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The intestinal epithelium during times of dynamic change : development and enteric infection Bhinder, Ganive 2017

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THE INTESTINAL EPITHELIUM DURING TIMES OF DYNAMIC CHANGE: DEVELOPMENT AND ENTERIC INFECTION by  Ganive Bhinder  B.Sc. Specialization, The University of Alberta, 2011  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Experimental Medicine)   THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)   August 2017  © Ganive Bhinder, 2017 ii  Abstract Enteric infections and neonatal intestinal development both represent times of significant change in the intestine.  Mode of delivery as well as food source, i.e. breast milk or formula, may impact later susceptibility to the development of allergies, asthma and other inflammatory diseases.  Similarly, the development of pathological conditions such as irritable bowel syndrome and inflammatory bowel disease have been reported as long-term sequelae of infection by various enteric pathogens.  The single cell layer of epithelial cells lining the intestine are in closest proximity to the luminal changes that occur, be it interaction with a bacterial pathogen or the introduction of breast milk.  This positions intestinal epithelial cells (IEC)s to serve as key players in shaping responses during these times.  To address the importance of IEC responses during enteric infections, we first examined the intricacies of IEC innate immune-mediated responses (MyD88-dependent) using in vivo models.   Following infection with the common food poisoning pathogen Salmonella Typhimurium, IEC-specific MyD88 signalling was required to limit pathogen penetration of intestinal crypts and mediate goblet cell/antimicrobial responses.  IECs lacking MyD88 also displayed decreased bactericidal capacity against Salmonella and Citrobacter rodentium, a close relative of enteropathogenic Escherichia coli.  Thus, IEC MyD88 signalling promotes early, protective responses during enteric infection.  Next, IEC changes during intestinal development, as well as the impact of food source during this process (i.e. mother’s milk versus formula) were explored using a neonate pup-in-a-cup model.   Specifically, the role of milk fats found in mammalian milk (and currently not widely used in formulas), referred to as milk fat globule membrane (MFGM), were assessed for their ability to normalize intestinal development to that seen with mother’s milk. Interestingly, MFGM addition to formula resulted in similar villus and crypt development, as well as goblet cell, Paneth cell and enterocyte iii  numbers and/or expression to that of mother’s milk fed pups.  Further, addition of MFGM protected the formula fed neonate from intestinal damage by bacterial toxins.    These studies highlight IECs as key players in shaping beneficial responses during enteric infection and intestinal development and have important implications for newborn health as well as for populations vulnerable to enteric infections.      iv  Lay summary  Times of change in the intestinal environment, such as after birth or during food poisoning, impact the inner lining of the intestine.  This lining is only one cell-layer thick, yet it is responsible for preventing the contents within our intestines (bacteria and food) from leaking into our bodies and causing damage. Using animal models infected with food poisoning bacteria, I discovered that early immune signalling within this lining is important for releasing protective factors, which when absent results in greater intestinal damage and a leaky gut.    Further, using a newborn rat model, I tested the impact of adding breast milk fat to formula on development of the intestinal lining. I found that breast milk fat supplemented formula results in similar intestinal development as mother’s milk, and protects against intestinal damage by bacterial toxins. These findings have implications for newborn health, especially premature births, as breast milk is not always available.  v  Preface Chapter 2 A version of this chapter has been published in the journal Infection and Immunity as: Bhinder G, Stahl M, Sham HP, Crowley SM, Morampudi V, Dalwadi U, Ma C, Jacobson K, Vallance BA. Intestinal epithelial specific MyD88 signalling impacts host susceptibility to infectious colitis by promoting protective goblet cell and antimicrobial responses. Infection & Immunity, 2014 September:82(9):3753-63. doi: 10.1128/IAI.02045-14.    I designed and conducted the majority of the studies reported in this chapter, analyzed the data, prepared all of the figures and wrote the manuscript under the supervision of Dr. Bruce A. Vallance.  Dr. Ho Pan Sham and Ms. Caixia Ma helped to generate the IEC-Myd88-/- mice, as well as in pathology scoring (H. P. S.) and collection of blood for serum analysis (C. M.) leading to Figure 2.2B and Figure 2.5A.   Dr. Martin Stahl performed the intestinal microbe assessment via quantitative PCR leading to Figure 2.4.  Ms. Shauna M. Crowley assisted in immunofluorescent staining leading to Figure 2.3C and in enumeration of Ki-67 positive cells in Figure 2.5C. Dr. Vijay Morampudi and Mr. Udit Dalwadi assisted in performing the crypt killing assay presented in Figure 2.10, while Dr. Kevan Jacobson provided reagents and input to the study design.  Ethics approval was required for this research and was obtained from the University of British Columbia Animal Care Committee certificate numbers: A11-0253 and A11-0290.    vi  Chapter 3 A version of this chapter has been published in the journal Scientific Reports as:  Bhinder G, Allaire JM, Garcia C, Lau JT, Chan JM, Ryz NR, Bosman ES, Graef FA, Crowley SM, Celiberto LS, Berkmann JC, Dyer, RA, Jacobson K, Surette M, Innis SM, Vallance BA. Milk Fat Globule Membrane Supplementation in Formula Modulates the Neonatal Gut Microbiome and Normalizes Intestinal Development. Scientific Reports, 2017 (7) 45274; doi: 10.1038/srep45274.  Dr. Bruce A. Vallance, Dr. Sheila M. Innis, Dr. Cyrielle Garcia and I conceived and designed the experiments for this study, with input from Mr. Roger A. Dyer and Dr. Kevan Jacobson.  I analyzed the majority of the data, prepared all but one of the figures, and wrote the manuscript under the supervision of Dr. Bruce A. Vallance.  Dr. Joannie M. Allaire performed immunofluorescent staining, enumeration of commensal microbes and histological analysis leading to Figure 3.10A, 3.11A-C and 3.12.  Ms. Jennifer T. Lau and Dr. Michael Surette performed the in-depth commensal analysis and generated the graphs for Figure 3.8B-E and 3.9. Mr. Justin M. Chan provided input for the appropriate statistical analysis of data generated for this study.  Dr. Natasha R. Ryz and Ms. Else S. Bosman assisted in tissue collection in addition to monitoring of rat pups with which Ms. Franziska A. Graef, Ms. Shauna M. Crowley, Ms. Larissa S. Celiberto, Ms. Julia C. Berkmann and Mr. Roger A. Dyer also assisted.  Mr. Roger A. Dyer prepared the formula used in this study.   Ethics approval was required for this research and was obtained from the University of British Columbia Animal Care Committee certificate number: A13-0257. vii  Table of contents Abstract .......................................................................................................................................... ii	Lay summary ................................................................................................................................ iv	Preface ............................................................................................................................................ v	Table of contents .......................................................................................................................... vii	List of tables ................................................................................................................................. xii	List of figures .............................................................................................................................. xiii	List of symbols and abbreviations ............................................................................................. xvi	Acknowledgements ...................................................................................................................... xx	Dedication ................................................................................................................................... xxii	Chapter 1: Introduction ................................................................................................................ 1	1.1	 The gastrointestinal tract ................................................................................................ 2	1.1.1	 Lower GI tract: jejunum, ileum and large intestine ................................................. 5	1.1.1.1	 Histological characteristics ............................................................................... 5	1.1.1.2	 Distribution of IECs ......................................................................................... 6	1.1.1.3	 Regional distribution of lymphoid tissues and immune cells ........................... 8	1.1.2	 Intestinal bacteria ................................................................................................... 10	1.1.2.1	 Host factors influencing bacterial colonization in the intestine ..................... 12	1.1.2.2	 Impact of bacterial factors on host cells ......................................................... 13	1.2	 Intestinal epithelial barrier and cell subtypes ............................................................... 15	1.2.1	 Tight junctions and the intestinal barrier ............................................................... 16	1.2.2	 Intestinal epithelial cell lineages ........................................................................... 19	1.2.2.1	 Absorptive cell types ...................................................................................... 24	viii  1.2.2.2	 Secretory cell types ......................................................................................... 24	1.2.3	 Goblet cell secreted proteins and the mucus layer ................................................ 26	1.3	 Innate immunity in intestinal epithelial cells ................................................................ 29	1.3.1	 MyD88 dependent signalling pathways ................................................................ 30	1.3.1.1	 Interleukin-1 receptors .................................................................................... 32	1.3.1.2	 Toll-like receptors .......................................................................................... 33	1.3.2	 Innate immune responses to infection ................................................................... 37	1.4	 Enteric infections and disease models .......................................................................... 38	1.4.1	 Salmonella enterica serovar Typhimurium induced gastroenteritis ...................... 40	1.4.2	 Murine model of attaching and effacing pathogens: Citrobacter rodentium ........ 43	1.4.3	 Clostridium difficile infection ................................................................................ 44	1.4.4	 Necrotizing enterocolitis ....................................................................................... 46	1.5	 Changes in the intestinal environment at birth ............................................................. 47	1.5.1	 Introduction of food and intestinal development ................................................... 48	1.5.2	 Breast milk composition and benefits ................................................................... 49	1.6	 Milk fat globule membrane .......................................................................................... 51	1.6.1	 Composition .......................................................................................................... 51	1.6.2	 Modeling neonate development: rat pup-in-a-cup model ...................................... 53	1.7	 Research objectives ...................................................................................................... 54	Chapter 2: Intestinal epithelial specific MyD88 signalling impacts host susceptibility to enteric infection through goblet cell and antimicrobial responses ......................................... 57	2.1	 Introduction .................................................................................................................. 57	2.2	 Experimental procedures .............................................................................................. 60	ix  2.2.1	 Mice ....................................................................................................................... 60	2.2.2	 Bacterial strains and infection of mice .................................................................. 61	2.2.3	 Tissue collection and bacterial counts ................................................................... 62	2.2.4	 Histology scoring ................................................................................................... 62	2.2.5	 Resident microbe analysis ..................................................................................... 63	2.2.6	 FITC-dextran intestinal permeability assay ........................................................... 63	2.2.7	 RNA extractions and quantitative real-time PCR ................................................. 64	2.2.8	 Immunofluorescence ............................................................................................. 64	2.2.9	 Fluorescence intensity measurements ................................................................... 65	2.2.10	 Crypt killing assay ............................................................................................... 65	2.2.11	 Statistical analysis ............................................................................................... 66	2.3	 Results .......................................................................................................................... 66	2.3.1	 IEC-Myd88-/- mice suffer exaggerated S. Typhimurium induced gastroenteritis .. 66	2.3.2	 IEC-Myd88-/- mice show altered localization of S. Typhimurium within ceca ..... 70	2.3.3	 IEC specific MyD88 signalling does not overtly alter the gut microbiome .......... 72	2.3.4	 MyD88 signalling in IECs protects barrier integrity during infection .................. 73	2.3.5	 IEC-Myd88-/- mice are impaired in antimicrobial and goblet cell specific responses   ............................................................................................................................... 76	2.3.6	 IEC-Myd88-/- mice also show early susceptibility to C. rodentium infection ........ 80	2.3.7	 IEC-Myd88-/- crypt epithelial cells show decreased bactericidal activity ............. 84	2.4	 Discussion ..................................................................................................................... 86	Chapter 3: Milk fat globule membrane supplementation in formula modulates the neonatal gut microbiome and normalizes intestinal development ......................................................... 91	x  3.1	 Introduction .................................................................................................................. 91	3.2	 Experimental procedures .............................................................................................. 94	3.2.1	 Animals and provision of formula ......................................................................... 94	3.2.2	 Milk fat supplementation ....................................................................................... 95	3.2.3	 Tissue collection .................................................................................................... 96	3.2.4	 Crypt depth and villus height measurements ......................................................... 97	3.2.5	 FITC-dextran barrier permeability assay ............................................................... 97	3.2.6	 Immunofluorescence ............................................................................................. 97	3.2.7	 Fluorescence intensity measurements and cell counts .......................................... 98	3.2.8	 Intestinal microbe counts ....................................................................................... 99	3.2.9	 DNA extraction and 16S rRNA gene sequencing, processing and analysis ....... 100	3.2.10	 Antibiotic treatments ......................................................................................... 101	3.2.11	 Clostridium difficile toxin induced intestinal inflammation .............................. 101	3.2.12	 Statistical analysis ............................................................................................. 102	3.3	 Results ........................................................................................................................ 103	3.3.1	 Rat pup growth and gross intestinal characteristics ............................................ 103	3.3.2	 Impact of MFGM on intestinal epithelial architecture and barrier ...................... 104	3.3.3	 Changes in IEC subtypes ..................................................................................... 109	3.3.3.1	 Paneth cells ................................................................................................... 109	3.3.3.2	 Goblet cells ................................................................................................... 110	3.3.3.3	 Enteroendocrine cells ................................................................................... 112	3.3.3.4	 Enterocytes ................................................................................................... 114	3.3.4	 Contribution of microbiota to intestinal development ......................................... 115	xi  3.3.5	 MFGM supplementation protects against Clostridium difficile toxin ................. 123	3.4	 Discussion ................................................................................................................... 126	Chapter 4: Discussion ................................................................................................................ 132	4.1	 The suitability of rodent models for the study of the human intestine ....................... 133	4.2	 Pup-in-a-cup model: advantages and potential improvements ................................... 137	4.3	 Changes in the intestinal environment during enteric infection and neonate development– potential influence on later health ................................................................ 139	4.3.1	 Future direction: exploring the long-term health consequences of MFGM supplementation in formula ............................................................................................. 141	4.3.2	 Enteric infections and their associated complications ......................................... 142	4.3.2.1	 Future direction: exploring the long term effects of increased susceptibility to early time-points of enteric infection .......................................................................... 145	4.4	 The impact of resident microbes shaped by food sources on intestinal development           .................................................................................................................................... 146 4.4.1	 Future direction: clarify the mechanisms through which MFGM impacts intestinal development .................................................................................................................... 149	4.5	 Similarities in IEC responses during enteric infection and neonate intestinal development ........................................................................................................................ 151	4.6	 Antibiotic induced disruption of the intestinal environment ...................................... 154	4.7	 Final remarks .............................................................................................................. 157	References .................................................................................................................................. 158	 xii  List of tables Table 1.1: Conventional CD4 T cell and ILC subsets found in the intestine ................................ 10	Table 3.1: Composition of formulas .............................................................................................. 96  xiii  List of figures Figure 1.1: Epithelial tight junctions and Claudin distribution along the length of the intestine  ....................................................................................................................................................... 18	Figure 1.2:  Maintenance of the stem cell niche and epithelial cell lineage commitment in the intestine .......................................................................................................................................... 22	Figure 1.3:  Muc2 structure and assembly within the goblet cell and upon luminal secretion ..... 28	Figure 1.4:  MyD88 dependent signalling pathway ....................................................................... 31	Figure 1.5:  Toll-like Receptors and their ligands ......................................................................... 35 Figure 1.6: Milk fat globule membrane secretion and structure ................................................... 52 Figure 2.1: Genotyping of mice ..................................................................................................... 61 Figure 2.2: IEC-MyD88 -/- mice are more susceptible to S. Typhimurium enteric infection ......... 67	Figure 2.3: IEC-MyD88 -/- mice suffer accelerated tissue damage during S. Typhimurium infection ......................................................................................................................................... 69	Figure 2.4: Disease severity of IEC-Myd88-/- mice is associated with altered S. Typhimurium localization .................................................................................................................................... 71	Figure 2.5: Similar bacterial populations at phylum level in stool samples of WT and IEC-Myd88-/- mice ................................................................................................................................. 73	Figure 2.6: IEC-MyD88 -/- mice display impaired barrier integrity at early infection time points ....................................................................................................................................................... 75	Figure 2.7: Gene transcripts for antimicrobial peptides and goblet cell mediators in WT and IEC-Myd88-/- mice ......................................................................................................................... 77	Figure 2.8: Muc2 and Relmβ production is impaired in infected IEC-Myd88-/- mice ................... 79	 xiv  Figure 2.9: IEC-Myd88-/- mice suffer accelerated tissue damage during C. rodentium infection  ....................................................................................................................................................... 82	Figure 2.10: IEC-Myd88-/- mice show defects in Muc2 staining and altered C. rodentium localization in tissue ...................................................................................................................... 84	Figure 2.11: Crypt epithelial cells lacking MyD88 have impaired antimicrobial capacity .......... 85	Figure 3.1: Formula feeding does not alter overall growth of rat pups compared to MM fed by pn day 15 .......................................................................................................................................... 104	Figure 3.2: MFGM (6g/L) supplementation in formula normalizes intestinal architecture and epithelial proliferation at pn day 15 ............................................................................................ 106	Figure 3.3: Localization of barrier proteins, Claudin-3 and Claudin-4, are similar in MM and 6 g/L MFGM pups .......................................................................................................................... 108	Figure 3.4: Addition of 6g/L MFGM to formula normalizes Paneth cell numbers at pn day 15  ..................................................................................................................................................... 110	Figure 3.5: Addition of 6 g/L MFGM to formula increases Muc2 positive staining at pn day 15 ..................................................................................................................................................... 112	Figure 3.6: Enteroendocrine numbers are similar in 6 g/L MFGM and MM pups ..................... 113	Figure 3.7: Enterocyte staining is similar in 6 g/L MFGM and MM pups at pn day 15 ............. 115	Figure 3.8: Food source impacts the composition of the intestinal microbiome ......................... 117	Figure 3.9: Differences in most abundant OTUs between the three groups ............................... 119	Figure 3.10: Microbes of MM pups can be efficiently depleted with antibiotic exposure resulting in changes to the intestinal architecture ..................................................................................... 121	Figure 3.11: Intestinal development is partially dependent on the presence of intestinal microbes in MM pups .................................................................................................................................. 123	xv  Figure 3.12: MFGM supplementation protects the formula fed neonate intestine from C. difficile toxin induced damage .................................................................................................................. 125	Figure 4.1: qPCR analysis of rat pup colonic tissues at pn day 15 ............................................. 138	Figure 4.2:  Impact of intestinal microbe depletion (45%) on intestinal development in MFGM formula fed pups .......................................................................................................................... 148	Figure 4.3:  Antibiotic treatment of genetically susceptibility (Tlr2-/-) mice results in altered responses ..................................................................................................................................... 156	 xvi  List of symbols and abbreviations α alpha β beta γ gamma κ kappa µ micro ° degree C Celsius < less than ≤ less than or equal to > greater than + positive ± plus or minus -/- deficient 5-HT 5-hyroxytyrtamine (serotonin) A/E attaching and effacing AMP antimicrobial peptide AP-1 activator protein 1 BMP bone morphogenetic protein CA carbonic anhydrase CBC crypt base columnar CD cluster of differentiation CDAD Clostridium difficile associated disease CFU colony forming units CpG cytidine-phosphate-guanosine CTL control DAPI 4',6-diamidino-2-phenylindole  DC dendritic cell DNA deoxyribose nucleic acid xvii  DSS dextran sodium sulfate EHEC Enterohemorrhagic Escherichia coli e.g. exempli gratia (for example) ER endoplasmic reticulum EPEC Enteropathogenic Escherichia coli FITC fluorescein isothiocyanate g gram GI Gastrointestinal tract GF germ free GPR G protein coupled receptor IBD Inflammatory Bowel Disease IBS irritable bowel syndrome i.e. id est (that is) IEC intestinal epithelial cell IEC-Myd88-/- IEC specific deletion of MyD88 signalling  IEL intraepithelial lymphocyte IFN interferon Ig immunoglobulin IκB  inhibitor of κB IL interleukin IL-1R interleukin-1 receptor iNOS inducible nitric oxide synthase IRAK IL-1 receptor associated kinase iLC innate lymphoid cell ILF isolated lymphoid follicles KC mouse keratinocyte derived chemokine  L litre Lgr5 leucine-rich repeat containing GPR5 LI large intestine LPS lipopolysaccharide xviii  m meters M cells microfold cells MAMP microbe associated molecular pattern MAPK mitogen activated protein kinase MFGM milk fat globule membrane MIP macrophage inflammatory protein MLN mesenteric lymph nodes MM mother’s milk Muc2 mucin 2 MyD88 myeloid differentiation primary response gene 88 NEC Necrotizing Enterocolitis NEMO NF-κB essential modulator NOD nucleotide-binding oligomerization domain NF-κB Nuclear factor kappa-B pi post infection pn post natal PP Peyer’s patches qPCR quantitative PCR Reg III γ regenerating islet-derived protein 3 gamma Relmβ resistin like molecule beta RNA ribonucleic acid rRNA ribosomal RNA SCFA short chain fatty acid SI small intestine SIGIRR Single Immunoglobulin Interleukin-1 related receptor SPI Salmonella pathogenicity island S. Typhimurium Salmonella enterica serovar Typhimurium T3SS type 3 secretion system TAB TAK1 binding proteins TAK1 TGF β activated kinase 1 xix  Tcd Clostridium difficile toxin TCR T cell receptor TFF3 trefoil like factor 3 TGF β transforming growth factor beta Th T helper Tir translocated intimin receptor TIR Toll/IL-1R TJ tight junction TLR toll like receptor TNF tumor necrosis factor TRAF6 TNF receptor associated factor 6 Treg T regulatory WT  wildtype  ZO zonula occludens  xx  Acknowledgements I would not be at this point today if it were not for the unreserved support, empathy and guidance of my supervisor, Dr. Bruce Vallance.  Thank you for knowing what I needed during the ups and downs, even when I did not.  I would also like to thank my committee members, Dr. Megan Levings and Dr. Kelly McNagny, who provided encouragement, advice and nothing but accommodation when the unexpected challenges of life arose. Dr. Sheila Innis for her dedication, throughout her career, to furthering our understanding of maternal and infant health and for inviting us to explore the world of neonate intestinal development. Dr. Kevan Jacobson and Dr. Laura Sly, for always providing a source of wonderful discussions.  To all the members of the Vallance Lab, past and present, thank you for creating such a wonderful environment where a love of science and life is equally fostered.  Dr. Andy Sham, Dr. Natasha Ryz, Justin Chan, Dr. Martin Stahl and Shauna Crowley – I am grateful that the overlap of our scientific journeys led to such great friendships.  To Dr. Joannie Allaire, Franzi Graef, Else Bosman, Larissa Celiberto, Carly Aspden, Dr. Vijay Morampudi, Dr. Kiran Bhullar, Dr. Hyungjun Yang and Dr. Hongbing Yu thank you all for the conversations, encouragement and help with the long experiments over the years.  I would also like to thank Caixia Ma, Tina Huang and Dr. Shelley Wu for always being so helpful and generous with their time.        To my brother, Amuel, thank you for everything: for leaving me with such love, understanding, and life-changing memories. Your stay on this earth may have been short, but the impact you’ve left is forever. To my partner Matthew, thank you for always believing in me and providing endless love and support through all of the good times as well as the challenges that life has thrown our way.  Thank you to my parents, Permi and Charan, your incredible strength and belief that I can do anything is my greatest strength.  Rob, Louise and the entire xxi  Fox/Silzer/MacDonald clan, I am thankful for all of your love and encouragement.  To my Vancouver friends who became family: Daniel, Danielle, Paul, Tash, Dave, Marc, Tessa: thank you for being there whenever I needed it.  Finally, I would like to thank all of the individuals affected by intestinal diseases that shared their stories with me; you are an incredible source of inspiration.      I would like to acknowledge the Canadian Institutes of Health Research Scholarships and the University of British Columbia Four Year Fellowship for funding.    xxii  Dedication To my mom and dad: if I asked for the moon you would give the earth, and had I asked for the earth you would give me the sun.1  Chapter 1: Introduction At birth, we emerge from the controlled environment of the womb to a shocking new, more complex, and constantly changing environment.  The gastrointestinal (GI) tract, like other mucosal sites such as the respiratory tract and eyes, is in continuous and direct interaction with this ever-changing environment(1).  Two major environmental components upon which we are in constant contact with from birth are our vast numbers of intestinal bacteria and the food from which we acquire nutrients. Adequate nutrient acquisition and establishment of a healthy microbial community are especially important for proper development during the neonatal period.  During this time, breast milk fuels growth, protects developing infants from infections, and also shapes the makeup of our intestinal microbes, which can have long lasting effects on our susceptibility to disease(2).  In addition to its roles in nutrient acquisition, development, and housing beneficial microbes, the GI tract serves an equally important function as a major protective physical and immune barrier.    The intestinal epithelium is best known for facilitating nutrient absorption, however it also plays a key role (along with the overlaying mucus layer) in creating a protective barrier that separates the microbes and food antigens found within the intestinal lumen from the rest of body.  Generating and maintaining this essential barrier between luminal contents and the systemic immune system helps avoid inordinate activation of inflammatory/immune signalling.  Being located at this interface between the host and gut microbes, it is of utmost importance that the GI epithelium is able to respond to enteric pathogens, while remaining tolerant to the resident luminal microbiota.  The necessity of this intricate balance is highlighted during conditions where it has gone awry.  For example, aberrant inflammatory activation and break down of the 2  epithelial/mucus barrier in response to the intestinal microbiota is a contributing factor to chronic GI diseases such as Inflammatory Bowel Diseases (IBD)(3).  In contrast, an inability to mount a rapid immune response to effectively clear enteric pathogens can clearly increase susceptibility and lead to exacerbated infections resulting in excessive morbidity and even mortality(4-7).  The following chapter will discuss these concepts in detail, with the goal of contextualizing how my research has aided in our understanding of the changes occurring at the GI epithelial surface during periods of development, infection and inflammation.      1.1 The gastrointestinal tract With a length of 9 meters (m) and a mucosal surface area of 200-300 m2 in adult humans the GI tract constitutes the largest organ system in contact with the external environment(1, 8-10).  It forms a continuous passageway through the body, extending from the mouth to the anus, and is associated with several glandular organs (salivary glands, liver and pancreas), which empty secretions into the tract to aid in digestion(1).  A major function of the GI system is nutrient acquisition from ingested food through the processes of digestion, absorption, and, ultimately, elimination of wastes.  This is achieved through a combination of physical and chemical breakdown of ingested materials as they move through the different regions of the tract.  Depending on the complexity of the animal, the GI tract can range from a rudimentary system, formed by a simple tube extending from mouth to anus as seen in lancelets (fish-like marine animals from a subphylum predating vertebrates), to more differentiated regions with specialized functions in the digestive process of higher vertebrates(11).  The latter is the case for humans and 3  commonly used laboratory animals, such as rats and mice, where the GI tract can be divided into two different sections, i.e. the upper and lower regions(12).  As food is ingested it first passes through the upper GI tract (mouth, pharynx, esophagus, stomach and duodenum) before entering the lower regions of the small intestine (SI) (jejunum, ileum) and large intestine (LI) (appendix, cecum, colon)(1, 10). In the mouth food is mechanically broken down by mastication while chemical digestion begins with amylases secreted by accessory salivary glands.  The food bolus, i.e. ball of food, then enters the esophagus via the pharynx, through which it is transported to the stomach.  Here the bolus undergoes further mechanical and chemical digestion, mediated by contracting of the muscularis externa within the stomach wall to mix digestive juices in the highly acidic glandular stomach environment.  These digestive juices contain hydrochloric acid that cleaves the zymogen pepsinogen to its active enzymatic form pepsin, which breaks down food proteins. From here the partially digested food, or chyme, enters the first of three segments of the 6-7 m long SI, known as the duodenum.  Upon receiving the acidic chyme, cells within the duodenum secrete the hormones cholecystokinin and secretin, which stimulate release of pancreatic enzymes and an alkaline bicarbonate buffer to aid in digestion and in the neutralization of luminal acidity. The duodenum also receives bile produced by the liver to aid in the digestion of fats.      Following these progressive digestive processes, the majority of nutrients are absorbed in the remaining regions of the SI (the jejunum and ileum)(1, 10, 13).  Nutrient absorption is optimized in these regions by maximizing the absorptive surface area of the intestinal epithelium through folds of the underlying submucosa, termed plicae circulares, as well as finger like projections of 4  the epithelium that rise into the lumen, termed villi.  The surface area of absorptive enterocytes within the epithelium is further increased by the presence of microvilli in the apical (luminal facing) cell membrane.  Between the villi, there exist invaginations of the epithelium termed crypts of Lieberkühn, which are also present in the LI.  Finally, the digested chyme enters the LI, which lacks villi and is much shorter (1.5 m), though considerably wider than the SI. Remaining water, electrolytes and vitamins are absorbed from the chyme, leaving compact feces that are eliminated from the body through the anus.  In the lower GI tract reside large populations of bacteria that can digest substances within the chyme that our digestive systems are unable to (on their own), producing vital metabolites as byproducts(14). These metabolites can be absorbed into the body, or used locally by intestinal epithelial cells (IECs), and can also alter immune function.   The GI tract – as seen in cross-section, is comprised of four major layers: the mucosa, submucosa, muscularis externa and serosa, with some regional differences in these structures found along its length(1, 8, 10).  The innermost layer, termed the mucosa, is composed of a single layer of columnar IECs, as well as the underlying lamina propria (connective tissue with capillaries and lymphoid follicles) and a thin double layer of smooth muscle referred to as the muscularis mucosa.  The surrounding submucosal layer contains larger blood vessels, lymphatic vessels as well as nerves (submucosal or Meissner’s plexus), which play a role in controlling blood flow and glandular secretions, as well as connective tissue.  The third layer, the muscularis externa drives food down the GI tract through a series of wave-like contractions (i.e. peristalsis) of its inner circular and outer longitudinal layers of smooth muscle. The actions of the muscularis externa are modulated by the myenteric plexus, which is located between the two muscle layers, 5  as well as by a network of pacemaker cells, termed the interstitial cells of Cajal.  The outermost layer of the GI tract is the serosa, composed of connective tissue and covered by the visceral peritoneum, which is a membrane that wraps around visceral organs within the abdominal cavity.  As a more thorough description of the entire GI tract is outside the scope of this thesis, moving forward I will focus my discussion on the intestinal epithelia and luminal contents of the large and lower small intestine (jejunum and ileum), which are most relevant to my research.    1.1.1 Lower GI tract: jejunum, ileum and large intestine 1.1.1.1 Histological characteristics The SI, at 6-7 m in length in adult humans, constitutes the longest portion of the GI tract.  At this site and in the LI as well, the mucosa, or innermost layer in direct contact with the lumen, is composed of a single layer of simple columnar epithelial cells(8, 15). Although the two regions of the lower SI (jejunum and ileum) are quite similar histologically, slight variations in the anatomical structure within which epithelial cells are arranged do exist.  In the jejunum, the plicae circulares are larger and the luminal projecting villi longer as compared to the ileum, likely reflecting the greater nutrient absorption occurring at this more proximal site.  In mice and rats, plicae circulares are absent in the SI and the surface area available for digestion and absorption of nutrients is reliant on increased villus length alone(16).  Located within the apical microvilli (or “brush border”) of the epithelial cells comprising the villi in the SI are several enzymes.  In addition to the luminal secretions from the stomach, pancreas, salivary glands and liver, these enzymes aid in digestion of carbohydrates, proteins and fats to an absorbable form(17, 18). These brush border enzymes include several peptidases, lactases and sucrases, among others.  Differential distribution of many of these enzymes exists between the jejunum 6  and ileum, with some displaying higher concentrations in the jejunum (e.g. sucrase, maltase, endopeptidase-24.11, angiotensin-converting enzyme) and others higher in the ileum (e.g. lactase, aminopeptidase W, caroboxypeptidase P)(19, 20).  Luminal nutrients in the SI can be taken up by the intestinal epithelium via simple passive diffusion, facilitated diffusion (mediated by a carrier, such as sodium), active transport, pinocytosis, or alternatively via a paracellular route (between cells).  The jejunum, along with the duodenum, is the site of absorption for most digested carbohydrates and proteins, while fats bound to bile salts, fat-soluble vitamins, as well as some fluids and electrolytes are absorbed in the ileum(21).  In contrast, the mucosal layer of the LI is quite distinct from that of the SI.  Plicae circulares and villi are completely absent at this site, with epithelial cells arranged exclusively within crypts.  Further, epithelial microvilli are much less pronounced than in the SI, and the majority of nutrient processing occurring at this site is dependent on the luminal bacteria, rather than specific enzymes present within the epithelium.  These differences likely reflect that the role of the LI is mainly in absorption of water and electrolytes in comparison to the SI, where there is a need for greater absorptive surface area and digestive capacity.      1.1.1.2 Distribution of IECs In addition to differences in histology and in the presence of digestive enzymes, the proportions of IEC subtypes within the mucosal layer also differ along segments of the lower GI tract. The majority (>80%) of cells comprising the SI epithelium are absorptive enterocytes(1, 8).  These cells are characterized by very long and numerous apical microvilli, their absence of secretory granules, and their localization along the top of crypts along with the entire villus.  In the LI, absorptive enterocytes are found at the top of crypts where they have migrated from the crypt 7  base, and they display less prominent apical microvilli than enterocytes in the SI. Secretory goblet cells, which create and maintain the intestinal mucus layer, constitute the second most numerous epithelial cell subtype, with an increase in their numbers observed from the jejunum to ileum (6% to 12%)(8).  Goblet cells are found scattered along the crypt-villus axis, from mid-crypt to villus tips and display an accumulation of mucus containing cytoplasmic granules.  They are more numerous in the proximal colon (16%) and most frequent in the distal colon (25%), where they are evenly distributed throughout the crypt, apart from the very top and base where they are not often observed(8).  