colonisation and succession of the gut microbiota in suckling- and newly weaned piglets · 3....

Colonisation and succession of the gut microbiotain suckling- and newly weaned piglets

A study on the influence of specific nutritional, probiotic, antibiotic, andgenotypic factors

Ph.D. thesis byAnn-Sofie Riis Poulsen

Section for Immunology and MicrobiologyDepartment of Animal Science

Faculty of Science and TechnologyAarhus University, Foulum

Denmark

September 2016

SupervisorsSenior Scientist Charlotte Lauridsen, Ph.D., Department of Animal Science,Faculty of Science and Technology, Aarhus University, Foulum, Denmark.

Senior Scientist Nuria Canibe, Ph.D., Department of Animal Science, Faculty ofScience and Technology, Aarhus University, Foulum, Denmark.

Assessment committeéProfessor Tine Rask Licht, National Food Institute, Technical University ofDenmark, Denmark.

Senior Research Scientist Claire Rogel-Gaillard, Génétique Animale et BiologieIntégrative (GABI, UMR 1313), INRA, France.

Senior Scientist Stig Purup (chairman), Department of Animal Science, Faculty ofScience and Technology, Aarhus University, Foulum, Denmark.

iii

PrefaceThis Ph.D. thesis has been submitted to the Faculty of Science and Technology,Aarhus University. It comprises experimental work conducted within the groupof Immunology and Microbiology, Department of Animal Science, in the periodFebruary 2013 to September 2016. Included is a three months stay at Centerfor Microbial Communities, Department of Chemistry and Bioscience, AalborgUniversity, with primary focus on next-generation sequencing. The presentedwork contributes to the pool of data generated in the NEOMUNE consortium(http://neomune.ku.dk).

The Ph.D. project has been supervised by senior scientist Charlotte Lauridsenas main supervisor, and senior scientist Nuria Canibe as co-supervisor; both fromthe research group in Immunology and Microbiology.

The project has been funded partly by the Graduate School of Science andTechnology and partly by the NEOMUNE consortium through Innovation FundDenmark.

Ann-Sofie Riis PoulsenSeptember, 2016Foulum

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AcknowledgementsFirst and foremost I want to thank my supervisors, Nuria Canibe and CharlotteLauridsen, who have been supportive in so many ways. Not only have you educatedme in the difficult art of scientific thinking and writing, you have also always beenthere when moral support and encouragement was needed. It has been a pleasurebeing under your supervision.

Experimental tasks would not have been completed without help from thelaboratory technicians in the group of Immunology and Microbiology. I want tothank Karin Durup, Trine Poulsen, Thomas Rebsdorf and Mona Dinsen. I amgrateful to all of you for teaching me microbiological techniques and for yourinvaluable help during experimental periods. Warm thanks to Helle Handll forhelping me with hundreds of DNA extractions and qPCR tasks during the finalstage of my project. Also, heartfelt thanks to Inger Marie Jepsen, Lene RosborgDal and Hanne Purup for your help during my final animal experiment. Thanksto the staff of SB for taking good care of the piglets and for your help duringweekends and holidays.

I also want to thank Nadieh de Jonge and Jeppe Lund Nielsen for havingme in their molecular laboratory in Aalborg, teaching me about next-generationsequencing and data analysis. Thank you, Nadieh, for putting so much hard workinto my project and for never being further away than an e-mail.

I want to thank everyone in the group of Immunology and Microbiology. Youhave been great colleagues during the last three years and I have enjoyed gettingto know every single one of you. Your kindness and openness has been encouragingand deliberating. A very special thanks to my office mate Samantha Noel. Thankyou for your technical and moral support and our talks about everything andnothing. Also warm thanks to Ditte Søvsø Gundelund Nielsen. I truly appreciateyour friendship, your always kind support and for our many talks about frustratingPh.D. stuff, but also about other more joyful things as our children and families.

Lastly, I want to thank my family for putting up with all the ’pig-talk’, theirunderstanding and support. Especially, I want to thank my husband and daughter,Rasmus and My, for always being there, especially when the Ph.D. life just wasn’tthat fun and for their understanding during busy periods.

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SummaryDenmark produces close to 30 million pigs annually. The production is enabled byintensive production systems, where newborn and newly weaned piglets frequentlyshow clinical symptoms of gastrointestinal disease. Such clinical symptoms oftenrequire antibiotic therapy, and large amounts of antibiotics are used especially inthe post-weaning period. The high consumption of antibiotics is of great concerndue to development of antibiotic resistant bacteria. In addition, a compromised guthealth is associated with reduced animal welfare. Hence, the need for improvinggut health of piglets is immense.

The gut is home to a community of bacteria known as the gut microbiota. Thegut microbiota is a complex and highly diverse community and is nowadays ap-preciated as an additional ’organ’ of the body, serving various functions in relationto host health. Numerous internal and external factors influence the compositionof the gut microbiota, especially during its development. These factors can po-tentially be used for improving gut health of suckling and newly weaned piglets.The primary aim of this Ph.D.-thesis was therefore to investigate, whether ma-nipulation of specific dietary and probiotic factors could shape the gut microbiotain a direction that favours healthier piglets. The aim was fulfilled by character-ising the colonisation and succession of the gut microbiota of piglets from birthuntil two weeks after weaning, using classical microbial culture and 16S rRNAgene sequencing. The investigated factors were specifically: Bovine colostrum andBacillus spp. spores combined with early antibiotic therapy. The effect of alpha-(1,2)-fucosyltransferase (FUT1 ) genotype variants was investigated as well.

The animal experiments showed, that: 1) The early milk-based diet had asignificant influence on the microbial community in the gut. When comparedto piglets fed a regular milk replacer, bovine colostrum reduced the incidence ofdiarrhoea and enhanced the growth of specific lactic acid bacteria genera. Pigletssuckling their dams performed expectedly better than the experimental groups. 2)Administrating gentamicin and Bacillus spp. spores influenced the microbiota ofthe distal part of the gut and was especially evident by a change in species richnessand diversity. Yet, administration of Bacillus spores had a negative effect on thefrequency of diarrhoea. 3) FUT1 genotype did not influence the colonisation asseen by analysing the faecal microbiota, though susceptible piglets did house ahigher number of haemolytic bacteria and enterobacteria in their guts. 4) Pigletage, and weaning, had a significant influence on the composition of the microbialcommunity of the gut.

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In conclusion, the investigated external factors, and especially diet, influencedthe colonisation of the piglet gut microbiota. No effect on the overall microbialcommunity was seen when FUT1 genotype was manipulated. Bovine colostrumshowed the most promising results in shaping the gut microbiota in a directionfavouring healthier piglets. We did not find a beneficial effect of Bacillus sporeswhen administered before, under and after gentamicin administration.

Future studies focusing on the gut microbial community in pigs should besupported by extensive functional analyses, and measures of clinical and subclinicalcondition. Such studies may provide a deeper insight into the actual significanceof microbial community changes in the gut.

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ResuméDanmark producerer årligt omkring 30 millioner svin. Denne høje produktionmuliggøres af intensive produktionssystemer, hvor der hos de helt unge grise idagene umiddelbart efter fødslen, og omkring fravænning, hyppigt forekommermavetarmlidelser og kliniske symptomer med diarré. Sådanne kliniske symptomerer ofte årsag til antibiotikabehandling, og en stor del af forbruget anvendes iforbindelse med fravænning. Det høje forbrug af antibiotika i svineproduktio-nen har skabt stor bekymring i forbindelse med opståen af antibiotikaresistentebakterier, og ydermere kompromitterer hyppigt forekommende mavetarmlidelserdyrevelfærden. Behovet for at forbedre tarmsundheden hos smågrise er derforstort.

Tarmen huser et samfund af bakterier, som udgør størstedelen af det der kendessom mikrobiotaen. Tarmens mikrobiota er et komplekst og diverst samfund. I dagfindes der utallige beviser for, at mikrobiotaen er et yderst vigtigt organ, der haren stor indflydelse på værtens sundhed. Adskillelige interne og eksterne faktorerpåvirker tarmens mikrobiota, især på etablerings- og udviklingsstadiet. Sådannefaktorer kan potentielt udnyttes til at fremme tarmsundheden hos diende- og nyligtfravænnede grise. Det primære formål med dette Ph.D.-studie var derfor at un-dersøge, om man ved at manipulere specifikke diætetiske og probiotiske faktorerkan påvirke mavetarmkanalens mikrobiota i en retning som favoriserer sunderedyr. Formålet blev opfyldt ved at karakterisere koloniseringen og udviklingenaf mavetarmkanalens mikrobiota, hos grise i perioden mellem fødsel og indtil touger efter fravænning, ved hjælp af klassisk mikrobiologisk dyrkning og 16S rRNAgensekventering. De specifikke undersøgte faktorer var: Bovin råmælk og Bacillusspp. sporer kombineret med tidlig antibiotikatildeling. Effekten af to forskelligealpha-(1,2)-fucosyltransferase (FUT1 ) genotyper blev ligeledes undersøgt.

Dyreforsøgene viste, at: 1) Den mælkebaserede diæt havde en betydelig ind-flydelse på det mikrobielle samfund i mave og tarm. I forhold til grise fodret medalmindelig mælkeerstatning, reducererede bovin råmælk forekomsten af diarré, oghavde en fordrende effekt på fremvæksten af visse slægter af mælkesyrebakterier.De somælksfodrede grise præsterede som forventeligt bedst. 2) Tildeling af gen-tamicin og Bacillus sporer påvirkede mikrobiotaen i den distale del af tarmkanalen,som især viste sig ved et ændret antal af arter og en ændret mikrobiel diversitet.Imidlertid havde tildelingen af Bacillus sporer en negativ effekt på forekomstenaf diarré. 3) FUT1 genotype havde ingen effekt på den mikrobielle kolonisering,pÃěvist ved analyse af den af den fæcale mikrobiota. Der blev dog fundet en be-

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tydelig højere forekomst af hæmolytiske bakterier og enterobakterier i tarmen hosmodtagelige grise. 4) Grisenes alder, og fravænning, havde en betydelig indflydelsepå sammensætningen af tarmkanalens mikrobiota.

På baggrund af de fremstillede resultater kunne det konkluderes, at de under-søgte eksterne faktorer, især diet, påvirkede tarmkanalens mikrobielle koloniseringhos smågrisene. Manipulering af FUT1 genotypen viste sig ikke at have nogeneffekt på det overordnede samfund. Bovin råmælk udviste størst potentiale til atpåvirke mavetarmkanalens mikrobiota i en gunstig retning, som ydermere var led-saget af sundere dyr. Derimod kunne der ikke påvises en gavnlig effekt af Bacillussporer i forbindelse med, og efter, indgivelse af gentamicin.

Fremtidige studier med fokus på grisens mikrobiota i mavetarmkanalen børunderstøttes af vidtgående funktionelle analyser og mål for grisene kliniske og sub-kliniske tilstand. Sådanne studier kan give en dybere indsigt i den reelle betydningaf ændringer i mavetarmkanalens mikrobiota.

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Abbreviations

AGP : Antibiotic Growth PromotersBC : Bovine Colostrumcfu : colony-forming unitsETEC : Enterotoxigenic Escherichia coliFAO : Food and Agriculture Organisation of the United NationsFUT1 : alpha-(1,2)-fucosyltransferaseIg : ImmunoglobulinLPS : LipopolysaccharideMR : Milk ReplacerNEC : Necrotising enterocolitisNGS : Next-generation sequencingOTU : Operational Taxonomical UnitPCR : Polymerase Chain ReactionrRNA : ribosomal RNASCFA : Short-Chain Fatty AcidWHO : World Health Organisation

x

List of original manuscriptsThe presented thesis is based on the following three original manuscripts:

1. Ann-Sofie Riis Poulsen, Nadieh de Jonge, Sugiharto Sugiharto, Jeppe LundNielsen, Charlotte Lauridsen and Nuria Canibe.

The microbial community of the gut differs between piglets fed sowmilk, milk replacer or bovine colostrum.

Submitted to British Journal of nutrition.

2. Ann-Sofie Riis Poulsen, Sugiharto Sugiharto, Nuria Canibe and CharlotteLauridsen.

Investigating the potential effects of alpha-(1,2)-fucosyltransferasegenotype on the establishment and succession of the gastrointesti-nal microbiota of young piglets.

Prepared for submission to Veterinary Microbiology.

3. Ann-Sofie Riis Poulsen, Nadieh de Jonge, Jeppe Lund Nielsen, Ole Højberg,Charlotte Lauridsen and Nuria Canibe.

Influence of Bacillus spp. spores and gentamicin on the gut mi-crobiota of suckling and newly weaned piglets.

In preparation.

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Other scientific presentations

1. Ann-Sofie Riis Poulsen, Nadieh de Jonge, Sugiharto Sugiharto, JeppeLund Nielsen, Charlotte Lauridsen, Nuria Canibe. Gut microbiota profileof piglets fed sow milk, bovine colostrum or milk replacer. Abstract/posterpresentation. Gut Microbiology - 20 years and counting. 10th INRA-RowettSymposium. France, 2016. Page 143.

2. Ann-Sofie Riis Poulsen, Nuria Canibe, Charlotte Lauridsen, SugihartoSugiharto, Lone Bruhn Madsen, Emøke Bendixen, Jamal Momeni. Alpha-(1,2)-fucosyltransferase genotype influences the gastrointestinal microbiota ofpiglets. Abstract/poster presentation. Digestive Physiology in Pigs Sympo-sium. Kliczkow, 2015. Page 272.

3. Ann-Sofie Riis Poulsen, Sugiharto Sugiharto, Charlotte Lauridsen, NuriaCanibe. Comparison of bovine colostrum, milk replacer and sow milk on gas-trointestinal microbiota and mucosal E.coli F18 attachment of piglets. Oralpresentation. International workshop on nutrition and intestinal microbiotahost interaction in the pig. 2013. Page 30.

xii Contents

Contents

General introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

Scope of thesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

1 Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31.1 Defining gut health . . . . . . . . . . . . . . . . . . . . . . . . . . . 31.2 The intestinal barrier . . . . . . . . . . . . . . . . . . . . . . . . . . 41.3 Functions of the gut microbiota . . . . . . . . . . . . . . . . . . . . 6

1.3.1 Host nutrition . . . . . . . . . . . . . . . . . . . . . . . . . . 71.3.2 Defence against pathogens . . . . . . . . . . . . . . . . . . . 8

1.4 Composition of the pig gut microbiota . . . . . . . . . . . . . . . . 101.4.1 Colonisation and succession . . . . . . . . . . . . . . . . . . 10

1.5 Factors influencing the gut microbiota . . . . . . . . . . . . . . . . 121.5.1 Diet . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121.5.2 Antibiotics . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151.5.3 Probiotics . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191.5.4 Host genetics . . . . . . . . . . . . . . . . . . . . . . . . . . 251.5.5 Investigated factors in the current thesis . . . . . . . . . . . 25

1.6 Methods for studying the gut microbiota . . . . . . . . . . . . . . . 301.6.1 Culture-based methods . . . . . . . . . . . . . . . . . . . . . 301.6.2 Next-generation sequencing . . . . . . . . . . . . . . . . . . 30

2 Objectives and hypotheses . . . . . . . . . . . . . . . . . . . . . . . 332.1 Project aim and objectives . . . . . . . . . . . . . . . . . . . . . . . 332.2 Experimental objectives and hypotheses . . . . . . . . . . . . . . . 33

3 Methodological approach . . . . . . . . . . . . . . . . . . . . . . . 353.1 Gastrointestinal samples . . . . . . . . . . . . . . . . . . . . . . . . 353.2 Characterising the gut bacterial community . . . . . . . . . . . . . 36

3.2.1 Sequencing of the 16S rRNA gene . . . . . . . . . . . . . . . 36

Contents xiii

3.2.2 Microbial plate culture . . . . . . . . . . . . . . . . . . . . . 37

4 Manuscript 1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39

5 Manuscript 2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75

6 Manuscript 3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .107

7 General discussion . . . . . . . . . . . . . . . . . . . . . . . . . . .1537.1 Manipulating the gut microbiota . . . . . . . . . . . . . . . . . . . 153

7.1.1 Milk-based dietary intervention . . . . . . . . . . . . . . . . 1547.1.2 ETEC F18 susceptibility . . . . . . . . . . . . . . . . . . . . 1557.1.3 Bacillus spores and gentamicin . . . . . . . . . . . . . . . . 156

7.2 Age-related succession of the gut microbiota . . . . . . . . . . . . . 1577.3 Sampling the gut microbial communities . . . . . . . . . . . . . . . 1587.4 High-throughput sequencing versus microbial culture . . . . . . . . 1597.5 Linking gut microbial community composition to gut health and

animal robustness . . . . . . . . . . . . . . . . . . . . . . . . . . . . 160

8 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .161

9 Perspectives and future work . . . . . . . . . . . . . . . . . . . . .163

Appendix A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .181

General introduction 1

General introduction

Intestinal disorders are highly prevalent in pigs reared in modern production sys-tems, and diarrhoea is a common clinical condition, especially in newborn andnewly weaned piglets (Kongsted et al., 2013; Lalles et al., 2007). The high preva-lence of diarrhoea in piglets poses a major challenge as it results in decreasedanimal welfare and high use of antibiotics, with the consequent risk of bacterialantibiotic resistance development (WHO, 2014).

Antibiotic resistance causes a major threat to human health. The ongoingincrease in multi-drug resistant bacterial infections (Blair et al., 2015) has empha-sised an increased need for prudent use of antibiotics, and has put major focuson the antibiotic consumption in production animals. In Denmark, pigs consume76 % of the total antibiotic mass consumed by production animals, horses, petsand fish, while only contributing with 43 % to the live biomass. Large amountsof antibiotics (approximately 42 %) are consumed by piglets in the post-weaningperiod (DANMAP, 2014).

From being perceived as a collection of disease promoting organisms, the gutmicrobiota has changed into a structure appreciated as an additional ’organ’ of thebody, serving numerous functions in relation to host biology and health (Sommerand Baeckhed, 2013). The early gut colonisation of the young pig is essential asa local protective mechanism, but also in maturing the adaptive immune system(Guarner and Malagelada, 2003). Intestinal disorders disrupting the gut microbialcommunity in early life may be jeopardising both the short- and long-term healthof the animal.

If the use of antibiotics is to be brought down, robustness of piglets againstinfectious diseases, especially those encountered immediately after weaning, hasto be improved. This may be accomplished by using specific dietary or probioticmeans to improve piglet gut health by beneficially influencing the colonisation ofthe gastrointestinal microbiota.

2 Scope of thesis

Scope of thesis

Gut health and piglet robustness are highly dependent on the gut microbiota,as well as the intestinal immune system and the epithelial barrier function. Theincluded studies and reviewed literature in this thesis focus on the gut microbiota,but a general introduction is also given to the concept of gut health, the barrierfunction of the intestinal wall, public health consequences of the high consumptionof antibiotics in the pig production, and the influencing factors investigated in thecurrent Ph.D.-study.

The present thesis investigates the impact of four factors on the gut microbialcomposition of piglets, with the end-point aim of obtaining healthy pigs with areduced use of antibiotics. The original studies included focus on the microbialcomposition and some fermentation metabolites. Functional studies (e.g. tran-scriptomics or proteomics) of the gut microbiota are not included. Wheneverpossible, background literature originating from pig studies is included. However,as current scientific literature on the gut microbiota comprises relevant studies inother mammalian species, these will be included when needed.

Chapter 1. Background 3

Chapter 1

Background

1.1 Defining gut health

’Gut health’ is a term often encountered in the scientific literature on human andanimal health. Although understood and appreciated by most, a precise definitionof a healthy gut is lacking (Bischoff, 2011). Due to the rather unspecific definitionson gut health, Bischoff (2011) proposed five major criteria for a healthy gastroin-testinal tract in humans, being 1) effective digestion and absorption of food, 2)absence of gastrointestinal illness, 3) normal and stable intestinal microbiota, 4)effective immune status, and 5) a status of well-being.

In veterinary medicine, as opposed to human medicine, only objective parame-ters are usable in the assessment of gut health status. The experimental veterinaryliterature proposes a variety of parameters as markers of gut health. In pigs, com-monly used markers are intestinal histomorphology, such as intestinal villous heightand crypt depth (e.g. Le Bon et al., 2010) and gene expression analyses on e.g.tight junction proteins, inflammatory mediators, oxidative stressors, and prolifer-ation and apoptosis markers (e.g. Alizadeh et al., 2015). From a microbiologicalpoint of view, a healthy gut is generally regarded as one with a balanced micro-biota, free from dysbiotic hallmarks as expansion of pathogens, loss of beneficialbacteria, and/or a reduced microbial diversity (Petersen and Round, 2014). Stud-ies focusing on the weaning process of piglets have contributed to our knowledgeabout which bacteria are supportive of a healthy animal, and which are poten-tial disease-causing organisms. The healthy pig gut has a high number of lacticacid bacteria, especially lactobacilli, and a low number of potential pathogenicbacteria. Such potential pathogenic bacteria are especially those belonging to theEnterobacteriaceae family, as for example some E. coli types and Salmonella. The

4 Chapter 1. Background

lactobacilli to enterobacteria ratio has traditionally been used as a measure of guthealth in pigs (Castillo et al., 2007).

An absolute state of optimal gut health is practically impossible to define, asgut health is a dynamic and relative concept. In addition, gut health status isthe result of a fine balance between gut microbiology, immunology and physiol-ogy (Lalles et al., 2007). The evaluation of selected gut health markers is oftendependent on the experimental set-up and subject in question. A definition of ahealthy gut has to be accompanied by a measure of the overall health conditionof the animal. An overall healthy pig is suggestively one that has an undisturbedgeneral condition, is in no need for antibiotic or analgesic therapy, and is able tocope with the given housing/environmental conditions.

The present thesis seeks to find gut health-promoting interventions. Success-fulness of these interventions is evaluated primarily from their effect on the gutmicrobiota and the occurrence of clinical symptoms originating from the gut. Asnot all animal experiments included a large number of animals, the studies werenot meant as diarrhoea studies per se. Hence, the microbiological aspect of guthealth is the focus of the current thesis.

1.2 The intestinal barrier

The intestinal mucosa is constantly bombarded with foreign antigens from ingestedmaterial and the gut microbiota, and defensive mechanisms are thus very well de-veloped, constituting physical, cellular, and chemical barriers. Optimal function ofthese barriers is pivotal to animal health. Figure 1.1 shows a schematic illustrationof the microscopic layers of the small intestine and the barrier structures presentedin the next paragraphs.

The most important barrier against antigens and microbes is the intestinalepithelial lining, consisting of an epithelial monolayer. Individual enterocytes arejoined by three groups of junctional proteins; 1) adherens junctions, 2) desmo-somes, and 3) tight junctions. While the first two groups of proteins in particularare responsible for mechanically linking neighbouring enterocytes, tight junctionsare responsible for limiting the intercellular space, being a crucial regulator ofparacellular permeability and only allowing certain electrolytes and substrates toby-pass the epithelial lining and enter the submucosal layers. Tight junction com-plexes consist of occludin, claudins, junctional adhesion molecules, and tricellulinproteins, which are spanning the cellular membranes near the apical membrane


(Groschwitz and Hogan, 2009). An increased intestinal permeability, also knownas a ’leaky gut’, may be caused by a down-regulation in the expression of tightjunction proteins, and has been associated with increased levels of inflammatorycytokines (Mankertz et al., 2000).

ç

Lumen

Peritoneal cavity

Epithelial monolayer

Superficial connective tissue layer (lamina propria)

Thin muscle layer= nerve plexus= blood vessel= lymph vessel= glands= dendritic cell= T-lymphocyte= B-lymphocyte= mucus

Tight junction

Microvilli

Crypt of LieberkühnOrigin of new enterocytes

Deep connective tissue layer

Peritoneum

Circular and longitudinal muscle layers

Figure 1.1: Schematic illustration of the microscopic structure of the small intestine

The intestinal luminal content is separated from the epithelial lining by aninner and an outer mucus layer. The outer layer (closest to the lumen) is looselyorganised and populated by bacteria, while the inner layer is firmly attached, andunder normal circumstances free from bacteria. Mucus is made up by glycosy-lated mucins (especially muc2 mucins) secreted by goblet cells (Johansson et al.,2011), acting as adhesins that enable binding of commensal and pathogenic bac-teria. For bacteria to be able to colonise and proliferate in the gut, the generalbelief is that the bacteria have to be able to adhere to the mucus layer, which re-quires the presence of surface-bound mucin-binding proteins (Juge, 2012). Besidesconstituting a physical barrier and acting as bacterial binding sites, mucins are


also important energy substrates to commensal bacteria (Johansson et al., 2011).Much research on this topic stems from human and mice studies. However, studiesin pigs confirm that MUC2 genes are expressed also in the pig intestine (Liu et al.,2014b). The importance of muc2 mucins as a protective barrier in the gut hasbeen demonstrated in mice, where muc2-deficient mice were more susceptible tointestinal infections, and experienced more mucosal damage upon infection, thandid wild-type mice (Bergstrom et al., 2010). Other mice studies show that theabsence of a gut microbiota results in a thinner intestinal mucus layer (Sharmaet al., 1995), indicating that the gut microbiota stimulates the intestinal mucusproduction.

The gut associated lymphoid tissue (GALT) consists of Peyer’s patches, lym-phoid follicles, intraepithelial lymphocytes and mesenteric lymph nodes, and pro-vide a cellular barrier against antigens and microorganisms. Peyer’s patches arewell-organised lymphoid structures located in the superficial connective tissue layerof the gut (lamina propria). These structures are dominated by T-lymphocytesand dendritic cells. The overlying epithelium is represented by microfold cells,which are specialised epithelial cells responsible for taking up luminal antigensand microbes, and delivering them to the underlying antigen-presenting dendriticcells (Min and Rhee, 2015). B-lymphocytes of the lamina propria are responsiblefor producing immunoglobulin A (IgA), the most abundant immunoglobulin of thegut. IgA is taken up by enterocytes and secreted to the intestinal lumen, where itcan interact with antigens and microbes by ’immune exclusion’, hence performingits chemical barrier function (Ramanan and Cadwell, 2016).

