Investigation of Clostridia Isolated from Animal Feces for Potential Use As a Probiotic

INVESTIGATION OF CLOSTRIDIA ISOLATED FROM ANIMAL FECES FOR

POTENTIAL USE AS A PROBIOTIC

A Thesis

Presented to the

Faculty of

California State Polytechnic University, Pomona

In Partial Fulfillment

Of the Requirements for the Degree

Master of Science

Biological Sciences

Danielle K. Valencia

2020

SIGNATURE PAGE

THESIS: INVESTIGATION OF CLOSTRIDIA ISOLATED FROM ANIMAL FECES FOR POTENTIAL USE AS A PROBIOTIC

AUTHOR: Danielle K. Valencia

DATE SUBMITTED: Fall 2020

Department of Biological Sciences

Dr. Wei-Jen Lin Thesis Committee Chair Biological Sciences

Dr. Gregory Barding Chemistry and Biochemistry

Dr. Christos Stathopoulos Biological Sciences

ACKNOWLEDGEMENTS

I will always appreciate my time spent at Cal Poly Pomona from my first year as an undergraduate through my Master’s degree graduation and continued work in the lab. I am sincerely thankful for my advisor, Dr. Wei-Jen Lin, who was always patient in allowing me to work independently while still providing support at every turn. I would like to thank

Dr. Gregory Barding, as a member of my committee, along with his student Jocelyn for both taking the time to collaborate on my project. I would also like to thank Dr. Christos

Stathopoulos as a committee member and for his time and support.

Thank you to Cindy Reich of the Arabian Horse Center, Anthony Estep of the beef unit, and Brad Foyil and Alicia Seaman of the sheep, goat, and swine units for their collaboration in my sample collection. Moreover, I would like to give a special thanks to

Dr. Paul Beardsley for inspiring me to be forever kind, helpful, and hardworking ever since the beginning of my undergraduate career.

I am grateful to every friend that I met and for every invitation to conferences, dinners, parties, and the BRIC. Thank you to Liana Ab Samad, Jonathan Guo, Ann

Nasongkla, Justin Lee, Marina De Leon, Ashley Magin, Luis Torres, Huiying Hu, and the many others who worked in the lab; the TAs which I spent every week with; and anyone who took the time to ask me how I was doing. To Brent Wasilow, for unconditionally supporting me when I needed it the most. And to my family, I cannot express my gratitude for your love and support both personally and financially. You have all always been there to hold me up to reach my potential and to catch me whenever I fall.

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ABSTRACT

Clostridium species make up a large portion of the intestinal microbiota in humans and animals with strong implications that they are necessary for overall gut function.

Dysbiosis due to decreased abundances and diversity of Clostridium often cause major health consequences due to the resulting lack of intestinal short chain fatty acids such as butyrate provided by these organisms. Safe and effective results have been reported upon oral administration of Clostridium probiotic supplements for symptom reduction and disease prevention indicating their prospective use as a therapeutic agent. We hypothesized that intestinal butyrate producing Clostridium can be isolated from the feces of animals and that some of these isolates will have the potential to be used safely as a probiotic dietary supplement.

Enrichment and isolation of Clostridium from horse, cattle, sheep, pig, and goat fecal samples were identified and characterized by their colony morphology on egg yolk agar, microscopic morphology by staining, biochemical reactions, and 16S rRNA sequencing results. A total of 6 species from 134 Clostridia isolates were identified and the butyrate kinase gene was confirmed in all of these isolates implicating the ability to produce butyrate. Based on hemolysis and antibiotic susceptibility tests outlined by the

European Food and Safety Authority, 6 isolates were considered to be potentially safe as a probiotic and were further biochemically characterized by API microbial identification kits. Of these probiotic candidates, Paeniclostridium sordellii isolates P1F and P2E were determined to be unsafe for use as a probiotic due to the high risk of virulence factors and horizontal gene transfer of toxins. Clostridium senegalense isolate C4H was potentially the safest for use as a probiotic followed by Clostridium tepidum isolates C4B, P2D, and P4D.

TABLE OF CONTENTS

SIGNATURE PAGE ...... ii

ACKNOWLEDGEMENTS ...... iii

ABSTRACT ...... iv

LIST OF TABLES ...... x

LIST OF FIGURES ...... xii

LIST OF ABBREVIATIONS ...... xiv

1. INTRODUCTION ...... 1

1.1. Probiotics ...... 1

1.1.1. Characteristics of probiotics ...... 1

1.1.2. Beneficial probiotic effects:...... 2

1.1.3. Safety and efficacy of probiotics: ...... 3

1.2. Clostridium ...... 4

1.2.1. Characteristics of Clostridium ...... 4

1.2.2. Clostridium in the human microbiome ...... 5

1.2.3. Probiotic Clostridium ...... 7

1.2.4. Therapeutic applications of Clostridium ...... 8

1.3. Beneficial effects of butyrate ...... 11

1.3.1. Intestinal butyrate ...... 11

1.3.2. Modulation of the inflammatory and immune response ...... 12

1.3.3. Improvement of intestinal barrier function ...... 13

1.3.4. Preferential inhibition of cancerous colonocytes ...... 14

1.3.5. Modulation of ion transport ...... 15

1.3.6. Improvement of oxidative status ...... 16

1.4. Animal digestive systems and butyrate production ...... 16

1.4.1. Monogastric digestive system ...... 16

1.4.2. Microbiome and butyrate production in monogastric animals ...... 17

1.4.3. Polygastric digestive system ...... 19

1.4.4. Microbiome and butyrate production in polygastric animals ...... 20

2. OBJECTIVES ...... 22

3. MATERIALS AND METHODS ...... 23

3.1. Animal demographics ...... 23

3.2. Labeling of bacterial isolates...... 25

3.3. Fecal sample collection ...... 26

3.4. Bacterial culture media and reagents ...... 26

3.4.1. Anaerobic conditions ...... 26

3.4.2. Thioglycolate broth...... 27

3.4.3. Cooked meat media (CMM) ...... 27

3.4.4. Egg yolk agar (EYA) ...... 27

3.4.5. Blood agar plates (BAP) ...... 28

3.4.6. Trypticase peptone glucose yeast extract (TPGY) ...... 28

3.4.7 Phosphate buffered saline (PBS) ...... 28

3.4.8. Brucella broth ...... 29

3.4.9. Vitamin K1 (1 mg/ml) ...... 29

3.4.10. Hemin (5 mg/ml) ...... 29

3.4.11. Laked sheep blood ...... 30

3.4.12. Antibiotic solutions...... 30

3.4.13. Modified Brucella blood agar ...... 30

3.4.14. Nitric acid (3 N) ...... 31

3.5. Enrichment of Clostridium species ...... 31

3.6. Isolation of Clostridium species ...... 32

3.7. Identification and characterization of Clostridium isolates...... 32

3.8. DNA purification...... 33

3.8.1. DNA purification from fecal samples ...... 33

3.8.2. DNA purification from presumed Clostridium isolates ...... 34

3.9. Polymerase Chain Reaction (PCR) ...... 34

3.9.1 PCR primers ...... 34

3.9.2. PCR assay ...... 35

3.9.3. Gel electrophoresis ...... 36

3.10. Crude sample gene analysis ...... 37

3.11. Clostridium species determination ...... 37

3.11.1. Sequencing...... 37

3.11.2. Species determination ...... 37

3.11.3. Limiting Clostridium sporogenes isolates ...... 38

3.12. Antibiotic susceptibility ...... 38

3.13. Biochemical testing ...... 39

3.14. Butyrate production ...... 40

3.14.1. Nitric acid wash ...... 40

3.14.2. Acidogenesis end point ...... 40

vii

4. RESULTS ...... 41

4.1. Presence of the BUK gene in fecal samples ...... 41

4.2. Colony morphology of fecal samples enriched for Clostridium species ...... 43

4.3. Staining, biochemical, and molecular characterization of bacterial isolates ...... 49

4.4. Presence of the BUK gene in presumptive Clostridium isolates ...... 55

4.5. Identification of presumptive Clostridium isolates by 16S RNA sequence

analysis ...... 59

4.6. Phylogenetic relationships of sequenced isolates and related Clostridia

species ...... 61

4.7. Antibiotic susceptibility of sequenced isolates ...... 63

4.8. Biochemical characterization by API of potential probiotic isolates ...... 69

4.9. Endpoint determination of acidogenesis in Clostridium sporogenes ...... 74

5. DISCUSSION ...... 78

5.1. Overview ...... 78

5.2. Comparison of characteristics in isolated Clostridia species ...... 79

5.3. Antibiotic resistance mechanisms in potential probiotic Clostridia ...... 82

5.4. Relation of isolates, commensal gut Clostridium, and probiotic Clostridium ...... 84

6. FUTURE STUDIES...... 86

6.1. Further characterization of newly published species ...... 86

6.2. Determining toxicity in potential probiotic isolates ...... 86

6.3. Determining the cause of antibiotic resistance in potential probiotic isolates ...... 87

6.4. Determining and optimizing the protocol for butyrate production analysis ...... 87

REFERENCES ...... 88

viii

APPENDIX A ...... 108

APPENDIX B ...... 112

APPENDIX C ...... 116

LIST OF TABLES

Table 3.1. Details of horses from which fecal samples were collected...... 23

Table 3.2. Details of cattle from which fecal samples were collected...... 24

Table 3.3. Details of sheep from which fecal samples were collected...... 24

Table 3.4. Details of pigs from which fecal samples were collected...... 25

Table 3.5. Details of goats from which fecal samples were collected...... 25

Table 3.6. Isolate labeling system...... 26

Table 3.7. Variable nucleotide symbols...... 35

Table 3.8. Clostridium species used in BUK gene multiple sequence alignment...... 35

Table 3.9. Primer sequences for PCR amplification...... 35

Table 3.10. PCR reaction settings for the 16S rRNA and BUK genes ...... 36

Table 3.11. Selected bacterial cut-off values (mg/L) for antimicrobials of human and

veterinary importance as required by the EFSA ...... 39

Table 4.1. Statistical significance for the relative amounts of buk in crude fecal

samples using the Mann-Whitney test ...... 43

Table 4.2. Total number of bacterial isolates and presumptive Clostridium isolates

from horse, cattle, sheep, pig, and goat samples...... 50

Table 4.3. Staining, biochemical, and molecular characterization of isolates from horse

samples ...... 51

Table 4.4. Staining, biochemical, and molecular characterization of isolates from cattle

samples ...... 52

Table 4.5. Staining, biochemical, and molecular characterization of isolates from sheep

samples ...... 53

Table 4.6. Staining, biochemical, and molecular characterization of isolates from pig

samples ...... 54

Table 4.7. Staining, biochemical, and molecular characterization of isolates from goat

samples ...... 55

Table 4.8. Species determination of presumptive Clostridium isolates based on 16S

rRNA sequencing ...... 61

Table 4.9. Clostridium sporogenes isolates limited to one representative isolate for

antibiotic susceptibility testing ...... 64

Table 4.10. Antibiotic susceptibility of sequenced isolates ...... 65

Table 4.11. Biochemical characterization of potential probiotic Clostridia isolates by

the API 20 A microbial identification kit ...... 71

Table 4.12. Biochemical characterization of potential probiotic Clostridia isolates by

the API 50 CH microbial identification kit ...... 71

Table 1. Endpoint determination of acidogenesis in Clostridium sporogenes by OD.... 116

LIST OF FIGURES

Figure 1.1. Pathways of butyrate biosynthesis from butyryl-CoA in the large intestine .. 12

Figure 1.2. An example of the monogastric digestive system ...... 17

Figure 1.3. An example of the polygastric digestive system ...... 20

Figure 4.1. Agarose gel electrophoresis of the 16S rRNA gene in crude fecal samples .. 41

Figure 4.2. Agarose gel electrophoresis of the BUK gene in crude fecal samples ...... 42

Figure 4.3. Relative amount of buk in crude fecal samples ...... 43

Figure 4.4. Growth from heat shocked and enriched horse fecal samples ...... 44

Figure 4.5. Growth from heat shocked and enriched cattle fecal samples ...... 45

Figure 4.6. Growth from heat shocked and enriched sheep fecal samples ...... 46

Figure 4.7. Growth from heat shocked and enriched pig fecal samples ...... 48

Figure 4.8. Growth from heat shocked and enriched goat fecal samples ...... 49

Figure 4.9. Agarose gel electrophoresis of the BUK gene in presumptive Clostridium

isolates from horse samples...... 56

Figure 4.10. Agarose gel electrophoresis of the BUK gene in presumptive Clostridium

isolates from cattle samples ...... 57

Figure 4.11. Agarose gel electrophoresis of the BUK gene in presumptive Clostridium

isolates from sheep samples ...... 57

Figure 4.12. Agarose gel electrophoresis of the BUK gene in presumptive Clostridium

isolates from pig samples ...... 58

Figure 4.13. Agarose gel electrophoresis of the BUK gene in presumptive Clostridium

isolates from goat samples ...... 59

xii

Figure 4.14. Phylogenetic tree based on 16S rRNA gene sequences of Clostridia

isolates and related Clostridia species...... 63

Figure 4.15. Representative modified Brucella blood agar plates for antibiotic

susceptibility ...... 67

Figure 4.16. Percentage of sequenced isolates by animal susceptible to antibiotics ...... 68

Figure 4.17. Percentage of sequenced isolates by species susceptible to antibiotics ...... 69

Figure 4.18. Analytical profile index ...... 73

Figure 4.19. Endpoint determination of acidogenesis in Clostridium sporogenes ...... 76

Figure 5.1. Isolate screening procedure ...... 79

Figure 1. Multiple sequence alignment using T-Coffee ...... 108

Figure 2. Agarose gel electrophoresis of the 16S rRNA gene in presumptive Clostridium

isolates from horse samples ...... 112

Figure 3. Agarose gel electrophoresis of the 16S rRNA gene in presumptive Clostridium

isolates from cattle samples ...... 112

Figure 4. Agarose gel electrophoresis of the 16S rRNA gene in presumptive Clostridium

isolates from sheep samples ...... 113

Figure 5. Agarose gel electrophoresis of the 16S rRNA gene in presumptive Clostridium

isolates from pig samples ...... 114

Figure 6. Agarose gel electrophoresis of the 16S rRNA gene in presumptive Clostridium

isolates from goat samples ...... 114

xiii

LIST OF ABBREVIATIONS

ABE Acetone-butanol-ethanol

ATCC American Type Culture Collection

BAP Blood agar plate

BF Bacteroides fragilis

BLAST Basic local alignment search tool

BoNT Botulinum neurotoxin

BUK Butyrate kinase

CLSI Clinical and Laboratory Standards Institute

CMM Cooked meat media

CP Clostridium perfringens

CPP California State Polytechnic University, Pomona

EFSA European Food and Safety Authority

EYA Egg yolk agar

FAO Food and Agriculture Organization of the United Nations

FDA Food and Drug Administration

GRAS Generally Recognized as Safe

IBD Inflammatory bowel disease

ISAPP International Scientific Association for Probiotics and Prebiotics

NCBI National Center for Biotechnology Information

NMR Nuclear magnetic resonance

OD Optical density

xiv

PBS Phosphate buffered saline

PCR Polymerase chain reaction

SCFA Short-chain fatty acid

TPGY Trypticase peptone glucose yeast

WHO World Health Organization

1 INTRODUCTION

1.1. Probiotics

1.1.1. Characteristics of probiotics: The Food and Agriculture Organization of the United Nations (FAO) and World Health Organization (WHO) defined probiotics in

2002 as “live strains of strictly selected microorganisms which, when administered in adequate amounts, confer a health benefit to the host” which was further maintained in

2013 by the International Scientific Association for Probiotics and Prebiotics (ISAPP)

(Markowiak and Slizewaka 2017). While this definition is internationally accepted, there is no legal definition or requirements to meet these fundamentals; although, the FAO,

WHO, and European Food and Safety Authority (EFSA) suggest that probiotic organisms meet certain criteria based on safety, functionality, and technological usefulness

(Markowiak and Slizewaka 2017, Sanders 2008). These suggested attributes include a lack of antibiotic resistance, nonpathogenic, ability to survive the digestive tract and proliferate in the gut, antagonistic towards pathogens, genetic stability, easy production, high viability and stability during storage, and documented health benefits (Markowiak and Slizewaka

2017, Vandenplas et al 2015). Probiotic organisms are also suggested to originate from the gastrointestinal tract of healthy humans or animals and are commonly from the genera

Lactobacillus and Bifidobacterium as well as Saccharomyces, Lactococcus, Streptococcus,

Enterococcus, Bacillus, and Escherichia (Azad et al 2018, Fijan 2014, Sanders 2008).

However, probiotic organisms of the same genus and species act through different mechanisms indicating strain specific safety profiles and clinical effects with differing strains needed to treat multiple health conditions (Doron and Snydman 2015, Gogineni et al 2013).

1.1.2. Beneficial probiotic effects: Probiotics have a high therapeutic potential providing many beneficial effects on the host’s health with both a preventative and curative nature (Markowiak and Slizewaka 2017, Vandenplas et al 2015). They are implicated in improving the host’s overall health status and decreasing susceptibility to pathogens as well as treating or preventing various ailments including colorectal cancer, diarrheal diseases, constipation, irritable bowel syndrome, inflammatory bowel disease, necrotizing enterocolitis, Clostridium difficile infections, Helicobacter pylori infections, allergic responses, lactose intolerance, and atopic dermatitis (Bermudez-Brito et al 2012, Fijan

2014, Gogineni et al 2013, Islam 2015, Markowiak and Slizewaka 2017). New biological effects by probiotic organisms are frequently studied although the mechanisms by which these effects are exerted are largely unknown and likely to be multifaceted (Bermudez-

Brito et al 2012, Vandenplas et al 2015). The strain-specific nature of probiotics also indicates that it is unlikely that an individual strain will exhibit all probiotic mechanisms to constitute a universal remedy and that a combination of strains will provide increased beneficial effects. (Markowiak and Slizewaka 2017).

Probiotics are reported to maintain intestinal microbial homeostasis, stimulate and modulate various biological functions, and contribute to host metabolic processes (Azad et al 2018, Gogineni et al 2013, Vandenplas et al 2015). Microbial homeostasis is maintained through the colonization of probiotic organisms on the mucus layer of the intestinal epithelium to form a protective barrier which blocks out pathogens through competitive exclusion (Markowiak and Slizewaka 2017, Wan et al 2016). Pathogen activity is further reduced by direct antagonism through interference with quorum sensing, induction of host cell defensins and cathelicidins, and production of organic acids, de-conjugated bile salts,

and bacteriocins. All of which create a hostile environment for invasive pathogenic organisms (Bermudez-Brito et al 2012, Markowiak and Slizewaka 2017, Gogineni et al

2013, Vandenplas et al 2015). Adhesion is also important for interactions between probiotic organisms and the host as it allows for probiotics to influence intestinal defense responses by stabilizing intestinal barrier function and modulating the innate and adaptive immune responses (Bermudez-Brito et al 2012, Wan et al 2016). Probiotics stabilize epithelial barrier function by regulating gene expression to decrease intestinal epithelial cell apoptosis, enhance tight junction proteins, and increase mucin production (Bermudez-

Brito et al 2012, Gogineni et al 2013). The effect on gene expression is extended to the immune system where probiotics maintain immunological tolerance to environmental antigens, modulate inflammatory responses, and inhibit autoimmunity (Markowiak and

Slizewaka 2017, Wan et al 2016). Furthermore, probiotic organisms positively affect host metabolism and nutrition by direct production or stimulating host pathways to produce certain vitamins, enzymes, and co-enzymes as well as to reduce toxic compounds

(Markowiak and Slizewaka 2017, Vandenplas et al 2015).

1.1.3. Safety and efficacy of probiotics: In the United States, the Food and Drug

Administration (FDA) classifies probiotics as dietary supplements which does not require premarket approval for claims of safety and efficacy by manufacturers (Gogineni et al

2013, Sanders 2008). As a result, most probiotics are often used for a wide range of scenarios where their efficacy is not well established by scientific evidence and clinical data (Fijan 2014, Islam 2015). General health claims are allowed for probiotic products with the FDA challenging their labelling and safety only in instances where the product is marketed as a drug while lacking drug approval and sufficient support from scientific

evidence (Sanders 2008, Vandenplas et al 2015). Moreover, the FDA has given probiotics the designation Generally Recognized as Safe (GRAS) which assesses safety but does not evaluate claims of efficacy (Gogineni et al 2013). GRAS status was obtained by probiotics due to their long history of safe use and current data from clinical trials, animal studies, and in vitro studies (Doron and Snydman 2015). While probiotics have shown to be safe in healthy individuals, some populations are at a high risk for adverse effects including the immunocompromised, critically ill, and pregnant as well as premature infants (Doron and

Snydman 2015, Fijan 2014, Islam 2015). Rare cases of bacteremia, fungemia, sepsis, gastrointestinal ischemia, endocarditis, and liver abscesses have been reported; although, these cases have only been seen in vulnerable individuals with severe comorbidities and no reports in otherwise healthy individuals (Fijan 2014, Gogineni et al 2013, Islam 2016).

Translocation of probiotic organisms into systemic circulation from the gastrointestinal tract and transfer of antibiotic resistance genes between organisms in the gut have not been reported despite their theoretical possibility (Doron and Snydman 2015, Vandenplas et al

2015). Nevertheless, research into the safety and efficacy of probiotics in the prophylaxis and treatment of medical conditions is moving forward through laboratory and clinical studies (Gogineni et al 2013).

1.2. Clostridium

1.2.1. Characteristics of Clostridium: Commonly found in the natural environment including soil, water, bottom sediments, and in human and animal digestive systems, bacteria in the genus Clostridium have traditionally been defined by similar morphological and phenotypical characteristics (Rainey et al 2015, Samul et al 2013).

Cells are rod shaped, stain Gram positive in young cultures, may be motile by peritrichous

flagella, and form oval or spherical endospores that typically distend the cell. They are obligately anaerobic with varying oxygen tolerances, oxidase negative as they lack a cytochrome system, and catalase negative. Species vary metabolically; however, they are usually chemoorganotrophic, produce acids and alcohols from carbohydrates or peptones, do not reduce sulfate to sulfide, and can be saccharolytic, proteolytic, neither or both

(Rainey et al 2015). Low G and C contents are typical in this genus and range from 22 to

55 mol% with toxigenic species at 24 to 29 mol% (Stevens et al 2015). Clostridium is most well known by its pathogenic organisms, but most play important roles in industry and medicine through their ability to ferment organic compounds to produces acids and solvents (Samul et al 2013). Phylogenetically, Clostridium is a diverse, polyphyletic group composed of 168 species divided into 19 16S rRNA gene clusters. Cluster I forms the basis of the genus representing Clostridium sensu stricto and contains 77 species including the type strain. Species in the remaining clusters require reclassification into novel or other existing genera (Rainey et al 2015).