Another secretory cell type present within the SI epithelium, located at the base of crypts and containing apical granules loaded with antimicrobial peptides (AMPs) and growth factors, is the Paneth cell(1, 8, 22). In contrast to goblet cells, which increase in number at more distal locations of the lower GI tract, Paneth Cells are numerous in the jejunum and ileum (~7%), but are completely absent in healthy large intestinal tissue. Moreover, several other cell types are found within the small and large intestinal epithelium in much smaller numbers including hormone producing enteroendocrine cells, antigen sampling microfold (M) cells that overlie lymphoid follicles, cup cells (described to date in guinea pigs, monkeys and rabbits), and tuft cells, which have recently been found to play a role in regulating host immune responses during parasitic infection(8, 23-26).   Coinciding with the increase in goblet cell numbers from the proximal to distal end of the lower GI tract, the mucus layer overlaying the intestinal epithelium also changes. In the jejunum and ileum, mucus exists as a loose layer that is not firmly attached to the underlying epithelium, whereas in the colon it forms a thick, adherent structure with two distinct layers(27).  The looser mucus layer found in the SI serves an antimicrobial role, forming a matrix within which AMPs 8  and antibodies produced at this site can adhere.  In the colon, the dense inner mucus layer is attached to the underlying epithelium and is at its thickest at the distal end (near the rectum), providing a charged gel barrier that is generally devoid of bacteria.  The outer mucus layer in the colon is unattached and similar to that found in the SI, with bacteria readily able to penetrate this layer.  Regional differences in lymphoid tissue, as well as innate and adaptive immune cells also exist along the GI tract(28), and although a thorough discussion on the topic is outside the scope of this thesis, there are several differences that are of interest to note.   1.1.1.3 Regional distribution of lymphoid tissues and immune cells Gut associated lymphoid tissues, including Peyer’s patches (PP) and isolated lymphoid follicles (ILF), are distributed in the mucosa and submucosa underlying the intestinal epithelium in the SI and LI, respectively(1, 28).  These sites comprise the primary locations for the induction of adaptive immune responses within the GI tract.  PP increase in number and size from the jejunum to the ileum and are overlaid by a follicle associated epithelium(28, 29). This specialized epithelial region primarily consists of M cells, is covered by a comparatively thin mucus layer, and lacks microvilli allowing the epithelium to directly interact with and transcytose bacteria, as well as other particles from the gut lumen across to their basolateral side(30). Subepithelial dendritic cells (DCs) are concentrated in this region and can process the transcytosed particles for antigen presentation to adaptive immune cells in the PP(28).  Similarly, the number of ILF in the LI increases along its length with the largest concentration found in the distal colon(28, 31).  Immune cells are also located throughout the lamina propria while specific subsets of T cells are found within the intestinal epithelium, with distinct differences between the two.  The lamina propria is host to a plethora of cells, including B cells, T cells, and several 9  innate immune cell populations such as DCs, macrophages and eosinophils. T cells, on the other hand, make up the majority of immune cells found within the epithelium (referred to as intraepithelial lymphocytes (IELs)) of mice and humans(1, 28).  These cells can be classified into two major groups: ‘induced’ (type a) IELs derived from conventional T cells, expressing an αβ T cell receptor (αβTCR) and cluster of differentiation (CD)8αβ heterodimer or CD4, or ‘natural’ (type b) IELs, which do not express CD4, but instead express a CD8αα homodimer and either a γδTCR or αβTCR(28).  IELs are more numerous in the SI than in the LI and decrease in number from the proximal to distal end of each(28, 32).  Type b IELs are more numerous in the murine SI, while type a IELs comprise the majority in the LI.  Conventional CD4 T cells can further be separated into several distinct types including T helper (Th)1, Th2, Th17, T regulatory (Treg), and Treg type 1 cells(1, 28).  These cell types can be distinguished by the expression of characteristic transcription factors and their cytokine production (summarized in Table 1.1) that can affect an array of cellular responses in various cell types.  In recent years, another important cell type, innate lymphoid cells (ILCs) have been discovered and researched (Table 1.1).  These cells have been found to be potent sources of many of the cytokines previously attributed solely to CD4 T cells in the intestinal immune compartment.        10  Cell Type Associated  Major Transcription Factors Cytokines Produced Th1 STAT1, STAT4, T-bet IFN-γ Th2 STAT6, GATA-3 IL-4. IL-5, IL-13 Th17 RORγt IL-6, IL-17A, IL-17F, IL-21, IL-22 Treg Foxp3 TGF-β, IL-10 Tr1 Foxp3- IL-10 ILC1 T-bet TNF, IFN-γ ILC2 GATA-3 IL-5, IL-13, IL-4, IL-6, IL-9, amphiregulin ILC3 RORγt, Ahr IL-17A, IL-22, TNF, GM-CSF Ahr:   aryl hydrocarbon receptor Foxp3:  forkhead box protein 3 GM-CSF:  granulocyte colony stimulating factor RORγt:  retinoic acid-related orphan receptor gamma t STAT:   signal transducer and activator of transcription T-bet:   T-box transcription factor  Table 1.1:  Conventional CD4 T cell and ILC subsets found in the intestine Summary of CD4+ T cell and ILC subsets, along with their associated transcription factors and cytokines   Due to the variation in the mucus layer and regional differences in host immunity, along with several other factors including nutrient availability, transit time, acidity, oxygen levels, and the presence of bile acids and antimicrobials, the diversity and number of bacteria along the length of the GI tract greatly varies. The next section of this thesis will focus on our current understanding of the intestinal microbiota and their implications for host health.    1.1.2 Intestinal bacteria Of the microbes present in the lower GI tract, including eukaryotes, archaea, bacteria and viruses, bacteria constitute that largest portion of this population and are composed of at least 1000 different species.  The rapid colonization of mucosal sites, including the GI tract, occurs at 11  birth and is initiated by the bacteria present at the sites of first contact with the newborn, which varies depending on mode of delivery.  With vaginal birth, babies are first colonized with vaginal and fecal bacteria, including Lactobacillus and Bifidobacterium.  In comparison, cesarean birth results in initial colonization by bacteria from the skin and hospital environment, such as Staphylococcus and Acinetobacter(33, 34).   These resident bacteria provide an arsenal of digestive enzymes that are not found within the host genome, such as glycoside hydrolase and polysaccharide lyase, which are able to breakdown complex carbohydrates otherwise indigestible by the host(35).  This increases digestive efficiency, as is evidenced in early studies on germ free (GF) rats that required a 30% increase in caloric intake to maintain body weight as compared to conventionally raised animals(36).  Intestinal bacteria are also able to synthesize essential nutrients for the host, such as B-group vitamins and vitamin K(35).  Furthermore, microbes occupy specific niches along the GI tract and, in a healthy gut, are able to inhibit colonization by invading enteric pathogens.  Use of GF mice as well as animals that have been treated with antibiotics have revealed increased susceptibility to several enteric bacterial pathogens including Shigella flexneri, Clostridium difficile, Salmonella enteritidis, enterohaemorrhagic Escherichia coli and Citrobacter rodentium in the absence (or following the reduction) of intestinal commensal bacteria(37-43).   Studies examining the microbiota in the first three years of life have found that bacterial diversity increases with age(2, 44-47).  From birth to a few days following, the intestinal lumen turns from an aerobic environment to largely anaerobic due to the expansion of the resident bacteria and their consumption of oxygen(2).  The bacteria able to grow under these changing conditions also shift from exclusively facultative anaerobes, such as members of the 12  Enterobacteriaceae family, to include obligate anaerobes such as Bifidobacteria, Clostridia and Bacteroides(2, 48).  In the following four to six months, the composition of the intestinal bacteria is greatly affected by the available food sources, with a higher proportion of Bifidobacteria and Lactobacilli in breast milk fed infants as compared to higher Clostridia, Bacteroides, and Streptococci in formula fed babies(2, 49-51). The introduction of solid food further impacts the makeup of the intestinal bacteria.  Several other environmental factors such as contact with the mother and other children, introduction of hands, feet and other objects into the mouth, and hygiene measures, also influence shifts in intestinal bacterial populations until around three years of age(46).  Around this time, intestinal bacterial composition stabilizes, leaving members of the Bacteroidetes and Firmicutes comprising the majority of gut microbes, along with minor portions of Verrucomicrobia, Actinobacteria and Proteobacteria(52, 53).  This succession of colonization in the infant intestine is essential in promoting proper gut development (discussed further in section 1.5), as well as for the education of the immune system.  Moreover, microbial colonization may also have ramifications later in life in terms of regulating susceptibility to the development of asthma, allergies and other inflammatory conditions(2, 34, 54).  Interestingly, early development is not the only timepoint during which dynamic changes within the makeup of the intestinal bacteria precede its stabilization are observed.  Intestinal disturbances in adult life, such as enteric pathogen infection, the introduction of antibiotics, or significant dietary changes can also lead to changes in intestinal bacterial composition.      1.1.2.1 Host factors influencing bacterial colonization in the intestine In the lumen of the healthy adult intestine, the number and types of microbes vary in different regions.  As mentioned previously, this is mostly due to changes in the mucosal and luminal 13  environment along its length.  In the SI, bacterial density is limited by the presence of Paneth cells (resulting in a greater concentration of AMPs), as well as bile salts(53, 55).  This results in bacterial concentrations of <105 microbes per mL in the jejunum and ≤107 in the ileum, as compared to the LI, where numbers can reach up to 1012 microbes per mL(53, 55). Moreover, the shorter transit time of luminal contents through the SI, regardless of its much greater length than the LI, results in an environment favouring more persistent colonization by species that are able to adhere to the loose mucus layer or underlying tissue.  Fast growing bacteria that are mostly facultative anaerobes, competing for simple carbohydrates abundant in the SI lumen, such as members of the Lactobacillaceae and Enterobacteriaceae families are able to better tolerate the AMP and bile salt concentrations in these regions(56, 57).  The diversity of bacteria, along with the total number of bacteria present, increases in the terminal ileum and LI.  The bacteria at these sites are responsible for the breakdown of indigestible food products, such as resistant polysaccharides, that are not broken down in the SI(56).  In addition, there are fewer AMPs, a denser and thicker mucus layer, differences in food sources (complex vs. simple carbohydrates), and slower transit time through this region.  Bacteria from the Bacteroidaceae and Clostridiaceae families, which are fermentative polysaccharide-degrading anaerobes, are the most abundant in the LI(56, 57).   1.1.2.2 Impact of bacterial factors on host cells Just as host factors are able to impact bacterial concentrations and diversity along the GI tract, interactions between host cells (IECs and immune cells) and bacteria, and the nutrients and metabolites produced by them, can in turn influence host cell growth and activation states.  Extensive studies examining the role of microbe-associated molecule pattern (MAMP) sensing 14  host receptors, such as Toll like receptors (TLRs) and nucleotide-binding oligomerization domain (NOD) like receptors, in maintaining intestinal homeostasis have been done and will be covered in further detail in section 1.3.  In addition, bacterial fermentation of dietary fibers produces short chain fatty acids (SCFA), such as butyrate, acetate and propionate(58-60), which can act on IECs and immune cells(61-64).  MAMPs and SCFAs have been hypothesized as major factors driving the differences observed in the GI tract of GF mice, as compared to conventionally raised mice.  Differences observed in GF mice include altered mucus layers that are penetrable in the LI and unable to concentrate AMPs in the SI, enlarged ceca, decreased SI and LI crypt depths, and decreased abundance of intestinal immune cell types(65, 66).       With regards to immune function, SCFA have been hypothesized to exert their effects through several pathways including activation of G protein coupled receptors (GPRs), inhibition of histone deacetylases, and autophagy regulation(61, 62, 67-72). For instance, propionate and acetate are able to induce neutrophil chemotactic responses through the activation of GPR43(73, 74), with an associated increase in intracellular calcium.  Introduction of acetate to neutrophils was further able to modulate reactive oxygen species production and phagocytic activity, although this effect was absent in GPR43 deficient (-/-) neutrophils(73).  In addition, SCFA are also able to impact the function of T cells, macrophages, DCs and IECs(75, 76).  Stimulation of T cells by SCFA induces increases in apoptosis and a decrease in proliferation(77-80).  In macrophages and DCs, SCFA suppress the production of inflammatory cytokines (such as tumor necrosis factor (TNF) and interferon (IFN)-γ) as well as chemotactic factors(28, 75, 76). Finally in IECs, in addition to serving as an energy source, SCFAs decrease autophagy(62) as well as the 15  expression of the neutrophil chemokine interleukin (IL)-8 (keratinocyte-derived chemokine (KC) in mice) (61), and suppress intestinal stem cell proliferation(81).  The vast numbers of resident intestinal bacteria require a highly specialized barrier to maintain intestinal homeostasis. Though the immune cells of the intestine are important in maintaining a tolerant relationship between the resident bacteria and host intestinal tissue, there is a growing appreciation for the contribution of the cell types in closest contact with these organisms, the IECs, in these processes.  IECs have the remarkable ability to respond to infection or perturbation of the mucosal layer, while also maintaining an environment permissive to commensal microbe colonization by integrating microbial signals through innate immune signalling pathways.  1.2 Intestinal epithelial barrier and cell subtypes The intestinal epithelial barrier represents the body’s largest interface with the external environment and the microbial world(82-84).  Only a single cell layer in thickness, it is in a unique position to provide physical separation of the underlying immune system and host tissue from the entirety of the luminal contents, while facilitating absorption of nutrients. As such, the intestinal epithelial barrier has the remarkable responsibility of initiating intrinsic responses to infection or perturbation of its single cell layer, while maintaining tolerance to the trillions of microorganisms and food antigens found in such close proximity. Though this homeostasis is achieved in concert with lymphoid, myeloid and stromal cells, there are many key features of the barrier itself, along with the specialized epithelial cells found within it, that optimize this role.   16  1.2.1 Tight junctions and the intestinal barrier The intestinal epithelium maintains permeability to ions and fluids while forming a barrier against potentially harmful luminal materials including bacteria (and their associated products), undigested food components, and environmental pollutants.  This balance is maintained through several structures generated between adjacent epithelial cells, the most apical being tight junctions (TJs), followed by adherens junctions, and finally desmosomes(85, 86) (Figure 1.1A).  TJs are composed of several groups of proteins including occludins, members of the claudin family, and junction-adhesion molecules.  Occludins play a key role in the stability of TJs and in maintaining barrier integrity by acting as anchors between transmembrane claudin proteins and actin filaments of neighboring cell’s cytoskeletons(87, 88). Peripheral membrane proteins referred to as zonula occludens (ZO) act as scaffolds between the cytoskeletal components and the proteins of the TJ complex(89).  These transmembrane TJ proteins form a selective seal in the paracellular space between adjacent cells limiting solute flux, therefore acting as the main determinants of epithelial barrier permeability(85, 90, 91).  Additionally, they play a crucial role in regulating cell polarity by preventing membrane receptors on the apical surface of the epithelial cell from diffusing to the basal membrane, which is oriented away from the lumen(92, 93).  Underlying the TJs at the apical regions of the epithelial cell are adherens junctions.  Adherens junctions, along with desmosomes, provide strong bonds between adjacent cells and keep them in close proximity to one another while also facilitating cell-to-cell communication(85).  Adherens junctions, composed of cadherin and catenin proteins, are connected to actin filaments of the cell’s cytoplasm, serving as anchors(85, 94).  In contrast, desmosomes anchor adjacent cells by interacting with intermediate filaments of the 17  cytoskeleton(85). Collectively, TJs, adherens junctions, and desmosomes form a structure referred to as the apical junction complex.               The proteins comprising TJs can vary in composition depending on the region of the intestine, as well as the stage of development, and whether the tissue is in a homeostatic versus an inflamed state(85, 86, 88, 90, 95, 96). There are 24 members of the claudin family of proteins that have been described in mice and humans, with at least 7 expressed abundantly along the GI tract in distinct patterns(86, 97, 98) (Figure 1.1B).  For instance, Claudins -2 and -3 display greater expression in the ileum and colon, as compared to the jejunum, whereas Claudin-4 is more highly expressed in the jejunum and ileum than the colon.  Localization of claudin protein expression along the crypt-villus axis also varies along the GI tract (e.g. Claudin-4 expression is restricted to the tops of villi in SI tissue, whereas in the colon it is expressed only along the tops of crypts).  Expression and localization of various claudin proteins during murine intestinal development has also been reported(95).  At days 1, 14, 28 and 90 after birth, several claudins showed major fluctuations in expression and changes in localization along the crypt-villus axis. Interestingly, expression and localization of other TJ components including ZO-1, occludins and junction-adhesion molecules remained unchanged over this same period.  It is hypothesized that charge and size selectivity of the different claudin proteins may, in part, explain their differential expression during development, and along the length of the adult GI tract(86, 95).  Components of the TJ can also be altered under inflammatory conditions(88, 90).  This is evidenced in IBD, as well as in some models of enteric infection, where internalization of TJ proteins, rearrangements in the actin cytoskeleton to which they are anchored, or changes in their gene expression can be initiated by exposure to cytokines such as TNF-α and IFN-γ leading to a 18  disruption of barrier integrity(96, 99-103).  Further, loss or alteration in other components of the apical junction complex, such as the adherens junction, can lead to disruptions in cell-to-cell contacts, perturbations in cell polarization and differentiation, and premature cell death.                    Figure 1.1: Epithelial tight junctions and Claudin distribution along the length of the intestine A) Diagram depicting the arrangement of junctions between polarized epithelial cells.  The apical junction complex is comprised of the tight junction, adherens junction, and desmosome. B) Schematic distribution of Claudins 2, 3, 4, 7, 8, 12 and 15 in regions of the SI and colon. (Image reproduced from (85) and (86) with permission)  Together, the components of the apical junction complex form a highly regulated, selectively permeable barrier that is effective at blocking most microbes and other antigens found within the lumen, from reaching the underlying intestinal tissues and activating resident immune cells.     A B Duodenum  Jejunum       Ileum        Colon  Cldn2 Cldn3 Cldn4 Cldn7 Cldn8 Cldn12 Cldn15 19  1.2.2 Intestinal epithelial cell lineages In addition to the barrier that is formed between epithelial cells by the apical junction complex, a region rich in secreted factors such as mucins and AMPs overlies the intestinal epithelium along the lower GI tract.  These factors are produced by several specialized epithelial cell types found within the single cell layer, and are able to limit direct contact with potential damaging stimuli found in the intestinal lumen(82-84, 104).  There are four distinct terminally differentiated cell types that comprise the majority of the intestinal epithelium throughout the SI that can be divided into two classes: absorptive and secretory cells(1, 8, 28, 104).  Absorptive enterocytes are the most numerous cell type present, followed by the secretory cell types including goblet cells, Paneth cells, and enteroendocrine cells.  In the LI, Paneth cells are absent under normal conditions; therefore the epithelium at this site is composed primarily of the remaining three terminally differentiated cell types.  Limited numbers of other specialized cell types are also present (<1%), including M cells and tuft cells, which are involved in luminal antigen sampling and in mounting responses to parasitic helminth infections, respectively(1, 8, 23).      Along the entirety of the lower GI tract enterocytes, goblet cells, Paneth cells, and enteroendocrine cells arise from pluripotent intestinal epithelial stem cells located at the base of crypts, where stromal and hematopoietic cells maintain a stem cell niche(8, 105).  These cells also referred to as crypt base columnar (CBC) stem cells, have been identified as Leucine-rich repeat-containing GPR 5 positive (Lgr5+) stem cells(105-107), which has allowed more thorough investigation of this cell type in recent years.  Asymmetric division of Lgr5+ CBC cells gives rise to one stem cell and one transit amplifying daughter cell, which rapidly proliferates and matures as it migrates up the crypt.  The only exception to this upwards migration is the 20  Paneth cell, which migrates towards the bottom of the crypt instead(22, 105, 108, 109). As cells leave the stem cell niche they progressively differentiate until they become irreversibly mature.  Maintenance of the stem cell niche as well as differentiation of transit amplifying cells into the absorptive or secretory cell types is controlled by the highly conserved Wnt and Notch developmental signalling pathways(104, 105, 107, 110, 111). Subepithelial myofibroblasts, along with Paneth cells in the SI, are hypothesized as the main sources of Wnt ligands in the intestine, which are highly expressed in close proximity to the stem cell niche (Figure 1.2A).  Wnt ligands bind to Frizzled and the Lrp receptor complexes, thereby inducing target genes that regulate proliferation and maintenance of the undifferentiated status of CBC cells (Figure 1.2A).  Further, R-spondin stimulation of Lgr5 interrupts negative feedback mechanisms and enables sustained activation of Wnt signalling. As a result, increased numbers of CBC cells as well as crypt hyperplasia, i.e. elongation of crypts, are observed in R-spondin treated mice(112).    In contrast, CBC cell numbers are negatively regulated by bone morphogenetic proteins (BMPs), which are growth factors belonging to the transforming growth factor (TGF)-β superfamily that induce epithelial cell differentiation(105, 107, 113-115).  A recent study examining the effects of BMPs revealed their ability to constrain CBC cell numbers by transcriptional repression of several signature stem cell genes, including Lgr5(113).  This effect was achieved through phosphorylation and nuclear translocation of Smad proteins, which are key regulators of gene expression.  Interestingly, similar to Wnt ligands, a gradient for some BMP has been described within the intestinal epithelium(114).  BMP2, which is produced by enterocytes within the epithelium, is most highly expressed by mature enterocytes at the tops of crypts.  These increasing BMP concentrations up the crypt–villus axis, as well as transcriptional activation of 21  specific target genes of the Notch pathway facilitates differentiation into the absorptive and secretory cell types (Figure 1.2B).  To counteract the stem cell repressive actions of BMP, several proteins concentrated at the base of crypts, such as Noggin and Gremlin, are able to bind and inhibit BMP with high affinity, thereby maintaining the stem cell niche(105, 116, 117) (Figure 1.2A). 22    Figure 1.2:  Maintenance of the stem cell niche and epithelial cell lineage commitment in the intestine A B 23  Figure 1.2:  Maintenance of the stem cell niche and epithelial lineage commitment in the intestine A) Paneth cells and subepithelial myofibroblasts produce Wnt ligands in the SI to maintain the stem cell nice. B) Overview of epithelial cell lineage commitment in the intestine. (Image reproduced from (107) and (105) with permission)  Resident stem cells at the crypt base allow for continuous and rapid renewal of the intestinal epithelial cell layer, up to 90% of which is replaced every 2-3 days in mice and within 4-5 days in humans(8, 118). In contrast to the other three epithelial cell types, Paneth cells in the SI are long-lived (>3 weeks) differentiated cells. Under normal conditions, IECs undergo cell death and shedding into the gut lumen to facilitate cell renewal and turnover of terminally differentiated cells(119).  During infection, or following specific inflammatory stimuli, increased turnover and IEC proliferation, apoptosis and shedding into the intestinal lumen may be observed.  Continuous turnover of IECs can serve as a defense strategy and similarly, epithelial cell proliferation is increased during infection(82-84, 120, 121).  This process keeps pathogens from close contact with intestinal crypts and stem cells as it increases the rate of IEC sloughing, along with any pathogens that may have already infected those cells.  An increase in IEC apoptosis and shedding at the tops of crypts requires an increase in intestinal stem cell proliferation and IEC migration up the crypt axis to maintain the integrity of the barrier.  The regulation behind this phenomenon is poorly understood, and appears to often lead to greater proliferation than required to replace sloughed cells, resulting in the characteristic elongated crypts, or hyperplasia, observed during many forms of intestinal infection and inflammation. Although TLR signalling is involved in the induction of IEC proliferative responses, many cytokines produced by immune cells, of note IL-22, are also potent inducers of this response(122, 123).           24  1.2.2.1 Absorptive cell types Enterocytes are the sole absorptive cell type of the intestinal epithelium and make up the majority of the cells found within this layer.  Besides serving an essential role in absorption of nutrients, enterocytes are also capable of producing several factors that contribute to maintaining intestinal homeostasis(82-84, 104).  Though Paneth cells are the primary producers of AMPs, enterocytes also produce several including β-defensins, cathelicidins, and regenerating islet-derived protein (Reg)IIIγ.  Enterocytes can also produce and release chemotactic factors such as IL-8  (or KC)(61, 124) as well as macrophage inflammatory protein (MIP)3α(125, 126), anti-inflammatory mediators such as prostaglandin(127, 128), an array of cytokines, and metalloproteases(129).  In addition to producing these factors under baseline conditions, enterocyte activation and upregulation of the aforementioned products occurs during enteric infection and inflammation (further discussed in section 1.3.2).      1.2.2.2 Secretory cell types As discussed above, secretory cell types of the intestinal epithelium include goblet cells (covered in detail in section 1.2.3), Paneth cells, and enteroendocrine cells.  Commitment towards a secretory cell type requires expression of the Math1 transcription factor, as Math1-/- mice contain only enterocytes within their intestinal epithelium(130, 131).  Paneth cells have apical granules packed full of proteins supported by an extensive endoplasmic reticulum (ER) and Golgi network. The antimicrobials released into the crypt lumen via exocytosis from Paneth cell granules include AMPs that can target different types of bacteria.(22, 132, 133)  Some AMPs work more broadly and can target both Gram- negative and positive bacteria such as α-defensins (cryptdins in mice) and lysozyme(133).  Other AMPs target Gram-positive bacteria alone, 25  including the C-type lectin RegIIIα (RegIIIγ in mice) and secretory phospholipase A2 (133).  Mouse Paneth cell granules contain additional AMPs such as angiogenin 4(134).  Granule release is triggered by a variety of stimuli.  Cholinergic agonists, such as carbamylcholine, released by the enteric nervous system, inflammatory cytokines (e.g. IFN-γ), the presence of bacteria, and bacterial products such as cell wall components lipopolysaccharide (LPS) and lipoteichoic acid can all stimulate granule release(135-138).  As many of these stimuli are present in the healthy intestinal environment, it is likely that Paneth cell granule release occurs at a low baseline rate and is upregulated in response to increased stimulation, as likely occurs during enteric infections. Defects in Paneth cells have been observed in several diseased states including Crohn’s Disease (one of the two main forms of IBD)(139-141) and Necrotizing Enterocolitis (NEC)(142, 143), a disease characterized by uncontrolled inflammation and damage in regions of the premature infant intestine, often resulting in the necrosis of affected intestinal tissues.  The final secretory IEC type, enteroendocrine cells, are well known for their production of hormones such as cholecystokinin and 5-hydroxytryptamine (5-HT; serotonin) that influence digestive processes(144).  5-HT, though commonly referred to as the “happiness hormone” is most highly expressed in the GI tract where it regulates intestinal motility, among other functions(145).  In addition to this role, enteroendocrine cells are also able to respond to the TLR ligands LPS and flagellin (proteins composing the flagella of motile bacteria) both in vitro and in vivo, resulting in the production of the AMP β-defensin2, and the neutrophil attractant KC(146).          26  1.2.3 Goblet cell secreted proteins and the mucus layer Goblet cells are the most numerous secretory IEC type found along the GI tract and are responsible for the secretion of mucins providing an additional layer of separation between the intestinal epithelium and gut luminal contents(27, 147, 148). As discussed in section 1.1.1.2, the mucus layer is not uniform along the length of the GI tract.  The properties of intestinal mucus and the mechanisms through which these regional differences are achieved will be further discussed here.    Goblet cells are named for their cup-like (goblet) shape formed by the concentration of secretory granules within their cytoplasm (Figure 1.3).  Granules are packed full of the large Muc2 glycoprotein, with a loss of its expression leading to an absence of full granules and goblet cell shape(149).  Muc2, the major gel-forming mucin comprising the intestinal mucus layer, is a large glycoprotein that dimerizes, via its C-terminus, in the ER and becomes highly glycosylated within the Golgi where it also trimerizes via its N-terminus(150-152).  These large polymers are packed tightly into apical secretory granules facilitated by high calcium levels and low pH(153).  Upon secretion, the densely packed polymers expand (>1000 fold) into large net-like structures, which is dependent on an increase in pH outside of the cell(154).  In addition, there are several transmembrane mucin proteins (Muc1, Muc3, Muc4, Muc13 and Muc17) expressed at the apical membrane of both absorptive and secretory IECs, which are structurally distinct from Muc2(155, 156).             In the SI, although the mucus is present in a loose layer suggesting it may be easily penetrable by bacteria, it in fact creates a largely sterile zone allowing the AMPs released by Paneth Cells to 27  reach concentrated levels at the crypt base and along the crypt-villus axis(27, 157, 158).  By trapping AMPs in such close proximity to the epithelium, mucus in the SI restricts their spread and dilution into the intestinal lumen.  Though Muc2 is a transmembrane protein, cleavage by the metalloprotease meprin β in the SI separates Muc2 from the goblet cell(159).  As luminal contents move down the intestinal tract through peristalsis the loose SI mucus layer in which many bacteria are sequestered moves towards the LI, aiding in flushing of bacteria down the GI tract.  In contrast, Muc2 in the LI remains anchored to goblet cells forming the inner mucus layer with small pore sizes that inhibit penetration by bacteria, mediating a layer of separation between the IEC barrier and luminal intestinal bacteria(160).  It is currently hypothesized that endogenous proteases reside at the periphery of the inner mucus layer, with their proteolytic cleavage of Muc2 responsible for the transformation of the firm inner mucus into the outer loose layer.     28   Figure 1.3:  Muc2 structure and assembly within the goblet cell and upon luminal secretion Diagram of the assembly and structure of Muc2 within the endoplasmic reticulum (ER), Golgi, trans-Golgi network (TGN), secretory granule and lumen.  (Image reproduced from (27) with permission)  Not surprisingly, lack of Muc2 (i.e. Muc2-/-) leads to the loss of almost all intestinal mucus, as well as increased direct interactions of bacteria with IECs, ultimately leading to the development of spontaneous intestinal inflammation(149, 160-162).  In addition, animals with a penetrable inner mucus layer also develop spontaneous gut inflammation(163).  Bacterial penetration of intestinal mucus layers is also observed in patients with chronic intestinal inflammation, such as Ulcerative Colitis, as compared to control individuals or patients in remission(163).  Goblet cells 29  also produce several other factors that, in addition to the AMPs (β-defensin and lysozymes) produced by Paneth cells and secretory immunoglobulin A from enterocytes, are concentrated within the intestinal mucus layer.  Additional goblet cell factors include trefoil factor 3 (TFF3), which is a barrier protective factor promoting migration of IECs to repair breaches in the epithelial layer(164), Fc-γ binding protein, which binds immunoglobulin G(165, 166), and resistin-like molecule (Relm)β, which is implicated in immunity to nematodes, induction of Paneth cell specific AMPs, and recruitment of CD4+ T cells to the colon during enteric infection(167-172).   The specialized IEC types comprising the epithelium, described above, through the production and release of their specific factors contribute to the maintenance of the intestinal barrier.  The factors they produce can be influenced by modulation from resident immune cell derived factors, but there is growing evidence that IEC-intrinsic signalling also plays an active role in these processes. In the next section of this thesis, the capacity of IECs to act as functional cells of the innate immune system will be discussed.  1.3 Innate immunity in intestinal epithelial cells In addition to the aforementioned factors, effective maintenance of the intestinal barrier also relies on the ability of IECs to initiate functional responses via innate immune signalling pathways under both homeostatic and inflammatory conditions.  In fact, microbial sensing via PRRs on IECs is able to influence several of these barrier-strengthening components, such as modulation of TJ proteins, AMP release, and increased expression of goblet cell factors(137, 158, 170, 173-175).  Immune cells, of course, play an integral role during infection and 30  inflammation. However, in concert with these cells, IEC-intrinsic innate immune signalling plays a key role in mounting appropriate responses and maintaining the intestinal barrier. In the studies outlined in this thesis, one of my primary goals was to examine one specific innate immune pathway, Myeloid differentiation primary response gene (MyD)88 dependent signalling, in IECs and its contribution to the maintenance of a functional gut barrier during enteric infection.   1.3.1 MyD88 dependent signalling pathways MyD88 is an integral adaptor protein utilized by the majority of receptors in the IL-1 receptor (IL-1R) superfamily to initiate their signalling(176-178).  These receptors include all of the TLRs, with the exception of TLR3, which are activated by a variety of MAMPs, as well as the IL-1 and IL-18 receptors, which respond to their respective cytokine ligands.  All members of this receptor superfamily are transmembrane in nature containing an extracellular ligand binding domain and a single membrane-spanning region linked to a cytoplasmic component known as the Toll/IL-1R (TIR) domain.  Activation of receptors by their respective ligands results in dimerization of TIR domains, which allows interaction with the TIR domain of MyD88, or in certain instances in concert with other TIR containing adaptor molecules (described in section 1.3.1.2).  MyD88 stimulation leads to recruitment and phosphorylation of IL-1R associated kinases (IRAK) and its subsequent association with TNF receptor associated factor (TRAF)6, creating a large receptor complex.  IRAK-1 and TRAF6 dissociation from the receptor complex recruits TGF-β-associated kinase (TAK)1 to TRAF6 along with the TAK1 binding proteins TAB1 and TAB2. Activated TAK1 in turn activates the Nuclear Factor (NF)-κB essential modulator (NEMO) complex leading to Inhibitor of-κB (IκB) degradation and release of the NF-κB p65 subunit from the cytoplasm into the nucleus. Once in the nucleus, the p65 subunit 31  induces transcription of NF-κB dependent genes.  TAK1 is also able to activate mitogen activated protein kinases (MAPK), which can further induce genes dependent on the activator protein (AP)-1 transcription factor (Figure 1.4).  The end result of transcription factor induction by microbial sensing TLRs (within IECs) is the production of several cell signalling molecules as well as the modulation of a variety of cellular processes.     Figure 1.4:  MyD88 dependent signalling pathway Activation of specific TLRs and IL-1Rs results in initiation of MyD88 dependent downstream signalling cascades.  Ultimately this leads to the translocation of transcription factors such as NF-κΒ and AP-1 into the nucleus.  32  The role of NF-κB signalling in controlling a variety of pathways within IECs, in addition to its classically appreciated involvement in pro-inflammatory responses, was highlighted by studies using mice lacking NEMO specifically within their IECs (IEC-NEMO-/-)(179).  These mice developed spontaneous and severe intestinal inflammation.  Further characterization revealed massive colonic IEC death, bacterial penetration of the epithelial barrier, and reduced production of AMPs.  Inflammation was perpetuated by the early accumulation of innate immune cells and then T cells at later time points.     1.3.1.1 Interleukin-1 receptors Members of the IL-1R subgroup of the IL-1R superfamily respond to cytokine ligands rather than MAMPs, such as IL-1α and β, IL-18 and IL-33(180, 181).  Though there are currently 10 identified receptors expressed by an array of cell types throughout the body, IL-1R1, IL-1R8 (or Single Immunoglobulin (Ig) IL-1R Related Molecule - SIGIRR) and IL-18R are the ones predominantly expressed by IECs(182).  IL-1Rs, with the exception of SIGIRR, contain three extracellular Ig domains and heterodimerize upon ligand binding to initiate MyD88 dependent signalling.  SIGIRR, on the other hand, inhibits MyD88 dependent signalling (of both IL-1Rs and TLRs) by sequestering IRAK and TRAF6 or through formation of a ligand-receptor complex with IL-37(183).  Its absence in vivo leads to extensive damage during both dextran sodium sulfate (DSS) as well as infection induced intestinal inflammation(184, 185).  Moreover, SIGIRR expression is decreased in inflamed intestinal tissues, such as those from Ulcerative Colitis patients (one of the two main forms of IBD)(186).  Through its inhibitory effects SIGIRR, along with other negative regulators of IL-1R/TLRs, helps maintain intestinal homeostasis through dampening potential inflammatory responses to intestinal bacteria by IECs.     33   The cytokines that activate the IL-1R are produced by a variety of cell types. In the intestine IECs and immune cells (including DCs and macrophages) can produce IL-1α, IL-1β, and IL-18, while IL-33 is produced by endothelial cells, smooth muscle cells, fibroblasts and IECs(187).  IL-1β and IL-18 are secreted as inert propeptides and require cleavage of their N-terminal by the cysteine protease caspase-1, which in turn is dependent on activation by inflammasomes.  IL-1β is a potent inflammatory cytokine with a plethora of functions including activation and survival of T cells, skewing T cells to a Th17 type (in concert with other cytokines), activation of effector functions of macrophages, DCs and neutrophils, and induction of IEC chemokine expression(188-193).  IL-18 can also influence T cells by inducing development and activation of the Th1 subset, along with activation of NK cells(194).  With regards to IECs, IL-18 has been found to induce proliferation, and expression of chemokines and AMPs in vitro.  However, its role in vivo has been more difficult to discern as conflicting results have been reported. During DSS induced colitis in mice unable to produce active IL-18, administration of exogenous IL-18 is protective against intestinal damage and weight loss(195, 196).  In contrast, sequestering of IL-18 (by administration of neutralizing antibodies or the inhibitory IL-18 binding protein) can also lead to protection from DSS induced intestinal inflammation and damage(197, 198).   1.3.1.2 Toll-like receptors The protein structures of TLRs are characterized by an extracellular domain containing leucine-rich repeats, resulting in their characteristic horseshoe shape(176, 178, 199).  The human genome encodes ten TLRs (TLR1 to 10), while there are twelve reported in mice (TLR 1 to 9, and 11 to 13).  Each TLR responds to specific ligands derived from bacteria, viruses, parasitic worms or 34  fungi, usually in the form of lipids, proteins, lipoproteins, or nucleic acids.  For the purposes of this thesis, only those TLRs recognizing bacterial ligands will be described further.    TLRs 2, 4, 5, and 9 recognize various bacterial ligands (Figure 1.5) and are expressed by IECs, as determined by RNA and/or immunohistochemical analysis of IECs isolated via laser capture microdissection(200, 201).  