1.3 Functions of the gut microbiota

The gut microbiota is the total collection of bacteria, archeae, protozoa, vira, andyeasts in the stomach and intestines (Mackie et al., 1999; Mikkelsen et al., 2004).In the following text, the gut microbiota refers to the bacterial community only.

The commensal gut microbiota interacts with the host through numerous path-ways. The major mechanisms whereby the microbiota serves its beneficial effectsare by 1) contributing to host nutrition by synthesising vitamins and providing en-ergy through short-chain fatty acid (SCFA) production (Cummings et al., 2004);2) defending the host against enteric pathogens (Cummings et al., 2004; Sarkerand Gyr, 1992); and 3) stimulating local immune cell proliferation, thus playinga key role in maturing the adaptive immune system (Guarner and Malagelada,


2003)

1.3.1 Host nutrition

Feed components escaping digestion and absorption in the small intestine are fer-mented by the intestinal microbiota (Jha and Berrocoso, 2016). Carbohydratesincluding oligosaccharides, non-starch polysaccharides, and starch are the mostimportant substrates for bacterial fermentation, yet proteins can act as substratesas well (Williams et al., 2005). Figure 1.2 provides an overview of the producedmetabolites from carbohydrates and proteins, and figure 1.3 gives an example ofthe approximate concentrations of selected compounds.

Lactic acid Ethanol Formic acid Succinic acid

Proteins Amino acids

Carbohydrates Starch, non-starch polysaccharides

oligosaccharides

Acetic acid Propionic acid Butyric acid

Butyric acid CH4 Acetic acid

Primary metabolites

H2S

Acetic acid

Propionic acid CO2 H2 Butyric acid

BCFA Ammonia Amines Phenols Indols

CH4

Acetaldehyde

Propionic acid

Secondary metabolites

Figure 1.2: Overview of metabolites originating from carbohydrate and protein sources. Modifiedfrom Rist et al. (2013) and Venema and Do Carmo (2015). BCFA=branched chain fatty acids

Carbohydrates are fermented to organic compounds as lactic acid, succinicacid, and straight SCFAs (Macfarlane and Macfarlane, 2003). The major SCFAsin the gut are acetic, propionic and butyric acid, and these are generally regardedas being beneficial to the host (Williams et al., 2005). Acetic, propionic, and bu-tyric acid are converted to glucose by gluconeogenesis in liver cells, while skeletaland cardiac muscle cells are able to use acetic acid for energy production (Jha andBerrocoso, 2016). Some SCFAs are used as bacterial substrates and metabolisedto secondary metabolites (Venema and Do Carmo, 2015). SCFAs furthermore pro-


vide a substantial amount of energy to colonocytes, that especially prefer butyricacid. SCFAs have a trophic effect on intestinal cells (Burrin and Stoll, 2002),and butyrate has been shown to have anti-inflammatory potentials (Andriamihajaet al., 2010). Therefore, dietary interventions resulting in high concentrations ofSCFAs are generally considered beneficial to gut health (Williams et al., 2005).

Intestinal bacteria metabolise undigested and endogenous proteins by aminoacid deamination and decarboxylation (Jha and Berrocoso, 2016). End-products,besides those mentioned above for carbohydrates, include specific metabolites likebranched (short) chain fatty acids, as iso-butyric and iso-valeric acid, and po-tentially toxic compounds as ammonia, phenols, indols, and biogenic amines asputrescine and cadaverine (Williams et al., 2005). While iso-butyric acid has beensuggested to be a potential energy source for colonocytes (Blachier et al., 2007),some of the latter nitrogenous compounds (e.g. ammonia) are believed to havenegative effects on the host by impairing the mucosal barrier and interfering withcolonocyte metabolism (Blachier et al., 2007; Pieper et al., 2014). Thus, diets re-sulting in extensive protein fermentation, and high concentrations of nitrogenouscompounds, are considered as having a negative effect on gut health and shouldpreferably be avoided.

Both carbohydrate and protein metabolites are found in different concentra-tions along the length of the gut, and the highest fermentation activity is generallyseen in the caecum and colon (see figure 1.3).

1.3.2 Defence against pathogens

Local defence mechanisms of the gut microbiota include competing with pathogensfor mucosal binding sites and nutrients, production of antimicrobials, and elimi-nation of potential harmful/toxic substances (Cummings et al., 2004; Sarker andGyr, 1992). Production of organic acids by the gut microbiota (as described above)decreases gut pH, which inhibits growth of pathogens as Escherichia coli andSalmonella spp. (Canibe and Jensen, 2012). Lactic acid bacteria, especially Lac-tobacilli, are found in high numbers in the gut and are important in maintaining ahealthy gut (Daly et al., 2014). They are capable of reducing pH by lactic acid pro-duction, compete with potential pathogens, and produce bacteriocins (Hou et al.,2015), thereby reducing the number of pathogens and hindering their attachmentto the intestinal mucosa.

Human and mice studies have shown that the development of the gut mu-cosal immune system starts prenatally. Unlike the prenatal development, proper


Organic acids and ammoniam

mol

/l di

gest

a

0

15

30

45

60

Stomach Ileum Caecum Prox. colon Dist. colon

Lactic acid Acetic acid Propionic acidButyric acid Ammonia

Protein metabolites

umol

/l di

gest

a

0

400

800

1200

1600

Stomach Ileum Caecum Prox. colon Dist. colon

Amines Putrescine Cadaverine Phenol

Figure 1.3: Example of carbohydrate and protein metabolite concentrations along the length ofthe gut. Samples originate from six weeks old piglets three weeks post-weaning. Original datais from Pieper et al. (2014)

(i.e. resulting in a state of homeostasis) postnatal development and maturation oflymfoid structures in the gut is dependent on the presence of microbes (Sommerand Baeckhed, 2013). Germ-free mice have been reported to have underdevelopedmesenteric lymph nodes, intestinal lymphoid follicles and Peyer’s patches (reviewedby Round and Mazmanian (2009)), and on the cellular level, lines of natural killercells, and T- and B- lymphocytes are all modulated by the gut microbiota (re-viewed by Sommer and Baeckhed (2013)). These observations are supported bystudies conducted on pigs. Mulder et al. (2009) reported that manipulation ofthe gut microbiota, by rearing piglets in different environments, changed the ex-pression of genes involved in immunological processes. A similar observation wasdone when the gut microbiota of four-day-old piglets was manipulated by antibi-otic administration (Schokker et al., 2014). Thus, the gut microbiota is crucialin educating the intestinal immune system, thereby playing a major part in theanimal’s long-term ability to effectively launch an immunological response againstpathogens and pathological processes.


1.4 Composition of the pig gut microbiota

The microbial composition is highly influenced by intrinsic factors as pH, oxygentension, transit time, and hepatic and pancreatic secretions, which vary betweendifferent compartments (Buddington and Sangild, 2011), and the gut microbialcommunity thus consists of different niches along the length and width of the gut(Looft et al., 2014a). When moving from the proximal duodenum to the colon,microbial richness and diversity increase (Jensen, 1998; Looft et al., 2014a). Thenumber of culturable bacteria range from 103 per gram content in the stomach to1011 in the colon (Pluske et al., 2002)

Firmicutes and Bacteroidetes are the most abundant phyla in the gut (Kimand Isaacson, 2015; Looft et al., 2014a). Clostridium and Lactobacillus are amongthe most abundant genera in the Firmicutes phylum (Slifierz et al., 2015). As de-scribed in earlier sections, Lactobacillus and other lactic acid bacteria are generallyregarded as being beneficial to the host. Clostridium perfringens is a commonlyencountered Clostridium species, whereof some types are potential pathogens.Neonatal piglets are highly susceptible to C. perfringens type A and C, whichcause necrotic enteritis and ultimately result in high mortality rates (Songer andUzal, 2005). Prevotella and Bacteroides are among the highest abundant generaof the Bacteroidetes phylum, and are beneficial bacteria involved in the breakdownof polysaccharides and milk glycans (Frese et al., 2015).

1.4.1 Colonisation and succession

The process of acquiring a fully colonised gastrointestinal tract is a gradual andcomplex successional process beginning during (or immediately after) birth, whenbacteria originating from the mother (vaginal secretions, faeces and skin) and sur-rounding environment are ingested by the young (Mackie et al., 1999). Somebacteria are able to establish themselves and become permanent residents withoutthe need for further reintroduction (indigenous), while others are only temporaryresidents (non-indigenous) (Pluske et al., 2002). Species diversity and richnessincrease with age (Slifierz et al., 2015) and when fully established, the gut micro-biota constitutes a complex micro-environment existing in symbiosis with the host,and generally accepted as having a crucial influence on host physiology and health(Sommer and Baeckhed, 2013). The establishment of a gut microbiota is pivotalto the adaptation of the gut to the extra-uterine life (Buddington and Sangild,2011). Appendix A gives an overview of the dominating taxa along the piglet


gut during suckling and the first weeks after weaning. Data from sequencing andculture is included, and comprise mainly samples from the lower part of the gut.The below description, however, is impeded by the lack of sufficient comparablepapers originating from pig studies. Also there is a gap in the available literatureon the colonisation of the small intestine.

Immediately after birth, the piglet gut microbiota is dominated by bacteria be-longing to the Firmicutes and Proteobacteria phyla (Petri and Hill, 2010; Slifierzet al., 2015). Sequencing pooled samples from the stomach, small intestine, andcolon, Petri and Hill (2010) found the dominating genera to belong to Clostridi-aceae and Enterobacteriaceae. Swords et al. (1993) cultured specific microbialgroups and found Clostridium spp. and E. coli to be the most numerous gen-era in distal colonic digesta up until 4 days of age. From 1 day of age, bacteriabelonging to Streptococcaceae dominated the pooled community investigated byPetri and Hill (2010), until Lactobacillaceae and Lactobacillus appeared from 1week of age (Petri and Hill, 2010; Swords et al., 1993). Studies on the successionalpattern in faeces generally support the pattern seen in digesta. Sequencing datashow that the faecal microbiota is dominated by Clostridium, Escherichia, andFusobacterium immediately after birth. Nevertheless, as mentioned earlier, themicrobial communities are not identical along the gut, and these differences areapparent shortly after birth. Using microbial culturing, Gancarcikova et al. (2008)investigated specific groups in digesta from the jejunum, ileum and caecum, andfound differences as early as 2 days after birth, which were especially apparent be-tween the small intestinal compartments and the caecum, with more Lactobacilliin jejunum and ileum, and more Enterobacteriaceae in the caecum. From 1 weekof age, the relative abundances of Clostridium and Escherichia in faeces decrease,while Lactobacillus increases in relative abundance and stabilises around 4 weeksof age (Slifierz et al., 2015). The pig gut microbiota appears to have become arelatively stable community 3 to 4 weeks after weaning. See Appendix A for bac-terial fluctuations in different gut compartments between 1 and 3 weeks of age.Weaning at 21 days of age has been associated with an increase in bacteria belong-ing to Prevotellaceae and Ruminococcaceae (Frese et al., 2015). As these taxa areprimarily identified by sequencing, these findings are not supported by culture.

During early life, the gut microbiota is thus a highly dynamic and unstablecommunity. According to Thompson et al. (2008), the gut microbiota is most sus-ceptible to perturbations between 2-3 weeks of age. Stress caused by weaning isa major cause of bringing the piglet gut microbiota out of balance (Konstantinov


et al., 2006; Thompson et al., 2008), and such imbalances are believed to im-pair host-defensive mechanisms and increase the susceptibility to enteric disease(Castillo et al., 2007; Heo et al., 2013). Gut microbial perturbations during earlylife are potentially problematic as the early colonisers are active players in shapingthe ’adult’ microbiota (Guarner and Malagelada, 2003). Also, several studies in-volving germ-free and specific-pathogen-free mice suggest, that there is a ’windowof opportunity’ during early life, where the gut microbes present at the time havelasting effects on immune cell functions (Olszak et al., 2012) and stress responses(Sudo et al., 2004). It can thus be speculated that some microbial profiles in earlylife may be more beneficial than others.

1.5 Factors influencing the gut microbiota

Numerous studies in pigs, rodents, and humans have illustrated that the gut mi-crobiota is a dynamic ecosystem, which is influenced by a variety of internal (host)and external factors. These influencers include: Genetics, diet, stress, environ-ment, antibiotics, probiotics, and age. Different specific factors can potentially beutilised for beneficially manipulating the gut microbiota in favour of an improvedanimal health.

The impact of different dietary components, breed, antibiotics, and probioticson the pig gut microbiota and, when available, intestinal immunology, is describedin the following subsections.

1.5.1 Diet

Non-digested dietary components act as bacterial substrates (Jha and Berrocoso,2016) and vary between different diets. Table 1.1 summarises information on timeof intervention, type of sample analysed, microbial analysis, and diets from thepapers cited in the next paragraphs.

The neonatal piglet’s intestinal microbiota is dependent on the origin of themilk provided. Yeruva et al. (2016) found that the ileal microbiotas of 21-day-oldpiglets were different between those fed sow milk and those fed milk formula (cowmilk and soy based). The piglets fed sow milk had the highest relative abundanceof Lactobacillus and Clostridiaceae, while those fed milk formula had the highestabundance of Enterobacteriaceae and Lactococcus. The authors furthermore foundthat the lymphoid follicle sizes in the ileum and jejunum of piglets fed milk formula


were smaller than those from piglets fed sow milk, and thus indicated an effect ofa changed microbiota on the intestinal mucosal immune system.

The abrupt transition from milk to solid feed, occurring at weaning, is as-sociated with a change from a microbiota utilising primarily milk sugars to oneutilising plant fibers. Frese et al. (2015) reported an increase in Prevotellaceae,Ruminococcaceae and Lactobacillaceae, and a decrease in Enterobacteriaceae andBacteroidaceae after weaning. These changes were accompanied by a relative in-crease in bacterial genes encoding plant-degrading enzymes in the post-weaningperiod. These changes may not be a reflection of the isolated effect of diet, butmay also be caused by age and weaning.

Studies in grower and finisher pigs have shown that different protein and car-bohydrate types influence the intestinal microbial community. As these groups ofpigs are not within the scope of this thesis, they will not be addressed further. Seetable 1.1 for more details.


Tabl

e1.

1:K

eyin

form

atio

non

tim

eof

inte

rven

tion

,die

tary

com

pone

nts

inve

stig

ated

,ana

lyse

dsa

mpl

e(s)

,and

type

ofan

alys

isus

edin

the

cite

dst

udie

sin

vest

igat

ing

the

influ

ence

ofdi

eton

the

pig

gut

mic

robi

ota

Per

iod

offe

edin

terv

ention

Die

tary

com

pone

nts

Sam

ple(

s)A

naly

sis

Fin

ding

sR

efer

ence

One

mon

thfr

om14

.6kg

Soyb

ean

mea

lC

otte

nsee

dm

eal

Fis

hm

eal

Smal

lint

esti

nal

dige

sta

V4-

V5

16S

rRN

Age

nese

q.Illu

min

aFis

hm

eal)

"co

mm

unity

rich

ness

dom

inat

edby

Pro

teob

acte

ria.

Lactobacillus

and

Clostridium

wer

em

ore

abun

dant

inpi

glet

sfe

dso

y-be

anor

cott

onse

edm

eal.

Cao

etal

.(20

16)

Wea

ning

Nur

sing

Wea

ning

diet

Faec

esV

416

SrR

NA

gene

seq.

Illu

min

aSe

ete

xt.

Fres

eet

al.(

2015

)

3to

5m

onth

sof

age

Hig

h-fa

t/lo

w-fi

ber

Low

-fat

/hig

h-fib

erFa

eces

qPC

R16

SrR

NA

gene

regi

onLow

-fat

/hig

h-fib

erdi

et)

"ab

unda

nce

ofLactobacillus

spp.

,Clostridium

leptum

,Faecalibacterium

prausnitzii

and

Bifidobacterium

spp.

Hig

h-fa

t/lo

w-fi

ber

diet

)"

abun

danc

eof

Enterobacteriaceae

and#

[ace

tate

],[b

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ate]

and

[pro

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ate]

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heeff

ect

ofdi

etw

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nrit

zet

al.(

2016

)

70to

170

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ofag

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gest

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gene

seq.

Illu

min

aR

PS)

#co

loni

cm

icro

bial

rich

ness

and

#re

lati

veab

unda

nce

ofTuricibacter,

Rum

inococcus,B

lautia,C

oprococcus

and

Lachnospiraceae.

Diff

eren

tst

arch

type

sin

fluen

ced

the

expr

essi

onof

imm

une

rela

ted

gene

sdi

ffere

ntly

.

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etal

.(20

15)

Gro

wer

pigs

from

31kg

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ley

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atFa

eces

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ldig

esta

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NA

gene

regi

onB

arle

y)

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spp.

toEnterobacteriaceae

rati

o.W

eiss

etal

.(20

16)

2to

21da

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age

Sow

milk

Milk

form

ula

Ilea

ldig

esta

V4

16S

rRN

Age

nese

q.Illu

min

aSe

ete

xt.

Yer

uva

etal

.(20

16)


1.5.2 Antibiotics

Antibiotics have since their discovery been an invaluable tool in combating bac-terial infections. Furthermore, many countries still allow the use of antibiotics asgrowth promoters (AGPs). Table 1.2 summarises information on period of antibi-otic intervention, type of antibiotic(s), sample types, and analysis from the paperscited in the next paragraphs.

Several studies have shown that antibiotics in sub-therapeutic doses, i.e. whenadded as AGPs, influence the composition of the gut microbiota. Holman andChenier (2014) showed that the effect on the gut microbiota was temporal anddiffered between tylosin and chlortetracycline. Their results indicated that theobserved microbial differences were due to changes in the relative abundancesof the species present, and not due to whether the species were present or not.On the contrary, Looft et al. (2014b) reported both short- and long-term effectsin the faecal microbiota after feeding carbadox. In the first week of antibioticadministration, the relative abundances of Prevotella, Roseburia, Faecalibacteriumand Asteroleplasma were increased, while Lactobacillus was increased in the controlgroup. Even though the most clear separation between carbadox-treated and non-treated piglets was seen in the early period of carbadox administration, an effecton the microbial community was still observed 3 and 6 weeks after the end ofcarbadox administration. AGPs targeting especially gram postive bacteria havebeen shown to reduce the number of lactic acid bacteria (Jensen, 1998).

Changes to the gut microbiota have also been observed when pigs are adminis-tered parenteral therapeutic antibiotics at an early age. Benis et al. (2015) showedthat therapeutic administration of tulathromycin on day 4 after birth had a tem-poral effect on the jejunal microbiota, but did not observe any long-term effects.Long-term effects were, however, seen in the expression of genes related to im-munological functions and cell proliferations in the jejunal tissue. Using a similarexperimental set-up, Schokker et al. (2014) and Schokker et al. (2015) showed thatthe jejunal microbial diversity was increased in tulathromycin-treated (at 4 days ofage) piglets 8 days after birth and decreased on day 176. The increased diversityon day 8 was associated with an increased relative abundance in Bifidobacterium,Faecalibacterium, Eubacterium, and Solobacterium, and a decreased relative abun-dance of Bacillus and Staphylococcus. The decreased diversity on day 176 wasassociated with an increased relative abundance of Actinomyces, Allofustis andEggerthella, and a decreased abundance of Bifidobacterium, Fusobacterium, Neis-seria, and Oxalobacter. However, no effect on the microbial diversity was seen on


day 55, yet effects on intestinal gene expressions related to metabolic and immuno-logical functions were seen at this time.

Hence, though the microbiota in some circumstances apparently is able to re-establish itself, disturbances in the microbial community may in some cases belong-lasting and have lasting functional effects. Disturbances in the gut micro-biota in early life, and the long-term effect on the expression of genes involved inimmunological processes, are all suggesting that perturbations to the gut micro-biota in early life change the way the intestinal immunological functions develop.


Tabl

e1.

2:K

eyin

form

atio

non

tim

eof

anti

biot

icad

min

istr

atio

n,an

tibi

otic

(s)

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type

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mpl

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alys

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the

cite

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sin

vest

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ing

the

influ

ence

ofan

tibi

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son

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mic

robi

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=bo

dyw

eigh

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=su

bcut

aneo

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Peri

odof

antibi

otic

adm

inis

trat

ion

Ant

ibio

tic(

s)Sa

mpl

e(s)

Ana

lysi

sR

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ence

3to

19w

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ofag

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in44

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11m

g/kg

feed

Chl

orte

trac

yclin

e5.

5m

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feed

Faec

essa

mpl

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3,6,

9,12

and

19w

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V4

16S

rRN

Age

nese

q.Ill

umin

a

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man

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nier

(201

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ton

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Xti

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al.(

2014

b)

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ledo

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in2.

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sam

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d17

6da

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PIT

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pG

ene

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onSc

hokk

eret

al.(

2014

)Sc

hokk

eret

al.(

2015

)B

enis

etal

.(20

15)


Antibiotic use and public health consequences

Before 2006, in-feed AGPs were used as preventative measures in coping with post-weaning enteric diseases (DANMAP, 2014). However, as the association betweenin-feed antibacterials and antibiotic resistance became clear, this practice had tobe abandoned and use of in-feed AGPs in the European Union was banned (Heoet al., 2013). In the years following this ban, the amount of prescribed antibioticsfor use in pigs rose, and the pool of antibacterial resistant genes remained high(DANMAP, 2014). As practically no new antibiotics (or antibacterials) have beendiscovered since the late 1980’s (Silver, 2011), it is crucial that the antibiotics oftoday remain effective for many years to come.

Development of antibiotic resistance is a natural evolutionary process and thusinevitable. However, the tremendous doses of antibiotics used in animal medicineare significant risk factors in selecting single and multi-drug resistant bacteria(WHO, 2014). The most commonly used antibiotics for pigs are tetracyclines withnearly 29 tonnes active compound, followed by penicillins and macrolides. Morethan 50 % of the tetracyclines and macrolides are used for pigs in the post-weaningperiod. Even though the amount of antibiotics used in pigs decreased from 2013 to2014 (DANMAP, 2014), the amounts of antibiotics consumed by pigs in Denmarkare still too high. Hence continuous efforts are made to bring the amounts useddown.

Antibiotic resistance arising in bacteria from pigs poses major problems in re-lation to human and animal health. Humans are in risk of being infected withresistant bacteria originating from pigs. Methicillin-Resistant Staphylococcus Au-reus (clonal complex 398; MRSA CC398) is an example of this. Pigs are consideredthe primary reservoir of MRSA CC398 and the number of infected humans in Den-mark rose in the period from 2004 to 2011. The vast majority of infections wererelated to the skin and soft tissues (Larsen et al., 2015), though fatal cases withsepticaemia have been reported as well (Nielsen et al., 2016). Another exampleis Salmonella typhimurium, which is an important zoonotic food-borne pathogen(DANMAP, 2014). DANMAP (2014) reported 64 % of isolates from pigs to beresistant against ampicillin, sulfonamide, and tetracycline, and hence categorisedas multi-resistant. Isolates from humans showed the same pattern of resistance.Furthermore, there is a horisontal transfer of antibiotic resistance genes among bac-teria. Transposable elements as plasmids can be transferred from one bacteriumto another, thus enabling transfer of resistance genes (Penesyan et al., 2015). Thisis especially worrying as otherwise harmless bacteria can transfer resistance genes


to pathogenic bacteria (Miller et al., 2016). Thus, whether it is the pathogenic orcommensal bacteria that carry the genes encoding resistance against specific an-tibiotics, they pose a major threat to the modern treatment of bacterial infectionsas we know it.

1.5.3 Probiotics

Oral probiotics have for several years been defined as ’live microbial feed supple-ments which beneficially affect the host animal by improving its intestinal micro-bial balance’ (Fuller, 1989). A never definition, proposed by WHO and FAO in2002, says that probiotics are ’live microorganisms which, when administered inadequate amounts, confer a health benefit on the host’ (FAO/WHO, 2002), andthus covers all probiotics.

Probiotics are believed to exert their beneficial effects by competing withpathogens for mucosal binding sites and nutrients in the intestine, and by pro-ducing antimicrobial compounds and organic acids thereby inhibiting pathogengrowth (Vondruskova et al., 2010). A considerable number of studies have beencarried out in pigs to test the effect of different probiotics on parameters as intesti-nal microbiota composition, immunological components, occurrence of diarrhoeaand growth performance, but results have been ambiguous, probably as a result ofdifferences in the studied organism, strain, dose, experimental setup and duration,environment, husbandry, and genotypes (Kenny et al., 2011; Vondruskova et al.,2010). Thus, general recommendations on the use of probiotics are so far lacking.

The most studied probiotic bacteria in pigs are Streptococcus thermophilus,Bifidobacterium animalis subsp. lactis and B. thermophilus, Enterococcus fae-cium, Lactobacillus bulgaricus, L. fermentum, L. murinus, L. acidophilus, L. pen-tosus, L. casai, L. paracasai, L. plantarum, L. salivarius and L. reuteri, Pedio-coccus acidilactici and P. pentasaceus, Bacillus subtilis, B. cereus var. Toyoi andB. licheniformis, Escherichia coli Nissle, and Propionibacterium freudenreichii.Saccharomyces cerevisiae and Aspergillus oryzae are examples of probiotic fungi.Three of the most commonly used probiotics belong to the Lactobacillus, Entero-coccus and Bacillus genera. Some of the studies focusing on the gut microbiota aredescribed in the next paragraphs. Table 1.3 summarises information on period ofprobiotic administration, type of probiotic(s), dose(s), sample type(s) and effectsfrom the cited and additional relevant papers.