1.2.2. Clostridium in the human microbiome: Commensal Clostridium are reported to start colonizing the intestines during the first month of a neonate’s life and can be detected within the first postnatal week (Guo et al 2020, Leputso et al 2013). Optimal conditions and a more stable environment arise as oxygen is consumed by aerobic bacteria during the first few days following birth leading to colonization by facultative and strictly anaerobic bacteria including Clostridium (Cassir et al 2016, Leputso et al 2013). A distinctly higher proportion of Clostridium cluster I is found in infants including mostly C. butyricum and C. paraputrificum whereas higher proportions of Clostridium clusters IV and XIVa stably colonize the gut of adults (Guo et al 2020). In the elderly, a decrease in

gut diversity and stability is seen and indicated by decreasing numbers of strict anaerobes and increasing numbers of facultative anaerobes. However, significantly higher counts of

Clostridium cluster I are found while counts of Bacteroides and Bifidobacteria are lower

(Biagi et al 2012, Leputso et al 2013). This consistent colonization of Clostridium species in the gut suggest that they play an indispensable role in maintaining intestinal homeostasis throughout life (Leputso et al 2013).

The adult human gut microbiota is dominated by bacteria with about 1013-1014 cells which are predominately strict anaerobes (Kho and Lal 2018, Rinninella et al 2019). While there is broad diversity at the species level, only 5 phyla are represented in the gut including

Firmicutes, Bacteroidetes, Actinobacteria, Proteobacteria, and Verrucomicrobia (Luu et al 2017). Furthermore, 90% of the community is restricted to the phyla Firmicutes (50-

80%) and Bacteroidetes (10-30%) with the genus Clostridium representing about 95% of

Firmicutes in the gut (Biagi et al 2012, Luu et al 2017, Rinninella et al 2019). Species within Clostridium clusters IV (Clostridium leptum group) and XIVa (Clostridium coccoides group) are the most numerically present and are the richest bacterial groups in the intestine containing 4 members and 21 members, respectively (Guo et al 2020, Nagano et al 2012). Making up 10-40% of the total gut bacteria, Clostridium clusters IV and XIVa are commensal organisms strongly implicated in playing crucial roles in overall gut maintenance and function (Leputso et al 2013).

Clostridium species are necessary regulators of intestinal homeostasis as a result of their close relationship with the intestinal epithelial cells and production of essential digestive enzymes and metabolites (Guo et al 2020, Nagano et al 2012). Commensal

Clostridium are not randomly distributed in the gut and primarily colonize the mucin layer

between the mucosal folds of the ascending colon (Ayua et al 2020, Nava et al 2011). It has been hypothesized that the close proximity of Clostridium to the epithelium allows for their strong influence on colonocyte energy metabolism as well as the host’s immune system (Leputso et al 2013, Nagano et al 2012). Clostridium species are responsible for fermenting large amounts of undigestible dietary fiber to produce short-chain fatty acids with butyrate exerting the greatest effect (Ayua et al 2020, Nava et al 2011). Butyrate functions as an essential fuel source for colonocytes while also influencing gene expression as a non-competitive inhibitor of histone deacetylases (Leputso et al 2013). Consequently, commensally produced butyrate has shown to be a potent inducer of colonic Treg cell de novo generation, differentiation, and functional maturation; moreover, these Treg cells have been reported to migrate to distant tissues to reduce inflammation and allergic responses throughout the body (Furusawa et al 2015, Kunisawa and Kiyono 2011, Luu et al 2017).

While Clostridium prefers fermentable carbohydrates, some species are also capable of fermenting amino acids into indolepropionic acid which has been reported to improve intestinal permeability (Guo et al 2020). Additionally, Clostridium play a role in converting primary bile acids into secondary bile acids through the production of bile salt hydrolases where this process also functions to improve resistance to C. difficile infections (Buffie et al 2015, Kho and Lal 2018).

1.2.3. Probiotic Clostridium: Both toxigenic and non-toxigenic species and strains of Clostridium form part of the normal gut microbiota (Cassir et al 2016). Careful consideration should be used when determining potential probiotic Clostridium due to the possibility of vertically and horizontally transferable toxin genes, virulence factors, and antibiotic resistance genes (Cassir et al 2016, Guo et al 2020). Currently, Clostridium

butyricum is the only species of Clostridium to be approved for the use as a probiotic and is commonly used throughout Asia (Cassir et al 2016). Commercialized strains of C. butyricum include MIYAIRI 588 from Miyarisan Pharmaceuticals and TO-A from TOA

Pharmaceuticals and Advanced Orthomolecular Research; furthermore, the C. butyricum

TO-A probiotics are formulated as a symbiotic supplement that also includes the lactic acid bacteria Enterococcus faecalis T-110 and the amylolytic bacteria Bacillus mesentericus

TO-A or Bacillus subtilis TO-A (BIO-THREE, Clostridium butyricum MIYAIRI strain,

Probiotic 3 2019). C. butyricum MIYAIRI 588 and TO-A have been used as probiotic in

Japan since the 1960s and have shown to be nontoxic, nonmutagenic, and genetically stable with no known toxins or virulence factors (BIO-THREE, Clostridium butyricum MIYAIRI strain, Pen and Tec Consulting 2012). C. butyricum is also beneficial as a probiotic because it is already known to be a nonpathogenic gut commensal, it readily forms shelf stable spores which are resistant to digestion and germinate in the gut, and it produces high amounts of butyrate which has known beneficial effects on cellular metabolism and intestinal homeostasis (Cassir et al 2016, Pen and Tec Consulting 2012, Wang et al 2018).

1.2.4. Therapeutic applications of Clostridium: Inflammatory bowel disease

(IBD) is a class of chronic, relapsing inflammatory disorders that affects the gastrointestinal tract (Fava and Danese 2011, Kumari et al 2013). With no known cause,

IBD is believed to be triggered by immunological abnormalities due to a combination of genetic and environmental factors (Kaakoush et al 2012, Takaishi et al 2008). Animal models have suggested that the indigenous intestinal microbiota is the most important environmental factor as IBD sensitive knockout mice do not develop colitis when raised under germ free conditions (Takaishi et al 2008, Yoshimatsu et al 2015). Furthermore, an

abnormal composition of the gut microbiota is characteristic of IBD as demonstrated by a reduced abundance of the phyla Firmicutes and Bacteroides along with an increase of

Proteobacteria and Actinobacteria (Fava and Danese 2011). The decrease in Firmicutes is attributed to the significantly decreased abundance and diversity of Clostridium clusters IV and XIVa (Fava and Danese 2011, Kabeerdoss et al 2013). Decreases in these dominant butyrate producers directly causes decreased concentrations of butyrate in the gut thus contributing to greater inflammation and reduced proliferation of colonic epithelial cells

(Kabeerdoss et al 2013, Kumari et al 2013, Takaishi et al 2008). Sodium butyrate is considered to have therapeutic value in the treatment of IBD, but can cause noninflammatory colonic hypersensitivity; moreover, other conventional treatments provide limited results and often have serious side effects (Zhang et al 2009). Recently, novel probiotic therapeutics have been reported to have promising results as a safe and effective treatment which improves clinical symptoms and gut dysbiosis as well as inducing and maintaining remission (Fava and Danese 2011, Yoshimatsu et al 2015).

In experimental colitis induced animal models, Clostridium butyricum reduces symptoms and histological damage or prevents the development of colitis by improving intestinal barrier function, modulating the immune system, and reducing oxidative stress

(Hayashi et al 2013, Liu et al 2020, Zhang et al 2009). Similar results were found in colitis induced germ-free mice colonized with 46 strains of Clostridium primarily from clusters

IV and XIVa or with 17 strains of Clostridium from clusters IV, XIVa, and XVII (Atarashi et al 2011, Atarashi et al 2013). In both mice and humans, Fusicatenibacter saccharivorans from Clostridium cluster XIVa was able to suppress a type of IBD known as ulcerative colitis (Takeshita et al 2016). Treatment with C. butyricum MIRAIRI prevented the

development of pouchitis in ulcerative colitis patients who had undergone a total proctocolectomy with ileal pouch anastomosis with minimal side effects by decreasing the amounts of putrefying bacteria and increasing beneficial bacteria (Yasueda et al 2016). A combination of C. butyricum and specific immunotherapy ameliorated the clinical symptoms of food allergy, a major causative factor in IBD, in patients with ulcerative colitis through modulation of the immune system more effectively than either treatment alone (Cai et al 2016, Lan et al 2016). Patients with IBD are more susceptible to colorectal cancer and treatment with C. butyricum indicated a potential preventative effect on carcinogenesis in animal models by improving IBD symptoms and increasing apoptosis of tumor cells which reduced tumor size, number, and invasiveness (Liu et al 2020).

C. butyricum has been implicated in other gastrointestinal conditions with reports to improve quality of life and stool frequency in diarrhea dominant irritable bowel syndrome, prevent gut dysbiosis and intestinal damage from antibiotic associated diarrhea, reduce chemotherapy induced diarrhea, protect against gastric ulcers, and act as a preventative and therapeutic for opportunistic pathogens such as Clostridium difficile,

Salmonella enteritidis, Escherichia coli O157:H7, and Helicobacter pylori (Chen et al

2019, Hagihara et al 2020, Oka et al 2018, Sun et al 2018, et al 2004, Takahashi, Tian et al 2018, Wang et al 2015, Zhao et al 2020). Studies on the effects outside of the gastrointestinal tract are limited and mostly restricted to animal models, but results have shown that C. butyricum may protect against type 1 and type 2 diabetes, ameliorate metabolic and inflammatory symptoms associated with obesity, regulate the gut-brain axis to exert neuroprotective effects against Alzheimer’s disease and vascular dementia, and

treat bacterial vaginosis (Jia et al 2017, Liu et al 2015, Shang et al 2016, Sun et al 2020,

Zhou et al 2019).

1.3. Beneficial effects of butyrate

1.3.1. Intestinal butyrate: Short-chain fatty acids (SCFA) are produced within the intestinal lumen by microbial fermentation of undigested dietary carbohydrates and, to a lesser degree, dietary and endogenous proteins (Canani et al 2011, Hamer et al 2007, Wang et al 2018). The production of SCFAs allows for energy to be produced from dietary fiber that has not been digested in the small intestine and is estimated to contribute about 60-

70% of the energy required for colonic epithelial cells as well as about 5-15% of the total caloric requirements of humans (Hamer et al 2011, Wang et al 2018). Although, the total and relative molar concentrations of SCFAs in the colonic lumen can vary depending on the site of fermentation, diet, and intestinal microbiota composition, they are generally produced at a ratio of 60% acetate, 25% propionate, and 15% butyrate (Canani et al 2011,

Hamer et al 2007, Wang et al 2018). Intestinal butyrate is produced by a functional group of Gram-positive anaerobic bacteria widely distributed across several Clostridium clusters with clusters IV and XIVa making up 10-40% of the total adult human gut microbiota

(Lopetuso et al 2013, Wang et al 2018). This community produces butyrate through fermentation by 2 metabolic pathways: the phosphorylation of butyryl-CoA to butyryl- phosphate which is converted to butyrate by butyrate kinase or the transfer of CoA from butyryl-CoA to acetate by butyryl-CoA:acetate-CoA transferase to form butyrate and acetyl-CoA (Figure 1.1). Uptake of butyrate across the apical membrane of colonocytes occurs by different mechanisms including diffusion of the undissociated form,

SCFA/HNO3 ̄ exchange, and active transport of the dissociated form (Louis and Flint

2009, Wang et al 2018). After absorption, colonocytes can then utilize it as a major source of energy while cytoplasmic butyrate also maintains many beneficial aspects of colonic health (Canani et al 2011).

Figure 1.1. Pathways of butyrate biosynthesis from butyryl-CoA in the large intestine. (A) The phosphorylation of butyryl-CoA to butyryl-phosphate which is converted to butyrate via butyrate kinase. (B) The transfer of CoA from butyryl-CoA to external acetate via butyryl-CoA:acetate-CoA transferase (Wang et al 2018).

1.3.2. Modulation of the inflammatory and immune response: Inflammatory responses are normally well-regulated by feedback mechanisms and signaling cascades, but excessive inflammatory responses can cause irreparable damage to the host and possibly lead to disease (Knudsen et al 2018). Chronic inflammation can occur locally causing intestinal inflammatory diseases such as inflammatory bowel disease, ulcerative colitis, and Crohn’s disease or it can occur systemically which is associated with an increased risk of inflammatory autoimmune disorders such as multiple sclerosis and rheumatoid arthritis (Chen et al 2018, Knudsen et al 2018, Saemann et al 2000). Butyrate modulates the immune system by acting as a histone deacetylase (HDAC) inhibitor on the inflammation regulating transcription factor Foxp3 gene locus to increase the expression and development of T cells (Furusawa et al 2013, Liu et al 2016). As a result, Foxp3+ CD4+

Treg cells are preferentially induced and significantly increase the production of the potent anti-inflammatory cytokine IL-10 while also significantly suppressing IL-12 which decreases the production of the pro-inflammatory cytokines IFN-γ and TNF-α (Furusawa et al 2013, Hui et al 2019, Saemann et al 2000). SCFAs, including butyrate, activate the G protein-coupled receptors GPR41 and GPR43 leading to a decrease in the production of pro-inflammatory cytokines and chemokines through the MAPK signaling pathway

(Chang et al 2019, Pirozzi et al 2018). Only butyrate activates GPR109a to downregulate the activation of the pro-inflammatory mediating transcription factor NF-κB by inhibiting the phosphorylation of the NF-κB p65 signaling pathway (Chen et al 2018, Singh et al

2014). NF-κB is further inhibited by the resulting decreases in the pro-inflammatory cytokines as TNF-α and IL-1β stimulate the nuclear translocation of NF-κB (Luhrs et al

2001, Segain et al 2000).

1.3.3. Improvement of intestinal barrier function: The intestinal barrier is composed of epithelial cells, a mucus layer, and adhered bacteria which allow for the absorption of nutrients while also providing the first line of defense against pathogens and toxins (Jung et al 2015, Yan and Ajuwon 2017). Moreover, intestinal permeability caused by a defective intestinal barrier is associated with gastrointestinal diseases including infections from pathogenic bacteria, diarrheal diseases, inflammatory bowel disease, and ulcerative colitis (Cobo et al 2017, Yan and Ajuwon 2017). Butyrate has been reported to improve the intestinal barrier by regulating the expression and assembly of tight junction proteins which anchor adjacent intestinal epithelial cells together (Feng et al 2018, Peng et al 2009, Yan and Ajuwon 2017). The expression of the main tight junction proteins claudin,

13nglish13d, and ZO-1 are increased in association with butyrate’s effect on the

Akt/mTOR mediated protein synthesis pathway (Feng et al 2018, Yan and Ajuwon 2017).

While the underlying mechanism of tight junction protein assembly is unclear, the activation of AMPK is necessary and is facilitated by butyrate as it increases AMPK activity (Feng et al 2018, Peng et al 2009, Yan and Ajuwon 2017). In addition to increasing tight junction protein expression, butyrate also increases the gene expression of the membrane associated mucin genes MUC1, MUC3, MUC4, and MUC12 as well as the secretory mucin gene MUC2 (Jung et al 2015, Feng et al 2018). Mucin acts as a defense for the host as MUC1 and MUC4 support the adhesion of commensal bacteria, MUC3 inhibits the adherence of pathogenic bacteria, and MUC2 stimulates the expression and secretion of the broadly antimicrobial peptides known as cathelicidins (Cobo et al 2017,

Jung et al 2015).

1.3.4. Preferential inhibition of cancerous colonocytes: For many years, a high intake of dietary fiber and resistant starches has been reported to reduce the risk of colon cancer (Zeng et al 2017, Zhang et al 2016). It is believed that this effect is due to the known anti-tumorigenic agent butyrate which is produced by the gut microbiota through fermentation of these prebiotics (Li et al 2017, Zeng et al 2017, Zhang et al 2016). Referred to as “the butyrate paradox,” butyrate characteristically stimulates the growth of noncancerous colonocytes while simultaneously inducing proliferation arrest and apoptosis of colorectal cancer cells (Li et al 2017, Li et al 2018). In cancerous colonocytes, the

Warburg effect prevents the metabolism of butyrate allowing for preferential accumulation in the cell where it acts as an HDAC inhibitor (Li et al 2018, Zhang et al 2016). HDAC3 is known to be overexpressed in colon tumors and plays a role in the metastasis of several cancer cells through the cytoskeleton and myosin assembling Akt1 and ERK1/2 signaling

pathways; however, butyrate significantly inhibits HDAC3 to block the activation of these pathways thus inhibiting the migration and invasion of colorectal cancer cells (Li et al

2017). The ERK1/2 pathway is also important in cell proliferation and the inhibition of this pathway by butyrate induces cell cycle arrest in cancerous colonocytes. The most crucial effector in butyrate-induced cell cycle arrest is the protein p21, a known tumor suppressor and negative regulator of cell proliferation, where the butyrate related histone acetylation induces expression of p21 causing apoptosis (Zeng et al 2017). Additionally, butyrate has been reported to preferentially inhibit cell proliferation of colorectal cancer cells by binding to and activating the PKM2 protein which reverses the metabolic advantages of cancerous colonocytes (Li et al 2018).

1.3.5. Modulation of ion transport: SCFAs, particularly butyrate, stimulate ion and water absorption and play a fundamental role in keeping the intestinal mucosa hydrated

(Canani et al 2004, Vidyasagar and Ramakrishna 2002). Butyrate modulates ion transport by stimulating the electroneutral NaCl absorptive mechanism and by limiting Cl ̄ secretion leading to implications for the treatment of enterotoxin or genetic secretory diarrheal diseases and cystic fibrosis (Canani et al 2004, Matthews et al 1998). During electroneutral

NaCl absorption, butyrate modulates transepithelial ion transport through activation of parallel Cl ̄ /butyrate and Na+/H+ exchangers NHE2 and NHE3 and secondarily through

+ + the upregulation of Na /H and Cl ̄ /HCO3 ̄ exchangers (Canani et al 2004, Canani et al

2013). The anti-secretory effect on Cl ̄ is due to butyrate inhibition of intestinal cAMP- dependent Cl ̄ secretion in the short term and by downregulation of Na-K-2Cl cotransporter expression and function in the long term (Matthews et al 1998).

1.3.6. Improvement of oxidative status: Oxidative stress causes cellular damage and occurs when there is an imbalance between the production of reactive oxygen species and host antioxidant defense mechanisms (Hamer et al 2009, Ma et al 2018). While the exact mechanism of butyrate’s effect on oxidative status is unknown, it is clear that butyrate does not act as primary antioxidant and instead affects gene expression for enzymatic antioxidant activity (Hamer et al 2009). Most importantly, butyrate increases the antioxidant capacity and colonic concentration of the antioxidant glutathione where low levels of glutathione cause severe degradation of intestinal epithelial cells (Hamer et al

2009, Vanhoutvin et al 2009). The expression of glutathione peroxidases GPX1 and GPX3 as well as glutathione reductase are increased in the glutathione oxidative stress pathway thus increasing its turnover capacity (Vanhoutvin et al 2009). Butyrate has been reported to increase the colonic concentrations of glutathione by increasing the gene expression of glutathione and the catalytic subunit of glutamine-cysteine ligase which is the rate limiting step in glutathione production (Hamer et al 2009).

1.4. Animal digestive systems and butyrate production

1.4.1. Monogastric digestive system: Monogastric digestion refers to a digestive tract with a single chambered stomach and is found in animals such as humans, horses, and pigs (Figure 1.2.; Baker 2017, Rowan et al 2015). Food enters the digestive tract through the mouth where the grinding of teeth mechanically breaks down food into smaller pieces and enzymes contained within saliva begin chemical digestion. The food then moves down the esophagus and into the stomach where it is further broken down by hydrochloric acid and gastric enzymes. Most nutrients are absorbed in the next part of the digestive tract known as the small intestine which is lined with many small villi to increase its surface

area and allow for a greater absorptive area. Enzymatic digestion of food also continues in the small intestine to further break down carbohydrates, proteins, and fats. Any remaining food particles and secretions pass into the large intestine. The beginning of the large intestine contains the cecum or “blind gut” containing large amounts microorganisms. In most animals the cecum has little to no effect on digestion, but in other animals that utilize hind-gut fermentation to break down plant material it is a crucial structure. The next part of the large intestine is the colon which functions to absorb water and to mold the waste into a solid form which then exits the digestive tract as feces through the rectum and the anus (Rowan et al 2015).

Figure 1.2. An example of the monogastric digestive system (QA International 2003).

1.4.2. Microbiome and butyrate production in monogastric animals:

Monogastric animals are comprised of herbivores, carnivores, and omnivores with the

diversity of their gut microbiota influenced by diet type; nevertheless, there are similarities in the gastrointestinal ecosystems and its underlying principles across animal species

(Koboyashi et al 2020, O’Donnell et al 2017, Richards et al 2005). Fermentation occurs primarily in the hindgut of the monogastric digestive system as the stomach and proximal small intestine are challenging environments for bacterial growth due to high acidity and rapid movements (Koboyashi et al 2020, Richards et al 2005). The microbiome of the large intestine consists of over 99% strict anaerobes where at least 80% of organisms are from the phyla Bacteroidetes and Firmicutes with Clostridia and Bacteroidia as the predominant classes and Clostridiales and Bacteroidales as the predominant orders (Koboyashi et al

2020, O’Donnell et al 2017, Richards et al 2005). Bacterial utilization of fermentable carbohydrates in the large intestine produces primarily the short chain fatty acids acetate, propionate, and butyrate which are absorbed from the apical membrane for use as energy and nutrients as well as stimulating the development of intestinal physiology and host defenses (Jha et al 2019, Richards et al 2005, Williams et al 2019). Although SCFAs are produced at a ratio of about 60% acetate, 25% propionate, and 15% butyrate, butyrate is particularly vital in maintaining the gut ecosystem as well as improving gut mucosal health and the immune system (Jha et al 2019). Intestinal epithelial cells preferentially utilize butyrate as an energy source to supplying 60-70% of colonocyte energy requirements as well as up to 15% of the maintenance energy required by the host. Butyrate has been shown to influence gut metabolic pathways by affecting cellular growth and metabolism to enhance the epithelial barrier by promoting proliferation of the intestinal mucosal epithelium, increase mucus production by stimulating goblet cell differentiation, and promote immune cell differentiation and functioning (Jha et al 2019, Williams et al 2019).

The production of butyrate is also important in maintaining intestinal homeostasis by keeping the environment anaerobic to prevent gut dysbiosis resulting in increased susceptibility to various diseases (Jha et al 2019).

1.4.3. Polygastric digestive system: Ruminant animals such as cattle, sheep, and goats have a polygastric digestive system defined by a compartmentalized stomach specialized for obtaining energy from fibrous plant material (Figure 1.3) (Parish 2017).

