control group (p < 0.05).
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CHAPTER V Antivirulence effect of select lactic acid bacteria against Clostridium difficile
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Abstract
Lactic acid bacteria (LAB) as probiotic therapy against C. difficile infection (CDI) may provide an alternative or adjunct strategy to antibiotics for controlling CDI in humans. Since the activity of many LAB is strain specific, characterization of LAB’s efficacy as monocultures or multi-strain cocktails is required for future development as effective treatment formulations against CDI. The objective of this study was to screen LAB from our laboratory culture collection for investigating their putative inhibitory activity against hypervirulent C. difficile. Five LAB isolates were selected for further experiments based on preliminary research. The efficacy of selected LAB to inhibit C. difficile toxin production was performed by culturing them along with the vegetative cells of two hypervirulent C. difficile isolates separately at an inoculation level of 5:5 log CFU/ml. Similarly, spores of C. difficile strains were also cocultured with LAB to assess the effect of beneficial bacteria on spore outgrowth. In the coculture experiments with vegetative C. difficile, although C. difficile growth was minimally inhibited, a significant reduction of at least 50-70% at 24 h and 70-
90% at 48 h in toxin production was observed (p<0.05). Additionally, coculture supernatants with all LAB produced negligible cytopathic effects on Vero cells as compared to supernatants from C. difficile monocultures (p<0.05). Moreover, co-culture of C. difficile spores with LAB isolates showed a significant reduction in spore outgrowth compared to untreated controls (p<0.05).
Among the five LAB isolates, two L. plantarum isolates were found to be most effective against both C. difficile isolates. Overall, the results indicate that all the five tested LAB isolates significantly inhibited the two critical C. difficile virulence factors, namely toxin production and spore outgrowth. However, further studies are needed to characterize the anti-C. difficile components produced by the LAB and validate their efficacy in controlling CDI using suitable animal models.
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1. Introduction
Clostridium difficile (CD) is an anaerobic, Gram-positive spore-former responsible for causing antibiotic-associated diarrhea in humans. Bacterial spore transmission via the feco-oral route with prolonged antibiotic therapy leading to the disruption of gastrointestinal microbiota provide an ideal environment for pathogen colonization resulting in C. difficile infection (CDI) (Bartlett,
1997; Spigaglia, 2016; Weese, 2010). C. difficile exotoxins, TcdA and TcdB, are critical virulence factors possessing glucosyltransferase activity capable of modifying host GTPases, which are required for actin polymerization and cytoskeletal assembly. Although both exotoxins have similar enzymatic activity, TcdA is an enterotoxin, whereas TcdB is a cytotoxin that exerts its biological effect at lower doses than TcdA (Castagliuolo et al., 1998). The CDI symptoms range from mild self-limiting to severe diarrhea, with up to a quarter of the affected individuals experiencing recurrent infections. In severe cases of CDI, the affected colon undergoes pseudomembrane formation progressing to toxic megacolon, mostly leading to severe disease in approximately one- third of the cases (Hookman & Barkin, 2009; Keel & Songer, 2006; Kyne, 2010; McDonald et al.,
2006; Sunenshine & McDonald, 2006; von Eichel-Streiber et al., 1999).
The recent emergence of hypervirulent C. difficile in the United States is related to antibiotic- resistant strains capable of inducing increased cytotoxicity, particularly the North American isolate
BI/NAP1/027 (Blossom & McDonald, 2007; Hookman & Barkin, 2009; Sunenshine & McDonald,
2006). Despite the increased epidemiology of antibiotic-resistant C. difficile strains, antibiotics ironically remain to be the primary treatment option against CDI (Bartlett, 1992; O'Connor et al.,
2004), with metronidazole, vancomycin and fidaxomicin are the most commonly used drugs
(Bagdasarian et al., 2015). Although these antibiotics have been found effective against C. difficile, the treatment especially for prolonged course could potentially lead to gut dysbiosis predisposing
117 patients to recurrence of infection. Further, C. difficile strains resistant to one or more of these antibiotics could emerge. In light of these concerns, there is a need for novel, alternate/adjuncts to antibiotics for controlling CDI.
Probiotics are defined as live microorganisms, which when consumed in appropriate amounts confer a health benefit on the host (Araya et al., 2002; Hill et al., 2014). The US Food and Drug
Administration has classified probiotic microorganisms as generally recognized as safe (Hotel &
Cordoba, 2001). Lactic acid bacteria (LAB), especially Lactobacillus spp., and Bifidobacteria are the most commonly used probiotic bacteria since they are considered an integral and desirable member of the intestinal microbiota (Kailasapathy & Chin, 2000; Schrezenmeir & de Vrese, 2001;
Soccol et al., 2010). Probiotic microbes provide multiple health benefits to the host, including improved nutrient digestion and assimilation, potentiating host immune function, and protection against enteric pathogens (Fukuda et al., 2011; Hill et al., 2014; Olszak et al., 2012; Soccol et al.,
2010; Sonnenburg et al., 2005). Probiotics potentially constitute an alternate approach to prevent and/or treat CDI, with several reports on the correction of gut dysbiosis by administration of probiotics or prebiotics (Colombel, 1987; Kondepudi et al., 2014; Kotowska et al., 2005; Lewis et al., 2005; Plummer et al., 2004; Rätsep et al., 2017; Segarra-Newnham, 2007). Antagonistic mechanisms of probiotics are suggestive of C. difficile virulence interference, which include toxin inactivation and modulation of inflammatory responses (Chen et al., 2006; Ripert et al., 2016;
Trejo et al., 2010). Based on the recent Cochrane systematic review on the preventive efficacy of probiotics against CDI in adults and children, short-term use of probiotics appeared to be safe and effective when used along with antibiotics in patients, who are not immunocompromised or severely debilitated (Goldenberg et al., 2017). Since the activity of many LAB is strain specific
(Mills et al., 2018), in vitro and in vivo characterization of LAB’s efficacy as monocultures or
118 multi-strain cocktails (Hell et al., 2013; Kondepudi et al., 2014) need to be investigated for future development as safe treatment formulations. The objective of this study was to screen LAB for investigating their putative inhibitory activity against hypervirulent C. difficile.
2. Materials and methods
2.1.Bacterial isolates and culture conditions
Five lactic acid bacteria selected from preliminary screening for their ability to grow well in
BHIS under anerobic conditions were used in this study. These isolates included Lactobacillus plantarum 900B, Lactobacillus rhamnosus 400B (Nebraska Cultures, Inc. - currently UAS Labs,
Wausau, WI), Lactobacillus plantarum 42-3, Lactobacillus rhamnosus NRRL-B 442 (USDA-
ARS, Peoria, IL) and Lactobacillus paracasei DUP 13076 (Dr. Amalaradjou’s culture collection).
The selected cultures were separately grown in de Man Ros Sharpe (MRS) broth (Difco, Sparks,
MD, USA) under anaerobic conditions at 37°C for 24 h. The bacterial count of these cultures was determined by plating 0.1 ml portions of appropriate dilutions on MRS agar (Difco) and incubated at 37°C for 24 h. The cultures were sedimented by centrifugation (3600 x g, 15 min, 4°C), and the pellets were washed twice, and resuspended in sterile phosphate-buffered saline (pH 7.3) to be used as the inoculum.
Two hypervirulent C. difficile isolates (ATCC BAA 1870 and BAA 1803) were grown in brain heart infusion broth (Difco) supplemented with 5% yeast extract (BHIS) in a Don Whitley A35 anaerobic workstation (Don Whitley Scientific, West Yorkshire, UK) in the presence of 80% nitrogen, 10% carbon dioxide and 10% hydrogen at 37°C for 24 h (Mooyottu et al., 2014). The bacterial population was determined by plating 0.1 ml portions of appropriate dilutions on BHIS
119 agar and Clostridium difficile moxalactam norfloxacin (CDMN) agar (Oxoid, Hampshire, UK) supplemented with 5% horse blood, under strict anaerobic conditions at 37°C for 24 h.
C. difficile spores were prepared as previously described with slight modifications (Sorg &
Dineen, 2009). Briefly, single colonies of ATCC BAA 1870 and 1803 were inoculated separately into BHIS broth and cultured overnight at 37°C under anaerobic conditions. A 150 µl aliquot of the overnight culture was spread onto anaerobically pre-reduced BHIS agar (Oxoid, Hampshire,
UK) in six-well plates and incubated for 10 days at 37°C in an anaerobic workstation to induce sporulation. The spores were harvested from the wells by flooding 2 ml of ice-cold sterile water.
The spore suspension was heat-treated at 60°C for 20 min to kill any vegetative cells and washed six times in dH2O by centrifuging at 16,000 x g for 5 min. Spore suspensions were examined for purity by phase-contrast microscopy before storage at −20 °C (Baines et al., 2009; Mooyottu et al.,
2017).