B. cereus var. Toyoi has been associated with a decreased number of enterotox-igenic E. coli (ETEC) and morbidity in weaned piglets (Papatsiros et al., 2011). In


a study by Scharek et al. (2007a), B. cereus var. Toyoi was fed to sows during ges-tation and to piglets from 15 days of age, and showed a decrease in the number ofpathogen-associated E. coli serogroups. In a similar study, feeding sows, and laterpiglets, B. cereus var. Toyoi resulted in a decrease in the number of pigs showingsigns of diarrhoea (Taras et al., 2005). Feeding E. faecium to piglets 1-14 days ofage decreased the number of E. coli in faeces, decreased pH in the duodenum, andincreased the concentration of lactic and propionic acid in the colon (Strompfovaet al., 2006). Scharek et al. (2005) found that feeding E. faecium to sows duringgestation and to piglets from 14 days of age resulted in a reduced frequency ofb-hemolytic E. coli and E. coli O:141. Feeding weaned piglets L. acidophilus hasalso been shown to reduce the coliform numbers, but also the diversity and richnessof the gut microbiota (Wang et al., 2012). Also B. pumilus has been reported todecrease the ileal number of E. coli after daily administration for 3 weeks in newlyweaned piglets (Prieto et al., 2014). The immunological effects are seen from table1.3.

Probiotic administration in periods of antibiotic therapy, as a way of diminish-ing unwanted side effects (e.g. antibiotic-associated diarrhoea) has been investi-gated in humans and mice, and has been shown to have beneficial effects (Cresciet al., 2013; Pirker et al., 2013). Administration of L. casei concomitantly withantibiotics resulted in a lower reduction in gut microbial diversity compared withsubjects treated with antibiotics alone. Moreover, the incidence of developingantibiotic-associated diarrhoea decreased from 18.6 % to 5 % when L. casei andantibiotics were given together (Pirker et al., 2013). A mouse study by Cresciet al. (2013) found that the negative effects of tributyrin on a butyrate transporterand receptor, ion exchangers, and water channels were reduced when mice weresimultaneously fed L. GG. A study by Plummer et al. (2005) reported that, whenpatients were administered strains of L. acidophilus and Bifidobacterium spp. con-comitantly with antibiotics, there was a significant decrease in the development ofantibiotic resistance among enterococci.

Hence, numerous studies have reported several probiotic bacteria to have health-promoting effects in pigs. However, the modes of actions are generally still un-known and, according to our knowledge, the consensus on which probiotics to usenon-existing.


Tabl

e1.

3:St

udie

son

the

use

ofpr

obio

tics

and

thei

reff

ect

onth

egu

tm

icro

biot

aan

din

test

inal

imm

une

para

met

ers

ofpi

glet

s.D

=di

gest

a;F=

faec

es;

M=

muc

osa;

T=

tiss

ue.

jeju

=je

junu

m;i

l=ile

um;c

ae=

caec

um;c

ol=

colo

n.[..]

=co

ncen

trat

ion.

ExP

EC

=ex

trai

ntes

tina

lpat

hoge

nic

E.coli

Age

and

peri

odof

prob

ioti

cad

min

istr

a-ti

on

Pro

biot

ic(s

)D

ose

Sam

ple(

s)E

ffect

sR

efer

ence

Sow

s:28

days

pre-

part

umto

wea

ning

.P

igle

ts:

14-3

9da

ysof

age.

Bacillus

cereus

var.

Toy

oiSo

ws:

3.5x

108

cfu/

gfe

ed.

Pig

lets

:1.

5x10

9cf

u/g

feed

.T

(jej

u)"

Per

mea

bilit

yof

inte

stin

alti

ssue

.#

CD

45+

toen

tero

cyte

rati

o.A

ltm

eyer

etal

.(20

14)

Sow

s:28

days

pre-

part

umto

wea

ning

.P

igle

ts:

12-3

4da

ysof

age.

Enterococcus

faecium

NC

IMB

1041

5So

ws:

4.2-

4.3x

106

cfu/

gfe

ed.

Pig

lets

:5.

1x10

6cf

u/g

(pre

-st

arte

r)an

d3.

6x10

6cf

u/g

(sta

rter

).

D+

M(c

ol)

F#

Num

ber

ofis

olat

esw

ith

ExP

EC

-typ

ical

viru

lenc

ege

nes

inco

lon

asce

nden

sB

edno

rzet

al.(

2013

)

Sow

s:28

days

pre-

part

umto

wea

ning

.P

igle

ts:

2-28

days

ofag

e.

Pediococcus

acidilactici

(PA

)an

d/or

Saccharo-

myces

cerevisiae

subs

p.boulardii

(SC

)

Sow

s:2.

5x10

9-

6x10

9cf

upe

rda

y.P

igle

ts:

1x10

9cf

upe

rda

y(s

uckl

ing)

and

2x10

9cf

u/kg

feed

(wea

ned)

.

F D(il+

col)

PA#

ileal

mic

robi

ota

dive

rsity.

PApr

omot

edes

tabl

ishm

ent

ofFir

mic

utes

,an

dSC

Porphyrom

onadaceae

and

Ru-

minococcaceae

inth

eco

lon.

Bro

usse

auet

al.(

2015

)

Sow

s:7

days

pre-

part

umun

tilb

irth

Bifidobacterium

lactis

BI-07,

Lactobacillus

acido-

philus

WN

0074

and

NC

FM

1.0x

1010

cfu/

day

ofea

chpr

obio

tic

D+

FP

robi

otic

sid

enti

fied

indi

gest

aan

dfa

eces

from

the

offsp

ring

.N

umbe

rof

prob

ioti

csde

crea

sed

wit

hti

me.

Bud

ding

ton

etal

.(20

10)

Pre

-ter

mpi

glet

s(9

0%

gest

atio

n).

5da

ys.

Lactobacillusparacasei,

Bifidobacterium

ani-

malis,

Streptococcus

therm

o-

philus

3.0x

109

cfu/

pig.

Tes

ted

both

live

and

dead

pro-

biot

ics.

D+

T"

NE

Cin

cide

nce.

#M

icro

bial

dive

rsity

(liv

e).

#SC

FAco

ncen

trat

ion

inco

lon

(liv

e).

"E

xpre

ssio

nsof

imm

une

med

iato

rs.

Cili

ebor

get

al.(

2011

)

Continues

on

the

next

page

22 Chapter 1. BackgroundTa

ble

1.3:

Stud

ies

onth

eus

eof

prob

ioti

csan

dth

eir

effec

ton

the

gut

mic

robi

ota

and

inte

stin

alim

mun

epa

ram

eter

sof

pigl

ets.

D=

dige

sta;

F=fa

eces

;M

=m

ucos

a;T

=ti

ssue

.je

ju=

jeju

num

;il=

ileum

;cae

=ca

ecum

;col

=co

lon.

[..]

=co

ncen

trat

ion.

ExP

EC

=ex

trai

ntes

tina

lpat

hoge

nic

E.coli

Age

and

peri

odof

prob

ioti

cad

min

istr

a-ti

on

Pro

biot

ic(s

)D

ose

Sam

ple(

s)E

ffect

sR

efer

ence

Sow

s:28

days

pre-

part

umto

21da

yspo

st-p

artu

m.

Pig

lets

:1-

31da

ysof

age.

Pediococcus

acidilactici

MA

18/5

M,

Saccharom

yces

cere-

visiae

subs

p.boulardii.

Alo

neor

com

bine

d.

Sow

s:2.

5-6.

9x10

9cf

u.P

igle

ts:

1.0-

2.0x

109

cfu.

ET

EC

F4

chal

leng

eda

y25

T(i

l)

Com

bine

d:"

Exp

ress

ion

ofIL

-6.

Sing

lepr

obio

tic:

#E

TE

CF4

atta

chm

ent.

Dau

delin

etal

.(20

11)

35-4

0da

ysol

d.4

wee

ks.

Bacillus

subtilis

MA

139

2.2x

105

cfu/

gfe

ed(l

ow)

2.2x

106

cfu/

gfe

ed(m

ediu

m)

2.2x

107

cfu/

gfe

ed(h

igh)

F"

Num

ber

ofla

ctob

acill

ion

day

28ir

resp

ecti

veof

prob

ioti

cdo

seG

uoet

al.(

2006

)

Wea

ned

pigl

ets

17-1

8da

ysof

age.

17da

ys.

Escherichia

coli

UM

-2an

d-7

3.1x

109

cfu/

ml;

50m

l/pi

g/da

yD

(il+

col)

"M

icro

bial

dive

rsity

inile

uman

dco

lon.

#E.coli

K88

num

bers

inile

uman

dco

lon.

Kra

use

etal

.(20

10)

Wea

ned

pigl

ets

26-2

7da

ysof

age.

4w

eeks

.

Enterococcus

faecium

,Lactobacillus

salivar-

ius,

Lactobacillus

reuteri,

Bifidobacterium

ther-

mophilus

1.0x

109

cfu/

kgfe

edD

(il+

col)

Ileu

m:

"Tot

alae

robe

san

den

tero

cocc

i.C

olon

:"

Gra

m-n

egat

ive

anae

robe

s,la

ctob

acill

iand

ente

roco

cci.

"[a

ceti

cac

id].

"Lac

toba

cilli

:ent

erob

acte

ria

ratio.

Mai

ret

al.(

2010

)

Wea

ned

pigl

ets

28da

ysof

age.

5w

eeks

.

Bacillus

cereus

var.

Toy

oisp

ores

1.0x

109

spor

es/g

feed

F#

Dia

rrho

easc

ore.

#N

umbe

rof

ET

EC

.Pap

atsi

ros

etal

.(20

11)

Continues

on

the

next

page


Tabl

e1.

3:St

udie

son

the

use

ofpr

obio

tics

and

thei

reff

ect

onth

egu

tm

icro

biot

aan

din

test

inal

imm

une

para

met

ers

ofpi

glet

s.D

=di

gest

a;F=

faec

es;

M=

muc

osa;

T=

tiss

ue.

jeju

=je

junu

m;i

l=ile

um;c

ae=

caec

um;c

ol=

colo

n.[..]

=co

ncen

trat

ion.

ExP

EC

=ex

trai

ntes

tina

lpat

hoge

nic

E.coli

Age

and

peri

odof

prob

ioti

cad

min

istr

a-ti

on

Pro

biot

ic(s

)D

ose

Sam

ple(

s)E

ffect

sR

efer

ence

1-21

days

afte

rw

eani

ngBacillus

Pum

ilus

5.0x

1010

spor

espe

rpi

g(o

rala

dmin

istr

atio

n)an

d>

1010

spor

espe

rpi

g(t

opdr

ess)

F D(il+

cae)

#N

umbe

rsof

E.coli

inile

um.

"Ilea

l[pr

opio

nic

acid

].P

riet

oet

al.(

2014

)

26da

ysol

d.4

wee

ks.

Enterococcus

faecium

,Lactobacillus

salivar-

ius,

Lactobacillus

reuteri,

Bifidobacterium

ther-

mophilum

1x10

9cf

u/kg

feed

D(i

l+ca

e+co

l)"

Mic

robi

alri

chne

ssin

ileum

."

Rel

ativ

eab

unda

nce

ofEnterococcaceae

inile

um.

Satt

ler

etal

.(20

15)

Sow

s:25

days

afte

rin

sem

inat

ion

unti

lw

eani

ng.

Pig

lets

:15

-56

days

ofag

e.

Bacillus

cereus

var.

Toy

oiSo

ws:

2.6-

4.0x

105

cfu/

gfe

ed.

Pig

lets

:1.

3-1.

4x10

6cf

u/g

feed

.T

(jej

u)Pey

er’s

Pat

ches

"C

D3/

CD

8nu

mbe

ron

day

28in

the

ep-

ithe

liall

ayer

."

Num

ber

ofC

D8+

lym

phoc

ytes

day

35.

"N

umbe

rof

CD

25+

andgdT

cells

inla

m-

ina

prop

ria.

#N

umbe

rof

part

hoge

n-as

soci

ated

E.

coli

sero

type

s.

Scha

rek

etal

.(20

07a)

Sow

s:25

days

afte

rin

sem

inat

ion

unti

lw

eani

ng.

Pig

lets

:1-

56da

ysof

age.

Bacillus

cereus

var.

Toy

oiEnterococcus

faecium

B.cereus

Sow

s:2.

6-4.

0x10

5cf

u/g

feed

.P

igle

ts:

1.3-

1.4x

106

cfu/

gfe

ed.

E.faecium

Sow

s:1.

6-1.

2x10

6cf

u/g

feed

.P

igle

ts:

1.7-

2.0x

105

cfu/

gfe

ed.

F Seru

m"

Faec

alIg

Ain

sow

san

dpi

glet

s(d

ay28

)(B

.cereus).

#Se

rum

IgG

inpi

glet

sda

y42

-56

(bot

hpr

o-bi

otic

s).

Scha

rek

etal

.(20

07b)

Continues

on

the

next

page

24 Chapter 1. BackgroundTa

ble

1.3:

Stud

ies

onth

eus

eof

prob

ioti

csan

dth

eir

effec

ton

the

gut

mic

robi

ota

and

inte

stin

alim

mun

epa

ram

eter

sof

pigl

ets.

D=

dige

sta;

F=fa

eces

;M

=m

ucos

a;T

=ti

ssue

.je

ju=

jeju

num

;il=

ileum

;cae

=ca

ecum

;col

=co

lon.

[..]

=co

ncen

trat

ion.

ExP

EC

=ex

trai

ntes

tina

lpat

hoge

nic

E.coli

Age

and

peri

odof

prob

ioti

cad

min

istr

a-ti

on

Pro

biot

ic(s

)D

ose

Sam

ple(

s)E

ffect

sR

efer

ence

1-14

days

ofag

eEnterococcus

faecium

EK

132.

0x10

9cf

u/pi

glet

D+

FFa

eces

day

7:"

Num

bers

ofen

tero

cocc

i.#

Num

bers

ofE.coli.

Dig

esta

:"

Num

bers

ofen

tero

cocc

i."

[Lac

tic

and

prop

ioni

cac

id]i

nco

lon.

Stro

mpf

ova

etal

.(20

06)

Wea

ned

pigl

ets

18-2

1da

ysol

d.5

wee

ks.

Lactobacillus

acido-

philus

orPediococcus

acidilactici.

Del

iver

edin

ahi

ghm

oist

ure

ferm

ente

dm

aize

feed

.

Bef

ore

ferm

enta

tion

:1.

7x10

7cf

u/m

lD

(il+

co)

#M

icro

bial

rich

ness

and

dive

rsity.

#N

umbe

rof

colif

orm

sw

hen

fed

L.

aci-

dophilus

com

pare

dto

P.acidilactici.

Wan

get

al.(

2012

)

Wea

ned

pigl

ets

21-3

6da

ysof

age

Bacillus

licheniform

is

DSM

5749

,Bacillus

subtilis

DSM

5750

3.9x

108

cfu/

day

(low

dose

)7.

8x10

8cf

u/da

y(h

igh

dose

)T

(jej

u)A

mel

iora

ted

path

ophy

siol

ogic

alch

ange

sca

used

byE.coli

F4

infe

ctio

n.In

duce

dT

-reg

type

1ce

lls.

Zhou

etal

.(20

15)


1.5.4 Host genetics

Different pig breeds have different genetic make-up and hence differ in certainphenotypic traits. Such traits may be related to intestinal physiology. Yorkshire,Landrace and Duroc are widely used pig breeds in Denmark. Pajarillo et al.(2015) characterised the faecal microbiota of these three pig breeds at 15 weeksof age, and found breed to have a significant effect on the overall gut microbiota.This breed effect was still apparent after 12 weeks of housing the three breedstogether, though the dissimilarity between the microbial communities decreased.A similar effect of breed has been reported in the faecal microbiota of neonatalYorkshire and Meishan piglets (Bian et al., 2016). Interestingly, the breed effectwas significant only on day 14 and 49 of age and hence, not detectable immediatelyafter birth. At 14 days of age, Meishan piglets had the highest relative abundanceof Fusobacteriales and lowest abundance of Erysipelotrichaceae, while they on day49 had the highest abundance of Lactobacilllus. Despite the influential effect ofgenotype on the composition of the faecal microbiota, it appears that this effectis minor when compared to the effect of age, weaning, and environment as thesetend to even out the effects of genotype (Bian et al., 2016; Pajarillo et al., 2015).

1.5.5 Investigated factors in the current thesis

Bovine colostrum

Bovine colostrum (BC) is the first milk secreted by the dairy cow after partu-rition. A precise definition of colostrum is lacking as authors generally disagreeabout when the mammary secretion switch from colostrum to non-colostrum milk(McGrath et al., 2016).

The essential function of colostrum is to provide the neonate with nutrients, inaddition to growth-promoting and protective compounds. The latter is especiallyimportant in mammals with epitheliochorial and synepitheliochoral placentas, e.g.pigs and cows, which do not allow transfer of immunoglobulins in utero. Hence,ingestion of colostrum immediately after birth is pivotal for the survival of theseneonates (Pakkanen and Aalto, 1997).

BC is generally regarded as unsuited for human consumption and produced inhigher amounts than required by the calf. Hence, it is a by-product from the dairyindustry and thus available in excess amounts (McGrath et al., 2016). Colostrumhas a chemical composition that varies significantly from that of ’mature’ (non-colostrum) milk, and especially the high content of antimicrobial components and


growth factors has led to an increased interest in bovine colostrum as a nutraceu-tical during recent years (Pandey et al., 2011). In addition to these bio-activecomponents, BC is also constituted of notable amounts of numerous oligosaccha-rides (Gopal and Gill, 2000), which have been suggested to have growth-promotingeffects on probiotic bacteria (Champagne et al., 2014).

Major antimicrobial components in BC are immunoglobulins (Igs) (Pakkanenand Aalto, 1997) and non-specific components as lactoferrin and lactoperoxidase(van Hooijdonk et al., 2000). The major Igs are IgG, IgA, and IgM, with IgG(1 and 2) being the primary Ig in colostrum (Hurley and Theil, 2011). Ingestedimmunoglobulins provide the neonate with local passive immunity by binding tobacteria and toxins, hereby limiting their ability to bind to the enterocyte surface(Hurley and Theil, 2011). The antimicrobial mode of action of lactoferrin is largelyunknown, but is generally believed to be related to the iron-binding capacity of themolecule (van Hooijdonk et al., 2000). Lactoperoxidase is an enzymatic antioxidantinhibiting bacterial metabolism (Pandey et al., 2011).

Growth factors are important to the growth and development of the newbornyoung. Numerous growth factors are found in BC, with the most abundant onesbeing Insulin-like Growth Factor 1 and 2. Then follows growth factors as Epider-mal Growth Factor and Transforming Growth Factor-b. These growth factors areimportant in stimulating cellular growth and cell differentiation (Boudry et al.,2008).

The beneficial effects of BC have been investigated in both pre-term, new-born, and newly weaned piglets (see table 1.4). Several studies suggest that BCexerts its function by inhibiting/limiting intestinal inflammatory processes andpathogen-induced cellular destruction. However, the conglomerate of studies hasprovided varying results. Discrepancies between studies are likely due to differ-ences in doses administered, health status of the target group, period and lengthof administration, and end-points. Also, the chemical composition of BC is influ-enced by processing (Elfstrand et al., 2002), cow breed, feeding, parity, length ofdry period, and time of collection (Gopal and Gill, 2000), hence different productsmay result in varying biological effects.


Table 1.4: Reported effects of bovine colostrum. BC = Bovine Colostrum; NEC = NecrotisingEnterocolitis; PP = Peyer’s Patches. IFN = Interferon. IL = Interleukin

Experimental set-up Effect(s) Reference

Newly weaned piglets

BC orally administered.Fed for three weeks.

No effect on feed intake or growth." serum IgA.Stimulated ileal PP.PP: # IFN-g and " IL-2, IL-4 and IL-10expression.Lymph nodes: " IL-2, IL-10 and IL-12expressionIntestinal tissue: " IL-12 expression.

Boudry et al. (2007)

BC supplemented diet.Fed for four weeks.

Improved weight gain (highest effectduring week 1)." Systemic IgA during week 1.

Boudry et al. (2008)

BC supplemented diet.Fed for 7 or 14 days.

" Duodenal villi perimeter. Huguet et al. (2007)

BC supplemented diet.Fed for 12 days post-weaning.

" Feed intake and daily weight gain.Improved faecal consistency.Effects lasted beyond the study period.

Huguet et al. (2012)

BC supplemented diet.Challenged with E. coli K88.

No beneficial effect on intestinalmorphology reported.

King et al. (2007)

BC supplemented diet.Fed for 7 days.

No significant effect on weight gain. King et al. (2008)

Fed solely BC for eight days.Relative to milk formula.

Fewer E. coli colonising jejunal and ilealtissue.Jejunum: # expression of TLR.Ileum: # expression of TLR and IL-2

Sugiharto et al. (2015)

Neonatal piglets

BC orally administered.Fed from 1-10 days of age.

" Lifespan of low birth-weight piglets.No effect on body weight or weight gain.

Viehmann et al. (2015)

Pre-term born piglets piglets

BC orally administered.Relative to milk formula.

# NEC incidence and severity." Lactose absorption.# Intestinal permeability.

Shen et al. (2015)

BC orally administered.Relative to milk formula.

# NEC severity." Small intestinal enzyme activities e.g.lactase.

Stoy et al. (2014)

In vitro

IPEC-J2 cells.Stimulated with heat-treatedE. coli and Salmonella entericaserovar Typhimurium.

# Expression of genes related to immune,defence and inflammatory responses." Expression of genes related to proliferation,wounding and migration processes.# Induction of NF-kB transcription factor.

Blais et al. (2015)

Bacterial cultures Stimulated probiotic bacteria.Inhibited growth of E. coli.

Champagne et al. (2014)


Bacillus spp. spores

Generally, there exists no consensus on which probiotic is considered the best, orwhen and how it should be administered. Therefore, due to the list of studiesreporting beneficial effects of Bacillus spp., and its availability, a total of nineBacillus strains belonging to B. subtilis, B. licheniformis and B. amyloliquefaecenswere included in the animal experiment presented in this thesis.

Bacillus spp. is a spore-forming aerobic/facultative anaerobic bacterial genus,which forms endospores when the surrounding environmental conditions are un-favourable to the continued survival of the vegetative cell (Cutting, 2011). Theproduced spores are in a state of dormancy, and their physical structure makesthem highly resistant to otherwise intolerant pH values, temperatures, irradiation,and desiccation (Nicholson et al., 2000). Figure 1.4 depicts the schematic structureof an endospore. These characteristics make Bacillus spp. able to survive pelletingprocesses, and hence ideal as an in-feed probiotic. Bacillus species are primarilyfound in soil and are not normally regarded as inhabitants of the pig gut micro-biota (Tam et al., 2006). Studies have shown that B. subtilis and B. licheniformisare able to germinate and re-sporulate in the pig gut, but that their ability to growis limited. Also, when spores are no longer administered, they disappear from themicrobial community, indicating that they are not able to colonise the gut (Leseret al., 2008). Studies suggest that both spores and vegetative cells have probioticpotentials (Tam et al., 2006).

Core

Inner membrane

Cortex

Coat

Thick peptidoglycan layer

Lamellar proteinaceous structure

Plasma membrane of the vegetative cell

Dehydrated cytoplasm, stabilised DNA and RNA. Ions in complexes.

Figure 1.4: Illustration of the basic structure of a Bacillus endospore. Adapted from Venemaand Do Carmo (2015)


Gentamicin

In Denmark, neonatal piglets with diarrhoea are commonly administered gentam-icin, an antibiotic sold as a watery formulation intended for oral administration.Gentamicin is a bactericidal concentration-dependant antibiotic acting primarilyagainst gram-negative aerobes by irreversible binding to the 30S ribosomal sub-unit, thereby inhibiting bacterial protein synthesis (Plumb, 2011). The effectsof gentamicin on the gut microbiota of piglets are, according to our knowledge,unknown.

Alpha-(1,2)-fucosyltransferase

Escherichia coli is a heterogeneous bacterial group. It is a gram-negative rod nor-mally inhabiting the gut, but is also found in several pathotypes causing intestinaland extra-intestinal disease (Croxen and Finlay, 2010). ETEC is an importantintestinal pathotype in piglets, where it causes intestinal disease (enteric colibacil-losis) in both newborn and newly-weaned piglets (Fairbrother et al., 2005; Nagyand Fekete, 1999). To be able to cause disease, ETEC is dependant on two majorvirulence factors; surface adhesins and enterotoxins. Adhesins are mainly consti-tuted of fimbriae. Fimbriae are antigenic proteins protruding from the bacterial cellwall and come in numerous variants. In pigs, especially the F4 and F18 fimbriaehave been studied intensively. Bacterial colonisation is dependent on the presenceof fimbriae-specific receptors on the small intestinal microvilli (Nagy and Fekete,2005). F4 receptors are expressed in the intestinal tissue from birth, while F18receptors are not fully expressed in piglets younger than three weeks of age (Fair-brother et al., 2005). ETEC produces heat-stable and heat-labile enterotoxins,and both are produced by ETECs able to infect piglets (Nagy and Fekete, 2005).The enterotoxins increase the cellular electrolyte and fluid secretion (Croxen andFinlay, 2010) leading to hyper-secretory diarrhoea. The expression of both F4 andF18 receptors are genetically inherited in an autosomal dominant manner, andpigs resistant toward F4 and F18 have been bred.

The alpha-(1,2)-fucosyltransferase (FUT1 ) gene product is involved in the for-mation of blood-group antigen sugar structures on cell membranes, and is ex-pressed in the small intestine of pigs (Bao et al., 2012; Frydendahl et al., 2003).Due to its chromosomal location in the ETEC F18 receptor region, the FUT1 genehas been proposed as a candidate gene for manipulating the adhesion of ETECF18 to the specific intestinal receptor (Meijerink et al., 1997). Frydendahl et al.(2003) showed that a guanine to adenine mutation at nucleotide position 307 (chro-


mosome 6), resulting in an alanine to threonine substitution at position 103, inboth loci of the FUT1 gene (FUT1-307A/A), significantly decreased the suscepti-bility of weanling piglets to ETEC F18 infection when compared to piglets with theFUT1-307A/G and FUT1-307G/G genotypes. The FUT1-307A/A genotype has beenassociated with a decreased FUT1 enzyme activity (Meijerink et al., 2000). Theeffect of FUT1 polymorphisms on colonisation of commensal bacteria is currentlyunknown.