Rumination begins when food enters the mouth where it is briefly mechanically crushed by teeth and mixed with saliva. Saliva begins the enzymatic breakdown of the food and, most importantly, functions to buffer the pH in the rumen and reticulum. The food then passes down the esophagus into the rumen; however, the esophagus functions bidirectionally in a process called rumination. During rumination, the food is forced back into the mouth where it can be further broken down into smaller pieces and mixed with more saliva before being swallowed again. Microbial digestion of food occurs in the rumen where papillae allow for increased surface area and better absorption of nutrients produced during fermentation. Small food particles are then collected in and pass through the reticulum where they enter the omasum which has a highly folded topography to increase surface area and remove water as well as to further grind down food. Next the food enters the abomasum or “true stomach” where hydrochloric acid and digestive enzymes break down food and kill the rumen microbes. Enzymatic digestion continues and most nutrient absorption occurs in the small intestine. Secondary fermentation of undigested fiber occurs in the large intestine along with the absorption of water. The remaining food particles are expelled from the digestive tract through the rectum and anus (Moran 2005, Parish 2017).

Figure 1.3. An example of the polygastric digestive system (Addison Wesley Longman INC. 1999).

1.4.4. Microbiome and butyrate production in polygastric animals: In polygastric animals, the rumen contains a diverse microbial ecosystem required to convert fibrous plant material into products necessary for host energy, maintenance, and growth

(Miguel et al 2019, Seshadri et al 2018). While the rumen contains organisms from a wide taxonomic range including bacteria, archaea, protozoa, fungi, and viruses, this anaerobic environment is dominated by fermentative bacteria (Aluwong et al 2010, Li et al 2012).

The rumen bacterial microbiome consists of more than 90% of organisms from the phyla

Bacteroidetes and Firmicutes with Clostridia and Bacteroidia as the predominant classes and Clostridiales and Bacteroidales as the predominant orders (Firkins et al 2015). These bacteria primarily produce the SCFAs acetate, butyrate, and propionate which are absorbed across the rumen epithelium as a nutrient and energy source as well as regulating and signaling molecules of host physiology (Aluwong et al 2010, Li et al 2012, Seshadri et al

2018). Acetate and propionate are produced in greater concentrations than butyrate with

butyrate representing 5-20% of the total proportion of SCFAs; however, butyrate functions the most dynamically and decreased concentrations are associated with dysfunction in gene regulation and disease prevention (Aluwong et al 2010, Gorka et al 2018). Butyrate significantly affects animal performance as it is required for fat production in milk and body composition, is the preferred energy source of ruminal epithelial cells, and influences postnatal development and function of the rumen (Li et al 2012, Miguel et al 2019, Moran

2005). As an energy source, it provides up to 70% of the host’s daily energy requirements and it provides a majority of the energy precursors required for metabolic processes (Li et al 2012, Miguel et al 2019). Butyrate is also required to stimulate and regulate the growth and function of the ruminal epithelium as it is important for the development of the rumen papillae and therefore effects the surface area of the rumen and nutrient absorption (Gorka et al 2018). This effect is due to butyrate’s function as a histone deacetylase inhibitor allows it to mediate growth associated genes and signaling pathways and influence the metabolic activity and blood flow of the ruminal epithelial cells, abundance of transcript proteins involved with SCFA absorption, and rumen motility (Gorka et al 2018, Lin et al

2019).

Much less is known about the microbiome and the effect of butyrate in the lower gastrointestinal tract of polygastric animals; although, it is believed to be similar to that of monogastric animals (Gorka et al 2018). It is reported that about 8.6-16.8% of ruminant

SCFAs are also produced in the large intestine by secondary fermentation of undigested carbohydrates and microbes surviving the upper gastrointestinal tract have an opportunity to self-colonize other gastrointestinal compartments (Aluwong et al 2010, Taschuk and

Griebel 2012).

2 OBJECTIVES

The aim of this study was to characterize and identify potential probiotic species of Clostridium from animal origin to find novel therapeutic strains for the maintenance of overall health or the treatment of disease. We hypothesized that intestinal butyrate producing Clostridium can be isolated from the feces of animals and that some of these isolates will have the potential to be used safely as a probiotic dietary supplement.

Our first objective was to analyze fecal samples from different animal species for the presence of butyrate producing Clostridium by enriching for, isolating, and identifying Clostridium species with anaerobic culture techniques, staining, biochemical testing, and 16S rRNA sequencing. Analysis by PCR was used to determine the ability of the Clostridium isolates to produce butyrate via the presence of the butyrate kinase gene.

Our second objective was to assess the safety of the Clostridium isolates through hemolysis results and antibiotic susceptibility testing based on standards outlined by the

European Food and Safety Authority. Our third objective was to biochemically characterize potentially safe Clostridium isolates using API microbial identification kits and to begin investigating butyrate production by determining the endpoint of acidogenesis.

3 MATERIALS AND METHODS

3.1. Animal demographics

Fecal samples were collected from the farm at California State Polytechnic

University, Pomona (CPP) during the year of 2018. Horse, cattle, sheep, pig, and goat samples were collected on June 25th, August 3rd, October 29th, November 25th, and

December 14th respectively. All animals were reported to be in good health and no animals received any probiotic supplements or antibiotic treatments at the time of collection. Horses were housed in individual pens with one adult horse or one adult horse and one foal per pen. Cattle were housed in individual pens containing one to three animals per pen. Sheep, pigs, and goats were housed in communal pens with one animal type and at least five animals per pen. Specific information for each animal is listed in Tables 3.1, 3.2, 3.3, 3.4, and 3.5.

Table 3.1. Details of horses from which fecal samples were collected. Animal Identification Gender Age Breed Feed (daily) Star Female 4 years Thoroughbred 9 flakes alfalfa 3 flakes orchard Horse 1 Purebred Starlit Dancer Male 3 months 8 lbs Growth Arabian supplement 2 flakes alfalfa Purebred 3 flakes orchard Horse 2 B’witched Female 14 years Arabian 2 lbs Growth supplement 2 flakes alfalfa Dancin’ Purebred 3 flakes orchard Horse 3 Female 7 years Shoes Arabian 2 lbs Growth supplement Socks Female 4 years Thoroughbred 9 flakes alfalfa 3 flakes orchard Horse 4 Purebred Fiona Female 3 months 6 lbs Growth Arabian supplement 2 flakes alfalfa Purebred 1 flake orchard Horse 5 Truly Yours Male 3 years Arabian 11 lbs Renew Gold supplement

Table 3.2. Details of cattle from which fecal samples were collected. Animal Identification Gender Age Breed Feed Cattle 1 (No ID) Male 1 year Cross breed Cattle 2 49 Male 7 years Angus Cattle 3 1455 Female 4 years Holstein Oat hay 45 Female 5 months Cross breed Corn silage Cattle 4 7543 Female 5 months Cross breed Mixed grain (No ID) Male 5 months Cross breed Trace mineral block 717 Male 1 year Angus Cattle 5 705 Male 1 year Angus

Table 3.3. Details of sheep from which fecal samples were collected. Animal Identification Gender Age Breed Feed D446 Female 4 years Dorset D511 Female 4 years Dorset D519 Female 3 years Dorset D524 Female 3 years Dorset D615 Female 2 years Dorset D702 Female 2 years Dorset D703 Female 2 years Dorset D726 Female 2 years Dorset 1601 Female unknown unknown 1703 Female unknown unknown 2512 Female 2 months Sulfix/ Alfalfa hay Hampshire Sheep Four-way grain 2517 Female 1 month Dorset Trace mineral block 2518 Female 1 month Dorset 2531 Female 1 month Dorset/ Hampshire 2534 Male 1 month Dorset 2536 Female unknown unknown 2552 Female 1 month Hair 2555 Female 1 month Dorset/Hair 13135 Female 5 years Sulfix/ Hampshire 13144 Female 5 years Dorset/Sulfix/ Hampshire

Table 3.4. Details of pigs from which fecal samples were collected. Animal Identification Gender Age Breed Feed 3 Female 5 years Yorkshire 8 Female 4 years Berkshire Hampshire/ Western Milling Pigs 10 Female 4 years Duroc commercial bred and

Hampshire/ nurse sow diet 21 Female 5 years Yorkshire 28 Female 3 years Hampshire

Table 3.5. Details of goats from which fecal samples were collected. Animal Identification Gender Age Breed Feed 006 Female 4 years Nubian/Boer 026 Female 1 year Boer 046 Female 1 year Boer Alfalfa hay 1174 Female unknown Nubian Goats Four-way grain 2051 Female 3 years Boer Trace mineral block 2520 Female 3 months Boer 2523 Female 3 months Boer 2530 Female 3 months Boer

3.2. Labeling of bacterial isolates

Each bacterial isolate was given a unique three-part label based on the animal type, sample number, and bacterial colony with the letters and numbers listed in Table 3.6. The different animal types were assigned a letter based on the first letter of their common name; the five samples collected from each animal type were assigned a number from 1-5; and the different colony morphologies seen in the initial streak to isolation were assigned a letter from A-J. For example: the label H1A denotes “H” for horse as the animal type, “1” as the horse sample number, and “A” as the bacterial colony.

Table 3.6. Isolate labeling system. Animal Type Label Horse H Cattle C Pig P Sheep S Goat G Sample Number 1-5 Bacterial Colony A-I

3.3. Fecal sample collection

Horse and Cow fecal samples were collected from five individual pens that the animals were housed in. Most pens housed a single animal; however, some pens contained two or three as there were no other pens available with only one animal. Each sample was collected by compiling three different fecal piles of varying age from one pen into a sterile

Whirl-Pak bag with the use of a sterile tongue depressor. For the horse sample collection, foal fecal piles were avoided if they were housed in the same pen as an adult horse.

Pig, sheep, and goat fecal samples were collected from one larger pen that housed at least five animals of the same type. The pens were broken up into five sections made up of the four corners plus the center of the pen. Each sample was collected by compiling three different fecal piles of varying age from one section of the pen into a sterile Whirl-

Pak bag with the use of a sterile tongue depressor. Samples were processed immediately after collection and then stored at -20°C.

3.4. Bacterial culture media and reagents

3.4.1. Anaerobic conditions were used for all culturing procedures unless otherwise noted. Broths were dispensed into Hungate tubes, flushed with nitrogen gas, and then capped. Hungate tubes utilize a gas-tight screw cap to maintain anaerobic conditions within the tube. The caps also contain a butyryl rubber stopper to allow for the use of a

needle and syringe when transferring cultures between tubes. Plates were placed within a gas-tight, anaerobic jar and reduced 24 hours prior to inoculation with the use of an

Anoxomat Mark II (Advanced Instruments; Norwood, Massachusetts) which creates an anaerobic environment by replacing the air inside of the jar with nitrogen gas. Inoculated plates were kept inside of an anaerobic jar and reduced with the use of the Anoxomat Mark

II. Only the inoculated plates used for spore staining were kept inside of an anerobic jar and reduced by placing an AnaeroPack (Mitsubishi Gas Chemical; Tokyo, Japan; catalog

#R681001) within the jar to chemically remove the oxygen.

3.4.2. Thioglycolate broth was purchased premade from Biomérieux (Marcy- l’Étoile, France; catalog #44001) and stored at 4°C.

3.4.3. Cooked meat media (CMM) was made by suspending 1 g of cooked meat media (HiMedia; Mumbai, India; catalog #M149S-500G) in 10 ml of deionized water in a

Hungate tube, flushed with nitrogen, and capped. The tubes were then autoclaved at 121°C for 20 minutes and stored at room temperature.

3.4.4. Egg yolk agar (EYA) was made by mixing 40 g of trypticase peptone (BD;

Franklin Lakes, New Jersey; catalog #211921), 5 g of sodium phosphate dibasic (Acros

Organics; Fair Lawn, New Jersey; catalog #20651500), 2 g of sodium chloride (Promega;

Madison, Wisconsin; catalog #H5273), 0.02 g of magnesium sulfate (Sigma-Aldrich; St.

Louis, Missouri; catalog #M-2773), 2 g of D-glucose (Sigma-Aldrich; St. Louis, Missouri; catalog #50-99-7), 5 g of yeast extract (Oxoid; Basingstoke, United Kingdom; catalog

#LP0021), and 15 g of agar (Fisher Scientific; Hampton, New Hampshire; catalog

#BP1423-500) with 900 ml of deionized water. The media, a bottle containing 50 ml of

PBS, and a 250 ml beaker containing a stir bar were then autoclaved at 121°C for 20

minutes. After autoclaving, the media and PBS were cooled in a 55°C water bath. The surface of three eggs was sterilized by spraying them with 70% ethanol and placing them in sterile petri plates under UV light and turning them every 30 minutes for 2 hours. Under a laminar flow hood, the egg shells were broken using a spatula and tongs sterilized with

70% ethanol. The yolk was separated and placed into the autoclaved 250 ml beaker. Once

50 ml of egg yolk was obtained, 50 ml of cooled PBS was added and then mixed using the stir bar. Then, the egg yolk and PBS mixture was added to the cooled autoclaved media, mixed, and poured into sterile petri plates. Once solidified, the plates were stored at 4°C.

3.4.5. Blood agar plates (BAP) were purchased premade as 5% sheep’s blood in a tryptic soy agar base (Ward’s Science; Rochester, New York; catalog #470180-680 and

Hardy Diagnostics; Santa Maria, California; catalog #89405-024) and stored at 4°C.

3.4.6. Trypticase peptone glucose yeast extract (TPGY) was made by mixing 50 g of trypticase peptone, 5 g of peptone (BD; Franklin Lakes, New Jersey; catalog #211677),

20 g of yeast extract, and 4 g of D-glucose with 1 L of 28nglish28d water. The recipe was modified only for NMR testing by increasing the amount of D-glucose to 15 g. For liquid media, 10 ml of the media was aliquoted per Hungate tube, flushed with nitrogen, and capped. The tubes were then autoclaved at 121°C for 20 minutes and stored at room temperature. For solid media, 15 g of agar was also added and the media was autoclaved at 121°C for 20 minutes. After cooling in a 55°C water bath, the media was poured into sterile petri plates. Once solidified, the plates were stored at 4°C.

3.4.7 Phosphate buffered saline (PBS) was made as a 10X stock by mixing 8 g of sodium chloride, 2 g of potassium chloride (Fisher Scientific; Hampton, New Hampshire; catalog #P217-500), 2.68 g of sodium phosphate dibasic, and 2.4 g of potassium phosphate

monobasic (Sigma-Aldrich; St. Louis, Missouri; catalog #10049-21-5) with 250 ml of deionized water. The pH was adjusted to 7.4 using hydrogen chloride and sodium hydroxide solutions then the volume was adjusted to 1 L with deionized water. When needed, the stock was diluted to 1X by mixing 50 ml of 10X PBS with 450 ml deionized water and autoclaving at 121°C for 20 minutes. The 10X PBS was stored at 4°C and the

1X PBS was stored at room temperature.

3.4.8. Brucella broth was made by dissolving 28 g of Brucella broth powder (BD;

Franklin Lakes, New Jersey; catalog #211088) in 1 L of deionized water. The broth was aliquoted at 10 ml per Hungate tube, flushed with nitrogen, and capped. The tubes were then autoclaved at 121°C for 20 minutes and stored at room temperature.

3.4.9. Vitamin K1 (1 mg/ml) was made by diluting a storage stock solution of 100 mg/ml vitamin K1. The storage stock solution was made by mixing 100 mg of vitamin K1

(Alfa Aesar; Haverhill, Massachusetts; catalog #L10575) with 1 ml of 100% ethanol

(Acros Organics; Fair Lawn, New Jersey; catalog #61509-0010). The working stock of 1 mg/ml vitamin K1 was made by mixing 0.1 ml of the storage stock with 9.9 ml of 100% ethanol and filter sterilized using a sterile syringe filter with a 0.2µm cellulose acetate membrane. Both the storage and working stocks were stored in a dark, sterile container at

-20°C.

3.4.10. Hemin (5 mg/ml) was made by mixing 40 mg of 1 N sodium hydroxide

(Fisher Scientific; Hampton, New Hampshire; catalog #S78605) in 1 ml of deionized water and then dissolving 50 mg of bovine hemin (Sigma-Aldrich; St. Louis, Missouri; catalog

#16009-13-5) into the sodium hydroxide solution. The volume was brought up to 10 ml by adding 9 ml of deionized water and filter sterilized using a sterile syringe filter with a

0.2µm cellulose acetate membrane. The solution was aliquoted into 1 ml volumes and stored at -20°C.

3.4.11. Laked sheep blood was made by freezing sterile sheep blood (Hemostat

Laboratories; Dixon, California; catalog #NC0581834) in 50 ml aliquots at -20°C for a minimum of 24 hours. Before use, the frozen sheep blood was thawed at room temperature then warmed to 55°C in a water bath for 5 minutes.

3.4.12. Antibiotic solutions for ampicillin (Shelton Scientific; catalog #IB02040), vancomycin (Alfa Aesar; Haverhill, Massachusetts; catalog #J62790), gentamicin (Fisher

Scientific; Hampton, New Hampshire; catalog #BP918-1), kanamycin (Shelton Scientific; catalog #IB02120), streptomycin (Sigma-Aldrich; St. Louis, Missouri; catalog #3810-74-

0), clindamycin (Alfa Aesar; Haverhill, Massachusetts; catalog #J61409), and tetracycline

(Sigma-Aldrich; St. Louis, Missouri; catalog #64-75-5) were each made by dissolving 100 mg of the appropriate powder in 10 ml of ultrapure water. Solutions for chloramphenicol

(Sigma-Aldrich; St. Louis, Missouri; catalog #56-75-7) and erythromycin (Sigma-Aldrich;

St. Louis, Missouri; catalog #114-07-8) were made by dissolving 100 mg of the appropriate powder into 10 ml of 100% ethanol. All antibiotic solutions were filter sterilized using a sterile syringe filter with a 0.2µm cellulose acetate membrane. All solutions were aliquoted into 1 ml volumes and stored at -20°C.

3.4.13. Modified Brucella blood agar was supplemented with 5 µg/ml hemin, 1

µg/ml vitamin K1, and 5% v/v laked sheep blood as recommended by the Clinical

Laboratory and Standards Institute (CLSI). A modification was made to the protocol by raising the agar concentration from 1.5% to 3% to slow swarming activity. The media was made by mixing 28 g of Brucella broth powder, 1 L of deionized water, 1 ml of 1 mg/ml

vitamin K1, 1 ml of 5 mg/ml hemin solution, and 30 g of agar. The media was aliquoted into 80 ml volumes and autoclaved at 121°C for 20 minutes then placed in a 55°C water bath. Once the media was cooled, 4 ml of warm laked sheep blood was added to each aliquot of autoclaved media and gently mixed. Antibiotics were added to the media at this time at the concentrations listed in Table 3.11, gently mixed again, and poured into sterile petri plates. Once solidified, the plates were placed in a 37°C incubator with the lids slightly ajar for 45 minutes to allow excess moisture to evaporate. The plates were stored up to 7 days at 4°C and were warmed to room temperature prior to use.

3.4.14. Nitric acid (3 N) was made in a fume hood by slowly mixing 187.5 ml of

16 N nitric acid (Fisher Scientific; Hampton, New Hampshire; catalog #A200-500) into

812.5 ml of ultrapure water in a glass bottle. The solution was stored in a flammable cabinet at room temperature.

3.5. Enrichment of Clostridium species

On the day of collection, fecal samples were homogenized within their sample collection bags by hand and 5 g were weighed into a sterile 50 ml conical tube using a sterile tongue depressor. This was followed by adding 10 ml of thioglycolate broth to the weighed fecal sample and vortexing until thoroughly mixed. Wide mouth pipette tips were made by cutting the end of a pipette tip with sterilized scissors. These pipette tips were then used to transfer 1 ml of the stool and thioglycolate mixture into a tube of cooked meat media which was then heat shocked by placing it in an 80°C water bath for 5 minutes. The inoculated media was incubated at 37°C with shaking at 200 rpm for 72 hours.

3.6. Isolation of Clostridium species

A loop full of the enriched cooked meat media broth was taken and streaked to isolation onto reduced EYA plates. The plates were incubated anaerobically at 37°C for 24 hours. Individual colonies with differing morphology were then taken from the plates, re- streaked onto reduced EYA plates, and incubated anaerobically at 37°C for 24 hours until a pure culture was obtained. Once well-isolated colonies were obtained, the colony morphology of the isolate was recorded. If at least five different colony morphologies were not seen on the plate, then different colonies with the same morphology were chosen. Stock cultures of each isolate were made by taking one colony from the pure culture plate and inoculating it into cooked meat media, incubating it at 37°C with shaking at 200 rpm for

24 hours, and then storing it at room temperature.

3.7. Identification and characterization of Clostridium isolates

Isolates were streaked onto reduced EYA plates and were incubated anaerobically at 37°C for 24 hours. One colony was Gram stained and the Gram reaction, cell shape, and cell arrangement of the organism were recorded. Lipase, lecithinase, and proteolytic activity were also determined at this time. Lipid hydrolysis through the production of lipase was determined by the presence of an iridescent sheen over the bacterial colonies; lecithinase activity was determined by a zone of white opaque precipitation within the media that surrounds the colonies; and proteolysis was determined by the appearance of a clear zone in the media surrounding the colonies. The streaked EYA plates were kept and incubated anaerobically at room temperature for 6 more days. One colony was spore stained and the spore reaction, spore shape, spore location, and presence of cell bulging were recorded.

Isolates were inoculated onto TPGY plates and were incubated aerobically at 37°C for 24 hours to determine if the organism could grow in the presence of oxygen. Isolates were inoculated onto reduced TPGY plates and were incubated anaerobically at 37°C for

24 hours. Afterwards, 3% H2O2 was added to the growth and the formation of bubbles indicated the production of catalase.

Hemolytic activity was determined by streaking the isolates onto reduced BAP plates and were incubated anaerobically at 37°C for 24 hours. Gamma-hemolysis was indicated by bacterial colonies with no change to the media; alpha-hemolysis was indicated by a green to brown zone in the media surrounding the colonies; and beta-hemolysis was indicated by a clear zone in the media surrounding the colonies. A double-zone of hemolysis is characteristic of C. perfringens and was indicated by an inner zone of beta hemolysis and an outer zone of alpha hemolysis surrounding the colony.

The bacterial isolates that were Gram positive, rod-shaped, spore formers, anaerobic, and catalase negative were presumed to be Clostridium. Furthermore, the isolates that were lecithinase positive and strongly beta hemolytic were presumed to be unsafe for use as a probiotic and did not continue for further testing.

3.8. DNA purification

3.8.1. DNA purification from fecal samples: Bacterial DNA was purified from fecal samples on the day of collection using the QIAamp PowerFecal DNA Kit (Qiagen;

Hilden, Germany; catalog #12830-50) following the manufacturer’s instructions. DNA was eluted using 50 µl of solution C6 and the concentration of the DNA was determined using a nanophotometer (Implen, Inc.; Westlake Village, California). The DNA samples were stored at -20°C.

3.8.2. DNA purification from presumed Clostridium isolates: Bacterial cultures were prepared by inoculating 0.1 ml of the cooked meat media stock culture into 10 ml of reduced TPGY broth and incubating at 37°C with shaking at 200 rpm for 12-18 hours.