TLR2 is located at the cell membrane surface and activated by several ligands derived from bacterial cell wall components including di- and tri- acylated lipoproteins, peptidoglycan, and lipoteichoic acid. Formation of heterodimers with TLR1 or 6 confers some ligand specificity with TLR2/1 detecting triacylated liproteins from Gram-negative bacteria, and TLR2/6 activated by diacylated lipoproteins from Gram-positive bacteria.  TLR2/6, as well as TLR4, require an additional adaptor protein to initiate downstream signalling referred to as MyD88-adaptor-like (or TIRAP), which links the activated receptor to MyD88.  TLR4 is another cell membrane surface receptor that recognizes the gram-negative cell wall component LPS. TLR4 forms a complex with MD2 to bind LPS at the cell surface, a process assisted by the soluble LPS binding protein or the TLR4-MD2 co-receptor CD14 on macrophages. TLR5 recognizes flagellin, which comprises the subunit structural proteins of the flagella of motile bacteria.  Finally, TLR9 responds to a component of bacterial deoxyribose nucleic acid (DNA) that is distinct from that of host genomes, namely unmethylated cytidine-phosphate-guanosine (CpG).    Though common in bacterial and viral genomes, unmethylated CpG motifs are rare in mammalian DNA.       35   Figure 1.5:  Toll-like Receptors and their ligands TLRs 2 (in association with 1 or 6), 4, 5 and 9 have been characterized to recognize bacterial components.   In the intestine, TLRs are expressed by macrophages, DCs, T and B cells, IECs, and stromal cells, although their levels of expression can be significantly modulated, for example, in IECs. Under normal conditions, TLRs 2 and 4 are expressed under low levels in IECs, however exposure of IEC to specific cytokines has been shown to impact their expression(200, 201).  For instance, expression of TLR4 and its co-receptor MD-2 can be induced by TNF-α and IFN-γ, whereas the Th2 cytokines IL-4 and IL-13 can decrease IEC responsiveness to LPS(202, 203).  Using GF mice, the impact of bacteria on IEC TLR expression has also been examined, revealing induction of TLR2, 4 and 5, along with apical expression of TLR9 on IEC being dependent on the presence of bacteria(204).  Some TLRs, such as TLRs 2, 4, 5, and 9, can be expressed on the apical and/or basolateral membranes of IECs.  This can lead to induction of differential responses by IEC, based on the location (apical versus basolateral) of receptor engagement.  The best characterized example of this is TLR9, where apical ligand binding results in dampening of the NF-κB response within IEC, presumably to limit pro-inflammatory 36  signalling in response to resident intestinal bacteria(205).  In contrast, basolateral TLR9 activation strongly stimulates pro-inflammatory chemokine induction via NF-κB.  Given the impact of cytokine regulation on TLR levels, it is not surprising that although expression by IEC of some TLRs under healthy conditions may be low, intestinal inflammation can lead to their increase; as is evidenced by increased TLR4 expression in IECs of IBD patients as compared to control individuals.    Activation of TLR signalling within IECs can modulate a variety of cellular signalling pathways.  In enteroendocrine cells, which are reported to express TLRs 1, 2 and 4 in vivo, ligand binding to specific TLRs may result in the secretion of different hormones.  For example, stimulation of enteroendocrine cell lines with LPS or flagellin and CpG results in the production of either somatostatin or cholecystokinin, respectively(146, 206). As these hormones can have effects on intestinal motility, this may have implications for how the host modulates the diarrheal response to enteric pathogens to aid in their elimination.  Moreover, activation of TLR2 stimulates the production of the barrier protective protein TFF3 by goblet cells, as well as the redistribution of the TJ component ZO-1 to enhance stabilization of the junctional complex between IECs and decrease intestinal permeability(173, 174, 207).  TLR4 and 5 signalling in IEC induces chemokine expression, in addition to increased TNF production, apoptosis, and phagocytosis and translocation of bacteria with TLR4 activation of enterocytes in vitro (208).  TLR9 ligand binding also leads to the degranulation and release of AMPs by Paneth Cells(209), in addition to its role in maintaining tolerance to resident bacteria as described above.                        37  Further, sensing of MAMPs from specific bacteria by various cell types can skew the host response towards certain immune cell types and alter immune receptor expression levels.  For instance, the presence of the resident bacterium Clostridium butyricum has been reported to drive TLR2-dependant induction of anti-inflammatory IL-10 producing macrophages, and decreased TLR4 expression by colonic IECs induced by bacterial production of SCFAs(210, 211). It is also now appreciated that the close interactions that occur between resident Segmented Filamentous Bacteria and IECs in the terminal ileum results in the production of serum amyloid A, which can act on DCs in the lamina propria, leading them to promote Th17 cell differentiation(212).  In addition to serum amyloid A, TLR signalling in IECs produces a variety of factors that in turn can influence classical immune cells.  These factors include, but are not limited to, thymic stromal lymphopoietin, TGF-β, IL-10, a proliferation-inducing ligand, and B cell activating factor(201).  Many of these factors can act on sentinel immune cells, such as DCs and macrophages in the intestinal lamina propria to maintain anti-inflammatory signalling and/or limit host immune responses to resident intestinal bacteria.   1.3.2 Innate immune responses to infection Initially, innate immune responses to infection were expected to function solely in deterring pathogen colonization and clearance of infections through the induction of pro-inflammatory and antimicrobial responses(213). However, over the last 12 years, MyD88 dependent signalling has also been found to play key roles in promoting intestinal epithelial integrity during infection, and in returning the intestine to homeostasis after infection(4-6, 200, 214, 215).  This role was initially recognized when the susceptibility of MyD88-/-, Tlr2-/- and Tlr4-/- mice to DSS induced intestinal 38  inflammation was examined(216).   Though the exact mechanism by which DSS induces inflammation is still not fully understood, it is able to act as a detergent and disrupt the epithelial barrier(217). This likely allows resident intestinal bacteria to translocate into the mucosa and activate local inflammatory responses, potentially mimicking a type of localized “infection”.  With this model Tlr2-/- and Tlr4-/- mice, as well as mice lacking MyD88, were expected to suffer reduced colitis, based on the well-defined pro-inflammatory roles of TLR signalling(216). Instead, these mice suffered exaggerated tissue damage and mucosal ulceration. Interestingly, their susceptibility was not due to altered inflammatory responses, but rather due to defects in IEC proliferation and barrier function.  These normally protective responses were found to depend on TLR recognition of the resident bacteria found in the intestine.  This suggested a role for innate signalling in limiting or repairing intestinal tissue damage, presumably with the goal of helping the host survive a noxious insult to the intestine.    Similar events occur during enteric infection, in that there is frequently a breach of the intestinal epithelial barrier by a pathogen that either itself, or through inducing enough damage in the epithelial layer allowing perturbation by resident bacteria, activates the immune system at the site of infection.  The events occurring at the intestinal epithelium, as well as the role of innate immune signalling, during an enteric bacterial infection will be discussed in the next section.      1.4 Enteric infections and disease models During the course of enteric infections there is significant change within the intestinal environment, from the density and composition of resident luminal microbes to alterations in expression of innate immune receptors by IECs(185, 218-220). Unlike resident intestinal bacteria 39  that are often beneficial to the host, enteric bacterial pathogens employ virulence factors to invade intestinal tissues resulting in barrier disruption, GI inflammation, intestinal pathology, and disturbances in resident intestinal bacterial composition(221-223).  How an individual’s GI tract responds to and recovers from such changes can have lasting implications for future health and disease, as has been highlighted by studies examining post-infectious irritable bowel syndrome (IBS)(224-227).  In addition, changes to intestinal bacterial composition post GI inflammation such as increased colonization with E. coli or C. difficile has been linked to a greater risk of developing allergic atopic disease or IBD(54, 228, 229).  This may reflect altered colonization succession by luminal microbes during recovery from enteric infection, as microbial repopulation is required after the initial loss in numbers and diversity of resident intestinal bacteria that typically occurs during infection.  IEC products released during this time, such as AMPs and goblet cell factors, in addition to those from immune cells help shape this microbial re-colonization.  There are several well-known enteric bacterial pathogens such as Salmonella, Vibrio, Shigella, Campylobacter, enterohemorrhagic and enteropathogenic E. coli (EHEC and EPEC).  These pathogens, upon ingestion by the host, enter the intestinal environment and are able to survive and/or evade luminal defenses including AMPs and the mucus barrier to reach and infect the intestinal epithelium and underlying tissues(230).  Other enteric infections are driven by bacteria referred to as opportunistic pathogens, which are only able to infect the host upon disturbance of its resident intestinal bacteria, such as post-antibiotic treatment, or in immunocompromised individuals. C. difficile represents a common health care associated pathogen in this category(231).  Advances in modeling of enteric bacterial infections have been integral towards 40  furthering our understanding of specific host responses that defend against, and clear such infections, as well as in studying the virulence mechanisms used by pathogens to colonize the host’s GI tract.  I have employed several models of enteric infection in my research to further understand the intricacies of early host responses exerted by, or at the intestinal epithelium, which will be further discussed in the following sections.              1.4.1 Salmonella enterica serovar Typhimurium induced gastroenteritis Nontyphoidal Salmonella infections in immunocompetent individuals are generally confined to the lower GI tract and mesenteric lymph nodes (MLN)(232, 233).  These include infections with the Gram-negative flagellated Enterobacteriaceae Salmonella enterica serovar Typhimurium (S. Typhimurium) and Salmonella enterica serovar Enteriditis.  Ingestion of contaminated food or water are the main sources of infection, with eggs and chickens posing the major concern due to their high carriage rates.   Salmonella GI infections produce a severe infection, resulting in a higher mortality rate than many other foodborne pathogens. Common symptoms of Salmonella infection include nausea, vomiting, diarrhea (in some cases bloody), abdominal pain and fever.  Recent global burden estimates are just under 100 million cases annually, resulting in approximately 155 000 deaths, highlighting a need for animal models to better understand these diseases.                   In the past, studies using murine models to examine host responses to nontyphoidal Salmonella and its infection dynamics were limited, since S. Typhimurium infection of mice resulted in a predominantly systemic infection via gut associated lymphoid tissues that produced a human typhoid-like disease(232, 233).  This murine typhoid like disease was observed in mice deficient 41  in natural-resistance associated macrophage protein 1 (or SLC11A1) such as the commonly used C57BL/6 strain(234).  As a result, bovine models were utilized for many years to gain insight into Salmonella induced enteric infection as infection of calves resulted in similar clinical and histological manifestation of gastrointestinal disease to that observed in humans(232, 233).  Due to the size of cows/calves, the technical specialties required, the lack of tools for immune or genetic manipulation, and the variability in host responses due to their comparatively outbred genetics, there were many practical limitations to this model.    Poor intestinal colonization in mouse models by S. Typhimurium and related species was believed to reflect colonization resistance exerted by the resident intestinal bacteria(233). A mouse model was established in 2003 that involved circumventing this resistance using a single dose of the antibiotic streptomycin, presumably clearing enough resident intestinal bacteria and changing the intestinal environment to provide a niche for S. Typhimurium colonization(235).  Streptomycin treatment followed by oral inoculation with S. Typhimurium 24 hours later, results in intestinal pathology with similarities to human gastroenteritis such as submucosal edema, crypt hyperplasia, goblet cell depletion, immune cell infiltration and epithelial ulceration.      Using bovine and murine models, our understanding of the mechanisms employed by Salmonella to infect their hosts has grown immensely.  Flagella are used to swim through, or otherwise penetrate the mucus layer and approach the intestinal epithelium(236).  Once at the epithelium, the salmonella pathogenicity island (SPI)-1 type III secretion system (T3SS) is induced, and utilized to inject toxins and effector proteins into the host cell’s cytoplasm(235).  This, in addition to the SPI-2 T3SS, allows for the manipulation of host responses and induction of 42  inflammation characterized by extensive colonic and luminal infiltration of neutrophils(237).    Interestingly, in recent years it has been found that intestinal inflammation allows S. Typhimurium to outgrow resident luminal bacteria(238).  This is in part enabled by the respiratory burst of neutrophils that generate tetrathionate by oxidizing endogenous sulfur compounds.   Salmonella are able to use energy generated by cellular respiration (by using tetrathionate as a respiratory electron receptor) instead of relying solely on fermentation, as is the case for resident bacteria in the anaerobic intestinal environment, providing Salmonella with a competitive growth advantage(239).             Though reports on the histopathology elicited during human S. Typhimurium intestinal infection are limited, a few authors have reported on their endoscopic and post mortem findings.  Similar to what is observed in the cecal tissues of pretreated and infected mice, human colon samples displayed diffuse colitis with mucosal ulceration, the presence of undifferentiated epithelial cells likely due to increased IEC proliferation in an attempt to repair damaged tissue, a coinciding lack of mature goblet cells, and only mild ileitis(240, 241).  Interestingly, Myd88-/- mice infected with Salmonella strains lacking SPI-1 and SPI-2 components display limited pathology as compared to wildtype (WT) littermates(242).  These results suggest that the S. Typhimurium model relies to some extent on rapid activation of the innate immune response to induce mucosal injury.  As Salmonella are motile and possess flagella, their presence activates TLR5 dependent pathways, in addition to activation of TLRs 2, 4, 9, NOD like receptors, and inflammasome complexes.   43   1.4.2 Murine model of attaching and effacing pathogens: Citrobacter rodentium C. rodentium is a Gram-negative, murine-specific bacterial pathogen that is a close relative of EHEC and EPEC, two human pathogens of significant clinical interest(243-245).  These bacteria belong to the family of attaching and effacing pathogens that attach to the apical host cell membrane of epithelial cells in the cecum and colon, forming a pedestal-like structure.  Oral challenge with C. rodentium results in intestinal infection and a host response characterized by colonic hyperplasia or elongation of the crypts, immune cell infiltration and goblet cell depletion.  In addition, alterations in the composition of the resident intestinal microbiota represented by a decrease in the number and diversity of intestinal microbes is observed during infection(185, 218). The cecal patch is the initial site of colonization a few hours after challenge, with infection of the distal colon achieved in the following 2 to 3 days(244).  In an immunocompetent host, clearance of the pathogen is achieved by 3 to 4 weeks after infection and is dependent on CD4+ T cells, without significant spread of the pathogen to extra-intestinal sites during the course of infection(246).    In addition to the adaptive T cell response required for mice to survive infection, MyD88 dependent signalling is also integral to mounting appropriate host responses as Myd88-/- mice rapidly succumb to C. rodentium infection(4, 7).  Lack of MyD88 resulted in severe and widespread colonic ulcerations attributed to impaired epithelial barrier function allowing deeper penetration of host colonic crypts by the pathogen.  Further, inadequate IEC proliferation resulted in an inability to repair mucosal injury(4).  Innate mediated pro-inflammatory responses were also affected, as Myd88-/- mice suffered impairments in inflammatory cytokine production and antimicrobial responses, resulting in a 10-100 fold increase in pathogen burdens compared to WT mice(4).  Further, TLR2 has been found 44  to play a significant role in pro-repair mechanisms during enteric infection(247).  For example, studies from our group have found that TLR2 promotes intestinal epithelial homeostasis during C. rodentium infection, even though pathogen burdens and inflammatory responses were similar to those in WT mice(5, 248). The basis for the poor outcomes of infected Tlr2-/- mice reflected significant IEC barrier dysfunction, and increased IEC apoptosis leading to mucosal ulcers and bloody diarrhea.  In contrast, TLR4 dependent responses were found to propagate much of the inflammation and tissue damage observed during C. rodentium infection.  This effect was mediated through induction of chemokines resulting in macrophage and neutrophil recruitment to colonic tissue.       1.4.3 Clostridium difficile infection C. difficile is a spore forming, anaerobic, Gram-positive bacillus associated with an array of clinical outcomes from asymptomatic carriage, to simple diarrhea or pseudomembranous colitis, which in rare cases can result in toxic megacolon, bowel perforation and ultimately death(249).  Unlike the aforementioned enteric pathogens asymptomatic colonization by C. difficile, though rare, does occur in healthy adults (prevalence: 1-7%).  Surprisingly these rates are much higher in pediatric populations (2-37%, up to 2 years of age) and are often, though not always, asymptomatic(250-253). However, what once was a hospital acquired (nosocomial) microorganism primarily affecting elderly patients has more recently emerged as a problem in pediatric and adult hospitalized populations. Heat resistant C. difficile spores can persist in the environment for months to years, most often in hospitals or long-term care facilities.  Upon ingestion these spores can survive through the acidic stomach to reach the LI.  For the development of colonic C. difficile associated disease (CDAD), a disturbance of the resident 45  intestinal bacteria by antibiotic therapy, most often broad spectrum or targeting a variety of bacteria, is required.  This reduces the existing colonization resistance within the gut, allowing C. difficile to flourish and produce toxins that cause mucosal damage and inflammation.  Clostridium difficile toxins (Tcd) A and B are the major toxins released once the microbe is established in the colon, and they are responsible for mediating much of the intestinal inflammation and pathology seen during infection(254).  TcdA targets the actin cytoskeleton of IECs, disrupting the barrier while TcdB enters cells via endocytosis, resulting in induction of apoptosis.  Stimulation of innate immune cells and the release of chemokines result in tissue infiltration by neutrophils.  The extent of symptoms suffered is hypothesized to largely depend on the host immune response.  Though much of the early work examining CDAD was carried out on hamster models, these are not ideal animal models given the few species-specific reagents, lack of genetically modified strains, and the lethal nature of the infection, which is not the normal course in humans(255).  This has led to the development of several mouse models of CDAD in recent years.  Earlier mouse models employed treatment with a mixture of antibiotics for several days before challenge with C. difficile, with more prevalent models injecting purified TcdA and TcdB into surgically generated ileal loops(255).  The first of these, though causing similar histopathology to human CDAD, requires inoculation with C. difficile spores, which poses a biosafety risk with regards to handling and housing of the infected animals.  The latter model is technically complex, involves risks associated with small-animal surgeries, and does not accurately recapitulate human CDAD, which is localized to the colon.  This recently led to the development of an intrarectal instillation model, whereby purified TcdA and TcdB are administered via a 46  catheter into the rectum and colon(256).  In two to four hours, epithelial barrier disruption, extensive tissue damage and a robust inflammatory response are achieved, providing a very efficient model for the study of CDAD susceptibility and progression.  Through the use of these models it was found that much of the damage observed during CDAD is driven by IL-1β and inflammasome dependent signalling(256, 257).                              As mentioned, the incidence of pediatric CDAD has been increasing, with recent data suggesting up to 26% of children hospitalized with CDAD were younger than 12 months of age, and 5% still neonates(252). Interestingly, breast fed infants have been reported to have lower C. difficile carriage rates than formula fed babies(250).  This is of interest as another complicated, bacterially driven disease referred to as Necrotizing Enterocolitis (NEC) which is restricted to the newborn population (specifically low birth weight premature infants) is reported to have improved outcomes with breastfeeding as well(258).   1.4.4 Necrotizing enterocolitis NEC is one of the leading causes of morbidity and mortality in neonatal intensive care units, with this disease affecting 7-10% of infants born very prematurely (≤32 weeks of gestation) and/or with extremely low birth weight (< 1500 g)(142, 258).  A high risk of mortality exists with approximately 30% of infants succumbing to NEC.  This disease is characterized by extensive inflammation, necrotic death of sections of small intestinal tissue, and is often associated with bacterial overgrowth.  It has long been hypothesized that inappropriate bacterial colonization of the premature intestine is a significant predisposing factor for NEC.  As 47  microbial colonization is difficult to avoid at birth, even for premature infants, much research has focused on interventions that can be introduced to prevent NEC development with early birth.  Several factors predisposing infants to NEC have been identified including: extent of prematurity - the more premature the greater likelihood of NEC development, exposure to antibiotics - longer exposure increases susceptibility, H2 blockers – changing the acidity of the GI tract favouring growth of Proteobacteria over Firmicutes, and formula feeding(258-261).  Conversely, breastfeeding has been consistently highlighted for its protective effects against the development of NEC, which can likely be attributed to specific properties of breast milk(262, 263).  Breast milk provides the ideal nutrient composition for proper development of the infant intestine (where there are extensive changes occurring post birth, described in the next section) and overall growth.  In addition, it contains immunoglobulins, protective antimicrobial factors such as lactoferrin, and beneficial bacteria from the mother.  Unfortunately breast milk is not always available, particularly when infants undergo premature delivery and a similar level of protection is not achieved when feeding infants with donor’s milk, which has been reported to result in a similar incidence of NEC as with formula feeding(264).  This has driven significant effort in recent years to create formulas that more accurately reflect breast milk to drive optimal development and protection during the neonatal period.   1.5 Changes in the intestinal environment at birth Birth signals the first incidence of dynamic change in the intestinal environment, which has major consequences for the development of the intestinal epithelial barrier and immune system.  In the womb, one of the major functions of the GI tract – digestion and absorption of nutrients, is 48  achieved through placental transfer.  This process does not involve absorption by the fetal intestine whatsoever; but rather, is dependent on maternal nutrient availability and transfer from the maternal circulation to fetal blood(265).  At birth, the newborn intestine takes over this task.  During this time the GI tract undergoes important developmental changes dependent on the colonizing bacteria, nutrients derived from breast milk or formula, and the ability of the food source to impact the composition of the developing bacterial community(262).  These changes include anatomical changes to increase the absorptive surface area of the epithelium, such as lengthening of villi and microvillus development, increased IEC proliferation and maturation of gut-associated lymphoid tissue(262, 266).                1.5.1 Introduction of food and intestinal development Outside of the presence of small amounts of swallowed amniotic fluid in the womb, the first exposure of a newborn’s intestines to a foreign substance from which it must derive nutrients occurs at birth.  The ideal nutrient source during this time for newborn mammals is breast milk, a complex biological fluid produced by the mother’s mammary gland.  Breast milk has evolved as the exclusive nutrient source required for neonate developmental needs and offers antimicrobial protection from environmental pathogens(267, 268).  Not surprisingly, it also influences the development of the bacterial community residing within the neonate’s intestines, through its antimicrobial properties and by providing nutrient components indigestible by infants such as oligosaccharides, glycolipids, glycoproteins and other milk glycans, that can instead be digested by the microbiota(262, 263).      49  With regards to fueling the development of the intestine, of which there are three phases: pre-natal, neonatal and post-weaning, breast milk (or a formula substitute) facilitates the neonatal phase, which will be the focus of this thesis. During the neonatal phase, many changes occur within the GI tract that have been studied in rodent, avian and piglet models including: significant increases in expression of brush border enzymes from 2.5 weeks to 5 months of age, changes in tight junction protein distribution (e.g. Claudin-15 progresses up the crypt-villus axis, while Claudin-2 moves down), up to a 10x increase in goblet cell numbers in some species, and other significant changes to epithelial architecture(17, 95, 266, 269, 270).  Changes to the epithelium in a rat model during the first 35 days post birth include increases in the number and depth of crypts at all sites of the lower GI tract, as well as increases in villus height at SI sites, with a decrease in the number of villi, likely due to villus fusion(266).    1.5.2 Breast milk composition and benefits Feeding exclusively with breast milk for the first 6 months of life, continued with food supplementation for 1 to 2 years post birth is recommended by most health care experts due to its numerous beneficial properties for infants(271).  Breast milk contains essential micro- and macro- nutrients, growth factors and other hormones, beneficial microbes, immune-protective components, as well as enzymes that aid the infant in its ability to digest and absorb nutrients(262, 263).  Interestingly, breast milk composition changes throughout lactation, corresponding to changes in the nutritional needs of the developing infant(263, 272).  The first milk secreted up to 4 days postpartum, the colostrum, has a high concentration of proteins, many of them antimicrobial. Transitional (5-14 days postpartum) and mature milk are characterized by much higher fat and lactose (sugar found in milk) concentrations than the colostrum.  50  Macronutrient concentrations for mature human milk have been approximated at 1.2 g/dL protein, 3.6 g/dL fat, and 7.8 g/dL lactose(263). The major proteins in breast milk include casein, α-lactalbumin (involved in lactose production, may be antimicrobial), serum albumin (source of amino acids), and the antimicrobial proteins lactoferrin, secretory IgA, and lysozyme(273).  Lactose comprises the main sugar in milk used for energy, while other important milk carbohydrates include oligosaccharides, which although indigestible by the infant, likely influence the establishment of the resident bacteria while providing antimicrobial protection against pathogens(272, 274).  Interestingly, milk fat provides around half (45-55%) of the energy in breast milk(275).   There are many immediate health benefits facilitated by breast milk, including its ability to protect the infant from developing diarrhea and GI infections.  In recent years there has been a growing appreciation for the potential implications of breast milk in long-term health.  Breastfeeding has been associated with a reduced risk of development of many diseases including obesity, type 1 and 2 diabetes, asthma, and atopic dermatitis(262, 276-278). As mentioned above, breast milk is not always available to the developing infant, and there are many differences that exist between its composition and that of formula. For instance, the protein portion of formula is composed of a much higher proportion of casein, which can be difficult to digest for some infants(279).  Many of the bioactive molecules of breast milk such as IgA, milk oligosaccharides, as well as its unique fat component are also not present in conventional formulas. Interestingly, the fat component of breast milk is also significantly altered during the pasteurization and freezing processes required for the storage and use of donor milk to limit transmission of infectious agents(280).  This is noteworthy as, in addition to serving as a major 51  energy source, the unique structure and components of milk fat have significant bioactive potential, which may confer some of the protective effects of breast milk.  This may also explain, to some extent, the inability of donor milk to confer the same level of protection against NEC as breast milk from the mother.        1.6 Milk fat globule membrane The molecules comprising the fat component of breast milk represent unique structures in nature.  The fat portion of breast milk contains a substantial number of proteins, reflecting its impressive bioactive potential(263, 280, 281).  This is due to the mechanism by which fat molecules are synthesized and secreted by the lactating mammary gland.  Surprisingly, the fat component of most infant formulas, which is derived from vegetable sources, does not resemble the structure or composition of milk fat(280).  1.6.1 Composition In mammary epithelial cells, droplets of triglycerides, comprising the main lipid component of breast milk (up to 80%), are synthesized in the ER and released into the cytoplasm surrounded by a single layer of ER membrane phospholipids (Figure 1.6)(267, 281).  These droplets fuse, increasing in size, and move towards the apical cell membrane where they are enveloped by the epithelial cell membrane and secreted.  This process results in a rare triple membrane structure composed of phospholipids, cholesterol, and cell membrane glycoproteins and proteins, referred to as Milk Fat Globule Membrane (MFGM), surrounding the triglyceride droplet core.    52   Figure 1.6:  Milk Fat Globule Membrane secretion and structure Schematic drawing of the process through which the milk fat globule is released from the mammary gland epithelial cell. The triple membrane structure of the MFGM (inlay) contains many transmembrane proteins.  (Image reproduced from (281) with permission)  The lipid portion of MFGM is composed largely of phospholipids (e.g. sphingomylein, phosphatidylcholine and phosphatidylethanolamine) and cholesterol, which in addition to providing a valuable source of energy can also affect several cell signalling pathways that impact absorption processes, molecular transport systems and stress responses(282-284). With a lipid to 53  protein ratio of 1:1, MFGM also harbours a significant protein component, which accounts for 1-4% of the total protein in milk(285). This includes several membrane glycoproteins that are able to act as decoy-receptors by binding potentially pathogenic bacteria and inhibiting their adhesion to the epithelium, similar to classic breast milk oligosaccharides.  An example of one such glycoprotein is Mucin-1, which is able to bind to, and inhibit rotavirus, Norwalk virus, as well as several pathogenic strains of E. coli and S. Typhimurium (286).  Studies examining the resistance of Mucin-1, along with several other MFGM proteins, to gastric digestion have revealed their ability to completely or partially resist degradation along the length of the GI tract, providing their potential beneficial effects along its length(287).  Lactadherin, another MFGM protein capable of surviving digestion, has been reported to interfere with enterotoxigenic E. coli binding to IECs as well as have antiviral effects(288).  Some proteins found in the protein component of breast milk are also found in the MFGM, such as lactoferrin, which outcompetes many bacteria for iron(289).  There are many other MFGM proteins with antimicrobial capacity through various mechanisms including, but not limited to, butyrophilin, secretory IgA, and glycoprotein 2, which is hypothesized to serve as a molecular decoy for E. coli Type 1 fimbriae (bacterial appendages used for adherence)(285, 288, 290).     1.6.2 Modeling neonate development: rat pup-in-a-cup model      To accurately understand and research the specific factors affecting neonate intestinal development, models using newborn animals must be employed.  These can be difficult and technically very challenging to carry out, but ultimately can lead to the improvement of nutritional guidelines in a vulnerable population undergoing a critical period of development.  Of those that exist, including mice, rats, guinea pigs, pigs, and non-human primates, neonatal rodent 54  and piglet models are the most widely used(291).  Rodents provide an advantage over piglets due to their low cost, greater potential for genetic manipulation, and larger selection of reagents.  To ensure complete control over nutrient intake and volume, artificial rearing must be utilized(292).  For rodents, this requires separation from the mother and littermates to individual housing units.  As this can be a stressful procedure in itself, adequate bedding and proper temperature maintenance is absolutely required.  This can be achieved by housing floating containers in warm water baths (i.e. pup-in-a-cup model).  Further, to administer milk substitute formula, a gastrostomy must be performed to place a feeding cannula into the stomach of 5 day old mouse or rat pups(292).  The cannulas are attached to low volume peristaltic pumps, which allow the flow of formula into the stomach in a pattern similar to that of natural suckling.  The use of artificial rearing models requires extensive monitoring of the animals (16 of 24 hours) in the form of health checks and stimulation of urination and defecation.  In addition, proper functioning of pumps and adequate replacement of formula must be ensured.  Although technically challenging and labour intensive, when carried out effectively rodent neonate models provide a valuable tool towards understanding the impact of specific nutrients on mammalian development and susceptibility, that could not otherwise be explored.  I employed the rat pup-in-a-cup model in my research to better understand the effect of MFGM supplementation in infant formula during times of dynamic change: neonate intestinal development and during C. difficile toxin induced disease.     1.7 Research objectives I have thus outlined multiple ways in which the mammalian intestine serves the extraordinary task of digestion and absorption of nutrients while maintaining an effective barrier from luminal 55  contents in concert with resident bacteria, specifically at the level of the IECs.  Further, our current understanding regarding the changes occurring at this level during times of flux in the intestinal environment, such as birth and infection, to achieve optimal intestinal health have been highlighted.          Although there are many differences during these two states, similarities also exist including low numbers and diversity of resident intestinal bacteria, as well as high IEC proliferation rates.  IEC responses during intestinal development and enteric infections likely impact microbial colonization succession, which is hypothesized to have important ramifications on long-term health(293).  This includes long term susceptibility to the development of diseases such as IBS, IBD, allergies, asthma, and diabetes among many others.  As every individual must undergo intestinal development and enteric bacterial infections impact people worldwide, better understanding the changes occurring at IECs and the intestinal barrier during these two important stages is highly relevant.  Further, better characterizing beneficial responses during these times and the factors that can be added to facilitate them is of particular interest for vulnerable populations such as premature infants, or immune-comprised individuals more likely to succumb to enteric infections.    The goal of my thesis work is therefore to better understand how times of significant change in the intestinal environment impact the IECs and the intestinal barrier. It is hypothesized that (1) IEC-intrinsic innate signalling is beneficial during enteric infection and (2) the addition of MFGM to formula positively impacts the intestinal epithelium during neonatal development. The 56  specific objectives of my thesis are 1) to examine the role of innate signalling, specifically MyD88 dependent pathways, in IECs during models of enteric infection, and 2) the impact of MFGM supplementation in formula on IECs and susceptibility to intestinal inflammation during neonate development.  Findings from these studies will help determine how we can modulate host responses at the IEC barrier during intestinal development and enteric infection, to ultimately define mechanisms that can benefit intestinal health.    57  Chapter 2: Intestinal epithelial specific MyD88 signalling impacts host susceptibility to enteric infection through goblet cell and antimicrobial responses  2.1 Introduction Each year, enteric infections cause more than 1.5 billion cases of diarrheal disease, along with more than 2 million deaths. Many of these cases reflect infection by bacterial pathogens such as Salmonella enterica serovar Typhimurium and enteropathogenic E. coli (EPEC)(294, 295).  Although bacterial virulence factors play a significant role in determining whether an infection is successful, the host’s innate immune response also plays an integral role in determining its overall susceptibility to infection, as well as regulating the course and severity of disease suffered during a successful infection.  This response to infection can have lasting implications for host health, as is evidenced by rates of post-infectious IBS.  IBS is amongst the most common gastrointestinal disorders in the Western world, associated with abdominal pain, bloating, diarrhea and/or constipation.  Though the mechanisms underlying IBS are incompletely understood, recent reports suggest 3.7 to 36% of individuals develop IBS after bacterial or viral intestinal infection.      A significant aspect of the host’s innate response to such infections is mediated by Toll-like Receptor (TLR) signalling pathways, most of which signal by recruiting the key adaptor protein MyD88(176). Ligation of most TLRs by conserved microbial-associated molecular patterns, as well as interleukin 1 and 18 receptors (IL1R, IL18R) by their respective ligands, results in 58  recruitment of MyD88 to the receptor complex initiating its activation and a cascade of signalling events leading to the activation of an array of inflammatory genes(176).    Although first recognized for its role in pro-inflammatory pathways, several studies in recent years have uncovered a critical protective role for MyD88 signalling in the promotion of mucosal homeostasis following inflammatory insult(4, 7, 215, 216, 296-299).  This notion was first identified when Myd88-/- mice were shown to be highly susceptible to DSS induced-colitis(216). MyD88 signalling was found to play a key protective role, promoting the production of tissue protective factors and the increased proliferation of IECs during colitis, with mice lacking MyD88 suffering widespread mucosal tissue damage and ulceration. Later work by our group and others found that MyD88 also played an essential protective role during the infectious colitis caused by the natural murine pathogen Citrobacter rodentium(4, 7), which is closely related to EPEC(243).    Bone marrow transplantation studies have shown that the protective effects of MyD88 during models of intestinal inflammation reflect signalling in both the hematopoietic as well as the non-hematopoietic cell compartments(7, 215, 299). While many of the MyD88 dependent changes that prove protective during enteric infection reflect changes in IEC function and/or proliferation, it remains uncertain whether any of the protective signalling occurs within the IECs themselves. Considering that the IEC layer is in constant contact with luminal microbes, these cells are known to be innately hypo-responsive to most bacterial products, as a way to prevent overt spontaneous inflammatory responses against resident microbes(185, 300, 301). In fact, several groups have studied the specific role of MyD88 signalling in IECs during DSS-induced colitis, 59  and most, but not all studies found little evidence for its involvement(298, 302). Correspondingly, in a recent study where we examined whether IECs played any role in responding to C. rodentium infection, we found no evidence that IEC-Myd88-/- mice were more susceptible to infection than WT mice, although in this study we focused solely on later stages of infection(185).    Considering that innate signalling within IECs might have a greater impact at early stages of infection, prior to the recruitment of large numbers of inflammatory cells, we decided to re-examine the role of MyD88 signalling in IECs. We therefore infected WT and IEC-Myd88-/- mice with the enteric pathogens S. Typhimurium and C. rodentium and examined responses at very early time points. We found that loss of MyD88 signalling in IECs increased early susceptibility to infection, impairing the induction of several key antimicrobial genes as well as the expression of the goblet cell specific factors Relmβ and Muc2 during infection. Correspondingly, the bactericidal activity of crypt supernatants from IEC-Myd88-/- mice was significantly reduced against both S. Typhimurium and C. rodentium when compared to crypts isolated from WT mice. This impairment in antimicrobial defenses appears to facilitate the ability of these bacterial pathogens to reach and infect the intestinal epithelium, resulting in accelerated intestinal tissue damage and barrier dysfunction. Our study thus demonstrates that MyD88 signalling in the intestinal epithelium in vivo confers protection to the host at the initial stages of an enteric bacterial infection.   