1.6 Methods for studying the gut microbiota

1.6.1 Culture-based methods

Microbial culture is generally regarded as the traditional method for studyingmicroorganisms. Using various growth media and fulfilling specific requirementsto substrates, pH, incubation temperatures and oxygen tension, bacteria can begrown, allowing their quantification and further biochemical, physiological andtaxonomical characterisation (Hoiby, 2008). The use of culture-based methods forstudying the pig gut microbiota dates back to the 1950’s, where focus was only on afew bacterial groups (Fewins et al., 1967). As more bacteria were isolated from thegut, researchers started realising the limitations of using selective growth mediaand growth conditions in characterising a microbial community as diverse as theone found in the pig gut (Allison et al., 1979; Salanitro et al., 1977). A study byPryde et al. (1999) showed that, compared to a molecular identification technique(using the 16S ribosomal DNA gene), bacterial culture was insufficient in describingthe microbial diversity of the gut. It was later suggested that approximately50 % of all bacterial species found in the human gut were yet to be cultured(Sears, 2005). Despite a change of focus towards newer molecular techniques,microbial culture should still be considered an invaluable tool in quantificationand functional characterisation of specific bacterial groups, as stressed by recentpublications on culturomics, a technique combining diverse culture conditions andmass spectrometry (Lagier et al., 2015).

1.6.2 Next-generation sequencing

During recent years, several molecular techniques taking advantage of the 16SrRNA gene have been employed in the investigation of the phylogenic diversity ofthe pig gut microbiota, these being; cloning and sequencing (Leser et al., 2002;


Pryde et al., 1999); Terminal Restriction Fragment Length Polymorphism (Ho-jberg et al., 2005; Liu et al., 2012); and Denaturing Gradient Gel Electrophoresis(Levesque et al., 2012; Liu et al., 2014a). However, due to the rapidly developingsequencing technologies and the appreciation of their potential in characterisingcomplex communities, focus has changed to the high-throughput next-generationsequencing (NGS) technologies. Currently, Illumina (HiSeq and MiSeq) (Kim andIsaacson, 2015; Slifierz et al., 2015) and 454-pyrosequencing (Hermann-Bank et al.,2015; Mach et al., 2015; Mann et al., 2014) are the dominating NGS technologiesin current literature investigating the pig gut microbiota.

The 16S ribosomal RNA (rRNA) gene has become the preferred gene for phy-logenetic classification of bacteria. It is found in all prokaryotes and made up bynine hypervariable (V1-V9) regions interspersed by highly conserved regions. Asclosely related bacterial species have more similar sequences in their hypervariableregions, than species more distantly related, these regions are used in the phyloge-netic classification (Kim et al., 2015). The conserved regions act as annealing sitesfor universal primers, allowing amplification of one or several of the hypervariableregions across species (Nocker et al., 2007). Before proceeding to sequencing, anamplicon library of a specific region of the 16S rRNA gene is produced by PCR(Kim and Isaacson, 2015). As current technologies do not allow sequencing of thefull 16S rRNA gene, only up to three variable regions are used as representativesof the full length. There exists no general agreement on which regions are the bestwhen studying the microbial community in the pig gut, but the most commonlyused regions are V1 to V6 (reviewed by Kim and Isaacson (2015)). Preparation ofsamples for sequencing, and the following data analyses, often vary considerablybetween studies, and hence make it difficult to directly compare results betweenstudies. These issues are further addressed in the general discussion.

Sequencing platforms differ not only in their sequencing technologies, but alsoin the number and length of reads produced in a single run. Generally, we wishto sequence as deep as possible. Sequencing depth is associated with the numberof reads produced, and should preferably be as high as possible. In addition, tobe able to assign taxonomy with high precision, sequencing reads should be aslong as possible. Hence, key features of a sequencing platform are the number andlength of reads it is able to produce at low error rates. This was demonstratedby Claesson et al. (2010), who reported the insufficiency of short reads with higherror rates in characterising a human faecal microbial community. Hence, severalparameters, including potential biases, have to be taken into consideration when


utilising these new sequencing platforms. Table 1.5 compares key features betweenNGS and traditional culture. Challenges of bioinformatic processing of sequencingdata are further described in the methodological approach.

Table 1.5: Comparing high-throughput next-generation sequencing and traditional bacterial cul-ture. NGS = next-generation sequencing

NGS Culture

PCR amplification step Yes NoCoverage of complex communities High LowQuantitative No YesPrice Expensive (decreasing) Lower expenseData output Large amounts Small amountsSpecial requirements for data analysis Yes NoWorkload Small Large

Chapter 2. Objectives and hypotheses 33

Chapter 2

Objectives and hypotheses

2.1 Project aim and objectives

The overall aim of the presented Ph.D. project is to investigate if it is possible, byspecific dietary and probiotic (in relation to antibiotic therapy) means, to shapethe piglet gut microbiota in a direction that favours healthier animals. In addition,the gut microbial effects of a genetic variation in ETEC F18 susceptibility is inves-tigated to see if this has an unintentional effect on the colonising gut microbiota.

The project objective is to taxonomically and quantitatively characterise thecolonisation and succession of the gut microbiota of piglets from birth until twoweeks after weaning within four scenarios, each representing a dietary, probiotic,antibiotic, or genetic factor. This objective will be accomplished by using classi-cal microbial culture (quantitative approach) accompanied by culture-independentNGS 16S rRNA gene sequencing (in-depth qualitative approach), and detection ofmajor microbial fermentation products.

2.2 Experimental objectives and hypotheses

Animal experiment 1Objective: To characterise the colonisation of the gut microbial community in 23-30 days old piglets fed BC, a porcine milk replacer (MR) or sow milk.

Hypothesis: When compared to a commercial porcine MR, BC is a more suitablesubstitute to sow milk in maintaining a health-promoting gut microbiota. Further-more, the gut microbial community of piglets fed BC will have a closer resemblanceto that of piglets fed sow milk than will MR-fed piglets.

34 Chapter 2. Objectives and hypotheses

Animal experiment 2Objective: To characterise how FUT1 genotype influences the establishment andsuccession of the gut microbial community in 5 to 34 days old piglets.

Hypothesis: Piglets with the FUT1-M307A/A genotype will have a gut microbiotadifferent from piglets with the FUT1-M307A/G genotype.

Animal experiment 3Objective: To characterise the gut microbial effects of administrating gentamicinand Bacillus subtilis, B. licheniformis, and B. amyloliquefaecens spores in pigletsbetween 0 and 42 days of age.

Hypothesis: Administering gentamicin to newborn piglets will change the microbialcommunity of the gut. Concurrent administration of Bacillus spp. spores willalleviate the expected negative effect(s) of gentamicin.

Chapter 3. Methodological approach 35

Chapter 3

Methodological approach

To obtain a thorough knowledge on the gut microbial diversity, the following ap-proaches were used for sample collection and community analyses.

3.1 Gastrointestinal samples

In the three animal studies, several sample types were collected from the lengthof the gut. In order to study the succession of the gut microbiota, sequentialfaecal samples were collected. Fresh faecal samples were collected directly fromthe rectum by gentle digital exploration. The advantage of this approach is that thesame animal can be sampled multiple times. As the microbial community differsbetween different gut segments (Looft et al., 2014a), luminal contents (digesta)were collected from several segments of the gut as well. Piglets were sacrificedfollowing a standard protocol using a captive bolt gun followed by bleeding, orby blunt trauma to the head (piglets < 5 kg). Immediately after sacrification,the stomach and intestines were located and excised. The stomach and caecumwere located and excised from the rest of the intestinal organs. The small andlarge intestine was removed from the intestinal mesenterium, measured in fulllength, and divided into two or three (dependent on piglet age) equally long parts.Stomach and intestinal digesta was immediately collected.

36 Chapter 3. Methodological approach

3.2 Characterising the gut bacterial community

3.2.1 Sequencing of the 16S rRNA gene

In-depth characterisation of the microbial community was achieved by ampliconsequencing of the 16S rRNA gene using high-throughput sequencing. A flowchartfor sample preparation and data analysis is seen in figure 3.1. Each of these steps,and different applied methods in each of these steps, potentially have an effect onthe final community diversity.

DNA extraction

Amplicon production by PCR

Clean-up of amplicon library

DNA quantification

Quality control

Quality control

Multiplex samples in equimolar

concentrations

Sequencing

Quality control

Step 1

Step 2

Step 3

Step 4

Step 5

Step 6

Quality filteringTrimming

Merge reads

OTU clustering

Chimera detection

Data visualisation

Assigning taxonomy

Figure 3.1: Flowchart on sample preparation for next-generation sequencing and data processing.OTU = Operational Taxonomical Unit

Sequencing of samples obtained from animal experiment 1 and 3 was doneusing the Illumina MiSeq (Illumina, USA). This platform provided 2x300 base-pairs (paired-end) reads and allowed us to use the V1-V3 regions. These regionshave been proposed to be the preferred representatives of the full-length 16S rRNAgene when analysing highly diverse communities as the gut microbiota (Kim et al.,2011). In animal experiment 2, the samples were sequenced on the Ion Torrent

Chapter 3. Methodological approach 37

PGM platform (Thermo Fisher Scientific, USA). However, as this platform onlyprovided 200 base-pairs reads, only the V3 region was sequenced. Using different16S rRNA regions potentially results in different microbial communities (Burbachet al., 2016), and the use of different approaches limits our ability to compare theresults of experiment 2 with those of experiment 1 and 3.

Aside from sequencing platform and the variable region(s) sequenced, DNAextraction procedure (Burbach et al., 2016; Clooney et al., 2016), PCR primers(Osborne et al., 2005), and DNA purity and concentration (Clooney et al., 2016)also influence the observed diversity. Bioinformatic processing of reads obtainedby high-throughput sequencing takes advantage of advanced computational al-gorithms. However, numerous pipelines using different algorithms (e.g. OTU-clustering algorithms) exist, which have a significant influence on the microbialdiversity (Schloss, 2010). Hence, the methodological approach has a significantimpact on the final community observed, and should be considered when analysingand interpreting sequencing data.

3.2.2 Microbial plate culture

Classical microbial enumeration by culture was included as a measure of absolutemicrobial numbers of selected microbial groups. The absolute numbers were givenas colony forming units (cfu) (log 10 transformed), and in order to obtain countablecolony numbers, a series of 10-fold dilutions were made from the digesta or faecalsample, followed by plating on different selective and non-selective media. Wechose to include six (seven in experiment 3) major microbial classes, which werecultured on the following media;

• Blood agar: Enumeration of haemolytic bacteria. Such colonies are gener-ally unwanted as they typically represent pathogenic bacteria, e.g. ETEC(Fairbrother et al., 2005). Further typing was not performed.

• Casein Soya Bean Digest agar: Enumeration of Bacillus spp. spores. Vege-tative cells were killed and spores activated by heat-treatment.

• Colon roll-tubes: Enumeration of total anaerobic bacteria in media contain-ing colon extract.

• MacConkey agar: Enumeration of bacteria belonging to the Enterobacteri-aceae family. Lactose fermentors grow as pink/red colonies (e.g. E. coli),

38 Chapter 3. Methodological approach

while non-lactose fermenters grow as white colonies (e.g. Salmonella andShigella).

• Malt Chloramphenicol (MCA) agar: Enumeration of yeasts and moulds.

• Man Rogosa and Sharpe (MRS) agar: Enumeration of lactic acid bacteria.Colonies were not further identified.

• Tryptose Sulfite Cycloserine (TSC) agar: Enumeration of C. perfringens.

Chapter 4. Manuscript 1 39

Chapter 4

Manuscript 1

The microbial community of the gut differs between piglets fed sowmilk, milk replacer or bovine colostrum.

Ann-Sofie Riis Poulsen, Nadieh de Jonge, Sugiharto Sugiharto, Jeppe Lund Nielsen,Charlotte Lauridsen, Nuria Canibe.

Submitted to British Journal of nutrition.

40 Chapter 4. Manuscript 1

Experimental set-up:

36 piglets from four sowsFollowed from 23 to 30 days of age

SM-fedPiglets fed sow milk

MR-fedPiglets fed milk replacer

BC-fedPiglets fed bovine

colostrum

Digesta Stomach, ileum, caecum and mid colon (day 30)

1. 16S rRNA gene sequencing• V1-V3 region• Illumina MiSeq

2. Classical culture3. Organic acid analysis

12 piglets 12 piglets12 piglets



Faeces Day 23, 25, 27 og 30



1

The microbial community of the gut differs between piglets fed sow milk, milk replacer or

bovine colostrum

Ann-Sofie R. Poulsen1*, Nadieh de Jonge2, Sugiharto Sugiharto3, Jeppe L. Nielsen2, Charlotte

Lauridsen1, Nuria Canibe1

1 Department of Animal Science, Faculty of Science and Technology, Aarhus University, Tjele,

Denmark

2 Center for Microbial Communities, Department of Chemistry and Bioscience, Aalborg University,

Aalborg, Denmark

3 Faculty of Animal and Agricultural Sciences, Diponegoro University, Semarang, Central Java,

Indonesia

* Corresponding author.

Aarhus University,

Faculty of Science and Technology, Department of Animal Science,

Blichers Allé 20, P.O. Box 50,

DK-8830 Tjele, Denmark

Tel.: +45 87 15 80 58

Fax.: +45 87 15 42 49

E-mail address: [email protected]

Short title: Bovine colostrum and gut microbiota.

Key words: Bovine colostrum. Gut microbiota. Undersized piglets. 16S rRNA gene sequencing.


2

Abstract

The aim of the current study was to characterise the gut microbiota of piglets fed bovine colostrum

(BC), milk replacer (MR) or sow milk (SM). Twenty-three days old piglets (n=36) were randomly

allocated to the three diets. Faecal samples were collected at 23, 25, 27 and 30 days of age. Piglets

were euthanized at 30 days of age and luminal content from the stomach, ileum, caecum and mid-

colon was collected. Bacterial DNA was subjected to amplicon sequencing of the 16S rRNA gene.

Bacterial enumerations by culture and short-chain fatty acid analysis were conducted as well. BC-

piglets had a high abundance of Lactococcus in the stomach (p<0.0001) and ileal (p<0.0001)

digesta, while SM-piglets had a high abundance of Lactobacillus in the stomach digesta (p<0.0001).

MR-piglets had a high abundance of Enterobacteriaceae in the ileal digesta (p<0.0001) and a

higher number of haemolytic bacteria in the ileal (p=0.0002) and mid-colon (p=0.001) digesta than

SM-piglets. BC-piglets showed the highest colonic concentration of iso-butyric and iso-valeric acid

(p=0.02). Both sequencing and culture indicated higher shedding of potential pathogenic

Escherichia coli in MR-piglets, while the gut microbiota of BC-piglets was characterised by a

change in lactic acid bacteria genera when compared to SM-piglets. We conclude that especially the

ileal microbiota of BC-piglets had a closer resemblance to that of SM-piglets in regard to the

abundance of potential enteric pathogens than did MR-piglets. The higher content of colonic

branched-chain fatty acids in BC-piglets, however, calls for further investigation.


3

Introduction

The increased litter sizes of the modern pig industry have negatively influenced piglet viability.

Large litters come with great variation in within-litter birth weights and are accompanied by an

increased number of low birth-weight piglets. Undersized piglets have difficulties competing with

heavier littermates, and experience reduced growth rates and increased morbidity and mortality (1).

To ensure adequate feed intake of undersized piglets, and hence improve weight gain and survival

chances, milk replacers have been implemented by many pig farmers (2). However, the

gastrointestinal tract of undersized piglets may be immature, influencing optimal nutrient digestion

and absorption. Also, as sow colostrum quality (and quantity) influences short- and long-term

survival (reviewed by Theil et al. (3)), there is an increasing interest in the nutritional and functional

quality of milk replacers and milk supplements for newborn piglets.

Diet is known to influence gut maturation (4). Besides maturing functional entities of the

intestinal tissues, gut maturation also includes the establishment of a gastrointestinal microbiota (5).

The gastrointestinal microbiota is appreciated as an important inhabitant of the body, being highly

involved in host defensive mechanisms. The local defensive mechanisms of the gut microbiota

include competing with pathogens for mucosal binding sites and nutrients, production of

antimicrobial-like agents and elimination of noxious substances (6; 7). Furthermore, microbial

colonisation of the gut stimulates local immune cell proliferation, hence playing a key role in the

maturation of the adaptive immune system. As the gut microbiota influences animal health,

choosing a diet in favor of a beneficial microbiota may be crucial. This could especially be

important at the time of microbiota establishment after birth, as the very first bacteria to colonise

the gut are able to influence the proceeding colonisation, and hence the composition of the 'adult'

gut microbiota (8).

Colostrum is the milk secreted during the first 24-48 hours following parturition (3; 9).

Bovine colostrum (BC) is a by-product from the dairy industry, available in excess amounts, and a

rich source of biologically active compounds (10; 11). The major bio-active compounds are growth

factors such as Insulin-like Growth Factor, Epidermal Growth Factor and Tissue Growth Factor β,

and antimicrobials as immunoglobulins (12). In preterm born piglets, BC has been reported to

increase the activity of specific brush-border enzymes, improve intestinal health and decrease

severity of necrotising enterocolitis when compared to feeding milk formula (13). In term born

piglets, provision of BC rather than milk replacer reduced the intestinal colonisation of

enterotoxigenic Escherichia coli and modulated the intestinal immune system, while no difference

between BC and natural rearing with the sow was observed (14). Furthermore, newly weaned piglets

have shown improved growth performance (15) and intestinal mucosal restoration (16) when

supplemented with BC.


4

The effect of BC on the gut microbiota remains to be explored. The aim of the current study

was therefore to taxonomically and quantitatively characterise the early colonisation of the

gastrointestinal microbial community in piglets fed sow milk, milk replacer (originating from

bovine milk) or BC. We hypothesised that the gastrointestinal microbial community of piglets fed

BC would have closer resemblance to that of piglets fed sow milk than would piglets fed milk

replacer.

Materials and methods

Study design

The present study was conducted according to the ethical license obtained from the Danish Animal

Experiments Inspectorate, Ministry of Food, Agriculture and Fisheries, Danish Veterinary and Food

administration. National guidelines on experimental animal housing, care and sacrificing procedure

were followed. The study was performed at the experimental facility at the Department of Animal

Science (Foulum, Aarhus University).

A total of 36 piglets (crossbred Danish Landrace x Yorkshire; mixed females and males)

from four different sows were included in the study. The sows originated from the herd at Foulum,

Aarhus University. Piglets were housed with their dams until 23 days of age. The 36 piglets (nine

piglets from each sow) were randomly assigned to one of the following treatment groups (three

piglets from each sow per treatment): (a) kept with the sow for the whole experimental period (SM-

fed); (b) separated from the sow and fed a commercial porcine milk replacer powder (Grifor,

Hatting KS) (MR-fed); (c) separated from the sow and fed powdered BC (European Colostrum

Industry S.A.) (BC-fed). Chemical compositions are shown in Table 1.

Piglets separated from the sow were transported to an experimental stable and housed in

pens (1.45 m x 1.70 m) in groups of three until 30 days of age (end of experiment). Piglets were

randomly allocated to the dietary treatments. All pens were padded with rubber mats and piglets

had access to rooting material. Stable temperature was 24˚C. Piglets were continuously exposed to

faecal material from the sow by daily collecting faecal matter from the dam pen and spreading it

into the piglet pens. Each pen was equipped with an automated wet feeder (Mambo Automix 25;

Wit-Mambo Inc.) from which the piglets received ad libitum feeding. Piglets had access to fresh

water. The powdered BC and MR were dissolved in approximately 45˚C warm water in the

automated feeder (approximate dry matter percentage: BC 20% and MR 15%). To get the piglets

accustomed to the feeding machine, they were fed one portion of sow milk in the trough of the

machine upon arrival to the pen. SM-fed piglets suckled their dams until 30 days of age (end of

experiment). In an attempt to minimize the impact of other factors than the planned dietary

intervention, SM-fed piglets were transported exactly as the BC-fed and MR-fed piglets before


5

returning to the sow. This ensured that all piglets were subjected to similar amount of stress due to

transportation.

Sample and data collection

Piglets were individually weighed at 23 and 30 days of age. BC and MR consumption was recorded

daily as powder provided minus leftovers from the automated wet feeder (determined by freeze

drying). The clinical condition of the piglets was evaluated daily, including occurrence of diarrhoea.

Faecal samples were collected from the rectum on day 23 (before transportation), 25, 27 and

30. All piglets were euthanised at 30 days of age; the abdomen was incised and the gastrointestinal

tract removed. Total luminal contents (digesta) from the stomach, small intestine (proximal and

distal), cecum and colon (proximal, mid and distal) were collected immediately after killing.

Digesta from the respective segments and faeces was mixed and subsamples were stored at -20˚C

for organic acid analysis (stomach, distal small intestine, caecum and mid colon) and snap-frozen in

liquid nitrogen and stored at -80˚C for 16S rRNA gene amplicon sequencing (stomach, distal small

intestine and mid colon). Bacterial enumeration by culture was performed on a fresh subsample of

faeces and digesta (stomach, distal small intestine, caecum and mid colon).

Dry matter and organic acid analysis

Dry matter content of digesta was determined by freeze-drying using a ScanVac Coolsafe 55

(Labogene ApS, Lynge, Denmark). Concentrations of the short-chain fatty acids (SCFAs) acetic,

propionic, butyric, isobutyric, valeric and isovaleric acid, and lactic acid in faeces and digesta were

quantified as previously described by Canibe et al. (17).

Microbiological enumerations

Approximately 1 g faecal material was suspended in 10 ml pre-reduced salt medium(18). The content

was homogenized in a Smasher paddle blender (bioMérieux Industry, USA) for 2 min.

Approximately 5 g digesta were suspended in a flask containing 50 ml pre-reduced salt medium.

The flask content was transferred to a CO2 flushed bag and homogenized for 2 minutes. 1 ml

homogenate was transferred to a Hungate tube containing 9 ml pre-reduced salt medium and 10-

fold dilutions were prepared using the technique previously described by Miller and Wolin (19). The

samples were plated on selective (and indicative) and non-selective agar plates.

Enterobacteriaceae were enumerated on MacConkey agar (Merck 1.05465) after aerobic

incubation for one day. Yeasts were enumerated on malt, chlortetracycline and chloramphenicol

agar (Merck 1.03753 (yeast extract), 1.05397 (malt extract), 1.07224 (bacto-pepton), 1.08337

(glucose), 1.01614 (agar-agar) and Oxoid Sr0177E) after aerobic incubation for two days.


6

Haemolytic bacteria were enumerated on blood agar (Oxoid Pb5039A) after aerobic incubation for

one day. Clostridium perfringens were enumerated using the pour-plate technique on tryptose sulfit

cycloserine agar (Merck 1.11972, 1.00888) after anaerobic incubation for one day. Lactic acid

bacteria were enumerated on de Man, Rogosa and Sharp agar (Merck 1.10660) after anaerobic

incubation for two days. Total anaerobic bacteria were enumerated in roll tubes containing pig

colon fluid-glucose-cellobiose agar (18) and incubated for seven days. Plates and roll-tubes were

incubated at 37˚C.

DNA extraction

Samples for DNA extraction included 47 faecal samples (one sample was missing from the SM-fed

group on day 25) from 12 piglets and 72 digesta (stomach, distal small intestine, mid colon)

samples from 24 piglets. DNA was extracted with the E.Z.N.A. Stool DNA Kit (Omega Bio-Tek,

inc., VWR international) following a standard protocol with the following exception; bead beating

was performed on a FastPrep FP120 (Bio 101 Savant/MP Biomedicals, USA) for 2x30 s. DNA

extract purity was evaluated with Nanodrop ND1000 (Thermo Scientific, USA) and quantified

fluorometrically with Qubit 3.0 HS dsDNA assay (Life Technologies, Thermo Fisher Scientific,

USA). DNA concentrations were normalized to 5 ng/µl.

16S rRNA gene amplicon sequencing

Amplicon libraries were generated by targeted amplification of the V1-V3 hypervariable regions of

the bacterial 16S rRNA gene. The PCR reaction (25 µl) contained 10 ng template DNA, Platinum®

High Fidelity buffer (x1), dNTP (400 uM of each), MgSO4 (1.5 mM) and Platinum® Taq DNA

polymerase High Fidelity (1U) and barcoded library adapters (400 nM). V1-V3 primers: 27F

AGAGTTTGATCCTGGCTCAG and 534R ATTACCGCGGCTGCTGG. Thermocycler settings:

Initial denaturation at 95˚C for 2 min, 30 cycles of 95˚C for 20 s, 56˚C for 30 s, 72˚C for 60 s and

final elongation at 72˚C for 5 min. PCR reactions were run in duplicate for each sample and pooled

afterwards. Purification of the amplicon libraries was performed using the Agencourt AMPure XP

bead protocol (Beckman Coulter, USA) and eluted in 23 µL nuclease-free water. Individual

libraries were quantified with Quant-iT HS dsDNA assay (Life Technologies, USA) and quality

checked on a Tapestation 2200 (Agilent, USA). Libraries were pooled in equimolar concentrations,

and diluted to 4 nM. The library pool was sequenced using an Illumina MiSeq (Illumina, USA) and

MiSeq reagent kit v3 (2x300 PE).


7

Bioinformatic processing and analysis

The obtained raw sequencing reads were quality filtered and trimmed using trimmomatic (v0.32) (20), only keeping reads with a minimum length of 275 bp. The trimmed reads were merged using

FLASH v. 1.2.7 (21) and read pairs between 425 and 525 bp in length were formatted for use with

the UPARSE workflow (22). Reads were dereplicated and clustered into Operational Taxonomical

Units (OTUs) using USEARCH7 at 97% sequence similarity. Taxonomy was assigned using the

RDP-classifier as implemented in QIIME (23) with a minimum confidence of 0.8 and Greengenes

(version 08-2013) as a reference database. Results were analysed in R studio (version 0.99.489 for

Mac) using the Ampvis package (24).

Statistical analyses

Principal Component Analysis was performed on square root transformed OTU abundances. A

constrained (by diet) redundancy analysis with plotting of bacterial culture and organic acid

parameters was performed to check for potential correlations between sequencing data, bacterial

enumerations and organic acid data. Significance of diet was tested on the first two principal

components (PCs) using the envfit parametric test and on the Bray-Curtis dissimilarity matrix using

the Adonis test (25). The parametric Wald-test (26) was used to test for significant OTU abundance

differences between the SM group and the MR and BC groups. OTUs with an adjusted p<0.001

were considered significantly different between the respective diets.