2.2. Effect of co-culturing of select LAB with C. difficile on toxin production
This experiment was performed based on a previously published protocol with slight modifications (Ratsep et al., 2014; Rätsep et al., 2017; Trejo et al., 2010). Approximately, 105
CFU/mL each of LAB and C. difficile (BAA 1870 or BAA 1803) were co-inoculated into 10 mL of anaerobically pre-reduced BHIS broth. As controls, BHIS broth was separately inoculated with
C. difficile or the respective LAB. The treatment and control tubes were incubated under anaerobiosis for 48 h at 37°C. The LAB and C. difficile cultures were serially diluted and enumerated separately on MRS agar and Clostridium difficile moxalactam norfloxacin (CDMN) agar (Oxoid, Hampshire, UK) supplemented with 5% horse blood, respectively at 0, 10, 24 and 48 h of incubation. At each of the specified times, the pH of control and treatment cultures was measured. The MRS plates were incubated aerobically for 48 h at 37°C (Ambalam et al., 2015),
120 whereas CDMN plates were incubated under anaerobic conditions for 48 h at 37°C. The cell-free supernatant of mono and co-cultures (at 24 and 48 h) was harvested by centrifugation at 14,000 x g for 10 min, and filter-sterilized using a 0.2 µm syringe filter. The amount of toxin in the culture supernatants was quantified using the Wampole Tox A/B II kit (TechLabs, Inc., Blacksburg, VA,
USA). The filter-sterilized supernatant was also assessed for its cytotoxicity on Vero cells.
2.3. ELISA for Total Toxin A and B
The amount of toxin in culture supernatants was quantitated using the Wampole ToxA/B II kit
(TechLabs, Inc.,VA, USA), as described previously (Merrigan et al., 2010; Mooyottu et al., 2014).
Purified toxin B (Sigma Aldrich) was used to plot a standard curve. The culture supernatants were appropriately diluted and ELISA was performed according to the manufacturer’s instructions. The optical density measured at 450 nm was compared with the linear range of the standard curve, and total toxin concentration was estimated (Mooyottu et al., 2014).
2.4. Cytotoxicity assay
The effect of selected LAB on the cytotoxicity of C. difficile culture supernatant was determined by Vero cell cytotoxicity assay (Baines et al., 2005; Mooyottu et al., 2014). The co- culture supernatants from the 24 h and 48 h time points were serially diluted (1:1) and added onto confluent Vero cell monolayers in 96-well microtiter plates. Subsequently, the plates were incubated in a 5% CO2 incubator at 37°C for 24 h and observed for cytopathic changes under an inverted microscope. Positive reactions were noted by characteristic Vero cell rounding accompanied by parallel neutralization of cytotoxicity with Clostridium sordellii antitoxin
(TechLabs, Inc., VA, USA). The cytotoxicity titer was considered as highest in wells dilution showing 80% cell rounding, and titer values were expressed as the reciprocal of the identified dilution.
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2.5. In vitro co-culturing of selected LAB on C. difficile spore outgrowth
This experiment was performed based on a previously published protocol with slight modifications (Rätsep et al., 2017; Woo et al., 2011). Approximately, 105 CFU/mL each of LAB and C. difficile spore were inoculated into 10 mL of anaerobically pre-reduced BHIS broth. As controls, BHIS broth was separately inoculated with C. difficile spores or the respective LAB. The treatment and control tubes were incubated under anaerobiosis for 48 h at 37°C. Immediately after inoculation, the levels for C. difficile spores and LAB were confirmed by serial dilution and plating on BHIS agar supplemented with 0.1% sodium taurocholate and MRS agar, respectively. In addition, C. difficile spores and LAB were enumerated on CDMN agar and MRS agar, respectively at 24 and 48 h of incubation.
2.6.Statistical Analysis
All experiments were performed in duplicate, and the study was repeated three times. The data were analysed using the PROC-MIXED procedure of SAS v. 9.3 (SAS Institute Inc., Cary, NC,
USA) and differences between the means were considered significantly different at p < 0.05.
3. Results
3.1. In vitro co-culturing of selected LAB isolates with vegetative C. difficile
The effect of co-culturing C. difficile isolates with LAB on pathogen growth and toxin production over a 48 h period was determined. The growth of both C. difficile isolates was reduced
(p<0.05) in presence of LAB, with the magnitude of reduction in counts ranging from 0.5-1.0 log10
CFU/mL when compared to C. difficile monoculture (p<0.05) (Figure 1 A & B). It was also observed the pH of cocultures was generally lower than that of the monoculture at each of the three
122 sampling points (P < 0.05) (Table 1). Further, no differences in the growth of LAB were observed when they were grown with C. difficile in comparison to monoculture controls (p < 0.05) (Figure
2 A & B).
3.2. Effect of LAB on C. difficile toxin production and cytotoxicity assay
C. difficile toxin production in the presence of LAB at 24 and 48 h was significantly decreased when compared to untreated C. difficile monoculture (p < 0.05) (Figure 3 A & B). In untreated C. difficile BAA 1870, ~ 2800 and 5,100 ng/ml of total toxin were detected at 24 and 48 h, respectively, compared to respective toxin concentrations of ~ 800 and 1200 ng/ml in the presence of LAB (Figure 3A). A similar trend was observed with C. difficile BAA 1803, where ~
2600 and 4500 ng/ml of toxin were produced at 24 and 48 h, respectively in comparison to 1000 and 600 ng/ml of toxins detected in the presence of LAB, respectively (Figure 3B). Concurring to these results, a reduced cytotoxicity on Vero cells was produced by C. difficile:LAB co-culture supernatants in comparison to supernatants derived from C. difficile monocultures (p < 0.05)
(Figure 4 A & B). The cytotoxicity titer in the presence of LAB-treated C. difficile supernatants was decreased by 80 to 100% when compared to monoculture C. difficile controls.
3.3. In vitro co-culturing of selected LAB on C. difficile spore outgrowth
In both C. difficile isolates, all five LAB isolates significantly inhibited spore outgrowth at 24 h and 48 h compared to untreated control (Figure 5 A & B) (p<0.05). Although the individual LAB isolates slightly differed in their effect, all of them decreased spore outgrowth in C. difficile BAA
1870 by more than 5.0 log CFU/ml at 48 h of incubation. However, in C. difficile BAA 1803, the two L. plantarum isolates (L. plantarum 900B and L. plantarum 42-3) decreased spore outgrowth by > 6.0 log CFU/ml at both time points, although the other three LAB brought about only 2.5 to
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3.0 log reduction in bacterial counts (p< 0.05). Moreover, there was no significant difference in growth of LAB in spore co-cultures when compared to their monoculture controls (p>0.05) (data not shown).
4. Discussion
Probiotic research has undergone significant advancements towards understanding their role in the prevention of various gastrointestinal infections and diseases. Some of the major antimicrobial attributes of probiotics on pathogens include: (a) elaboration of inhibitory products, including organic acids, metabolites, and antimicrobial peptides, (b) outcompeting pathogens for nutrients, (c) anti-invasive activity and colonization resistance (d) bacterial toxin inactivation and
(e) inhibition of quorum sensing (Nair et al., 2017). In particular, previous studies have documented multiple anti-C. difficile activities exerted by LAB, including toxin inactivation, colonization resistance, improving the balance of beneficial microbial population and potentially counteracting gut dysbiosis post-antibiotic treatment (Goldenberg et al., 2017; Hudson et al., 2017;
Theriot & Young, 2015). Although the current national guidelines for CDI treatment do not recommend routine probiotic administration (Mills et al., 2018), several clinical trials have identified that multi-strain probiotic administration in CDI patients receiving antibiotics is safe and effective (Goldenberg et al., 2017; Hell et al., 2013; Mills et al., 2018). Thus, identification of effective probiotics as adjunct therapeutics for alleviating CDI merits investigation for consideration as a potential treatment option. In our study, we screened and evaluated the probiotic potential of selected LAB against C. difficile vegetative cell growth, toxin production and spore outgrowth using published in vitro co-culture models.
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Since coculturing of LAB and C. difficile needs a non-selective medium that supports the growth of both bacteria, twelve LAB from our culture collection were initially screened for their ability to grow in BHIS anaerobically. Based on the results from this experiment (results not shown), five LAB, including Lactobacillus plantarum 900B, Lactobacillus rhamnosus 400B,
Lactobacillus plantarum 42-3 (canine feces isolate), Lactobacillus rhamnosus NRRL-B 442 and
Lactobacillus paracasei DUP 13076 were selected for studying their antagonistic activity against
C. difficile.
Since the toxins produced by C. difficile constitute the most important virulence factor that brings about pathogenesis in patients, we determined the efficacy of selected LAB to inhibit toxin production by culturing them along with C. difficile at an inoculation level of 5 log CFU/ml each in BHIS. Results revealed a significant reduction of at least 50-70% at 24 h and 70-90% at 48 h in toxin production in the co-culture supernatants from both C. difficile isolates (Figure 3 A & B). In parallel with these results, the coculture supernatants with all LAB produced negligible cytopathic effects on Vero cells as compared to supernatants from C. difficile monocultures (Figure 4 A &
B).