Bacterial DNA was purified from the cultures using the Easy-DNA Kit (Invitrogen;

Carlsbad, California; catalog #45-0424) following the manufacturer’s instructions. A modification was added to the procedure by adding 3-5 mg of lysozyme from chicken egg white (Sigma-Aldrich; St. Louis, Missouri; catalog #L6876-5G) to the mixture of the cell suspension and solution A, followed by incubating at 65°C for 10 minutes with vortexing every 3 minutes, and then incubating at 37°C for 10 minutes with vortexing every 5 minutes. The concentration of the DNA was determined using a nanophotometer. If the concentration was greater than 200 ng/µl, then the sample was diluted using TE buffer provided in the kit. The DNA samples were stored at -20°C.

3.9. Polymerase Chain Reaction (PCR)

3.9.1 PCR primers: Primers for the butyrate kinase (BUK) gene were designed based on the primers described by Vital et al (2012). The same primer regions were used and only the degeneracy was modified using the designators in Table 3.7. The degenerated bases were determined by a multiple sequence alignment of the species listed in Table 3.8 using T-Coffee (Appendix A). All bases that were not similar were replaced with the appropriate degenerate base. Primers for the 16S rRNA gene remained unchanged from their reference as outlined in Table 3.9. Primers were synthesized by Fisher Scientific

(Hampton, New Hampshire) and were reconstituted to a concentration of 100 µM with sterile ultrapure water. A 10 µM working stock was made by diluting the 100 µM storage stock with sterile ultrapure water. All primers were stored at -20°C.

Table 3.7. Variable nucleotide symbols. Designator Nucleotides Y C or T W A or T R A or G S C or G H A or C or T D A or G or T N A or C or G or T

Table 3.8. Clostridium species used in BUK gene multiple sequence alignment. Organism Strain GenBank ID Clostridium butyricum 60E.3 NZ_KB851133.1 Clostridium beijerinckii ATCC 35702 CP006777.1 Clostridium sporogenes ATCC 15579 ABKW02000002.1 Clostridium saccharoperbutylacetonicum ATCC 27021 CP004121.1 Clostridium saccharobutylicum ATCC BAA-117 CP006721.1 Clostridium acetobutylicum ATCC 824 NC_003030.1 Clostridium perfringens ATCC 3626 ABDV01000013.1 Clostridium botulinum NCTC 8550 CP010521.1

Table 3.9. Primer sequences for PCR amplification. Product Primer Sequence (5’-3’) Reference (bp) 16S27F AGAGTTTGATCCTGGCTCAG Lane, 1991 1465 16S1492R GGTTACCTTGTTACGACTT Turner et al, 1999 BUKF ATHAATCCHGGNTCDACHTCWACWAA Vital et al, 2013 464 BUKR ACHGCYTTTTGATTHARWGCATG and this study

3.9.2. PCR assay: The 16S rRNA gene was used as a DNA positive control and for species identification through sequencing. The reaction mix contained 200 ng of template DNA, 0.5 µM of each primer, 10 µl Taq 2X Master Mix (New England BioLabs;

Ipswich, Massachusetts; catalog #M0270S), and filled to 20 µl with sterile ultrapure water.

The BUK gene was used to detect butyrate producing organisms. The reaction mix contained 200 ng of template DNA, 0.75 µM of each primer, 10 µl Taq 2X Master Mix, and filled to 20 µl with sterile ultrapure water. Both genes used C. sporogenes ATCC 11437 as a positive control and sterile ultrapure water as a negative control. The PCR reaction

was performed in a GeneAmp PCR System 9700 thermocycler (Applied Biosystems;

Foster City, California) using the settings in Table 3.10.

Table 3.10. PCR reaction settings for the 16S rRNA and BUK genes Step Temperature (°C) Time (minutes) Cycles Initial denaturation 94 5 1 Denaturation 94 0.75 Annealing 55 0.75 30 Elongation 72 1 Final extension 72 7 1

3.9.3. Gel electrophoresis: All PCR products were visualized using agarose gel electrophoresis. Agarose gels were made by dissolving 1.5% w/v of Aquapor agarose

(National Diagnostics; Atlanta, Georgia; catalog #50-899-90020) in 1X TAE

(ThermoFisher Scientific; Waltham, Massachusetts; catalog #B49).

For determining PCR product size, the GeneRuler 1 kb Plus DNA Ladder (Thermo

Fisher Scientific; Waltham, Massachusetts; catalog #FERSM1331) and PCR products were mixed 1:6 with 6X DNA Loading Dye (Thermo Fisher Scientific; Waltham,

Massachusetts; catalog #FERSM1331). The gel was run in 1X TAE at 100 V for 25 minutes using a Gel XL Mini Gel Migration Tank (Labnet International) followed by post- staining in 1X SYBR Green with gentle agitation for 25 minutes. The stained gel was photographed using a Gel Doc XR+ with Image Lab Software (Bio-Rad Laboratories;

Hercules, California).

For determining PCR product concentration before sequencing, the GeneRuler 1 kb Plus DNA Ladder and purified PCR products were mixed 1:5 with 5X DNA Loading

Dye/SYBR Green. The gel was run in 1X TAE at 100 V for 15 minutes using a Gel XL

Mini Gel Migration Tank. The gel was analyzed using a Gel Doc XR+ with Image Lab

Software.

3.10. Crude sample gene analysis

The band intensities of the 16S rRNA and BUK genes amplified from the crude

fecal samples were measured using ImageJ with the 16S rRNA bands used to normalize

the data. GraphPad Prism was used for statistical analysis with P-values calculated by the

Mann-Whitney test. Statistical significance was defined as P < 0.05.

3.11. Clostridium species determination

3.11.1. Sequencing: After amplifying the 16S rRNA gene for all presumed

Clostridium isolates, PCR products were purified using the QIAquick PCR Purification Kit

(Qiagen; Hilden, Germany; catalog #28106) following the manufacturer’s instructions.

The concentration of the purified PCR products were determined by gel electrophoresis

and comparing the intensity of the bands to the GeneRuler 1 kb Plus DNA Ladder using

the Gel Doc XR+ with Image Lab Software. The concentration of purified PCR products

were adjusted to 20 ng/µl by diluting with buffer EB from the QIAquick PCR Purification

Kit. In a 96-well plate, 100 ng PCR product was mixed with 2 µl of 10 pmole 2200 16S-

27F primer and ultrapure water to a total of 12 µl. Samples were then sent to Retrogen, Inc.

(San Diego, California) for sequencing.

3.11.2. Species determination: The National Center for Biotechnology

Information (NCBI) basic local alignment search tool (BLAST) was used to align

sequences with an rRNA/ITS database of bacterial 16S rRNA nucleotide sequences in

order to determine the genus and species of the presumed Clostridium isolates.

EZBioCloud’s 16S-based ID was also used to align the sequences with a database of

quality controlled 16S rRNA to corroborate the results determined by NCBI BLAST. A

similarity of at least 95% was used as a cut-off for genus identification and 98.7% for species identification for both NCBI BLAST and EZBioCloud.

3.11.3. Limiting Clostridium sporogenes isolates: Due to the high number of isolates determined to be C. sporogenes, those that were from the same animal type and sample number that shared the same morphological, biochemical, and 16S rRNA sequence results were considered to be the same organism.

3.12. Antibiotic susceptibility

Susceptibility to antibiotics was determined using the Wadsworth agar dilution method referenced by the Clinical and Laboratory Standards Institute (CLSI). However, isolates were only tested at the microbiological cut-off values used for viable microorganisms as the active agent in feed additives set by the European Food Safety

Authority (EFSA) as outlined in Table 3.11. C. perfringens ATCC 13124 and B. fragilis

ATCC 25285 were used as quality control strains while sterile Brucella broth was used as a negative growth control.

Isolates were streaked onto reduced TPGY plates and were incubated anaerobically at 37°C for 24 hours. A bacterial suspension of each isolate in Brucella broth equivalent to a 0.5 McFarland standard density was made using the colonies on the TPGY plates. Each isolate was added in triplicate to a 96-well plate at 200 µl of bacterial suspension per well.

A multichannel pipette was used to inoculate 10 µl of each bacterial suspension onto the

Brucella agar plates. The antibiotic containing plates were inoculated starting with the bacteriostatic antibiotics first (clindamycin, erythromycin, tetracycline, and chloramphenicol) followed by the bactericidal antibiotics (gentamicin, streptomycin, kanamycin, ampicillin, and vancomycin). At the beginning and end of each set of antibiotic

plates, two plates containing no antibiotics were inoculated with one plate serving as a growth control and the other as an aerobic contaminate control. To allow the inoculum to absorb into the media, the inoculated plates were left upright for 10 minutes before inverting. The aerobic contaminate control plate was incubated aerobically at 37°C for 48 hours and all other plates were incubated anaerobically at 37°C for 48 hours.

Isolates were determined to be susceptible to an antibiotic if growth was inhibited at the cut-off value or resistant if growth was not inhibited at the cut-off value (EFSA

2020). Isolates that were resistant to any antibiotic and did not demonstrate intrinsic resistance were considered unsafe for use as a probiotic.

Table 3.11. Selected bacterial cut-off values (mg/L) for antimicrobials of human and veterinary importance as required by the EFSA (EFSA 2012).

nicol

Ampicillin Vancomycin Gentamicin Kanamycin Streptomycin Erythromycin Clindamycin Tetracycline Chloramphe Other Gram + 1 2 4 16 8 0.5 0.25 2 2

3.13. Biochemical testing

Sequenced isolates that were determined to be safe for use as a probiotic were biochemically characterized with the use of the microbial identification kits API 50CH

(Biomérieux; Durham, North Carolina; catalog #50300) with 50 CHB/E medium

(Biomérieux; Durham, North Carolina; catalog #50430) and API 20 A (Biomérieux;

Durham, North Carolina; catalog #20300) following the manufacturer’s instructions. API

50 CH was completed following the protocol for 50 CHB except PBS was reduced prior to use and the strips were incubated anaerobically.

3.14. Butyrate production

3.14.1. Nitric acid wash: In a fume hood, Hungate tubes were filled at least halfway with 3 N nitric acid and capped. They were stored at room temperature for 12 hours in an upright position followed by 12 hours in an inverted position. The nitric acid was poured from the tubes the original glass container and stored at room temperature to be reused. Ultrapure water was used to rinse the empty tubes and caps which were then left to air dry. Once dry, the tubes were filled with 10 ml of TPGY, flushed with nitrogen gas, capped, and autoclaved at 121°C for 20 minutes.

3.14.2. Acidogenesis end point: Cultures for C. perfringens ATCC 13124, E. coli

ATCC 25922, C. sporogenes ATCC 11437, C. sporogenes ATCC 19404, C. sporogenes

ATCC 15579, and C. sporogenes ATCC 25779 were prepared by inoculating 0.1 ml of the cooked meat media stock culture into 10 ml of reduced TPGY broth and incubating at 37°C with shaking at 200 rpm for 12-18 hours. Using a Spectronic 20D+ spectrophotometer

(Spectronic Instruments) set to a wavelength of 600nm, the fresh cultures were each inoculated into three new tubes of reduced TPGY broth to an optical density (OD) of 0.010.

The tubes were incubated at 37°C with shaking at 200 rpm for 12 hours. The OD of each culture was read every 1 hour and the ph was read using a pH 6+ meter (Oakton

Instruments; Vernon Hills, IL) every 2 hours.

4 RESULTS

4.1. Presence of the BUK gene in fecal samples

The fecal samples collected from all five animals were analyzed for the presence of the Clostridial BUK gene. All DNA purified directly from fecal samples was of good quality as indicated by a band at 1465 bp (Figure 4.1) after PCR amplification with the universal 16S rRNA primers. The BUK gene was amplified as an indicator of the presence and relative amount of butyrate producing Clostridium organisms. Each reaction contains

200 ng of DNA and the 16S rRNA bands serve as an internal control. All fecal samples indicated the presence of the BUK gene with varying band intensities at 464 bp after PCR amplification (Figure 4.2). While all PCR reaction mixes contained 200 ng of DNA, all fecal samples had lower band intensities than the positive control C. sporogenes ATCC

11437 indicating fewer gene copies in the fecal samples. The intensities of the BUK gene on the images were measured by ImageJ and normalized to the 16S rRNA bands to allow sample-to-sample comparison. Comparing the relative amount of BUK gene copies among the animals revealed the sheep had significantly fewer than all other animals and the goats had significantly fewer than the horses and cattle (Figure 4.3; Table 4.1).

A Horse B Cattle C Sheep 1 2 3 4 5 + - 1 2 3 4 5 + - 1 2 3 4 5 + -

Figure 4.1. Agarose gel electrophoresis of the 16S rRNA gene in crude fecal samples from (A) horse, (B) cattle, (C) sheep, (D) pig, and (E) goat. The 16srRNA gene presents at 1465bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control.

D Pig E Goat 1 2 3 4 5 + - 1 2 3 4 5 + -

Figure 4.1. (continued)

A Horse B Cattle C Sheep 1 2 3 4 5 + - 1 2 3 4 5 + - 1 2 3 4 5 + -

D Pig E Goat 1 2 3 4 5 + - 1 2 3 4 5 + -

Figure 4.2. Agarose gel electrophoresis of the BUK gene in crude fecal samples from (A) horse, (B) cattle, (C) sheep, (D) pig, and (E) goat. The BUK gene presents at 464 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control.

Figure 4.3. Relative amount of buk in crude fecal samples with band intensities measured using ImageJ. (*) represents P < 0.05 and (†) represents P < 0.05 from all.

Table 4.1. Statistical significance for the relative amounts of buk in crude fecal samples using the Mann-Whitney test. Grey boxes indicate P < 0.05. Pig Horse Cattle Sheep Goat Pig Horse 0.2222 Cattle 0.4206 0.3095 Sheep 0.0079 0.0079 0.0079 Goat 0.2222 0.0317 0.0317 0.0079 Mean 0.3661 0.6291 0.4769 0.0610 0.2764 Standard 0.1685 0.2611 0.1718 0.0148 0.1476 deviation

4.2. Colony morphology of fecal samples enriched for Clostridium species

Each fecal sample was inoculated into reduced CMM to select for and enhance the growth of anaerobic organisms and heat shocked before incubating to reduce the number of non-spore forming organisms. The CMM enriched culture was streaked onto reduced

EYA to obtain isolated colonies while also providing preliminary differential results for

Clostridium species. The resulting EYA plates for the horses, cattle, sheep, pigs, and goats are shown in Figures 4.4, 4.5, 4.6, 4.7, and 4.8, respectively. Overall, the horse, sheep, and goat samples displayed the lowest amount of colony diversity with the sheep and goat

samples giving very similar results and the cattle and pig samples displayed the highest amount of diversity with the pig sample being the most diverse.

All five horse samples (Figure 4.4) showed one predominate proteolytic, lipase positive colony morphology as shown by the clearance of the yolk protein around the colonies and the iridescent sheen on the surface, respectively. All samples show colonies in large, dry, slightly raised but flat colonies with a small, raised center, resembling colonies of Clostridium species. H1, H2, and H5 show the lowest diversity of all enriched fecal samples with all three appearing to have only one type of presumptive clostridial colonies. H3 and H4 both had an additional colony morphology that was small, raised, smooth, and round. H3 had the most diversity with two additional morphologies where one was small, flat, translucent, smooth, and round while the other was medium, wet, raised, translucent, smooth, and round. No horse samples had any lecithinase positive organisms.

A H1 B H2 C H3

D H4 E H5

Figure 4.4. Growth from heat shocked and enriched horse fecal samples (A) H1, (B) H2, (C) H3, (D) H4, and (E) H5 on EYA.

All five cattle samples (Figure 4.5) had varying diversity with some shared colony morphologies. All samples shared a proteolytic, lipase positive, small, dry, raised, smooth, and irregularly shaped but mostly round colony morphology that was predominant in C1,

C2, C3, and C5; however, the lipase reaction was weak for C3. C2 and C5 had a second common colony morphology that was lecithinase positive, small, raised, white, smooth, and round. C3 also shared this colony type, but only had a few colonies on the plate. In addition to the lipase positive and lecithinase positive colonies, C2 and C3 had a small number of small, wet, raised, translucently yellow, smooth, and round colonies. C3 had one more colony morphology in small numbers that were medium, wet, raised, translucent, smooth, and irregularly shaped. Besides the lipase positive and lecithinase positive colonies, C5 had an equally prevalent number of colonies that were medium, wet, raised, translucently white, smooth, and round. C4 showed the most diversity of all five samples including the following colony morphologies: medium, flat periphery with a raised center, translucently yellow, wavy, and round; medium, wet, raised, light yellow, smooth, and irregularly shaped; lecithinase positive, medium, wrinkly, raised, light yellow, smooth, and round; and medium, flat, cream, smooth, and irregularly shaped. All colony morphologies for C4 were present in relatively equal numbers when compared to each other.

A C1 B C2 C C3

Figure 4.5. Growth from heat shocked and enriched cattle fecal samples (A) C1, (B) C2, (C) C3, (D) C4, and (E) C5 on EYA.

D C4 E C5

Figure 4.5 (continued)

All 5 sheep samples (Figure 4.6) showed one predominant lecithinase positive organism with a medium sized, raised, smooth, and round colony morphology. All samples also showed another prevalent colony morphology that was proteolytic, lipase positive, small, raised, dry, smooth, and irregularly shaped but mostly round. S4 was the only one to have more diversity. S4 had a colony morphology that was small, raised, wet, smooth, and round; one that was small, raised, translucent, smooth, and round; and another lecithinase positive organism that was slightly more yellow when compared to the predominate morphology.

A S1 B S2 C S3

Figure 4.6. Growth from heat shocked and enriched sheep fecal samples (A) S1, (B) S2, (C) S3, (D) S4, and (E) S5 on EYA.

D S4 E S5

Figure 4.6 (continued)

All five pig samples (Figure 4.7) had a high level of diversity with similar colony morphologies. All samples had predominantly lecithinase positive colonies although the morphology varied between samples: P1, P2, and P3 showed a medium, raised with a flat top, cream, wavy, and irregularly shaped morphology; P1, P2, and P5 showed another similar morphology, but it was raised to a point and light yellow; P5 also showed a medium, raised, white, smooth, and irregularly shaped but mostly round morphology; and P4 showed a medium, dry, flat, translucent, smooth, and irregularly shaped morphology. All samples had a less prevalent number of proteolytic, lipase positive, small, dry, raised, smooth, and irregularly shaped but mostly round colonies. Another organism that was similar was seen in P2 and P4, but it was more proteolytic, weakly lipase positive, and medium sized. P1 and P3 also had another similar organism that was flat instead of raised.

P2, P3, and P4 showed one more similar morphology, but it was not proteolytic, lipase negative, and white. P2 showed a small number of medium, wet, flat, translucent, smooth, and irregularly shaped colonies while P3 shared a similar morphology, but it was small and raised. A small amount of medium, raised, white, smooth, and round colonies were seen in

P2 and P5. P3 had two unique colony morphologies that were both small, raised, and cream colored while one was wavy and irregularly shaped and the other was smooth and round.

A P1 B P2 C P3

D P4 E P5

Figure 4.7. Growth from heat shocked and enriched pig fecal samples (A) P1, (B) P2, (C) P3, (D) P4, and (E) P5 on EYA.

All 5 goat samples (Figure 4.8) displayed two of the same colony morphologies.

One was lecithinase positive, medium, smooth, and mostly round while the other was proteolytic, lipase positive, small, dry, raised, smooth, and irregularly shaped but mostly round. G1 showed the most diversity with two more colony morphologies. Both morphologies were proteolytic, medium, dry, flat with a raised center, and irregularly shaped, but one was lipase positive and the other was negative. G2 was the only other sample to have more diversity with one other morphology being very small, dry, raised, smooth, and round.

A G1 B G2 C G3

D G4 E G5

Figure 4.8. Growth from heat shocked and enriched goat fecal samples (A) G1, (B) G2, (C) G3, (D) G4, and (E) G5 on EYA.

4.3. Staining, biochemical, and molecular characterization of bacterial isolates

At least five colonies were further purified on a fresh EYA plate for isolation and characterization. The total number of bacterial isolates and the presumptive Clostridium isolates from the fecal samples collected from all animal types are summarized in Table

4.2. A total of 168 bacterial isolates were obtained and 134 of those were presumptive

Clostridium after characterization by Gram staining, spore staining, and oxygen tolerance as well as lecithinase, lipase, proteolytic, and hemolytic activities (Tables 4.3, 4.4, 4.5, 4.6, and 4.7). Of the 134 presumptive Clostridium isolates, 15 were presumptive C. perfringens.

All 15 of these isolates did not form spores under our growth conditions, but they fit all other criteria and demonstrated a double-zone of hemolysis which is characteristic of the species. These isolates were also determined to be unsafe for the use of a probiotic due to the possibility of α-toxins indicated by lecithinase production on EYA and the outer zone

of alpha hemolysis on BAP as well as θ-toxins indicated by the inner zone of beta hemolysis on BAP.

Table 4.2. Total number of bacterial isolates and presumptive Clostridium isolates from horse, cattle, sheep, pig, and goat samples. Total number Number of presumptive Animal of isolates Clostridium isolates Horse 33 24 Cattle 35 28 Sheep 31 29 Pig 39 26 Goat 30 27 Total 168 134

Table 4.3. Staining, biochemical, and molecular characterization of isolates from horse samples.*

Isolate stain Gram shape Cell Arrangement Spores shape Spore location Spore Bulging Aerobic Lecithinase Lipase Proteolysis Catalase Hemolysis buk H1A + rod S/Ch + oval ST + - - + w - wβ + H1B + rod S/Ch + oval ST + - - + w - wβ + H1C + rod S/Ch + oval ST + - - + w - wβ + H1D + rod S/Ch + oval ST + - - + w - wβ + H1E + rod S/Ch + oval ST + - - + w - wβ + H1F + rod S/Ch + oval ST + - - + w - wβ + H2A + rod S/Ch + oval ST + - - + w - wβ + H2B + rod S/Ch + oval ST + - - + w - wβ + H2C + rod S/Ch + oval ST + - - + w - wβ + H2D + rod S/Ch + oval ST + - - + w - wβ + H2E + rod S/Ch + oval ST + - - + w - wβ + H2F + rod S/Ch + oval ST + - - + w - wβ + H3A + rod S - + - - w + wβ H3B + rod S/Ch + oval ST + - - + w - wβ + H3C + rod S/Ch + oval ST + - - + w - wβ + H3D + rod S - + - - - - wβ H3E + cocci Cl - + - - - - wβ H3F + rod S - + - - - + γ H3G + rod S/Ch + oval ST + - - + w - wβ + H4A + rod S/Ch + oval ST + - - + w - wβ + H4B + cocci Cl - + - - - - γ H4C + rod S/Ch + oval ST + - - + w - wβ + H4D + cocci Cl - + - - - - γ H4E + cocci Cl - + - - - - γ H4F + rod S/Ch + oval ST + - - + w - wβ + H4G + cocci Cl - + - - - - γ H4H + cocci Cl ------γ H5A + rod S/Ch + oval ST + - - + w - wβ + H5B + rod S/Ch + oval ST + - - + w - wβ + H5C + rod S/Ch + oval ST + - - + w - wβ + H5D + rod S/Ch + oval ST + - - + w - wβ + H5E + rod S/Ch + oval ST + - - + w - wβ + H5F + rod S/Ch + oval ST + - - + w - wβ + *Abbreviations in Tables 4.3, 4.4, 4.5, 4.6, and 4.7: (+) positive, (w) weak positive, (-) negative, (S) single, (Ch) chain, (Cl) cluster, (ST) subterminal, (T) terminal, € central, (dzβ) double zone of beta and alpha, (β) beta, (wβ) weak beta, (γ) gamma, and (v) variable weak beta and gamma. Grey boxes in staining and biochemical columns indicate properties that are not found in Clostridium and Grey boxes in the isolate column indicates isolates that are not presumptive Clostridium.