60  2.2 Experimental procedures 2.2.1 Mice MyD88flox/flox mice were a kind gift from Dr. Xiaoxia Li (Lerner Research Institute, Cleveland, OH, USA) and Villin-cre mice were purchased from Jackson Laboratory (Bar Harbor, ME, USA), both mouse strains were on a C57BL/6 background.  Mice lacking MyD88 signalling specifically within IECs (IEC-Myd88-/-) were generated by crossing MyD88flox/flox mice with Villin-cre.  For experiments, co-housed littermates from IEC- Myd88-/- mice (hemizygous for Villin-cre) bred with MyD88flox/flox were used, with the resulting litters genotyped to identify controls (example provided in Figure 2.1).  MyD88flox/flox mice were used as wildtype controls for all studies. All mice were kept in sterilized, filter-topped cages and fed autoclaved food (PicoLab Rodent Diet 20 #5053, Lab Diet, Brentwood, MO, USA) and water under specific-pathogen-free conditions at the British Columbia Children’s Hospital Research Institute (BCCHRI). Sentinel animals were routinely tested for common pathogens at the BCCHRI animal facility.  The protocols employed were approved by the University of British Columbia’s Animal Care Committee and were in direct accordance with guidelines provided by the Canadian Council on the Use of Laboratory Animals.  61   Figure 2.1: Genotyping of mice Agarose gel electrophoresis photo for IEC-MyD88-/- genotyping.  MyD88flox/flox allele specific primer PCR products appear at 353 bp and Villin-cre specific primer amplicons occur at 1.1 Kbp.   2.2.2 Bacterial strains and infection of mice Salmonella enterica serovar Typhimurium SL1344 ΔaroA(303) and streptomycin resistant Citrobacter rodentium DBS100 (formerly C. freundii biotype 4280, strain DBS100) strains were grown, shaking (200 rpm) at 37°C in lysogeny broth supplemented with 100g/mL streptomycin. 24 h prior to SL1344 ΔaroA infection, 6-10 week-old mice were treated with 20 mg of streptomycin (in 100µL) by oral gavage.  Mice were infected with 3 x 106 CFU of Salmonella in 100µL phosphate buffered saline (pH 7.2) by oral gavage. Mice infected with C. rodentium were inoculated by oral gavage with 100 µL of overnight culture (~2.5 x 108 colony forming unites (CFU)).  Mice were weighed daily and assessed for any signs of stress for the duration of the study.        62  2.2.3 Tissue collection and bacterial counts Mice were anesthetized with isofluorane and euthanized via cervical dislocation at various time points over the course of infection.  For bacterial cell counts, the cecum, colon, and luminal contents were each collected and homogenized separately in 1 mL of sterile phosphate buffered saline.  Samples were weighed and homogenized in a MixerMill 301 bead miller (Retsch, Haan, Germany).  Homogenates were serially diluted in phosphate buffered saline and plated on streptomycin supplemented lysogeny broth agar plates, incubated at 37°C overnight and colonies enumerated and normalized to the weight of the tissues.  For histology, cecal samples were fixed in 10% neutral buffered formalin (Thermo Fischer Scientific, Waltham, MA, USA) overnight, transferred to 70% ethanol, embedded in paraffin and cut into 5 µm sections.    2.2.4 Histology scoring 5 µm tissue sections were stained with hematoxylin and eosin (by the histology laboratory at the BCCHRI) and were examined by two blinded observers to assess histological damage.  Tissue sections were assessed for: (i) submucosal edema (0-no change, 1- mild, 2- moderate, 3- severe) (ii) hyperplasia (0-no change, 1: 1-50%, 2: 51-100%, 3: >100%) (iii) goblet cell depletion (0-no change, 1-mild depletion, 2-severe depletion, 3-absence of goblet cells) (iv) epithelial integrity (0-no pathological changes detectable, 1-epithelial desquamation (few cells sloughed, surface rippled, 2-erosion of epithelial surface (epithelial surface rippled, damaged), 3-epithelial surface severely disrupted/damaged, large amounts of cell sloughing, 4-ulceration (with an additional score of 1 added for each 25% of tissue in the cross-section affected, e.g. a large ulcer affecting 70% of the tissue section would score 4+3)) (v) mucosal mononuclear cell infiltration (per 400x magnification field) (0-no change, 1- <20, 2- 20 to 50, 3- >50 cells/ field) (vi) submucosal 63  polymorphonuclear leucocytes and mononuclear cell infiltration (per 400x magnification field) (1- <5, 2- 21 to 60, 3- 61 to 100, 4- >100 cells/ field).  The maximum possible score was 22.     2.2.5 Resident microbe analysis   Microbial composition analysis was performed by quantitative PCR (qPCR) as described previously(43).  Briefly, DNA was extracted from at least two fecal pellets per animal using the Qiagen DNA stool extraction kit (Qiagen, Hilden, Germany).  50 ng of extracted DNA per reaction was used for qPCR on a MJ Mini-Opticon Real-Time PCR System (Bio-Rad, Hercules, CA, USA) using IQ SYBR Green Supermix (Bio-Rad) and specific primers listed.  16s rRNA group specific primers were used to determine the relative abundance of the selected bacterial phyla: Bacteroidetes (5′-GAG AGG AAG GTC CCC CAC-3′, 5′-CGC TAC TTG GCT GGT TCA G-3′)(304), Firmicutes (5′-GGA GYA TGT GGT TTA ATT CGA AGC A-3′, 5′-AGC TGA CGA CAA CCA TGC AC-3′)(305), and γ-Proteobacteria (5′-TCG TCA GCT CGT GTY GTG A-3′, 5′-CGT AAG GGC CAT GAT G-3′)(306).  Universal Eubacteria primers (5′-ACT CCT ACG GGA GGC AGC AGT-3′, 5′-ATT ACC GCG GCT GCT GGC - 3′)(307) were used to determine total bacterial 16S rRNA in each sample, and relative abundance of each taxonomic group was determined by calculating the average cycle threshold value relative to this number, normalized to each primer’s determined efficiency. 2.2.6 FITC-dextran intestinal permeability assay Uninfected mice or mice at D1, D3 (S. Typhimurium) or D4 (C. rodentium) pi were gavaged with 150 µl of 80 mg/ml 4 kDa fluorescein isothiocyanate (FITC)-dextran (Sigma-Aldrich, St. Louis, MO, USA) in phosphate buffered saline, 4 hrs prior to sacrifice. Mice were anaesthetized with isofluorane until withdrawal reflex was absent, and blood (~500 µl) collected by cardiac 64  puncture then added immediately to a final concentration of 3% acid-citrate dextrose (20 mM citric acid, 100 nM sodium citrate, 5 mM dextrose). FITC-dextran concentration in serum was measured using a fluorometer (Perkin-Elmer Life Sciences, Waltham, MA, USA) (excitation λ 485 nm, emission λ 530 nm).  2.2.7 RNA extractions and quantitative real-time PCR Immediately following euthanization of mice, cecal tissues were placed in RNA-later (Qiagen) and stored at -80°C.  Total RNA was extracted using a Qiagen RNeasy kit, according to manufacturer’s instructions.  Total RNA was quantified using a NanoDrop spectrophotometer and complementary DNA was synthesized using 1 µg of RNA with an Omniscript reverse transcription kit (Qiagen, Hilden, Germany).  For the qPCR reaction, 5 µL of 1/5 dilution of complementary DNA was added to 10 µL Bio-Rad SYBR green supermix with primers [300nM final concentration] (final volume = 20 µL), and qPCR was carried out using a Bio-Rad MJ Mini-Opticon machine. Quantitation of data was carried out using Gene Expression Macro OM 3.0 Software (Bio-Rad).      2.2.8 Immunofluorescence 5 µm paraffin section were deparaffinized by heating to 60°C for 15 minutes, cleared with xylene, rehydrated through an ethanol gradient to water and steamed for 30 minutes in citrate buffer for antigen retrieval.  Tissues were then blocked using blocking buffer (Goat or Donkey serum in phosphate buffered saline containing 1% bovine serum albumin, 0.1% Triton-X100, and 0.05% Tween 20, and 0.05% sodium azide).  The primary antibodies used were anti-Muc2 (1:500, Santa Cruz Biotechnology, Dallas, TX, USA), anti-Relmβ (1:400, Peprotech, Rocky Hill, 65  NJ, USA), anti-Ki-67 (1:200, Thermo Fisher Scientific), anti-Salmonella LPS (1:50, BD Biosciences, Franklin Lakes, NJ, USA), anti-β-actin (1:200, Santa Cruz Biotechnology) or anti-Tir (1:2000, kind gift from Dr. Wanyin Deng) and the secondary antibodies (all 1:2000, Molecular Probes, Eugene, OR, USA) were AlexaFluor 568- or 488- conjugated goat anti-rabbit or goat anti-rat IgG and AlexaFluor 568- or 488- conjugated donkey anti-rabbit or donkey anti-goat IgG . ProLong Gold Antifade reagent with DAPI (Invitrogen, Carlsbad, CA, USA) to stain DNA was used to mount tissues.  Tissues were viewed on a Zeiss AxioImager microscope and images taken using AxioVision software and an AxioCam HRm camera (Zeiss, Oberkochen, Germany).  2.2.9 Fluorescence intensity measurements Fluorescence intensities of immunostained samples were assessed using ImageJ (open source image processing software) to determine the ratio of Muc2 or Relmβ in each tissue section relative to DAPI staining.  This was done using the Integrated Density measurement tool.  Integrated Density for each image was assessed on its separate channels to determine the pixel intensity of DAPI and Muc2 or Relmβ for each section.  The fluorescence intensity was then represented as Integrated Density value of Muc2 or Relmβ, relative to total DAPI Integrated Density values.  2.2.10 Crypt killing assay As described previously(185, 308), cecal and colonic crypts were isolated from uninfected WT (C57Bl/6) and IEC-Myd88-/- mice.  Ceca and colons were extracted from mice following euthanization and placed in 50 µg/ml gentamycin in sterile phosphate buffered saline follwing 66  removal of fecal contents.  Tissues were washed three times in the Gentamycin phosphate buffered saline solution, then cut into small (0.5 cm) sections and placed in a petri dish containing 10 mL of cell recovery solution (BD Biosciences) at 4°C  for 2 hours.  Crypts were then dislodged by gently flicking with forceps in the dish.  Following centrifugation, crypts were resuspended in iPIPEs buffer (10 mM PIPES; pH 7.4 and 137 mM NaCl) at a ratio of 2000 crypts per 40 µL buffer and incubated at 37°C for 30 minutes.  Samples were then centrifuged and supernatant collected.  11 µL of supernatant was added to 103 ΔaroA S. Typhimurium, 103 C. rodentium, or 20 µM RegIII-γ (MyBioSource, San Diego, CA, USA) as a positive control and incubated at 37°C for 2 hours.  Antimicrobial activity was assessed by counting overnight growth of crypt supernatant treated ΔaroA S. Typhimurium or C. rodentium and expressed as % bacterial growth relative to overnight growth observed in cultures treated with sterile iPIPEs alone.    2.2.11 Statistical analysis All results presented in this study are expressed as the mean value ± Standard Error of the Mean (SEM). Nonparametric Mann–Whitney t-tests, or Student’s t tests were performed using Graph-Pad Prism Software for Mac.  A P value of 0.05 or less was considered significant.  2.3  Results 2.3.1  IEC-Myd88-/- mice suffer exaggerated S. Typhimurium induced gastroenteritis To assess whether IEC specific MyD88 signalling plays any role in regulating host defense and intestinal inflammation in response to S. Typhimurium, we infected Myd88flox/flox (WT) and IEC-Myd88 -/- mice with ΔaroA S. Typhimurium and assessed body weights over the following seven 67  days. This Salmonella strain was chosen because it does not kill mice on a C57BL/6 genetic background, which lack the natural resistance-associated macrophage protein 1, whereas it still causes severe gastroenteritis in infected mice(309, 310). Although 6-8 week old WT and IEC-MyD88-/- mice displayed similar initial body weights (19.94±1.141 g and 19.2±0.3742 g, respectively) (Figure 2.2A), infected WT mice showed an early weight gain at D1-3 pi followed thereafter by a modest weight loss, while IEC-Myd88 -/- mice exhibited significantly greater (*p<0.05) weight loss (~5%) over the first six days of infection (Figure 2.2B). Following their euthanization at specific time points, infected IEC-Myd88 -/- mice showed greater macroscopic intestinal damage than WT mice, with severely shrunken ceca that were devoid of stool content as early as D1 pi (Figure 2.2C).    Figure 2.2: IEC-MyD88 -/- mice are more susceptible to S. Typhimurium enteric infection  68  Figure 2.2: IEC-MyD88-/- mice are more susceptible to S. Typhimurium enteric infection A) WT and IEC-MyD88 -/- mice display similar initial body weights (n=7-8). B) Body weights of WT and IEC-MyD88 -/- mice from D0 to D7 pi, plotted as the percentage of starting weight.  IEC-MyD88 -/- mice exhibited rapid weight loss by D1 pi that remained significantly below that of WT mice until D6 pi (n≥7, from at least 2 experiments).  C) Unlike WT mice, at D1 pi the IEC-MyD88 -/- intestinal tissues displayed severe damage, with severely shrunken ceca devoid of stool contents.  Black arrow indicates inflamed ceca, error bars indicate SEM, Asterisks show significance at *p<0.05 and **p<0.005.    As the majority of the pathology seen in this model occurred in the cecum, we focused our subsequent analysis on this region. At both D1 and D3 pi, IEC-Myd88 -/- mice displayed significantly greater cecal histopathology scores compared to WT mice, represented by increased goblet cell depletion, epithelial damage and inflammatory cell infiltration (Figure 2.3A&B, p<0.001). Specifically, we noted the presence of more macrophages and neutrophils infiltrating the cecal tissues of the IEC-Myd88 -/- mice, although infected WT and IEC-Myd88 -/- mice showed similarly increased gene transcript levels for the chemokines MCP-1 and MIP-2α (Figure 2.3C).  While IEC-Myd88 -/- mice showed decreased induction of gene transcript levels for the inflammatory cytokine TNF-α compared to WT, IFN-ϒ, IL-1β and IL-17A were induced to similar levels at D1 pi (Figure 2.3C, *p<0.05).       69   Figure 2.3: IEC-MyD88 -/- mice suffer accelerated tissue damage during S. Typhimurium infection A) Representative hematoxylin and eosin staining of uninfected, D1 and D3 pi cecal tissues from WT and IEC-MyD88 -/- mice. Inflammatory cell infiltrate is marked as follows: (*mucosal infiltration, #submucosal infiltration), and damage to IEC integrity (arrows) at D1 and D3 pi. B) Comparative histological damage scores of uninfected, D1 and D3 pi IEC-MyD88 -/- and WT mice.  Cecal tissues of IEC-MyD88 -/- mice displayed significantly higher damage scores at both D1 and D3 pi. (C) Infected WT and IEC-MyD88 -/- mice show similar increases in gene transcript levels for the chemokines MCP-1 and MIP2-α, as well as IFN-γ, IL-1β, and IL-17A. IEC-MyD88 -/- mice showed significantly decreased transcript levels for TNF-α after infection with ΔaroA S. Typhimurium. Results are representative of at least 3 independent infections, each with 3 to 5 mice.  Error bars indicate SEM, asterisks show significance at **p< 0.005 and ***p<0.001. Original magnification: 200X.   70  2.3.2 IEC-Myd88-/- mice show altered localization of S. Typhimurium within ceca As increased intestinal pathology is often associated with higher pathogen burdens during infection, we enumerated ΔaroA S. Typhimurium within cecal tissues and within the lumen, as well as in the spleen and liver tissues of mice at D1 and D3 pi. Surprisingly, no significant differences in pathogen burdens were found at any of these sites at either time point (Figure 2.4A). Since exaggerated inflammation could also reflect deeper penetration of tissues by the pathogens, we next used immunostaining to visualize the location of S. Typhimurium within the intestine. Immunostaining for Salmonella LPS revealed that the majority of S. Typhimurium remained sequestered within the cecal lumen in WT mice, whereas, in IEC-Myd88 -/- mice, the S. Typhimurium were found in close proximity to the epithelial surface as outlined by staining for β-actin (Figure 2.4B).  71   Figure 2.4: Disease severity of IEC-Myd88-/- mice is associated with altered S. Typhimurium localization A) No differences in ΔaroA S. Typhimurium pathogen burdens (CFU/gram) were identified among the ceca, luminal contents, spleens or livers from IEC-MyD88 -/- and WT mice at D1 or D3 pi.  Error bars indicate SEM. Results are representative of at least 3 independent infections, each with 3 to 5 mice. B) Immunofluorescence staining for Salmonella LPS (red), β-actin (green) and DNA (blue) in cecal tissues at D1 pi. Presence of MyD88 in IECs (WT) prevents ΔaroA S. Typhimurium (location indicated by white arrow) from associating closely with the epithelium, sequestering them to the lumen, whereas S. Typhimurium in the IEC-MyD88 -/- ceca were found in close proximity to the epithelial surface.  Original magnification: 630X.  72  2.3.3 IEC specific MyD88 signalling does not overtly alter the gut microbiome Host susceptibility to enteric bacterial infections can be influenced by differences in the makeup of the intestinal microbiota. A recent study reported that the resident microbe populations of IEC-Myd88 -/- mice differed from that of WT mice under baseline conditions(302). While our pretreatment of mice with the antibiotic streptomycin likely minimized any impact of the microbiota in our infections, we decided to still assess the intestinal microbiomes of the mice at our facility, both pre- and post-streptomycin treatment.  Using qPCR analysis of fecal pellets we found that members of the phylum Bacteroidetes were the dominant bacteria present under uninfected conditions in both groups (~50-70%), whereas smaller numbers of Firmicutes (~5-10%) and other bacteria represented the remainder of luminal bacteria.  There was a substantial shift in bacterial populations post-streptomycin treatment at D1 pi, with significant decreases seen in both Bacteroidetes and Firmicutes and a significant increase in γ-Proteobacteria (~35-60%, likely largely representing Salmonella) in both WT and IEC-Myd88 -/- mice (Figure 2.5). Despite modest differences in the exact percentages of the different phyla between WT and IEC-Myd88 -/- mice, no overt/significant differences in bacterial phyla were noted between strains under uninfected, or post-streptomycin infected conditions (Figure 2.5).  73   Figure 2.5: Similar bacterial populations at phylum level in stool samples of WT and IEC-Myd88-/- mice The resident intestinal bacteria found in fecal pellets from WT and IEC-MyD88 -/- mice under uninfected conditions and after streptomycin treatment at D1 pi, as measured by qPCR.  No significant differences were found between the intestinal bacterial populations (at phylum level) present in WT and IEC-MyD88 -/- mice. Results are representative of 2 independent experiments, each with 3 to 5 mice.  2.3.4 MyD88 signalling in IECs protects barrier integrity during infection Several previous studies have implicated MyD88 dependent signalling as driving profound protective changes in IEC proliferation and barrier function(4, 216) during infectious or chemical induced models of colitis. Moreover defects in IEC responses could be responsible for the exaggerated cecal inflammation suffered by the IEC-Myd88 -/- mice. We therefore examined whether the barrier maintenance and increased IEC proliferation that develops during S. Typhimurium infection was mediated by MyD88 signalling within the IECs.  74  Following oral gavage of FITC-dextran, we found IEC specific MyD88 signalling had no effect on baseline barrier function, as serum FITC dextran levels were similar in uninfected WT and IEC-Myd88 -/- mice (Figure 2.6A). In contrast, although ΔaroA S. Typhimurium was able to cause a modest impairment in epithelial barrier function in WT mice by D1 that lasted until D3 pi (*p<0.05), infected IEC-Myd88 -/- mice demonstrated significantly greater (2-fold) intestinal permeability at D1 (*p<0.05) compared to WT mice. By D3 pi, while the FITC dextran levels in IEC-Myd88 -/- mice were still significantly higher than those of uninfected mice (Figure 2.6A, **p<0.005), they were no longer significantly higher than the levels in WT mice.   We next immunostained tissues for Ki-67, a nuclear factor marking cell proliferation (Figure 2.6B) and noted similar levels of baseline IEC proliferation in uninfected WT and IEC-Myd88 -/- mice. In contrast, at D1 pi IEC proliferation was significantly increased (measured as % of Ki-67 positive cells per crypt) in IEC-Myd88 -/- crypts compared to WT (Figure 2.6C, ***p<0.0005). By D3 pi, IEC proliferation continued to increase in both groups, such that there were no significant differences between the two groups (Figure 2.6C). These data indicate that MyD88 signalling within IECs plays a critical role in promoting protective barrier responses early during ΔaroA S. Typhimurium infection. In contrast, MyD88 signalling within IECs is not required for the increased proliferation of IECs seen during infection, with the more rapid induction of proliferation in the IEC-Myd88 -/- mice likely reflecting the accelerated inflammation these mice suffer.  75   Figure 2.6: IEC-MyD88 -/- mice display impaired barrier integrity at early infection time points A) FITC-dextran based intestinal permeability assay performed on WT and IEC-MyD88 -/- mice, under uninfected as well as D1 and D3 pi conditions.  IEC-MyD88 -/- mice showed significantly increased barrier permeability compared to WT mice at D1 pi.  Both groups experienced increased barrier permeability due to infection at D1 and D3 pi as compared to uninfected conditions. Bars represent the average values for at least 7 mice per group, from 3 independent experiments.  B) Immunostaining for the proliferation marker Ki-67 (red) and DNA (blue) revealed that WT and IEC-MyD88 -/- mice had increased proliferation in cecal tissue beginning at D1 pi. C) Quantification of % Ki-67 positive cells per crypt showed there were significantly more proliferating cells in IEC-MyD88 -/- mice at D1 pi than in WT.  Results are representative of at least 3 independent infections, each with 3 to 5 mice.  Error bars indicate SEM, Asterisks indicate significance *p<0.05, **p<0.005 and ***p<0.0005.  76  2.3.5 IEC-Myd88-/- mice are impaired in antimicrobial and goblet cell specific responses While impaired IEC barrier function could help drive the exaggerated inflammation suffered by infected IEC-Myd88 -/- mice, we examined whether there were other defects in IEC function/defense that could be involved. Interestingly, IEC-dependent MyD88 signalling has previously been found to maintain a microbial free zone above the host’s mucosal surface in the small intestine (SI)(158). Upon assessment of several antimicrobial factors by qPCR, we found that they were similarly expressed in cecal tissues of the two mouse strains under uninfected conditions (Figure 2.7A-C).  As Vaishnava et al.(158) have previously reported a significant reduction of RegIII-γ gene transcript levels in the terminal ileum of uninfected IEC-MyD88-/- mice, we assessed its levels in the terminal ileum of our IEC-MyD88-/- mice and confirmed that compared to WT mice, RegIII-γ gene transcript levels in the ilea of IEC-MyD88-/- mice were impaired (Figure 2.7D, *p=0.03). By D1 pi, transcription was significantly increased for several of these antimicrobial genes in WT mice; however, this induction was dramatically impaired for RegIII-γ in the IEC-Myd88 -/- mice (Figure 2.7A, *p<0.05). The induction of RegIII-β also seemed impaired, although the difference with WT mice did not reach significance (Figure 2.7C).  Aside from antimicrobial factors, the mucosal surface of the large bowel is protected from luminal bacteria by a thick mucus layer that is largely comprised of the goblet cell derived mucin Muc2. We therefore examined whether intrinsic MyD88 signalling impacted on goblet cell specific factors. We assessed three goblet cell mediators including Muc2, the pro-inflammatory mediator Relmβ, and the reparative TFF3 and at the gene transcript level, all three were similar between IEC-Myd88 -/- and WT mice under uninfected conditions. However by D1 pi the 77  transcript levels for all three factors were significantly lower in IEC-Myd88 -/- mice when compared to WT mice (Figure 2.7E-G, *p<0.05, **p<0.005).    Figure 2.7: Gene transcripts for antimicrobial peptides and goblet cell mediators in WT and IEC-Myd88-/- mice Infection of WT mice led to increased gene transcript levels for RegIII- γ (A) (significant) and RegIII-β (C), not significant, in cecal tissues by D1 pi, as compared to IEC-MyD88 -/- mice. β-defensin transcript levels (B) were similar between the two groups.  D) Ileal RegIII- γ levels under uninfected conditions are significantly lower in IEC-MyD88 -/- mice.  IEC-MyD88 -/- mice were also impaired in gene transcription levels in cecal tissues for the goblet cell mediators Relmβ (E), Muc2 (F) and TFF3 (G) at D1 pi with ΔaroA S. Typhimurium, in comparison to WT mice.  Error bars represent SEM from three independent experiments (at least 9 mice per group).  Asterisks indicate significant differences at *p<0.05 and **p<0.005.       78  Based on the impaired transcriptional induction of goblet cell specific factors in IEC-MyD88-/- mice, along with the ability of Relmβ to induce antimicrobial genes(311), i.e. the RegIII proteins, and the role played by Muc2 in generating the protective mucus barrier(312), we wanted to further assess changes in these factors during infection. We used immunostaining to determine whether Relmβ and Muc2 were differentially expressed in the two mouse strains at the protein level during ΔaroA S. Typhimurium infection.  Whereas similar levels of Relmβ staining was seen in uninfected cecal tissues of WT and IEC-MyD88-/- mice (Figure 2.8A), by D1 up to D7 pi there was a significant increase in detection of this protein within the goblet cells in WT mice, whereas there was little to no positive staining seen in tissues from infected IEC-Myd88 -/- mice (Figure 2.8B-D).    To semi-quantify the expression of these proteins, Relmβ fluorescence intensity relative to DAPI in each section was assessed using the ImageJ Integrated Density Analysis tool.  By D1 pi, there was 2-fold greater fluorescence intensity of Relmβ (relative to DAPI) in WT tissues compared to IEC-Myd88 -/- tissues. This increased to 6-fold by D3, which continued (and was significant) at D7 pi (Figure 2.8I).  Similarly, although comparable Muc2 staining was observed in uninfected WT and IEC-Myd88 -/- tissues (Figure 2.8E), by D1 up to D7 pi, WT tissues showed significantly increased Muc2 staining compared to IEC-Myd88 -/- mice (Figure 2.8F-H).  Fluorescence intensity of Muc2 relative to DAPI was significantly greater in WT tissues at D1 pi (4-fold increase, *p<0.05), as well as at D3 (2-fold, **p<0.005) and D7 (2-fold, *p<0.05) pi (Figure 2.8J). 79   Figure 2.8: Muc2 and Relmβ production is impaired in infected IEC-Myd88-/- mice  I J 80   Figure 2.8:  Muc2 and Relmβ production is impaired in infected IEC-Myd88-/- mice Representative immunostaining for the goblet cell specific factors (A-D) Relmβ (red) and (E-H) Muc2 (red) in uninfected, D1, D3 and D7 pi cecal tissues with DNA stained in blue.  Fluorescence intensity measurements for Relmβ (I) and Muc2 (J) relative to total DNA staining using ImageJ software revealed WT tissues had significantly greater Relmβ and Muc2 positive staining at D1, D3 and D7 pi. Bars represent the average fluorescence intensity values with three measurements per mouse, with at least 7 mice per group from 3 independent experiments.  Asterisks indicate significance at *p<0.05 and **p<0.005. Original magnification: 200X.   2.3.6 IEC-Myd88-/- mice also show early susceptibility to C. rodentium infection While our results clearly showed that IEC-Myd88 -/- mice display increased susceptibility to early stages of S. Typhimurium infection, in an earlier study we had found that MyD88 signalling within IECs played little role in regulating host responses to C. rodentium infection(185). Notably the earliest time point we had studied with these mice was D6 pi. We therefore examined whether IEC-Myd88 -/- mice showed any defects in promoting host defense against C. rodentium at an earlier time point, i.e. at D4 pi.  Following euthanization and removal of the entire large bowel, IEC-Myd88 -/- mice showed increased macroscopic damage in the form of shrunken ceca and shortened colons compared to WT mice (Figure 2.9A). Interestingly, no differences in pathogen burdens were found between WT and IEC-Myd88 -/- mice upon enumeration of C. rodentium in cecal tissue, luminal contents and colonic tissues of infected mice (Figure 2.9B).       To ensure consistency in our examination of responses, we once again focused our analysis on the cecum.  Although the ceca of WT and IEC-Myd88 -/- mice appeared similar under uninfected 81  conditions, at D4 pi the IEC-Myd88 -/- mice exhibited significantly increased histological damage scores compared to WT mice (Figure 2.9C&D). Specifically, IEC-Myd88 -/- mice displayed increased crypt hyperplasia, edema, and greater inflammatory cell infiltration at D4 pi (Figure 2.9D), but by D6 pi, pathology was similar between the two strains.  FITC-dextran was again used to assess intestinal barrier permeability, and although there was a greater increase in FITC-dextran serum levels in IEC-Myd88 -/- mice compared to WT at D4 pi, it did not reach statistical significance (Figure 2.9E).   82    Figure 2.9: IEC-Myd88-/- mice suffer accelerated tissue damage during C. rodentium infection  83  Figure 2.9:  IEC-Myd88-/- mice suffer accelerated tissue damage during C. rodentium infection A) After C. rodentium infection, IEC-MyD88 -/- mice displayed shrunken ceca compared to WT at D4 pi. B) No differences in C. rodentium tissue pathogen burdens (CFU/gram) were identified among the ceca, colons, or luminal contents from IEC-MyD88 -/- and WT mice at D4 pi.  C) Representative H&E staining of cecal tissues from D4 pi WT and IEC-MyD88 -/- mice.  IEC-MyD88 -/- mice exhibit increased edema, hyperplasia, and damage to IEC integrity at D4 pi. D) Comparative histological damage scores of uninfected, D4 and D6 pi IEC-MyD88 -/- and WT mice.  Cecal tissues of IEC-MyD88 -/- mice had significantly higher histological damage scores at D4 pi.  E) Both WT and IEC-MyD88 -/- mice experience increased barrier permeability at D4 pi.  Error bars indicate SEM from at least 6 mice, from ≥ 2 independent experiments. Asterisks show significance at **p<0.005, ***p<0.0005. Original magnification: 100X.  Consistent with our assessment of S. Typhimurium infection, we next examined whether goblet cell derived factors as well as antimicrobial responses were impaired in IEC-Myd88 -/- mice during C. rodentium infection.  Immunostaining revealed that at both D4 and D6 pi, WT tissues had greater Muc2 positive staining compared to IEC-Myd88 -/- mice (Figure 2.10A).  To examine the effects of these changes on C. rodentium localization during infection we stained tissues for the C. rodentium translocated effector Tir.  In WT tissues, most C. rodentium were found segregated within the cecal lumen at D4 pi, however at the same timepoint in IEC-Myd88 -/- mice, C. rodentium had successfully infected most of the cecal crypt epithelium (Figure 2.10B).  84   Figure 2.10: IEC-Myd88-/- mice show defects in Muc2 staining and altered C. rodentium localization in tissue A) Representative immunostaining for the goblet cell specific factor Muc2 (red) in uninfected, D4 and D6 pi cecal tissues with DNA stained blue.  IEC-MyD88 -/- mice have decreased positive Muc2 staining at D4 and D6 pi in cecal tissues compared to WT.  B) Immunostaining for C. rodentium effector Tir (red) and DNA (blue) at D4 pi reveals that lack of MyD88 signalling in IECs allows C. rodentium to more readily infect cecal crypts compared to WT mice, where they are sequestered to the lumen. Original magnification: 100X (A), 200X (B).  2.3.7 IEC-Myd88-/- crypt epithelial cells show decreased bactericidal activity Based on the impaired induction of goblet cell mediators and antimicrobial factors in IEC-Myd88 -/- mice, as well as much closer epithelial association of bacteria during both S. Typhimurium and C. rodentium infection, we examined whether the antimicrobial activity at the intestinal mucosal surface was impaired in these mice. We isolated crypts from uninfected IEC-Myd88 -/- and WT mice and collected their stimulated crypt supernatants to examine their bactericidal activity against ΔaroA S. Typhimurium and C. rodentium. As shown in Figure 2.11, relative to the 85  growth of bacteria exposed to the negative control of iPIPEs buffer alone, WT mouse crypt supernatants had direct bactericidal activity against Salmonella as well as C. rodentium, similar to that seen with purified RegIII-γ (positive control).  When incubated with supernatants from IEC-Myd88 -/- crypts, although some bactericidal activity was observed, it was significantly decreased against both bacterial strains when compared to WT supernatants (Figure 2.11, ***p<0.0005). Thus MyD88 signalling within IECs plays a key role in promoting antimicrobial defenses at the intestinal mucosal surface. Taken together, these results indicate that MyD88 signalling in IECs can actually determine host susceptibility to infection, potentially by controlling the ability of the pathogen to reach and survive at the surface of the intestinal epithelium and avoid subsequent accelerated intestinal tissue damage.   Figure 2.11: Crypt epithelial cells lacking MyD88 have impaired antimicrobial capacity Cecal crypts isolated from WT mice possess greater bactericidal activity against ΔaroA S. Typhimurium and C. rodentium than crypts from IEC-MyD88 -/- mice.  Results are plotted as average growth of bacteria relative to the negative control iPIPES buffer (100% growth) with 20 µM RegIII-γ incubation presented as a positive control, representative of 2 independent experiments with 3 mice per group. Error bars indicate SEM, Asterisks show significance at ***p<0.0005.   86  2.4 Discussion MyD88 dependent signalling has been previously shown to play a critical role in host defense against a number of bacterial pathogens(4, 7, 299), however its role in the GI tract appears to be unique, not only in controlling pathogen burdens, but also in limiting pathogen access to mucosal tissues and limiting the tissue damage resulting from these infections. As many of these protective MyD88 dependent responses involve changes in the level of proliferation or function of the IEC layer, it was of significant interest to clarify whether these changes arose from intrinsic MyD88 signalling within the IECs themselves.  In this study we clearly showed that loss of MyD88 signalling in IECs leads to increased susceptibility to infection and to worsened intestinal tissue damage at the early stages of infection by two different bacterial pathogens. Mice lacking MyD88 signalling within their IECs were impaired in their ability to maintain barrier integrity or upregulate the expression of goblet cell or antimicrobial mediators during infection, allowing enteric pathogens to come into closer proximity with the intestinal epithelium and more rapidly cause infection.  Through crypt killing assays we found that crypts from mice lacking IEC-MyD88 signalling displayed significantly decreased antimicrobial capacity against both S. Typhimurium and C. rodentium These findings thus expand our understanding of the role played by innate IEC signalling during in vivo enteric bacterial infections.      Several previous studies have shown that the global loss of MyD88 signalling in all cell types dramatically increased susceptibility of mice to intestinal inflammation and tissue damage, leading to rapid mortality following challenge with DSS or C. rodentium (4, 7, 216, 296). This susceptibility was found to reflect the loss of microbial driven responses that promoted epithelial homeostasis and protection from epithelial injury. Striking defects observed in  87  Myd88 -/- mice included an inability to protect their IEC barrier function, resulting in exaggerated gut leakiness(4, 216). Moreover these mice were unable to undergo the typical protective increases in colonic epithelial cell proliferation that develop during colitis, leaving them unable to readily replace damaged cells, resulting in widespread mucosal ulceration. In the current study, we clarify that at least in part, it is MyD88 signalling within the IECs themselves that protects/promotes IEC barrier function during infection, whereas this signalling is not required for the induction of increased IEC proliferation. In contrast, our results suggest that the proliferative status of IECs is driven by MyD88-dependent signalling by other cell types, rather than IECs themselves.  There are three main subsets of IECs present within the large bowel, namely enterocytes (80%), goblet cells (20%), and enteroendocrine cells (~1%). The intestinal epithelial monolayer acts as a physical barrier between the external environment, including the luminal microbes, and host tissues(300).  In addition, a thick mucus layer is present, composed primarily of mucins secreted by goblet cells, which separates potentially hazardous luminal bacteria from the mucosal surface to avoid spontaneous and/or maladaptive inflammatory activation.  The underlying enterocytes and goblet cells produce several types of antimicrobial peptides (RegIII-γ, β-defensin-II, RegIII-β, mCRAMP), as well as tissue protective factors (Relmβ, Muc2, TFF3). Many of these proteins are thought to localize within the overlying mucus layer, at an optimal location to impact invading pathogens, or repair damage to underlying tissues. At least in the small bowel, it has been shown that the mucus layer and the apical release of antimicrobials create a sterile zone overlying the epithelium that helps limit contact between the microbiota and the epithelial barrier(158).  Impaired function of this barrier, either due to reduced mucus production or 88  dysfunctional antimicrobial responses can potentially allow bacteria to reach the epithelial surface where they can trigger detrimental and prolonged inflammatory responses. Our results suggest that MyD88 also regulates this zone of separation in the large bowel, at least during infection, and that in its absence, S. Typhimurium and C. rodentium were able to achieve much closer association with the mucosal surface and more quickly invade tissues, causing more rapid tissue damage.    Breakdown of this zone of separation resulting in closer association of the pathogen with the epithelium may in part be driving the increased histological damage observed in the IEC-MyD88-/- mice at early time points, as pathogen burdens were similar between the two groups.  Upon initial observation these similar pathogen burdens were a surprise as WT mice had significantly greater gene transcript levels of several AMPs, goblet cell factors and crypt killing ability as compared to IEC-MyD88-/- mice.  However these IEC products may play a greater role in limiting pathogen proximity to the epithelium rather than overall burdens.  Interestingly, a recent study using a knock-in mouse model to express functional MyD88 only within IECs found that they harboured similar pathogen burdens to complete MyD88-/- mice, suggesting that MyD88 signalling in other cell types plays a greater role in controlling C. rodentium burdens(313).  Further, the AMPs that were decreased in the IEC-MyD88-/- mice during infection (such as RegIII-γ and -β) are thought to more readily target Gram-positive bacteria, which may have a greater effect on the resident bacteria rather than the Gram-negative pathogens tested in vivo.  Our results also noted overt impairment in the infection-induced expression of goblet cell mediators in IEC-Myd88 -/- mice as compared to WT mice. At present it is unclear whether these 89  results indicate that MyD88 signalling in IECs preferentially controls goblet cell responses or alternatively whether they are simply the easiest changes to detect in our system. As goblet cells have a secretory phenotype, immunostaining techniques allowed us to easily visualize the changes occurring in several goblet cell specific mediators, however these techniques are not as easily employed to assess changes in antimicrobial factors.  Future work using mice lacking MyD88 signalling within the goblet cell subset alone would aid in clarifying the role of goblet cell MyD88-dependent innate signalling in conferring host susceptibility to infection.    It is also notable that IEC intrinsic MyD88 signalling appears to only play a significant role in host defense at early stages of infection. When we previously assessed the role of MyD88 signalling in IECs during C. rodentium infection at a later time point (D6 pi), we were unable to identify a significant defect in the protective host responses elicited during colitis(185).  This was attributed to the previously described innate hypo-responsive status of IECs that is maintained by negative regulators, including Single Ig IL-1 related receptor (SIGIRR), a negative regulator of TLR and IL-1R signalling(185, 301).  However, we report here that during the initial stages of enteric infection in vivo, MyD88 signalling within IECs plays a critical role in maintaining barrier integrity and limiting tissue damage through induction of several goblet cell specific protective factors.  We hypothesize that this early role for MyD88 signalling in IECs is lost at later time points due to increasing involvement of MyD88 signalling in infiltrating inflammatory cells such as macrophages which likely also regulate IEC proliferation and function.   90  Although we found no differences between WT and IEC-Myd88 -/- mice under uninfected conditions, a recent study published by Frantz et al. reported baseline differences in the expression of RegIII-γ and Muc2.  Our inability to detect these differences may simply reflect differences in the resident microbes present in the different facilities. Correspondingly, Frantz et al, noted baseline differences in the microbe composition of IEC-Myd88 -/- mice compared to WT mice after extensive and detailed analysis.  They also observed a baseline increase in the translocation of luminal bacteria as well as compromised epithelial barrier integrity in the IEC-Myd88 -/- mice.  In contrast, we noted only modest differences in bacterial composition between strains at the phylum level and using the FITC-dextran permeability assay, found similar barrier integrity in our IEC-Myd88 -/- mice and WT mice.  The lack of baseline differences noted in our facility may reflect our practice of co-housing IEC-Myd88 -/- mice - and WT (MyD88flox/flox) mice since bringing them into our facility (>10 generations co-housed), likely minimizing strain specific differences in resident bacteria.   In conclusion, our findings illuminate a novel role for IEC-specific MyD88 signalling in promoting host defense and tissue tolerance during early enteric infection, one that may have been previously overlooked due to the concept that IECs are innately hypo-responsive, particularly in vivo. We also demonstrate that MyD88-signalling may be particularly important in a certain subset of IECs, goblet cells, to limit significant early tissue damage and clarify which MyD88-dependent protective responses require IEC-intrinsic signalling. Notably, although conflicting results on the innate responsive status of IECs in vitro have been reported(314-316), we show here for the first time that MyD88-dependent innate signalling within IECs can actually determine host susceptibility to an enteric bacterial infection.   91  Chapter 3: Milk fat globule membrane supplementation in formula modulates the neonatal gut microbiome and normalizes intestinal development  3.1 Introduction During gestation, the GI tract is immature, and possesses limited functions as most nutrients are obtained via placental transfer. Following birth, there is a switch to nutrient acquisition from ingested food, with a corresponding maturation of the intestine and significant change in the intestinal luminal environment. The source, makeup as well as the quantity of these nutrients are important in overall development of the infant, and can act locally in regulating the maturation of the intestine.  Moreover, these nutrients also likely impact the makeup and development of the gut microbes that colonize the neonate’s GI tract(262, 282, 283).  