Besides the toxins, C. difficile spore germination and outgrowth are critical for CDI transmission and relapse (Deakin et al., 2012). C. difficile spores have the ability to survive in the gut for a prolonged period of time (Paredes-Sabja et al., 2014), leading to spore germination and colonization of C. difficile vegetative cells, which in turn causes new infections or relapse of the infection (Riggs et al., 2007; Sun et al., 2011). Since relapse of CDI has been reported in approximately 25% of the patients undergoing anti-C. difficile therapy (Rupnik et al., 2009), the effect of selected LAB on C. difficile spore outgrowth was determined. As observed in Figure 5
(A & B), co-culture of C. difficile spores with LAB isolates showed a significant reduction in spore
125 outgrowth compared to untreated controls (p<0.05). Among the five LAB, the two L. planatarum isolates were found to be most effective against both C. difficile isolates.
When C. difficile was grown as a monoculture, the bacterial counts increased over time reaching ~ 8.0 log CFU/ml in BAA 1870 and ~ 7.5 log CFU/ml in BAA 1803 at 48 h (Figure 1 A
& B). However, when C. difficile was cocultured with LAB, a marginal decrease in growth was observed, which ranged in magnitude from 0.5 to 1.0 log CFU/mL depending on the LAB present.
Probiotic bacteria are known to produce organic acids that possess inhibitory action against gut pathogens (Fooks & Gibson, 2002; Cook & Sellin, 1998; Makras & De Vuyst, 2006). Previously short-chain fatty acids such as lactate, butyrate, acetate and propionate were documented to exert antimicrobial activity against C. difficile (Kondepudi et al., 2012), and this may have partially contributed to the growth inhibition of C. difficile in cocultures observed in our study. This was further supported by the small decline in the pH of cocultures compared to monocultures of C. difficile (Table 1).
Previously, Woo and co-workers reported that co-culturing C. difficile with the Japanese probiotic strain, Clostridium butyricum MIYAIRI 588 (CBM588), inhibited toxin production and the researchers attributed the low-pH contributed by secreted organic acids to the inhibitory effects on C. difficile growth, toxicity and metabolism (Woo et al., 2011). Likewise, Kolling and group observed that lactic acid produced by Streptococcus thermophilus LMD-9 induced a bactericidal effect on C. difficile. Additionally, sub-inhibitory concentrations of lactic acid were found to significantly downregulate tcdA expression and exotoxin release in C. difficile (Kolling et al.,
2012). Recently, researchers identified that a secretory component harboring protease activity produced by Bacillus clausii strain O/C was capable of inactivating purified C. difficile toxin B
(Ripert et al., 2016). In our study, although the low pH ranging from 5..5-6.0 in the co-culture
126 samples may have contributed in part to the C. difficile growth inhibition and toxin production, the observed pH was not low enough to denature the toxins as the isoelectric pH of the toxins ranges between 4.7-4.8 (Eichel-Streiber et al., 1987). To conclude, results indicate that despite reducing C. difficile growth minimally, the five tested LAB significantly inhibited two critical virulence factors, namely toxin production and spore outgrowth. Therefore, further studies are needed to characterize the antimicrobial components elaborated by the LAB against C. difficile, and validate their efficacy in controlling CDI using in vivo models.
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Table 1. pH values in co-culture experiments when vegetative CD was incubated with LAB isolates in 5 log: 5log inoculation ratio. * Treatments significantly differed from control (p<0.05)
Treatment BAA 1870 BAA 1803 10 hours 24 hours 48 hours 10 hours 24 hours 48 hours C. difficile 5.959+0.035 6.060+0.023 6.010+0.072 (CD) monoculture 6.103+0.022 6.150+0.069 6.069+0.026 CD+ 5.871+0.022 5.718+0.026* 5.789+0.026* Lactobacillus plantarum 900B 5.716+0.053* 5.678+0.026* 5.578+0.116* CD+ 5.856+0.014* 5.758+0.050* 5.918+0.033 Lactobacillus rhamnosus 400B 5.656+0.068* 5.487+0.077* 6.0361+0.055 CD+ 5.654+0.016* 5.733+0.024* 5.763+0.008* Lactobacillus plantarum 42-3 5.927+0.038* 5.640+0.091* 5.681+0.06* CD+ 5.932+0.015 5.919+0.049* 5.934+0.096 Lactobacillus rhamnosus NRRL-B 442 5.984+0.030 5.639+0.095* 5.791+0.06* CD+ 5.858+0.032 5.796+0.033* 5.894+0.062 Lactobacillus paracasei DUP 13076 5.888+0.037 5.736+0.058* 5.890+0.063*
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Figure 1. Effect of co-culturing vegetative CD with LAB on C. difficile growth at 10, 24 and 48 h. Pre-reduced BHIS broth was inoculated in 1:1 ratio with respective CD [(A) BAA 1870; (B) BAA 1803] and LAB isolates and incubated anaerobically for 48 h. The CD counts are expressed as log CFU/mL at time points of 10, 24, and 48 h. *Treatments significantly differed from control (p<0.05)
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Figure 2. LAB counts in co-culture experiment using vegetative CD at 10, 24 and 48 h. Pre- reduced BHIS broth was inoculated in 1:1 ratio with respective CD and LAB isolates and incubated anaerobically for 48 h. (A) LAB count when co-cultured with BAA 1870; (B) LAB counts when co-cultured with BAA 1803. The LAB counts are expressed as log CFU/mL at time points of 10, 24, and 48 h. Treatments did not significantly differ from control (p>0.05)
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Figure 3. Effect of LAB on C. difficile toxin production. Co-culture supernatants harvested from 24 and 48 h of incubation were subjected to total toxin A/B quantitation by ELISA. Supernatants from (A) BAA 1870 and (B) 1803 when co-cultured with LAB isolates. * Treatments significantly differed from the control (p < 0.05).
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Figure 4. Effect of co-culture supernatants of C. difficile isolates BAA 1870 (A) and BAA 1803 (B) on CD induced cytotoxicity on Vero cells. The co-culture supernatants were harvested at 24, and 48 h of incubation and the cytotoxicity titer on Vero cells was determined. Serially diluted (1:2) co-culture supernatants were added to the monolayers in 96-well plates and incubated at 37 °C under 5% CO2 for 24 h. The characteristic cell rounding observed during Vero cell cytotoxicity was analyzed using an inverted microscope.
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Figure 5. Effect of LAB isolates on spore outgrowth of C. difficile isolates (A) BAA 1870 and (B) BAA 1803. Pre-reduced BHIS broth was inoculated in 1:1 ratio with respective CD spores [(A) BAA 1870; (B) BAA 1803] and LAB isolates and incubated anaerobically for 48 h. The CD counts are expressed as log CFU/mL at time points of 24, and 48 h. *Treatments significantly differed from control (p<0.05)
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CHAPTER VI
Effect of baicalin in reducing Clostridium difficile infection in a mouse model
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Abstract This study investigated the prophylactic and therapeutic effect of baicalin (BC), a plant-derived flavone glycoside, in reducing C. difficile infection in a mouse model. Five to six-week-old
C57BL/6 mice were randomly divided into eight treatment groups (challenge and control) of twelve mice each. Mice were fed with irradiated feed and supplemented with baicalin (0, 0.11, and
0.22% w/v) in sterile drinking water. The challenge groups were made susceptible to C. difficile by orally administering an antibiotic cocktail in water and an intra-peritoneal injection of clindamycin. Both challenge and control groups were infected with 106CFU/ml of hypervirulent BAA 1803 spores or PBS, and observed for clinical signs for 10 days. Respective control groups for baicalin, antibiotic, and combination were included for investigating their effect on mouse enteric microbiota. The prophylactic administration of baicalin was initiated twelve days before infection, whereas, therapeutic administration began from day 1 post-infection. Mouse body weight and clinical and diarrhea scores were recorded daily post infection. In the prophylactic study, fecal samples were collected on day 2 post-infection for microbiome analysis using rRNA sequencing in MiSeq platform. Baicalin supplementation significantly reduced the incidence of diarrhea and improved the clinical and diarrhea scores in mice in both prophylactic and therapeutic intervention studies (p < 0.05). Microbiome analysis revealed a significant increase in
Proteobacteria and reduction in the abundance of protective microbiota in antibiotic-treated and C. difficile-infected mice compared to controls (p < 0.05). However, baicalin supplementation positively altered the microbiome composition, as revealed by an increased abundance of beneficial bacteria, especially Lachnospiraceae and Akkermansia. Results justify follow up investigations for using baicalin as an adjunct to antibiotic therapy to control gut dysbiosis and reduce C. difficile infection.