The impact of diet and age on bacterial and organic acid parameters, microbial richness, and

diversity index were investigated by fitting the data to a linear mixed model using the lmer function

from the lme4 package (27) using R studio (Version 0.99.489 for Mac). Diet and age/intestinal

segment were included as fixed effects, while pig and sow were included as random effects (by

including random intercept terms) to account for multiple observations made on the same litter and

on the same pig. When analysing the body weight variable, the piglets’ body weights at the

beginning of the experiment (day 23) were included as a co-variate. The fixed effects were tested

using an F-test with Kenward-Roger approximation, where the reduced model was tested against

the full model. This was done using the KRmodcomp function in the pbkrtest package (28). When a

fixed effect was found to be significant, a post-hoc test was performed using the multcomp package

and Bonferroni adjustment to correct for multiple comparisons (29). Effects were considered

significant when p<0.05 and as trends when 0.05≤p<0.10.


8

Results

During the course of the experiment, one BC-fed piglet was euthanised due to vomitus and general

weakness and one MR-fed piglet died. At 30 days of age, SM-fed piglets weighed more than BC-

fed (p=0.016) and MR-fed piglets (p=0.011) (Table 2). Diarrhoea incidence rate and the consumed

amounts of milk replacer and BC are also listed in Table 2.

Microbiome composition – 16S rRNA gene amplicon sequencing

Sequencing of 119 samples yielded a total of 2,090,874 sequences. A sequencing depth of 5000

sequences was considered appropriate from rarefaction curves, excluding four samples from

analysis (data not shown). Recovered sequences clustered into 2485 OTUs, which were classified

into 34 bacterial phyla, 154 families and 271 genera. Eight phyla had an overall relative abundance

above 1%.

Faecal microbiota

The relative abundance of the eight most abundant phyla (relative abundance > 1%) and 20 most

abundant genera are presented in Fig. 1A and B. Irrespective of diet and age, Firmicutes and

Bacteroidetes dominated the communities. Prevotella and Oscillospira were the most abundant and

stable genera both regarding diet and age. Of the 20 most abundant genera, ten belonged to the

phylum Bacteroidetes and nine to the phylum Firmicutes. The microbial community richness did

not differ between diet or age (Fig. 1C and S1A). The diversity, expressed as the Shannon index,

was higher in MR-fed compared to BC-fed piglets (p=0.015; Fig. 1D). There was a significant

effect of diet on the overall faecal microbial community composition of the three dietary groups on

days 25 (padonis=0.01) and 30 (padonis=0.008) (Fig. 2). However, no OTUs were found to differ

significantly in their read abundances between diets.

Digesta microbiota

Fig. 3A and B presents the eight most abundant phyla and 20 most abundant genera of the microbial

communities of the stomach, distal small intestine and mid-colon. Irrespective of diet, the microbial

communities of the stomach and mid-colon were dominated by Firmicutes, followed by

Bacteroidetes. The distal small intestinal community was dominated by Firmicutes in SM-fed

piglets, Firmicutes and Proteobacteria in MR-fed piglets, and Firmicutes followed by Proteobacteria

and Actinobacteria in BC-fed piglets. Overall, Lactobacillus and Prevotella were the most

dominating genera in the stomach and mid colon. In addition, Mitsuokella was the third most

dominating genus in the stomach of BC-fed piglets. The microbial community of the distal small

intestine was dominated by Lactobacillus (most pronounced in SM-fed piglets) and


9

Enterobacteriaceae in MR-fed piglets. As for the faecal samples, community richness did not differ

between diets; nor did the Shannon index (Fig. 3C and D, and S1B). Diet had a significant effect on

the overall microbial community of the stomach (padonis=0.001; Fig. 4A), distal small intestine

(padonis=0.001; Fig. 5A) and mid colon (padonis=0.001; Fig. 6A).

In the stomach, when comparing BC- to SM-fed piglets, six out of 551 OTUs were found to

have significantly different read abundances (Fig. 4B), all with a higher read abundance in BC-fed

piglets. OTU_72, belonging to Lactococcus, was the most significantly changed OTU, having a

higher read abundance in BC-fed piglets (p=5.3E-6). Nine out of 566 OTUs were found to have

significantly different read abundances between MR- and SM-fed piglets (Fig. 4C). OTU_66,

belonging to Lactobacillus, was the most significantly changed OTU, having a higher read

abundance in SM-fed piglets (p=2.7E-11).

In the distal small intestine, the three most influential taxa were responsible for the observed

grouping seen from the Principal Component Analysis (Fig 5A). Clustering of MR-fed piglets was

explained by the higher relative abundance of Enterobacteriaceae, clustering of SM-fed piglets was

explained by Lactobacillus, and clustering of BC-fed piglets was explained by Lactococcus.

Comparing BC- and MR-fed piglets with SM-fed piglets resulted in 11 out of 433 (BC-fed) and 435

(MR-fed) OTUs having significantly different read abundances (Fig. 5B and C). Again, OTU_72

belonging to Lactococcus was found to be the most significantly changed OTU, having a higher

read abundance in BC-fed piglets (p=2.2E-15). OTU_6, belonging to Enterobacteriaceae, was the

most significantly changed OTU when comparing MR- to SM-fed piglet (p=4.0E-10), having a

higher read abundance in MR-fed piglets.

Mid colon samples showed distinct grouping according to diet on PC3 (when plotted as a

function of PC1) with MR-fed piglets clustering by themselves (Fig. 6A). 14 out of 802 OTUs were

found to have significantly different read abundances when comparing BC- and SM-fed piglets

(Fig. 6B). The most significantly changed OTU was OTU_1 (p=4.7E-12) belonging to Lactobacillus,

and was found to have a higher read abundance in SM-fed piglets. Comparing MR- and SM-fed

piglets resulted in 13 out of 840 OTUs being significantly different (Fig. 6C). OTU_47, belonging

to Blautia, was the most significantly changed OTU (p=1.1E-13) and was found to have a higher

read abundance in MR-fed piglets.

Dry matter and pH

There was no difference in pH of digesta between diets. Dry matter content of digesta varied

between diets, being dependent on gut segment (Table S1). There was no difference in dry matter

content of digesta from the proximal small intestine and caecum between diets. Dry matter content

of digesta from the stomach (p<0.0001), proximal colon (p≤0.002), and mid colon (p≤0.006) was


10

highest in SM-piglets, while being higher in digesta from the distal small intestine (p=0.049) in BC-

piglets compared to MR-piglets. The lowest dry matter content of digesta from the distal colon was

found in MR-piglets (p≤0.0003).

Microbiological enumerations and concentration of organic acids

Faeces

The number of C. perfringens was lower on day 30 compared to day 23 (p=0.0006) and day 25

(p=0.0008) for all diets, but there was no difference in any of the investigated microbial groups

between diets (Table S2). Results of haemolytic bacteria have not been included due to the majority

of counts being below detection level.

Faecal concentrations of acetic (p≤0.004), propionic (p≤0.013), butyric (p≤0.0034) and the

sum of acetic, propionic and butyric acid (p≤0.001) were higher in BC-fed compared to SM- and

MR-fed piglets (Table 3). The concentration of the sum of iso-butyric and iso-valeric acid was

highest in BC-fed piglets (p≤0.009) on day 25, 27 and 30. The concentrations of propionic

(p≤0.037), butyric (p≤0.045), and the sum of acetic, propionic and butyric (p≤0.045) acid were

higher on day 25, 27 and 30 compared to day 23. The acetic acid concentration was higher on day

25 (p=0.014) and day 30 (p=0.001) compared to day 23. In BC-piglets, the concentration of the sum

of iso-butyric and iso-valeric acid was lowest on day 23 (p≤0.003).

Digesta

Haemolytic bacterial counts in digesta from the distal small intestine (p=0.0002), caecum

(p=0.003), and mid colon (p=0.001) were higher in MR-fed compared to SM-fed piglets (Table 4).

The number of C. perfringens was higher in all segments of BC-fed (p=0.041) compared to MR-fed

piglets but similar to those in the SM-fed group.

The concentration of the sum of iso-butyric and iso-valeric acid in digesta from the colon

(p≤0.02) was higher in BC-fed compared to MR-fed piglets, while the caecal concentration was

higher in SM-fed (p=0.036) compared to MR-fed piglets (Table 5).

Correlation between 16S rRNA gene sequences, organic acids and bacterial enumerations

The constrained redundancy analysis performed on all samples (16S rRNA amplicon data) with

fitted microbial enumerations and SCFA data (lactic acid omitted as detectable amounts were only

found in digesta) showed a clear separation between the different diets with positive correlations to

microbial enumerations and SCFA concentrations (Fig. 7). Samples from the SM-fed piglets

correlated with the number of lactic acid bacteria (r2=0.11), while samples from the MR-fed piglets

correlated with the number of haemolytic bacteria (r2=0.30) and yeast (r2=0.25) and to a lesser


11

degree with Enterobacteriaceae (r2=0.07). Samples from the BC-fed piglets correlated with the

concentration of iso-butyric (r2=0.38), iso-valeric (r2=0.32), the sum of iso-butyric and iso-valeric

acid (r2=0.36) and to a lesser degree with the number of C. perfringens (r2=0.10) and concentration

of acetic (r2=0.18), propionic (r2=0.15), butyric (r2=0.16), valeric (r2=0.21) and the sum of acetic,

propionic and butyric acid (r2=0.18).

Discussion

Several studies have investigated the effects of supplementary BC feeding on a variety of host

protective functions in pigs (e.g. (16), (13) and (30)). However, according to our knowledge no studies

have focused on the effect of BC on the gut microbiota when fed as sole nutrition. This should be

considered an important step in the process of optimising supplementary feeding of undersized

piglets, and in developing dietary strategies for optimal nutrition of preterm infants prone to

necrotising enterocolitis (31).

The piglets in the SM- and BC-fed dietary groups were weaned at 23 days of age. As

weaning is a highly stressful experience resulting in a decreased feed intake and nutrient digestion

capacity (32), the piglets in the current study were used as models for undersized piglets in regards to

having an unstable (immature) intestinal microbiota, suboptimal nutrient digestion and impaired

immune status.

The microbial community is known to change according to gut segment (33), and multiple

samples from the length of the gastrointestinal tract were therefore sampled. In addition, to be able

to follow the development of the microbiota from the same pigs over the course of the experiment,

faecal samples were included as well. Using 16S rRNA gene sequencing, we showed that there

were clear differences in the microbial communities from piglets fed different milk-based diets. As

expected, the microbial communities were gut region dependent, why differences between diets

varied in the different gut segments. The present results showed a clear influence of diet on the

microbial communities of the stomach, small intestine and colon. Diet did not influence the faecal

microbial community to the same level as the digesta communities, suggesting that the faecal

microbiota might need longer time to adjust in order to become diet specific.

Compared to the SM-fed piglets, the microbial communities of the stomach and small

intestine of BC-fed piglets indicated a shift in lactic acid bacteria (LAB) genera. OTUs belonging to

Lactococcus, Leuconostoc, Streptococcus and Carnobacterium were more abundant in BC-fed

piglets, whereas the LAB of SM-fed piglets was mainly dominated by Lactobacillus. Lactococcus

sp. is traditionally not found to be part of the commensal gut microbiota, but has been reported to

have bacteriocin producing properties (34). Lactococcus and Leuconostoc have most frequently been

associated with fermented dairy products, Carnobacterium have been found in dairy products, meat


12

and fish (35) and Streptococcus sp. has previously been isolated from the gastrointestinal tract of pigs (36). BC-fed piglets furthermore had a higher abundance of Mitsuokella in the stomach digesta,

which previously has been isolated from the stomach and colon digesta of pigs (36; 37). A previous

study reported an inhibiting effect of Mitsuokella jalaludinii on Salmonella typhimurium. Levine et

al. (38) found that the combined effect of acetic, lactic and succinic acid produced by M. jalaludinii

during fermentation, and the accompanied decrease in pH, was able to inhibit the growth of and

cellular invasion by S. typhimurium. The authors therefore proposed M. jalaludinii to have potential

as a probiotic species. In our study, we were not able to determine the Misuokella species in

question, but assuming that it was M. jalaludinii, it could be speculated on the mentioned beneficial

effects. Hence, the finding of potential probiotic bacteria in BC-fed piglets may have enhanced

pathogen resistance and intestinal mucosal integrity, as demonstrated by the immunological results

obtained by Sugiharto et al. (14) from the piglets included in the present study.

The shift in LAB genera seen in BC-fed piglets was not observed in MR-fed piglets. An in

vitro study showed a stimulating effect of BC on LAB growth rates, suggestively due to the

oligosaccharides found in colostrum (39). As BC has a higher content of oligosaccharides than

mature milk (40), such oligosaccharides might be the reason why BC-fed and not MR-fed piglets

were found to have a higher read abundance of LAB as Lactococcus, Mitsuokella and Leuconostoc

in their stomach and small intestinal digesta than SM-fed piglets.

16S rRNA gene sequencing on digesta from the distal small intestine revealed that MR-fed

piglets had a higher abundance of Enterobacteriaceae when compared to SM-fed piglets.

Enterotoxigenic Escherichia coli (ETEC) belongs to the Enterobacteriaceae family and is an

intestinal pathogen frequently observed after weaning causing post-weaning diarrhoea. ETEC

produces haemolysin, a virulence factor enabling haemolysis of red blood cells (41). By bacterial

culture, we observed a higher number of haemolytic bacteria in digesta from MR-fed compared to

SM- and BC-fed piglets. The higher abundance of Enterobacteriaceae observed using 16S rRNA

gene sequencing could therefore represent a higher abundance of potential intestinal pathogens as

ETEC. This is further supported by the higher diarrhoea incidence observed in MR- compared to

BC- and SM-fed piglets. The improved faecal consistency in BC-fed piglets is in accordance with

results by Huguet et al. (15) who found an improved sanitary status and faecal consistency in

weanling pigs supplemented with BC. The fact that BC-fed piglets in the present study experienced

less diarrhoea than MR-fed piglets, could be due to the effect of immunoglobulins and growth

promoting factors present in colostrum. The high content of IgG provides passive immune

protection to the newborn calf (42) and growth factors, as insulin-like growth factor, stimulate

enterocyte proliferation (9) potentially resulting in a less disturbed intestinal barrier. BC has

furthermore been reported to inhibit E. coli growth in vitro (39). De Vos et al. (30) investigated the


13

effect of feeding three-day-old piglets a milk replacer supplemented with a BC whey fraction on the

intestinal permeability, and found an increased occludin gene expression and decreased mannitol

absorption, thus indicating an effect on the enterocyte-to-enterocyte adherence and hence gut barrier

function. Thus, BC seems to have various bioactive factors of importance to intestinal immunity

and integrity.

Despite the high content of growth factors reported in BC, we found no difference in growth

performance between BC-fed and MR-fed piglets. The present study, however, was not designed as

a performance study per se. The study by De Vos et al. (30) did not obtain any effect on growth

performance when feeding BC-supplemented milk replacer to 3-10 days old piglets. Other studies

have reported an increased growth performance when piglets were fed weaning diets supplemented

with BC compared to un-supplemented diets (15; 43).

The chemical compositions of the diets were very different and with the most noticeably

difference being the protein content. The high protein content of BC was attributed to the high

concentration of immunoglobulins (12). As branched SCFAs are indicators of protein fermentation (44; 45), the higher concentrations of iso-valeric and iso-butyric acid in faeces and digesta from BC-

fed piglets most likely reflect that these piglets were fed a high-protein diet. Protein fermentation,

however, also produces unwanted and potentially toxic compounds such as ammonia (46), amines,

phenols and indols (45). Other reports have shown an increased shedding of E. coli from pigs fed

high-protein diets (47) and a decreased ileal cytokine response upon LPS-stimulation in pigs fed a

high-protein milk formula in early life (48). In the present study, we did not see any association

between the higher protein content in the BC diet and an increased shedding of pathogenic bacteria

as E. coli. Besides a higher concentration of branched SCFAs, feeding BC also resulted in higher

faecal concentrations of the straight SCFAs acetic, propionic and butyric acid, which are considered

to be beneficial to the host (45).

Conclusion

In conclusion, sequencing and bacterial culture indicated a higher shedding of potential pathogenic

ETEC in MR-fed piglets, while the microbiota of BC-fed piglets was characterised by a change in

LAB genera when compared to SM-fed piglets. Especially the ileal (distal small intestinal)

microbiota of BC-piglets had a closer resemblance to that of SM-piglets with regard to the

abundance of potential enteric pathogens than had the MR-fed piglets. Dietary treatment had less

influence on the faecal microbial community than on the digesta community, and this should be

taken into account when studying the impact of nutrition on the microbial community in other

species (e.g. humans) as well. However, our study does not enable us to account for the long-term

effects of feeding BC. Furthermore, read abundances obtained from 16S rRNA gene amplicon


14

sequencing should be supported by absolute quantifications. Additional studies on the potential

bioactivity of BC are needed to investigate whether the positive results on the intestinal microbiota

observed in this study are reproducible in groups of undersized weak piglets. Also, the effect of

higher branched SCFA concentrations in BC-fed piglets needs further investigation, as these

components indicate an increased protein fermentation in the gut.

Acknowledgements

The authors would like to thank Karin Durup, Mette Lykkegaard, Mette Kvist and Karin Johansen

for their skillful technical assistance during the course of the experiment.

Financial support

The Danish Council for Strategic Research funded the experiment through the NEOMUNE

consortium together with the Graduate School of Science and Technology, Aarhus University,

Denmark. Funders had no role in the design, analysis or writing of this article.

Conflict of interest

The bovine colostrum powder was partly sponsored by The European Colostrum Industry S.A. The

European Colostrum Industry S.A. had no role in the design, analysis or writing of this article.

Authorship

Authors contributed as follows: A.-S.R.P. conducted the animal experiment, performed sample and

data analyses and drafted the manuscript. S.S. conducted the animal experiment. N.C. designed the

experiment, contributed to the conduction of the animal experiment and assisted with data analysis.

N.d.J and J.L.N performed data analysis. C.L. designed the experiment. All authors have revised

and accepted the final manuscript.

(49)


15

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17

Tables:

Table 1

Chemical compositions of sow milk (SM), milk replacer (MR) and bovine colostrum (BC).

Item Dietary group SM MR BC

Dry matter (%) 17.9* 95.0 96.1

Protein (%) 5.1* 22.1 68.2

Fat (%) 6.5* 13.2 2.0

Ash (%) 1.0† 6.8 6.0

Immunoglobulins (% of DM)‡

IgG IgA IgM

0.11 2.18 0.56

0.05 0.01 ND

38.4 3.59 2.52

Chemical analyses (drymatter, protein, fat and ash) were performed by Eurofins Steins Laboratory A/S,

Odense, Denmark.

DM = dry matter.

ND = not detected.

* Adopted from Lauridsen and Danielsen (50).

† Adopted from Aguinaga et al. (51).

‡ Immunoglobulin concentrations are adopted from Sugiharto et al. (14).


18

Table 2

Body weight at 23 (initial body weight) and 30 days of age, milk replacer and bovine colostrum

powder intake, and diarrhoea incidence rate*

Item Dietary group† p-value SM MR BC

Body weight day 23 (kg) 8.9 (8.5-9.4) 8.1 (7.7-8.5) 8.2 (7.8-8.6) Body weight day 30 (kg) 10.2b (9.8-10.6) 9.4a (8.9-9.8) 9.4a (9.0-9.9) 0.005 Powder intake (g) ND 3980 (2168-5791) 2653 (842-4465) 0.25 Diarrhoea incidence rate‡ 0.038 0.172 0.054

ND = not determined. a,b Values with different superscripts within a row are significantly different (p<0.05).

* Values are presented as least square means and 95% confidence intervals (in parentheses).

† SM = Sow Milk; MR = Powdered porcine milk replacer; BC = Spray-dried bovine colostrum powder.

Number of piglets: SM=12; MR=11; BC=11.

‡ Defined as the ratio between the number of new cases of diarrhoea in the study period and the total number

of days the piglets have been in risk (i.e. the number of days from the animal enters the study until the

animal (a) shows clinical signs of diarrhoea, (b) dies or (c) the study ends) (49).


19

Tab

le 3

Sh

ort-c

hain

fatty

aci

d (S

CFA

) con

cent

ratio

ns (m

mol

/kg

sam

ple)

in fa

eces

from

pig

lets

at 2

3, 2

5, 2

7 an

d 30

day

s of

age

*

D

ieta

ry g

roup

†

p-va

lue

Item

SM

M

R

BC

#

D‡

A§

DxA

Ace

tic a

cid

y

y

z

0.

0005

0.

0008

0.

06

Day

23

30.5

(2

2.8-

38.3

) 28

.0

(20.

2-35

.7)

46.3

(3

8.5-

54.0

) a

D

ay 2

5 42

.0

(34.

1-49

.8)

39.4

(3

1.5-

47.3

) 57

.7

(49.

9-65

.5)

b

Day

27

Day

30

37.6

(2

9.8-

45.4

) 35

.1

(27.

2-42

.9)

53.4

(4

5.3-

61.5

) ab

45.3

(3

7.5-

53.1

) 42

.8

(34.

8-50

.7)

61.1

(5

3.1-

69.0

) b

Pr

opio

nic

acid

y

y

z

0.00

2 <0

.000

1 0.

41

Day

23

7.4

(5.6

-9.7

) 6.

6 (5

.0-8

.7)

11.9

(9

.0-1

5.6)

a

D

ay 2

5 10

.9

(8.2

-14.

4)

9.8

(7.4

-12.

9)

17.5

(1

3.2-

23.0

) b

D

ay 2

7 13

.4

(10.

2-17

.7)

12.0

(9

.1-1

5.9)

21

.5

(16.

1-28

.7)

b

Day

30

15.4

(1

1.7-

20.3

) 13

.8

(10.

4-18

.3)

24.7

(1

8.6-

32.7

) b

B

utyr

ic a

cid

y

y

z

0.

0008

0.

002

0.11

D

ay 2

3 1.

6 (0

.8-2

.9)

0.9

(0.4

-1.7

) 3.

9 (2

.2-6

.8)

a

Day

25

3.9

(2.2

-6.8

) 2.

3 (1

.2-4

.1)

9.0

(5.2

-15.

4)

b

Day

27

3.5

(2.0

-6.1

) 2.

1 (1

.1-3

.7)

8.2

(4.6

-14.

3)

b

Day

30

4.5

(2.5

-7.7

) 2.

7 (1

.4-4

.8)

10.3

(5

.9-1

7.7)

b

A

+P+B�||

y

y

z

<0

.000

1 0.

0002

0.

09

Day

23

41.8

(2

8.4-

55.2

) 34

.8

(21.

4-48

.2)

71.3

(5

8.5-

85.4

) a

D

ay 2

5 61

.8

(48.

1-75

.4)

54.8

(4

1.1-

68.5

) 91

.9

(78.

4-10

5.5)

b

D

ay 2

7 59

.6

(46.

0-73

.1)

52.6

(3

9.0-

66.2

) 89

.7

(75.

6-10

3.8)

b

D

ay 3

0 70

.8

(57.

2-84

.4)

63.8

(4

9.9-

77.7

) 10

1.0

(87.

2-11

4.8)

b

IB

+IV

�

<.

0001

0.

002

0.00

04

Day

23

3.5

(2.2

-5.5

) 2.

0 (1

.2-5

.5)

3.7A

(2

.4-5

.7)

Day

25

3.3b

(2.1

-5.3

) 3.

5b (2

.2-5

.3)

10.8

aB

(7.0

-16.

4)

Day

27

3.8b

(2.4

-5.9

) 2.

0b (1

.2-5

.9)

11.9

aB

(7.3

-19.

4)

Day

30

5.0b

(3.2

-7.7

) 3.

0b (1

.8-7

.7)

12.9

aB

(8.3

-20.

0)


20

a,b V

alue

s w

ith d

iffer

ent s

uper

scrip

ts w

ithin

a ro

w a

re s

igni

fican

tly d

iffer

ent.

y, z

: Col

umns

with

diff

eren

t let

ters

with

in a

SC

FA g

roup

are

sig

nific

antly

diff

eren

t. A

,B: V

alue

s w

ith d

iffer

ent s

uper

scrip

ts w

ithin

a c

olum

n ar

e si

gnifi

cant

ly d

iffer

ent.

* V

alue

s ar

e pr

esen

ted

as le

ast s

quar

e m

eans

and

95%

con

fiden

ce in

terv

al (i

n pa

rent

hese

s).

† SM

= s

ow m

ilk, M

R =

milk

repl

acer

, BC

= b

ovin

e co

lost

rum

. Num

ber o

f pig

lets

: BC

=12,

exc

ept d

ay 5

(n=9

) and

day

8 (n

=11)

; MR

=12,

exc

ept d

ay 3

(n=1

1) a

nd

day

8 (n

=10)

; SM

=12,

exc

ept d

ay 3

(n=1

0).

#: R

ows

with

diff

eren

t let

ters

with

in a

SC

FA g

roup

are

sig

nific

antly

diff

eren

t.

‡ D

= D

iet.

§ A

= A

ge.

|| A

+P+B

= a

cetic

+ p

ropi

onic

+ b

utyr

ic a

cid.

¶ IB+I

V =

iso-

buty

ric +

iso-

vale

ric a

cid.


21

Tab

le 4

C

ount

s (lo

g cf

u/g

sam

ple)

of s

elec

ted

mic

robi

al g

roup

s in

dig

esta

from

the

gast

roin

test

inal

trac

t of 3

0 da

ys o

ld p

igle

ts (e

nd o

f exp

erim

ent)*

Die

tary

gro

up†

p-va

lue

Item

SM

M

R

BC

D

‡ S§

D

xS

Ente

roba

cter

iace

ae

0.

42

<.00

01

0.08

Stom

ach

<4.9

(1)

(3.9

-5.8

) <5

.5 (3

) (4

.5-6

.4)

<5.1

(2)

(4.1

-6.0

)

Dis

tal s

mal

l int

estin

e 7.