Using rodent, piglet and avian models, several groups have explored postnatal intestinal development from birth to one year of age, revealing major changes in epithelial architecture along the GI tract(95, 266, 269, 270, 317).  Within the small intestine, crypt depths and villus lengths significantly increase during the first month, and the number of crypts within the large intestine doubles(266).  Others have reported increases in innervation of submucosal ganglia(317), changes in localization of epithelial TJ proteins(95), and significant increases in goblet cell numbers and mucins(269, 270) during the neonatal period.  Proper intestinal development clearly facilitates the overall development of the neonate, but is also important in providing appropriate defense against noxious stimuli that pass through the intestine.  This is particularly important for premature babies who are born with an immature GI tract, leaving 92  them highly susceptible to infections and NEC, a leading cause of GI morbidity and mortality in premature infants(258).   Neonate development is fueled by breast milk, the ideal nutrient source during this stage of life.  Further, breastfeeding has been reported to lower risk of infection and diarrhea(268, 271, 278, 318), and to protect against development of asthma, allergies and immune mediated diseases(262, 271, 277, 319).  Its protective functions have been attributed to antibodies, enzymes (e.g. lysozyme, alkaline phosphatase) and growth factors (e.g. TGF-βand insulin like growth factor)(263, 320).  Unfortunately, breast milk is often unavailable in sufficient quantities, if at all, to satisfy the nutrient requirements of newborns, particularly with premature births. In addition, the prevalence of breastfeeding, although increasing, is negatively impacted in different regions of the world by hygiene, as well as social and economic status(262).  As a result, formula feeding has taken on a significant role in neonatal development and health care(321). Therefore, we and others have proposed that optimal formula composition should match that of breast milk as closely as possible(280, 283).   The lipid fraction of breast milk, representing a major energy source for the newborn, is composed of a triglyceride core surrounded by a unique triple membrane structure: the MFGM(275, 280).  MFGM, derived from the mammary gland epithelium, is composed primarily of polar lipids with interspersed membrane-bound proteins, glycoproteins, enzymes and cholesterol resulting in a bioactive molecule that likely confers some of the protective effects of breast milk(282, 283).  Most available infant formulas do not contain MFGM, but rather derive their lipids from vegetable sources, which differ greatly in size (1/10th the diameter) and 93  composition(280). Specifically, MFGM and vegetable derived lipids differ in triglyceride composition and internal structure, while the bioactive molecules present in MFGM are largely absent from formula lipids(280).  Recent breakthroughs in manufacturing technologies permit the concentration of bovine MFGM, making it feasible to add into infant formula.   Previous studies examining MFGM supplementation in piglets and human infants have predominantly focused on neurodevelopment, with its addition increasing cognitive scores compared to control formula, and similar to those of breastfed infants(281, 322, 323). Interestingly, a study examining the incidence of acute otitis media (AOM) and antipyretic use in human infants found that MFGM supplementation in formula resulted in decreased AOM and fewer days with fever compared to infants consuming control formula(324).  Additional studies using rodent models (>6 weeks old) have examined the effects of MFGM supplementation on infection and inflammation in vivo and in vitro (325-327). Components of MFGM display in vitro bactericidal activity against several foodborne pathogens, including Campylobacter jejuni, Salmonella enteriditis, and Listeria monocytogenes(325). In vivo, rats supplemented with MFGM and then infected with L. monocytogenes were protected against pathogen colonization and translocation(325).  During lipopolysaccharide-induced systemic inflammation in mice, MFGM supplementation significantly reduced gut barrier disruption and inflammatory cytokines(327).  Finally, in a rat model of dimethylhydrazine induced colon cancer, MFGM offered protection from aberrant crypt foci development, as compared to diets containing corn oil as their fat source(326).   94  Given the aforementioned protective effects of MFGM, we hypothesized that its supplementation in formula might prove beneficial within the developing intestine. To test this, we utilized the unique pup-in-a-cup model of artificial rearing, allowing exclusive formula feeding of rat pups starting at postnatal (pn) day 5.  Pups were provided either control (CTL) formula, with fat derived exclusively from vegetable sources, or with an identical formula with MFGM comprising part of the fat component. Rat pups left with mothers, and fed mother’s milk (MM), served as positive controls. Interestingly, rats fed CTL formula showed delayed intestinal growth as compared to MM fed pups at pn day 15. Notably, the addition of MFGM normalized most readouts to the levels seen in MM littermates, including intestinal crypt depths, epithelial cell proliferation, makeup of IEC subsets, and similar make-up of intestinal microbes at the phylum level. Lastly, upon challenge with Clostridium difficile toxins, MFGM supplementation afforded significant protection from mucosal damage as compared to CTL rat pups. Our study thus demonstrates that MFGM supplementation promotes intestinal epithelial and microbiome development and confers significant protection against noxious inflammatory stimuli.    3.2 Experimental procedures 3.2.1 Animals and provision of formula Pregnant female Sprague Dawley rats at 12 days of gestation were ordered from Charles River Laboratories (Wilmington, MA, USA).  Pregnant females were housed individually in sterilized, filter-topped cages and fed autoclaved food and water under specific-pathogen-free conditions at the BC Children’s Hospital Research Institute (BCCHRI).  At pn day 5, rat pups from each litter were randomly assigned to three different formula supplementation groups (control formula, or formula containing either 1.2 or 6 g/L bovine MFGM), with age-matched mother milk (MM) 95  reared littermates used as reference controls for intestinal development. In brief, gastrostomy feeding of the formula groups commenced on pn day 5 via PE 10 polyethylene tubing cannulas inserted into the stomach, as previously described in detail(328).  Pups were anaesthetized using halothane for the duration of the cannulation procedure. Gastric cannulas were connected to PE 50 tubing attached to Ismatec peristaltic pumps, which delivered volume controlled amounts of formula over 24 hours (2 cycles/hour, 10 minutes on, 20 minutes pause) to pups housed in an incubated water bath to maintain body temperature (40 to 42°C, depending on pup age).  The protocols employed were approved by the University of British Columbia’s Animal Care Committee and were in direct accordance with guidelines provided by the Canadian Council on the Use of Laboratory Animals.  3.2.2 Milk fat supplementation Cannulated rat pups were provided either a control (CTL) formula rat’s milk substitute (details of formula provided in Table 3.1) reflecting the vegetable derived fat source currently available in infant formulas, or alternatively, formula supplemented with bovine Milk Fat Globule Membrane (Lacprodan MFGM-10®, kindly provided by Arla Foods Ingredients (Aarhus, Denmark) and Mead Johnson Nutrition (Evansville, IN, USA)). MFGM concentrations used for this study were calculated based on the amount of phospholipid in breast milk, as MFGM is the sole source of phospholipids in breast milk. Human breast milk ranges from 3-4 % fat, therefore to reflect this content, a concentration of 1.2 g/L of MFGM was used.  As rat milk contains a higher % of fat (13-15%) a concentration of 6g/L of MFGM was also used, to more accurately reflect this fat content.  Formula was prepared and stored at -20°C, and thawed and mixed with a polytron prior to use. 96   Table 3.1: Composition of formulas Amounts are listed per 1L of formula preparation  3.2.3 Tissue collection Rat pups were anesthetized with isofluorane and euthanized via cervical dislocation from (1) MM, MFGM and CTL formula groups at pn day 15 (8-10 pups/group), (2) after 7 days of antibiotic treatment at pn d15 in the MM group along with untreated MM and CTL formula pups (5 pups/group), or (3) at pn day 15 following 2 hours of Clostridium difficile TcdA/TcdB toxin exposure (6-8 pups/group).  For histology and immunofluorescent staining, jejunal, ileal and distal colonic samples were fixed in 10% neutral buffered formalin (Fisher Scientific, Waltham, MA, USA) or Carnoy’s solution (60% ethanol, 30% chloroform, 10% glacial acetic acid).  Samples were allowed to fix overnight (formalin) or for 2 hours (Carnoy’s) at 4°C, transferred to 97  70% or 100% ethanol respectively, embedded in paraffin and cut into 5-µm sections.  Tissue sections with >50% of cross section intact on slides were included in immunofluorescent analysis, resulting in 3-4 images per tissue section assessed for 6-10 tissues per group.  Stool samples were also collected for intestinal microbe analysis.  3.2.4 Crypt depth and villus height measurements Hematoxylin and eosin stained paraffin sections from the jejunum, ileum, and colon were viewed under brightfield on a Zeiss Axio Imager microscope as outlined previously(161). Villus heights and crypt depths were measured in images using the measurement tool on AxioVision software at 200X magnification. At least 10 height or depth measurements were made per tissue section imaged.     3.2.5 FITC-dextran barrier permeability assay Barrier permeability was assessed as previously described(185, 329). Briefly, rat pups were gavaged at pn day 15 (4 hours before euthanization) with 12 mg of 4 kDa FITC-dextran in phosphate buffered saline. At sacrifice, blood was collected by cardiac puncture and added to 3% acid-citrate dextrose. FITC-dextran concentration in serum was measured using a fluorometer (excitation λ 485 nm, emission λ 530 nm).  3.2.6 Immunofluorescence Paraffin embedded sections (~5-µm thick) were deparaffinized by heating to 60°C for 15 min, cleared with xylene, followed by an ethanol gradient (75%, 95%, 100%) and water and steamed for 30 min in citrate buffer for antigen retrieval. Tissues were then treated with blocking buffer 98  (goat or donkey serum in PBS containing 1% bovine serum albumin [BSA], 0.1% Triton X-100, 0.05% Tween 20, and 0.05% sodium azide). The primary antibodies used were anti-Ki-67 (1:200, Thermo Scientific, Waltham, MA, USA), anti-Lysozyme (1:200, Santa Cruz, Dallas, TX, USA), anti-Muc2 (1:500, Santa Cruz), anti-CA-1 (1:100, Santa Cruz), anti-5HT (1:100, Antibodies Incorporated, Davis, CA, USA), anti-Claudin-3 (1:200, Invitrogen, Carlsbad, CA, USA), anti-Claudin-4 (1:150, Invitrogen). The secondary antibodies used (all 1:2000) were Alexa Fluor 568- or 488-conjugated goat anti-rabbit IgG, and Alexa Fluor 568- or 488-conjugated donkey anti-rabbit or donkey anti-goat IgG (Life Technologies, Carlsbad, CA, USA). ProLong gold antifade reagent with 4′,6-diamidino-2-phenylindole (DAPI; Invitrogen, Carlsbad, CA, USA) to stain DNA was used to mount tissues. Tissues were viewed on a Zeiss Axio Imager microscope, and images were taken using AxioVision software and an AxioCam HRm camera.  3.2.7 Fluorescence intensity measurements and cell counts To semi-quantify positive staining in tissue sections, the fluorescence intensity of immunostained samples was assessed using the integrated density measurement tool on ImageJ software.  For Muc2 (all sites) and CA-1 (small intestinal sections only) fluorescence intensity in each tissue section was presented relative to DAPI staining.  Integrated density for each image was assessed on separate channels to determine the pixel intensity of DAPI and Muc2 or CA-1 for each section. CA-1, Claudin-3 and Claudin-4 colonic sections were analyzed for fluorescence intensity at the top of crypts relative to the rest of crypts by using the selection tool prior to integrated density measurements on ImageJ.  Claudin-3 and Claudin-4 small intestinal sections were analyzed for fluorescence intensity in villi relative to crypts, again using the selection tool 99  prior to integrated density analysis.  Note that slides for all three groups from each experiment were stained at the same time and under the same conditions.  For Ki-67 cell counts, pictures were taken at 200X magnification for each stained tissue section under the same conditions.  The number of Ki-67 positive cells were then counted per crypt.  At least 4 crypts were counted per section (only in tact crypts in the cross section were counted).  5-HT positive cells were counted directly on the Zeiss Axio Imager microscope at 200X magnificent as a total number of positive cells for each whole cross section of tissue.    3.2.8 Intestinal microbe counts Total microbes in the colonic luminal contents were enumerated as previously described(161, 185).  In brief, following euthanization, colonic contents were collected, weighed, and homogenized in phosphate buffered saline. Homogenized samples were fixed in 10% Neutral Buffered Formalin to a final concentration of 3%. Samples were further diluted 1:10 in phosphate buffered saline, vortexed briefly, then 5 to 10 µl of these diluted samples were diluted in phosphate buffered saline to a final volume of 1 mL and filtered onto Anodisc 25 filters (Whatman International Ltd, Maidstone, UK) with a pore size of 0.2 µM and 2.5 cm diameter. Filters were then mounted onto glass slides with DAPI ProLong gold antifade reagent after drying and viewed at 1000X magnification on a Zeiss Axio Imager microscope. The number of DAPI-positive microbes were counted in 3 to 6 randomly chosen fields per disc, by two blinded scorers. The total number of intestinal microbes were calculated based on the mean of all counted fields per sample and the dilution factor used.  100  3.2.9 DNA extraction and 16S rRNA gene sequencing, processing and analysis Bacterial DNA extractions were performed as previously described in detail(330, 331).  Briefly, colonic contents were mechanically homogenized with glass beads (MoBio, Carlsbad, CA, USA) in 200 mM NaPO4, pH 8 and guanidine thiocyanate-EDTA-N-lauroyl sarcosine. Enzymatic lysis was performed for 1 h at 37 °C with 50 µl lysozyme (100 mg/ml), 50 µl mutanolysin (10 U/µl), and 10 µl RNase A (10 mg/ml), followed by another incubation for 1 hr at 65 °C after addition of 25 µl 25 % SDS, 25 µl Proteinase K (20 mg/ml,) and 75 µl 5 M NaCl. Supernatants were collected and DNA extracted with phenol-chloroform-isoamyl alcohol (25:24:1; Sigma, St. Louis, MO, USA) and purified using DNA Clean and Concentrator-25 columns (Zymo, Irvine, CA, USA) as per manufacturer’s instructions (purified DNA was stored at −20 °C).  PCR amplification of the V3 region of the 16S rRNA gene was performed as previously described(332), with the following modifications: a 50 µl reaction containing 1.25 mM MgCl2, 2.5 mM of each dNTP, 100 nM of each barcoded primer, and 0.25 U Taq was divided into 3 × 20 µl reactions for amplification. PCR conditions consisted of an initial denaturation at 94 °C for 2 min, 25 cycles of 94 °C for 30 s, 50 °C for 30 s, 72 °C for 30 s, followed by a final elongation at 72 °C for 10 min. Purified PCR products were sequenced using the Illumina MiSeq platform at the McMaster Genome Facility (Hamilton, ON, Canada). 16S rRNA gene sequence processing was completed as previously described (331, 333, 334). Sequences were filtered for quality with Sickle using a threshold of 30(335) and chimera checking was performed using USEARCH(336). OTUs were binned at 97 % similarity using AbundantOTU(337) and taxonomy was assigned using the Ribosomal Database Project (RDP) classifier(338) against the Greengenes reference database (4 February 2011 release)(339) using Quantitative Insights Into 101  Microbial Ecology (QIIME) (340). Unassigned OTUs and singletons were not included. The total number of sequence reads after filtering was 766, 232 (average 51, 082 reads/sample; range: 25, 673 – 71, 590 reads). Relative abundance taxonomic summaries, alpha diversity and beta diversity were performed using QIIME.  3.2.10 Antibiotic treatments Rat pups (MM reared) were treated with an antibiotic cocktail daily from pn day 8 through to 15 by oral gavage.  The cocktail consisted of 1 g/L ampicillin sodium, 1 g/L neomycin sulfate, 1g/L metronidazole and 0.5 g/L vancomycin hydrochloride administered at a volume of 10 mL/kg body weight.  Crypt depths, villus heights, Paneth cell numbers, Ki-67 positive cells and Muc2 fluorescence intensity for antibiotic treated groups were expressed relative to their untreated groups, as described above.    3.2.11 Clostridium difficile toxin induced intestinal inflammation C. difficile toxins TcdA and TcdB were kindly provided by Dr. Paul Beck, University of Calgary (Calgary, AB, Canada). Intra-rectal instillation of C. difficile toxins was performed as previously described(256, 341).  Briefly, a PE-50 polyethylene catheter was attached to a 19G needle on a 1 mL syringe and lubricated with a water-soluble tear gel lubricant (Novartis, Basel, Switzerland).  The catheter was inserted 2.5 cm into the colon of isofluorane anesthetized pups resting on a heating pad, and 50 μg TcdA/TcdB toxin in 100 μL phosphate buffered saline was slowly injected over 30 seconds.  Upon removal of the catheter, pressure was applied to the anal area for a further 30 seconds to prevent the diluted toxin from leaking out. Pups were placed in incubated water bath housing to recover, and observed for the following 2 hours until euthanization. 102  Histological damage was scored by 2 blinded observers using the following scoring criteria: (a) architectural changes: 0 = normal, 1 = vacuolation /blebbing, 2 = loss of epithelium, 3 = complete loss of crypt architecture; (b) inflammatory cell infiltrate: 0 = normal, 1 = increased inflammatory cell infiltrate in lamina propria, 2 = inflammatory cell infiltrate in submucosa, 3 = dense collection of inflammatory cells, but not transmural, 4 = transmural inflammatory cell infiltrate; (c) goblet cell depletion: 0 = no change, 1 = <50% depletion, 2 = > 50% depletion, 3 = >90% depletion. For each section assessed, parameters (a) and (b) were multiplied by the percentage of the colonic tissue section exhibiting changes to more accurately reflect the damage for each tissue.    3.2.12 Statistical analysis Results presented in this study, unless otherwise noted, were expressed as the mean value ± standard error of the mean (SEM) and analyzed using GraphPad Prism 6 for Mac OS X.  Shapiro-Wilk tests were performed on all groups, where possible, to determine normality prior to analysis.  Differences between means were calculated using Student’s t-tests or analysis of variance (ANOVA) (one- or two- way), followed by a post hoc Tukey test where appropriate.  If normality was not achieved, non-parametric Kruskal-Wallis or Mann-Whitney tests were performed to determine differences.  Statistical significance of phyla abundance was determined using ANOVA with Tukey’s multiple comparison test, while alpha diversity significance was determined using Kruskal-Wallis with Dunn’s multiple comparison. Statistically significant OTUs between MM, MFGM, 103   and CTL groups were identified using LEfSe(342). Adonis, implemented in QIIME, was used to determine statistical significance of groups using beta-diversity distances. A p value of <0.05 was considered statistically significant.  3.3 Results 3.3.1 Rat pup growth and gross intestinal characteristics To determine if MFGM supplementation altered overall growth of the animals, pups were weighed daily beginning at pn day 5 until day 15. There were no significant baseline differences noted in initial weights between the pups, and all groups displayed similar body weight after 10 days of supplementation (Figure 3.1A).  In addition, average daily weight gain was similar between the three formula groups (Figure 3.1B). Upon euthanization at pn day 15, the overall length of the small intestine and liver weights were similar between all groups (Figure 3.1C&D).        104   Figure 3.1: Formula feeding does not alter overall growth of rat pups compared to MM fed by pn day 15 A) Body weights (g) of rat pups in the 4 different diet groups were similar after ten days of supplementation. B) Average daily weight gain from pn day 5 to 15 did not differ between pups supplemented with CTL formula, 1.2 g/L or 6 g/L MFGM.  Small intestinal length (C) and liver weight (D) were similar in the four groups at pn day 15. n>10, from 3 independent experiments.  3.3.2 Impact of MFGM on intestinal epithelial architecture and barrier As villus lengths and crypt depths are overt markers of intestinal health, they were next assessed to determine if intestinal architecture was impacted by MFGM supplementation. In the jejunum, CTL formula, MM and 6 g/L MFGM pups displayed similar villus lengths, with MFGM fed pups displaying a dose dependent increase in villus lengths in both the jejunum and the ileum (Figure 3.2A).  In the ileum, CTL formula, MM and 1.2 g/L MFGM pups displayed similar 105  villus lengths, while 6 g/L MFGM supplementation resulted in significantly longer villi at this site (Figure 3.2A).       Although the CTL formula group showed normal villus lengths, they displayed significantly shorter crypt depths as compared to MM and both MFGM groups in the jejunum and ileum (Figure 3.2B).  In the distal colon, CTL formula pups again displayed significantly shorter crypts than MM and both MFGM groups (Figure 3.2C).  A dose dependent increase in distal colonic crypt depths was also recorded in the MFGM fed pups, where 6g/L supplementation restored crypt depths to levels similar to MM pups. As these assessments noted greater effects on intestinal villus and crypt architecture following 6g/L MFGM supplementation, subsequent analysis focused on this dose.  Next, we examined the contribution of IEC proliferation to the changes observed in intestinal architecture by immunostaining tissues collected at pn day 15 for the nuclear proliferation marker Ki-67.  In both the jejunum and colon, MFGM and MM fed pups displayed similar Ki-67 positive staining, whereas the CTL pups had significantly fewer positive cells/crypt (Figure 3.2D-I).  In the ileum, similar numbers of Ki-67 positive cells/crypt were recorded in all three groups (Figure 3.2F-G).   106   Figure 3.2: MFGM (6g/L) supplementation in formula normalizes intestinal architecture and epithelial proliferation at pn day 15 A)  Villus lengths, measured in μm, in the jejunum (left) and ileum (right). Dose dependent increases in length in the MFGM supplemented group were observed. n=8-10. Crypt depth in the jejunum (left) and ileum (right) (B) and colon (C) with CTL formula fed pups showing significantly decreased depths compared to all other groups. n=8-10. Representative images of immunostaining for the proliferation marker Ki-67 (red) and DNA (blue) in the jejunum (D), ileum (F), and colon (H) of MM, 6 g/L MFGM and CTL formula fed pups, with corresponding quantification of number of Ki-67 positive cells per crypt in (E), (G) and (I). n>6. The graphed data presented are the mean ±  107  Figure 3.2: MFGM (6g/L) supplementation in formula normalizes intestinal architecture and epithelial proliferation at pn day 15 SEM from 3 independent experiments, analyzed by One-way ANOVA followed by Tukey’s multiple comparisons test. *p<0.05, **p<0.005, ***p<0.0005, ****p<0.0001. Original magnification: 200X.  To clarify whether formula feeding altered the intestinal epithelial barrier, we next immunostained for the TJ proteins Claudin-3 and Claudin-4. At all three sites (jejunum, ileum and colon), MM and MFGM fed pups displayed similar positive staining for Claudin-3, as compared to its limited staining in CTL formula littermates (Figure 3.3A-C).  At the small intestinal sites, Claudin-3 was broadly expressed along the entire length of the villi in MM and MFGM pups, whereas its staining was sporadic and limited to the crypts in CTL fed pups (Figure 3.3A&B). In the colon, MM and MFGM pups showed greater positive staining along the tops of the crypts compared to CTL fed pups (Figure 3.3C).  In contrast, Claudin-4 displayed similar positive staining in all three groups at both small intestinal sites (Figure 3.3D&E), however within the colon CTL fed pups displayed greater positive staining at the tips of crypts as compared to MM and MFGM groups (Figure 3.3F).  Claudin-3 and 4 staining was semi-quantified by examining their fluorescence intensity in villi relative to crypts at small intestinal sites and at the top of crypts relative to the rest of the crypts in colonic tissues, confirming the results observed in A-F (Figure3.3G-L).  Finally, to determine whether these differences in expression of TJ proteins had any functional impact, we assessed barrier permeability using the FITC-dextran assay.  Interestingly, no significant differences in barrier permeability were observed between the three groups at pn day15 (Figure 3.3M).   108  Figure 3.3: Localization of barrier proteins, Claudin-3 and Claudin-4, are similar in MM and 6 g/L MFGM pups  109  Figure 3.3: Localization of barrier protein, Claudin-3 and Claudin-4, are similar in MM and 6 g/L MFGM pups Representative images from jejunal (A, D), ileal (B, E) and colonic (C,F) tissues of pn day 15 MM, 6 g/L MFGM and CTL formula fed pups immunostained for the TJ proteins (red) Claudin-3 (A-C), Claudin-4 (D-F) and DNA (blue). n=5-8. Original magnification: 200X.   Fluorescence intensity ratios in villi relative to crypts in small intestinal sections (G-H, J-K) and intensity at top of crypts relative to the rest of crypts (I&L) at pn day 15, n>6, from 3 independent experiments.  (M) Assessment of intestinal barrier integrity using the FITC-dextran barrier permeability assay revealed similar permeability between MM, 6 g/L MFGM and CTL formula fed pups. n=4-6. The graphed data presented are the mean ± SEM, analyzed by One-way ANOVA followed by post hoc Tukey’s test, *p<0.05, ***p>0.0005, ****p>0.0001.     3.3.3 Changes in IEC subtypes Rapid changes in the development of the various IEC subtypes have been documented during early intestinal development in several animal models(109, 269, 270, 343, 344).  Therefore, we next examined the impact of MFGM supplemented formula on the development of IECs.  3.3.3.1 Paneth cells Paneth cells are a secretory cell type found at the base of crypts in the small intestine and a major source of antimicrobial peptides, such as defensins, cathelicidins and lysozyme.  During murine intestinal development, Paneth cells have been found in small intestinal tissues by pn day 7, with rapid increases observed in following weeks(109).  We examined Paneth cell numbers in the jejunum and ileum of our pn day 15 rat pups by immunostaining for lysozyme.  At both small intestinal sites, MM and MFGM supplemented pups displayed similar numbers of lysozyme-positive cells/crypt, whereas significantly fewer lysozyme-positive cells/crypt were identified in CTL formula tissues (Figure 3.4A-D).  110    Figure 3.4: Addition of 6g/L MFGM to formula normalizes Paneth cell numbers at pn day 15 Representative images of immunostaining for the Paneth cell antimicrobial protein Lysozyme (red) and DNA (blue) in the jejunum (A) and ileum (C), quantified as number of lysozyme positive cells per crypt in (B) and (D). White arrowheads indicate lysozyme positive Paneth cells. n>6, from 3 independent experiments. The graphed data presented are the mean ± SEM, analyzed by Kruskal-Wallis test (B, D) followed by multiple comparisons tests. **p<0.005, ***p<0.0005, ****p<0.0001. Original magnification: 630X.   3.3.3.2 Goblet cells Goblet cells are the most numerous secretory cell type found within the intestinal epithelium, where they synthesize and secrete mucins, such as Muc2, and other protective bioactive molecules(161, 167, 329, 345). Goblet cell numbers have also been found to significantly increase within the intestinal epithelium during postnatal development(269, 270).  We assessed goblet cells by immunostaining for Muc2, and measuring Muc2 fluorescence intensity relative to DAPI (nuclear stain). Staining in the small intestine showed individual Muc2 positive goblet cells scattered along the villus lengths and within crypts (Figure 3.5A&C). Although similar 111  levels of Muc2 positive staining were observed between the three groups in the jejunum, in the ileum, MM tissues displayed significantly more Muc2 positive staining than CTL formula tissues (Figure 3.5B&D). Interestingly MFGM tissues showed Muc2 levels similar to that seen in MM tissues (Figure 3.5A-D). In the colon, Muc2 staining was much greater than in the small intestine with a mean ratio of Muc2/DAPI positive staining for all three groups of 0.16 ± 0.01 in the ileum and 0.30 ± 0.025 in the colon (****, p>0.0001, Mann-Whitney test), reflecting the presence of numerous goblet cells. Interestingly, while Muc2 staining was modestly, but not significantly greater in the MM group, as compared to CTL tissues, MFGM tissues displayed significantly increased Muc2 positive staining as compared to both MM and CTL formula groups (Figure 3.5E-F).     112   Figure 3.5: Addition of 6 g/L MFGM to formula increases Muc2 positive staining at pn day 15 Representative immunostaining of Muc2 (red), with DNA in blue, in jejunal (A), ileal (C) and colonic (E) tissues of MM, 6 g/L MFGM and CTL formula fed rat pups.  Fluorescence intensity measurements for Muc2 relative to total DNA staining in sections in (B), (D) and (F). n>6, from 3 independent experiments. The graphed data presented are the mean ± SEM, analyzed by One-way ANOVA (B) or Kruskal-Wallis test (D, F) followed by multiple comparisons tests. *p=0.01, ***p<0.0005. Original magnification: 200X.   3.3.3.3 Enteroendocrine cells Enteroendocrine cells in small intestinal and colonic tissues were examined by immunostaining for 5-Hydroxytryptophan (5-HT). In the jejunum, similar amounts of 5-HT-positive cells/tissue cross section were observed in MM and MFGM pups, which was increased, though not significantly, compared to CTL formula pups (Figure 3.6A-B).  In the ileum this was reversed as the CTL formula fed group carried significantly more 5-HT-positive cells/tissue cross section 113  compared to MM (Figure 3.6C-D).  In both tissues, the addition of MFGM brought the numbers of 5-HT-positive cells/tissue cross section closer to that seen in the MM pups, although the numbers did not reach significance. Interestingly all three groups displayed similar numbers of 5-HT-positive cells/tissue cross section in the colon (Figure 3.6E-F).     Figure 3.6: Enteroendocrine numbers are similar in 6 g/L MFGM and MM pups Representative immunostained images for 5-HT (red) in the jejunum (A), ileum (C) and colon (E) to assess enteroendocrine cells with DNA in blue.  Similar numbers of enteroendocrine cells per tissue section were counted in the jejunum (B) and colon (F) of all three groups, with CTL formula fed ileal sections displaying increased 5-HT positive cells. n>6, from 3 independent experiments. The graphed data presented are the mean ± SEM, analyzed by One-way ANOVA followed by Tukey’s multiple comparisons test. *p<0.05. Original magnification: 200X.   114  3.3.3.4 Enterocytes Enterocytes were examined at the three intestinal sites by immunostaining for carbonic anhydrase-1 (CA-1), an electrolyte transporter(343, 344).  At both small intestinal sites, CA-1 staining patterns were quite similar between the three groups (Figure 3.7A-D).  However, in the colon, the CTL formula fed pups displayed a very distinct and thick layer of CA-1 positive staining along the surface of the crypts, whereas MFGM and MM pups displayed patchier positive staining (Figure 3.7E&F), potentially reflecting the presence of numerous non-enterocytes (i.e. goblet cells) in these pups.         115   Figure 3.7: Enterocyte staining is similar in 6 g/L MFGM and MM pups at pn day 15 Representative images of CA-1 (red) and DNA (blue) immunostaining to assess enterocytes in the jejunum (A), ileum (C) and colon (E). All groups displayed similar positive staining at SI sites, whereas in the colon MM and 6 g/L MFGM pups displayed similar staining patterns compared to CTL. Fluorescence intensity measurements for CA-1 positive staining relative to DAPI in whole sections (B&D) or at the surface of colonic crypts is presented (F).   n>6, from 3 independent experiments. The graphed data presented are the mean ± SEM, analyzed by One-way ANOVA followed by Tukey’s multiple comparisons test. *p<0.05. Original magnification: 200X.  3.3.4 Contribution of microbiota to intestinal development As many studies and reviews have highlighted the importance of intestinal microbes in postnatal intestinal development(2, 283, 346), we next assessed total microbe numbers in the feces of the three groups (Figure 3.8A).  Interestingly, MM pups harboured the greatest number of microbes  (5.5x1010CFU/g feces), at least 10-fold greater than MFGM and CTL pups, likely reflecting their continued contact with their mother and her microbiome.  CTL pups carried the lowest number 116  of microbes (6.2x109CFU/g feces), with MFGM pups carrying slightly more (9.7x109CFU/g feces) at pn day 15 (Figure 3.8A).  To further assess differences in the microbiome we next analyzed their makeup in the three groups using 16S rRNA gene sequencing, finding distinct differences between the three groups. At the phylum level, the MM and MFGM supplemented pups displayed similar abundances of Firmicutes and Proteobacteria, as compared to CTL formula fed littermates who carried significantly lower levels of Firmicutes and higher levels of Proteobacteria (Figure 3.8B). However, at the genus level, major differences in taxa between all three groups were detected, including greater populations of Enterbacteriaceae with CTL formula and increased Lactobacillus species found in MM fed pups(Figure 3.8C).  Upon examining the diversity of bacterial species present within each diet group, MFGM and MM pups displayed greater species richness and evenness within their samples than the CTL formula group; with CTL formula leading to a significantly decreased Shannon Diversity index as compared to MM pups  (Figure 3.8D).  Next, beta-diversity between the three groups was examined using Weighted Unifrac analysis of all OTUs and clustering of MM, MFGM and CTL formula pups was found to be significant, suggesting the three groups harboured a distinct pattern of taxa from one another (Figure 3.8E). 117    Figure 3.8: Food source impacts the composition of the intestinal microbiome    118  Figure 3.8:  Food source impacts the composition of the intestinal microbiome A) Intestinal microbes were enumerated in colonic luminal contents of MM, 6 g/L MFGM and CTL formula fed pups at pn day 15, with MM pups harbouring ten-fold greater microbes than formula fed groups. The graphed data presented are the mean ± SEM, analyzed by One-way ANOVA. n=5. Microbial communities in fecal pellets at pn day 15 were assessed by 16S rRNA gene sequencing at the Phylum (B) and Genus (C) levels. Comparison of relative abundance of bacterial taxa revealed similar abundance of Firmicutes and Proteobacteria in MM (blue) and 6 g/L MFGM (red) pups as compared to CTL (green) formula fed pups (B). Two-way ANOVA followed by Tukey’s multiple comparisons test.  (D) MM pups harboured microbiota with significantly higher richness and evenness than CTL formula pups (Shannon Diversity index, Kruskal-Wallis with Dunn’s Multiple Comparison) and clustering of MM, 6 g/L MFGM and CTL formula fed pups was significant (E) Weighted UniFrac adonis R2=0.6036, p = 0.001.  n=5 *p<0.05, ***p<0.0005, ****p<0.0001.   Finally, we performed LEfSe analysis to determine which OTUs were significantly different between the three groups (Figure 3.9).  Taking into account only the most abundant OTUs (greater than 1% relative abundance), we observed that Enterobacteriaceae were significantly higher in the CTL group, decreased in the MFGM and absent in the MM group. Lactobacilli were significantly greater in the MM group, decreased in the MFGM and absent in the CTL group. Additional OTUs that were significantly higher in the MM group included Lachnospiraceae, Ruminococcaceae, Blautia and Parabacteroides. OTUs that were significantly increased in the MFGM group were Enterococcus, Clostridiales, Streptococcus and Morganella. 119   Figure 3.9: Differences in most abundant OTUs between the three groups LEfSs analysis to determine differences in OTUs (greater than 1% relative abundance) between the three groups, significantly different OTUs are shown. n=5. *p<0.05.  We next tested the impact of microbes on intestinal development, using an antibiotic cocktail to deplete the microbes in the MM group, which displayed the greatest abundance and diversity of 120  microbes, as well as in the MFGM formula group. Enumeration of fecal microbe numbers revealed a 10-fold depletion of microbes at pn day 15 in MM pups (4.4x109CFU/g feces) resulting in microbial numbers roughly similar to those observed in untreated CTL and MFGM formula fed pups (Figure 3.10A). In contrast, antibiotic treatment of MFGM formula fed pups only depleted intestinal microbes by 45%  (from 9.7x109 to 5.1x109CFU/g feces) (Figure 3.10A). Based on the limited depletion seen in the formula fed pups, further analysis regarding the impact of microbiome depletion on intestinal development was focused on the MM antibiotic treated pups.   Notably, antibiotic treatment of MM pups did not cause any significant changes in body weight, or small intestinal/colonic length as compared to untreated MM littermates (Figure 3.10B-D). To test whether the loss of microbes had any impact on intestinal development, villus lengths in both the jejunum and ileum were measured. They showed a significant decrease in MM pups post-antibiotic treatment as compared to untreated MM littermates (Figure 3.10E). Interestingly, although crypt depths were significantly decreased at all three intestinal sites in antibiotic treated MM pups as compared to untreated MM littermates, the depths were similar to, if not higher than, those recorded in untreated CTL formula fed pups (Figure 3.10F&G). Similar to untreated MM littermates, untreated MFGM crypt depths were significantly greater than antibiotic treated pups (jejunum: MM antibiotic: 59.7µm ± 1.7, MFGM: 82.2µm ± 2.8 **** (p<0.0001), ileum: MM antibiotic: 55.9µm ± 2.4, MFGM: 71.8µm ± 2.8 *** (p=0.0001), colon: MM antibiotic: 170.2µm ± 4.1, MFGM: 190.2µm ± 4.6 **(p=0.003), n=5, Student’s t-test).  121   Figure 3.10: Microbes of MM pups can be efficiently depleted with antibiotic exposure resulting in changes to the intestinal architecture A) Daily oral gavage of 10 mL/kg antibiotic cocktail from pn day 8 to 15, led to a significant decrease (>90%) of intestinal microbes in MM fed pups, compared to approximately 45% depletion in 6 g/L MFGM fed pups.  Weight (B), small intestinal length (C) and colon length (D) are similar between antibiotic treated and untreated MM fed pups at pn day 15. Quantification of cell proliferation (D) as number of Ki-67 positive cells per crypt, Paneth cells (E) as Lysozyme positive cells per crypt and (F) Muc2 fluorescence intensity relative to DNA in the jejunum, ileum and colon of Antibiotic cocktail treated and untreated MM pups.  Antibiotic treatment led to a decrease in cell proliferation at all sites, decreased Paneth cells in the ileum, and an increase in Muc2 intensity in the colon. Measurements of untreated CTL formula fed pups are presented as a reference (☐).  n=5 The graphed data presented are the mean ± SEM, analyzed by unpaired Student’s t-test. *p<0.05, **p<0.005, ***p<0.0005, ****p<0.0001.  We next examined epithelial cell proliferation by again immunostaining intestinal tissues for Ki-67. Antibiotic treatment resulted in significantly fewer Ki-67-positive cells/crypt in MM pups at all three sites when compared to untreated littermates (Figure 3.11A).  When compared to CTL 122  formula supplemented pups, antibiotic treatment of MM pups resulted in a similar number of Ki-67 positive cells at all three sites (Figure 3.11A).   IEC subtypes were again quantified by immunostaining tissue sections, to assess the effects of antibiotic treatment.  No differences in Paneth cell numbers with antibiotic treatment were noted in the jejunum (Figure 3.11B), however, in ileal tissues treated MM pups showed a significant reduction in Paneth cells compared to untreated pups, similar to numbers observed in CTL formula fed pups (Figure 3.11B).  Assessment of goblet cells in the jejunum and ileum revealed MM antibiotic treated pups had similar levels of Muc2 fluorescence intensity when compared to untreated MM pups, whereas in the colon MM pups surprisingly displayed an increase in Muc2 intensity after exposure (Figure 3.11C).       123   Figure 3.11: Intestinal development is partially dependent on the presence of intestinal microbes in MM pups Quantification of cell proliferation (A) as number of Ki-67 positive cells per crypt, Paneth cells (B) as Lysozyme positive cells per crypt and (C) Muc2 fluorescence intensity relative to DNA in the jejunum, ileum and colon of Antibiotic cocktail treated and untreated MM pups at pn day 15.  Antibiotic treatment led to a decrease in cell proliferation at all sites, decreased Paneth cells in the ileum, and an increase in Muc2 intensity in the colon. Measurements of untreated CTL formula fed pups are presented as a reference (☐).  n=5 The graphed data presented are the mean ± SEM, analyzed by unpaired Student’s t-test. *p<0.05, **p<0.005, ***p<0.0005, ****p<0.0001.  3.3.5 MFGM supplementation protects against Clostridium difficile toxin  While MFGM supplementation positively impacts the structure and makeup of the neonatal intestinal epithelium, the effect of these changes on overall gut protection was unclear. Neonates are vulnerable to a variety of enteric infections, as well as idiopathic causes of intestinal inflammation such as NEC. Unfortunately there are few models for these conditions that have been established in rat pups. To overcome this, we tested the susceptibility of pups to C. difficile toxin induced colitis through intra-rectal challenge, as previously described in adult mouse models(256, 341).  As in the mouse model of toxin induced colitis, 2 hours of exposure in pn day 15 rat pups resulted in mucosal inflammation as evidenced by inflammatory cell infiltration, 124  submucosal edema, goblet cell depletion and breakdown of epithelial architecture compared to unchallenged littermates (Figure 3.12A-B).  Furthermore, C. difficile toxin exposure led to significant colonic shortening in the formula fed groups as compared to unchallenged groups (Figure 3.12C-D), with more pronounced shortening in the toxin treated CTL formula group, significantly more than that in toxin challenged MM pups.  Assessment of histological damage by blinded observers revealed significantly greater overall damage scores in the CTL formula group after toxin challenge when compared to both the MFGM and MM pups (Figure 3.12E). Thus, supplementation of formula with MFGM yields a state of protection against the colitis caused by C difficile toxins.  125   Figure 3.12: MFGM supplementation protects the formula fed neonate intestine from C. difficile toxin induced damage  Representative hematoxylin and eosin stained distal colonic sections from MM, 6 g/L MFGM and CTL formula fed pups under (A) unchallenged conditions, and (B) after intra-rectal exposure to C. difficile toxins TcdA and TcdB for 2 hours at pn day 15. C) Representative macroscopic images of toxin induced colonic shortening and inflammation, quantified in (D). E) Comparative histological damage scores after 2 hours toxin exposure in the three groups. n=6-7, from 2 independent experiments. The graphed data presented are the mean ± SEM, analyzed by One-way ANOVA followed Tukey’s multiple comparisons test. *p<0.05, **p<0.005. Original magnification: 200X.    126  3.4 Discussion Breast milk is considered the optimal food source for neonates, however it is not always available to developing infants.  