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1. Introduction
Clostridium difficile is an important cause of nosocomial, antibiotic-associated diarrhea around the world (Hookman and Barkin 2009; McFarland 2008). The pathogen causes a toxin-mediated colitis in individuals of all age groups, with more severity observed in elderly and immunocompromised patients (Weese et al. 2011). In the United States, more than 453,000 cases of C. difficile infection (CDI) with 29,000 deaths are reported annually, which incur an economic burden ranging between US$ 0.4-3.0 billion as health-care associated costs (Leffler and Lamont
2015; Napolitano and Edmiston Jr 2017). The increased incidence of CDI in humans is primarily attributed to the emergence of a highly toxigenic and hypervirulent C. difficile strain
NAP1/ribotype 027 (Blossom and McDonald 2007; Hookman and Barkin 2009; Spigaglia 2016;
Sunenshine and McDonald 2006).
Generally, CDI predisposition is observed in individuals requiring long-term antibiotic therapy and gastric acid suppressing agents (Bartlet et al. 1992; Dial et al. 2006; Kelly & LaMont, 1998).
This condition results in disturbances in the normal enteric microbial balance, creating a microbiome shift encompassing reduced Bacteriodetes and Firmicutes in the gut (Ling et al. 2014;
Seekatz and Young 2014; Shahinas et al. 2012; Theriot et al. 2014). The resultant dysbiotic environment is believed to support spore germination, thereby favorably selecting for the proliferation and colonization of C. difficile (Voth and Ballard 2005). The colonized vegetative C. difficile produces potent exotoxins namely, toxin A and toxin B, which harbor glucosyl transferase activity capable of inactivating the host Rho family GTPases associated with F-actin regulation
(Voth and Ballard 2005). This process triggers disruption of cytoskeletal and tight junctions in the gut epithelium, thereby initiating a severe inflammatory response along with the release of cytokines and leukotrienes, eventually leading to severe diarrhea and pseudomembrane formation
141 in the colon (Hookman and Barkin 2009; McDonald et al. 2005; McDonald et al. 2006; Sunenshine and McDonald 2006).
Although extended antibiotic therapy predisposes individuals to CDI, antibiotics are still considered the primary line of treatment against the disease, where metronidazole, vancomycin and fidaxomicin are commonly used for treatment (Cohen et al. 2010; Debast et al. 2014; Leffler and Lamont 2015; Spigaglia 2016). However, recent incidences of emerging C. difficile isolates with significantly reduced susceptibility or even resistance to clinically recommended antibiotics potentially raises a critical concern with the continued use of these agents as a treatment option
(Peng et al. 2018; Spigaglia 2016). With global emergence of antibiotic resistant, hypervirulent C. difficile strains, The Centers for Disease Control and Prevention (CDC) has categorized the pathogen as one among the three urgent threats to public health (Centres for Disease Control and
Prevention, 2013). With these concerns, there is a need to identify alternative therapeutic agents that can reduce C. difficile virulence without adversely affecting the gastrointestinal microbiota.
Phytochemicals represent a natural group of molecules that have been used for treating various diseases in traditional medicine (Wollenweber, 1988). Baicalin (5,6-dihydroxy-7-O-glucuronide flavone) is a flavone glycoside present in the plant, Scutellaria baicalensis Georgi, known to possess antimicrobial, anti-sepsis, antioxidant and anti-inflammatory activities (Chen et al. 2001;
Liu et al. 2000; Novy et al. 2011; Tsou et al. 2016; Wang and Liu 2014; Zhang et al. 2017; Zhu et al. 2012). Previously, our laboratory documented the potential use of baicalin as an anti-C. difficile therapeutic agent due to its inhibitory effect on C. difficile toxin production without affecting the growth of selected beneficial microbiota in vitro (Pellissery et al. 2018). Hence, this study aimed at investigating the prophylactic and therapeutic effect of baicalin against C. difficile in an in vivo model by focusing on the clinical course and host microbiome changes encountered in a murine
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CDI model. Mouse models for CDI are well established, and antibiotic-induced gut dysbiosis in mice can be simulated by administering antibiotics orally and intraperitoneally, followed by inoculation of C. difficile spores (Chen et al. 2008; Sun et al. 2011).
2. Materials and methods
2.1. Ethics statement, animals, and housing
The study was performed with the approval of the Institutional Animal Care and Use
Committee (IACUC) at the University of Connecticut, following the endorsed guidelines for animal care and use. Six-week old C57BL/6 mice were obtained from Charles River (Boston, MA), housed in a biohazard level II AALAC-accredited facility, and monitored for health status twice daily. Mice were provided with irradiated feed, autoclaved water and bedding, along with 12-h light/dark cycles. The procedures that required animal handling (spore administration, cage changes and sample collection) were done under a biosafety cabinet (class II) using proper personal protective equipment. Decontamination and sterilization of the biosafety cabinet was done using
10% bleach to prevent cross-contamination between experimental treatment groups. The mice were singly housed in a cage and twelve cages were included for each treatment in each of the experiments.
2.2. Prophylactic and therapeutic administration of baicalin in a mouse model of C. difficile infection
The in vivo infection model was based on a previously established protocol with minor modifications (Chen et al. 2008). Five to six-week old female animals were randomly assigned to one of the following eight treatment groups of twelve animals each (Table 1). In the prophylactic model, animals were provided irradiated pellet feed and incorportated baicalin in drinking water
143 containing 0%, 0.11% and 0.22% w/v of the compound for a period of twenty two days (Fig. 1).
As equated from the average daily water consumed by each mouse (~ 5-7 mL per day), baicalin treated water was expected to deliver approximately 250 mg/kg and 500 mg/kg of the compound per day in the 0.11% and 0.22% treatments, respectively. Previous researchers have indicated that baicalin dosage of 400 mg/kg is well tolerated by mice (Xi et al. 2015). Subsequently, an antibiotic cocktail comprising of kanamycin (0.4 mg/mL), gentamicin (0.03 mg/mL), colistin (850 U/mL), metronidazole (0.215 mg/mL) and vancomycin (0.045 mg/mL) was added in drinking water for 3 days. After antibiotic supplementation, the mice were switched back to their prior treatment regimens, and all animals in the challenge groups (CD, CD+PBS, CD+BC1 and CD+BC2), and the antibiotic control group (Ant) received a single intraperitoneal injection of clindamycin (10 mg/kg, with a maximum of 0.5 mL/mouse using a 27G needle and syringe) a day prior to C. difficile challenge. Pre-treatment of mice with antibiotics was intended to induce gastrointestinal dysbiosis and enable C. difficile colonization following spore challenge. Mice proposed for C. difficile infection were orally administered 106 spores (CFU) per 0.1 mL total volume of hypervirulent C. difficile ATCC BAA 1803 using a straight 18G gavage needle (1” shaft length), and were observed for signs of CDI, including diarrhea, wet tail and hunched posture using a mouse clinical score sheet (Table 2 a & b).
Individual weight of each mouse was measured every day, fecal samples were collected on alternate days post-infection (DPI), and all animals were observed twice daily for ten days for morbidity and mortality. At the end of the experiment (10th day after challenge), all animals were euthanized. In the therapeutic model, the only difference from the aforementioned procedure is that baicalin was administered from day 1 post-C. difficile spore challenge (1 DPI). In addition, microbiome analysis was not performed in the therapeutic study (Fig. 1).
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2.3. DNA extraction, PCR amplification, and sequencing of taxonomic markers
Fecal samples from day 2 post-infection from all treatment groups (from eight animals per treatment group) of the prophylactic baicalin study were subjected to DNA extraction using the
MoBio PowerMag Soil 96 well kit (MoBio Laboratories, Inc), according to the manufacturer’s protocol for the Eppendorf ep Motion liquid handling robot. Quantification of DNA was performed using the Quant-iT PicoGreen kit (Invitrogen, ThermoFisher Scientific) and DNA was subjected to amplification of partial bacterial 16S rRNA genes (V4 region) from 30 ng of extracted DNA as template, using 515F and 806R primers bound with Illumina adapters and dual indices (8 basepair golay in 3’ and 5’) (Caporaso et al. 2012; Kozich et al. 2013). Amplification was performed in triplicates with the addition of 10 µg BSA (New England BioLabs) using Phusion High-Fidelity
PCR master mix (New England BioLabs). The reaction mixes were incubated at 95°C for 3.5min, and then subjected to PCR reaction for 30 cycles of 30 s at 95.0°C, 30s at 50.0°C, and 90s at
72.0°C, followed by a final extension at 72.0°C for 1 min. Quantification and visualization of pooled PCR products were performed using the QIAxcel DNA Fast Analysis (Qiagen). DNA concentration of the PCR products were normalized to 250-400 bp and pooled using the QIAgility liquid handling robot. Pooled PCR products were cleaned up using the Gene Read Size Selection kit (Qiagen) according to the manufacturer’s protocol and the cleaned pool was subjected to sequencing on MiSeq using v2 2 x 250 base pair kit (Illumina, Inc).