7 (6

.8-8

.7)

8.3

(7.4

-9.3

) 7.

9 (7

.0-8

.9)

C

aecu

m

8.1

(7.1

-9.0

) 8.

7 (7

.7-9

.6)

<8.3

(1)

(7.3

-9.2

)

Mid

col

on

8.2

(7.2

-9.1

) 8.

8 (7

.8-9

.7)

8.4

(7.4

-9.3

)

Hae

mol

ytic

bac

teri

a

0.

003

<.00

01

<0.0

001

Stom

ach

<4.0

(3)

(3.2

-4.9

) <4

.3 (2

) (3

.5-5

.2)

<4.5

(4)

(3.6

-5.3

)

Dis

tal s

mal

l int

estin

e <5

.1a (3

) (4

.2-5

.9)

8.3b

(7.4

-9.1

) <6

.8b (2

) (5

.9-7

.6)

C

aecu

m

<6.2

a (4)

(5.3

-7.0

) 8.

6b (7

.7-9

.4)

<7.4

ab (2

) (6

.6-8

.3)

M

id c

olon

<6

.0a (3

) (5

.1-6

.8)

8.6b

(7.8

-9.5

) <7

.4ab

(2)

(6.5

-8.2

)

Clo

stri

dium

per

frin

gens

ab

a

b

0.03

<.

0001

0.

15

Stom

ach

4.3

(3.7

-5.0

) 3.

3 (2

.7-3

.9)

4.6

(4.0

-5.2

)

Dis

tal s

mal

l int

estin

e 5.

2 (4

.5-5

.8)

<4.1

(4)

(3.5

-4.8

) 5.

4 (4

.8-6

.0)

C

aecu

m

5.5

(4.9

-6.1

) <4

.5 (2

) (3

.8-5

.1)

<5.7

(1)

(5.1

-6.4

)

Mid

col

on

5.4

(4.7

-6.0

) <4

.3 (2

) (3

.7-5

.0)

5.6

(5.0

-6.2

)

Lact

ic a

cid

bact

eria

0.

07

<.00

01

0.16

Stom

ach

8.5

(8.0

-9.0

) 7.

8 (7

.3-8

.3)

8.1

(7.6

-8.6

)

Dis

tal s

mal

l int

estin

e 8.

3 (7

.8-8

.8)

7.6

(7.1

-8.1

) 7.

9 (7

.4-8

.4)

C

aecu

m

Mid

col

on

8.6

(8.1

-9.1

) 7.

9 (7

.4-8

.4)

8.2

(7.7

-8.7

)

9.0

(8.5

-9.5

) 8.

3 (7

.8-8

.8)

8.6

(8.1

-9.1

)

Yea

st

0.12

0.

001

0.30

St

omac

h <4

.1 (2

) (3

.2-5

.1)

5.5

(4.5

-6.4

) <4

.4 (1

) (3

.4-5

.3)

D

ista

l sm

all i

ntes

tine

4.1

(3.2

-5.1

) 5.

5 (4

.5-6

.5)

4.4

(3.4

-5.3

)

Cae

cum

<4

.3 (1

) (3

.3-5

.2)

5.6

(4.7

-6.6

) 4.

5 (3

.6-5

.5)

M

id c

olon

4.

6 (3

.7-5

.6)

6.0

(5.1

-7.0

) 4.

9 (3

.9-5

.9)

To

tal a

naer

obic

bac

teria

0.

47

<.00

01

0.34

St

omac

h 8.

5 (8

.2-8

.9)

8.3

(7.9

-8.7

) 8.

5 (8

.1-8

.9)

D

ista

l sm

all i

ntes

tine

8.9

(8.5

-9.3

) 8.

7 (8

.3-9

.0)

8.9

(8.5

-9.3

)

Cae

cum

9.

4 (9

.0-9

.8)

9.2

(8.8

-9.6

) 9.

4 (9

.0-9

.8)

M

id c

olon

9.

8 (9

.4-1

0.2)

9.

5 (9

.1-9

.0)

9.8

(9.4

-10.

2)


22

a,b:

Col

umns

with

diff

eren

t let

ters

with

in a

mic

robi

al g

roup

are

sig

nific

antly

diff

eren

t. a,

b Val

ues

with

diff

eren

t sup

ersc

ripts

with

in a

row

are

sig

nific

antly

diff

eren

t.

<: In

dica

tes

that

at l

east

one

of t

he o

bser

vatio

ns u

sed

to c

alcu

late

the

leas

t squ

are

mea

n w

as b

elow

det

ectio

n le

vel.

Num

bers

in b

rack

ets

indi

cate

how

man

y

sam

ples

wer

e be

low

det

ectio

n le

vels

.

* Sa

mpl

es fr

om th

e st

omac

h, d

ista

l sm

all i

ntes

tine,

cae

cum

and

mid

col

on w

ere

anal

ysed

. Val

ues

are

pres

ente

d as

leas

t squ

are

mea

ns a

nd 9

5% c

onfid

ence

inte

rval

(in

pare

nthe

ses)

.

† SM

= s

ow m

ilk, M

R =

milk

repl

acer

, BC

= b

ovin

e co

lost

rum

. Num

ber o

f pig

lets

: BC

=4; M

R=4

; SM

=4.

‡ D

= D

iet.

§ S =

Inte

stin

al s

egm

ent.


23

Tab

le 5

O

rgan

ic a

cid

conc

entra

tions

(m

mol

/g s

ampl

e) i

n di

gest

a fr

om f

our

segm

ents

of

the

gast

roin

test

inal

tra

ct o

f 30

day

s ol

d pi

glet

s (e

nd o

f

expe

rimen

t)*

D

ieta

ry g

roup

† p-

valu

e Ite

m

SM

MR

B

C

D‡

S§

DxS

La

ctic

aci

d ||

0.

11

0.64

0.

09

Stom

ach

9.0

(4.1

-17.

5)

7.2

(3.2

-14.

5)

2.8

(0.7

-6.6

)

Dis

tal s

mal

l int

estin

e 8.

7 (4

.0-1

7-1)

7.

1 (3

.1-1

4.2)

2.

8 (0

.7-6

.5)

A

cetic

aci

d

!0.

42

<.00

01

0.46

Stom

ach

3.4

(1.6

-6.3

) 2.

7 (1

.2-5

.2)

4.1

(2.0

-7.5

)

Dis

tal s

mal

l int

estin

e 7.

1 (3

.8-1

2.5)

5.

8 (3

.0-1

0.4)

8.

3 (4

.6-1

4.6)

Cae

cum

52

.5

(31.

0-88

.5)

44.1

(2

5.7-

75.2

) 61

.0

(36.

1-10

2.7)

Mid

col

on

44.0

(2

6.0-

73.9

) 36

.9

(21.

7-62

.3)

51.2

(3

0.3-

85.8

)

Prop

ioni

c ac

id

0.10

<.

0001

0.

30

Stom

ach

2.5

(1.2

-4.8

) 1.

5 (0

.6-3

.0)

3.3

(1.7

-6.2

)

Dis

tal s

mal

l int

estin

e 0.

8 (0

.2-1

.8)

0.4

(0.0

-1.0

) 1.

2 (0

.4-2

.4)

C

aecu

m

14.6

(8

.0-2

6.3)

9.

4 (5

.0-1

7.4)

18

.7

(10,

3-33

.6)

M

id c

olon

17

.4

(9.7

-31.

1)

11.3

(6

.1-2

0.3)

22

.3

(12.

4-39

.7)

B

utyr

ic a

cid �

0.

56

0.64

0.

13

Cae

cum

6.

9 (2

.6-1

1.3)

5.

6 (1

.1-1

0.1)

8.

2 (3

.8-1

2.5)

Mid

col

on

6.8

(2.5

-11.

2)

5.5

(1.1

-9.9

) 8.

1 (3

.7-1

2.4)

A+P

+B**

0.

26

<.00

01

0.04

St

omac

h 6.

8 (3

.7-1

2.4)

4.

9 (2

.6-8

.9)

7.7

(4.2

-14.

2)

D

ista

l sm

all i

ntes

tine

8.4

(4.6

-15.

4)

6.0

(3.3

-11.

1)

9.6

(5.2

-17.

6)

C

aecu

m

76.9

(4

1.7-

141.

6)

55.0

(2

9.4-

102.

8)

87.6

(4

7.6-

161.

3)

M

id c

olon

71

.9

(39.

2-13

1.7)

51

.4

(28.

0-94

.5)

81.9

(4

4.7-

150.

0)

IB

+IV

��†

†

0.

02

0.65

0.

03

Cae

cum

5.

8a (3

.3-1

0.2)

1.

7b (0

.8-3

.2)

4.7ab

(2

.6-8

.3)

M

id c

olon

4.

1ab

(2.3

-7.3

) 2.

2a (1

.2-4

.0)

10.0

b (5

.7-1

7.5)


24

a,b V

alue

s w

ith d

iffer

ent s

uper

scrip

ts w

ithin

a ro

w a

re s

igni

fican

tly d

iffer

ent.

* V

alue

s ar

e pr

esen

ted

as le

ast s

quar

e m

eans

and

95%

con

fiden

ce in

terv

al (i

n pa

rent

hese

s). S

ampl

es fr

om th

e st

omac

h, d

ista

l sm

all i

ntes

tine,

cae

cum

and

mid

colo

n w

ere

anal

ysed

.

† SM

= s

ow m

ilk, M

R =

milk

repl

acer

, BC

= b

ovin

e co

lost

rum

. Num

ber o

f pig

lets

: BC

=4; M

R=4

; SM

=4, e

xcep

t seg

men

t 8 (n

=2).

‡ D

= D

iet.

§ S

= In

test

inal

seg

men

t.

|| Sa

mpl

es fr

om th

e ca

ecum

and

mid

col

on h

ad v

alue

s be

low

det

ectio

n le

vel.

¶ Sa

mpl

es fr

om th

e st

omac

h an

d di

stal

sm

all i

ntes

tine

had

valu

es b

elow

det

ectio

n le

vel.

** A

+P+B

= a

cetic

+ p

ropi

onic

+ b

utyr

ic a

cid.

†† IB

+IV

= is

o-bu

tyric

+ is

o-va

leric

aci

d.


25

Figures:

Fig. 1. Heatmaps, estimated species richness and Shannon diversity index of faecal samples

collected at 23, 25, 27, and 30 days of age from piglets fed sow milk (SM; n=15), milk replacer

(MR; n=15) or bovine colostrum (BC; n=16). Heatmaps show the relative abundances (%) of (A)

the eight most abundant phyla and (B) the 20 most abundant genera in faecal samples. Colours

represent the relative abundances. Boxplots show the (C) estimated species richness and (D)

Shannon diversity index.

Fig. 2. Principal Component Analysis of square root transformed OTU abundances in faeces (n=46)

displaying PC1 and PC2. Points are coloured for diet and grouped according to age.

Fig. 3. Heatmaps, estimated species richness and Shannon diversity index of stomach, distal small

intestinal and mid colon digesta samples from piglets fed sow milk (SM; n=23), milk replacer (MR;

n=23) or bovine colostrum (BC; n=23). Heatmaps show the relative abundances (%) of (A) the

eight most abundant phyla and (B) 20 most abundant genera. Colours represent relative abundances.

Boxplots show the (C) estimated species richness and (D) Shannon diversity index.

Fig. 4. (A) Principal Component Analysis of square root transformed OTU abundances displaying

PC1 and PC2; stomach content (n=21). Points are coloured for diet. Boxplots show the OTUs

significantly different between (B) sow milk fed and bovine colostrum fed piglets and (C) sow milk

fed and milk replacer fed piglets.


PC1 and PC2; distal small intestinal content (n=24). Points are coloured for diet. Boxplots show the

OTUs significantly different between (B) sow milk fed and bovine colostrum fed piglets and (C)

sow milk fed and milk replacer fed piglets.


PC1 and PC3; mid colon content (n=24). Points are coloured for diet. Boxplots show the OTUs

significantly different between (B) sow milk fed and bovine colostrum fed piglets and (C) sow milk

fed and milk replacer fed piglets.


26

Fig. 7. Constrained redundancy analysis of square root transformed OTU abundances fitted with

microbial culture and short-chain fatty data (n=115), displaying RDA1 and RDA2. The arrows

point toward the highest values and the length of the arrow indicates the parameters significance.

Ana = total anaerobic bacteria; Clos = Clostridium perfringens; Entero = Enterobacteriaceae; Hem

= haemolytic bacteria; Lab = lactic acid bacteria. Ace = Acetic acid; Apb = Acetic + Propionic +

Butyric acid; But = Butyric acid; Pro = Propionic acid; Iso_But = Iso-butyric acid; Iso_Val = Iso-

valeric acid; Ibiv = Iso-butyric + iso-valeric acid; Val = Valeric acid.

Fig. S1. Boxplots showing the observed species richness in (A) faecal samples (n=46) collected at

23, 25, 27 and 30 days of age and (B) stomach (n=21), distal small intestinal (n=24) and mid colon

(n=24) digesta from piglets fed sow milk (SM), milk replacer (MR) or bovine colostrum (BC).


2325

2730

0.1

1.5

1.7

0.4

0.9

1.1

66.8

26.1

0.5

32.4 20.8

0.8

0.2

0.5

61.4

0.3

1.1

40.4

1.3

1.250 0.5

4.3

0.4

41.9

0.5 055.1

0.3

0.4

0.6

0.9

26.8

65.1 1 0.1

0.5

1.3

3.1

0.3

38.4

54.1

3.6 1 1.4

0.5

0.1

45.4

0.445 5.6

1.6

0.8 00.9

46 1.6

1.7

47.3

1.4

0.4

0.1

0.5

11.6

40.9

1.6

0.7

43.4

0.3

0.1

0.4

1.3

0.4

0.6

0.441 0.1

55.5

0.6

15.1

45.6

1.1

0.1

44.7

0.3

0.5

0.3

0.1

2.2

53.3

39.1

2.1

1.6

0.7

Syne

rgis

tete

s

Actin

obac

teria

Tene

ricut

es

Spiro

chae

tes

Prot

eoba

cter

ia

Fuso

bact

eria

Bact

eroi

dete

s

Firm

icut

es

23 colostrum

23 milk replacer

23 sow milk

25 colostrum

25 milk replacer

25 sow milk

27 colostrum

27 milk replacer

27 sow milk

30 colostrum

30 milk replacer

30 sow milk

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Day

23

Day

25

Day

27

Day

30

BC

M

R

S

M

BC

M

R

S

M

BC

M

R

S

M

BC

M

R

S

M

2325

2730

1.2 55.4

0.4

0.2

9.5

1.4

1.2

0.4

3.9

1.7

4.3

2.3 31.1

1.8

15.4

0.1

0.3

0.6

0.9

0.1

0.5

5.7

1.5

9.7

1.2

1.6

6.1

1.33 1.7 3 0.2

0.67 20.5

10.7

1.3

1.3 0 1.8

3.3

0.3

0.8

1.2

2.1

4.6

1.4

0.684.9

1.19 4.3

8.8

14.3

0.2

1.4

2.8

1.1

0.4

1.9

0.8

0.4

3.8

1.4

0.1

2.3

17.7

0.3

1.7

15.6

2.6

12.2

0.3

1.8 40.4

0.5

2.2

1.4

3.2

4.1

0.1

3.8

0.8

0.7

7.4

1.5

2.2

1.6

2.6

0.9

2.3

0.5

2.2

5.3

0.1

0.4

2.2

2.6

0.901.7

3.1

6.5

2.5 2 0.4

13.9

1.1

4.9

3.6

0.5

1.513.6

6.8

0.6

4.3

7.5

0.81 0.6

1.3

1.2

1.5

0.4

1.1

4.5

0.4

2.7

6.2

2.6

15.2

5.6

0.2

10.7

1.2

0.4

4.7

8.5 10 2.3

0.2

1.5

1.3

0.9

0.5

0.2

0.1

17.8

1.6

1.2

2.7

0.3

3.8

0.7

5.2

0.6

0.8

0.9

1.1

5.6 12.2

0.8

24.6

0.501.6

1.7

6.3

0.5

0.8

11.6

2.4

0.29 0.1

0.5

6.5

1.7

9.8

5.5

0.6

0.7

0.7

18.7

0.7

0.3

8.7

0.3

2.1

1.9

4.5

0.4

0.3

0.9

3.6 026.1

4.2

0.7

1.64 00.1

2.2

1.3

2.9

0.1

1.3

0.5

17.9

0.1

02 0.1 03.3

2.5

1.864.8

8.4

1.2

1.6

7.6

2.3

1.7

0.4 01.2

18.5

1.6

Firm

icute

s; f_

_Rum

inoc

occa

ceae

_OTU

_20

Bact

eroi

dete

s; f_

_BS1

1_OT

U_12

4Fi

rmicu

tes;

Meg

asph

aera

Bact

eroi

dete

s; P

arab

acte

roid

esFi

rmicu

tes;

f__R

umin

ococ

cace

ae_O

TU_1

605

Firm

icute

s; f_

_Rum

inoc

occa

ceae

_OTU

_29

Firm

icute

s; f_

_Rum

inoc

occa

ceae

_OTU

_21

Bact

eroi

dete

s; f_

_p−2

534−

18B5

_OTU

_52

Bact

eroi

dete

s; f_

_S24−7

_OTU

_100

Bact

eroi

dete

s; o

__Ba

cter

oida

les_

OTU_

4Ba

cter

oide

tes;

f__S

24−7

_OTU

_27

Bact

eroi

dete

s; B

acte

roid

esFi

rmicu

tes;

Rum

inoc

occu

sFu

soba

cter

ia; F

usob

acte

rium

Bact

eroi

dete

s; [P

revo

tella

]Fi

rmicu

tes;

Lac

toba

cillu

sBa

cter

oide

tes;

f__[

Para

prev

otel

lace

ae]_

OTU_

19Fi

rmicu

tes;

p−7

5−a5

Firm

icute

s; O

scillo

spira

Bact

eroi

dete

s; P

revo

tella

23 colostrum

23 milk replacer

23 sow milk

25 colostrum

25 milk replacer

25 sow milk

27 colostrum

27 milk replacer

27 sow milk

30 colostrum

30 milk replacer

30 sow milk

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Day

23

Day

25

Day

27

Day

30

Bact

eroi

dete

s; fa

mily

[Par

apre

vote

llace

ae] O

TU 1

9

Bact

eroi

dete

s; fa

mily

S24

-7 O

TU 2

7

Bact

eroi

dete

s; o

rder

Bac

tero

idal

es O

TU 4

Bact

eroi

dete

s; fa

mily

S24

-7 O

TU 1

00

Bact

eroi

dete

s; fa

mily

p-2

534-

16B5

OTU

52

Firm

icut

es; f

amily

Rum

inoc

occa

ceae

OTU

21

Firm

icut

es; f

amily

Rum

inoc

occa

ceae

OTU

29

Firm

icut

es; f

amily

Rum

inoc

occa

ceae

OTU

160

5

Bact

eroi

dete

s; fa

mily

BS1

1 O

TU 1

24

Firm

icut

es; f

amily

Rum

inoc

occa

ceae

OTU

20

BC

MR

S

M

BC

MR

S

M

BC

MR

S

M

BC

MR

S

M

2325

2730

250

375

500

625

750

875

1000

colos

trum

milk re

place

rso

w milk

colos

trum

milk re

place

rso

w milk

colos

trum

milk re

place

rso

w milk

colos

trum

milk re

place

rso

w milk

Diet

Estimated number of OTUs Estimated number of OTUs

Day

23

Day

25

Day

27

Day

30

BC

MR

S

M

BC

MR

S

M D

iet

BC

MR

S

M

BC

MR

S

M

2325

2730

2.00

2.25

2.50

2.75

3.00

3.25

3.50

3.75

4.00

4.25

4.50

4.75

5.00

5.25

colos

trum

milk

repla

cer

sow

milk

colos

trum

milk

repla

cer

sow

milk

colos

trum

milk

repla

cer

sow

milk

colos

trum

milk

repla

cer

sow

milk

Die

t

Diversity index Diversity index

Day

23

Day

25

Day

27

Day

30

BC

MR

S

M

BC

MR

S

M

BC

MR

S

M

BC

MR

S

M

Die

t

(A)

(B)

(C)

(D)

Figu

re 1


23 25 27 30

−5.0

−2.5

0.0

2.5

5.0

−5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5PC1 [14.8%]

PC2

[13.

3%] Diet

colostrum

milk replacer

sow milk

PC

2 - v

aria

tion

expl

aine

d 13

.3%

PC1 - variation explained 14.8%

Day 23 Day 25 Day 27 Day 30

23 25 27 30

−5.0

−2.5

0.0

2.5

5.0

−5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5PC1 [14.8%]

PC2

[13.

3%] Diet

colostrum

milk replacer

sow milk

23 25 27 30

−5.0

−2.5

0.0

2.5

5.0

−5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5PC1 [14.8%]

PC2

[13.

3%] Diet

colostrum

milk replacer

sow milk

PC

2 - v

aria

tion

expl

aine

d 13

.3%



23 25 27 30

−5.0

−2.5

0.0

2.5

5.0

−5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5 −5.0 −2.5 0.0 2.5PC1 [14.8%]

PC2

[13.

3%] Diet

colostrum

milk replacer

sow milk

Figure 2


Dis

tal S

mal

l Int

estin

eM

id C

olon

Stom

ach

10.6

1.8 0069.9

16.7

0.6 0

046 0.2 0 051.5

1.1

0.9

03.2 000.1

92.9

1.2

2.1

2 1.2

1.4

1.2

1.1

1.2

55.8

35.6

0.7 13.9

57.6

7.5

0.4

27.6

0.3

0.4

0.6

1.6

0.9

61.9 132.1

0.8

005.4 01.1

20.1

68.3

3.9

0 0.1

5.7

2.7

0.6 082.6

7.3

0.2 077.7

3.4

1.5

16.7 0 0

Spiro

chae

tes

Tene

ricut

es

Plan

ctom

ycet

es

Fuso

bact

eria

Actin

obac

teria

Prot

eoba

cter

ia

Bact

eroi

dete

s

Firm

icut

es

colostrum Distal Small Intestine

milk replacer Distal Small Intestine

sow milk Distal Small Intestine

colostrum Mid Colon

milk replacer Mid Colon

sow milk Mid Colon

colostrum Stomach

milk replacer Stomach

sow milk Stomach

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Dis

tal S

mal

l Int

estin

eM

id C

olon

Stom

ach

10.6

1.8 0069.9

16.7

0.6 0

046 0.2 0 051.5

1.1

0.9

03.2 000.1

92.9

1.2

2.1

2 1.2

1.4

1.2

1.1

1.2

55.8

35.6

0.7 13.9

57.6

7.5

0.4

27.6

0.3

0.4

0.6

1.6

0.9

61.9 132.1

0.8

005.4 01.1

20.1

68.3

3.9

0 0.1

5.7

2.7

0.6 082.6

7.3

0.2 077.7

3.4

1.5

16.7 0 0

Spiro

chae

tes

Tene

ricut

es

Plan

ctom

ycet

es

Fuso

bact

eria

Actin

obac

teria

Prot

eoba

cter

ia

Bact

eroi

dete

s

Firm

icut

es




colostrum Mid Colon


sow milk Mid Colon

colostrum Stomach


sow milk Stomach

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Dis

tal S

mal

l Int

estin

eM

id C

olon

Stom

ach

10.6

1.8 0069.9

16.7

0.6 0

046 0.2 0 051.5

1.1

0.9

03.2 000.1

92.9

1.2

2.1

2 1.2

1.4

1.2

1.1

1.2

55.8

35.6

0.7 13.9

57.6

7.5

0.4

27.6

0.3

0.4

0.6

1.6

0.9

61.9 132.1

0.8

005.4 01.1

20.1

68.3

3.9

0 0.1

5.7

2.7

0.6 082.6

7.3

0.2 077.7

3.4

1.5

16.7 0 0

Spiro

chae

tes

Tene

ricut

es

Plan

ctom

ycet

es

Fuso

bact

eria

Actin

obac

teria

Prot

eoba

cter

ia

Bact

eroi

dete

s

Firm

icut

es




colostrum Mid Colon


sow milk Mid Colon

colostrum Stomach


sow milk Stomach

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Dis

tal S

mal

l Int

estin

eM

id C

olon

Stom

ach

10.6

1.8 0069.9

16.7

0.6 0

046 0.2 0 051.5

1.1

0.9

03.2 000.1

92.9

1.2

2.1

2 1.2

1.4

1.2

1.1

1.2

55.8

35.6

0.7 13.9

57.6

7.5

0.4

27.6

0.3

0.4

0.6

1.6

0.9

61.9 132.1

0.8

005.4 01.1

20.1

68.3

3.9

0 0.1

5.7

2.7

0.6 082.6

7.3

0.2 077.7

3.4

1.5

16.7 0 0

Spiro

chae

tes

Tene

ricut

es

Plan

ctom

ycet

es

Fuso

bact

eria

Actin

obac

teria

Prot

eoba

cter

ia

Bact

eroi

dete

s

Firm

icut

es




colostrum Mid Colon


sow milk Mid Colon

colostrum Stomach


sow milk Stomach

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Sto

mac

hD

ista

l sm

all i

ntes

tine

Mid

col

on

BC

MR

SM

B

C

M

R

S

M

BC

MR

SM

Dist

al S

mal

l Int

estin

eM

id C

olon

Stom

ach

66.2

5.4

4.8

1.23 08.3 0 07.4

8.6

9.2

0.1

3.700.3

0.1

13.4 0

09.6

34.8

0.2

0.1

0.7

1.9

4.8 00.2

5.1

1.3

0.1 325.5

0.8

1.9 0 00

0.3

0.4

1.2

0.5011.6

0.8 02.8

4.401.6

0.164 1.9 00 0 0.20

0.1

0.1 0 40 13.4

1.5 01.4

8.8

0.42 0 0.1

2.4 00.7

3.8

1.3

11.9

4.8

3.7

0.4

1.4

8.1

0.2 0 2.1 00.4 00.6

0.1

5.2

1.3

0.7

0.9 00.1

4.3

0.1

0.1

2.8

5.8 400.1

1.9

1.7 0014.1

2.1

1.5

16.6

0.2 00.1 54.4

6.2

0.2

1.6

2.3

0.1

0.4

0.2

0.2

16.4

14.8 011 0.1016.5

0.7

0.9

0.2

1.6

0.8

11.3

0.1

2.9

5.4

0.8

0.1

0.21 00.5

1.3

3.2

0.2

3.5

1.6

0.8

37.5

0.1

1.4

0.5

0.701.4

0.4 0 0 0015.8

9.3 053.4

0.3

0.7

2.80 00.3

3.2

0.1

Firm

icute

s; B

acillu

sAc

tinob

acte

ria; f

__Co

rioba

cter

iace

ae_O

TU_1

0Ba

cter

oide

tes;