Table 4.4. Staining, biochemical, and molecular characterization of isolates from cattle samples.*

robic

Isolate stain Gram shape Cell Arrangement Spores shape Spore location Spore Bulging Ae Lecithinase Lipase Proteolysis Catalase Hemolysis buk C1A + rod S/Ch + oval ST + - - + w - wβ + C1B + rod S/Ch + oval ST + - - + w - wβ + C1C + rod S/Ch + oval ST + - - + w - wβ + C1D + rod S/Ch + oval ST + - - + w - wβ + C1E + rod S/Ch + oval ST + - - + w - wβ + C1F + rod S/Ch + oval ST + - - + w - wβ + C1G + rod S/Ch + oval ST + - - + w - wβ + C2A + rod S/Ch + oval ST + - - + w - wβ + C2B + rod S/Ch + oval ST + - - + w - wβ + C2C + rod S/Ch ------wβ C2D + rod S/Ch + oval ST + - - + w - wβ + C2E + rod S/Ch + oval ST + - - + w - wβ + C2F + rod S/Ch - + + - - + β C2G + rod S/Ch + oval ST + - - + w - wβ + C3A - rod S/Ch ------wβ C3B + rod S/Ch - + + - - + β C3C - rod S/Ch ------wβ C3D + rod S/Ch + oval ST + - - + w - wβ + C3E + rod S/Ch + oval ST + - - + w - wβ + C3F + rod S/Ch + oval ST + - - + w - wβ + C4A + rod S/Ch + oval ST + - - + w - wβ + C4B + rod S + oval ST + - - - + - wβ + C4C + rod S/Ch + oval ST + - - - - - wβ + C4D + rod S/Ch + oval ST + - - + w - wβ + C4E + rod S/Ch + oval ST + - - + w - wβ + C4F + rod S - - + - - - dzβ C4G + rod S/Ch + oval ST + - - + w - wβ + C4H + rod S + oval ST + - - - - - γ + C4I + rod S/Ch + oval ST + - - + w - wβ + C5A + rod S - - - - - + γ C5B + rod S/Ch + oval ST + - - + w - wβ + C5C + rod S/Ch + oval ST + - - + w - wβ + C5D + rod S/Ch + oval ST + - - + w - wβ + C5E + rod S/Ch + oval ST + - - + w - wβ + C5F + rod S/Ch - + + - - + β *Please refer to the footnote in Table 4.3 for abbreviations.

Table 4.5. Staining, biochemical, and molecular characterization of isolates from sheep samples.*

Isolate stain Gram shape Cell Arrangement Spores shape Spore location Spore Bulging Aerobic Lecithinase Lipase Proteolysis Catalase Hemolysis buk S1A + rod S/Ch + oval ST + - - + w w v + S1B + rod S/Ch + oval ST + - - + w w v + S1C + rod S/Ch + oval ST + - - + w w v + S1D + rod S/Ch + oval ST + - - + w w v + S1E + rod S/Ch + oval ST + - - + w w v + S1F + rod S/Ch + oval ST + - - + w w v + S1G + rod S - - + - - - dzβ S2A + rod S/Ch + oval ST + - - + w w v + S2B + rod S/Ch + oval ST + - - + w w v + S2C + rod S/Ch + oval ST + - - + w w v + S2D + rod S/Ch + oval ST + - - + w w v + S2E + rod S/Ch + oval ST + - - + w w v + S2F + rod S - - + - - - dzβ S3A + rod S/Ch + oval ST + - - + w w v + S3B + rod S/Ch + oval ST + - - + w w v + S3C + rod S/Ch + oval ST + - - + w w v + S3D + rod S/Ch + oval ST + - - + w w v + S3E + rod S/Ch + oval ST + - - + w w v + S3F + rod S - - + - - - dzβ S4A + rod Ch - - - - w - v S4B + rod S/Ch + oval T + - - + w - wβ + S4C + rod S/Ch + oval T + - - + w - wβ + S4D + rod S - - + - - - dzβ S4E + rod S - - + - - - dzβ S4F + rod S/Ch ------γ S5A + rod S/Ch + oval ST + - - + w - v + S5B + rod S/Ch + oval ST + - - + w - v + S5C + rod S/Ch + oval ST + - - + w w v + S5D + rod S/Ch + oval C - - - - w - wβ w S5E + rod S/Ch + oval ST + - - + w w v + S5F + rod S - - + - - - dzβ *Please refer to the footnote in Table 4.3 for abbreviations.

Table 4.6. Staining, biochemical, and molecular characterization of isolates from pig samples.*

atalase

Isolate stain Gram shape Cell Arrangement Spores shape Spore location Spore Bulging Aerobic Lecithinase Lipase Proteolysis C Hemolysis buk P1A + rod S/Ch + oval ST + - - + w - γ + P1B + rod S/Ch + oval ST + - - + w - γ + P1C + rod S ------γ P1D + rod S/Ch + oval ST + - - + w - wβ + P1E + rod S/Ch + oval ST + - - + - - wβ + P1F + rod S/Ch + oval C - - + - - - γ + P1G + rod S - - + - - - dzβ P2A + rod S/Ch + oval ST + - - - - - γ w P2B - rod S ------γ P2C + cocci Cl - + - - - - γ P2D + rod S/Ch + oval ST + - - + + - wβ + P2E + rod S/Ch + oval ST + - + - - - γ w P2F + rod S - - + - - - γ P3A + rod S - + - - - + γ P3B + rod S/Ch + oval ST + - - - - - γ w P3C + rod S - + - - - + γ P3D + rod S/Ch + oval ST + - - - - - γ w P3E + rod S - + - - - + γ P3F + cocci Cl - + - - - - γ P3G + rod S/Ch + oval ST + - - + w - wβ + P3H + rod S/Ch + oval T + - - + w - wβ + P3I + rod S/Ch - + - - - + γ P4A + rod S/Ch + oval ST + - - - - - γ w P4B + rod S/Ch + oval ST + - - - - - γ w P4C + rod S/Ch + oval ST + - - + w - wβ + P4D + rod S/Ch + oval ST + - - + + - wβ + P4E - rod S/Ch ------γ P4F + rod S/Ch + oval T ------γ + P4G + rod S - - + - - - dzβ P4H + rod S/Ch + oval ST + - - + + - wβ + P5A - rod S ------γ P5B + rod S/Ch ------γ P5C + rod S/Ch + oval ST + - - + w - wβ + P5D + cocci Cl - + - - - - γ P5E + rod S/Ch + oval ST + - - + w - wβ + P5F + rod S/Ch + oval ST + - - + w - wβ + P5G + rod S/Ch + oval ST + - - + w - wβ + P5H + rod S + oval ST + - + - - - γ w P5I + rod S - - + - - - dzβ

Table 4.7. Staining, biochemical, and molecular characterization of isolates from goat samples.*

Isolate stain Gram shapeCell Arrangement Spores shapeSpore location Spore Bulging Aerobic Lecithinase Lipase Proteolysis Catalase Hemolysis buk G1A + rod S/Ch - - - - + - β G1B + rod S/Ch + oval ST + - - + w - v + G1C + rod S/Ch - - - - w - wβ G1D + rod S/Ch + oval ST + - - + w - v + G1E + rod S/Ch + oval ST + - - + w - v + G1F + rod S - - + - - - dzβ G2A + rod S/Ch + oval ST + - - + w - v + G2B + rod S/Ch + oval ST + - - + w - v + G2C + rod S/Ch + oval ST + - - + w - v + G2D + rod S/Ch + oval ST + - - + w - v + G2E + rod S ------γ G2F + rod S - - + - - - dzβ G3A + rod S/Ch + oval ST + - - + w - v + G3B + rod S/Ch + oval ST + - - + w - v + G3C + rod S/Ch + oval ST + - - + w - v + G3D + rod S/Ch + oval ST + - - + w - v + G3E + rod S/Ch + oval ST + - - + w - v + G3F + rod S - - + - - - dzβ G4A + rod S/Ch + oval ST + - - + w - v + G4B + rod S/Ch + oval ST + - - + w - v + G4C + rod S/Ch + oval ST + - - + w - v + G4D + rod S/Ch + oval ST + - - + w - v + G4E + rod S/Ch + oval ST + - - + w - v + G4F + rod S - - + - - - dzβ G5A + rod S/Ch + oval ST + - - + w - v + G5B + rod S/Ch + oval ST + - - + w - v + G5C + rod S/Ch + oval ST + - - + w - v + G5D + rod S/Ch + oval ST + - - + w - v + G5E + rod S/Ch + oval ST + - - + w - v + G5F + rod S - - + - - - dzβ *Please refer to the footnote in Table 4.3 for abbreviations.

4.4. Presence of the BUK gene in presumptive Clostridium isolates

Genomic DNA was purified from all presumptive Clostridium isolates and was analyzed for the presence of the BUK gene by PCR using degenerated BUK primers. All

DNA was of good quality as indicated by a band at 1465 bp (Appendix B) after PCR

amplification. The BUK gene was amplified as an indicator of butyrate producing

Clostridium and gene similarity. All presumptive Clostridium isolates indicated the presence of the BUK gene with a band at 464 bp after PCR amplification (Figure 4.9, 4.10,

4.11, 4.12, and 4.13). All presumptive Clostridium horse, cattle, and goat as well as most pig and sheep isolates displayed bands of the same intensity as the control C. sporogenes

ATCC 11437. Presumptive Clostridium isolates S5D, P2A, P2E, P3B, P3D, P4A, P4B, and P5H had lower band intensities and therefore lower sequence similarity than the control. A second faint band was seen for P3D, P4A, and P4D at about 2500 bp.

A H1 B H2 C H3 A B C D E F + - A B C D E F + - B C G + -

D H4 E H5 A C F + - A B C D E F + -

Figure 4.9. Agarose gel electrophoresis of the BUK gene in presumptive Clostridium isolates from horse samples (A) H1, (B) H2, (C) H3, (D) H4, and (E) H5. The BUK gene presents at 464 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control.

A C1 B C2 C C3 A B C D E F G + - A B D E G + - D E F + -

D C4 E C5 A B C D E G H I + - B C D E + -

Figure 4.10. Agarose gel electrophoresis of the BUK gene in presumptive Clostridium isolates from cattle samples (A) C1, (B) C2, (C) C3, (D) C4, and (E) C5. The BUK gene presents at 464 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control.

A S1 B S2 A B C D E F + - A B C D E + -

Figure 4.11. Agarose gel electrophoresis of the BUK gene in presumptive Clostridium isolates from sheep samples (A) S1, (B) S2, (C) S3, (D) S4, and (E) S5. The BUK gene presents at 464 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control.

C S3 D S4 E S5 A B C D E + - B C + - A B C D E + -

Figure 4.11. (continued)

A P1 B P2 C P3 A B D E F + - A D E + - B D G H + -

D P4 E P5 A B C D F H + - C E F G H + -

Figure 4.12. Agarose gel electrophoresis of the BUK gene in presumptive Clostridium isolates from pig samples (A) P1, (B) P2, (C) P3, (D) P4, and (E) P5. The BUK gene presents at 464 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control.

A G1 B G2 C G3 B D E + - A B C D + - A B C D E + -

D G4 E G5 A B C D E + - A B C D E + -

Figure 4.13. Agarose gel electrophoresis of the BUK gene in presumptive Clostridium isolates from goat samples (A) G1, (B) G2, (C) G3, (D) G4, and (E) G5. The BUK gene presents at 464 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control.

4.5. Identification of presumptive Clostridium isolates by 16S RNA sequence analysis

The 16S rRNA gene was sequenced for all presumptive Clostridium isolates and potentially safe to be used as probiotic based on staining, biochemical, and molecular results (Tables 4.3, 4.4, 4.5, 4.6, and 4.7). A total of 6 species were identified for the 119 sequenced isolates including 2 strains of Clostridium sporogenes, Clostridium tepidum,

Clostridium senegalense, Clostridium subterminale, Paeniclostridium sordellii, and

Paraclostridium benzoelyticum (Table 4.8). The 2 distinct strains of C. sporogenes differed by 2 base pairs where one exhibited 5’-TATG-3’ and the other was 5’-CATA-3’ at the same location. Of all the sequenced isolates, there were 47 C. sporogenes (TATG), 56 C. sporogenes (CATA), 5 C. tepidum, 1 C. senegalense, 4 C. subterminale, 4 P. sordellii, and

2 P. benzoelyticum. Although C4C was sequenced to be C. sporogenes (CATA), this was due to contamination and the original organism was unable to be isolated again.

C. sporogenes was the most prevalent species in all animal types with some showing a bias towards one strain over the other. The sequenced horse and goat isolates were only C. sporogenes where all 24 horse isolates were C. sporogenes (TATG) and all

22 goat isolates were C. sporogenes (CATA). The cattle and sheep isolates were mostly C. sporogenes where 24 cattle and 5 sheep isolates were determined to be C. sporogenes

(CATA) and 2 cattle and 15 sheep isolates were C. sporogenes (TATG). The remaining 2 cattle isolates were identified as C. tepidum and C. senegalense with C4H being the only

C. senegalense isolate identified. Of the remaining sheep isolates, 2 were C. tepidum and

1 was P. benzoelyticum. The pig isolates showed the most diversity with 6 C. sporogenes

(TATG), 5 C. sporogenes (CATA), 2 C. tepidum, 4 C. subterminale, 4 P. sordellii, and 1

P. benzoelyticum isolate.

Table 4.8. Species determination of presumptive Clostridium isolates based on 16S rRNA sequencing. Separate strains of C. sporogenes are indicated by their differing base pairs.

nse

sporogenes sporogenes tepidum senegale subterminal sordellii benzoelyticum

...... Sample C (TATG) C (CATA) C. C C P P H1 ABCDEF** H2 ABCDEF H3 BCG H4 ACF H5 ABCDEF C1 ABCDEFG C2 ABCDEG C3 DEF C4 EG ACDI B H C5 BCDE S1 ABCDEF S2 ABCDE S3 ABCDE S4 BC S5 ABCE D P1 ABDE F P2 D A E P3 GH D B P4 C D AB F P5 G CEF H G1 BDE G2 ABCD G3 ABCDE G4 ABCDE G5 ABCDE Total no. 47 56 5 1 4 4 2 of isolates **Each letter represents one isolate. For example: “ABCDEF” in sample row H1 represents isolates H1A, H1B, H1C, H1D, H1E, and H1F.

4.6. Phylogenetic relationships of sequenced isolates and related Clostridia species

A phylogenetic tree was generated to compare the relationships of the sequenced isolates to their type strain and other Clostridium species using B. subtilis NCIB 3610 as

an outgroup (Figure 4.14). Due to the high number of isolates with the same sequence, species were grouped together using the following representative sequences: H1A for C. sporogenes (TATG), C1A for C. sporogenes (CATA), C4B for C. tepidum, C4H for C. senegalense, P2A for C. subterminale, P1F for P. sordellii, and S5D for P. benzoelyticum.

Other Clostridia species in the tree include and Paraclostridium bifermentans; common toxin producers Clostridium perfringens ATCC 13124, Clostridium tetani NCTC 279, and

Clostridioides difficile DSM 1296; and high acid producers Clostridium butyricum DSM

10702, Clostridium beijerinckii DSM 791, Clostridium saccharobutylicum DSM 13864, and Clostridium acetobutylicum ATCC 824.

All isolated species and other included Clostridia were from clostridial clusters I and XI. In cluster I , C. sporogenes and C. tepidum are closely related to each other and both form a clade with the toxin producer C. tetani; C. subterminale and C. senegalense are related to each other and both form a clade with the acid producer C. acetobutylicum; and the acid producers C. butyricum, C. beijerinckii, and C. saccharobutylicum form a clade with the toxin producer C. perfringens. In cluster XI, P. benzoelyticum is very closely related to P. bifermentans while P. sordellii is related to the toxin producer C. difficile. All isolates except for C. subterminale were closely related to their type strain with a 16S rRNA sequence match of 100% for C. tepidum and P. benzoelyticum; at least 99% for C. sporogenes, C. senegalense, and P. sordellii; and 98.12% for C. subterminale. When comparing the 2 isolated strains of C. sporogenes, strain CATA was slightly more closely related to the type strain DSM 795 than strain TATG.

Cluster I I Cluster Cluster XI XI Cluster

Figure 4.14. Phylogenetic tree based on 16S rRNA gene sequences of Clostridia isolates and related Clostridia species with B. subtilis IAM 12118 used as an outgroup. Created by PhyML and aLRT, rendered by TreeDyn, and hosted by LIRMM.

4.7. Antibiotic susceptibility of sequenced isolates

A limited number of C. sporogenes isolates (Table 4.9) and all other sequenced isolates including C. tepidum, C. senegalense, C. subterminale, P. sordellii, and P. benzoelyticum were tested for susceptibility to selected antibiotics at the cut-off values outlined by the EFSA (Table 3.11). Antibiotic susceptibility was tested for a total of 45 isolates (Table 4.10 and Figure 4.15). No isolates were susceptible to all 9 antibiotics and none were susceptible to gentamycin, streptomycin, and kanamycin as anaerobes are intrinsically resistant to aminoglycosides. Because of this natural resistance, the aminoglycosides will not be considered in the following analysis. The number of isolates susceptible to other antibiotics were as follows: 35 to tetracycline, 17 to erythromycin, 11 to vancomycin, 6 to ampicillin, 6 to chloramphenicol, and 5 to clindamycin. The quality

control strain C. perfringens ATCC 13124 should have been completely inhibited by ampicillin, but weak growth indicates that the concentration may have been lower than the expected 1 mg/L (Figure 4.15C). Taking this into account, the 6 isolates with weak growth to ampicillin may be considered susceptible.

Table 4.9. Clostridium sporogenes isolates limited to one representative isolate for antibiotic susceptibility testing based on animal type and sample number as well as shared morphological, biochemical, and 16S rRNA sequencing results. C. sporogenes Representative Sample isolates isolate H1 ABCDEF** A H2 ABCDEF A H3 BCG B H4 ACF A H5 ABCDEF A C1 ABCDEFG A C2 ABCDEG A C3 DEF D C4 ADI A C4 EG E C5 BCDE B S1 ABCDEF A S2 ABCDE A S3 ABCDE A S5 AB A S5 CE C P1 AB A P1 D D P1 E E P3 G G P3 H H P4 C C P5 CEF C P5 G G G1 BDE B G2 ABCD A G3 ABCDE A G4 ABCDE A G5 ABCDE A **Please refer to the footnote in Table 4.7 for abbreviations

Table 4.10. Antibiotic susceptibility of sequenced isolates based on cut-off values required by the EFSA

(Table 3.11).***

illin

tamicin

Isolate Species (1) control Aerobic (2) control Aerobic (1) control (+) (2) control (+) Ampic Vancomycin Gen Kanamycin Streptomycin Erythromycin Clindamycin Tetracycline Chloramphenicol (-) ------BF B. fragilis - - + + + + + + + + + - + CP C. perfringens - - + + w - + + + + - - + C. sporogenes H1A (TATG) - - + + + + + + + + + - + C. sporogenes H2A (TATG) - - + + + + + + + w + - + C. sporogenes H3B (TATG) - - + + + + + + + + + - + C. sporogenes H4A (TATG) - - + + + + + + + + + - + C. sporogenes H5A (TATG) - - + + + + + + + w + - + C. sporogenes C1A (CATA) - - + + w + + + + + + - + C. sporogenes C2A (CATA) - - + + + + + + + - + - + C. sporogenes C3D (CATA) - - + + w + + + + + + - + C. sporogenes C4A (CATA) - - + + + + + + + + + - + C4B C. tepidum - - + + + + + + + - - - - C. sporogenes C4E (TATG) - - + + + + + + + + + + + C4H C. senegalense - - + + - - + + + - - - + C. sporogenes C5B (CATA) - - + + + + + + + + + - - C. sporogenes S1A (TATG) - - + + + + + + + + + - + C. sporogenes S2A (TATG) - - + + + + + + + + + - + C. sporogenes S3A (CATA) - - + + + + + + + - + - + S4B C. tepidum - - + + + + + + + + w - - S4C C. tepidum - - + + + + + + + w + - + C. sporogenes S5A (TATG) - - + + + + + + + + w - - C. sporogenes S5C (TATG) - - + + + + + + + + + - + S5D P. benzoelyticum - - + + w - + + + - w + +

Table 4.10.(continued)

racycline

Isolate Species (1) control Aerobic (2) control Aerobic (1) control (+) (2) control (+) Ampicillin Vancomycin Gentamicin Kanamycin Streptomycin Erythromycin Clindamycin Tet Chloramphenicol C. sporogenes P1A (TATG) - - + + + + + + + w + - + C. sporogenes P1D (TATG) - - + + w + + + + w + - + C. sporogenes P1E (TATG) - - + + + + + + + - w - + P1F P. sordellii - - + + - - + + + - + - + P2A C. subterminale - - + + + - + + + + + + + P2D C. tepidum - - + + - + + + + - - - - P2E P. sordellii - - + + - - + + + - + - + P3B P. sordellii - - + + w - + + + - + + + P3D C. subterminale - - + + w - + + + + + - + C. sporogenes P3G (CATA) - - + + + + + + + - + - + C. sporogenes P3H (CATA) - - + + + + + + + + + - + P4A C. subterminale - - + + + - + + + + + + + P4B C. subterminale - - + + + - + + + + + + w C. sporogenes P4C (TATG) - - + + + + + + + w + + + P4D C. tepidum - - + + - + + + + - - - - P4F P. sordellii - - + + - - + + + - + + + C. sporogenes P5C (CATA) - - + + + + + + + w + - + C. sporogenes P5G (TATG) - - + + + + + + + + + - + P5H P. benzoelyticum - - + + + - + + + - - + + C. sporogenes G1B (CATA) - - + + + w + + + - + - + C. sporogenes G2A (CATA) - - + + + w + + + - + - w C. sporogenes G3A (CATA) - - + + + + + + + w + + w C. sporogenes G4A (CATA) - - + + + + + + + + + - + C. sporogenes G5A (CATA) - - + + + + + + + - + - + ***Sterile reduced Brucella broth was used as a negative control. B. fragilis ATCC 25285 (BF) and C. perfringens ATCC 13124 (CP) were used as quality control strains. Grey boxes highlight negative growth results.

A Aerobic control (1) B (+) control (1) C Ampicillin

Figure 4.15. Representative modified Brucella blood agar plates for antibiotic susceptibility including (A) the aerobic control and (B) positive control plates inoculated before the antibiotic plates and (C) an inoculated ampicillin plate with weak growth for the quality control strain C. perfringens ATCC 13124 boxed.