2.4. Sequence Analysis
Microbiome analysis was set up as a completely randomized design with treatments done in replicates of eight. Filtering and clustering of sequences were performed using Mothur 1.36.1 based on a published protocol (Kozich et al. 2013). The operational taxonomic units (OTUs) of samples were clustered at 97% sequence similarity and downstream analysis was done using R
145 version 3.2. The richness and evenness of sample OTUs were calculated by estimating alpha diversity using inverse Simpson diversity index, which were analyzed using Tukey’s Test.
Permutational multivariate analysis (PERMANOVA, adonis function, 75 permutations) was performed to analyze differences in bacterial community composition in the various treatment groups. Test for significance in alpha diversity was determined by ANOVA followed by Tukey’s honest significant differences adjusting for multiple comparisons (p=0.05). NMS ordinations were run in R (v 3.3.0) using metaMDS in the vegan (v 2.3-5) package after calculating the stress scree plots to determine the number of axes required to achieve stress below 0.2, plotted using ggplot2
(v 2.1.0). In addition, the relative abundance of OTUs of major phyla, order and genera was determined to assess the effect of treatment. Tukey’s Test was used to identify changes in groups of bacteria based on treatment and the significance was detected at p<0.05.
2.5. Statistical Analysis
The differences between means between experimental groups across the days were compared by two-way mixed ANOVA using PROC GLM of SAS (v 9.4). “N1” Chi-squared test was used to compare diarrhea incidence rate between to different treatments. The statistical significance level was set at p<0.05.
3. Results
3.1. Effect of baicalin supplementation on the incidence of diarrhea and severity of C. difficile
infection in mice
The prophylactic efficacy of baicalin against CDI in mice was assessed by supplementing the phytochemical in drinking water at two different concentrations (0.11% and 0.22%). Oral administration of 106 CFU/mL C. difficile spores (ATCC BAA 1803) resulted in high morbidity
146 with low mortality in infected mice. In C. difficile infected control groups (Ant+CD), 61% and
85% of animals showed severe diarrhea on 1 DPI and 2DPI, respectively (Fig. 2 A). On 7 DPI, one animal from the Ant+CD group died and no further mortality was recorded in this group.
Although diarrhea continued for five days in the Ant+CD group, there was no increase in percent incidence thereafter. However, there was much lower incidence of diarrhea in the CD+BC1 group with 38% and 31% incidence on 1 DPI and 2 DPI, respectively, with absence of diarrhea on the subsequent days (Fig. 2 A). Moreover, there was no diarrhea in the CD+BC2 group, although there were two mortalities recorded in this group each on days 2 and 3 post-infection.
In the therapeutic model, baicalin supplementation was provided similar to the prophylactic model, but was initiated only from day-1 post-infection. Interestingly, C. difficile positive control group (Ant+CD) did not show any diarrhea on 1 DPI, but an incidence of 62.5% and 87.5% was observed on 2 DPI and 3 DPI, respectively (Fig 2 B). Diarrhea was observed until 5th day post- infection in this group with no increase in percent incidence after 3 DPI. Diarrhea was observed from 1 DPI in CD+BC1 and CD+BC2 groups, although, a significantly reduced incidence of diarrhea was observed for both BC treated groups compared to positive control (p<0.05). An incidence of 25% was observed on both 2 DPI and 3 DPI in CD+BC1 group with no diarrhea thereafter. In CD+BC2 group, the incidence of diarrhea stayed at 14% for days 1-3 post-infection, with no more diarrhea observed for the remainder of experiment duration (Fig 2 B). In addition, there was only one mortality recorded in C. difficile positive control group on 6 DPI.
There were no symptoms of diarrhea observed in the control groups (i.e., negative control
[NC], baicalin control [BC2], antibiotic control [Ant] and antibiotic with baicalin control
[Ant+BC2]) in both the prophylactic and therapeutic BC studies.
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3.2. Effect of baicalin supplementation on clinical score and body weight of C. difficile
infected mice
Clinical scores of animals from different treatment groups were individually recorded using a standard score chart, from 1 DPI to 10 DPI (Table 2a) (Chen et al. 2008). Mice groups receiving prophylactic supplementation of baicalin (CD+BC1 and CD+BC2) had a significantly reduced average clinical score compared to C. difficile positive control (CD) (p<0.05) (Fig 3A). The recovery of surviving morbid animals in C. difficile positive control group was much slower compared to baicalin-treated groups (p<0.05), with clinical resolution observed by 9 DPI.
However, the baicalin supplemented groups showed a dose-dependent reduction in severity with complete recovery observed by 6 DPI (p<0.05). In addition, CD+BC2 group had a significantly lower clinical score compared to CD+BC1 (p<0.05). Interestingly, a similar trend in the average clinical scores was also observed in the therapeutic baicalin study. The clinical scores in CD+BC1 and CD+BC2 groups also followed a dose-dependent reduction in severity (Fig 2 B). However, the recovery rate was much slower compared to the prophylactic study, with complete recovery observed by 9 DPI. Mice in C. difficile positive control group showed a delayed recovery, with no clinical resolution observed even by 10 DPI.
Body weights were recorded on a daily basis post-infection, and the relative percentage weight with respect to the initial weight prior to C. difficile challenge was calculated. In the prophylactic study, baicalin control group [BC2] and Ant+BC2 group showed no significant weight loss compared to negative control. However, mice in the C. difficile positive control [CD] showed significant and progressive weight loss from 1 DPI to 5 DPI compared to negative control
(p<0.05), with animals regaining their pre-challenge body weights by 9 DPI. Although there was no significant difference observed in the average body weights of mice from the BC treated
148 challenge groups (CD+BC1 and CD+BC2) compared to positive control (except for 2DPI for CD vs. CD+BC2, p<0.05), baicalin treated animals were able to rapidly regain their pre-challenge body weights by 5 DPI compared to the C. difficile positive control (9 DPI) (Fig 4A).
In the therapeutic study, C. difficile positive control (CD) also showed a significant weight loss compared to negative controls (p<0.05). Mice in positive control group showed weight reduction 3 DPI through 6 DPI, which returned to their initial body weights by 7 DPI. In addition, there was no significant difference in average percent body weights between the CD group and
CD+BC1 group. However, a significant difference was observed in the average percent body weights of the CD+BC2 groups compared to the CD group from 3 DPI through 6 DPI (p<0.05)
(Fig 3B). Moreover, the CD+BC2 group attained their pre-challenge body weight by 4 DPI, however, a slight delay was observed in the CD+BC1 group, with animals attaining their initial body weight by 6 DPI (Fig. 4B).
3.3. Effect of baicalin supplementation on the gut microbiome of C. difficile infected and non-
infected mice
Prophylactic administration of baicalin revealed distinctive patterns in bacterial taxa composition in the different treatment groups. In negative control group (NC), the predominant phyla groups consisted of Firmicutes and Bacteroidetes in a ratio of 1.05:1, with a minimal proportion of other phyla, including Proteobacteria (Fig 5). In baicalin control group (BC2), we observed a higher proportion of Firmicutes compared to Bacteroidetes having a ratio of 1.79:1.
Antibiotic control group (Ant) had a significantly higher proportion of Proteobacteria compared to negative control and baicalin control group (p<0.05). The supplementation of baicalin along with the antibiotic (Ant+BC2) was not able to the reverse the increase in Proteobacteria, however, there was an increase in the proportion of the phylum Verrucomicrobia compared to antibiotic
149 control group (p<0.05) (Fig 5). The baicalin untreated challenge groups (CD and CD+PBS) had a predominantly higher proportion of Firmicutes and Proteobacteria compared to uninfected controls
(p<0.05). However, baicalin administration to spore challenged groups (CD+BC1 and CD+BC2) reduced the abundance of Firmicutes, and increased the proportion of Proteobacteria compared to antibiotic control, positive control groups (CD and CD+PBS) (p<0.05). A notably distinct phylum that prevailed among baicalin treated, spore challenged (CD+BC1 and CD+BC2) and unchallenged (BC2 and Ant+BC2) groups was Verrucomicrobia, specifically the genus
Akkermansia (p<0.05) (Fig 5).