Bac

tero

ides

Firm

icute

s; R

umin

ococ

cus

Firm

icute

s; S

arcin

aFi

rmicu

tes;

Stre

ptoc

occu

sBa

cter

oide

tes;

[Pre

vote

lla]

Bact

eroi

dete

s; o

__Ba

cter

oida

les_

OTU

_4Fi

rmicu

tes;

02d

06Pr

oteo

bact

eria

; Acin

etob

acte

rFi

rmicu

tes;

Osc

illosp

iraFi

rmicu

tes;

Lac

toco

ccus

Firm

icute

s; p−7

5−a5

Firm

icute

s; S

MB5

3Fi

rmicu

tes;

Vei

llone

llaFi

rmicu

tes;

Meg

asph

aera

Firm

icute

s; M

itsuo

kella

Prot

eoba

cter

ia; f

__En

tero

bact

eria

ceae

_OTU

_6Ba

cter

oide

tes;

Pre

vote

llaFi

rmicu

tes;

Lac

toba

cillu

s




colostrum Mid Colon


sow milk Mid Colon

colostrum Stomach


sow milk Stomach

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Dist

al S

mal

l Int

estin

eM

id C

olon

Stom

ach

66.2

5.4

4.8

1.23 08.3 0 07.4

8.6

9.2

0.1

3.700.3

0.1

13.4 0

09.6

34.8

0.2

0.1

0.7

1.9

4.8 00.2

5.1

1.3

0.1 325.5

0.8

1.9 0 00

0.3

0.4

1.2

0.5011.6

0.8 02.8

4.401.6

0.164 1.9 00 0 0.20

0.1

0.1 0 40 13.4

1.5 01.4

8.8

0.42 0 0.1

2.4 00.7

3.8

1.3

11.9

4.8

3.7

0.4

1.4

8.1

0.2 0 2.1 00.4 00.6

0.1

5.2

1.3

0.7

0.9 00.1

4.3

0.1

0.1

2.8

5.8 400.1

1.9

1.7 0014.1

2.1

1.5

16.6

0.2 00.1 54.4

6.2

0.2

1.6

2.3

0.1

0.4

0.2

0.2

16.4

14.8 011 0.1016.5

0.7

0.9

0.2

1.6

0.8

11.3

0.1

2.9

5.4

0.8

0.1

0.21 00.5

1.3

3.2

0.2

3.5

1.6

0.8

37.5

0.1

1.4

0.5

0.701.4

0.4 0 0 0015.8

9.3 053.4

0.3

0.7

2.80 00.3

3.2

0.1

Firm

icute

s; B

acillu

sAc

tinob

acte

ria; f

__Co

rioba

cter

iace

ae_O

TU_1

0Ba

cter

oide

tes;

Bac

tero

ides

Firm

icute

s; R

umin

ococ

cus

Firm

icute

s; S

arcin

aFi

rmicu

tes;

Stre

ptoc

occu

sBa

cter

oide

tes;

[Pre

vote

lla]

Bact

eroi

dete

s; o

__Ba

cter

oida

les_

OTU

_4Fi

rmicu

tes;

02d

06Pr

oteo

bact

eria

; Acin

etob

acte

rFi

rmicu

tes;

Osc

illosp

iraFi

rmicu

tes;

Lac

toco

ccus

Firm

icute

s; p−7

5−a5

Firm

icute

s; S

MB5

3Fi

rmicu

tes;

Vei

llone

llaFi

rmicu

tes;

Meg

asph

aera

Firm

icute

s; M

itsuo

kella

Prot

eoba

cter

ia; f

__En

tero

bact

eria

ceae

_OTU

_6Ba

cter

oide

tes;

Pre

vote

llaFi

rmicu

tes;

Lac

toba

cillu

s




colostrum Mid Colon


sow milk Mid Colon

colostrum Stomach


sow milk Stomach

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Dist

al S

mal

l Int

estin

eM

id C

olon

Stom

ach

66.2

5.4

4.8

1.23 08.3 0 07.4

8.6

9.2

0.1

3.700.3

0.1

13.4 0

09.6

34.8

0.2

0.1

0.7

1.9

4.8 00.2

5.1

1.3

0.1 325.5

0.8

1.9 0 00

0.3

0.4

1.2

0.5011.6

0.8 02.8

4.401.6

0.164 1.9 00 0 0.20

0.1

0.1 0 40 13.4

1.5 01.4

8.8

0.42 0 0.1

2.4 00.7

3.8

1.3

11.9

4.8

3.7

0.4

1.4

8.1

0.2 0 2.1 00.4 00.6

0.1

5.2

1.3

0.7

0.9 00.1

4.3

0.1

0.1

2.8

5.8 400.1

1.9

1.7 0014.1

2.1

1.5

16.6

0.2 00.1 54.4

6.2

0.2

1.6

2.3

0.1

0.4

0.2

0.2

16.4

14.8 011 0.1016.5

0.7

0.9

0.2

1.6

0.8

11.3

0.1

2.9

5.4

0.8

0.1

0.21 00.5

1.3

3.2

0.2

3.5

1.6

0.8

37.5

0.1

1.4

0.5

0.701.4

0.4 0 0 0015.8

9.3 053.4

0.3

0.7

2.80 00.3

3.2

0.1

Firm

icute

s; B

acillu

sAc

tinob

acte

ria; f

__Co

rioba

cter

iace

ae_O

TU_1

0Ba

cter

oide

tes;

Bac

tero

ides

Firm

icute

s; R

umin

ococ

cus

Firm

icute

s; S

arcin

aFi

rmicu

tes;

Stre

ptoc

occu

sBa

cter

oide

tes;

[Pre

vote

lla]

Bact

eroi

dete

s; o

__Ba

cter

oida

les_

OTU

_4Fi

rmicu

tes;

02d

06Pr

oteo

bact

eria

; Acin

etob

acte

rFi

rmicu

tes;

Osc

illosp

iraFi

rmicu

tes;

Lac

toco

ccus

Firm

icute

s; p−7

5−a5

Firm

icute

s; S

MB5

3Fi

rmicu

tes;

Vei

llone

llaFi

rmicu

tes;

Meg

asph

aera

Firm

icute

s; M

itsuo

kella

Prot

eoba

cter

ia; f

__En

tero

bact

eria

ceae

_OTU

_6Ba

cter

oide

tes;

Pre

vote

llaFi

rmicu

tes;

Lac

toba

cillu

s




colostrum Mid Colon


sow milk Mid Colon

colostrum Stomach


sow milk Stomach

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Dist

al S

mal

l Int

estin

eM

id C

olon

Stom

ach

66.2

5.4

4.8

1.23 08.3 0 07.4

8.6

9.2

0.1

3.700.3

0.1

13.4 0

09.6

34.8

0.2

0.1

0.7

1.9

4.8 00.2

5.1

1.3

0.1 325.5

0.8

1.9 0 00

0.3

0.4

1.2

0.5011.6

0.8 02.8

4.401.6

0.164 1.9 00 0 0.20

0.1

0.1 0 40 13.4

1.5 01.4

8.8

0.42 0 0.1

2.4 00.7

3.8

1.3

11.9

4.8

3.7

0.4

1.4

8.1

0.2 0 2.1 00.4 00.6

0.1

5.2

1.3

0.7

0.9 00.1

4.3

0.1

0.1

2.8

5.8 400.1

1.9

1.7 0014.1

2.1

1.5

16.6

0.2 00.1 54.4

6.2

0.2

1.6

2.3

0.1

0.4

0.2

0.2

16.4

14.8 011 0.1016.5

0.7

0.9

0.2

1.6

0.8

11.3

0.1

2.9

5.4

0.8

0.1

0.21 00.5

1.3

3.2

0.2

3.5

1.6

0.8

37.5

0.1

1.4

0.5

0.701.4

0.4 0 0 0015.8

9.3 053.4

0.3

0.7

2.80 00.3

3.2

0.1

Firm

icute

s; B

acillu

sAc

tinob

acte

ria; f

__Co

rioba

cter

iace

ae_O

TU_1

0Ba

cter

oide

tes;

Bac

tero

ides

Firm

icute

s; R

umin

ococ

cus

Firm

icute

s; S

arcin

aFi

rmicu

tes;

Stre

ptoc

occu

sBa

cter

oide

tes;

[Pre

vote

lla]

Bact

eroi

dete

s; o

__Ba

cter

oida

les_

OTU

_4Fi

rmicu

tes;

02d

06Pr

oteo

bact

eria

; Acin

etob

acte

rFi

rmicu

tes;

Osc

illosp

iraFi

rmicu

tes;

Lac

toco

ccus

Firm

icute

s; p−7

5−a5

Firm

icute

s; S

MB5

3Fi

rmicu

tes;

Vei

llone

llaFi

rmicu

tes;

Meg

asph

aera

Firm

icute

s; M

itsuo

kella

Prot

eoba

cter

ia; f

__En

tero

bact

eria

ceae

_OTU

_6Ba

cter

oide

tes;

Pre

vote

llaFi

rmicu

tes;

Lac

toba

cillu

s




colostrum Mid Colon


sow milk Mid Colon

colostrum Stomach


sow milk Stomach

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

BC

MR

SM

B

C

M

R

S

M

BC

MR

SM

Act

inob

acte

ria; f

amily

Cor

ioba

cter

iace

ae O

TU 1

0

Bac

tero

idet

es; o

rder

Bac

tero

idal

es O

TU 4

Pro

teob

acte

ria; f

amily

Ent

erob

acte

riace

ae O

TU 6

Sto

mac

hD

ista

l sm

all i

ntes

tine

Mid

col

on

Stom

ach

Dist

al S

mal

l Int

estin

eM

id C

olon

125

250

375

500

625

750

875

1000

1125

colos

trum

milk re

place

rso

w milk

colos

trum

milk re

place

rso

w milk

colos

trum

milk re

place

rso

w milk

Diet

Estimated number of OTUs Estimated number of OTUs

BC

MR

S

M

BC

MR

S

M

Die

t

BC

MR

S

M

Stom

ach

Dis

tal s

mal

l int

estin

e M

id c

olon

Stom

ach

Dis

tal S

mal

l Int

estin

eM

id C

olon

0.50

0.75

1.00

1.25

1.50

1.75

2.00

2.25

2.50

2.75

3.00

3.25

3.50

3.75

4.00

4.25

4.50

4.75

5.00

colos

trum

milk

repla

cer

sow

milk

colos

trum

milk

repla

cer

sow

milk

colos

trum

milk

repla

cer

sow

milk

Die

t

Diversity index Diversity index

BC

M

R

SM

B

C

MR

S

M

Die

t

BC

M

R

SM

Sto

mac

hD

ista

l sm

all i

ntes

tine

Mid

col

on

(A)

(B)

(C)

(D)

Figu

re 3


Firmicutes; Leuconostoc; OTU_209

Firmicutes; Megasphaera; OTU_1106

Firmicutes; f__[Mogibacteriaceae]_OTU_103; OTU_103

Firmicutes; Lactobacillus; OTU_58



Firmicutes; Lactococcus; OTU_31

Bacteroidetes; Prevotella; OTU_152


0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)

Firmicutes; family [Mogibacteriaceae] OTU_103

Firmicutes; Oscillospira; OTU_3

Actinobacteria; Collinsella; OTU_79


Firmicutes; Mitsuokella; OTU_1282



0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)

Figure 4










0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)








0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)










0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)








0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)










0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)








0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)


Firmicutes; Dorea; OTU_207


Firmicutes; Weissella; OTU_880


Firmicutes; Geobacillus; OTU_220


Firmicutes; Streptococcus; OTU_163



Firmicutes; Anoxybacillus; OTU_83

Proteobacteria; f__Enterobacteriaceae_OTU_6; OTU_6

0.01 0.10 1.00 10.00Read Abundance (%)

Diet

milk replacer

sow milk

Distal small intestine

Read abundance (%)

Proteobacteria; family Enterobacteriaceae OTU_6

Firmicutes; Bulleidia; OTU_505



Firmicutes; Anoxybacillus; OTU_83

Firmicutes; Carnobacterium; OTU_670

Firmicutes; Streptococcus; OTU_703






0.01 0.10 1.00 10.00Read Abundance (%)

Diet

colostrum

sow milk


Read abundance (%)

−5.0

−2.5

0.0

2.5

−5.0 −2.5 0.0 2.5 5.0PC1 [26%]

PC2

[15.

8%] Diet

colostrum

milk replacer

sow milk


PC

2 - v

aria

tion

expl

aine

d 15

.8%


Distal small intestine(A)

(B)

(C)

Enterobacteriaceae Lactobacillus

Lactococcus

Figure 5










0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)








0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)Firmicutes; Leuconostoc; OTU_209









0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)








0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)










0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)








0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)










0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)








0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)


Firmicutes; o__Clostridiales_OTU_1273; OTU_1273


Deferribacteres; Mucispirillum; OTU_462

Firmicutes; f__Lachnospiraceae_OTU_603; OTU_603


Firmicutes; Ruminococcus; OTU_11


Actinobacteria; f__Coriobacteriaceae_OTU_41; OTU_41

Firmicutes; [Ruminococcus]; OTU_128



Firmicutes; Blautia; OTU_305

Firmicutes; Blautia; OTU_47

0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Mid colon

Read abundance (%)

Actinobacteria; family Coriobacteriaceae OTU_41

Firmicutes; family Lachnospiraceae OTU_603

Firmicutes; order Clostridiales OTU_1273

Spirochaetes; Sphaerochaeta; OTU_295

Firmicutes; c__Firmicutes_OTU_85; OTU_85


Firmicutes; f__Ruminococcaceae_OTU_119; OTU_119




Firmicutes; f__Christensenellaceae_OTU_17; OTU_17


Fusobacteria; Fusobacterium; OTU_7

Proteobacteria; Campylobacter; OTU_2348




0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Mid colon

Read abundance (%)

Firmicutes; family Ruminococcaceae OTU_21

Firmicutes; family Christensenellaceae OTU_17




Firmicutes; OTU_85

−5.0

−2.5

0.0

2.5

5.0

−6 −3 0 3PC1 [13.3%]

PC3

[10%

] Diet

colostrum

milk replacer

sow milk

Mid colon

PC

3 - v

aria

tion

expl

aine

d 10

.0%


Mid colon

(A)

(B)

(C)

Figure 6










0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)








0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)










0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)








0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)










0.01 0.10 1.00 10.00Read Abundance (%)

Dietmilk replacer

sow milk

Stomach

Read abundance (%)








0.01 0.10 1.00 10.00Read Abundance (%)

Dietcolostrum

sow milk

Stomach

Read abundance (%)

−3

0

3

6

−5.0 −2.5 0.0 2.5 5.0PC1 [26.4%]

PC2

[17.

2%] Diet

colostrum

milk replacer

sow milk

StomachStomach

PC

2 - v

aria

tion

expl

aine

d 17

.2%


(A)

(B)

(C)


−10

−50

5

−6−4−2024

RDA

1

RDA2

colo

stru

m

milk

repl

acer

sow

milk

Dis

tal S

mal

l Int

estin

e

Faec

es

Mid

Col

on

Stom

ach

Ace

Pro

Iso_

But

But

Iso_

Val

Val

Ibiv

Apb

Ente

ro

Hem

Clo

sLa

b

Yeas

t

Ana

RDA2

RD

A1

Sto

mac

h D

ista

l sm

all i

ntes

tine

Mid

col

on

Faec

es

Col

ostru

m

Milk

repl

acer

S

ow m

ilk

−10

−50

5

−6−4−2024

RDA

1

RDA2

colo

stru

m

milk

repl

acer

sow

milk

Dis

tal S

mal

l Int

estin

e

Faec

es

Mid

Col

on

Stom

ach

Ace

Pro

Iso_

But

But

Iso_

Val

Val

Ibiv

Apb

Ente

ro

Hem

Clo

sLa

b

Yeas

t

Ana

−10

−50

5

−6−4−2024

RDA

1

RDA2

colo

stru

m

milk

repl

acer

sow

milk

Dis

tal S

mal

l Int

estin

e

Faec

es

Mid

Col

on

Stom

ach

Ace

Pro

Iso_

But

But

Iso_

Val

Val

Ibiv

Apb

Ente

ro

Hem

Clo

sLa

b

Yeas

t

Ana

−10

−50

5

−6−4−2024

RDA

1

RDA2

colo

stru

m

milk

repl

acer

sow

milk

Dis

tal S

mal

l Int

estin

e

Faec

es

Mid

Col

on

Stom

ach

Ace

Pro

Iso_

But

But

Iso_

Val

Val

Ibiv

Apb

Ente

ro

Hem

Clo

sLa

b

Yeas

t

Ana

−10

−50

5

−6−4−2024

RDA

1

RDA2

colo

stru

m

milk

repl

acer

sow

milk

Dis

tal S

mal

l Int

estin

e

Faec

es

Mid

Col

on

Stom

ach

Ace

Pro

Iso_

But

But

Iso_

Val

Val

Ibiv

Apb

Ente

ro

Hem

Clo

sLa

b

Yeas

t

Ana

−10

−50

5

−6−4−2024

RDA

1

RDA2

colo

stru

m

milk

repl

acer

sow

milk

Dis

tal S

mal

l Int

estin

e

Faec

es

Mid

Col

on

Stom

ach

Ace

Pro

Iso_

But

But

Iso_

Val

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Figure S1



Chapter 5

Manuscript 2

Investigating the potential effects of alpha-(1,2)-fucosyltransferase geno-type on the establishment and succession of the gastrointestinal micro-biota of young piglets.

Ann-Sofie Riis Poulsen, Sugiharto Sugiharto, Nuria Canibe and Charlotte Laurid-sen.

Prepared for submission to Veterinary Microbiology.



17 piglets from two sowsFollowed from 5 to 34 days of age

FUT1-307AA piglets= E. coli F18 susceptible

Faeces Day 5, 7, 14, 21,

28 and 34


FUT1-307AG piglets= E. coli F18 resistant


Faeces Day 5, 7, 14, 21,

28 and 34

10 piglets 7 piglets

1. 16S rRNA gene sequencing

• V3 region• Ion Torrent



! 1

Investigating the potential effects of alpha-(1,2)-fucosyltransferase genotype on the

establishment and succession of the gastrointestinal microbiota of young piglets

Ann-Sofie Riis Poulsena, Sugiharto Sugihartob, Nuria Canibea, Charlotte Lauridsena*

a Aarhus University, Faculty of Science and Technology, Department of Animal Science, Blichers

Allé 20, 8830,Tjele, Denmark

b Diponegoro University, Faculty of Animal and Agricultural Sciences, Semarang, Central Java,

Indonesia

* Corresponding author

Aarhus University,


Blichers Allé 20, P.O. Box 50


Tel.: +45 87 15 80 19

Fax.: +45 87 15 42 49



! 2

Abstract

Piglets with the alpha-(1,2)-fucosyltransferase (FUT1) genotype M307A/A have decreased

susceptibility to enterotoxigenic Escherichia coli F18 infection. The effect of FUT1 polymorphisms

on the establishing gut microbiota, however, is unknown. This study aimed at characterising the

gastrointestinal microbiota of FUT1-M307A/A and FUT1-M307A/G piglets and the influence of age

using high-throughput sequencing and classical culture techniques.

Ten sensitive (M307G/A) and seven resistant (M307A/A) piglets were included. Piglets suckled

their mother-sow from birth until weaning at 28 days of age. Faecal samples collected on day 5, 7,

14, 21, 28 and 34 of age were analysed using 16S rRNA gene sequencing. All piglets were

euthanized at 34 days of age. Bacterial enumerations by culture and short-chain fatty acid analysis

were performed on faeces and digesta. Multivariate statistics showed similar faecal communities

between genotypes. A numerical higher relative abundance of Prevotella was observed in sensitive

piglets at 7, 14, 21 and 28 days of age, though. Bacterial enumeration by plating showed a higher

number of haemolytic bacteria in faeces (p=0.003) and digesta (p=0.02) in 34 days old sensitive

piglets compared to resistant piglets. The colonic acetic acid concentration was higher in sensitive

piglets (p=0.01). Age was found to have an effect on the faecal communities (padonis=0.001). In

conclusion, faecal microbiota of sensitive piglets was significantly different from that of resistant

piglets in the number of potentially pathogenic E. coli. However, 16S rRNA gene sequencing also

indicated an effect of FUT1 genotype on Prevotella. Our data do not allow us to definitively rule

out an effect of FUT1 genotype on other bacterial groups in the gastrointestinal tract as 16S rRNA

gene sequencing was only performed on faecal samples.

Keywords: Piglets. Gastrointestinal microbiota. 16S rRNA amplicon sequencing. Alpha-(1,2)-

fucosyltransferase (FUT1). Escherichia coli F18. Post-weaning diarrhoea.


! 3

Introduction

Diarrhoea is a common clinical condition in pigs post-weaning (Lalles et al., 2007). The vast

majority of pigs having diarrhoea are treated with antibiotics, and in Denmark clinical diarrhoea is

the main cause of antibiotic treatment of pigs (DANMAP, 2014). The growing concern over

antibiotic resistance has increased the demand for alternative therapeutic or preventative options

and underscores the need for improving the general health condition of pigs.

The gut microbiota is a complex micro-environment existing in symbiosis with the host and

recognised as a major player in host health (Sommer and Baeckhed, 2013). Microbial gut

colonisation of the newborn is essential in stimulating local immune cell proliferation and is critical

to the maturation of the adaptive immune system (Chung et al., 2012; Guarner and Malagelada,

2003). Microbial gut colonisation is a gradual and complex process starting at birth (Mackie et al.,

1999). During early life the microbiota is characterised by instability, resulting in a highly dynamic

community with frequent changes in dominating taxa (Petri et al., 2010; Slifierz et al., 2015). The

developing gut microbiota is easily disturbed and in pigs, microbial imbalances due to e.g. weaning

stress (Konstantinov et al., 2006; Thompson et al., 2008) are believed to affect host-defense

mechanisms and increase susceptibility to enteric disease and hence, diarrhoea (Castillo et al., 2007;

Heo et al., 2013). In relation to weaning, enterotoxigenic Escherichia coli (ETEC) F18 is an

important pathogen causing infectious diarrhoea during the first week post-weaning (Fairbrother et

al., 2005). Breeding ETEC F18-resistant piglets has been studied as a potential preventative strategy

in coping with post-weaning diarrhoea (Frydendahl et al., 2003). Host genetics, however, has been

suggested as one of the main factors influencing gut microbiota composition (Scott et al., 2013).

Different breeds of pigs have been reported to harbor different gut microbial communities (Pajarillo

et al., 2014b) and in chickens, different genetic variants of disease-determining genes have been

found to influence the gut colonisation process of the young animal (Schokker et al., 2015).

The FUT1 gene, encoding the alpha-(1,2)-fucosyltransferase enzyme, has been proposed as

a candidate gene for manipulating adhesion of ETEC F18 to its intestinal receptor (Meijerink et al.,

1997). It has been confirmed that a guanine-to-adenine mutation at nucleotide 307 (M307) in both

loci of the FUT1 gene does result in resistance to ETEC F18 adherence to small intestinal

enterocytes (Meijerink et al., 2000). Furthermore, the M307A/A genotype has been associated with

decreased susceptibility of weanling piglets to ETEC F18 infection compared to piglets with

genotypes M307A/G and M307G/G (Frydendahl et al., 2003).


! 4

As ETEC F18-resistant and sensitive piglets are genetically different and the resistance

mechanism is believed to involve intestinal receptors, FUT1 polymorphisms might influence the

establishment of the gut microbiota of young piglets beyond the establishment of ETEC F18. As the

gut microbiota is highly important to host health, this knowledge is of great relevance. Hence, the

primary aim of the present study was to investigate how FUT1 genotype influences the

establishment of the gastrointestinal microbiota in piglets aged 5 to 34 days combining high-

throughput next generation sequencing and classical culture techniques. A secondary aim was to

investigate how the microbiota changes with age. We hypothesise that piglets with the FUT1-

M307A/A genotype will have a gut microbiota different from piglets with the FUT1-M307A/G

genotype and that the gut microbial community will change as the piglet ages.


Animals, study design and FUT1 genotyping

The present study was conducted according to the license obtained from the Danish Animal


administration. The study was performed at the experimental facility at the Department of Animal

Science (Foulum, Aarhus University). Sows and piglets were housed according to Danish

guidelines.

A total of 17 vaginally born piglets were included in the study. They were the result of

breeding two M307G/A sows with the same M307A/A boar (Danish Landrace x Yorkshire breed x

Duroc). Only FUT1-M307G/A and FUT1- M307A/A piglets were included in the study because FUT1-

M307G/A and FUT1-M307G/G piglets were considered equally susceptible to ETEC F18 (Frydendahl

et al., 2003). An ear tissue sample was collected shortly after birth and used for FUT1 genotyping.

In short, the genotyping procedure included a PCR amplification of the FUT1 gene product

harboring the mutation of interest. Subsequently, the PCR fragments were digested with the HinP1I

restriction enzyme and run on a 1.5% agarose gel.

All piglets were kept with their mother-sow until weaning at 28 days of age. Piglets suckled

their dams and had free access to water and creep feed (no added zinc oxide) from three weeks of

age. After weaning, piglets were housed together in pens according to litter and fed a standard post-

weaning diet (no added zinc oxide). The temperature in the weaning unit was 23°C and piglets had

free access to rooting material.


! 5

Sample collection

Faecal samples were collected directly from the rectum at 5, 7, 14, 21, 28 and 34 days of age.