Antibiotic susceptibility varied by animal with the cattle and pig isolates exhibiting the highest rate of susceptibility followed by the sheep, goat, and then horse isolates (Figure

4.16). Only the pig and cattle isolates showed variable degrees of susceptibility to all antibiotics analyzed and isolates from all animals were susceptible to tetracycline. The pig isolates displayed the highest rate of susceptibility to ampicillin, vancomycin, and erythromycin, but had the lowest rate of susceptibility to tetracycline. The cattle isolates showed the highest susceptibility to erythromycin and clindamycin and tied for chloramphenicol with the sheep. The sheep isolates also shared the same amount of susceptibility with the cattle for tetracycline and vancomycin. The sheep, goat, and horse isolates were all resistant to ampicillin and clindamycin. The goat isolates were also resistant to vancomycin and chloramphenicol but had the highest rate of susceptibility to erythromycin. The horse isolates had the highest susceptibility to tetracycline but were resistant to all other antibiotics.

Figure 4.16. Percentage of sequenced isolates by animal susceptible to antibiotics based on cut-off values required by the EFSA (Table 3.11). Gentamycin, streptomycin, and kanamycin are not included due to intrinsic resistance.

Antibiotic susceptibility varied by species although no species were susceptible to all antibiotics (Figure 4.17). While C4H was the only C. senegalense isolate, it displayed the highest amount of susceptibility as it was only resistant to chloramphenicol. P. sordellii,

P. benzoelyticum, and C. subterminale isolates were also resistant to chloramphenicol.

Isolates from these 4 species were the most susceptible to vancomycin and, except C. subterminale, to erythromycin. C. subterminale isolates were the least susceptible overall and displayed the lowest rate of susceptibility to tetracycline. C. tepidum isolates were only resistant to vancomycin and displayed the highest rate of susceptibility to tetracycline along with C. senegalense and to chloramphenicol. Both strains of C. sporogenes isolates were also resistant to vancomycin as well as ampicillin and clindamycin. C. sporogenes (CATA) isolates were more susceptible to erythromycin, tetracycline, and chloramphenicol when compared to C. sporogenes (TATG).

Figure 4.17. Percentage of sequenced isolates by species susceptible to antibiotics based on cut-off values required by the EFSA (Table 3.11). Gentamycin, streptomycin, and kanamycin are not included due to intrinsic resistance.

Among individual isolates, C. senegalense isolate C4H and C. tepidum isolates P2D and P4D exhibited the highest number of antibiotic susceptibility as they were inhibited by

5 of the antibiotics tested. C. tepidum isolate C4B and P. sordellii isolates P1F and P2E followed as they were inhibited by 4 antibiotics. These six isolates were considered potentially safe to be used as a probiotic under the condition that their resistance can be proven to be intrinsic. All other isolates were considered unsafe to be used as a probiotic due to antibiotic resistance.

4.8. Biochemical characterization by API of potential probiotic isolates

Potential probiotic isolates C4B, C4H, P1F, P2D, P2E, and P4D were further biochemically characterized by the API 20 A and 50 CH microbial identification kits

(Tables 4.11 and 4.12). These tests help to reassure the species identity and to reveal the

metabolic diversity of these isolates for potential effects in the gut. Some results differed between the kits as all strong acidification results seen in the API 20 A kit were corroborated by the API 50 CH kit, but the weak results were not. This may be due to the different mediums used for each kit and because the API 50 CHB media is not intended for the use with anaerobes.

All 6 isolates tested positive for the acidification of D-glucose and D-maltose in both kits. Biochemical results for C. tepidum isolates C4B, P2D, and P4D were very similar as all 3 were positive for gelatinase and weakly acidified D-mannose and D-sorbitol; however, only C4B was able to form acid with D-trehalose. All C. tepidum isolates were the only ones to ferment D-sorbitol and only C4B showed acidification of N- acetylglucosamine. C. senegalense isolate C4H was the only isolate to test negative for gelatinase and positive for the acidification of D-lactose and D-saccharose. P. sordellii isolates P1F and P2E shared the same results. Both were able to produce indole, urease, and gelatinase and were the only isolates to ferment D-fructose and L-fuccose.

Table 4.11. Biochemical characterization of potential probiotic Clostridia isolates by the API 20 A microbial identification kit Test C4B C4H P1F P2D P2E P4D Indole formation - - + - + - Urease - - + - + - Gelatin hydrolysis + - + + + + Esculin hydrolysis ------Acidification: D-glucose + + + + + + D-mannitol ------D-lactose - + - - - - D-saccharose - + - - - - D-maltose + + + + + + Salicin ------D-xylose ------L-arabinose ------Glycerol ------D-cellobiose ------D-mannose w w - w - w D-melezitose ------D-raffinose ------D-sorbitol w - - w - w L-rhamnose ------D-trehalose + - - - - -

Table 4.12. Biochemical characterization of potential probiotic Clostridia isolates by the API 50 CH microbial identification kit and API 50 CHB medium.**** Test C4B C4H P1F P2D P2E P4D (-) control ------Glycerol ------Erythritol ------D-arabinose ------L-arabinose ------D-ribose ------D-xylose ------L-xylose ------Adonitol ------Methyl-βD------xylopyranoside D-galactose ------D-glucose + + + + + + D-fructose - - + - + - D-mannose ------L-sorbose ------L-rhamnose ------Dulcitol ------

Table 4.12. (continued) Test C4B C4H P1F P2D P2E P4D Inositol ------D-mannitol ------D-sorbitol ------Methyl-αD------mannopyranoside Methyl-αD------glucopyranoside N-acetylglucosamine + - - - - - Amygdalin ------Arbutin ------Esculin ------Salicin ------D-cellobiose ------D-maltose + + + + + + D-lactose - + - - - - D-melibiose ------D-saccharose - + - - - - D-trehalose + - - - - - Inulin ------D-melezitose ------D-raffinose ------Amidon ------Glycogen ------Xylitol ------Gentiobiose ------D-Turanose ------D-Lyxose ------D-tagatose ------D-fuccose ------L-fuccose - - + - + - D-arabitol ------L-arabitol ------Potassium gluconate ------Potassium ------2-ketogluconate Potassium ------5-ketogluconate ****Grey boxes indicate tests in common with the API 20 A kit.

Although the apiweb software was not used to identify the isolates, the results do agree with the 16S rRNA sequence results (Figure 4.18). C. senegalense isolate C4H had a 93.2% match and good identification to C. perfringens, but C. senegalense is not yet in

the database. C. tepidum is also not in the database but is closely related to C. sporogenes which is supported by a 98.9% match for C4B and a 77.6% match for P1F and P2D to C. botulinum/C. sporogenes. P. sordellii isolate P1F and P2E were in the database with a

99.9% match to P. sordellii.

Figure 4.18. Analytical profile index of (A) C. senegalense isolate C4H, (B) C. tepidum isolate C4B, (C) C. tepidum isolates P2D and P4D, and (D) P. sordellii isolates P1F and P2E using the API 20 A microbial identification kit and apiweb software.

Figure 4.18. (continued)

4.9. Endpoint determination of acidogenesis in Clostridium sporogenes

To determine the peak of butyrate production, the OD and pH for 4 strains of C. sporogenes were monitored during the growth in TPGY medium (Appendix C and Figure

4.19). The endpoint of acidogenesis should correlate to the beginning of stationary phase and a pH of 4.5-5.0. The positive control C. perfringens was the only organism to reach a pH of about 5.00 which then started to increase; however, this occurred at 10 h which did not correlate to the beginning of stationary phase at 7 h. C. sporogenes ATCC 11437 and

ATCC 25779 were the only strains to lower and then raise in pH although neither strain reached a pH close to 4.5-5.0. C. sporogenes ATCC 11437 reached a pH of only 6.45 which did not correlate to the beginning of stationary phase and the pH for C. sporogenes ATCC

25779 remained constant until it increased at the beginning of stationary phase. The pH continued to drop for both C. sporogenes ATCC 15579 and ATCC 19404 passed the end of exponential phase and passed 12 h at the end of the time study. Out of the 4 C. sporogenes strains, ATCC 15579 reached the lowest pH of 5.58 and only ATCC 25779 showed an increase of pH at the beginning of stationary phase.

Further studies are needed to determine the endpoint of acidogenesis. Originally the concentration of glucose was too low and presented as a limiting factor, so it is possible that other critical nutrients are insufficient in the current media. It is also possible that

stationary phase is occurring later than what was measured since OD readings above 1.000 were taken and the cultures were not diluted.

A B

C D 76

Figure 4.19. Endpoint determination of acidogenesis in Clostridium sporogenes by OD at a wavelength of 600 nm and pH plotted over 12 hours where OD is represented by ( ) and pH is represented by ( ). (A) C. perfringens ATCC 13124 was used as a positive control and (B) E. coli ATCC 25922 was used as a negative control. C. sporogenes strains test included (C) C. sporogenes ATCC 11437, (D) C. sporogenes ATCC 15579, (E) C. sporogenes ATCC 19404, and (F) C. sporogenes ATCC 25779.

E F

Figure 4.19. (continued)

5 DISCUSSION

5.1. Overview

Clostridium species are abundant in soil environments and as commensal gut organisms in humans and animals. In this study, we isolated 134 Clostridia isolates (Table

4.2) from 5 different farm animals including horses, cattle, sheep, pigs, and goats (Tables

3.1, 3.2, 3.3, 3.4, and 3.5) for the potential use as a probiotic supplement. Among these isolates, 6 species were identified along with 2 distinct strains of C. sporogenes where 47 were C. sporogenes (TATG), 56 were C. sporogenes (CATA), 5 were C. tepidum, 1 was

C. senegalense, 4 were C. subterminale, 4 were P. sordellii, and 2 were P. benzoelyticum

(Table 4.8 and Figure 4.20B). These isolates were all confirmed to have the butyrate kinase gene (Figures 4.9, 4.10, 4.11, 4.12, and 4.13) and were characterized for their phenotypic and metabolic diversity (Tables 4.3, 4.4, 4.5, 4.6, and 4.7 and Figure 4.20A) as well as antibiotic susceptibility (Table 4.10 and Figure 4.20C). Based on these results, isolates

C4H, C4B, P1F, P2D, P2E, and P4D were selected for further biochemical characterization

(Tables 4.11 and 4.12). P. sordellii isolates P1F and P2E were later determined to be unsafe as a probiotic and the remaining C. senegalense isolate C4H and C. tepidum isolates C4B,

P2D, and P4D where considered potential probiotic candidates pending future studies into butyrate production, presence of virulence factors and toxins, and cause of antibiotic resistance.

A Characterization of isolates EYA: Staining: TPGY: BAP: PCR: lipase, Gram (+), aerobic hemolysis buk (+) lecithinase, spore (+) growth (-), (γ, α, v, wβ) proteolysis catalase (-) ➔ ➔ ➔ ➔

B Species identification C Antibiotic susceptibility 16S rRNA NCBI Wadsworth EFSA cut- sequencing BLAST and agar dilution off values EZBioCloud (no growth) ➔ ➔

Figure 5.1. Isolate screening procedure with any required results noted. (A) Phenotypic and metabolic characterization of 168 initial isolates resulting in 134 presumptive Clostridium isolates. (B) 16S rRNA sequencing of 134 presumptive Clostridium isolates resulting in 6 species of Clostridia. (C) Antibiotic susceptibility of the identified Clostridia species resulting in 6 potential probiotic candidates.

5.2. Comparison of characteristics in isolated Clostridia species

C. sporogenes is a known commensal mutualist in animal and human gut microbiomes and plays a beneficial role in producing high levels of short chain fatty acids.

Moreover, it can stably colonize germ-free mice with no negative effects from mono- association (Guo et al 2019). While C. sporogenes has been isolated from infections, they are generally polymicrobial and the role of C. sporogenes as a pathogen has not been established. C. sporogenes was the most frequently isolated species with 2 distinct 16S rRNA gene sequences identified (Table 4.8). When compared to documented strains, only the sheep and goat isolates differ by displaying weak catalase and variable hemolysis results (Tables 4.5 and 4.7; Rainey et al 2015).

The newly published species and type strain C. tepidum IEH 97212 was isolated in

2013 from bloated bottles containing non-dairy protein shake drinks indicating its’ role as a food spoilage organism. If isolates show no toxicity, then this role should not affect this species’ potential as a probiotic. Although C. tepidum IEH 97212 is closely related to C. sporogenes and C. botulinum, it showed no evidence of BoNT or other toxin genes by PCR or genome sequencing. Of the tests performed in this study, all isolates shared similar results to the type strain except for small differences in sugar fermentation (Tables 4.11 and 4.12; Dobritsa et al 2017). When comparing isolates, the sheep strains differ from the cattle and pig strains by exhibiting weaker proteolysis and higher antibiotic resistance

(Tables 4.4, 4.5, 4.6, and 4.10).

C4H was the only C. senegalense isolate to be found sharing a 99.18% 16S rRNA gene sequence similarity to the type strain C. senegalense JC122. The type strain was isolated in 2010 from the stool of a healthy 16-year-old male living in Senegal thus showing that C. senegalense is already a commensal organism of the human gut microbiome. Of the tests performed in this study, both C4H and JC122 share the same characteristics except

C4H did not produce acid from N-acetylglucosamine which may be a result of strain variation or the less than optimal use of the API 50CH kit with anaerobes (Tables 4.4, 4.11, and 4.12; Mishra et al 2012). The only other published paper for C. senegalense reports a draft genome sequence of a newly isolated strain, AGRFS4, from a soil sample on dairy farm similar to the isolation of C4H from cattle at CPP’s farm (Gupta et al 2020).

C. subterminale isolates shared a16S rRNA gene sequence similarity of 98.1% with the type strain C. subterminale DSM 6970 and 100% with C. culturomicsence CL-6. C. subterminale is part of the healthy gut microbiome and is rarely associated with disease

(Trepani et al 2017). In a few cases, this species presented as an opportunistic pathogen causing bacteremia or septicemia after injury typically in immunocompromised patients

(Cook et al 2018, Maria et al 2016). Of the tests performed in this study, C. subterminale isolates only differ from documented strains by displaying γ-hemolysis and much higher rate of antibiotic resistance (Tables 4.6 and 4.10; Rainey et al 2015). C. culturomicsence

CL-6 is not validly published and was isolated from liquid stool of an obese Saudi Arabian patient from an unpublished study seeking toxigenic C. difficile. No conclusions can be drawn regarding the similarity of CL-6 to the isolates in this study (Dubourg et al 2018).

P. sordellii is commonly found in the gastrointestinal tract of humans and animals and rarely causes infections; although, mortality rates are extremely high even in less pathogenic strains. All P. sordellii strains carry several virulence factors and some encode for the potent toxins. Both potent toxins TcsL and TcsH are closely related to C. difficile toxins and are located on plasmids (Reddy et al 2013, Couchman et al 2015). When compared to documented P. sordellii strains, isolates share similar characteristics except spores do not swell the cells for P1F and P4F, lecithinase is not produced by P2E and P3B, and antibiotic resistance is higher in all isolates in our study (Tables 4.6, 4.10, 4.11, and

4.12; Rainey et al 2015). All P. sordellii isolates were determined to be unsafe for use as a probiotic due to the ubiquity of virulence factors and high risk of horizontal gene transfer of toxins.

Both P. benzoelyticum isolates S5D and P5H shared a 100% 16S rRNA gene sequence similarity to the type stain P. benzoelyticum JC272 and 99.8% to the closely related species P. bifermentans ATCC 628. P. benzoelyticum JC272 was isolated from marine soil samples in India in 2013 with minimal information relating to this study

(Jyothsna et al 2016). No conclusions can be made regarding the similarity of JC272 to

S5D and P5H. All results for S5D and P5H match those of P. bifermentans but each isolate individually varied slightly when comparing the results with this published species (Tables

4.5, 4.6, and 4.10; Rainey et al 2015).

5.3. Antibiotic resistance mechanisms in potential probiotic Clostridia

A growing trend of decreased susceptibility and increased resistance rates to antibiotics have been observed due to the indiscriminate and improper use of antibiotics encouraging bacteria to adapt to selective pressures (Imperial and Ibana 2016). Antibiotic resistance in probiotics are of concern because they share an environment with gastrointestinal bacteria and pose a risk of horizontal gene transfer to pathogens (Broaders et al 2013). Increasing safety concerns of transferable antibiotic resistance genes in probiotic organisms has led to well-documented profiles of probiotic strains of food and biological sources; however, antibiotic resistance profiles of probiotic strains from commercial dietary supplements are scarce although recent research has found resistance in these organisms as well (Wong et al 2015, Zheng et al 2017). Furthermore, it is important to discriminate between intrinsic and extrinsic resistance. Intrinsic resistance presents minimal potential for horizontal gene transfer. As seen in this study, all isolates demonstrated resistance to gentamicin, kanamycin, and streptomycin (Table 4.10) as anaerobes inherently lack the oxygen dependent mechanisms required for cellular uptake of aminoglycosides. These antibiotics also pose little risk of causing C. difficile associated colitis due to their lack of impact on other protective anaerobic gut flora (Khanafer et al

2018). Extrinsic resistance requires further confirmation on whether the acquired resistance

was through genomic mutation which is generally acceptable or through added genes which pose the greatest risk for horizontal gene transfer (EFSA 2012).

After antibiotic susceptibility testing, C. senegalense isolate C4H is potentially the safest for use as a probiotic as it was only resistant to chloramphenicol. The use of chloramphenicol in the United States is rare and is now limited to treating only a few life- threatening infections due to a risk of significant toxicity (Brook et al 2013). Resistance to chloramphenicol is also rare; although, resistance by chloramphenicol acetyl transferases encoded by cat genes can be found in Clostrdium. Chromosomal genes catQ and catP can be seen in C. perfringens, catB in C. butyricum, and catD in C. difficile. The plasmid pIP501 is known to carry catP in Clostridium and poses a risk due to its’ ability to replicate in a wide variety of Gram-positive bacteria (Schwarz et al 2004). C. tepidum isolates P2D and P4D may also be safe to use as a probiotic as they were only resistant to vancomycin.

Resistance to vancomycin is most often caused by the acquisition of transposable elements which encode for cell wall synthesizing enzymes that change the structure of cell wall amino acids preventing vancomycin from binding (Czepiel et al 2019). Resistance can also be caused by amino acid changes in peptidoglycan associated peptides, but this mechanism is not well understood (Peng et al 2017).

C. tepidum isolate C4B and P. sordellii isolates P1F and P2E may still be considered prospective probiotic candidates, but they pose more risk since they show resistance to 2 of the antibiotics tested. In addition to vancomycin, C4B displays resistance to ampicillin despite considering the lower concentration of ampicillin in the media. The production of β-lactamases are the most common mechanism of ampicillin resistance in anaerobes with some species of Clostridium found to produce one or more β-lactamases

(Brook et al 2013). P1F and P2E were resistant to clindamycin as well as chloramphenicol.

Clindamycin resistance is increasing in anaerobes and is typically associated with ribosomal methylase erm genes that determine the polymorphism of the clindamycin binding site (Banawas 2018). Furthermore, care should be taken when assessing resistance to ampicillin and clindamycin as both are known to cause C. difficile associated colitis

(Brook et al 2013, Peechakara et al 2020). Confirmation of the absence of these genes will help address the concern of any potential risks.

5.4. Relation of isolates, commensal gut Clostridium, and probiotic Clostridium

Firmicutes are the dominant organisms in the adult human gut microbiome consisting of more than 200 genera including Lactobacillus, Bacillus, and Clostridium

(Vrieze et al 2010). Clostridium clusters XIVa and IV constitute the predominate genera making up 10-40% of the total gut microbiota; although, clusters I, XI, and XVI are also common (Lopetuso et al 2013, Ohashi and Fujisawa 2018). In this study, C. sporogenes,

C. tepidum, C. senegalense, and C. subterminale from cluster I and P. sordellii and P. benzoelyticum from cluster XI were isolated (Table 4.7 and Figure 4.14). Species from cluster XI consist of important harmful bacteria including C. difficile, P. sordellii, and P. bifermentans with increased numbers associated with colitis and carcinogenesis (Ohashi and Fujisawa 2019). Cluster I, or the authentic Clostridium, are generally associated with the development of disease since this cluster contains the most toxin producing pathogens including C. perfringens (Yang et al 2019); however, this cluster also includes the most industry relevant species (Cruz-Morales et al 2019) including the type strain, C. butyricum, which is sold as a probiotic supplement (Pen and Tec Consulting 2012).

Currently, C. butyricum is the only species of Clostridium approved for use as a probiotic and is available in 2 different strains: C. butyricum MIYAIRI 588 from Miyarisan

Pharmaceutical in Japan (Pen and Tec Consulting 2012) or C. butyricum TO-A from TOA

Biopharma in Japan (BIO-THREE) and Advanced Orthomolecular Research in Canada

(Probiotic 3 2019). Phylogenetically, C. butyricum and the potential probiotic isolates C. senegalense and C. tepidum are in cluster I. Neither C. senegalense nor C. tepidum are closely related to C. butyricum, but C. tepidum is more closely related than C. senegalense to C. butyricum (Figure 4.14; Rainey et al 2015). Biochemically, the C. senegalense isolate is more similar to C. butyricum as both do not produce indole, lecithinase, and lipase; both are not proteolytic; and C. senegalense can ferment some similar sugars (Tables 4.3, 4.10, and 4.11; Rainey et al 2015). C. butyricum has been indicated as both a food spoilage organism and a gut commensal similar to C. tepidum and C. senegalense isolates respectively (Dobritsa et al 2017, Rainey et al 2015, Mishra et al 2012). Regarding antibiotic susceptibility, C. tepidum isolates are more similar to C. butyricum as only the

C. senegalense isolate was resistant to chloramphenicol (Table 4.9; Dobritsa et al 2017,

Rainey et al 2015).

6 FUTURE STUDIES

6.1. Further characterization of newly published species

C. tepidum, C. senegalense, and P. benzoelyticum isolates are newly defined species which were validly published in 2017, 2012, and 2016, respectively. As a result, minimal information regarding these species is available and further characterization would generate novel data. C. tepidum and C. senegalense isolates also require extensive biochemical and genetic research to validate their safe use as probiotics since they do not already have an established history of stability. Moreover, further information on spore production and viability is needed to determine the efficacy of C. tepidum and C. senegalense to produce viable spores that can be stored as a tablet and pass through the digestive system until they germinate in the gut.

6.2. Determining toxicity in potential probiotic isolates

While some Clostridium species are a vital part of the human microbiome, there are other pathogenic species which produce toxins and virulence factors which can be fatal.

Neither C. tepidum or C. senegalense have been profiled for any potential toxicity and should be screened for the most potent toxins including: C. perfringens toxins α, β, ε, and

ι; C. difficile toxins TcdA and TcdB; C. botulinum non-toxic non-hemagglutinin (NTNH) and BoNT A-G; and C. tetani toxin TeNT. Due to the lack of information for these two isolate species, it may be necessary to screen for the less common toxins and virulence factors typically found in the clinically relevant species C. chauvoei, C. novyi, C. septicum, and P. sordellii.