At the family/genus level, abundance of Enterobacteriaceae was higher in antibiotic control (Ant), C. difficile positive control and PBS control (CD and CD+PBS) groups compared to negative control and baicalin control (BC2) groups (Fig 6B). In addition, Enterobacteriaceae proportion in baicalin treated antibiotic control group (Ant+BC2) was significantly increased compared to antibiotic control (Ant) (p<0.05). The relative abundance of Peptostreptococcaceae was negligible and showed no significant difference in negative control (NC), baicalin control
(BC2) and antibiotic controls (Ant and Ant+BC2 groups) (p>0.05) (Fig 6B). However, in baicalin treated spore challenged groups (CD+BC1 and CD+BC2), Peptostreptococcaceae was significantly reduced compared to the C. difficile positive control and PBS control groups, which had a higher abundance (CD and CD+PBS) (p<0.05). With regards to Lachnospiraceae and
Akkermansia, although not significant, baicalin treated control (BC2) marginally increased their relative abundance compared to the negative control (p>0.05) (Fig 6A). In untreated spore challenge groups (CD and CD+PBS), the abundance of both Lachnospiraceae and Akkermansia was significantly reduced compared to the negative control (NC), baicalin control (BC2) and the baicalin treated antibiotic control (Ant+BC2) (p<0.05). However, with the exception of CD+BC1
150 group, there was a significant increase in the relative abundance of Lachnospiraceae in CD+BC2 group compared to untreated spore challenge groups (CD and CD+PBS) (p<0.05). In terms of relative abundance of Akkermansia, there was a significant increase in both baicalin treated challenge groups compared to untreated spore challenge groups (p<0.05) (Fig 6A). The relative abundance of Lactobacillaceae did not show any significant difference amongst the negative control (NC), baicalin control (BC2) and baicalin treated antibiotic control groups (Ant+BC2)
(p>0.05). In contrast, the antibiotic control and untreated spore challenge groups (CD and
CD+PBS) had a significantly higher abundance of Lactobacillaceae compared to aforementioned controls (p<0.05). In addition, baicalin treated spore challenge groups had a much lower abundance of Lactobacillaceae compared to positive controls (Fig 6A).
The NMDS plot indicating the differential pattern of bacterial diversity revealed a close clustering of baicalin control (BC2) and negative control, suggesting that the species abundance in BC2 group is comparable to untreated negative control. However, the other treatment groups
(antibiotic treated groups, challenged or unchallenged with C. difficile, and with or with BC treatment) did not indicate a typical relationship pattern for the abundance of species present in each sample (Fig 7). The inverse Simpson plot representing the differential pattern of bacterial diversity revealed that BC2 group did not alter the diversity of the gut bacterial community compared to negative control (NC) (p>0.05). However, irrespective of the baicalin treatment, there was a marked reduction in diversity of bacterial communities in C. difficile infected groups and antibiotic controls (Fig 8).
4. Discussion
In the current study, we investigated the prophylactic and therapeutic efficacies of baicalin as alternative antimicrobial agent that can ameliorate CDI without compromising the gut microbial
151 population. Previous research conducted in our laboratory revealed that sub-inhibitory concentration of baicalin reduced C. difficile toxin production and cytotoxicity on Vero cells in vitro. Additionally, baicalin inhibited C. difficile spore germination and outgrowth. The current study demonstrating an improved clinical outcome in baicalin-supplemented mice both in the prophylactic and therapeutic models accentuate the in vitro results. Concurring with the reduced incidence of diarrhea in baicalin-treated C. difficile infected mice (p<0.05) (Fig 2 A & B), a significant reduction in average clinical scores was also observed compared to infected control group (CD) (p<0.05) (Fig 3 A & B). Although the percentage weight loss between positive controls
[CD and CD+PBS] and baicalin treated challenge groups [CD+BC1 and CD+BC2] in the prophylactic study was not significant (p>0.05), baicalin treated mice had an improved and rapid weight gain (Figure 4 A). Moreover, baicalin treated infected mice attained their pre-challenge weights much earlier compared to baicalin untreated positive controls in both prophylactic and therapeutic study (Fig 4 A & B). The reduced CDI severity in baicalin treated mice compared to positive controls could be attributed to the inhibitory effect of baicalin on C. difficile toxin production as observed in our in vitro studies. In addition, baicalin possesses anti-inflammatory and anti-diarrheal properties (Chen et al. 2014; Dou et al. 2012; Ishimaru et al. 1995), which may have contributed to the enhanced clinical outcome observed in the current study.
A normal and healthy gastrointestinal microbiota is key for preventing pathogen colonization, including C. difficile (Britton and Young 2014). Disruption of host gut microbiota as a result of antibiotic therapy is the most important predisposing factor for CDI (Hookman and
Barkin 2009). Antibiotic administration significantly alters microbiome diversity and composition, the effects of which can persist even after the withdrawal of antibiotics (Antonopoulos et al. 2009;
Dethlefsen et al. 2008). The increased risk for CDI susceptibility in the elderly is attributed to the
152 reduction of protective bacterial population such as Firmicutes with simultaneous increase in
Bacteroidetes and undesirable Protoebacteria groups in the gut (Biagi et al. 2010; Claesson et al.
2011; Hopkins et al. 2001).
Microbiome analyses of human CDI patients by previous researchers have identified that
Ruminococcaceae and Lachnospiraceae, as well as butyrate-producing bacteria were significantly depleted in patients with CDI compared to healthy subjects, whereas
Enterococcus and Lactobacillus were more abundant in CDI patients (Antharam et al. 2013). In addition, other researchers have demonstrated a decrease in Enterococcaceae along with an increase in Peptostreptococcaceae, Lactobacillaceae and Enterobacteriaceae in C. difficile positive patients (Buffie et al. 2012; Crobach et al. 2018; Perez-Cobas et al. 2014; Rea et al. 2012;
Schubert et al. 2014; Skraban et al. 2013; Zhang et al. 2015). C. difficile colonization resistance in humans is mainly contributed by abundance of Lachnospiraceae, Ruminococcaceae, and
Bacteroidaceae families (Antharam et al. 2013; Schubert et al. 2014; Schubert et al. 2015). Similar observations in gut microbiome of mice were also observed, wherein an increase in
Lactobacillaceae and Enterobacteriaceae families was noted in susceptible mice that were treated with antibiotics, whereas Lachnospiraceae dominated in animals that remained resistant to CDI
(Reeves et al. 2012). In addition, it has been collectively implicated from several research findings that that a decrease in Lachnospiraceae and Barnesiella, with an increase in
Lactobacillaceae and Enterobacteriaceae is responsible for the loss of colonization resistance against C. difficile (Schubert et al. 2015). Akkermansia genus (phylum Verrucomicrobia) is a strictly anaerobic, Gram-negative bacterium detected in the intestine of most healthy individuals, representing 1–4% of the total microbiota, that is capable of utilizing gut secreted mucin as a sole source of carbon and nitrogen (Collado et al. 2007; Derrien et al. 2011). The only species in this
153 genus, Akkermansia muciniphila, has beneficial effects on metabolism and gut health by exhibiting anti-inflammatory and immunostimulant properties (Derrien et al. 2011; Naito et al. 2018).
Recently, studies have revealed that co-administration of A. muciniphila with polyphenols or prebiotics resulted in improvement of gut barrier function and reduced endotoxemia (Anhê et al.
2016).
In this study, baicalin did not reduce bacterial diversity of the mouse gut microbiome compared to untreated negative control (Fig 7 and 8). Baicalin treatment alone significantly increased the abundance of Firmicutes, especially the members of Lachnospiraceae and to a modest extent, the Lactobacillaceae group, compared to negative control (Fig 5). Antibiotic induced microbiome dynamics observed in the current study are in agreement with the findings reported by previous researchers. Antibiotic pre-treatment significantly increased the abundance of the Lactobacillaceae and Proteobacteria, with a drastic reduction in the Lachnospiraceae (Fig
5, Fig 6 A and B). This change in the microbial composition could be correlated to an increased susceptibility of mice to C. difficile challenge. Although co-administration of baicalin with antibiotics (Ant+BC2) was not able to reverse the abundance of Proteobacteria, a significant increase in Lachnospiraceae and Akkermansia was observed (p<0.05) (Fig 6 A). Therefore, in the
CD+BC1 and CD+BC2 groups, the microbiome shift observed during co-administration of baicalin and antibiotics may have contributed to the colonization resistance against C. difficile.
The untreated spore challenged mice groups (CD and CD+PBS) had invariably showed an increased abundance of Lactobacillus and Proteobacteria due to antibiotic administration, along with an increase in the abundance of Peptostreptococcaceae, the family under which the pathogenic C. difficile are classified (Yutin and Galperin 2013). However, in baicalin treated spore challenge groups, we observed a dose dependent increase in the abundance of Lachnospiraceae
154 and Akkermansia along with a significant reduction in Peptostreptococcaceae (p<0.05) (Fig 6 A
& B). These results suggest that the reduced clinical symptoms and infection in baicalin treated animals could be attributed in part to the beneficial shift in the gut microbiome, especially with the improved abundance of Lachnospiraceae and Akkermansia.