Manufacturing formulas that more closely reflect the composition and function of breast milk is therefore of great importance to optimize outcomes among formula fed infants.  In this study we tested the effect of formula supplemented with bovine MFGM, which contains similar components to human MFGM, on intestinal development. By utilizing the pup-in-a-cup model, we showed that although CTL formula led to impaired intestinal development as compared to MM fed pups, the addition of MFGM to formula beneficially aligned intestinal readouts to levels similar to those observed with MM.  Tissues were examined at pn day 15, as previous studies have reported significant increases in villus lengths and crypt depths by this time point during development (347, 348).  In addition, due to the size restriction of rat pups in cups, studies utilizing this model have generally terminated on pn days 12-18 (328, 347-349).  As no differences were seen in weight gain between the three diet groups, and the CTL and MFGM formula fed groups were administered identical volumes of formula daily, the differences noted in intestinal mucosal development are not due to differences in the volume of food consumed or to differences in weight gain.    Through histological and immunofluorescence analysis, we found that MFGM supplementation led to similar intestinal mucosal architecture at small intestinal and colonic sites to that seen in MM pups.  In addition, MFGM supplementation led to similar numbers of proliferating cells, Paneth cells, and goblet cells as those observed in MM raised pups.  The increase in secretory cells (Paneth cells, goblet cells) with MFGM supplemented formula compared to CTL is noteworthy as previous studies have reported significantly decreased numbers of these cell types 127  in the resected tissues of children with NEC, as compared to children with other non-inflammatory colonic diseases(142).  In addition, MFGM and MM pups displayed similar TJ protein staining at all intestinal sites, whereas CTL formula fed pups showed altered staining patterns for Claudin-3 at all three sites, and Claudin-4 in the colon compared to the other two groups.  Upon assessing the effect of these alterations on barrier function, using the FITC-dextran assay, no differences in permeability were observed; suggesting the differences in TJ protein staining in the CTL formula group did not result in overt barrier disruption under homeostatic conditions.   In addition to serving as an important nutrient source, the composition of breast milk and/or formula can also impact the makeup of the gut microbes that colonize the neonate GI tract(262, 282, 283).  As the colon harbours the highest density of intestinal microbes, this may help explain why the effects of MFGM supplementation compared to CTL formula appeared greatest in the colon. In this study, we found that MFGM supplementation of formula altered the makeup of the gut microbiota, resulting in a microbiome that at the phylum level resembled that found in MM pups.  In contrast, CTL formula pups harboured a significantly higher percentage of Proteobacteria and a lower percentage of Firmicutes as compared to both MM and MFGM fed pups.  These results are similar to what some studies have found for phylum level differences in the microbiota of formula fed versus breastfed infants, with formula feeding resulting in higher Proteobacteria and lower Firmicutes(350-352). However, likely due to the fluctuations that occur in the gut microbiota in early life, other studies examining the effect of breast milk vs. formula feeding on the infant microbiome have reported anywhere from no major differences(353) to the reverse of those found in our studies at the phylum level(274, 283).   128  Upon assessing the Shannon diversity of the groups, we found that MM fed pups displayed increased microbial species richness and evenness when compared to CTL formula.  While formula feeding has previously been reported to increase alpha-diversity more than breastfeeding in the first months of life(274, 350, 351, 353, 354), this result may reflect that within our model, formula fed pups are separated from their mother at pn day 5 and no longer exposed to maternal microbes. Of note, however, is that the MFGM formula fed pups displayed similar species richness and evenness to their MM littermates.  As oligosaccharides within breast milk have been implicated in driving the diversity of the gut microbiome, components within the mammary epithelial membrane that form the outer membrane of the MFGM may also be involved. Assessment of lower taxonomic levels revealed differences in diversity between the MM, MFGM and CTL formula groups. It has previously been shown that formula feeding results in increased numbers of Clostridiales, Streptococcus, Enterococcus and Enterobacteriaceae(2, 51), which was also observed in this study, as these groups were higher in MFGM and CTL formula pups. Breast-feeding has been associated with increased abundance of Lactobacilli, which was highest in the MM group.  Notably, Lactobacilli were also detected in the MFGM group, although at lower levels, and were absent in the CTL group.    Interestingly, MM pups had significantly higher levels of bacterial groups commonly associated with the healthy adult gut microbiota, including Lachnospiraceae, Ruminococcaceae, Blautia and Parabacteroides in addition to harbouring 10-fold more intestinal microbes than formula fed littermates.  This likely reflects their housing (and close contact) with their mothers, siblings, and their fecal matter. Future studies will be necessary to define the significance of MFGM mediated alterations in the gut microbiome on intestinal development and health in the long term.  As 129  breast milk has been associated with not only early protection from infection and diarrhea, but also with long lasting protection against allergies, asthma and other immune mediated diseases(268, 278, 318, 319), it is of significant interest to clarify whether MFGM supplemented formula will also have the ability to confer these long lasting effects.         To examine the role of intestinal microbes in intestinal development, an antibiotic cocktail was used to deplete gut bacteria. Surprisingly, antibiotic exposure in MFGM formula fed pups only depleted 45% of microbes, potentially due to the effects of formula delivery via gastric cannula on gut motility, or perhaps because the formula fed pups carried a smaller number of starting microbes.  Due to this limited depletion in the artificially reared pups, we decided to focus our antibiotic studies on MM fed pups where antibiotic exposure led to a 10-fold depletion of intestinal microbes. Our analysis confirmed previous studies in germ free mice, that many aspects of intestinal development are partially dependent on the presence of gut microbes(134, 355).  Upon depletion of the gut microbes within MM fed pups, significant decreases in the previously mentioned histological parameters were observed, similar to those seen in untreated CTL formula fed littermates. Interestingly, although antibiotic treatment of MM pups also resulted in similar numbers of gut microbes to that carried by untreated MFGM formula fed pups, villus lengths and crypt depths were significantly greater in the MFGM fed pups. This suggests that in addition to exerting some of its beneficial effects via changes to the microbiome, MFGM likely contributes to normalizing intestinal development through additional mechanisms, potentially acting directly on the intestinal epithelium itself.    130  As diarrhea and enteric infections present at higher rates in formula fed infants, we next tested whether MFGM supplementation promoted a protective effect against intra-rectal C. difficile toxin challenge. There is growing evidence that C. difficile is associated with diarrhea and increased lengths of hospital stays in children under 2 years of age(250-252). C. difficile infections may be underreported in this population, as it is often not associated with the severe outcomes observed in infected adults such as colon resection and mortality(250). In our neonate model, C. difficile toxins induced a rapid colitis, albeit to a lesser degree than previously described in adult mice. Using this model we showed that the accelerated intestinal development in MFGM formula supplemented pups was protective, leading to significantly reduced colitis as compared to the CTL formula fed group. As Engevik et al., have recently reported that patients with C. difficile associated intestinal disease exhibit decreased Muc2 and an impaired mucus barrier(356), the ability of MFGM supplementation to increase Muc2 levels in the colon may be one mechanism by which MFGM conferred protection in this model.  Furthermore, intracellular and membrane proteins within MFGM comprise 2-4% of the proteins found within breast milk, with many of them having antibacterial or immune-modulatory functions(285).  Therefore, the proteins found within MFGM may also be contributing to the protection conferred in this model.     Taken together, this study has shown that ingestion of MFGM supplemented formula by rat pups normalizes many aspects of their intestinal development to levels similar to those seen in MM pups. We have also shown, for the first time, that addition of MFGM in formula promotes intestinal epithelial cell proliferation, TJ protein patterns, and development of IEC subsets (Paneth cells, goblet cells and enterocytes) to levels above that seen with CTL formula, and similar to those in MM fed pups. Interestingly, MFGM formula also impacted the intestinal 131  microbiota of formula fed pups, leading to a microbiome more similar to that of MM pups, which likely has important implications for long-term health.  Moreover, the addition of MFGM to formula protected pups against intestinal challenge with C. difficile toxins, as compared to the colitis suffered by CTL formula fed pups. From these results we suggest that MFGM supplementation in formula has many beneficial effects on intestinal development and promotes protection against noxious intestinal stimuli. Therefore its supplementation in formula may prove beneficial in human populations that have limited access to breast milk.   132  Chapter 4: Discussion Enteric infections and/or changes within the intestine after birth, such as the introduction of certain bacteria and formula versus breast milk, can impact later risk for the development of several diseases.  Development of irritable bowel syndrome (IBS) and reactive arthritis are complications that have been reported as long-term sequelae of acute infection by various enteric pathogens(224-226, 357, 358).  Similarly, increased susceptibility to the later development of allergies, asthma and several other inflammatory diseases has been reported for infants fed formula as compared to breast milk(262, 276, 277). The intestinal immune system and how it changes during these events is widely regarded to play a role in later susceptibilities.  However, one often overlooked cell type - the IECs, being in closest contact with these major luminal stimuli, are poised to serve as key players in shaping responses that maintain a homeostatic intestinal environment, as well as in modulating susceptibility to the development of diseases at later ages.  As such, we examined the intricacies of IEC-intrinsic innate immune (MyD88 dependent) mediated responses during the early stages of enteric infection using in vivo models in Chapter 2.  Following infection with S. Typhimurium and C. rodentium, we demonstrated that MyD88 signalling within IECs was required to limit pathogen penetration of intestinal crypts as well as mount appropriate goblet cell and antimicrobial responses to infection.  Further, lack of MyD88 signalling within IECs resulted in decreased bactericidal capacity of IECs, as studied using crypt epithelial supernatants.  In Chapter 3, intestinal development and optimal IEC maturation and responses were explored in formula feeding as compared to mother’s milk (MM).  In addition, the role of milk fats (MFGM) found specifically in mammalian milk was assessed for its ability to normalize intestinal development parameters similar to those observed with MM.  Interestingly, addition of MFGM to formula promoted intestinal development as well as the 133  maturation/differentiation of key IEC subtypes to levels similar to those seen in MM fed pups.  Susceptibility to intestinal damage by bacterial toxins revealed that the addition of MFGM to formula was protective against toxin-induced damage.  These studies aimed to detail the intricacies of IEC responses during enteric infection and development, with future studies proposed to examine how these responses may impact long-term health.    Much of the research conducted that aims to further our understanding of the intestinal epithelium and the intricate balance maintained with the luminal environment, including in this thesis, is done using rodent models.  Therefore a discussion regarding the suitability of rodent models in the study of intestinal health and disease is warranted and presented in the following section.   4.1 The suitability of rodent models for the study of the human intestine Since the times of Aristotle in ancient Greece, animal models have been used to further our understanding of human anatomy, physiology, and disease(359).  In more recent times, rodent models, particularly those utilizing mice and rats, have taken on a fundamental role in medical research for demonstrating biological significance.  Mice represent the most widely used organism in animal research studies today, with the number of publications using mouse models indexed on PubMed more than tripling over the last three decades(359). In contrast, studies using other mammalian models have remained relatively stagnant during that time with only slight increases or decreases for specific species. This is not surprising given that the laboratory mouse shares the majority of its protein-coding genes with humans(360) and is anatomically similar in the makeup and organization of its internal structures.  Many breakthroughs have also been made 134  in genetic manipulation of mice over the last century, with the first knockout mouse generated in 1987 and more than 7000 genetically defined strains of mice recognized to date(359, 361).  In addition, their small size, convenient housing arrangements, short reproduction time, and life span (1 mouse year is equivalent to approximately 30 human years) make mice an ideal and economical choice to study complex diseases.  Unquestionably, for the aforementioned reasons and many more, laboratory mice provide an indispensible research tool.  However, there are of course many ethical considerations associated with their use, with their capacity to feel pain of paramount concern.  This has resulted in the coinciding creation of regulatory bodies in virtually all jurisdictions employing animal models for research(362).  In Canada, this led to the formation of the Canadian Council on Animal Care (CCAC) in 1968, which sets the standards for the care and use of animals in research in this country(363).  CCAC guidelines aim to reduce the number of animals being used in research, as well as refine procedures to mitigate pain and stress incurred during experimentation.  Given the importance of well thought out experiments to justify the use of animals in research, a discussion highlighting similarities and differences between mice, rats, and humans, specifically in the GI system, is of interest to put into scope the significance of research done using these animal models.    The overall anatomy of the GI tract in mice, rats and humans is surprisingly similar.  Each organism’s GI tract is comprised of a mouth, esophagus, stomach, SI, LI and anus, with several supporting organs including similar salivary glands, the liver, gallbladder (absent in rats), and pancreas(364).  The stomachs of mice and rats, unlike in humans, are separated into two distinct 135  regions: the forestomach and the glandular stomach(12, 364).  The rodent forestomach is able to store well-chewed and partially digested food for upwards of three hours, emptying according to the host’s energy needs rather than the degree of filling. This controls for the bulk intake of food characteristic of rodent feeding in their natural environment allowing for a more steady state digestion.  The rodent glandular stomach is similar to the human stomach, which is connected to the duodenum of the SI.  The anatomical regions of the LI also differ slightly between humans and laboratory rodents(364).  The first portion of the LI (the cecum) is proportionally much larger in mice and rats in comparison to humans to facilitate the fermentation of indigestible plant materials that rodents consume much greater quantities of.  Due to their coprophagic lifestyle, much of what is produced in the rodent cecum is absorbed as ingested fecal pellets pass through their GI tract.  The remaining regions of the LI are relatively similar between mice, rats and humans.   With regards to the intestinal immune system, although there are many similarities between rodents and humans, slight differences do exist(28, 365).  For instance, the lymphoid tissues associated with the intestinal mucosa, such as PPs and ILFs, are present in varying numbers and differ in their development between mice and humans.  In the adult human ileum the concentration of PPs is much greater than that found in adult mice(365).  ILF development in mice occurs postnatally, though their presence has also been described in the human fetal intestine(366, 367).  PPs, on the other hand, develop in utero in both species.  Interestingly, mice display greater numbers of antigen sampling M-cells and antimicrobial peptides(365).  They also produce IEL B cells that express TLR4 and the IgD immunoglobulin isotype, whereas human IEL B cells tend not to(368).  As rodents have a coprophagic lifestyle with close proximity to the 136  ground, likely resulting in much great exposure to MAMPs, this may partially explain the presence of greater antimicrobial defenses at the level of the intestinal epithelium.   Though differences exist between mouse and human GI systems, it is important to note that the goals of their systems are the same: maintaining a state of controlled responsiveness to luminal antigens, tolerance to food and resident bacteria, and preserving the integrity of the gut barriers, all while the system absorbs adequate nutrients.  The extent of similarities within their systems results in accurate recapitulation of many characteristics of human intestinal disease in mouse and rat models.  Hallmarks of intestinal inflammation often observed in the intestinal biopsies of IBD patients such as crypt hyperplasia, depletion of secretory Paneth cells and goblet cells, a perturbed mucus layer, and decreased numbers and diversity of intestinal bacteria are also observed in chemical, bacterial, and T cell induced rodent models of colitis(217, 244).           In summary, mice display impressive genetic and biological similarities to humans providing an invaluable model organism that the research community is privileged to work with.  Although differences exist, the ability to utilize multiple models, in the context of intestinal disease research, allows for a robust analysis of the in vivo intricacies of epithelial and immune cell responses, inflammatory pathways, and infection dynamics.  When used in concert with other research tools, such as molecular biology techniques, intestinal organoid cultures, and large-scale human studies, mouse models are integral to the discovery and efficacious testing of novel therapeutics. 137  4.2 Pup-in-a-cup model: advantages and potential improvements In addition to mice, rats represent another well-used rodent model in research including studies exploring dynamics of infectious diseases, nutrition, and hypertension(359).  Though not as widely used as mice, rats also have a long history in modern biomedical research, particularly in nutrition research, with the first genetic knockout rat generated in 2009.  They have especially served as a valuable model for exploring the effects of newborn nutrition on health status and development since the introduction of the pup-in-a-cup model by Messer et al. in 1969.  There are many advantages to using the rat pup-in-a-model over those utilizing mouse pups, which are much smaller at the same age and therefore harder to handle and manipulate, or pigs, which have a much longer gestational length (115 days versus 21 days in rats) and larger size resulting in significantly greater costs(291).  However, there are also limitations to the model and potential improvements that can be made that would expand upon the studies conducted in Chapter 3, which are noted below.  Maternal separation is a requirement for the pup-in-a-cup model, and all other currently employed neonate nutrition models, to appropriately control nutrient intake and the pup’s environment.  Previous studies have highlighted the short and long term effects of maternal separation on inflammatory responses and adult sensitivity to pain(369).  The currently employed artificial rearing protocol does take steps to alleviate the stresses associated with separation such as housing the pups in a hot water bath to provide ample warmth and through constant handling, stimulation of urination and defecation, and grooming of the pups by researchers(370).  These actions help to mimic the tactile stimulation that would otherwise be received from the mother during these early stages of postnatal development.  Though increases 138  in inflammatory cytokines have been reported in some adult tissues (pn day 75) of artificially reared rats(371), we did not find any differences in intestinal tissues as measured by gene transcript levels for TNF-α or IL-1β in artificially reared and mother reared pups at pn day 15 using our model (Figure 4.1).  Further, other studies have shown that early maternal separation results in increased colonic permeability at 12 weeks of age, suggesting that early psychological factors may influence the functioning of the intestine later in life(372). As the pup-in-cup model provides a valuable tool to study the long-term effects of infant nutrition on health and susceptibility to disease, it is of interest to clarify whether our artificially reared pups also have altered baseline inflammatory cytokine levels or increased intestinal permeability as adults.  Moreover, it is prudent to consider steps that can be taken in future experiments to further mitigate the potential stress of separation for long-term studies.   Figure 4.1: qPCR analysis of rat pup colonic tissues at pn day 15 Gene expression levels of the inflammatory cytokines IL-1β and TNF-α in artificially reared MFGM and CTL formula fed pups relative to MM fed pups (kept with mother).  Artificially reared pups were separated from mother and housed individually starting from pn day 5.  No differences in gene transcript levels of the two inflammatory cytokines (IL-1β and TNF-α) were observed in the colonic tissue of pups separated from (MFGM and CTL) or kept with their mothers (MM) by pn day 15. qPCR Results 139  One such step may be to provide sensory stimuli that would normally be found in the mother’s nest.  Studies examining olfactory or odour cues, which are used by rats for social contact behaviour as early as pn day 5, have shown that neonatal rats will readily orient themselves towards and approach their home nest(373).  Not surprisingly, behavioural development in pups is linked to sensing of odours and chemical cues through the olfactory system(374).  Given this link, daily addition of bedding from the mother’s nest may beneficially impact long-term readouts that are proposed in the discussion of future studies (section 4.3.1).  Beneficial effects of daily bedding addition will include providing pups with adequate olfactory cues, supplemented by the current protocol of pup handling and stimulation by researchers, as well as providing an inoculum of bacteria from mother and siblings from the home nest.  This is especially important in long-term studies given the hypothesized importance of resident bacterial composition in shaping health.  4.3 Changes in the intestinal environment during enteric infection and neonate development– potential influence on later health In recent years, there has been growing evidence for an association between shifts in resident intestinal microbes and the presence/development of diseased states such as obesity, allergies, type II diabetes, autism spectrum disorders, and several autoimmune diseases.  For instance, in both mice and humans it has been found that stool samples from lean individuals contain a higher percentage of bacteria from the phylum Bacteroidetes, whereas obese samples have a higher percentage of Firmicutes(375).  These shifts can be influenced by diet along with host responses during the loss and re-establishment of homeostasis in the intestinal environment, such 140  as with the introduction of food after birth, enteric infections, antibiotic treatment, or the introduction of other environmental factors.  At birth, a major transition occurs in the intestinal environment where the initial introduction of substantial microbial communities (dependent on mode of delivery) has major effects on intestinal development. This includes proper education of the immune system along with maturation of IECs and the intestinal barrier, which are important steps in establishing intestinal homeostasis. The added variable of food, in the form of breast milk or formula, further alters the luminal environment and as a result, is able to influence the composition of the resident microbial community as well as influence intestinal maturation(2, 262, 274).  In addition, factors produced and released by the maturing IECs such as antimicrobial peptides and mucin proteins, which can serve as a food source for some bacteria, also influence the resident bacterial composition.  The long-term health implications of this multifactorial process have been a topic of interest in recent years, furthering our understanding of how differences in resident intestinal bacteria in infants can impact later health(2, 48, 281, 318).  For example, reports have identified differences in the resident intestinal microbes of infants who later go on to develop allergic disease, compared to those that do not.  Though inconsistencies between results in different studies exist, likely due to the great diversity and uniqueness of each individual’s microbiota, some interesting associations across several studies have been reported.  Low rates of Bifidobacteria and Lactobacilli during infancy and later development of allergy is one such association(376).  Interestingly, food sources (i.e. breast milk versus formula) and the composition of the intestinal mucus layer have both been shown to affect the presence of these specific microbes(2, 376, 377).  Not surprisingly, infants that are breastfed have been shown to 141  carry greater numbers of Bifidobacteria and Lactobacilli species as compared to formula fed infants.  Further, changes in intestinal mucus during post natal development and over the course of rotavirus infection (the most common cause of diarrhea in young children), impacts the ability of these bacteria to adhere to the mucus layer(378).     These studies demonstrate the ability of both environmental factors (food source) and IEC factors (mucus) to influence gut microbe composition and later development of disease (in this case allergies).   It is noteworthy to mention that other factors likely influence this process as well including mode of delivery, level of gestational development, enteric infections, antibiotic exposure, geographical location, and host genetics.   4.3.1 Future direction: exploring the long-term health consequences of MFGM supplementation in formula The work presented in this thesis provides valuable insight into the role of neonate nutrition, particularly the fat component of formulas, and its impact on intestinal epithelial development and susceptibility to toxin-induced intestinal injury.  However, this work also raises many questions as to what the long-term implications of these changes may be, along with how other aspects of intestinal health may be impacted.  Previous studies exploring the impact of MFGM addition in formula on later health have focused on cognitive development in infants(281, 322, 323).  Although this is of course a very important consideration, given the rise in incidence of allergic and autoimmune/inflammatory diseases it is also of great interest to determine the influence of neonate nutrition on later susceptibility to their development. Pursuing such questions would involve the rearing of rat pups past the pn day 15 time points used in these studies.  As size constraints are of concern by pn day 18 within the cups, pups would need to be 142  moved to conventional housing at this point after closure of their cannulas.  Introduction of oral based solid food would also need to commence at this point.  Examination of pups in all three diet groups at 1 and 2 months of age in terms of gut health and resident intestinal microbes would provide valuable insights into the long term effects of neonate nutrition.  Further analysis of more in depth parameters of intestinal health, both at early (pn day 15) and later (1-2 months) time points would also provide a more thorough understanding of the changes occurring within this complex biologic system.  This analysis should include examination beyond the intestinal epithelium including the makeup of immune subsets in the GI tract (T cells, B cells, ILCs) using fluorescence-associated cell sorting analysis, examination of immunoglobulin levels, and a thorough analysis of the structure and function (e.g. bacterial adhesion qualities) of the intestinal mucus layer.  Finally, determination of the cumulative effects of these changes on later susceptibility to models of colitis, such as that induced by administration of DSS or dinitrobenzene sulfonic acid would greatly contribute to the impact of this research on neonate intestinal health.         4.3.2 Enteric infections and their associated complications Likewise, perturbations in the intestinal environment as a result of enteric infection can also have long-term health consequences.  This is most obviously illustrated by complications such as post-infectious Guillain-Barré (where the body’s immune system attacks the nervous system), reactive arthritis, or hemolytic uremic syndrome, which may arise from infections with Campylobacter, Yersinia or Salmonella, and EHEC, respectively(224).  These complications may be driven in part by alterations in the beneficial responses (i.e. maladaptive responses) arising from multiple cell types during and after infection that are essential for maintaining and/or returning the 143  infected gut to a healthy state.  This includes responses at the level of IECs, immune cells, as well as in the changes and succession of bacterial communities in the intestine.  As many of the factors produced by each of these cell types and bacterial communities can impact the activation states and factors produced by others, this results in a complex interdependent system that in a susceptible individual may be easily perturbed.    For instance, bacterial metabolites such as short chain fatty acids (SCFAs), in addition to serving as a key energy source for IECs, can also significantly influence the immune system within the GI tract(61).  The induction and maintenance of Tregs in the colon is also promoted by SCFAs, a process which can limit intestinal tissue damage in models of experimentally induced intestinal inflammation(379).   Retinoic acid, a vitamin A metabolite whose production is also partly controlled by bacteria, can influence immune function such as the production of immunoglobulins by B cells(380).  The immune system can be influenced by bacterially derived metabolites indirectly through their activation of other cell types as well.   For instance, SCFA activation of G protein coupled receptors on IECs results in the production of pro-IL-18 and NLRP3 activation, cleaving IL-18 into its activated form(381).  IL-18 can in turn act on CD4+ T cells to limit Th17 cell differentiation and promote Treg effector molecule expression, as well as promote the production of AMPs(382).  Further, SCFAs can also inhibit the production of some chemokines by IECs, such as the neutrophil chemoattractant CXCL8, influencing immune cell recruitment to colonic tissue(61).  These beneficial responses from a variety of cell types are interdependent on one another.  When there is an altered response by one specific cell type in this interdependent relationship or in the reconstitution of the resident bacterial composition after 144  an enteric infection, this can have a significant impact on the responses generated by the others as outlined above.     Interestingly many studies exploring long term health outcomes post enteric infection have reported an elevated risk (up to 6-fold greater) for the development of IBS(220, 224-227, 358), although the intricacies of the pathways involved in the more susceptible individuals were not examined.  In one such study of a Swedish cohort, within 1 year of enteric infection by Campylobacter or Salmonella the observed number of ulcerative colitis and IBS cases was significantly elevated as compared to expected population risk levels(224).  In another study, the risk of developing ulcerative colitis was reported to double subsequent to an enteric infection(358).  Moreover, long-term implications of childhood enteric infections have been examined, and have shown an associated higher risk for developing inflammatory bowel disease in following years(357, 383-385).  It is important to consider for these cohort studies that many of the patients selected are those that have presented at a hospital with bacterial enteric infection.  This skews results to some extent, in that only patients that have presented with the most severe clinical symptoms will seek hospitalization.  As these post-infectious complications seem to arise only in some of the individuals infected and not in all, it is of interest to characterize what aspects of their responses (and by what cell types) may be contributing to their increased susceptibility.    145  4.3.2.1 Future direction: exploring the long term effects of increased susceptibility to early time-points of enteric infection In Chapter 2 we examined one key player, the IECs and how alterations in their ability to respond to intestinal bacteria and their products (through deletion of MyD88 within IECs) changed short-term susceptibility to infection.  Further studies in this area should examine whether the resulting increased sensitivity to early infection represented by significant intestinal damage, crypt infiltration of pathogenic bacteria and development of a leaky gut seen in the IEC-Myd88-/- mice may have implications in later susceptibility to disease.  This is of interest to clarify as the clinical studies discussed above see increased risk of developing IBS or ulcerative colitis up to one year after the initial infection, a time point by which the enteric pathogen would have been long cleared from the individuals’s system.  One hypothesized mechanism for these increased susceptibilities even after clearance of the pathogen are long lasting alterations to the intestinal mucosa-associated bacterial communities initiated during the infection(386).   In healthy subjects the bacterial communities present in the lumen are distinct from the mucosa-associated microbes that exist in a biofilm phenotype(52, 387).  These bacteria form complex colonies growing in a slime-enclosed film on intestinal mucus, protecting them from various AMPs, competing microbes and luminal viscous forces(386).  Further, bacteria released from the biofilm to a planktonic state into the lumen may display increased virulence as they differ metabolically from their biofilm counterparts and encounter different stressors(388).  As biofilm bacteria live in close proximity to the IECs, they may be the bacterial population most likely to impact host responses through sensing of their MAMPs or production of SCFAs.  Most current research focuses on changes in resident bacterial populations in luminal samples.  However, developing a clear understanding of intestinal bacterial biofilm communities in the healthy state, 146  how they change during infection and factors influencing their successful return to a pre-infectious state are of significant interest.    Given that IECs are in closest proximity to these biofilm communities, the factors they produce likely influence their composition.  For instance, the intestinal mucin glycoproteins produced by goblet cells offer a significant source of carbohydrates for saccharolytic bacteria in the colon.  Alterations to the mucus layer can affect the intestinal bacterial community, as is evidenced in mice undergoing chronic pharmacological Notch inhibition (skewing their IECs to a secretory subtype) infected with C. rodentium.  These mice display alterations in their mucin glycosylation with subsequent increases in the amount of mucin degrading bacterium Akkermansia muciniphila(389).  As reported in Chapter 2, goblet cell responses during infection in IEC-MyD88-/- mice are also impaired, which may influence biofilm communities. Therefore, in addition to assessing later susceptibility to intestinal inflammation after clearance of enteric pathogens, further studies in this area should focus on characterizing the biofilm communities of IEC-MyD88-/- mice before and after infection.  Recent advancements in intestinal mucus harvesting techniques, whereby aspirates of the outer mucus layer can be collected and analyzed(387), will allow for the collection of mucus adherent bacteria in our model.  Importantly, the impact of IEC-intrinsic innate signalling on these important bacterial communities can be assessed under homeostatic conditions as well as post-infection. 4.4 The impact of resident microbes shaped by food sources on intestinal development In addition to impacting the resident microbial community, neonate nutrition also influences the maturation of intestinal tissue.  The presence and composition of resident microbes has also been observed to influence intestinal maturation(48, 262, 283). Therefore, in Chapter 3 we aimed to 147  discern the contribution of the intestinal bacterial community shaped by neonate nutrition source on the development of the intestinal epithelium.  We achieved this in MM fed pups by gavaging them with an antibiotic cocktail to deplete intestinal bacteria before assessing parameters of intestinal development. Microbes were effectively depleted (>90%) after treatment with the antibiotic cocktail for 7 days.  Decreased microbe numbers impacted several parameters of intestinal development including villus heights, crypt depths, number of Ki-67 positive (proliferating) cells, lysozyme positive cells and Muc2 positive staining, suggesting the impact of MM on the resident intestinal bacteria contributes to the differences in intestinal development observed.    It would also be of interest to clarify to what extent MFGM supplementation in formula impacts intestinal development by influencing bacterial composition or by being absorbed and interacting with the epithelium itself.  We therefore attempted to deplete resident intestinal microbes in the MFGM formula fed group through a similar protocol to that employed on MM pups, but were only able to achieve a 45% decrease in microbes.  Upon assessing histological and IEC changes with the antibiotic treatment, we did find some read outs were affected with this amount of depletion (Figure 4.2).  In contrast to MM pups with antibiotic depleted microbes, crypt depths at the three intestinal sites with antibiotic treatment did not undergo as significant a decrease  (jejunum: MM antibiotic: 59.7μm ± 1.7, MFGM antibiotic: 69.8μm ± 2.3, ileum: MM antibiotic: 55.9μm ± 2.4, MFGM antibiotic: 70.5μm ± 2.0, colon: MM antibiotic: 170.2μm ± 4.1, MFGM antibiotic: 175μm ± 2.7, n=5) (Figure 4.2).  This may of course be in part due to the reduced depletion achieved in MFGM versus MM fed pups.    148   Figure 4.2:  Impact of intestinal microbe depletion (45%) on intestinal development in MFGM formula fed pups Developmental (A-C), histological (D-F) and immunofluorescent (G-I) analysis of antibiotic treated (7 days) or untreated MFGM pups at pn day 15.  Major developmental readouts including weight (A), overall small intestinal   (B), and colonic (C) length after antibiotic treatment were similar to untreated pups.  Microbe depletion significantly decreased villus length (D) and crypt depth (E) in the jejunum and colon (F) as compared to untreated MFGM intestinal tissues.  Number of proliferating cells (G) were decreased with antibiotic treatment in ileal and colonic tissues.  Lysozyme positive Paneth cells (H) were decreased in the ileum and Muc2 positive staining (I) was decreased in jejunal tissue of antibiotic treated MFGM pups as compared to untreated.  n=5  The graphed data presented are mean ± SEM, analyzed by unpaired Student’s t-test.  *p<0.05, **p<0.005, ***p<0.0005, ****p<0.0001.  149  4.4.1 Future direction: clarify the mechanisms through which MFGM impacts intestinal development   Since we were only able to achieve partial microbial depletion as compared to that observed with antibiotic treatment in MM pups, further studies should be conducted to more accurately clarify the role of resident microbes in the intestinal development of MFGM fed pups.  This can be attempted in two different ways.  One way would be to optimize depletion of the intestinal microbes in the formula fed groups that are administered their nutrition via gastric cannulas connected to isametic pumps.  Feeding through these pumps may result in overall faster transit time of the formula (and therefore the antibiotics) through their GI system.  As a single gavage dose of antibiotics was given to both MM and MFGM formula fed pups, the fast transit time of the antibiotics through the formula fed system may account in part for the reduced microbe depletion observed.  Rather than a single gavage dose, addition of antibiotics to the formula itself would presumably lead to the continued presence of the antibiotics in the formula fed GI tract over a longer period and may be advantageous in achieving greater depletion.  Alternatively, another way to examine the formula fed microbe effects on intestinal development would be to collect stool from formula fed pups and perform fecal transplants with these samples resuspended in phosphate buffered saline, into germ free mice, with subsequent analysis of gut development performed on the previously germfree mice.    Further work focusing on characterizing luminal bacterial metabolites and how these may differ with formula supplementation (and the addition of MFGM) as compared to MM is also of interest.  Epithelial cells can use SCFAs, produced by bacterial fermentation of indigestible carbohydrates in the colon, as a major source of energy.  For example, one specific colonic 150  SCFA, butyrate has been reported to provide approximately 60% of the energy needs of IECs(14).  Using a metabolomics approach, the bacterial metabolites present within luminal contents and an assessment of how formula feeding and the introduction of MFGM to formula can change these concentrations could be examined.  In addition, characterizing IEC expression of the monocarboxylate transporter 1, which plays a role in transport of butyrate into IECs in vitro and is reported to be upregulated by its presence, may provide insights into the mechanism through which SCFAs (specifically butyrate) may be affecting IECs(390).  