6.3. Determining the cause of antibiotic resistance in potential probiotic isolates

C. senegalense isolate C4H and C. tepidum isolates C4B, P2D, and P4D require further study to determine the nature of their antibiotic resistance. The minimal inhibitory concentration for chloramphenicol should be determined to access the rate of resistance as the CLSI defined breakpoint of ≤ 8 mg/L is higher than the EFSA defined cut-off value of

≤ 2 mg/L (EFSA 20120, Jousimies-Somer et al 2002). Due to the high risk of horizontal gene transfer, C4H should be screened for the presence of chromosomal and plasmid cat genes while C4B, P2D, and P4D should be screened for transposable elements encoding for genes affecting cell wall synthesis. C4B should be also screened for the presence of β- lactamases since it shows resistance to ampicillin.

6.4. Determining and optimizing the protocol for butyrate production analysis

It is imperative to determine the time point at which acids produced during the first phase of ABE fermentation become re-assimilated and converted into solvents to accurately measure and compare butyrate production between different species of

Clostridium. Standard batch fermentation methods indicate a clear metabolic shift from acidogenesis to solventogenesis linked with the beginning of stationary phase; however, some species and strains have shown to enter solventogenesis before stationary phase

(Millat and Winzer 2017). In future studies, the growth phase can still be determined using

OD, but measurements above 1.000 should be diluted for more accurate results. Media composition should also be taken into consideration as the initial concentration of glucose presented as a limiting factor of acid production.

REFERENCES

Aluwong, T., P. I. Kobo, and A. Abdullahi. Volatile fatty acids production in ruminants

and the role of monocarboxylate transporters: A review. African Journal of

Biotechnology 9:6229-6232.

Atarashi, K., T. Tanoue, K. Oshima, W. Suda, Y. Nagano, H. Nishikawa, S. Fukuda, T.

Saito, S. Narushima, K. Hase, S. Kim, S. Kim, J. V. Fritz, P. Wilmes, S. Ueha, K.

Matsushima, H. Ohno, B. Olle, S. Sakaguchi, T. Taniguchi, H. Morita, M. Hattori,

and K. Honda. 2013. Treg induction by a rationally selected mixture of Clostridia

strains from the human microbiota. Natuer 500:232-236.

Atarashi, K., T. Tanoue, T. Shimo, A. Imaoka, T. Kuwuhara, Y. Momose, G. Cheng, S.

Yamasaki, T. Saito, Y. Ohba, T. Taniguchi, K. Takeda, S. Hori, I. I. Ivanov, Y.

Umesaki, K. Itoh, and K. Honda. 2011. Induction of regulatory T cells by

indigenous Closridium species. Science 331:337-341.

Ayua, E. O., A. E. Kazem, B. R. Hamaker. 2020. Whole grain cereal fibers and their

support of the gut commensal Clostridia for health. Bioactive Carbohydrates and

Dietary Fibre 24:1-33.

Azad, A. K., M. Sarker, and D. Wan. 2018. Immunomodulatory effects of probiotics on

cytokine profiles. Biomedical Research International 2018:1-10.

Addison Wesley Longman INC. 1999. Ruminant digestion in Bos taurus

https://www.mun.ca/biology/88ngl/Ruminant_Digestion.html

Baker, S. D. 2017. Digestive systems. University of Idaho Extension.

Banawas, S. S. 2018. Clostridium difficile infections: a global overview of drug sensitivity

and resistance mechanisms. BioMed Research International 2018:1-9.

Bermudez-Brito, M., J. Plaza-Diaz, S. Munoz-Quezada, C. Gomez-Llorente, and A. Gil.

2012. Probiotic mechanisms of action. Annals of Nutrition and Metabolism 61:160-

174.

Biagi, E., and M. Candela. 2012. Ageing of the human metaorganism: the microbial

counterpart. Age 34:247-267.

BIO-THREE. TOA Biopharma Co. Ltd. https://www.toabio.co.jp/cms/toa/forlign/

item_index_e.html

Broaders, E., C. G. M. Gahan, and J. R. Marchesi. 2013. Mobile genetic elements of the

human gastrointestinal tract: potential spread of antibiotic resistance genes. Gut

Microbes 4:271-280.

Brook, I., H. M. Wexler, and E. J. C. Goldstein. 2013. Antianaerobic antimicrobials:

spectrum and susceptibility. Clinical Microbiology Reviews 26:526-546.

Buffie, C. G, V. Bucci, R. R. Stein, P. T. McKenney, L. Ling, A. Gobourne, B. No, H. Liu,

M. Kinnebrew, A. Viale, E. Littmann, M. R. M. van den Brink, R. R. Jenq, Y. Taur,

C. Sander, J. Cross, N. C. Toussaint, J. B. Xavier, and E. G. Pamer. 2015. Precision

microbiome restoration of bile acid-mediated resistance to Clostridium difficile.

Nature 517:205-208.

Cai, M., L. Zeng, L. H. Mo, R. D. Xie, B. S. Feng, P. Y. Zheng, Z. G. Liu, Z. J. Liu, and

P. C. Yang. 2016. Specific immunotherapy ameliorates ulcerative colitis. Allergy,

Asthma and Clinical Immunotherapy 12:1-7.

Canani, R. B., G. Terrin, P. Cirillo, G. Castaldo, F. Salvatore, G. Cardillo, A. Coruzzo, and

R. Troncone. 2004. Butyrate as an effective treatment of congenital chloride

diarrhea. Gastroenterology 127:630-634.

Canani, R. B., M. Di Costanzo, L. Leone, M. Pedata, R. Meli, and A. Calignano. 2011.

Potential beneficial effects of butyrate in intestinal and extraintestinal diseases.

World Journal of Gastroenterology 17:1519-1528.

Canani, R. B., G. Terrin, A. Elce, V. Pezzella, P. Heinz-Erian, A. Pedrolli, C. Centenari,

F. Amato, R. Tomaiuolo, A. Calignano, R. Troncone, and G. Castaldo. 2013.

Genotype-dependency of butyrate efficacy in children with congenital chloride

diarrhea. Orphanet Journal of Rare Diseases 8:194-202.

Cassir, N., S. Benamr, and B. La Scola. 2016. Clostridium butyricum: from beneficial to a

new emerging pathogen. Clinical Microbiology and Infection 22:37-45.

Chang, G., N. Ma, H. Zhang, Y. Wang, J. Hurang, J. Liu, H. Dai, and Z. Shen. 2019.

Sodium butyrate modulates mucosal inflammation injury mediated by GPR41-43

in the cecum of goats fed a high concentration diet. Frontiers in Physiology 10:1-

10.

Chen, G., X. Ran, B. Li, Y. Li, D. He, B. Huang, S. Fu, J. Liu, W. Wang. 2018. Sodium

butyrate inhibits inflammation and maintains epithelium barrier integrity in a

TNBS-induced inflammatory bowel disease mice model. EbioMedicine 30: 317-

325.

Chen, L., W. Xu, A. Lee, J. He, B. Huang, W. Zheng, T. Su, S. Lai, Y. Long, H. Chu, Y.

Chen, L. Wang, K. Wang, J. Si, and S. Chen. 2018. The impact of Helicobacter

pylori infection, eradication therapy and probiotic supplementation on gut

microenvironment homeostasis: an open-label, randomized clinical trial.

EbioMedicine 35:87-96.

Clostridium butyricum MIYAIRI strain. Miyarisan Co., Ltd. http://www.miyarisan.com/

91nglish_index.htm

Cobo, E. R, V. Kissoon-Singh, F. Moreau, R. Holani, and K. Chadee. 2017. MUC2 mucin

and butyrate contribute to the synthesis of the antimicrobial peptide cathelicidin in

response to Entamoeba histolytica- and dextran sodium sulfate-induced colitis.

Infection and Immunity 85:1-14.

Cook, P. A., A. Miishler, D. Quan, and A. Parrish-Garcia. 2018. Wound botulism caused

by Clostridium subterminale after heroin injection 10:42-44.

Couchman, E. C., H. P. Browne, M. Dunn, T. D. Lawly, J. G. Songer, V. Hall, L. Petrovska,

C. Vidor, M. Awad, D. Lyras, and N. F. Fairweather. 2015. Clostridium sordellii

genome analysis reveals plasmid localized toxin genes encoded within

pathogenicity loci. BMC Genomics 16:1-13.

Czepiel, J., M. Drozdz, H. Pituch, E. J. Kuijper, W. pericki, A. Mielimonka, S. Goldman,

D. Wultanska, A. Garlicki, G. Biesiada. 2019. Clostridium difficile infection:

review. European Journal of Clinical Microbiology and Infectious Diseases

38:1211-1221.

Dobritsa, A. P., K. K. Kutumbaka, K. Werner, M. Wiedmann, A. Asmus, and M.

Samadpour. 2017. Clostridium tepidum sp. nov., a close relative of Clostridium

sporogenes and Clostridium botulinum group I. International Journal of Systematic

and Evolutionary Microbiology 67:2317-2322.

Doron, S. and D. R. Snydman. 2015. Risk and safety of probiotics. Clinical Infectious

Diseases 60:S129-134.

Douek, D. C., B. Hill, T. Smith, and S. Arnon. 2008. Method for rapid detection of

Clostridium botulinum. World Intellectual Property Organization International

Bureau 1-41.

Dubourg, G., S. Baron, F. Cadoret, C. Couderc, P. E. Fournier, J. C. Lagier, and D. Raoult.

2018. From culturomics to clinical microbiology and forward. Emerging Infectious

Diseases 24:1683-1690.

EFSA. 2012. Guidance on the assessment of bacterial susceptibility to antimicrobials of

human and veterinary importance. EFSA Journal.

Fava, F., and S. Danese. 2011. Intestinal microbiota in inflammatory bowel disease: friend

or foe? World of Gastroenterology 17:557-566.

Feng, W., Y. Wu, G. Chen, S. Fu, B. Li, B. Huang, D. Wang, W. Wang, and J. Liu. 2018.

Sodium butyrate attenuates diarrhea in weaned piglets and promotes tight junction

protein expression in colon in a GPR109A-dependent manner. Cellular Physiology

and Biochemistry 47:1617-1629.

Fijan, S. 2014. Microorganisms with claimed probiotic properties: an overview of recent

literature. International Journal of Environmental Research and Public Health

11:4745-4767.

Firkins, J. L., and Z. Yu. 2015. Ruminant nutrition symposium: How to use data on the

rumen microbiome to improve our understanding of ruminant nutrition. American

Society of Animal Science 93:1450-1470.

Furusawa, Y., Y. Obata, S. Fukuda, T. A. Endo, G. Nakato, D. Takahashi, Y. Nakanishi,

C. Uetake, K. Kato, T. Kato. M. Takahashi, N. N. Fukuda, S. Murakami, E.

Miyaushi, S. Hino, K. Atarashi, S. Onawa, Y. Fujimura, T. Lockett, J. M. Clasrke,

D. L. Topping, M. Tomita, S. Hori, O. Ohara, T. Morita, H. Koseki, J. Kikushi, K.

Honda, K. Hase, and H. Ohno. 2013. Commensal microbe-derived butyrate induces

the differentiation of colonic regulatory T cells. Nature 504:446-450.

Gogineni, V. K., L. E. Morrow, and M. A. Malesker. 2013. Probiotics: mechanisms of

action and clinical applications. Probiotics and Health1:1-11.

Gorka, P. Z. M. Kowalski, R. Zabielski, and P. Guilloteau. 2018. Invited review: Use of

butyrate to promote gastrointestinal trac development in calves. Journal of Dairy

Science 101:4785-4800.

Guo, C. J., B. M. Allen, K.J. Hiam, D. Dodd, W. Van Treuren, S. Higginbottom, K.

Nagashima, C. R. Fischer, J. L. Sonnenburg, M. H. Spitzer, and M. A. Fischbach.

2019. Depletion of microbiome-derived molecules in the host using Clostridium

genetics. Science 366:1-15.

Guo, P., K. Zhang, X. Ma, and P. He. 2020. Clostridium species as probiotics: potentials

and challenges. Journal of Animal Science and Biotechnology 11:1-10.

Gupta,T. B., R. Jauregui, A. S. Nawarathana, g. Brightwell, and P. Maclean. 2020. Draft

genome sequence of Clostridium senegalense strain AGRSF4, isolated from a dairy

farm in New Zealand. American Society for Microbiology 9:1-2.

Hagihara, M., Y. Kuroki, T. Ariyoshi, S. Higashi, K. Fukuda, R. Yamashita, A.

Matsumoto, T. Mori, K. Mimura, N. Yamaguchi, S. Okada, T. Nonogaki, T.

Ogawa, K. Iwasaki, S. Tomono, N. Asai, Y. Koizumi, K. Oka, Y. Yamagishi, M.

Takahashi, and H. Mikamo. 2020. Clostridium butyricum modulates the

microbiome to protect intestinal barrier function in mice with antibiotic-induced

dysbiosis. iScience 23:1-18.

Hamer, H. M., D. M. Jonkers, A. Bast, S. A. Vanhoutvin, M. A. Fischer, A. Kodde, F. J.

Troost, K. Venema, and R. M. Brummer. 2008. Burtyrate modulates oxidative

stress in the colonic mucosa of healthy humans. Clinical Nutrition 28:88-93.

Hamer, H. M., D. Jonkers, K. Venema, S. Vanhoutvin, F. J. Troost, and R. J. Brummer.

2007. Review article: the role of butyrate on colonic function. Alimentary

Pharmacology and Therapeutics 27:104-119.

Hayashi, A., T. Sato, N. Kamada, Y. Mikami, K. Matsuoka, T. Hisamatsu, T. Hibi, A.

Roers, H. Yagita, T. Ohteki, A. Yoshimura, and T. Kanai. 2013. A single strain of

Clostridium butyricum induces intestinal IL-10-producing macrophages to

suppress acute experimental colitis in mice. Cell Host and Microbe 13:711-722.

Hui, W., D. Yu, Z. Cao, and X. Zhao. 2019. Butyrate inhibit collagen-induced arthritis via

Treg/IL-10/Th17 axis. International Immunopharmacology 68:226-233.

Imperial, I. C. V. J., and J. A. Ibana. 2016. Addressing the antibiotic resistance problem

with probiotics: reducing the risk of its double-edged sword effect. Frontiers in

Microbiology 7:1-10.

Islam, S. U. 2016. Clinical uses of probiotics. Medicine: Systematic Review and Meta-

Analysis 95:1-5.

Jha, R., J. M. Fouhse, U. P. Tiwari, L. Li, and B. P. Willing. 2019. Dietary fiber and

intestinal health of monogastric animals. Frontiers in Veterinary Science 6:1-12.

Jia, L., D. Li, N. Feng, M. Shamoon, Z. Sun, L. Ding, H. Zhang, J. Sun, and Y. Q. Chen.

2017. Anti-diabetic effects of Clostridium butyricum CGMCC0313.1 through

promoting the growth of gut butyrate-producing bacteria in type 2 diabetes.

Scientific Reports 7:1-15.

Jia, L., K. Shan, L. Pan, N. Feng, Z. Lv, Y. Sun, J. Li, C. Wu, H. Zhang, W. Chen, J. Diana,

J. sun, and Y. Q. Chen. 2017. Clostridium butyricum CGMCC0313.1 protects

against autoimmune diabetes by modulating intestinal immune homeostasis and

inducing pancreatic regulatory T cells. Frontiers in Microbiology 8:1-20.

Jung, T. H., J. H. Park, W. M. Jeon, and K. S. Han. 2015. Butyrate modulates bacterial

adherence on LS174T human colorectal cells by stimulating mucin secretion and

MAPK signaling pathway. Nutrition, Research, and Practice 9:343-349.

Jousimies-soomer, H. R., P. Summanen, D. M. Citron, E. J. Baron, H. M. Wexler, and S.

M. Finegold. 2002. Susceptibility testing of anaerobic bacteria. Anaerobic

Bacteriology Manual. 6th edition. (eds S. Hoffman). Star Publishing. Chapter 8.

Jyothsna, T. S. S., L. Tushar, C. Sasikala, and C. V. Ramana. 2016. Paraclostridium

benzoelyticum gen. nov., sp. nov., isolated from marine sediment and

reclassification of Clostridium bifermentans as Paraclostridium bifermentans

comb. nov. Proposal of a new genus Paeniclostridium gen. nov. to accommodate

Clostridium sordellii and Clostridium ghonii. International Journal of Systematic

and Evolutionary Microbiology 66:1286-1274.

Kabeerdoss, J., V. Sankaran, S. Pugazhendhi, and B. S. Ramakrishna. 2013.Clostridium

leptum group bacteria abundance and diversity in the fecal microbiota of patients

with inflammatory bowel disease: a case-control study in India. BMC

Gastroenterology 13:1-8.

Kaakoush, N. O., A. S. Day, K. D. Huinao, S. T. Leach, D. A. Lemberg, S. E. Dowd, and

H. M. Mitchell. 2012. Microbial dysbiosis in pediatric patients with Crohn’s

disease. Journal of Clinical Microbiology 50:3258-3266.

Khanafer, N., N. Daneman, T. Greene, A. Simor, P. Vanhems, and M. Samore. 2018.

Susceptibilities of clinical Clostridium difficile isolates to antimicrobials: a

systematic review and meta-analysis of studies since 1970. Clinical Microbiology

and Infection 24:110-117.

Kho, Z. Y., and S. K. Lal. 2018. The human gut microbiome – A potential controller of

wellness and disease. Frontiers in Microbiology 9:1-23.

Knudsen, K. E. B., H. N. Laerke, M. S. Hedemann, T. S. Nielsen, A. K. Ingerslev, D. S.

G. Nielson, P. K. Theil, S. Purup, S. Hald, A. G. Schioldan, M. L. Marco, S.

Gregersen, and K. Hermansen. 2018. Impact of diet-modulated butyrate production

on intestinal barrier function and inflammation. Nutrients 10:1499-1518.

Kobayashi, R., K. Nagaoka, N. Nishimura, S. Koike, E. Takahashi, K. Niimi, H. Murase,

T. Kinjo, T. Tsukahara, and R. Inoue. 2020. Comparison of the fecal microbiota of

two monogastric herbivorous and five omnivorous mammals. Animal Science

Journal 91:1-10.

Kumari, R., V. Ahuja, and J. Paul. 2013. Fluctuations in butyrate-producing bacteria in

ulcerative colitis patients of North India. World Journal of Gastroenterology

19:3404-3414.

Kunisawa, J. and H. Kiyono. 2011. Peaceful mutualism in the gut: revealing key

commensal bacteria for the creation and maintenance of immunological

homeostasis. Cell Host and Microbe 9:83-84.

Lan, B., F. Yang, D. Lu, and Z. Lin. 2016. Specific immunotherapy plus Clostridium

butyricum alleviates ulcerative colitis in patients with food allergy. Scientific

Reports 6:1-7.

Lane, D.J. 1991. 16S/23S rRNA sequencing. In: Nucleic acid techniques in bacterial

systematics. Stackebrandt, E., and Goodfellow, M., eds., John Wiley and Sons,

New York, NY, pp. 115-175.

Leputso, L. R., F. Sca;daferri, V. Petito, and A. Gasbarrini. 2013. Commensal Clostridia:

leading players in the maintenance of gut homeostasis. Gut Pathogens 5:1-8.

Li, R. W., S. Wu, R. L. Baldwin VI, W. Li, C. Li. 2012. Perturbation dynamics of the

rumen microbiota in response to exogenous butyrate. PloS ONE 7:1-11.

Li, W., C. Ding, T. Meng, W. Lu, W. Liu, H. Hao, and L. Cao. 2017. Butyrate suppresses

motility of colorectal cancer cells via deactivating Akt/ERK signaling in histone

deacetylase dependent manner. Journal of Pharmacological Sciences 135:148-155.

Li, Q., L. Cao. Y. Tian, P. Zhang, C. Ding, W. Lu, C. Jia, C. Sao, W. Liu, D. Wang, H. Ye,

and H. Hao. 2018. Butyrate suppresses the proliferation of colorectal cancer cells

via targeting pyruvate kinase M2 and metabolic reprogramming. Molecular and

Cellular Proteomics 17:1531-1545.

Lin, L., F. Xie, D. Sun, J. Liu, W. Zhu, and S. Mao. 2019. Ruminal microbiome-host

crosstalk stimulated the development of the ruminal epithelium in a lamb model.

Microbiome 7:1-16.

Liu, J., J. Sun, F. Wang, X. Yu., Z. Ling, H. Li, H. Zhang, J. Jin, W. Chen, M. Pang, J. Yu,

Y. He, and J. Xu. 2015. Neuroprotective effects of Clostridium butyricum against

vascular dementia in mice via metabolic butyrate. BioMed Research International

2015:1-12.

Liu, Y., B. Upadhyaya, A. R. Fardin-Kia, R. M. Juenemann, and M. Dey. 2016. Dietary

resistant starch type 4-derived butyrate attenuates nuclear factor-kappa-B1 through

modulation of lysine 27 trimethylation of histone H3. Food and Function 14:3772-

3781.

Liu, M., W. Xie, X. Wan, and T. Deng. 2020. Clostridium butyricum modulates gut

microbiota and reduces colitis associated colon cancer in mice. International

immunopharmacology 88:1-10.

Liu, M., W. Xie, X. Wan, and T. Deng. 2020. Clostridium butyricum protects intestinal

barrier function via upregulation of the tight junction proteins and activation of the

Akt/MTOR signaling pathway in a mouse model of dextran sodium sulfate-induced

colitis. Experimental and Therapeutic Medicine 20:1-10.

Lopetuso, L. R., F. Scaldaferri, V. Petito, and A. Gasbarrini. 2013. Commensal Clostridia:

leading players in the maintenance of gut homeostasis. Gut Pathogens 5:23-31.

Louis, P. and H. J. Flint. 2009. Diversity, metabolism, and microbial ecology of butyrate-

producing bacteria from the human large intestine. Federation of European

Microbiological Societies 294:1-8.

Luhrs, H., T. Gerke, F. Boxberger, K. Backhaus, R. Melcher, W. Scheppach, and T.

Menzel. 2001. Butyrate inhibits interleukin-1-mediated nuclear factor-kappa B

activation in human epithelial cells. Digestive Diseases and Sciences 46:1968-

1973.

Luu, M., U. Steinhoff, and A. Visekruna. 2017. Functional heterogeneity of gut resident

regulatory T cells. Clinical and Translational Immunology 6:1-8.

Ma, N., J. A. Abaker, M. S. Bilal, H. Dai, and X. Shen. 2018. Sodium butyrate improves

antioxidant stability in sub-acute ruminal acidosis in dairy goats. BMC Veterinary

Research 14:275-288.

Markowiak, P. and K. Slizewaka. 2017. Effects of probiotics, prebiotics, and synbiotics on

human health. Nutrients 9:1021-1051.

Matthews, J. B., I. Hassan, S. Meng, S. Y. Archer, B. J. Hrnjez, and R. A. Hodin. 1998.

Na-K-2Cl cotransporter gene expression and function during enterocyte

differentiation. Journal of Clinical Investigation 101:2072-2079.