To conclude, our results suggest baicalin supplementation provided protection against C. difficile infection in mice. Baicalin supplementation significantly reduced the incidence of diarrhea as well as the severity of CDI clinical symptoms, besides inducing a favorable shift in the composition of gut microbiota without detrimentally affecting the gut microbiome diversity in mice. However, further functional and metabolomic investigations would be required to identify key association factors between baicalin intervention, metabolic status and the gut microbial community that aided in reducing CDI in mice.
155
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Table 1. Different treatment groups used in the experiment Abbreviations: Ant (Antibiotic); CD (C. difficile); BC (Baicalin); PBS (Phosphate buffered saline)
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Table 2a. Mouse clinical score sheet
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Table 2b. Mouse body condition chart
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Fig. 1. Antibiotic induced murine CDI model
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Fig. 2. Effect of baicalin supplementation on the incidence of diarrhea in mice after CDI Five to six-week-old C57BL/6 mice were randomly divided into eight treatment groups of 8 mice each. Mice were fed with irradiated feed and supplemented with baicalin (0%, 0.11% and 0.22%) in drinking water; the challenge groups were made susceptible to C. difficile by administering an antibiotic cocktail in water and an intra-peritoneal injection of clindamycin. Further, challenge and control groups were infected with 106 CFU/ml of a hypervirulent C. difficile (ATCC 1803) spores or PBS and observed for clinical signs for ten days. The incidence of clinical signs including diarrhea was recorded from 1 DPI to 10 DPI. Groups: (1) CD: administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (2) CD+PBS: phosphate buffered saline in drinking water, administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (3) CD+BC1: Mice provided with baicalin (0.11%), administered with antibiotic cocktail in water and an intra- peritoneal injection of clindamycin, and infected with C. difficile; (4) CD+BC2: Mice provided with baicalin (0.22%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (5) NC: Mice provided with no baicalin, no antibiotics and no C. difficile; (6) BC2 : Mice provided with 0.22% baicalin only; (7) Ant: Mice administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin; (8) Ant+BC2 control: Mice fed with baicalin (0.22%) supplemented water and administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin. (2A) Prophylactic study; (2B) Therapeutic study.
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Fig 2A
Fig 2B
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Fig. 3. Effect of baicalin supplementation on the clinical severity of mice after CDI Five to six-week-old C57BL/6 mice were randomly divided into eight treatment groups of 8 mice each. Mice were fed with irradiated feed and supplemented with baicalin (0%, 0.11% and 0.22%) in drinking water); the challenge groups were made susceptible to C. difficile by administering an antibiotic cocktail in water and an intra-peritoneal injection of clindamycin. Further, challenge and control groups were infected with 106 CFU/ml of a hypervirulent C. difficile (ATCC 1803) spores or PBS and observed for clinical signs for ten days. The incidence of clinical signs including diarrhea was recorded from 1DPI to 10DPI. Groups: (1) CD: administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (2) CD+PBS: phosphate buffered saline in drinking water, administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (3) CD+BC1: Mice provided with BC (0.11%), administered with antibiotic cocktail in water and an intra- peritoneal injection of clindamycin, and infected with C. difficile; (4) CD+BC2: Mice provided with BC (0.22%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (5) NC: Mice provided with no BC, no antibiotics and no C. difficile; (6) BC2 : Mice provided with 0.22% BC; (7) Ant: Mice administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin; (8) Ant+BC2 control: Mice fed with BC (0.22%) supplemented water and administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin (A) Prophylactic study; (B)Therapeutic study *Significant difference between the untreated challenge (only CD group) with the rest of the treatment groups.
167
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Fig. 4. Effect of baicalin supplementation on relative weight loss in C. difficile infected and non- infected mice Five to six-week-old C57BL/6 mice were randomly divided into eight treatment groups of 8 mice each. Mice were fed with irradiated feed and supplemented with baicalin (0%, 0.11% and 0.22%) in drinking water); the challenge groups were made susceptible to C. difficile by administering an antibiotic cocktail in water and an intra-peritoneal injection of clindamycin. Further, challenge and control groups were infected with 106 CFU/ml of a hypervirulent C. difficile (ATCC 1803) spores or PBS and observed for clinical signs for ten days. The body weights of the animals were recorded daily and the relative percentage weight with respect to the initial weight prior to the infection was calculated.. Groups: (1) CD: administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (2) CD+PBS: phosphate buffered saline in drinking water, administered with antibiotic cocktail in water and an intra- peritoneal injection of clindamycin, and infected by C. difficile; (3) CD+BC1: Mice provided with BC (0.11%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (4) CD+BC2: Mice provided with BC (0.22%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (5) NC: Mice provided with no BC, no antibiotics and no C. difficile; (6) BC2 : Mice provided with 0.22% BC; (7) Ant: Mice administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin; (8) Ant+BC2 control: Mice fed with BC (0.22%) supplemented water and administered with antibiotic cocktail in water and an intra- peritoneal injection of clindamycin (A) Prophylactic study; (B) Therapeutic study *Significant difference between the untreated challenge (only CD group) with the rest of the treatment groups. #Significant difference between the untreated challenge (only CD group) with the rest of the treatment groups except CD+BC1.
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Fig 4A.
Fig 4B.
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Fig. 5 . Effect of baicalin supplementation on the abundance of major gut microbiome taxa in the antibiotic treated and C. difficile challenged mice Five to six-week-old C57BL/6 mice were randomly divided into eight treatment groups of 8 mice each. Mice were fed with irradiated feed and prophylactically supplemented with baicalin (0%, 0.11% and 0.22%) in drinking water); the challenge groups were made susceptible to C. difficile by administering an antibiotic cocktail in water and an intra-peritoneal injection of clindamycin. Further, challenge and control groups were infected with 106 CFU/ml of a hypervirulent C. difficile (ATCC 1803) spores or PBS and observed for clinical signs for ten days. The fecal samples were collected 2 DPI from which DNA was extracted for microbiome analysis using Illumina MiSeq platform, and the relative abundance of OTUs of major phyla, order, family, and genera was determined using microbiome analysis. Groups: (1) CD: administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (2) CD+PBS: phosphate buffered saline in drinking water, administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (3) CD+BC1: Mice provided with BC (0.11%), administered with antibiotic cocktail in water and an intra- peritoneal injection of clindamycin, and infected with C. difficile; (4) CD+BC2: Mice provided with BC (0.22%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (5) NC: Mice provided with no BC, no antibiotics and no C. difficile; (6) BC2 : Mice provided with 0.22% BC; (7) Ant: Mice administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin; (8) Ant+BC2 control: Mice fed with BC (0.22%) supplemented water and administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin
171
Fig. 6. Effect of baicalin supplementation on the abundance of Lactobacillaceae (5A), Lachnospriaceae (6A), Akkermansia (6A), Enterobacteriaceae (6B) and Peptostreptococcaceae (6B) in the antibiotic treated and C. difficile challenged mice Five to six-week-old C57BL/6 mice were randomly divided into eight treatment groups of 8 mice each. Mice were fed with irradiated feed and prophylactically supplemented with baicalin (0%, 0.11% and 0.22%) in drinking water); the challenge groups were made susceptible to C. difficile by administering an antibiotic cocktail in water and an intra-peritoneal injection of clindamycin. Further, challenge and control groups were infected with 106 CFU/ml of a hypervirulent C. difficile (ATCC 1803) spores or PBS and observed for clinical signs for ten days. The fecal samples were collected 2 DPI from which DNA was extracted for microbiome analysis using Illumina MiSeq platform, and the relative abundance of OTUs of major families (Lactobacillaceae (5a), Lachnospriaceae (5a), Akkermansia (5a), Enterobacteriaceae (5b) and Peptostreptococcaceae (5b) ) . Groups: (1) CD: administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (2) CD+PBS: phosphate buffered saline in drinking water, administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (3) CD+BC1: Mice provided with BC (0.11%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (4) CD+BC2: Mice provided with BC (0.22%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (5) NC: Mice provided with no BC, no antibiotics and no C. difficile; (6) BC2 : Mice provided with 0.22% BC; (7) Ant: Mice administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin; (8) Ant+BC2 control: Mice fed with BC (0.22%) supplemented water and administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin
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Fig 6A.
Fig 6B.