Understanding differences in bacterial metabolite production based on the food source given to the neonate could provide insight into how neonate nutrition may be impacting the intestinal environment.  This may be one mechanism through which direct action of a bacterial byproduct on IECs is facilitating different rates of intestinal development.   In addition to microbial depletion by antibiotics in rat pups, fecal transplants into germ free mice, or examination of microbial metabolites effects on IECs, another method that could be employed to differentiate the potential direct effects of MFGM on IECs versus its influence on microbial composition would be through the use of intestinal organoids.  This in vitro model provides a 3D system of self-renewing tissue mimicking the histological and physiological characteristics of the intestinal epithelium(107). Generating organoids by isolating intestinal stem cells from various regions of the rat pup GI tract would provide a system isolated from resident intestinal bacteria to examine the potential direct effects of MFGM (as compared to CTL formula and MM) on IECs. One pathway of interest to examine within this system would be that governing intestinal stem cells and their differentiation into secretory and absorptive cell types (i.e. the Notch, Wnt, Bmp pathways).  As CTL formula colonic tissues displayed much greater CA-1 positive staining, this 151  may indicate the presence of more mature enterocytes.  Moreover, the decreased IEC proliferation and shorter crypts in the CTL formula pups may indicate greater amounts of BMP growth factors in their system resulting in greater differentiation into an absorptive cell phenotype along with decreased proliferation(114).  Analysis of these pathways in this system would allow us to discern whether these developmental changes are dependent on the presence of the bacteria alone, their metabolic byproducts, or some component found in the MFGM or MM.      4.5 Similarities in IEC responses during enteric infection and neonate intestinal development  As discussed in section 4.3, factors influencing the intestinal environment such as mode of delivery, neonate nutrient source, or enteric infections can have ramifications on long-term health.  Interestingly, there are many similarities in the changes occurring in the intestinal environment as well as IEC responses during these two states (development and infection) as well.    During enteric infection by pathogens such as C. rodentium, depletion of resident intestinal microbes by up to 80% occurs within one week, as well as significant changes to the make-up of the microbial communities(161, 185, 218).  The bacterial community shifts from a majority represented by the order Bacteroidales (Bacteroidetes) (>80%) to their significant depletion (down to approximately 27%) coinciding with a bloom in Enterobacteriales (>40%), represented by C. rodentium, by one-week post infection.  The re-establishment of the density and population diversity of bacteria to a pre-infection state occurs within four weeks of infection in an 152  immunocompetent host.  During development, although there is not depletion of resident bacteria occurring, we enter the world with a mostly sterile intestinal environment that is immediately colonized as the first step of establishing a resident microbial community.  Similar to the succession of bacteria that occurs after the infection induced depletion of microbes, there are shifts in the diversity and numbers of bacteria in the developing intestine until approximately three years of age in humans (approximately three weeks in mice) when the community reaches an adult-like profile(47, 391-393).   Similar to vaginally born humans, mice display similar initial intestinal bacteria to maternal vaginal communities. In the following days an increase in Streptococcus is followed by an increase in Lactobacillus species until weaning when members of the Bacteroidiales and Clostridiales represent the majority of the bacterial community, similar to that in adult mice(393).  Colonization succession similar to this also occurs in human infants and is discussed in section 1.1.2.   The establishment and/or reestablishment of bacterial communities occurring during both these circumstances are likely attributable to several factors both derived from the host and resident bacterial communities.      There are similarities present in the IEC responses observed during development and infection as well.  For instance, epithelial cell proliferation rates are higher than under homeostatic conditions during both circumstances as reported in Chapters 2, 3 and by several others(185, 201, 213, 274).  Notably, secretory cell responses were also variable during development and infection with food source (along with its influence on resident bacterial populations) and host immune status impacting these responses, respectively.   During intestinal development, lack of the milk fat component (present in both MFGM and MM fed pups) resulted in decreased positive staining of the major goblet cell secreted mucin Muc2 at pn day 15 in intestinal tissues (Figure 3.5).  As 153  there is some evidence that specific bacteria can influence transcription of intestinal mucin genes during neonate development(270), and our preliminary results show that antibiotic depletion of MFGM fed pups led to decreased staining at least in jejunal tissues, the influence of neonate food source on shaping intestinal bacteria may be one mechanism through which goblet cell responses are being impacted.  Decreased positive staining and gene transcript levels for Muc2, along with another goblet cell secreted protein Relmβ, were also observed in the cecal tissues of mice lacking the adaptor protein MyD88 specifically in IECs during Salmonella Typhimurium infection (Figure 2.7). Interestingly, studies examining defects in goblet cells as a result of impaired inflammasome (Nlrp6-/-, Caspase-1/11-/-, Asc-/-) or autophagy (IEC-Atg5-/-) signalling have revealed an inability of mice to clear C. rodentium infection when their signalling is impaired(394, 395).  In these aforementioned studies, although goblet cell secretion was compromised there were no differences detected in the transcription of goblet cell secreted factors.  Mucus production by the goblet cells in these studies was instead altered by their inability to effectively secrete their mucins, resulting in the aggregation of granules in the apical cytoplasm of the cell.  In our study, transcript levels of goblet cell factors were impacted by the lack of MyD88 signalling within IECs, resulting in decreased positive staining for goblet cell proteins during infection. Each of these studies highlights a major role for intestinal goblet cells, their secreted factors/mucus layer and their interactions with resident intestinal microbes during both homeostasis and intestinal challenge.  Future studies examining MyD88 deletion specifically within goblet cells would be of interest to clarify whether it is the lack of signalling and resulting responses within this IEC subset that controls the early susceptibility to infection observed in the IEC-MyD88-/- mice presented in Chapter 2.    154  4.6 Antibiotic induced disruption of the intestinal environment As discussed in the above sections, colonization succession after birth and the reassembly of resident bacterial communities post infection are highly influenced by IEC responses and their products.  This process can have implications for long-term health of the individual.  It is also an important concept for maintenance of health in the short term, as is the case with antibiotic therapy where disturbances in resident bacteria and altered re-colonization of the gut can increase susceptibility to infection with opportunistic pathogens such as C. difficile(249).  Further, antibiotic treatment, especially repeated exposures during childhood, increases the risk for the later development of intestinal inflammation, asthma and obesity(357, 396).   For instance, the early use of a specific class of antibiotics (macrolides) in a cohort of Finnish children was recently reported to significantly impact their resident intestinal bacteria, with decreased Actinobacteria and increases in Bacteroidetes and Proteobacteria observed(396).  These changes positively correlated with later development of either asthma or increased body mass index.  Moreover, a recent study exploring the role that single antibiotic doses may play on later susceptibly to inflammation in mice reported that intestinal inflammation was induced following antibiotic treatment in some strains of immunodeficient mice (Myd88 -/- and goblet cell-MyD88-/-)(397).  This inflammation was associated with significantly increased numbers of CX3CR1+ DCs trafficking bacteria to the MLN.  This study again illustrates the interdependence of immune cells, resident bacteria and the responses at the gut barrier in dictating susceptibility to the severity of intestinal disruption.  As my thesis work focused on changes at the intestinal epithelial layer during alterations in the intestinal environment, it was also of interest to explore 155  the effects of antibiotic treatment on the intestinal barrier and cells comprising it.  Preliminary studies found that antibiotic treatment in a genetically susceptibility host (Tlr2-/- mouse) resulted in acute cecitis (inflammation of the cecum), disruption of the intestinal barrier, as well as decreased positive staining for several goblet cell specific factors (Figure 4.3).  This preliminary data suggests that antibiotic treatment, through as yet undefined pathways, has the ability to affect responses at the level of IECs in addition to previous reports detailing the influence of antibiotic treatment at the level of immune cells.  To more fully understand the interdependence of these cellular responses and bacterial changes, further investigation is warranted.    156  Figure 4.3:  Antibiotic treatment of genetically susceptibility (Tlr2-/-) mice results in altered responses A) Representative hematoxylin and eosin staining of WT and Tlr2-/- cecal tissues after 1 dose of 20 mg streptomycin. B) Barrier permeability analysis via FITC-Dextran assay of untreated and streptomycin treated mice.  Tlr2-/- mice display significantly increased barrier disruption after streptomycin treatment.    Representative immunofluorescent images of the goblet cell factors TFF3 (C), Muc2 (D), and Relmβ (E) in cecal tissues of WT and  Tlr2-/- mice after streptomycin treatment. Tlr2-/- tissues display decreased positive staining for all three factors after treatment as compared to WT.  n≥4  The graphed data presented are mean ± SEM, analyzed by unpaired Student’s t-test.  **p<0.005.  100X magnification, Enlarged: 400X magnification.  157  4.7 Final remarks The findings presented in this thesis present evidence that IEC responses play a key role in maintaining homeostasis between the host and resident microbes, especially during times of dynamic change in the intestinal environment such as following birth or enteric infection.  An often-overlooked cell type in a physiologically very complex system, IECs can produce many factors such as mucins, cytokines and antimicrobial proteins that can influence luminal bacterial composition, intestinal immune cell makeup, and the strength of the intestinal barrier.  The production of these IEC factors, in turn can be influenced by bacterial MAMPs engaging IEC receptors, bacterial metabolites such as SCFAs, and cytokines and growth factors produced by immune cells.  Further, immune cells, which also play an irreplaceable role in intestinal health, can further be activated by bacterial MAMPs and their byproducts as well as these IEC factors.  This results in a complex interdependent system within the intestine that, under most circumstances, effectively maintains homeostasis between host tissue and luminal contents.  Understanding the intricacies of each contributing cell type (or bacterial community/byproduct) under both homeostatic conditions, as well during more tenuous times in the intestine, such as infection or immediately following birth, is valuable in identifying key pathways and cellular processes that potentiate beneficial responses.  Better characterizing these beneficial responses and the factors produced that facilitate them are of particular interest for vulnerable populations, such as premature infants or immune-compromised individuals who are generally more severely impacted by perturbations of the intestinal environment.  It would also help to highlight potential factors (or lack thereof) contributing to long-term health complications.  Taken together, it is hoped that the work presented in this PhD thesis contributes to our understanding of intestinal health and disease and provides unique insights into factors influencing intestinal development.  158  References 1. Yamada T AD, Kalloo A, Kaplowitz N, Owyang C, Powell D. Textbook of Gastroenterology New York, NY: John Wiley & Sons; 2009. 2. Arrieta MC, Stiemsma LT, Amenyogbe N, Brown EM, Finlay B. The intestinal microbiome in early life: health and disease. Front Immunol. 2014;5:427. 3. Khor B, Gardet A, Xavier RJ. Genetics and pathogenesis of inflammatory bowel disease. Nature. 2011;474(7351):307-17. 4. Gibson DL, Ma C, Bergstrom KS, Huang JT, Man C, Vallance BA. MyD88 signalling plays a critical role in host defence by controlling pathogen burden and promoting epithelial cell homeostasis during Citrobacter rodentium-induced colitis. Cell Microbiol. 2008;10(3):618-31. 5. Gibson DL, Ma C, Rosenberger CM, Bergstrom KS, Valdez Y, Huang JT, et al. Toll-like receptor 2 plays a critical role in maintaining mucosal integrity during Citrobacter rodentium-induced colitis. Cell Microbiol. 2008;10(2):388-403. 6. Lebeis SL, Powell KR, Merlin D, Sherman MA, Kalman D. Interleukin-1 receptor signaling protects mice from lethal intestinal damage caused by the attaching and effacing pathogen Citrobacter rodentium. Infect Immun. 2009;77(2):604-14. 7. Lebeis SL, Bommarius B, Parkos CA, Sherman MA, Kalman D. TLR signaling mediated by MyD88 is required for a protective innate immune response by neutrophils to Citrobacter rodentium. J Immunol. 2007;179(1):566-77. 8. Karam SM. Lineage commitment and maturation of epithelial cells in the gut. Front Biosci. 1999;4:D286-98. 9. Hao WL, Lee YK. Microflora of the gastrointestinal tract: a review. Methods Mol Biol. 2004;268:491-502. 10. Rao JN, Wang JY.  Regulation of Gastrointestinal Mucosal Growth. Integrated Systems Physiology: from Molecule to Function to Disease. San Rafael (CA)2010. 11. Hejnol A, Martin-Duran, J. M. Getting to the bottom of anal evolution. Zoologischer Anzeiger - A Journal of Comparative Zoology. 2015;256:61-74. 12. Kararli TT. Comparison of the gastrointestinal anatomy, physiology, and biochemistry of humans and commonly used laboratory animals. Biopharm Drug Dispos. 1995;16(5):351-80. 13. Goodman BE. Insights into digestion and absorption of major nutrients in humans. Adv Physiol Educ. 2010;34(2):44-53. 159  14. Cummings JH, Macfarlane GT. Role of intestinal bacteria in nutrient metabolism. JPEN J Parenter Enteral Nutr. 1997;21(6):357-65. 15. Reed KK, Wickham R. Review of the gastrointestinal tract: from macro to micro. Semin Oncol Nurs. 2009;25(1):3-14. 16. Mayhew TM, Middleton C. Crypts, villi and microvilli in the small intestine of the rat. A stereological study of their variability within and between animals. J Anat. 1985;141:1-17. 17. Jang I, Jung K, Cho J. Influence of age on duodenal brush border membrane and specific activities of brush border membrane enzymes in Wistar rats. Exp Anim. 2000;49(4):281-7. 18. Holmes R. The intestinal brush border. Gut. 1971;12(8):668-77. 19. Bai JP. Distribution of brush-border membrane peptidases along the rat intestine. Pharm Res. 1994;11(6):897-900. 20. Buts JP, Vijverman V, Barudi C, De Keyser N, Maldague P, Dive C. Refeeding after starvation in the rat: comparative effects of lipids, proteins and carbohydrates on jejunal and ileal mucosal adaptation. Eur J Clin Invest. 1990;20(4):441-52. 21. Jeejeebhoy KN. Short bowel syndrome: a nutritional and medical approach. CMAJ. 2002;166(10):1297-302. 22. Elphick DA, Mahida YR. Paneth cells: their role in innate immunity and inflammatory disease. Gut. 2005;54(12):1802-9. 23. Gerbe F, Jay P. Intestinal tuft cells: epithelial sentinels linking luminal cues to the immune system. Mucosal Immunol. 2016;9(6):1353-9. 24. Howitt MR, Lavoie S, Michaud M, Blum AM, Tran SV, Weinstock JV, et al. Tuft cells, taste-chemosensory cells, orchestrate parasite type 2 immunity in the gut. Science. 2016;351(6279):1329-33. 25. Gerbe F, Sidot E, Smyth DJ, Ohmoto M, Matsumoto I, Dardalhon V, et al. Intestinal epithelial tuft cells initiate type 2 mucosal immunity to helminth parasites. Nature. 2016;529(7585):226-30. 26. von Moltke J, Ji M, Liang HE, Locksley RM. Tuft-cell-derived IL-25 regulates an intestinal ILC2-epithelial response circuit. Nature. 2016;529(7585):221-5. 27. Birchenough GM, Johansson ME, Gustafsson JK, Bergstrom JH, Hansson GC. New developments in goblet cell mucus secretion and function. Mucosal Immunol. 2015;8(4):712-9. 28. Mowat AM, Agace WW. Regional specialization within the intestinal immune system. Nat Rev Immunol. 2014;14(10):667-85. 160  29. Cornes JS. Number, size, and distribution of Peyer's patches in the human small intestine: Part I The development of Peyer's patches. Gut. 1965;6(3):225-9. 30. Jang MH, Kweon MN, Iwatani K, Yamamoto M, Terahara K, Sasakawa C, et al. Intestinal villous M cells: an antigen entry site in the mucosal epithelium. Proc Natl Acad Sci U S A. 2004;101(16):6110-5. 31. O'Leary AD, Sweeney EC. Lymphoglandular complexes of the colon: structure and distribution. Histopathology. 1986;10(3):267-83. 32. Beagley KW, Fujihashi K, Lagoo AS, Lagoo-Deenadaylan S, Black CA, Murray AM, et al. Differences in intraepithelial lymphocyte T cell subsets isolated from murine small versus large intestine. J Immunol. 1995;154(11):5611-9. 33. Dominguez-Bello MG, Costello EK, Contreras M, Magris M, Hidalgo G, Fierer N, et al. Delivery mode shapes the acquisition and structure of the initial microbiota across multiple body habitats in newborns. Proc Natl Acad Sci U S A. 2010;107(26):11971-5. 34. Neu J, Rushing J. Cesarean versus vaginal delivery: long-term infant outcomes and the hygiene hypothesis. Clin Perinatol. 2011;38(2):321-31. 35. Morowitz MJ, Carlisle EM, Alverdy JC. Contributions of intestinal bacteria to nutrition and metabolism in the critically ill. Surg Clin North Am. 2011;91(4):771-85, viii. 36. Wostmann BS, Larkin C, Moriarty A, Bruckner-Kardoss E. Dietary intake, energy metabolism, and excretory losses of adult male germfree Wistar rats. Lab Anim Sci. 1983;33(1):46-50. 37. Osawa N, Mitsuhashi S. Infection of Germeree Mice with Shigella Flexneri 3a. Jpn J Exp Med. 1964;34:77-80. 38. Hentges DJ, Freter R. In vivo and in vitro antagonism of intestinal bacteria against Shigella flexneri. I. Correlation between various tests. J Infect Dis. 1962;110:30-7. 39. Lawley TD, Clare S, Walker AW, Goulding D, Stabler RA, Croucher N, et al. Antibiotic treatment of clostridium difficile carrier mice triggers a supershedder state, spore-mediated transmission, and severe disease in immunocompromised hosts. Infect Immun. 2009;77(9):3661-9. 40. Rupnik M, Wilcox MH, Gerding DN. Clostridium difficile infection: new developments in epidemiology and pathogenesis. Nat Rev Microbiol. 2009;7(7):526-36. 41. Bohnhoff M, Drake BL, Miller CP. Effect of streptomycin on susceptibility of intestinal tract to experimental Salmonella infection. Proc Soc Exp Biol Med. 1954;86(1):132-7. 161  42. Fukuda S, Toh H, Hase K, Oshima K, Nakanishi Y, Yoshimura K, et al. Bifidobacteria can protect from enteropathogenic infection through production of acetate. Nature. 2011;469(7331):543-7. 43. Wlodarska M, Willing B, Keeney KM, Menendez A, Bergstrom KS, Gill N, et al. Antibiotic treatment alters the colonic mucus layer and predisposes the host to exacerbated Citrobacter rodentium-induced colitis. Infect Immun. 2011;79(4):1536-45. 44. Lozupone CA, Stombaugh JI, Gordon JI, Jansson JK, Knight R. Diversity, stability and resilience of the human gut microbiota. Nature. 2012;489(7415):220-30. 45. Yatsunenko T, Rey FE, Manary MJ, Trehan I, Dominguez-Bello MG, Contreras M, et al. Human gut microbiome viewed across age and geography. Nature. 2012;486(7402):222-7. 46. Koenig JE, Spor A, Scalfone N, Fricker AD, Stombaugh J, Knight R, et al. Succession of microbial consortia in the developing infant gut microbiome. Proc Natl Acad Sci U S A. 2011;108 Suppl 1:4578-85. 47. Palmer C, Bik EM, DiGiulio DB, Relman DA, Brown PO. Development of the human infant intestinal microbiota. PLoS Biol. 2007;5(7):e177. 48. Matamoros S, Gras-Leguen C, Le Vacon F, Potel G, de La Cochetiere MF. Development of intestinal microbiota in infants and its impact on health. Trends Microbiol. 2013;21(4):167-73. 49. Adlerberth I, Wold AE. Establishment of the gut microbiota in Western infants. Acta Paediatr. 2009;98(2):229-38. 50. Bezirtzoglou E, Tsiotsias A, Welling GW. Microbiota profile in feces of breast- and formula-fed newborns by using fluorescence in situ hybridization (FISH). Anaerobe. 2011;17(6):478-82. 51. Guaraldi F, Salvatori G. Effect of breast and formula feeding on gut microbiota shaping in newborns. Front Cell Infect Microbiol. 2012;2:94. 52. Zoetendal EG, Rajilic-Stojanovic M, de Vos WM. High-throughput diversity and functionality analysis of the gastrointestinal tract microbiota. Gut. 2008;57(11):1605-15. 53. Gerritsen J, Smidt H, Rijkers GT, de Vos WM. Intestinal microbiota in human health and disease: the impact of probiotics. Genes Nutr. 2011;6(3):209-40. 54. Penders J, Stobberingh EE, van den Brandt PA, Thijs C. The role of the intestinal microbiota in the development of atopic disorders. Allergy. 2007;62(11):1223-36. 55. Booijink CC, Zoetendal EG, Kleerebezem M, de Vos WM. Microbial communities in the human small intestine: coupling diversity to metagenomics. Future Microbiol. 2007;2(3):285-95. 162  56. Donaldson GP, Lee SM, Mazmanian SK. Gut biogeography of the bacterial microbiota. Nat Rev Microbiol. 2016;14(1):20-32. 57. Gu S, Chen D, Zhang JN, Lv X, Wang K, Duan LP, et al. Bacterial community mapping of the mouse gastrointestinal tract. PLoS One. 2013;8(10):e74957. 58. Miller TL, Wolin MJ. Fermentations by saccharolytic intestinal bacteria. Am J Clin Nutr. 1979;32(1):164-72. 59. Cummings JH. Fermentation in the human large intestine: evidence and implications for health. Lancet. 1983;1(8335):1206-9. 60. Cummings JH, Macfarlane GT. The control and consequences of bacterial fermentation in the human colon. J Appl Bacteriol. 1991;70(6):443-59. 61. Brestoff JR, Artis D. Commensal bacteria at the interface of host metabolism and the immune system. Nat Immunol. 2013;14(7):676-84. 62. Donohoe DR, Garge N, Zhang X, Sun W, O'Connell TM, Bunger MK, et al. The microbiome and butyrate regulate energy metabolism and autophagy in the mammalian colon. Cell Metab. 2011;13(5):517-26. 63. Uchiyama K, Sakiyama T, Hasebe T, Musch MW, Miyoshi H, Nakagawa Y, et al. Butyrate and bioactive proteolytic form of Wnt-5a regulate colonic epithelial proliferation and spatial development. Sci Rep. 2016;6:32094. 64. Kaiko GE, Ryu SH, Koues OI, Collins PL, Solnica-Krezel L, Pearce EJ, et al. The Colonic Crypt Protects Stem Cells from Microbiota-Derived Metabolites. Cell. 2016;165(7):1708-20. 65. Johansson ME, Jakobsson HE, Holmen-Larsson J, Schutte A, Ermund A, Rodriguez-Pineiro AM, et al. Normalization of Host Intestinal Mucus Layers Requires Long-Term Microbial Colonization. Cell Host Microbe. 2015;18(5):582-92. 66. El Aidy S, van Baarlen P, Derrien M, Lindenbergh-Kortleve DJ, Hooiveld G, Levenez F, et al. Temporal and spatial interplay of microbiota and intestinal mucosa drive establishment of immune homeostasis in conventionalized mice. Mucosal Immunol. 2012;5(5):567-79. 67. Brown AJ, Goldsworthy SM, Barnes AA, Eilert MM, Tcheang L, Daniels D, et al. The Orphan G protein-coupled receptors GPR41 and GPR43 are activated by propionate and other short chain carboxylic acids. J Biol Chem. 2003;278(13):11312-9. 68. Le Poul E, Loison C, Struyf S, Springael JY, Lannoy V, Decobecq ME, et al. Functional characterization of human receptors for short chain fatty acids and their role in polymorphonuclear cell activation. J Biol Chem. 2003;278(28):25481-9. 163  69. Nilsson NE, Kotarsky K, Owman C, Olde B. Identification of a free fatty acid receptor, FFA2R, expressed on leukocytes and activated by short-chain fatty acids. Biochem Biophys Res Commun. 2003;303(4):1047-52. 70. Donohoe DR, Collins LB, Wali A, Bigler R, Sun W, Bultman SJ. The Warburg effect dictates the mechanism of butyrate-mediated histone acetylation and cell proliferation. Mol Cell. 2012;48(4):612-26. 71. Waldecker M, Kautenburger T, Daumann H, Busch C, Schrenk D. Inhibition of histone-deacetylase activity by short-chain fatty acids and some polyphenol metabolites formed in the colon. J Nutr Biochem. 2008;19(9):587-93. 72. Hinnebusch BF, Meng S, Wu JT, Archer SY, Hodin RA. The effects of short-chain fatty acids on human colon cancer cell phenotype are associated with histone hyperacetylation. J Nutr. 2002;132(5):1012-7. 73. Maslowski KM, Vieira AT, Ng A, Kranich J, Sierro F, Yu D, et al. Regulation of inflammatory responses by gut microbiota and chemoattractant receptor GPR43. Nature. 2009;461(7268):1282-6. 74. Cox MA, Jackson J, Stanton M, Rojas-Triana A, Bober L, Laverty M, et al. Short-chain fatty acids act as antiinflammatory mediators by regulating prostaglandin E(2) and cytokines. World J Gastroenterol. 2009;15(44):5549-57. 75. Berndt BE, Zhang M, Owyang SY, Cole TS, Wang TW, Luther J, et al. Butyrate increases IL-23 production by stimulated dendritic cells. Am J Physiol Gastrointest Liver Physiol. 2012;303(12):G1384-92. 76. Liu L, Li L, Min J, Wang J, Wu H, Zeng Y, et al. Butyrate interferes with the differentiation and function of human monocyte-derived dendritic cells. Cell Immunol. 2012;277(1-2):66-73. 77. Eftimiadi C, Stashenko P, Tonetti M, Mangiante PE, Massara R, Zupo S, et al. Divergent effect of the anaerobic bacteria by-product butyric acid on the immune response: suppression of T-lymphocyte proliferation and stimulation of interleukin-1 beta production. Oral Microbiol Immunol. 1991;6(1):17-23. 78. Gilbert KM, DeLoose A, Valentine JL, Fifer EK. Structure-activity relationship between carboxylic acids and T cell cycle blockade. Life Sci. 2006;78(19):2159-65. 79. Bailon E, Cueto-Sola M, Utrilla P, Rodriguez-Cabezas ME, Garrido-Mesa N, Zarzuelo A, et al. Butyrate in vitro immune-modulatory effects might be mediated through a proliferation-related induction of apoptosis. Immunobiology. 2010;215(11):863-73. 164  80. Zimmerman MA, Singh N, Martin PM, Thangaraju M, Ganapathy V, Waller JL, et al. Butyrate suppresses colonic inflammation through HDAC1-dependent Fas upregulation and Fas-mediated apoptosis of T cells. Am J Physiol Gastrointest Liver Physiol. 2012;302(12):G1405-15. 81. Kaiko GE, Ryu SH, Koues OI, Collins PL, Solnica-Krezel L, Pearce EJ, et al. The Colonic Crypt Protects Stem Cells from Microbiota-Derived Metabolites. Cell. 2016;167(4):1137. 82. Moens E, Veldhoen M. Epithelial barrier biology: good fences make good neighbours. Immunology. 2012;135(1):1-8. 83. Ramanan D, Cadwell K. Intrinsic Defense Mechanisms of the Intestinal Epithelium. Cell Host Microbe. 2016;19(4):434-41. 84. Zhang K, Hornef MW, Dupont A. The intestinal epithelium as guardian of gut barrier integrity. Cell Microbiol. 2015;17(11):1561-9. 85. Turner JR. Intestinal mucosal barrier function in health and disease. Nat Rev Immunol. 2009;9(11):799-809. 86. Chiba H, Osanai M, Murata M, Kojima T, Sawada N. Transmembrane proteins of tight junctions. Biochim Biophys Acta. 2008;1778(3):588-600. 87. Furuse M, Itoh M, Hirase T, Nagafuchi A, Yonemura S, Tsukita S, et al. Direct association of occludin with ZO-1 and its possible involvement in the localization of occludin at tight junctions. J Cell Biol. 1994;127(6 Pt 1):1617-26. 88. Lee SH. Intestinal permeability regulation by tight junction: implication on inflammatory bowel diseases. Intest Res. 2015;13(1):11-8. 89. Van Itallie CM, Fanning AS, Bridges A, Anderson JM. ZO-1 stabilizes the tight junction solute barrier through coupling to the perijunctional cytoskeleton. Mol Biol Cell. 2009;20(17):3930-40. 90. Marchiando AM, Graham WV, Turner JR. Epithelial barriers in homeostasis and disease. Annu Rev Pathol. 2010;5:119-44. 91. Anderson JM, Van Itallie CM. Physiology and function of the tight junction. Cold Spring Harb Perspect Biol. 2009;1(2):a002584. 92. Rehder D, Iden S, Nasdala I, Wegener J, Brickwedde MK, Vestweber D, et al. Junctional adhesion molecule-a participates in the formation of apico-basal polarity through different domains. Exp Cell Res. 2006;312(17):3389-403. 93. Satohisa S, Chiba H, Osanai M, Ohno S, Kojima T, Saito T, et al. Behavior of tight-junction, adherens-junction and cell polarity proteins during HNF-4alpha-induced epithelial polarization. Exp Cell Res. 2005;310(1):66-78. 165  94. Hermiston ML, Gordon JI. In vivo analysis of cadherin function in the mouse intestinal epithelium: essential roles in adhesion, maintenance of differentiation, and regulation of programmed cell death. J Cell Biol. 1995;129(2):489-506. 95. Holmes JL, Van Itallie CM, Rasmussen JE, Anderson JM. Claudin profiling in the mouse during postnatal intestinal development and along the gastrointestinal tract reveals complex expression patterns. Gene Expr Patterns. 2006;6(6):581-8. 96. Marchiando AM, Shen L, Graham WV, Weber CR, Schwarz BT, Austin JR, 2nd, et al. Caveolin-1-dependent occludin endocytosis is required for TNF-induced tight junction regulation in vivo. J Cell Biol. 2010;189(1):111-26. 97. Tsukita S, Furuse M, Itoh M. Multifunctional strands in tight junctions. Nat Rev Mol Cell Biol. 2001;2(4):285-93. 98. Morita K, Furuse M, Fujimoto K, Tsukita S. Claudin multigene family encoding four-transmembrane domain protein components of tight junction strands. Proc Natl Acad Sci U S A. 1999;96(2):511-6. 99. Mankertz J, Tavalali S, Schmitz H, Mankertz A, Riecken EO, Fromm M, et al. Expression from the human occludin promoter is affected by tumor necrosis factor alpha and interferon gamma. J Cell Sci. 2000;113 ( Pt 11):2085-90. 100. Zeissig S, Burgel N, Gunzel D, Richter J, Mankertz J, Wahnschaffe U, et al. Changes in expression and distribution of claudin 2, 5 and 8 lead to discontinuous tight junctions and barrier dysfunction in active Crohn's disease. Gut. 2007;56(1):61-72. 101. Prasad S, Mingrino R, Kaukinen K, Hayes KL, Powell RM, MacDonald TT, et al. Inflammatory processes have differential effects on claudins 2, 3 and 4 in colonic epithelial cells. Lab Invest. 2005;85(9):1139-62. 102. Heller F, Florian P, Bojarski C, Richter J, Christ M, Hillenbrand B, et al. Interleukin-13 is the key effector Th2 cytokine in ulcerative colitis that affects epithelial tight junctions, apoptosis, and cell restitution. Gastroenterology. 2005;129(2):550-64. 103. Guttman JA, Finlay BB. Tight junctions as targets of infectious agents. Biochim Biophys Acta. 2009;1788(4):832-41. 104. Peterson LW, Artis D. Intestinal epithelial cells: regulators of barrier function and immune homeostasis. Nat Rev Immunol. 2014;14(3):141-53. 105. van der Flier LG, Clevers H. Stem cells, self-renewal, and differentiation in the intestinal epithelium. Annu Rev Physiol. 2009;71:241-60. 106. Barker N, van Es JH, Kuipers J, Kujala P, van den Born M, Cozijnsen M, et al. Identification of stem cells in small intestine and colon by marker gene Lgr5. Nature. 2007;449(7165):1003-7. 166  107. Date S, Sato T. Mini-gut organoids: reconstitution of the stem cell niche. Annu Rev Cell Dev Biol. 2015;31:269-89. 108. Cheng H. Origin, differentiation and renewal of the four main epithelial cell types in the mouse small intestine. IV. Paneth cells. Am J Anat. 1974;141(4):521-35. 109. Bry L, Falk P, Huttner K, Ouellette A, Midtvedt T, Gordon JI. Paneth cell differentiation in the developing intestine of normal and transgenic mice. Proc Natl Acad Sci U S A. 1994;91(22):10335-9. 110. van de Wetering M, Sancho E, Verweij C, de Lau W, Oving I, Hurlstone A, et al. The beta-catenin/TCF-4 complex imposes a crypt progenitor phenotype on colorectal cancer cells. Cell. 2002;111(2):241-50. 111. de Lau W, Barker N, Low TY, Koo BK, Li VS, Teunissen H, et al. Lgr5 homologues associate with Wnt receptors and mediate R-spondin signalling. Nature. 2011;476(7360):293-7. 112. Kim KA, Kakitani M, Zhao J, Oshima T, Tang T, Binnerts M, et al. Mitogenic influence of human R-spondin1 on the intestinal epithelium. Science. 2005;309(5738):1256-9. 113. Qi Z, Li Y, Zhao B, Xu C, Liu Y, Li H, et al. BMP restricts stemness of intestinal Lgr5+ stem cells by directly suppressing their signature genes. Nat Commun. 2017;8:13824. 114. Hardwick JC, Van Den Brink GR, Bleuming SA, Ballester I, Van Den Brande JM, Keller JJ, et al. Bone morphogenetic protein 2 is expressed by, and acts upon, mature epithelial cells in the colon. Gastroenterology. 2004;126(1):111-21. 115. He XC, Zhang J, Tong WG, Tawfik O, Ross J, Scoville DH, et al. BMP signaling inhibits intestinal stem cell self-renewal through suppression of Wnt-beta-catenin signaling. Nat Genet. 2004;36(10):1117-21. 116. Davis H, Irshad S, Bansal M, Rafferty H, Boitsova T, Bardella C, et al. Aberrant epithelial GREM1 expression initiates colonic tumorigenesis from cells outside the stem cell niche. Nat Med. 2015;21(1):62-70. 117. Haramis AP, Begthel H, van den Born M, van Es J, Jonkheer S, Offerhaus GJ, et al. De novo crypt formation and juvenile polyposis on BMP inhibition in mouse intestine. Science. 2004;303(5664):1684-6. 118. van der Flier LG, van Gijn ME, Hatzis P, Kujala P, Haegebarth A, Stange DE, et al. Transcription factor achaete scute-like 2 controls intestinal stem cell fate. Cell. 2009;136(5):903-12. 119. Bertrand K. Survival of exfoliated epithelial cells: a delicate balance between anoikis and apoptosis. J Biomed Biotechnol. 2011;2011:534139. 167  120. Cliffe LJ, Humphreys NE, Lane TE, Potten CS, Booth C, Grencis RK. Accelerated intestinal epithelial cell turnover: a new mechanism of parasite expulsion. Science. 2005;308(5727):1463-5. 121. Schauer DB, Falkow S. The eae gene of Citrobacter freundii biotype 4280 is necessary for colonization in transmissible murine colonic hyperplasia. Infect Immun. 1993;61(11):4654-61. 122. Zheng Y, Valdez PA, Danilenko DM, Hu Y, Sa SM, Gong Q, et al. Interleukin-22 mediates early host defense against attaching and effacing bacterial pathogens. Nat Med. 2008;14(3):282-9. 123. Parks OB, Pociask DA, Hodzic Z, Kolls JK, Good M. Interleukin-22 Signaling in the Regulation of Intestinal Health and Disease. Front Cell Dev Biol. 2015;3:85. 124. Liboni KC, Li N, Scumpia PO, Neu J. Glutamine modulates LPS-induced IL-8 production through IkappaB/NF-kappaB in human fetal and adult intestinal epithelium. J Nutr. 2005;135(2):245-51. 125. Kwon JH, Keates S, Bassani L, Mayer LF, Keates AC. Colonic epithelial cells are a major site of macrophage inflammatory protein 3alpha (MIP-3alpha) production in normal colon and inflammatory bowel disease. Gut. 2002;51(6):818-26. 126. Caruso R, Fina D, Peluso I, Stolfi C, Fantini MC, Gioia V, et al. A functional role for interleukin-21 in promoting the synthesis of the T-cell chemoattractant, MIP-3alpha, by gut epithelial cells. Gastroenterology. 2007;132(1):166-75. 127. Ajuebor MN, Das AM, Virág L, Flower RJ, Szabó C, Perretti M. Role of resident peritoneal macrophages and mast cells in chemokine production and neutrophil migration in acute inflammation: evidence for an inhibitory loop involving endogenous IL-10. J Immunol. 1999;162(3):1685-91. 128. Ajuebor MN, Singh A, Wallace JL. Cyclooxygenase-2-derived prostaglandin D(2) is an early anti-inflammatory signal in experimental colitis. Am J Physiol Gastrointest Liver Physiol. 2000;279(1):G238-44. 129. Keates S, Han X, Kelly CP, Keates AC. Macrophage-inflammatory protein-3alpha mediates epidermal growth factor receptor transactivation and ERK1/2 MAPK signaling in Caco-2 colonic epithelial cells via metalloproteinase-dependent release of amphiregulin. J Immunol. 2007;178(12):8013-21. 130. Kim TH, Escudero S, Shivdasani RA. Intact function of Lgr5 receptor-expressing intestinal stem cells in the absence of Paneth cells. Proc Natl Acad Sci U S A. 2012;109(10):3932-7. 168  131. Durand A, Donahue B, Peignon G, Letourneur F, Cagnard N, Slomianny C, et al. Functional intestinal stem cells after Paneth cell ablation induced by the loss of transcription factor Math1 (Atoh1). Proc Natl Acad Sci U S A. 2012;109(23):8965-70. 132. Clevers HC, Bevins CL. Paneth cells: maestros of the small intestinal crypts. Annu Rev Physiol. 2013;75:289-311. 133. Bevins CL, Salzman NH. Paneth cells, antimicrobial peptides and maintenance of intestinal homeostasis. Nat Rev Microbiol. 2011;9(5):356-68. 134. Hooper LV, Stappenbeck TS, Hong CV, Gordon JI. Angiogenins: a new class of microbicidal proteins involved in innate immunity. Nat Immunol. 2003;4(3):269-73. 135. Satoh Y, Habara Y, Ono K, Kanno T. Carbamylcholine- and catecholamine-induced intracellular calcium dynamics of epithelial cells in mouse ileal crypts. Gastroenterology. 1995;108(5):1345-56. 136. Farin HF, Karthaus WR, Kujala P, Rakhshandehroo M, Schwank G, Vries RG, et al. Paneth cell extrusion and release of antimicrobial products is directly controlled by immune cell-derived IFN-gamma. J Exp Med. 2014;211(7):1393-405. 137. Ayabe T, Satchell DP, Wilson CL, Parks WC, Selsted ME, Ouellette AJ. Secretion of microbicidal alpha-defensins by intestinal Paneth cells in response to bacteria. Nat Immunol. 2000;1(2):113-8. 138. Tanabe H, Ayabe T, Bainbridge B, Guina T, Ernst RK, Darveau RP, et al. Mouse paneth cell secretory responses to cell surface glycolipids of virulent and attenuated pathogenic bacteria. Infect Immun. 2005;73(4):2312-20. 139. Kobayashi KS, Chamaillard M, Ogura Y, Henegariu O, Inohara N, Nunez G, et al. Nod2-dependent regulation of innate and adaptive immunity in the intestinal tract. Science. 2005;307(5710):731-4. 140. Wehkamp J, Stange EF. Paneth's disease. J Crohns Colitis. 2010;4(5):523-31. 141. Wehkamp J, Salzman NH, Porter E, Nuding S, Weichenthal M, Petras RE, et al. Reduced Paneth cell alpha-defensins in ileal Crohn's disease. Proc Natl Acad Sci U S A. 2005;102(50):18129-34. 142. McElroy SJ, Prince LS, Weitkamp JH, Reese J, Slaughter JC, Polk DB. Tumor necrosis factor receptor 1-dependent depletion of mucus in immature small intestine: a potential role in neonatal necrotizing enterocolitis. Am J Physiol Gastrointest Liver Physiol. 2011;301(4):G656-66. 143. Salzman NH, Polin RA, Harris MC, Ruchelli E, Hebra A, Zirin-Butler S, et al. Enteric defensin expression in necrotizing enterocolitis. Pediatr Res. 1998;44(1):20-6. 169  144. Furness JB, Rivera LR, Cho HJ, Bravo DM, Callaghan B. The gut as a sensory organ. Nat Rev Gastroenterol Hepatol. 2013;10(12):729-40. 145. Mawe GM, Hoffman JM. Serotonin signalling in the gut--functions, dysfunctions and therapeutic targets. Nat Rev Gastroenterol Hepatol. 2013;10(8):473-86. 146. Palazzo M, Balsari A, Rossini A, Selleri S, Calcaterra C, Gariboldi S, et al. Activation of enteroendocrine cells via TLRs induces hormone, chemokine, and defensin secretion. J Immunol. 2007;178(7):4296-303. 147. Atuma C, Strugala V, Allen A, Holm L. The adherent gastrointestinal mucus gel layer: thickness and physical state in vivo. Am J Physiol Gastrointest Liver Physiol. 2001;280(5):G922-9. 148. Kim YS, Ho SB. Intestinal goblet cells and mucins in health and disease: recent insights and progress. Curr Gastroenterol Rep. 2010;12(5):319-30. 149. Velcich A, Yang W, Heyer J, Fragale A, Nicholas C, Viani S, et al. Colorectal cancer in mice genetically deficient in the mucin Muc2. Science. 2002;295(5560):1726-9. 150. Xu G, Bell SL, McCool D, Forstner JF. The cationic C-terminus of rat Muc2 facilitates dimer formation post translationally and is subsequently removed by furin. Eur J Biochem. 2000;267(10):2998-3004. 151. Godl K, Johansson ME, Lidell ME, Morgelin M, Karlsson H, Olson FJ, et al. The N terminus of the MUC2 mucin forms trimers that are held together within a trypsin-resistant core fragment. J Biol Chem. 2002;277(49):47248-56. 152. Lidell ME, Johansson ME, Morgelin M, Asker N, Gum JR, Jr., Kim YS, et al. The recombinant C-terminus of the human MUC2 mucin forms dimers in Chinese-hamster ovary cells and heterodimers with full-length MUC2 in LS 174T cells. Biochem J. 2003;372(Pt 2):335-45. 153. Ambort D, Johansson ME, Gustafsson JK, Nilsson HE, Ermund A, Johansson BR, et al. Calcium and pH-dependent packing and release of the gel-forming MUC2 mucin. Proc Natl Acad Sci U S A. 2012;109(15):5645-50. 154. Ridley C, Kouvatsos N, Raynal BD, Howard M, Collins RF, Desseyn JL, et al. Assembly of the respiratory mucin MUC5B: a new model for a gel-forming mucin. J Biol Chem. 2014;289(23):16409-20. 155. Andrianifahanana M, Moniaux N, Batra SK. Regulation of mucin expression: mechanistic aspects and implications for cancer and inflammatory diseases. Biochim Biophys Acta. 2006;1765(2):189-222. 170  156. Gum JR, Byrd JC, Hicks JW, Toribara NW, Lamport DT, Kim YS. Molecular cloning of human intestinal mucin cDNAs. Sequence analysis and evidence for genetic polymorphism. J Biol Chem. 1989;264(11):6480-7. 157. Ermund A, Schutte A, Johansson ME, Gustafsson JK, Hansson GC. Studies of mucus in mouse stomach, small intestine, and colon. I. Gastrointestinal mucus layers have different properties depending on location as well as over the Peyer's patches. Am J Physiol Gastrointest Liver Physiol. 2013;305(5):G341-7. 158. Vaishnava S, Yamamoto M, Severson KM, Ruhn KA, Yu X, Koren O, et al. The antibacterial lectin RegIIIgamma promotes the spatial segregation of microbiota and host in the intestine. Science. 2011;334(6053):255-8. 159. Schutte A, Ermund A, Becker-Pauly C, Johansson ME, Rodriguez-Pineiro AM, Backhed F, et al. Microbial-induced meprin beta cleavage in MUC2 mucin and a functional CFTR channel are required to release anchored small intestinal mucus. Proc Natl Acad Sci U S A. 2014;111(34):12396-401. 160. Johansson ME, Phillipson M, Petersson J, Velcich A, Holm L, Hansson GC. The inner of the two Muc2 mucin-dependent mucus layers in colon is devoid of bacteria. Proc Natl Acad Sci U S A. 2008;105(39):15064-9. 161. Bergstrom KS, Kissoon-Singh V, Gibson DL, Ma C, Montero M, Sham HP, et al. Muc2 protects against lethal infectious colitis by disassociating pathogenic and commensal bacteria from the colonic mucosa. PLoS Pathog. 2010;6(5):e1000902. 162. Van der Sluis M, De Koning BA, De Bruijn AC, Velcich A, Meijerink JP, Van Goudoever JB, et al. Muc2-deficient mice spontaneously develop colitis, indicating that MUC2 is critical for colonic protection. Gastroenterology. 2006;131(1):117-29. 163. Johansson ME, Gustafsson JK, Holmen-Larsson J, Jabbar KS, Xia L, Xu H, et al. Bacteria penetrate the normally impenetrable inner colon mucus layer in both murine colitis models and patients with ulcerative colitis. Gut. 2014;63(2):281-91. 164. Dignass A, Lynch-Devaney K, Kindon H, Thim L, Podolsky DK. Trefoil peptides promote epithelial migration through a transforming growth factor beta-independent pathway. J Clin Invest. 1994;94(1):376-83. 165. Kobayashi K, Ogata H, Morikawa M, Iijima S, Harada N, Yoshida T, et al. Distribution and partial characterisation of IgG Fc binding protein in various mucin producing cells and body fluids. Gut. 2002;51(2):169-76. 166. Harada N, Iijima S, Kobayashi K, Yoshida T, Brown WR, Hibi T, et al. Human IgGFc binding protein (FcgammaBP) in colonic epithelial cells exhibits mucin-like structure. J Biol Chem. 1997;272(24):15232-41. 171  167. Bergstrom KS, Morampudi V, Chan JM, Bhinder G, Lau J, Yang H, et al. Goblet Cell Derived RELM-beta Recruits CD4+ T Cells during Infectious Colitis to Promote Protective Intestinal Epithelial Cell Proliferation. PLoS Pathog. 2015;11(8):e1005108. 