Miguel, M. A., S. S. Lee, L. L. Mamuad, Y. J. Choi, C. D. Jeong, A. Son, K. K. Cho, E. T.

Kim, S. B. Kim, and S. S .Lee. 2019. Enhancing butyrate production, ruminal

fermentation and microbial population through supplementation with Clostridium

saccharobutylicum. The Korean Society for Microbiology and Biotechnology

29:1083-1095.

Moran, J. 2005. How the rumen works. In Tropical Dairy Farming: Feeding and

Management for Small Holder Dairy Farmers in the Humid Tropics. Landlinks

Press.

Nagano, Y., K. Itoh, and K. Honda. 2012. The induction of Treg cells by gut-indigenous

Clostridium. Current Opinion in Immunology 24:392-397.

Nava, G. M., H. J. Friedrichssen, and T. S. Stappenbeck. 2011. Spatial organization of

intestinal microbiota in the mouse ascending colon. The ISME Journal 5:627-638.

O’Donnell, M. M., H. M. B. Harris, R. P. Ross, and P. W. O’Toole. 2017. Core fecal

microbiota of domesticated herbivorous ruminant, hindgut fermenters, and

monogastric animals. Microbiology Open 6:1-11.

Oka, K., T. Osaki, T. Hanawa, S. Kurata, E. Sugiyama, M. takahashi, M. Tanaka, H.

Tagushi, and S. Kamiya. 2018. Establishment of an endogenous Clostridium

difficile rat infection model and evaluation of the effects of Clostridium butyricum

MIRAIRI 588 probiotic strain. Frontiers in Microbiology 9:1-9.

Parish, J. A. 2017. Understanding the ruminant animal digestive system. Mississippi State

University Extension.

Probiotic 3. 2019. Advanced Orthomolecular Research https://aorhealth.com/products/

probiotic-3/

Peechakara, B. V., and M. Gupta. 2020. Ampicillin/sulbactam. StatPearls. Treasure Island

(FL): StatPearls Publishing.

Pen and Tec Consulting S. L. 2012. Public version – Clostridium butyricum MIRAIRI 588

novel food application. Mirarisan Pharmaceutical Co. Ltd.

Peng, L., Z. R. Li, R. S. Green, I. R. Holzman, and J. Lin. 2009. Butyrate enhances the

intestinal barrier by facilitating tight junction assemble via activation of AMP-

activated protein kinase in Caco-2 cell monolayers. The Journal of Nutrition

139:1619-1625.

Peng, Z., D. Jin, H. B. Kim, C. W. Stratton, B. Wu, Y. W. Tang, and X. Sun. 2017. Update

on antimicrobial resistance in Clostridium difficile: resistance mechanisms and

antimicrobial susceptibility testing. Journal of Clinical Microbiology 55:1998-

2008.

Pirozzi, C., V. Francisco, F. Di Guida, R. Gomez, F. Lago, J. Pino, R. Meli, and O. Gualillo.

2018. Butyrate modulates inflammation in chondrocytes via GPR43 receptor.

Cellular Physiology and Biochemistry 21:228-243.

Probiotic-3. 2020. Advanced Orthomolecular Research. https://aor.us/products/probiotic-

3/#

100

QA International. 2003. The horse’s digestive system. The Visual Dictionary. Hygain

https://hygain.com.au/blogs/library/horses-digestive-system

Rainey, F. A., B. J. Hollen, and A. M. Small. 2015. Clostridium. Bergey’s Manual of

Systematics of Archaea and Bacteria. (eds M. E. Trujillo, S. Dedysh, P. DeVos, B.

Helund, P. Kampfer, F. A. Rainey, and W.B. Whitman). John Wiley & Sons, Inc.,

in association with Bergey’s Manual Trust.

Reddy, A. S., B. P. Girinathan, R. Zapotocny, R. Govind. 2013. Identification and

characterization of Clostridium sordellii toxin gene regulator, Journal of

Bacteriology 195:4246-4256.

Richards, J. D., J. Gong, and C. F. M. de Lange. 2005. The gastrointestinal microbiota and

its role in monogastric nutrition and health with an emphasis on pigs: current

understanding, possible modulations, and new technologies for ecological studies.

Canadian Journal of Animal Science 85:421-435.

Rinninella, E., P. Raoul, M. Cintoni, F. Franceschi, G. A. D. Miggiano, A. Gasbarrini, and

M. C. Mele. 2019. What is the healthy gut microbiota composition? A changing

ecosystem across age, environment, diet, and diseases. Microorganisms 7:1-22.

Rowan, J. P., K. L Durrance, G. E. Combs, and L.Z. Fisher. 2015. The digestive tract of

the pig. University of Florida IFAS Extension.

Saemann, M. D., G. A. Bohmig, C/ H/ Osterreicher, H. Burtscher, O. Parolini, C. Diakos,

J. Stockl, W. H. Horl, and G. J. Zlabinger. 2000. Anti-inflammatory effects of

sodium butyrate on human monocytes: potent inhibition of IL-12 and up-regulation

of IL-10 production. 14:2380-2403.

101

Samul, D., P. Worztynowicz, K. Leja, and W. Grajek. 2013. Beneficial and harmful roles

of bacteria from the Clostridium genus. ACTA biochimica Polonica 60:515-521.

Sanders, M. E. 2008. Probiotics: definition, sources, selection, and uses. Clinical Infectious

Diseases 46:S58-61.

Schwarz, S., C. Kehrenberg, B. Doublet, and A. Cloeckaert. 2004. Molecular basis of

bacterial resistance to chloramphenicol and florfenicol. Federation of European

Microbiological Societies. 28:519-542.

Segain, J. P., D. Raingaerd de la Bletiere, A. Bourreille, V. Leray, N. Gervois, C. Rosales,

L. Ferrier, C. Bonnet, H. M. Blottiere, and J. P. Galmiche. 2000. Butyrate inhibits

inflammatory responses through NFκB inhibition: implications for Crohn’s

disease. Gut 47:397-403.

Seshadri, R., S. C. Leahy, G. T. Attwood, K. H. The, S. C. Lambie, A. L. Cookson, E. A.

Eloe-Fadrosh, G. A. Pavlopoulos, M. Hadjithomas, N. J. Varghese, D. Paez-

Espino, Hungate1000 project collaborators, R. Perry, G. Henderson, C. J. Creevey,

N. Terrapon, P. Lapebie, E. Drula, V. Lombard, E. Rubin, N. C. Kyrpides, B.

Henrissat, T. Woyke, N. N. Ivanova, and W. J. Kelly. 2018. Cultivation and

sequencing of rumen microbiome members from the Hungate1000 Collection.

Nature Biotechnology 36 359-371.

Shang, H., J. Sun, and Y. Q. Chen. 2016. Clostridium butyricum CGMCC0313.1 modulates

lipid profile, insulin resistance and colon homeostasis in obese mice. PloS ONE

11:1-15.

Singh, N., A. Gurav, D. Sivaprakasam, e. Brady, r. Padia, H. Shi, M. Thangaraju, P. D.

Prasad, S. Manicassamy, D. H. Munn, J. R. Lee, S. Offermanns, and V. Ganapathy.

102

2014. Activation of the receptor (Gpr109a) for niacin and the commensal

metabolite butyrate suppresses colonic inflammation and carcinogenesis. Immunity

40:128-139.

Stevens, D. L., A. E. Bryant, and K. Carroll. 2015. Clostridium. In Manual of Clinical

Microbiology (eds J. H. Jorgensen and M. A Pfaller). ASM Press.

Sun, J., J. Xu, B. Yang, K. Chen, Y. Kong, N. Fang, T. Gong, F. Wang, Z. Ling, and J. Liu.

2020. Effect of Clostridium butyricum against microglia-mediated

neuroinflammation in Alzheimer’s disease via regulating gut microbiota and

metabolites butyrate. Molecular Nutrition Food Research 64:1-11.

Sun, Y. Y., L. Ming, Y. Y. Li, L. X. Li, W. Z. Zhai, P. Wang, X. X. Yang, X. Gu, L. J.

Song, Z. Li, X, L, Zuo, and Y. Q. Li. 2018. The effect of Clostridium butyricum on

the symptoms and fecal microbiota in diarrhea-dominant irritable bowel syndrome:

a randomized, double-blind, placebo-controlled trial. Scientific reports 8:1-11.

Takaishi, H., T. Matsuki, A. Nakazawa, T. Takada, S. Kado, T. Asahara, N. Kamada, A.

Sakuraba, T. Yajima, H. Higuchi, N. Inoue, H. Ogata, Y. Iwao, K. Nomoto, R.

Tanaka, and T. Hibi. 2008. Imbalance in intestinal microflora constitution could be

involved in the pathogenesis of inflammatory bowel disease. International Journal

of Medical Microbiology 298:463-472.

Takeshita, K., S. Mizuno, Y. Mikami, T. Sujino, K. Saigusa, K. Matsuoka, M. Naganuma,

T. Sato, T. Takada, H. Tsuji, A. Kushiro, K. Nomoto, and T. Kanai. 2016. A single

of Clostridium subcluster XIVa decreased in ulcerative colitis. Inflammatory

Bowel Diseases 22:2802-2810.

103

Taschuk, R. and P. J. Griebel. 2012. Commensal microbiome effects on mucosal immune

system development in the ruminant gastrointestinal tract. Animal Health Research

Reviews 13:129:141.

Tian, Y., M. Li, W. Song, R. Jiang, and Y. Q. Li. 2019. Effects on chemotherapy in patients

with lung cancer. Oncology Letters 17:2836-2848.

Trepani, K. M., L. J. Boghssian, and E. Caskey. 2017. Clostridium subterminale septicemia

in a patient with metastatic gastrointestinal adenocarcinoma. Case Repots to

Infectious Diseases 2018:1-3.

Turner, S., Pryer, K.M., Miao, V.P.W., and Palmer, J.D. 1999. Investigating deep

phylogenetic relationships among cyanobacteria and plastids by small subunit

rRNA sequence analysis. Journal of Eukaryotic Microbiology 46: 327–338.

Vandenplas, Y., G. Huys, and G. Daube. 2015. Probiotics: an update. Jornal de Pediatria

91:6-21.

Vanhoutvin, S. A., F. J. Troost, H. M. Hamer, P. L. Lindsey, G. H. Koek, D. M. Jonkers,

A. Kodde, K. Venema, and R. J. Brummer. 2009. Butyrate-induced transcriptional

changes in human colonic mucosa. PloS ONE 4:1-7.

Vidyasagar, S., and B. S. Ramakrishna. 2002. Effects of butyrate on active sodium and

chloride transport in rat and rabbit distal colon. Journal of Physiology 539:163-173.

Vital, M., C. R. Penton, Q. Wang, V. B. Young, D. A. Antonopoulos, M. L. Sogin, H. G.

Morrison, L. Raffals, E. B. Chang, G. B. Huffngale, et al. 2013. A gene-targeted

approach to investigate the intestinal butyrate-producing bacterial community.

Microbiome 1:1-14.

104

Wan, L. Y. M., Z. J. Chen, N. P. Shah, and H. El-Nezami. 2016. Modulation of intestinal

epithelial defense responses by probiotic bacteria. Critical Reviewa in Food

Science and Nutrition 56:2628-2641.

Wang, F. Y., J. M. Liu, H. H. Luo, A. H. Liu, and Y. Jiang. 2015. Potential protective

effects of Clostridium butyricum on experimental gastric ulcers in mice. World

Journal of Gastroenterology 21:8340-8351.

Wang, H. L., T. He, S. Becker, G. Zhang, D. Li, and X. Ma. 2018. Butyrate: a double-

edged sword for health? American Society for Nutrition 9:21-29.

Williams, B. A., D. Mikkelsen, B. M. Flanagan, and M. K. Gidley. 2019. “Dietary fibre:”

moving beyond the “soluble/insoluble” classification for monogastric nutrition,

with an emphasis on human and pigs. Journal of Animal Science and Biotechnology

10:1-12.

Wong, A., D. Y. S. Ngu, L. A. Dan, A. Ooi, and R. L. Hong. 2015. Detection of antibiotic

resistance in probiotics of dietary supplements. Nutrition Journal 141:1-6.

Yan, H, and K. M. Ajuwon. 2017. Butyrate modifies intestinal barrier function in IPEC-J2

cells through a selective upregulation of tight junction proteins and activation of

the Akt signaling pathway. PloS ONE 12:1-20.

Yasueda, A.,T. Mizushima, T., R. Nezu, R. Sumi, M. Tanaka, J. Nishimura, Y. Kai, M.

Hirota, H. Osawa, K. Nakajima, M. Mori, and T. Ito. 2016. The effect of

Clostridium butyricum MIYAIRI on the prevention of pouchitis and alteration of

the microbiota profile in patients with ulcerative colitis. Surgery Today 46:939-

949.

105

Yoo, H. S., S. U. Lee, K. Y. Park, and Y. H. Park. 1997. Molecular typing and

epidemiological survey of prevalence of Clostridium perfringens types by

multiplex PCR. Journal of Clinical Microbiology 35:228-232.

Yoshimatsu, Y., A. Yamada, R. Furukawa, K. Sono, A. Osamura, K. Nakamura, H. Aoki,

Y. Tsuda, N. Hosoe, N. Takada, and Y. Suzuki. 2015. Effectiveness of probiotic

therapy for the prevention of relapse in patients with inactive ulcerative colitis.

World Journal of Gastroenterology 21:5986-5994.

Zeng, H., D. P. Taussig, W. H. Cheng, L. K. Johnson, and R. Hakkak. 2017. Butyrate

inhibits cancerous HCT116 colon cell proliferation but to a lesser extent in

noncancerous NCM460 colon cells. Nutrients 9:1-13.

Zhang, H. Q., T. T. Ding, J. S. Zhao, X. Yang, H. X. Zhang, J. J. Zhang, and Y. L. Cui.

2009. Therapeutic effects of Clostridium butyricum on experimental colitis induced

by oxazolone in rats. Worls Journal of Gastroenterology 15:1821-1828.

Zhang, J., M. Yi, L. Zha, S. Chen, Z. Li, C. Li, M. Gong, H. Deng, X. Chu, J. Chen, Z.

Zhang, L. Mao, and S. Sun. 2016. Sodium butyrate induces endoplasmic reticulum

stress and autophagy in colorectal cells: implications for apoptosis. PloS ONE 11:1-

25.

Zhao, X., J. Yang, Z. Ju, J. Wu, L. Wang, H. Lin, and S. Sun. 2020. Clostridium butyricum

ameliorates Salmonella enteritidis induced inflammation by enhancing and

improving immunity of the intestinal epithelial barrier at the intestinal mucosal

level. Frontiers in Microbiology 11:1-11.

106

Zheng, M., R. Zhang, X. Tian, X. Xhou, X. Pan, and A. Wong. 2107. Assessing the risk of

probiotic dietary supplements in the context of antibiotic resistance. Frontiers in

Microbiology 8:1-8.

Zhou, Y., W. Xu, K. Hong, H. Li, J. Zhang, X. Chen, Y. Zhu, Q. Zhang, F. Ding, and F.

Wang. 2018. Therapeutic effects of probiotic Clostridium butyricum WZ001 on

bacterial vaginosis in mice. Journal of Applied Microbiology 127:565-575.

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APPENDIX A

Figure 1. Multiple sequence alignment of the species listed in Table 3.8 using T-Coffee for the design of BUK gene primers with the primer region boxed.

108

Figure 1 (continued)

109

Figure 1 (continued)

110

Figure 1 (continued)

111

APPENDIX B

A H1 B H2 C H3 A B C D E F + - A B C D E F + - B C G + -

D H4 E H5 A C F + - A B C D E F + -

Figure 2. Agarose gel electrophoresis of the 16S rRNA gene in presumptive Clostridium isolates from horse samples (A) H1, (B) H2, (C) H3, (D) H4, and (E) H5. The 16S rRNA gene presents at 1465 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control. A C1 B C2 C C3 A B C D E F G + - A B D E G + - D E F + -

Figure 3. Agarose gel electrophoresis of the 16S rRNA gene in presumptive Clostridium isolates from cattle samples (A) C1, (B) C2, (C) C3, (D) C4, and (E) C5. The 16S rRNA gene presents at 1465 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control.

112

D C4 E C5 A B C D E G H I + - B C D E + -

Figure 3. (continued)

A S1 B S2 A B C D E F + - A B C D E + -

C S3 D S4 E S5 A B C D E + - B C + - A B C D E + -

Figure 4. Agarose gel electrophoresis of the 16S rRNA gene in presumptive Clostridium isolates from sheep samples (A) S1, (B) S2, (C) S3, (D) S4, and (E) S5. The 16S rRNA gene presents at 1465 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control.

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A P1 B P2 C P3 A B D E F + - A D E + - B D G H + -

D P4 E P5 A B C D F H + - C E F G H + -

Figure 5. Agarose gel electrophoresis of the 16S rRNA gene in presumptive Clostridium isolates from pig samples (A) P1, (B) P2, (C) P3, (D) P4, and (E) P5. The 16S rRNA gene presents at 1465 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control. A G1 B G2 C G3 B D E + - A B C D + - A B C D E + -

Figure 6. Agarose gel electrophoresis of the 16S rRNA gene in presumptive Clostridium isolates from goat samples (A) G1, (B) G2, (C) G3, (D) G4, and (E) G5. The 16S rRNA gene presents at 1465 bp in size with C. sporogenes ATCC 11437 used as a positive control and sterile ultrapure water used as a negative control.

114

D G4 E G5 A B C D E + - A B C D E + -

Figure 6. (continued)

115

APPENDIX C

Table 1. Endpoint determination of acidogenesis in Clostridium sporogenes by OD at a wavelength of 600 nm and pH readings taken over 12 hours. C. perfringens ATCC 13124 was used as a positive control and E. coli ATCC 25922 was used as a negative control. C. perfringens ATCC 13124 E. coli ATCC 25922 C. sporogenes ATCC 11437 1 2 3 average 1 2 3 average 1 2 3 average Time (h) OD (600 nm) 0 0.099 0.101 0.106 0.102 0.106 0.115 0.097 0.106 0.102 0.115 0.096 0.104 1 0.105 0.096 0.110 0.104 0.180 0.170 0.210 0.187 0.139 0.189 0.148 0.159 2 0.266 0.225 0.280 0.257 0.502 0.450 0.520 0.491 0.235 0.327 0.219 0.260 3 0.620 0.608 0.676 0.635 0.740 0.690 0.738 0.723 0.456 0.574 0.410 0.480 4 1.160 1.120 1.200 1.160 0.875 0.860 0.870 0.868 0.815 0.910 0.720 0.815 5 1.460 1.460 1.500 1.473 0.980 0.955 0.970 0.968 1.100 1.230 1.040 1.123 6 1.660 1.660 1.660 1.660 1.060 1.045 1.070 1.058 1.370 1.420 1.300 1.363 116 7 1.720 1.740 1.760 1.740 1.120 1.120 1.100 1.113 1.460 1.540 1.460 1.487

8 1.760 1.760 1.780 1.767 1.130 1.140 1.150 1.140 1.600 1.640 1.580 1.607 9 1.760 1.760 1.800 1.773 1.130 1.140 1.140 1.137 1.680 1.700 1.680 1.687 10 1.760 1.760 1.780 1.767 1.140 1.140 1.130 1.137 1.740 1.760 1.740 1.747 11 1.760 1.760 1.780 1.767 1.150 1.150 1.130 1.143 1.780 1.800 1.780 1.787 12 1.760 1.760 1.780 1.767 1.140 1.130 1.130 1.133 1.800 1.850 1.800 1.817 Time (h) pH 0 6.51 6.51 6.51 6.51 6.51 6.51 6.51 6.51 6.51 6.51 6.51 6.51 2 6.36 6.39 5.39 6.05 6.20 6.27 6.14 6.20 6.51 6.51 6.51 6.51 4 5.61 5.61 5.61 5.61 5.70 5.78 5.73 5.74 6.52 6.51 6.49 6.51 6 5.14 5.20 5.17 5.17 5.46 5.48 5.43 5.46 6.55 6.57 6.56 6.56 8 5.00 5.00 5.05 5.02 5.28 5.28 5.27 5.28 6.47 6.48 6.50 6.48 10 5.01 5.01 5.02 5.01 5.27 5.28 5.25 5.27 6.45 6.42 6.47 6.45 12 5.06 5.05 5.06 5.06 5.10 5.11 5.09 5.10 6.67 6.61 6.66 6.65

Table 1. (continued) C. sporogenes ATCC 15579 C. sporogenes ATCC 19404 C. sporogenes ATCC 25779 1 2 3 average 1 2 3 average 1 2 3 average Time (h) OD (600 nm) 0 0.097 0.107 0.100 0.104 0.099 0.112 0.101 0.104 0.105 0.113 0.113 0.110 1 0.143 0.207 0.185 0.178 0.125 0.130 0.130 0.128 0.127 0.126 0.113 0.122 2 0.197 0.345 0.302 0.281 0.267 0.258 0.274 0.266 0.131 0.127 0.110 0.123 3 0.267 0.672 0.572 0.504 0.482 0.490 0.508 0.493 0.174 0.150 0.127 0.150 4 0.434 1.060 1.035 0.843 0.920 0.915 0.905 0.913 0.240 0.208 0.184 0.211 5 0.750 1.400 1.310 1.153 1.350 1.330 1.350 1.343 0.408 0.338 0.300 0.349 6 1.150 1.580 1.560 1.430 1.560 1.560 1.540 1.553 0.732 0.580 0.582 0.631 7 1.480 1.600 1.600 1.560 1.700 1.700 1.700 1.700 1.025 0.885 0.850 0.920 117 8 1.640 1.620 1.620 1.627 1.800 1.800 1.800 1.800 1.280 1.180 1.170 1.210

9 1.640 1.620 1.620 1.627 1.850 1.900 1.850 1.867 1.440 1.350 1.350 1.380 10 1.640 1.620 1.620 1.627 1.900 1.950 1.850 1.900 1.560 1.520 1.520 1.533 11 1.620 1.620 1.620 1.620 1.950 1.950 1.900 1.933 1.620 1.580 1.580 1.593 12 1.640 1.620 1.620 1.627 1.950 1.950 1.950 1.950 1.680 1.640 1.640 1.653 Time (h) pH 0 6.51 6.51 6.51 6.51 6.51 6.51 6.51 6.51 6.51 6.51 6.51 6.51 2 6.50 6.51 6.53 6.51 6.52 6.52 6.52 6.52 6.56 6.51 6.53 6.53 4 6.60 6.60 6.60 6.60 6.60 6.61 6.63 6.61 6.56 6.49 6.52 6.52 6 6.57 6.12 6.09 6.26 6.38 6.41 6.41 6.40 6.56 6.55 6.57 6.56 8 5.81 5.77 5.94 5.84 6.10 6.08 6.06 6.08 6.57 6.57 6.57 6.57 10 5.81 5.65 5.65 5.70 5.97 5.98 5.98 5.98 6.65 6.67 6.68 6.67 12 5.70 5.55 5.50 5.58 5.87 5.87 5.85 5.86 6.85 6.85 6.84 6.85