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Fig. 7 . Effect of baicalin supplementation on the diversity of gut microbiota of antibiotic treated and C. difficile challenged mice – Inverse Simpson plot Five to six-week-old C57BL/6 mice were randomly divided into eight treatment groups of 8 mice each. Mice were fed with irradiated feed and prophylactically supplemented with baicalin (0%, 0.11% and 0.22%) in drinking water); the challenge groups were made susceptible to C. difficile by administering an antibiotic cocktail in water and an intra-peritoneal injection of clindamycin. Further, challenge and control groups were infected with 106 CFU/ml of a hypervirulent C. difficile (ATCC 1803) spores or PBS and observed for clinical signs for ten days. The fecal samples were collected 2 DPI from which DNA was extracted for microbiome analysis using Illumina MiSeq platform, and Alpha diversity was calculated by using inverse Simpson to measure the richness and evenness of the OTUs. Groups: (1) CD: administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (2) CD+PBS: phosphate buffered saline in drinking water, administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (3) CD+BC1: Mice provided with BC (0.11%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (4) CD+BC2: Mice provided with BC (0.22%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (5) NC: Mice provided with no BC, no antibiotics and no C. difficile; (6) BC2 : Mice provided with 0.22% BC; (7) Ant: Mice administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin; (8) Ant+BC2 control: Mice fed with BC (0.22%) supplemented water and administered with antibiotic cocktail in water and an intra- peritoneal injection of clindamycin
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Fig. 8 . Effect of baicalin supplementation on the diversity of gut microbiota of antibiotic treated and C. difficile challenged mice – Bray-Curtis plot Five to six-week-old C57BL/6 mice were randomly divided into eight treatment groups of 8 mice each. Mice were fed with irradiated feed and prophylactically supplemented with baicalin (0%, 0.11% and 0.22%) in drinking water; the challenge groups were made susceptible to C. difficile by administering an antibiotic cocktail in water and an intra-peritoneal injection of clindamycin. Further, challenge and control groups were infected with 106 CFU/ml of a hypervirulent C. difficile (ATCC 1803) spores or PBS and observed for clinical signs for ten days. The fecal samples were collected 2 DPI from which DNA was extracted for microbiome analysis using Illumina MiSeq platform. Relationships between treatment groups based on the abundance of species present in each sample were plotted. NMS ordinations were run in R (v 3.3.0) using metaMDS in the vegan (v 2.3-5) package after calculating the stress scree plots to determine the number of axes required to achieve stress below 0.2, plotted using ggplot2 (v 2.1.0). Groups: (1) CD: administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (2) CD+PBS: phosphate buffered saline in drinking water, administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected by C. difficile; (3) CD+BC1: Mice provided with BC (0.11%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (4) CD+BC2: Mice provided with BC (0.22%), administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin, and infected with C. difficile; (5) NC: Mice provided with no BC, no antibiotics and no C. difficile; (6) BC2 : Mice provided with 0.22% BC; (7) Ant: Mice administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin; (8) Ant+BC2 control: Mice fed with BC (0.22%) supplemented water and administered with antibiotic cocktail in water and an intra-peritoneal injection of clindamycin
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CHAPTER VIII
Summary
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Clostridium difficile is a nosocomial bacterial pathogen causing a toxin-mediated enteric disease in humans. In the United States, the severity and incidence of C. difficile infection (CDI) have increased along with the emergence of hypervirulent strains causing both nosocomial and community associated outbreaks. With the global emergence of antibiotic resistant, hypervirulent strains of C. difficile, the use of antibiotics for controlling the infection in humans raises concerns.
Since C. difficile toxins and spores are the major virulence factors contributing to C. difficile pathogenesis and disease recurrence, identification of drug targets that can reduce toxin production, sporulation and spore germination in C. difficile could be a viable approach for reducing severity of the disease. Hence, this Ph.D. dissertation investigated the antivirulence activity of an essential mineral (sodium selenite), a plant derived flavonoid (baicalin), and selected lactic acid bacteria (LAB) against C. difficile, especially toxin production, spore formation and outgrowth. In addition, the efficacy of selenium in increasing C. difficile sensitivity towards ciprofloxacin and vancomycin was determined.
In vitro experiments using sub-minimum inhibitory concentrations of sodium selenite reduced toxin production and inhibited spore outgrowth in hypervirulent C. difficile. Analysis of gene expression studies revealed significant transcriptional downregulation of toxin associated genes. In addition, no inhibitory effect of sodium selenite on beneficial gut bacteria in vitro was observed. Overall, results suggest the potential use of sodium selenite to attenuate C. difficile virulence in the host. Although sodium selenite has a narrow margin of safety in mammals, validation of its antivirulence efficacy as an adjunct therapeutic candidate could be considered.
Furthermore, sodium selenite significantly increased C. difficile sensitivity to ciprofloxacin, but not vancomycin. Ciprofloxacin targets bacterial replication by compromising the DNA gyrase activity. Increased sensitivity of sodium selenite treated C. difficile to ciprofloxacin may be
177 attributed to its DNA toxicity and subsequent adverse effect on gyrase, thereby enhancing efficacy of the antibiotic on the bacterium. On the other hand, sodium selenite was not consistently effective in increasing C. difficile sensitivity to vancomycin. The antibacterial mechanism of vancomycin involves disruption of the bacterial pentapeptide crosslinking during peptidoglycan synthesis.
Although bacteria such as Staphylococcus aureus are reported to display resistance to vancomycin by enhancing the peptidoglycan synthesis process, a similar phenomenon has not been documented in C. difficile.
Antivirulence studies in C. difficile using baicalin, a plant derived flavone glycoside, was investigated based on previous findings by several researchers on its antimicrobial and antivirulence activities. Our current study revealed that the compound substantially reduced C. difficile toxin production, sporulation and spore outgrowth. In addition, baicalin negatively influenced the expression of genes required for pathogen colonization in host. The results from the in vitro studies suggested the potential use of baicalin to reduce all three components of C. difficile virulence such as toxin production, sporulation and spore germination. Therefore, as a logical extension, the prophylactic and therapeutic effect of baicalin against C. difficile was investigated in a murine model. Baicalin supplementation significantly reduced the incidence of diarrhea as well as the severity of CDI clinical symptoms. Microbiome analysis in antibiotic treated and C. difficile infected mice showed a drastic shift from a protective microbiota to a less beneficial microbiome profile that favors pathogen colonization. However, baicalin supplementation altered the microbiota by increasing the abundance of beneficial bacteria such as Lachnospiraceae and
Akkermansia. These results suggest that baicalin could potentially be used to control CDI without deleteriously affecting the microbiome. Further studies are need to understand host response to
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CDI during baicalin intervention and perform clinical trails in humans to validate its efficacy as an adjunctive therapeutic agent against CDI.
Lactic acid bacteria with putative probiotic activity exert multiple health benefits to the host, including improved nutrient digestion and assimilation, potentiating host immune function, and protection against enteric pathogens. In the third objective, five selected LAB isolates, namely
Lactobacillus plantarum 900B, Lactobacillus rhamnosus 400B, Lactobacillus plantarum 42-3,
Lactobacillus rhamnosus NRRL-B 442 and Lactobacillus paracasei DUP 13076 were screened for their antagonistic activity against C. difficile. Coculturing vegetative C. difficile with LAB significantly reduced toxin production when compared to C. difficile monocultures. In addition, the cell-free supernatants from coculture experiments produced negligible toxic effects on Vero cells. Moreover, when LAB isolates were cocultured with C. difficile spores, there was a significant reduction in spore outgrowth compared to controls. In summary, the tested LAB isolates significantly reduced two critical C. difficile virulence factors, namely toxin production and spore outgrowth. Follow up studies are needed to characterize the antimicrobial components elaborated by the LAB isolates and validate their efficacy in controlling CDI using in vivo models.
To conclude, the results of this Ph.D. research identified the potential of sodium selenite, baicalin and select lactic acid bacteria against CDI in humans, thereby justifying further investigations to develop them as an adjunct/alternative to antibiotics for controlling C. difficile.
Among the three anti-C. difficile approaches investigated in this dissertation, sodium selenite possesses comparatively the lowest margin of safety, with an upper tolerable limit of 400 µg and no observed-adverse-effect level of 800 µg (Anonymous, 2000; Sunde, 2012). Although the sub-
MIC of selenium found effective against C. difficile falls below the aforementioned limits, its efficacy and safety for controlling CDI in humans need to be validated using pharmacokinetic and
179 toxicological studies in appropriate animal models. In addition, a combinatorial strategy incorporating all three aforementioned approaches against CDI could also explored to assess if there exists any additive effect or synergism among them. Finally, the development of adaptive resistance (Fernández et al. 2012) in C. difficile towards selenium and baicalin could raise concerns in their therapeutic efficacy against CDI. Although there are no reports of development of adaptive resistance to non-conventional antimicrobials in C. difficile, further studies are needed to determine the plausible development of this phenomenon in the bacterium under continuous exposure to selenium and baicalin.
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