168. Herbert DR, Yang JQ, Hogan SP, Groschwitz K, Khodoun M, Munitz A, et al. Intestinal epithelial cell secretion of RELM-beta protects against gastrointestinal worm infection. J Exp Med. 2009;206(13):2947-57. 169. Wang ML, Shin ME, Knight PA, Artis D, Silberg DG, Suh E, et al. Regulation of RELM/FIZZ isoform expression by Cdx2 in response to innate and adaptive immune stimulation in the intestine. Am J Physiol Gastrointest Liver Physiol. 2005;288(5):G1074-83. 170. Vaishnava S, Behrendt CL, Ismail AS, Eckmann L, Hooper LV. Paneth cells directly sense gut commensals and maintain homeostasis at the intestinal host-microbial interface. Proc Natl Acad Sci U S A. 2008;105(52):20858-63. 171. Nair MG, Guild KJ, Du Y, Zaph C, Yancopoulos GD, Valenzuela DM, et al. Goblet cell-derived resistin-like molecule beta augments CD4+ T cell production of IFN-gamma and infection-induced intestinal inflammation. J Immunol. 2008;181(7):4709-15. 172. Artis D, Wang ML, Keilbaugh SA, He W, Brenes M, Swain GP, et al. RELMbeta/FIZZ2 is a goblet cell-specific immune-effector molecule in the gastrointestinal tract. Proc Natl Acad Sci U S A. 2004;101(37):13596-600. 173. Ey B, Eyking A, Gerken G, Podolsky DK, Cario E. TLR2 mediates gap junctional intercellular communication through connexin-43 in intestinal epithelial barrier injury. J Biol Chem. 2009;284(33):22332-43. 174. Cario E, Gerken G, Podolsky DK. Toll-like receptor 2 enhances ZO-1-associated intestinal epithelial barrier integrity via protein kinase C. Gastroenterology. 2004;127(1):224-38. 175. Vora P, Youdim A, Thomas LS, Fukata M, Tesfay SY, Lukasek K, et al. Beta-defensin-2 expression is regulated by TLR signaling in intestinal epithelial cells. J Immunol. 2004;173(9):5398-405. 176. Akira S, Takeda K. Toll-like receptor signalling. Nat Rev Immunol. 2004;4(7):499-511. 177. Akira S, Yamamoto M, Takeda K. Role of adapters in Toll-like receptor signalling. Biochem Soc Trans. 2003;31(Pt 3):637-42. 178. Akira S, Uematsu S, Takeuchi O. Pathogen recognition and innate immunity. Cell. 2006;124(4):783-801. 179. Nenci A, Becker C, Wullaert A, Gareus R, van Loo G, Danese S, et al. Epithelial NEMO links innate immunity to chronic intestinal inflammation. Nature. 2007;446(7135):557-61. 172  180. Palomo J, Dietrich D, Martin P, Palmer G, Gabay C. The interleukin (IL)-1 cytokine family--Balance between agonists and antagonists in inflammatory diseases. Cytokine. 2015;76(1):25-37. 181. Boraschi D, Tagliabue A. The interleukin-1 receptor family. Semin Immunol. 2013;25(6):394-407. 182. Qin J, Qian Y, Yao J, Grace C, Li X. SIGIRR inhibits interleukin-1 receptor- and toll-like receptor 4-mediated signaling through different mechanisms. J Biol Chem. 2005;280(26):25233-41. 183. Nold-Petry CA, Lo CY, Rudloff I, Elgass KD, Li S, Gantier MP, et al. IL-37 requires the receptors IL-18Ralpha and IL-1R8 (SIGIRR) to carry out its multifaceted anti-inflammatory program upon innate signal transduction. Nat Immunol. 2015;16(4):354-65. 184. Xiao H, Gulen MF, Qin J, Yao J, Bulek K, Kish D, et al. The Toll-interleukin-1 receptor member SIGIRR regulates colonic epithelial homeostasis, inflammation, and tumorigenesis. Immunity. 2007;26(4):461-75. 185. Sham HP, Yu EY, Gulen MF, Bhinder G, Stahl M, Chan JM, et al. SIGIRR, a negative regulator of TLR/IL-1R signalling promotes Microbiota dependent resistance to colonization by enteric bacterial pathogens. PLoS Pathog. 2013;9(8):e1003539. 186. Kadota C, Ishihara S, Aziz MM, Rumi MA, Oshima N, Mishima Y, et al. Down-regulation of single immunoglobulin interleukin-1R-related molecule (SIGIRR)/TIR8 expression in intestinal epithelial cells during inflammation. Clin Exp Immunol. 2010;162(2):348-61. 187. Sims JE, Smith DE. The IL-1 family: regulators of immunity. Nat Rev Immunol. 2010;10(2):89-102. 188. Coccia M, Harrison OJ, Schiering C, Asquith MJ, Becher B, Powrie F, et al. IL-1beta mediates chronic intestinal inflammation by promoting the accumulation of IL-17A secreting innate lymphoid cells and CD4(+) Th17 cells. J Exp Med. 2012;209(9):1595-609. 189. Ben-Sasson SZ, Hu-Li J, Quiel J, Cauchetaux S, Ratner M, Shapira I, et al. IL-1 acts directly on CD4 T cells to enhance their antigen-driven expansion and differentiation. Proc Natl Acad Sci U S A. 2009;106(17):7119-24. 190. Sutton CE, Lalor SJ, Sweeney CM, Brereton CF, Lavelle EC, Mills KH. Interleukin-1 and IL-23 induce innate IL-17 production from gammadelta T cells, amplifying Th17 responses and autoimmunity. Immunity. 2009;31(2):331-41. 191. Sutton C, Brereton C, Keogh B, Mills KH, Lavelle EC. A crucial role for interleukin (IL)-1 in the induction of IL-17-producing T cells that mediate autoimmune encephalomyelitis. J Exp Med. 2006;203(7):1685-91. 173  192. Acosta-Rodriguez EV, Napolitani G, Lanzavecchia A, Sallusto F. Interleukins 1beta and 6 but not transforming growth factor-beta are essential for the differentiation of interleukin 17-producing human T helper cells. Nat Immunol. 2007;8(9):942-9. 193. Chung Y, Chang SH, Martinez GJ, Yang XO, Nurieva R, Kang HS, et al. Critical regulation of early Th17 cell differentiation by interleukin-1 signaling. Immunity. 2009;30(4):576-87. 194. Takeda K, Tsutsui H, Yoshimoto T, Adachi O, Yoshida N, Kishimoto T, et al. Defective NK cell activity and Th1 response in IL-18-deficient mice. Immunity. 1998;8(3):383-90. 195. Oficjalska K, Raverdeau M, Aviello G, Wade SC, Hickey A, Sheehan KM, et al. Protective role for caspase-11 during acute experimental murine colitis. J Immunol. 2015;194(3):1252-60. 196. Dupaul-Chicoine J, Yeretssian G, Doiron K, Bergstrom KS, McIntire CR, LeBlanc PM, et al. Control of intestinal homeostasis, colitis, and colitis-associated colorectal cancer by the inflammatory caspases. Immunity. 2010;32(3):367-78. 197. Siegmund B, Fantuzzi G, Rieder F, Gamboni-Robertson F, Lehr HA, Hartmann G, et al. Neutralization of interleukin-18 reduces severity in murine colitis and intestinal IFN-gamma and TNF-alpha production. Am J Physiol Regul Integr Comp Physiol. 2001;281(4):R1264-73. 198. Sivakumar PV, Westrich GM, Kanaly S, Garka K, Born TL, Derry JM, et al. Interleukin 18 is a primary mediator of the inflammation associated with dextran sulphate sodium induced colitis: blocking interleukin 18 attenuates intestinal damage. Gut. 2002;50(6):812-20. 199. Akira S. Pathogen recognition by innate immunity and its signaling. Proc Jpn Acad Ser B Phys Biol Sci. 2009;85(4):143-56. 200. Cario E. Toll-like receptors in inflammatory bowel diseases: a decade later. Inflamm Bowel Dis. 2010;16(9):1583-97. 201. Abreu MT. Toll-like receptor signalling in the intestinal epithelium: how bacterial recognition shapes intestinal function. Nat Rev Immunol. 2010;10(2):131-44. 202. Abreu MT, Arnold ET, Thomas LS, Gonsky R, Zhou Y, Hu B, et al. TLR4 and MD-2 expression is regulated by immune-mediated signals in human intestinal epithelial cells. J Biol Chem. 2002;277(23):20431-7. 203. Suzuki M, Hisamatsu T, Podolsky DK. Gamma interferon augments the intracellular pathway for lipopolysaccharide (LPS) recognition in human intestinal epithelial cells through coordinated up-regulation of LPS uptake and expression of the intracellular Toll-like receptor 4-MD-2 complex. Infect Immun. 2003;71(6):3503-11. 174  204. Lundin A, Bok CM, Aronsson L, Bjorkholm B, Gustafsson JA, Pott S, et al. Gut flora, Toll-like receptors and nuclear receptors: a tripartite communication that tunes innate immunity in large intestine. Cell Microbiol. 2008;10(5):1093-103. 205. Lee J, Mo JH, Katakura K, Alkalay I, Rucker AN, Liu YT, et al. Maintenance of colonic homeostasis by distinctive apical TLR9 signalling in intestinal epithelial cells. Nat Cell Biol. 2006;8(12):1327-36. 206. Bogunovic M, Dave SH, Tilstra JS, Chang DT, Harpaz N, Xiong H, et al. Enteroendocrine cells express functional Toll-like receptors. Am J Physiol Gastrointest Liver Physiol. 2007;292(6):G1770-83. 207. Podolsky DK, Gerken G, Eyking A, Cario E. Colitis-associated variant of TLR2 causes impaired mucosal repair because of TFF3 deficiency. Gastroenterology. 2009;137(1):209-20. 208. Neal MD, Leaphart C, Levy R, Prince J, Billiar TR, Watkins S, et al. Enterocyte TLR4 mediates phagocytosis and translocation of bacteria across the intestinal barrier. J Immunol. 2006;176(5):3070-9. 209. Rumio C, Besusso D, Palazzo M, Selleri S, Sfondrini L, Dubini F, et al. Degranulation of paneth cells via toll-like receptor 9. Am J Pathol. 2004;165(2):373-81. 210. Hayashi A, Sato T, Kamada N, Mikami Y, Matsuoka K, Hisamatsu T, et al. A single strain of Clostridium butyricum induces intestinal IL-10-producing macrophages to suppress acute experimental colitis in mice. Cell Host Microbe. 2013;13(6):711-22. 211. Isono A, Katsuno T, Sato T, Nakagawa T, Kato Y, Sato N, et al. Clostridium butyricum TO-A culture supernatant downregulates TLR4 in human colonic epithelial cells. Dig Dis Sci. 2007;52(11):2963-71. 212. Ivanov, II, Atarashi K, Manel N, Brodie EL, Shima T, Karaoz U, et al. Induction of intestinal Th17 cells by segmented filamentous bacteria. Cell. 2009;139(3):485-98. 213. Bergstrom KS, Sham HP, Zarepour M, Vallance BA. Innate host responses to enteric bacterial pathogens: a balancing act between resistance and tolerance. Cell Microbiol. 2012;14(4):475-84. 214. Brown SL, Riehl TE, Walker MR, Geske MJ, Doherty JM, Stenson WF, et al. Myd88-dependent positioning of Ptgs2-expressing stromal cells maintains colonic epithelial proliferation during injury. J Clin Invest. 2007;117(1):258-69. 215. Brandl K, Sun L, Neppl C, Siggs OM, Le Gall SM, Tomisato W, et al. MyD88 signaling in nonhematopoietic cells protects mice against induced colitis by regulating specific EGF receptor ligands. Proc Natl Acad Sci U S A. 2010;107(46):19967-72. 175  216. Rakoff-Nahoum S, Paglino J, Eslami-Varzaneh F, Edberg S, Medzhitov R. Recognition of commensal microflora by toll-like receptors is required for intestinal homeostasis. Cell. 2004;118(2):229-41. 217. Chassaing B, Aitken JD, Malleshappa M, Vijay-Kumar M. Dextran sulfate sodium (DSS)-induced colitis in mice. Curr Protoc Immunol. 2014;104:Unit 15 25. 218. Lupp C, Robertson ML, Wickham ME, Sekirov I, Champion OL, Gaynor EC, et al. Host-mediated inflammation disrupts the intestinal microbiota and promotes the overgrowth of Enterobacteriaceae. Cell Host Microbe. 2007;2(3):204. 219. Singh JC, Cruickshank SM, Newton DJ, Wakenshaw L, Graham A, Lan J, et al. Toll-like receptor-mediated responses of primary intestinal epithelial cells during the development of colitis. Am J Physiol Gastrointest Liver Physiol. 2005;288(3):G514-24. 220. Gradel KO, Nielsen HL, Schonheyder HC, Ejlertsen T, Kristensen B, Nielsen H. Increased short- and long-term risk of inflammatory bowel disease after salmonella or campylobacter gastroenteritis. Gastroenterology. 2009;137(2):495-501. 221. Khan MA, Bouzari S, Ma C, Rosenberger CM, Bergstrom KS, Gibson DL, et al. Flagellin-dependent and -independent inflammatory responses following infection by enteropathogenic Escherichia coli and Citrobacter rodentium. Infect Immun. 2008;76(4):1410-22. 222. Ruchaud-Sparagano MH, Maresca M, Kenny B. Enteropathogenic Escherichia coli (EPEC) inactivate innate immune responses prior to compromising epithelial barrier function. Cell Microbiol. 2007;9(8):1909-21. 223. de Kivit S, Tobin MC, Forsyth CB, Keshavarzian A, Landay AL. Regulation of Intestinal Immune Responses through TLR Activation: Implications for Pro- and Prebiotics. Front Immunol. 2014;5:60. 224. Ternhag A, Torner A, Svensson A, Ekdahl K, Giesecke J. Short- and long-term effects of bacterial gastrointestinal infections. Emerg Infect Dis. 2008;14(1):143-8. 225. Neal KR, Barker L, Spiller RC. Prognosis in post-infective irritable bowel syndrome: a six year follow up study. Gut. 2002;51(3):410-3. 226. Connor BA. Sequelae of traveler's diarrhea: focus on postinfectious irritable bowel syndrome. Clin Infect Dis. 2005;41 Suppl 8:S577-86. 227. Rodriguez LA, Ruigomez A. Increased risk of irritable bowel syndrome after bacterial gastroenteritis: cohort study. BMJ. 1999;318(7183):565-6. 228. Penders J, Thijs C, Mommers M, Stobberingh EE, Dompeling E, Reijmerink NE, et al. Host-microbial interactions in childhood atopy: toll-like receptor 4 (TLR4), CD14, and fecal Escherichia coli. J Allergy Clin Immunol. 2010;125(1):231-6 e1-5. 176  229. Loh G, Blaut M. Role of commensal gut bacteria in inflammatory bowel diseases. Gut Microbes. 2012;3(6):544-55. 230. Sherman PM, Ossa JC, Wine E. Bacterial infections: new and emerging enteric pathogens. Curr Opin Gastroenterol. 2010;26(1):1-4. 231. Baumler AJ, Sperandio V. Interactions between the microbiota and pathogenic bacteria in the gut. Nature. 2016;535(7610):85-93. 232. Tsolis RM, Xavier MN, Santos RL, Baumler AJ. How to become a top model: impact of animal experimentation on human Salmonella disease research. Infect Immun. 2011;79(5):1806-14. 233. Hapfelmeier S, Hardt WD. A mouse model for S. typhimurium-induced enterocolitis. Trends Microbiol. 2005;13(10):497-503. 234. Govoni G, Gros P. Macrophage NRAMP1 and its role in resistance to microbial infections. Inflamm Res. 1998;47(7):277-84. 235. Barthel M, Hapfelmeier S, Quintanilla-Martínez L, Kremer M, Rohde M, Hogardt M, et al. Pretreatment of mice with streptomycin provides a Salmonella enterica serovar Typhimurium colitis model that allows analysis of both pathogen and host. Infect Immun. 2003;71(5):2839-58. 236. Stecher B, Hapfelmeier S, Muller C, Kremer M, Stallmach T, Hardt WD. Flagella and chemotaxis are required for efficient induction of Salmonella enterica serovar Typhimurium colitis in streptomycin-pretreated mice. Infect Immun. 2004;72(7):4138-50. 237. Coburn B, Li Y, Owen D, Vallance BA, Finlay BB. Salmonella enterica serovar Typhimurium pathogenicity island 2 is necessary for complete virulence in a mouse model of infectious enterocolitis. Infect Immun. 2005;73(6):3219-27. 238. Stecher B, Robbiani R, Walker AW, Westendorf AM, Barthel M, Kremer M, et al. Salmonella enterica serovar typhimurium exploits inflammation to compete with the intestinal microbiota. PLoS Biol. 2007;5(10):2177-89. 239. Winter SE, Thiennimitr P, Winter MG, Butler BP, Huseby DL, Crawford RW, et al. Gut inflammation provides a respiratory electron acceptor for Salmonella. Nature. 2010;467(7314):426-9. 240. Boyd JF. Pathology of the alimentary tract in Salmonella typhimurium food poisoning. Gut. 1985;26(9):935-44. 241. Day DW, Mandal BK, Morson BC. The rectal biopsy appearances in Salmonella colitis. Histopathology. 1978;2(2):117-31. 242. Hapfelmeier S, Stecher B, Barthel M, Kremer M, Muller AJ, Heikenwalder M, et al. The Salmonella pathogenicity island (SPI)-2 and SPI-1 type III secretion systems allow Salmonella 177  serovar typhimurium to trigger colitis via MyD88-dependent and MyD88-independent mechanisms. J Immunol. 2005;174(3):1675-85. 243. Mundy R, MacDonald TT, Dougan G, Frankel G, Wiles S. Citrobacter rodentium of mice and man. Cell Microbiol. 2005;7(12):1697-706. 244. Eckmann L. Animal models of inflammatory bowel disease - Lessons from enteric infections. Inflammatory Bowel Disease: Genetics, Barrier Function, Immunologic Mechanisms, and Microbial Pathways. 2006:28-38. 245. Luperchio SA, Schauer DB. Molecular pathogenesis of Citrobacter rodentium and transmissible murine colonic hyperplasia. Microbes Infect. 2001;3(4):333-40. 246. Simmons CP, Clare S, Ghaem-Maghami M, Uren TK, Rankin J, Huett A, et al. Central role for B lymphocytes and CD4+ T cells in immunity to infection by the attaching and effacing pathogen Citrobacter rodentium. Infect Immun. 2003;71(9):5077-86. 247. Cario E. Barrier-protective function of intestinal epithelial Toll-like receptor 2. Mucosal Immunol. 2008;1 Suppl 1:S62-6. 248. Gibson DL, Montero M, Ropeleski MJ, Bergstrom KS, Ma C, Ghosh S, et al. Interleukin-11 reduces TLR4-induced colitis in TLR2-deficient mice and restores intestinal STAT3 signaling. Gastroenterology. 2010;139(4):1277-88. 249. Burke KE, Lamont JT. Clostridium difficile infection: a worldwide disease. Gut Liver. 2014;8(1):1-6. 250. Schutze GE, Willoughby RE, Committee on Infectious D, American Academy of P. Clostridium difficile infection in infants and children. Pediatrics. 2013;131(1):196-200. 251. Na JY, Park JM, Lee KS, Kang JO, Oh SH, Kim YJ. Clinical Characteristics of Symptomatic Clostridium difficile Infection in Children: Conditions as Infection Risks and Whether Probiotics Is Effective. Pediatr Gastroenterol Hepatol Nutr. 2014;17(4):232-8. 252. Kim J, Smathers SA, Prasad P, Leckerman KH, Coffin S, Zaoutis T. Epidemiological features of Clostridium difficile-associated disease among inpatients at children's hospitals in the United States, 2001-2006. Pediatrics. 2008;122(6):1266-70. 253. Rousseau C, Poilane I, De Pontual L, Maherault AC, Le Monnier A, Collignon A. Clostridium difficile carriage in healthy infants in the community: a potential reservoir for pathogenic strains. Clin Infect Dis. 2012;55(9):1209-15. 254. Voth DE, Ballard JD. Clostridium difficile toxins: mechanism of action and role in disease. Clin Microbiol Rev. 2005;18(2):247-63. 178  255. Chen X, Katchar K, Goldsmith JD, Nanthakumar N, Cheknis A, Gerding DN, et al. A mouse model of Clostridium difficile-associated disease. Gastroenterology. 2008;135(6):1984-92. 256. Hirota SA, Iablokov V, Tulk SE, Schenck LP, Becker H, Nguyen J, et al. Intrarectal instillation of Clostridium difficile toxin A triggers colonic inflammation and tissue damage: development of a novel and efficient mouse model of Clostridium difficile toxin exposure. Infect Immun. 2012;80(12):4474-84. 257. Ng J, Hirota SA, Gross O, Li Y, Ulke-Lemee A, Potentier MS, et al. Clostridium difficile toxin-induced inflammation and intestinal injury are mediated by the inflammasome. Gastroenterology. 2010;139(2):542-52, 52 e1-3. 258. Schnabl KL, Van Aerde JE, Thomson AB, Clandinin MT. Necrotizing enterocolitis: a multifactorial disease with no cure. World J Gastroenterol. 2008;14(14):2142-61. 259. Torrazza RM, Ukhanova M, Wang X, Sharma R, Hudak ML, Neu J, et al. Intestinal microbial ecology and environmental factors affecting necrotizing enterocolitis. PLoS One. 2013;8(12):e83304. 260. Terrin G, Passariello A, De Curtis M, Manguso F, Salvia G, Lega L, et al. Ranitidine is associated with infections, necrotizing enterocolitis, and fatal outcome in newborns. Pediatrics. 2012;129(1):e40-5. 261. Neu J, Pammi M. Pathogenesis of NEC: Impact of an altered intestinal microbiome. Semin Perinatol. 2017;41(1):29-35. 262. Le Huerou-Luron I, Blat S, Boudry G. Breast- v. formula-feeding: impacts on the digestive tract and immediate and long-term health effects. Nutr Res Rev. 2010;23(1):23-36. 263. Ballard O, Morrow, A. L. Human Milk Composition: Nutrients and Bioactive Factors. Pediatric Clinics of North America. 2013;60(1):49-74. 264. McGuire W, Anthony MY. Donor human milk versus formula for preventing necrotising enterocolitis in preterm infants: systematic review. Arch Dis Child Fetal Neonatal Ed. 2003;88(1):F11-4. 265. Brett KE, Ferraro ZM, Yockell-Lelievre J, Gruslin A, Adamo KB. Maternal-fetal nutrient transport in pregnancy pathologies: the role of the placenta. Int J Mol Sci. 2014;15(9):16153-85. 266. Viguera RM, Rojas-Castaneda J, Hernandez R, Reyes G, Alvarez C. Histological characteristics of the intestinal mucosa of the rat during the first year of life. Lab Anim. 1999;33(4):393-400. 267. Capuco AV, Akers RM. The origin and evolution of lactation. J Biol. 2009;8(4):37. 179  268. Howie PW, Forsyth JS, Ogston SA, Clark A, Florey CD. Protective effect of breast feeding against infection. BMJ. 1990;300(6716):11-6. 269. Smirnov A, Tako E, Ferket PR, Uni Z. Mucin gene expression and mucin content in the chicken intestinal goblet cells are affected by in ovo feeding of carbohydrates. Poult Sci. 2006;85(4):669-73. 270. Bergstrom A, Kristensen MB, Bahl MI, Metzdorff SB, Fink LN, Frokiaer H, et al. Nature of bacterial colonization influences transcription of mucin genes in mice during the first week of life. BMC Res Notes. 2012;5:402. 271. Nutrition ECo, Agostoni C, Braegger C, Decsi T, Kolacek S, Koletzko B, et al. Breast-feeding: A commentary by the ESPGHAN Committee on Nutrition. J Pediatr Gastroenterol Nutr. 2009;49(1):112-25. 272. Nommsen LA, Lovelady CA, Heinig MJ, Lonnerdal B, Dewey KG. Determinants of energy, protein, lipid, and lactose concentrations in human milk during the first 12 mo of lactation: the DARLING Study. Am J Clin Nutr. 1991;53(2):457-65. 273. Lonnerdal B. Human milk proteins: key components for the biological activity of human milk. Adv Exp Med Biol. 2004;554:11-25. 274. Donovan SM, Wang M, Li M, Friedberg I, Schwartz SL, Chapkin RS. Host-microbe interactions in the neonatal intestine: role of human milk oligosaccharides. Adv Nutr. 2012;3(3):450S-5S. 275. Uauy R, Castillo C. Lipid requirements of infants: implications for nutrient composition of fortified complementary foods. J Nutr. 2003;133(9):2962S-72S. 276. Saarinen UM, Kajosaari M, Backman A, Siimes MA. Prolonged breast-feeding as prophylaxis for atopic disease. Lancet. 1979;2(8135):163-6. 277. Mayer EJ, Hamman RF, Gay EC, Lezotte DC, Savitz DA, Klingensmith GJ. Reduced risk of IDDM among breast-fed children. The Colorado IDDM Registry. Diabetes. 1988;37(12):1625-32. 278. Hoddinott P, Tappin D, Wright C. Breast feeding. BMJ. 2008;336(7649):881-7. 279. Martin CR, Ling PR, Blackburn GL. Review of Infant Feeding: Key Features of Breast Milk and Infant Formula. Nutrients. 2016;8(5). 280. Lopez C, Cauty C, Guyomarc'h F. Organization of lipids in milks, infant milk formulas and various dairy products: role of technological processes and potential impacts. Dairy Sci Technol. 2015;95(6):863-93. 281. Hernell O, Timby N, Domellof M, Lonnerdal B. Clinical Benefits of Milk Fat Globule Membranes for Infants and Children. J Pediatr. 2016;173 Suppl:S60-5. 180  282. Bourlieu C, Michalski MC. Structure-function relationship of the milk fat globule. Curr Opin Clin Nutr Metab Care. 2015;18(2):118-27. 283. Bourlieu C, Bouzerzour, K., Ferret-Bernard, S., Bourgot, C. L., Chever, S., Ménard, O., Deglaire, A., Cuinet, I., Ruyet, P. L., Bonhomme, C., Dupont, D. and Huërou-Luron, I. L.   . Infant formula interface and fat source impact on neonatal digestion and gut microbiota. European Journal of Lipid Science and Technology. 2015. 284. Reinhardt TA, Lippolis JD. Bovine milk fat globule membrane proteome. J Dairy Res. 2006;73(4):406-16. 285. Murgiano L, Timperio AM, Zolla L, Bongiorni S, Valentini A, Pariset L. Comparison of milk fat globule membrane (MFGM) proteins of Chianina and Holstein cattle breed milk samples through proteomics methods. Nutrients. 2009;1(2):302-15. 286. Sando L, Pearson R, Gray C, Parker P, Hawken R, Thomson PC, et al. Bovine Muc1 is a highly polymorphic gene encoding an extensively glycosylated mucin that binds bacteria. J Dairy Sci. 2009;92(10):5276-91. 287. Le TT, Van de Wiele T, Do TN, Debyser G, Struijs K, Devreese B, et al. Stability of milk fat globule membrane proteins toward human enzymatic gastrointestinal digestion. J Dairy Sci. 2012;95(5):2307-18. 288. Novakovic P, Charavaryamath C, Moshynskyy I, Lockerbie B, Kaushik RS, Loewen ME, et al. Evaluation of inhibition of F4ac positive Escherichia coli attachment with xanthine dehydrogenase, butyrophilin, lactadherin and fatty acid binding protein. BMC Vet Res. 2015;11:238. 289. Bullen JJ. Iron-binding proteins and other factors in milk responsible for resistance to Escherichia coli. Ciba Found Symp. 1976(42):149-69. 290. Yu S, Lowe AW. The pancreatic zymogen granule membrane protein, GP2, binds Escherichia coli Type 1 fimbriae. BMC Gastroenterol. 2009;9:58. 291. Puiman P, Stoll B. Animal models to study neonatal nutrition in humans. Curr Opin Clin Nutr Metab Care. 2008;11(5):601-6. 292. Messer M, Thoman EB, Galofre A, Dallman T, Dallman PR. Artificial feeding of infant rats by continuous gastric infusion. J Nutr. 1969;98(4):404-10. 293. Clemente JC, Ursell LK, Parfrey LW, Knight R. The impact of the gut microbiota on human health: an integrative view. Cell. 2012;148(6):1258-70. 294. Mandeville KL, Krabshuis J, Ladep NG, Mulder CJ, Quigley EM, Khan SA. Gastroenterology in developing countries: issues and advances. World J Gastroenterol. 2009;15(23):2839-54. 181  295. Ohl ME, Miller SI. Salmonella: a model for bacterial pathogenesis. Annu Rev Med. 2001;52:259-74. 296. Araki A, Kanai T, Ishikura T, Makita S, Uraushihara K, Iiyama R, et al. MyD88-deficient mice develop severe intestinal inflammation in dextran sodium sulfate colitis. J Gastroenterol. 2005;40(1):16-23. 297. Malvin NP, Seno H, Stappenbeck TS. Colonic epithelial response to injury requires Myd88 signaling in myeloid cells. Mucosal Immunol. 2012;5(2):194-206. 298. Kirkland D, Benson A, Mirpuri J, Pifer R, Hou B, DeFranco AL, et al. B cell-intrinsic MyD88 signaling prevents the lethal dissemination of commensal bacteria during colonic damage. Immunity. 2012;36(2):228-38. 299. Asquith MJ, Boulard O, Powrie F, Maloy KJ. Pathogenic and protective roles of MyD88 in leukocytes and epithelial cells in mouse models of inflammatory bowel disease. Gastroenterology. 2010;139(2):519-29, 29.e1-2. 300. Shibata T, Takemura N, Motoi Y, Goto Y, Karuppuchamy T, Izawa K, et al. PRAT4A-dependent expression of cell surface TLR5 on neutrophils, classical monocytes and dendritic cells. Int Immunol. 2012;24(10):613-23. 301. Khan MA, Steiner TS, Sham HP, Bergstrom KS, Huang JT, Assi K, et al. The single IgG IL-1-related receptor controls TLR responses in differentiated human intestinal epithelial cells. J Immunol. 2010;184(5):2305-13. 302. Frantz AL, Rogier EW, Weber CR, Shen L, Cohen DA, Fenton LA, et al. Targeted deletion of MyD88 in intestinal epithelial cells results in compromised antibacterial immunity associated with downregulation of polymeric immunoglobulin receptor, mucin-2, and antibacterial peptides. Mucosal Immunol. 2012;5(5):501-12. 303. Chatfield SN, Strahan K, Pickard D, Charles IG, Hormaeche CE, Dougan G. Evaluation of Salmonella typhimurium strains harbouring defined mutations in htrA and aroA in the murine salmonellosis model. Microb Pathog. 1992;12(2):145-51. 304. Layton A, McKay L, Williams D, Garrett V, Gentry R, Sayler G. Development of Bacteroides 16S rRNA gene TaqMan-based real-time PCR assays for estimation of total, human, and bovine fecal pollution in water. Appl Environ Microbiol. 2006;72(6):4214-24. 305. Guo X, Xia X, Tang R, Zhou J, Zhao H, Wang K. Development of a real-time PCR method for Firmicutes and Bacteroidetes in faeces and its application to quantify intestinal population of obese and lean pigs. Lett Appl Microbiol. 2008;47(5):367-73. 306. Bacchetti De Gregoris T, Aldred N, Clare AS, Burgess JG. Improvement of phylum- and class-specific primers for real-time PCR quantification of bacterial taxa. J Microbiol Methods. 2011;86(3):351-6. 182  307. Fierer N, Jackson JA, Vilgalys R, Jackson RB. Assessment of soil microbial community structure by use of taxon-specific quantitative PCR assays. Appl Environ Microbiol. 2005;71(7):4117-20. 308. Hirota SA, Ng J, Lueng A, Khajah M, Parhar K, Li Y, et al. NLRP3 inflammasome plays a key role in the regulation of intestinal homeostasis. Inflamm Bowel Dis. 2011;17(6):1359-72. 309. Valdez Y, Grassl GA, Guttman JA, Coburn B, Gros P, Vallance BA, et al. Nramp1 drives an accelerated inflammatory response during Salmonella-induced colitis in mice. Cell Microbiol. 2009;11(2):351-62. 310. Barthel M, Hapfelmeier S, Quintanilla-Martinez L, Kremer M, Rohde M, Hogardt M, et al. Pretreatment of mice with streptomycin provides a Salmonella enterica serovar Typhimurium colitis model that allows analysis of both pathogen and host. Infect Immun. 2003;71(5):2839-58. 311. Hogan SP, Seidu L, Blanchard C, Groschwitz K, Mishra A, Karow ML, et al. Resistin-like molecule beta regulates innate colonic function: barrier integrity and inflammation susceptibility. J Allergy Clin Immunol. 2006;118(1):257-68. 312. Zarepour M, Bhullar K, Montero M, Ma C, Huang T, Velcich A, et al. The mucin Muc2 limits pathogen burdens and epithelial barrier dysfunction during Salmonella enterica serovar Typhimurium colitis. Infect Immun. 2013;81(10):3672-83. 313. Friedrich C, Mamareli P, Thiemann S, Kruse F, Wang Z, Holzmann B, et al. MyD88 signaling in dendritic cells and the intestinal epithelium controls immunity against intestinal infection with C. rodentium. PLoS Pathog. 2017;13(5):e1006357. 314. Melmed G, Thomas LS, Lee N, Tesfay SY, Lukasek K, Michelsen KS, et al. Human intestinal epithelial cells are broadly unresponsive to Toll-like receptor 2-dependent bacterial ligands: implications for host-microbial interactions in the gut. J Immunol. 2003;170(3):1406-15. 315. Otte JM, Cario E, Podolsky DK. Mechanisms of cross hyporesponsiveness to Toll-like receptor bacterial ligands in intestinal epithelial cells. Gastroenterology. 2004;126(4):1054-70. 316. Cario E, Rosenberg IM, Brandwein SL, Beck PL, Reinecker HC, Podolsky DK. Lipopolysaccharide activates distinct signaling pathways in intestinal epithelial cell lines expressing Toll-like receptors. J Immunol. 2000;164(2):966-72. 317. McKeown SJ, Chow CW, Young HM. Development of the submucous plexus in the large intestine of the mouse. Cell Tissue Res. 2001;303(2):301-5. 318. Dewey KG, Heinig MJ, Nommsen-Rivers LA. Differences in morbidity between breast-fed and formula-fed infants. J Pediatr. 1995;126(5 Pt 1):696-702. 319. Saarinen UM, Kajosaari M. Breastfeeding as prophylaxis against atopic disease: prospective follow-up study until 17 years old. Lancet. 1995;346(8982):1065-9. 183  320. Chatterton DE, Nguyen, D.N., Bering, S.B., Sanglid, P. T. Anti-inflammatory mechanisms of bioactive milk proteins in the intestine of newborns. The International Journal of Biochemistry & Cell Biology. 2013;45:1730– 47. 321. Boudry G, Morise A, Seve B, I LEH-L. Effect of milk formula protein content on intestinal barrier function in a porcine model of LBW neonates. Pediatr Res. 2011;69(1):4-9. 322. Mudd AT, Alexander LS, Berding K, Waworuntu RV, Berg BM, Donovan SM, et al. Dietary Prebiotics, Milk Fat Globule Membrane, and Lactoferrin Affects Structural Neurodevelopment in the Young Piglet. Front Pediatr. 2016;4:4. 323. Timby N, Domellof E, Hernell O, Lonnerdal B, Domellof M. Neurodevelopment, nutrition, and growth until 12 mo of age in infants fed a low-energy, low-protein formula supplemented with bovine milk fat globule membranes: a randomized controlled trial. Am J Clin Nutr. 2014;99(4):860-8. 324. Timby N, Hernell O, Vaarala O, Melin M, Lonnerdal B, Domellof M. Infections in infants fed formula supplemented with bovine milk fat globule membranes. J Pediatr Gastroenterol Nutr. 2015;60(3):384-9. 325. Sprong RC, Hulstein MF, Lambers TT, van der Meer R. Sweet buttermilk intake reduces colonisation and translocation of Listeria monocytogenes in rats by inhibiting mucosal pathogen adherence. Br J Nutr. 2012;108(11):2026-33. 326. Snow DR, Jimenez-Flores R, Ward RE, Cambell J, Young MJ, Nemere I, et al. Dietary milk fat globule membrane reduces the incidence of aberrant crypt foci in Fischer-344 rats. J Agric Food Chem. 2010;58(4):2157-63. 327. Snow DR, Ward RE, Olsen A, Jimenez-Flores R, Hintze KJ. Membrane-rich milk fat diet provides protection against gastrointestinal leakiness in mice treated with lipopolysaccharide. J Dairy Sci. 2011;94(5):2201-12. 328. Hall WG. Weaning and growth of artificially reared rats. Science. 1975;190(4221):1313-5. 329. Bhinder G, Stahl M, Sham HP, Crowley SM, Morampudi V, Dalwadi U, et al. Intestinal epithelium-specific MyD88 signaling impacts host susceptibility to infectious colitis by promoting protective goblet cell and antimicrobial responses. Infect Immun. 2014;82(9):3753-63. 330. Lau JT, Whelan FJ, Herath I, Lee CH, Collins SM, Bercik P, et al. Capturing the diversity of the human gut microbiota through culture-enriched molecular profiling. Genome Med. 2016;8(1):72. 184  331. Whelan FJ, Verschoor CP, Stearns JC, Rossi L, Luinstra K, Loeb M, et al. The loss of topography in the microbial communities of the upper respiratory tract in the elderly. Ann Am Thorac Soc. 2014;11(4):513-21. 332. Bartram AK, Lynch MD, Stearns JC, Moreno-Hagelsieb G, Neufeld JD. Generation of multimillion-sequence 16S rRNA gene libraries from complex microbial communities by assembling paired-end illumina reads. Appl Environ Microbiol. 2011;77(11):3846-52. 333. Martin M. Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnetjournal. 2011;17(1):10-2. 334. Masella AP, Bartram AK, Truszkowski JM, Brown DG, Neufeld JD. PANDAseq: paired-end assembler for illumina sequences. BMC Bioinformatics. 2012;13:31. 335. Joshi NA FJ. Sickle: A sliding-window, adaptive, quality-based trimming tool for FastQ files  (Version 1.33) [Software].  Available at https://github.com/najoshi/sickle. 2011. Date accessed: July 11, 2016. 336. Edgar RC. Search and clustering orders of magnitude faster than BLAST. Bioinformatics. 2010;26(19):2460-1. 337. Ye Y. Identification and Quantification of Abundant Species from Pyrosequences of 16S rRNA by Consensus Alignment. Proceedings (IEEE Int Conf Bioinformatics Biomed). 2011;2010:153-7. 338. Wang Q, Garrity GM, Tiedje JM, Cole JR. Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl Environ Microbiol. 2007;73(16):5261-7. 339. DeSantis TZ, Hugenholtz P, Larsen N, Rojas M, Brodie EL, Keller K, et al. Greengenes, a chimera-checked 16S rRNA gene database and workbench compatible with ARB. Appl Environ Microbiol. 2006;72(7):5069-72. 340. Caporaso JG, Kuczynski J, Stombaugh J, Bittinger K, Bushman FD, Costello EK, et al. QIIME allows analysis of high-throughput community sequencing data. Nat Methods. 2010;7(5):335-6. 341. Hansen A, Alston L, Tulk SE, Schenck LP, Grassie ME, Alhassan BF, et al. The P2Y6 receptor mediates Clostridium difficile toxin-induced CXCL8/IL-8 production and intestinal epithelial barrier dysfunction. PLoS One. 2013;8(11):e81491. 342. Segata N, Izard J, Waldron L, Gevers D, Miropolsky L, Garrett WS, et al. Metagenomic biomarker discovery and explanation. Genome Biol. 2011;12(6):R60. 343. Lonnerholm G, Wistrand P. Carbonic anhydrase in the human fetal gastrointestinal tract. Biol Neonate. 1983;44(3):166-76. 185  344. Amasaki T, Amasaki H, Nagasao J, Ichihara N, Asari M, Nishita T, et al. Immunohistochemical localization of carbonic anhydrase isoenzymes in salivary gland and intestine in adult and suckling pigs. J Vet Med Sci. 2001;63(9):967-70. 345. Chan JM, Bhinder G, Sham HP, Ryz N, Huang T, Bergstrom KS, et al. CD4+ T cells drive goblet cell depletion during Citrobacter rodentium infection. Infect Immun. 2013;81(12):4649-58. 346. Bullen CL, Tearle PV, Willis AT. Bifidobacteria in the intestinal tract of infants: an in-vivo study. J Med Microbiol. 1976;9(3):325-33. 347. Dvorak B, McWilliam DL, Williams CS, Dominguez JA, Machen NW, McCuskey RS, et al. Artificial formula induces precocious maturation of the small intestine of artificially reared suckling rats. J Pediatr Gastroenterol Nutr. 2000;31(2):162-9. 348. Yeh KY, Yeh M. Use of pup in a cup model to study gastrointestinal development: interaction of nutrition and pituitary hormones. J Nutr. 1993;123(2 Suppl):378-81. 349. Zhang L, Li N, des Robert C, Fang M, Liboni K, McMahon R, et al. Lactobacillus rhamnosus GG decreases lipopolysaccharide-induced systemic inflammation in a gastrostomy-fed infant rat model. J Pediatr Gastroenterol Nutr. 2006;42(5):545-52. 350. Fan W, Huo G, Li X, Yang L, Duan C, Wang T, et al. Diversity of the intestinal microbiota in different patterns of feeding infants by Illumina high-throughput sequencing. World J Microbiol Biotechnol. 2013;29(12):2365-72. 351. Lee SA, Lim JY, Kim BS, Cho SJ, Kim NY, Kim OB, et al. Comparison of the gut microbiota profile in breast-fed and formula-fed Korean infants using pyrosequencing. Nutr Res Pract. 2015;9(3):242-8. 352. Fan W, Tang Y, Qu Y, Cao F, Huo G. Infant formula supplemented with low protein and high carbohydrate alters the intestinal microbiota in neonatal SD rats. BMC Microbiol. 2014;14:279. 353. Azad MB, Konya T, Maughan H, Guttman DS, Field CJ, Chari RS, et al. Gut microbiota of healthy Canadian infants: profiles by mode of delivery and infant diet at 4 months. CMAJ. 2013;185(5):385-94. 354. Abrahamse E, Minekus M, van Aken GA, van de Heijning B, Knol J, Bartke N, et al. Development of the Digestive System-Experimental Challenges and Approaches of Infant Lipid Digestion. Food Dig. 2012;3(1-3):63-77. 355. Bry L, Falk PG, Midtvedt T, Gordon JI. A model of host-microbial interactions in an open mammalian ecosystem. Science. 1996;273(5280):1380-3. 186  356. Engevik MA, Yacyshyn MB, Engevik KA, Wang J, Darien B, Hassett DJ, et al. Human Clostridium difficile infection: altered mucus production and composition. Am J Physiol Gastrointest Liver Physiol. 2015;308(6):G510-24. 357. Wurzelmann JI, Lyles CM, Sandler RS. Childhood infections and the risk of inflammatory bowel disease. Dig Dis Sci. 1994;39(3):555-60. 358. Garcia Rodriguez LA, Ruigomez A, Panes J. Acute gastroenteritis is followed by an increased risk of inflammatory bowel disease. Gastroenterology. 2006;130(6):1588-94. 359. Ericsson AC, Crim MJ, Franklin CL. A brief history of animal modeling. Mo Med. 2013;110(3):201-5. 360. Yue F, Cheng Y, Breschi A, Vierstra J, Wu W, Ryba T, et al. A comparative encyclopedia of DNA elements in the mouse genome. Nature. 2014;515(7527):355-64. 361. Paigen K. One hundred years of mouse genetics: an intellectual history. II. The molecular revolution (1981-2002). Genetics. 2003;163(4):1227-35. 362. C. T. Streba CCV, C. Miscu, D. I. Gheonea, L. Sandulescu, T. Cuirea, I. Rogoveanu, A. Saftoiu. Of Mice and Ethics. Current Health Sciences Jounral. 2012;38(1):5-8. 363. Canadian Council on Animal Care.  About the CCAC [Internet]. Ottawa, ON. Available from  [http://www.ccac.ca/en_/about].  Date accessed: April 3, 2017. 364. Treuting PM, Dintzis, S. M. Lower Gastrointestinal Tract.  Comparative Anatomy and Histology: A Mouse and Human Atlas: Elsevier Inc.; 2012. p. 177-92. 365. Gibbons DL, Spencer J. Mouse and human intestinal immunity: same ballpark, different players; different rules, same score. Mucosal Immunol. 2011;4(2):148-57. 366. Bouskra D, Brezillon C, Berard M, Werts C, Varona R, Boneca IG, et al. Lymphoid tissue genesis induced by commensals through NOD1 regulates intestinal homeostasis. Nature. 2008;456(7221):507-10. 367. Spencer J, MacDonald TT, Finn T, Isaacson PG. The development of gut associated lymphoid tissue in the terminal ileum of fetal human intestine. Clin Exp Immunol. 1986;64(3):536-43. 368. Peng SL. Signaling in B cells via Toll-like receptors. Curr Opin Immunol. 2005;17(3):230-6. 369. de Medeiros CB, Fleming AS, Johnston CC, Walker CD. Artificial rearing of rat pups reveals the beneficial effects of mother care on neonatal inflammation and adult sensitivity to pain. Pediatr Res. 2009;66(3):272-7. 187  370. Lomanowska AM, Melo AI. Deconstructing the function of maternal stimulation in offspring development: Insights from the artificial rearing model in rats. Horm Behav. 2016;77:224-36. 371. Caicedo RA, Li N, Des Robert C, Scumpia PO, Hubsher CP, Wasserfall CH, et al. Neonatal formula feeding leads to immunological alterations in an animal model of type 1 diabetes. Pediatr Res. 2008;63(3):303-7. 372. Barreau F, Ferrier L, Fioramonti J, Bueno L. Neonatal maternal deprivation triggers long term alterations in colonic epithelial barrier and mucosal immunity in rats. Gut. 2004;53(4):501-6. 373. Alberts JR, May B. Ontogeny of olfaction: development of the rats' sensitivity to urine and amyl acetate. Physiol Behav. 1980;24(5):965-70. 374. Alberts JR. Olfactory contributions to behavioral development in rodents. New York: Academic Press; 1976. 375. Ley RE, Turnbaugh PJ, Klein S, Gordon JI. Microbial ecology: human gut microbes associated with obesity. Nature. 2006;444(7122):1022-3. 376. Rodriguez JM, Murphy K, Stanton C, Ross RP, Kober OI, Juge N, et al. The composition of the gut microbiota throughout life, with an emphasis on early life. Microb Ecol Health Dis. 2015;26:26050. 377. Marcobal A, Barboza M, Sonnenburg ED, Pudlo N, Martens EC, Desai P, et al. Bacteroides in the infant gut consume milk oligosaccharides via mucus-utilization pathways. Cell Host Microbe. 2011;10(5):507-14. 378. Juntunen M, Kirjavainen PV, Ouwehand AC, Salminen SJ, Isolauri E. Adherence of probiotic bacteria to human intestinal mucus in healthy infants and during rotavirus infection. Clin Diagn Lab Immunol. 2001;8(2):293-6. 379. Tao R, de Zoeten EF, Ozkaynak E, Chen C, Wang L, Porrett PM, et al. Deacetylase inhibition promotes the generation and function of regulatory T cells. Nat Med. 2007;13(11):1299-307. 380. Suzuki K, Maruya M, Kawamoto S, Sitnik K, Kitamura H, Agace WW, et al. The sensing of environmental stimuli by follicular dendritic cells promotes immunoglobulin A generation in the gut. Immunity. 2010;33(1):71-83. 381. Macia L, Tan J, Vieira AT, Leach K, Stanley D, Luong S, et al. Metabolite-sensing receptors GPR43 and GPR109A facilitate dietary fibre-induced gut homeostasis through regulation of the inflammasome. Nat Commun. 2015;6:6734. 188  382. Harrison OJ, Srinivasan N, Pott J, Schiering C, Krausgruber T, Ilott NE, et al. Epithelial-derived IL-18 regulates Th17 cell differentiation and Foxp3(+) Treg cell function in the intestine. Mucosal Immunol. 2015;8(6):1226-36. 383. Aujnarain A, Mack DR, Benchimol EI. The role of the environment in the development of pediatric inflammatory bowel disease. Curr Gastroenterol Rep. 2013;15(6):326. 384. Amre DK, Lambrette P, Law L, Krupoves A, Chotard V, Costea F, et al. Investigating the hygiene hypothesis as a risk factor in pediatric onset Crohn's disease: a case-control study. Am J Gastroenterol. 2006;101(5):1005-11. 385. Jakobsen C, Paerregaard A, Munkholm P, Wewer V. Environmental factors and risk of developing paediatric inflammatory bowel disease -- a population based study 2007-2009. J Crohns Colitis. 2013;7(1):79-88. 386. Buret AG. Enteropathogen-Induced Microbiota Biofilm Disruptions and Post-Infectious Intestinal Inflammatory Disorders. Current Tropical Medicine Reports. 2016;3(3):94-101. 387. Li H, Limenitakis JP, Fuhrer T, Geuking MB, Lawson MA, Wyss M, et al. The outer mucus layer hosts a distinct intestinal microbial niche. Nat Commun. 2015;6:8292. 388. Macfarlane S, Woodmansey EJ, Macfarlane GT. Colonization of mucin by human intestinal bacteria and establishment of biofilm communities in a two-stage continuous culture system. Appl Environ Microbiol. 2005;71(11):7483-92. 389. Ahmed I, Roy B, McFadden R-m, Anant S, Septer S, Umar S. Abstract 3297: Altered mucus composition and bacterial dysbiosis promote development of colitis following chronic Notch inhibition. Cancer Research. 2016;76(14 Supplement):3297-. 390. Hadjiagapiou C, Schmidt L, Dudeja PK, Layden TJ, Ramaswamy K. Mechanism(s) of butyrate transport in Caco-2 cells: role of monocarboxylate transporter 1. Am J Physiol Gastrointest Liver Physiol. 2000;279(4):G775-80. 391. Stark PL, Lee A. The microbial ecology of the large bowel of breast-fed and formula-fed infants during the first year of life. J Med Microbiol. 1982;15(2):189-203. 392. Backhed F, Roswall J, Peng Y, Feng Q, Jia H, Kovatcheva-Datchary P, et al. Dynamics and Stabilization of the Human Gut Microbiome during the First Year of Life. Cell Host Microbe. 2015;17(6):852. 393. Pantoja-Feliciano IG, Clemente JC, Costello EK, Perez ME, Blaser MJ, Knight R, et al. Biphasic assembly of the murine intestinal microbiota during early development. ISME J. 2013;7(6):1112-5. 394. Wlodarska M, Thaiss CA, Nowarski R, Henao-Mejia J, Zhang JP, Brown EM, et al. NLRP6 inflammasome orchestrates the colonic host-microbial interface by regulating goblet cell mucus secretion. Cell. 2014;156(5):1045-59. 189  395. Patel KK, Miyoshi H, Beatty WL, Head RD, Malvin NP, Cadwell K, et al. Autophagy proteins control goblet cell function by potentiating reactive oxygen species production. EMBO J. 2013;32(24):3130-44. 396. Korpela K, Salonen A, Virta LJ, Kekkonen RA, Forslund K, Bork P, et al. Intestinal microbiome is related to lifetime antibiotic use in Finnish pre-school children. Nat Commun. 2016;7:10410. 397. Knoop KA, McDonald KG, Kulkarni DH, Newberry RD. Antibiotics promote inflammation through the translocation of native commensal colonic bacteria. Gut. 2016;65(7):1100-9.   

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