Subsamples were stored at -20˚C for organic acid analysis and snap-frozen in liquid nitrogen and

stored at -80˚C for 16S rRNA amplicon sequencing. Bacterial enumerations by plating were

performed on a subsample of fresh faeces.

All piglets were euthanised at 34 days of age. The abdomen was incised and the

gastrointestinal tract removed. Luminal content (digesta) from stomach, distal small intestine,

caecum and middle part of the colon was sampled immediately and subsamples were stored at -

20˚C for organic acid analysis. Bacterial enumerations by plating were performed on fresh

subsamples.

DNA extraction

DNA was extracted using the MOBIO Power Soil DNA Isolation Kit (MOBIO Laboratories Inc.,

Carlsbad, CA) including an initial enzymatic treatment and otherwise following a standard protocol.

Extracted DNA concentrations were quantified fluorometrically with Qubit 3.0 HS dsDNA assay

(Life Technologies, Thermo Fisher Scientific, USA) and normalized to 5 ng/µL.

16S rRNA amplicon sequencing

Partial 16S rRNA sequences were amplified (at GenoSkan A/S, Tjele, Denmark) from extracted

DNA using ProBio_Uni/Probio_Rev primers (Milani et al., 2013), amplifying the V3 region of the

16S rRNA gene sequence. To establish the amplicon library, a PCR was conducted using the

following thermocycler settings: Initial denaturation at 95°C for 5 min, 35 cycles at 94°C for 30 s,

58°C for 30 s, and 72°C for 90 s, and final elongation at 72°C for 10 min. Individual amplicon

libraries were purified using the Agencourt AMPure XP DNA purification beads (Beckman Coulter

Genomics GmbH, Germany) and the Ion OneTouchTM 200 template Kit v2 DL (Life

Technologies) was employed for emulsion PCR according to the manufacturers recommendations.

Amplicon libraries were sequenced using the Ion Torrent PGM system applying the Ion Sequencing

200 kit (Life Technologies) according to the manufacturer’s recommendations. All PGM quality-

approved, trimmed and filtered data were exported as fastq files. Sequences were filtered using the

FastQ Quality software and sequences with length 150-220 bp and an average quality score above

19 were used for further analysis.


! 6

Amplicon bioinformatics processing

Bioinformatic processing was done using a pipeline from The Brazilian Microbiome Project for Ion

Torrent 16S rRNA sequence analysis (Pylro et al., 2014), UPARSE (USEARCH 8.1.1756) (Edgar,

2013) and QIIME (Caporaso et al., 2010). Reads were quality filtered using USEARCH –

fastq_filter with a maximum expected error of 1.0 and global trimming at bp 166 was performed in

the same step. Reads were dereplicated (i.e. identification of unique sequences) and formatted to the

UPARSE pipeline. Singletons were discarded and dereplicated reads were clustered into

Operational Taxonomical Units (OTUs) using USEARCH at 97% sequence similarity. An

additional chimera detection step was included using USEARCH with rpd_gold version 9 (Cole et

al., 2014) as reference database. Taxonomy was assigned using the UCLUST-classifier as

implemented in QIIME (Caporaso et al., 2010) with Greengenes version 13.8 (DeSantis et al.,

2006) as a reference database and default settings. Results were analysed in R studio (version

0.99.489 for Mac) using the Ampvis package (Albertsen et al., 2015).

Dry matter and organic acid analysis

Dry matter content of digesta was determined by freeze-drying (ScanVac Coolsafe 55, Labogene

ApS, Lynge, Denmark). The concentration of short-chain fatty acids and lactic acid in faeces and

digesta were measured as previously described by Canibe et al. (2007).

Microbiological enumerations by plating

Faecal samples (approximately 1 g) were transferred to plastic bags and 5 or 10 ml pre-reduced salt

medium was added. The content was homogenised in a Smasher paddle blender (bioMérieux

Industry, USA) for 2 minutes. Digesta samples (approximately 5 g) were transferred to flasks

containing 50 ml pre-reduced salt medium (Holdeman et al., 1977). The flask content was

transferred to a CO2 flushed bag and homogenized in a stomacher blender for 2 minutes. 1 ml


fold dilutions were prepared using the technique previously described by Miller and Wolin (1974).

Each sample was plated on selective (and indicative) and non-selective agar plates.


incubation for 1 day. Yeasts were enumerated on Malt, Chlortetracycline and -Amphenicol agar

(Merck 1.03753 (yeast extract), 1.05397 (malt extract), 1.07224 (bacto-pepton), 1.08337 (glucose),

1.01614 (agar-agar) and Oxoid Sr0177E) after aerobic incubation for 2 days. Haemolytic bacteria


! 7

were enumerated on blood agar (Oxoid Pb5039A) after aerobic incubation for 1 day. Clostridium

perfringens was enumerated using the pour-plate technique on Tryptose Sulfite Cycloserine agar

(Merck 1.11972 and 1.00888) after anaerobic incubation for 1 day. Lactic acid bacteria were

enumerated on de Man, Rogosa and Sharp agar (Merck 1.10660) after anaerobic incubation for 2

days. Total anaerobic bacteria were enumerated in roll tubes containing pig colon fluid-glucose-

cellobiose agar (Holdeman et al., 1977) and incubated for 7 days. All plates and roll-tubes were

incubated at 37˚C.

Porcine Intestinal Organ Culture (PIOC)

The PIOC procedure was included to investigate the adherence of E. coli O138:F18 to the mucosa

of resistant and sensitive piglets following the procedure previously described by Sugiharto et al.

(2012). In short, a distal small intestinal segment (15 cm) was sampled and incubated with a

solution containing E. coli O138:F18 for 1 hour. The tissue was then homogenized, diluted and E.

coli enumerated on MacConkey (Merck 1.05465) agar plates after aerobic incubation at 37˚C

overnight.

Statistical analyses

The effect of genotype and age on bacterial and organic acid parameters, pH, microbial community

richness and diversity and animal body weight were analysed by fitting the data to a linear mixed

model using the lmer function from the lme4 package (Bates, 2014) using R studio (Version

0.99.489 for Mac). Genotype and age/intestinal segment were included as fixed effects, while pig

and sow were included as random effects (by including random intercept terms) to account for

multiple observations within the same litter or within the same pig. When analysing the weight

variable, birth weight was included as co-variate. The fixed effects were tested using a F-test with

Kenward-Roger approximation, where the reduced model was tested against the full model. This

was done using the KRmodcomp function in the pbkrtest package (Halekoh, 2014). When age was

found to have a significant effect, a post-hoc test was performed using the multcomp package and

Bonferroni adjustment to correct for multiple comparisons (Hothorn, 2008). Effects were

considered significant when p<0.05 and as trends when 0.05≤p<0.10. Principal Component

Analysis (PCA) was performed on square root transformed OTU abundances. Significance of

genotype and age was tested on the first two principal components (PCs) using the envfit parametric

test and on the Bray-Curtis dissimilarity matrix using adonis (Oksanen, 2015). The parametric


! 8

Wald-test (Love et al., 2014) was used to test for significant differences in OTUs abundances

between genotypes and between the ages 5 and 28, and 28 and 34 days. OTUs with an adjusted

p<0.001 were considered significantly different between genotypes and/or ages.

Results

Genetic testing revealed that ten piglets were of the sensitive genotype (M307G/A) and seven piglets

were of the resistant genotype (M307A/A). Body weight data is presented in Table 1. Sensitive

piglets weighed significantly more than resistant piglets at day 28 (p=0.003) and 34 (p=0.01) of age.

Composition of faecal microbiota– 16S rRNA sequencing

Sequencing of 74 samples (sensitive n=39; resistant n=35) yielded a total of 6,060,572 sequences

after removal of poor quality reads. Due to the small size of the piglets, 28 samples are missing due

to either unsuccessful sampling or sampling an amount too small to allow successful DNA

extraction. Furthermore, nine of the sequenced samples had less than 5000 sequences and were

excluded from further analyses. Recovered sequences clustered into 2993 OTUs, which were

classified into 15 bacterial phyla, 74 families and 112 genera. Nine phylum level groups had an

overall relative abundance ≥1% (found in at least one of the two genotypes at each age), of which

two groups were taxonomically unassigned (OTU 57 and OTU 188) (Fig. 1a).

Firmicutes, followed by Bacteroidetes, dominated the communities irrespective of FUT1

genotype and age. The remaining phyla were present at frequencies lower than 6% in both

genotypes at all ages. The five most abundant families were Ruminococcaceae, Clostridiaceae,

Lactobacillaceae, Prevotellaceae and Lachnospiraceae (Fig. 1b). A decrease in relative abundance

was observed from day 7 for Lactobacillaceae and an increase on day 34 for Lachnospiraceae in

both genotypes. At the genus level, numerical differences (at least a doubling in the relative

abundance and at least one genotype having a relative abundance above 5%) between genotypes

were seen for Lactobacillus on day 7, Prevotella on day 7, 14, 21 and 28, Clostridium on day 5 and

14, Bacteroides on day 14, and Butyricimonas on day 7 (Fig S2). Based on the first two PCs of the

OTU abundances, the FUT1 genotype had no significant effect on the overall faecal microbial

community (Fig. 2, padonis>0.27), and accordingly no OTUs differed significantly between

genotypes. Species richness (Fig. S1 a and b; observed p=0.73; estimated p=0.75) and Shannon

diversity index (Fig. S1 c; p=0.11) were not significantly different between genotypes.


! 9

Based on the first and third PC of the OTU abundances, a significant effect of age on the

overall microbial community was observed (Fig. 3; padonis=0.001). A gradual change in the overall

bacterial community composition was seen from day 5 to 34 of age. Pairwise comparisons between

all sampling days revealed that the microbial community richness was higher on day 21 (observed

p=0.004; estimated p=0.003) and 28 (observed p=0.03; estimated p=0.01) compared to day 5 (Fig.

S1 a and b). Age did not have an effect on the Shannon diversity index. Comparing day 5 and day

28 (the first and last sampling day during suckling), 201 out of 1769 OTUs had significantly

different read abundances (Fig. 4a: the 25 most significantly different OTUs). The most

significantly different OTU (the OTU with the lowest p-value) was OTU_9 (p=3.3E-26) belonging

to the family Ruminococcaceae, having a higher read abundance on day 5. Of the 25 most

significantly different OTUs, 12 belonged to the order Clostridiales with 10 having a lower read

abundance on day 28 compared to day 5. When comparing day 28 and day 34 (weaning day and

one week later), 105 out of the 1790 OTUs differed significantly (Fig. 4b; the 25 most significantly

different OTUs). OTU_28, belonging to the order Clostridiales, was the most significantly different

OTU (p=6.1E-26), having a higher read abundance on day 28. Of the 25 most significantly changed

OTUs, 11 belonged to the order Clostridiales with nine having a lower read abundance on day 34.

Six OTUs belonged to the family Ruminococcaceae and had a lower read abundance on day 34.

Litter was found to have a significant effect on the overall microbial community on day 21

(padonis=0.01).

Microbiological enumerations by plating and organic acid concentrations

Due to the small size of the piglets, adequate quantities of faeces for bacterial enumeration by

plating and organic acid analysis could not be sampled from all animals at all ages.

Faeces. Higher faecal numbers of haemolytic bacteria at 34 days of age (p=0.003), and a tendency

to higher numbers of lactic acid bacteria (p=0.06), were observed in sensitive piglets (Table 2). The

number of haemolytic bacteria was constant between day 5 and 28, whereas a significant increase

was detected on day 34 (p≤0.003). An age-dependent decrease of C. perfringens was seen with

numbers being lowest on day 28 (p<0.0001) and 34 (p<0.0001). The same was observed for lactic

acid bacteria, with lower numbers on day 28 (p≤0.03) and day 34 (p≤0.005) compared to day 5 and

7. There was no significant effect of genotype or age on the number of Enterobacteriaceae or total

anaerobic bacteria.


! 10

Furthermore, no difference in the concentration of faecal organic acids between genotypes

was determined, except for a tendency to a lower concentration of butyric acid (p=0.09) in resistant

piglets (Table 3). The concentration of acetic acid was higher at 34 days of age (one week post-

weaning) compared to day 5, 7, 14 and 21 of age (suckling period) (p<0.05). The concentration of

propionic acid was constant between day 5 and 21, but at 34 days of age, a significant increase in

propionic acid was observed (p=0.0002). A tendency to a lower concentration of butyric acid

(p=0.09) with age was measured, whereas no effect of genotype or age was detected for valeric, iso-

butyric and iso-valeric acid in faeces (data not shown).

The age-dependent decrease in lactic acid bacteria and C. perfringens seen by culture were

in agreement with the results from the 16S rRNA gene sequencing. Both methods detected a

marked decrease in Clostridium (C. perfringens) from day 14. Regarding lactic acid bacteria, the

culture results were in accordance with the relative abundances of Lactobacillus as both methods

detected a decrease from day 7. In accordance with the 16S rRNA sequencing results, culture of

Enterobacteriaceae showed a higher number in resistant piglets at 5 and 7 days of age, while the

number was higher in sensitive piglets at 28 and 34 days of age. See Fig. 1b, Fig S2 and Table 2.

Digesta. Genotype had no significant effect on dry matter content or pH (Table S1). Sensitive

piglets had a higher number of Enterobacteriaceae (p≤0.02) in digesta from the distal small

intestine, caecum and mid colon; of haemolytic bacteria (p=0.02) in the stomach, small intestine,

caecum and colon; and of total anaerobic bacteria (p=0.004) in the distal small intestine compared

to resistant piglets (Table 4).

The concentration of acetic acid was higher in the colon of sensitive piglets (p=0.01, Table

5). Digesta concentrations of propionic, butyric, lactic, valeric, and the sum of iso-butyric and iso-

valeric acid were similar between genotypes.

Quantification of E. coli attachment to distal small intestinal tissue. The E. coli numbers in the

small intestinal tissue were similar between genotypes (p=0.41). However, sensitive piglets had

numerically higher E. coli numbers before (sensitive 6.3 log cfu/g; resistant 6.0 log cfu/g) and after

E. coli O138:F18 inoculation (sensitive 8.1 log cfu/g; resistant 7.8 log cfu/g).


! 11

Discussion

Newly weaned piglets are highly susceptible to enteric disease and diarrhoea caused by ETEC F18.

The FUT1 M307A/A genotype is believed to cause changes in the intestinal receptors responsible for

ETEC F18 adhesion (Meijerink et al., 1997); however, little scientific information is available on

the influence of FUT1 genotype on the gut microbiota of piglets beyond ETEC F18.

Applying 16S rRNA gene sequencing, the overall faecal microbial communities were found

to be similar between resistant and sensitive piglets. Bacterial culture showed a significant effect of

FUT1 genotype on the number of haemolytic bacteria after weaning, being higher in both faeces

and digesta from sensitive piglets. These results indicate that sensitive piglets have a higher

shedding of what could potentially be ETEC F18 in the post-weaning period and supports the

findings of Frydendahl et al. (2003), that FUT1 M307G/A piglets are indeed more susceptible to

ETEC F18 infection than are FUT M307A/A piglets. This is furthermore supported by an apparent

up-regulation in the humoral immune response in sensitive piglets. We were, however, not able to

detect significantly higher E. coli F18 numbers in the small intestinal tissue using an ex vivo porcine

intestinal model. As this ex vivo model has been successfully applied in a number of studies

(Sugiharto and Lauridsen, 2014; Sugiharto et al., 2015) and the fact that we in the present study

were able to detect numerical differences between genotypes, the lack of significance might be due

to the low number of replicates.

Ruminococcoceae, Clostridiaceae, Lactobacillaceae, Prevotellaceae and Lachnospiraceae

dominated the microbial community of ETEC F18-resistant and sensitive piglets in the present

study. Even though no OTUs were found to be significantly different between the two genotypes,

interesting trends were observed in especially Prevotella, as this genus was present in numerically

higher relative abundances in sensitive piglets at 7, 14, 21 and 28 days of age. Prevotella is

normally associated with the intake of fermentable fibers (Ivarsson et al., 2014) and its abundance

has been shown to increase after weaning (Pajarillo et al., 2014a) as piglets are normally weaned to

solid feed which, contrary to sow milk, is rich in fibers. These results are in contrast to the

observations in the present study, in which the relative abundance of Prevotella was highest before

weaning especially in sensitive piglets.

Mach et al. (2015) identified Prevotella and Ruminococcaceae as being two enterotype-like

clusters in piglets aged 14-70 days and reported that the body weight of piglets belonging to the

Prevotella cluster was lower during lactation but higher after weaning when compared to pigs

belonging to the Ruminococcaceae cluster. Bao et al. (2011) found ETEC F18-resistant Sutai pigs


! 12

to be marginally heavier at weaning, while Huang et al. (2008) found no effect of FUT1 genotype

on growth performance in Landrace and Duroc pigs. These observations do not agree with the

results of the present study, where sensitive piglets weighed significantly more than resistant piglets

on the day of weaning. It is unlikely that the higher body weight of sensitive piglets were due to the

higher fiber digestion capacity of Prevotella as piglets were not introduced to creep-feed until three

weeks of age. Furthermore, the limited intake of feed during the suckling period can probably not

explain the difference between genotypes on piglet weight gain until weaning. On the basis of data

from the present study, we were not able to suggest a mode of action of the effect of FUT1

genotype on growth parameters. It should be noted, however, that Prevotella is represented by

numerous species (Ley, 2016). As our data does not enable us to classify OTUs to species level, an

exact interpretation of the significance of the Prevotella genus is not possible.

Acetic acid, which is the major short-chain fatty acid in the gut, was found in higher

concentrations in colonic content of sensitive piglets. As the short-chain fatty acid profile reflects

bacterial metabolism (Williams et al., 2005), the higher colonic concentration of acetic acid in

sensitive piglets most likely reflect bacterial community differences between genotypes. Although a

higher number of Enterobacteriaceae was measured in sensitive piglets, and E. coli, belonging to

this family, has been shown to produce higher amounts of acetic acid at high growth rates (Han et

al., 1992), this alone is unlikely to explain the difference in acetic acid concentrations. Regardless

of genotype, the acetic acid concentration in the stomach was higher than anticipated. Also, a higher

concentration of butyric acid in the stomach than in the large intestine was unexpected, since the

opposite often is observed, e.g. as shown by Starke et al. (2014). An obvious explanation has not

been found.

In order to assess the age-dependent changes in the gut microbiota, faecal specimens were

used for 16S rRNA amplicon sequencing. Assessing the overall faecal microbiota, we found a

gradual change in the microbial community in the period from 5 to 34 days of age with an

increasing microbial diversity peaking at 21 days of age. When these results are compared with

previous studies, it is obvious that the dominating populations differ between studies. Frese et al.

(2015) investigated the effect of diet on the faecal microbiota in the pre- and post-weaning period

and found the dominating families to be Enterobacteriaceae, Lachnospiraceae, Bacteroidaceae,

Lactobacillaceae, Clostridiaceae, Ruminococcaceae and Prevotellaceae. In contrary to the results

of Frese et al. (2015) we did not find Enterobacteriaceae and Bacteroidaceae to be dominating.


! 13

Owing to differences between studies, such as housing conditions, dietary composition, in-feed

antibiotics, pig breed and analytical procedures, microbial community dissimilarities were expected.

As microbial species richness in the gut has been reported to increase with age (Slifierz et

al., 2015), and this study involved young piglets, the species richness, irrespective of age, was

higher than expected. As 16S rRNA sequence read length has been reported to influence the

observed microbial community richness (Engelbrektson et al., 2010), the high species richness is

most likely explained by the relatively short reads. This could also be the reason why numerous

OTUs were not taxonomically assigned, or only assigned to order level.

Gut microbiota composition varies between gastrointestinal region and location within the

gut (lumen vs. mucosa) (Looft et al., 2014). As the FUT1 gene product is believed to exert its effect

on ETEC F18 receptors in the small intestine (Meijerink et al., 1997), faecal samples might not be

the most appropriate sampling choice for in-depth microbial analysis. Supporting this, a recent

study from our group (Poulsen et al, in preparation) revealed a dietary treatment effect on the

microbial community composition of luminal contents in the small intestine, when analysed by 16S

rRNA gene sequencing, which was not detected in faecal samples. It could therefore be speculated

that different results would have been obtained if small intestinal luminal content and/or mucosal

samples rather than faecal samples had been analysed by 16S rRNA amplicon sequencing.

Conclusion

The data obtained from the present study contribute to the existing knowledge on FUT1

polymorphisms in pigs. In conclusion, the faecal microbiota of sensitive piglets significantly

differed from that of resistant piglets in only the number of potentially pathogenic ETEC, although

16S rRNA gene sequencing did indicate an effect of FUT1 genotype on Prevotella. Further studies

on the impact of FUT1 genotype on the microbial community composition in the small intestine are

needed, including absolute bacterial quantifications. The reported effect of FUT1 genotype on body

weight gain should also be investigated further.

Acknowledgements

The authors would like to thank laboratory technicians Trine Poulsen and Karin Durup, technicians

Mette Lykkegaard and Karin Johansen and the staff of SB32 at the Department of Animal Science

for their skillful technical assistance during the course of the experiment and Samantha Joan Noel

for proofreading the manuscript. The authors want to thank Lone Bruhn Madsen for FUT1


! 14

genotyping the animals. The authors furthermore want to thank Senior Scientist Emøke Bendixen,

Department of Molecular Biology and Genetics and collaborators at GenoSkan A/S, Niels

Pedersens Allé 2, DK-8830 Tjele. The paper drafting was primarily funded through a Ph.D.-

scholarship obtained from the Graduate School of Science and Technology, Aarhus University,

Denmark, for the main author. Funders had no influence on study design, collection, analysis or

interpretation of data, or drafting of manuscript.

Author contributions

Authors contributed as follows: A.-S.R.P. conducted the animal experiment, performed

bioinformatics and statistical data analyses and drafted the manuscript. S.S. conducted the animal

experiment and performed the PIOC experiment. N.C. designed the experiment, contributed in the

conduction of the animal experiment and assisted with the overall data analysis. C.L. designed the

experiment and contributed in the conduction of the animal experiment. All authors have revised

and accepted the final manuscript.

Conflict of interest statement

Conflicts of interest: None.


! 15

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Tables:

Table 1 Piglet weight (kg) at day 0, 7, 14, 21, 28 and 34 of age1

Days of age Genotype2 p-value

SENS RES G3 A4 G*A 0.002 <.0001 0.002

0 1.3 (0.0-4.4) 1.7 (0.0-4.1) 7 2.9 (0.0-5.4) 2.4 (0.0-4.8)

14 4.0 (1.0-7.1) 3.8 (1.3-6.2) 21 5.9 (2.9-9.0) 5.2 (2.7-7.6) 28 8.1a (5.1-11.3) 6.8b (4.3-9.2) 34 7.8a (4.7-10.9) 6.7b (4.2-9.1)

1 Values are presented as least square means and 95% confidence intervals (in parentheses). 2 SENS = M307GA; RES = M307AA. Number of piglets: SENS=10; RES=7. 3 G = Genotype. 4 A = Age. a,b Values with different superscripts within a row are significantly different (p<0.05).

! !


! 19

Table 2 Enumerations (log cfu/g sample) of selected microbial groups in faeces from piglets at 5, 7, 14, 21,

28 and 34 days of age1

Genotype2 p-value

Item SENS RES # G3 A4 G*A Enterobacteriaceae 0.30 0.38 0.08

5 8.5 (7.7-9.2) 8.7 (8.0-9.5) 7 8.3 (7.5-9.0) 8.6 (7.9-9.3) 14 8.3 (7.5-9.1) 8.1 (7.4-8.8) 21 8.2 (7.4-8.9) 8.1 (7.4-8.8) 28 8.4 (7.7-9.1) 7.9 (7.2-8.6) 34 8.9 (8.2-9.6) 8.0 (7.3-8.7)

Haemolytic bacteria 0.02 <.0001 0.03 5 <6.2A (2) (5.4-7.1) <6.3AB (1) (5.2-7.5) 7 <6.2A (9) (5.8-6.6) <6.3A (5) (5.8-6.8) 14 <6.3A (3) (5.7-6.9) <6.3A (7) (5.8-6.7) 21 <6.3A (7) (5.8-6.7) <6.2A (6) (5.8-6.7) 28 <6.3A (7) (5.9-6.7) <6.3A (6) (5.8-6.7) 34 8.7aB (8.3-9.1) <7.5bB (3) (7.1-7.9)

Clostridium perfringens 0.60 <.0001 0.97 5 7.7 3.3-12.0 7.5 3.5-11.5 ab 7 <8.0 (1) 3.2-12.7 7.8 3.5-12.1 b 14 7.5 3.6-11.4 <7.4 (1) 3.2-11.6 ab 21 <6.6 (1) 2.2-11.1 6.5 2.2-10.8 a 28 5.2 0.7-9.6 <5.0 (1) 0.7-9.3 c 34 <3.0 (6) 0-7.4 <2.8 (3) 0-7.2 d

Lactic acid bacteria 0.06 0.0002 0.25 5 9.7 9.3-10.0 9.4 9.0-9.8 a 7 9.6 9.3-10.0 9.4 9.0-9.7 a 14 8.9 8.5-9.3 8.7 8.3-9.0 b 21 9.1 8.8-9.4 8.8 8.5-9.2 a 28 9.0 8.6-9.3 8.7 8.4-9.0 b 34 8.8 8.5-9.2 8.6 8.2-8.9 b

Total anaerobic bacteria 0.86 0.43 0.36 5 9.9 9.7-10.2 9.9 9.7-10.2 7 9.7 9.4-9.9 9.7 9.4-9.9 14 9.7 9.5-10.0 9.7 9.5-10.0 21 9.7 9.5-10.0 9.8 9.5-10.0 28 9.8 9.6-10.0 9.8 9.6-10.0 34 9.8 9.5-10.0 9.8 9.6-10.0


! 20

1 Values are presented as least square means and 95% confidence intervals (in parentheses). 2 SENS = M307GA; RES = M307AA. Number of piglets: SENS=7, except day 5 (n=8), day 7 (n=9) and day 14

(n=4); RES=6, except day 5 (n=4), day 7 (n=5) and day 14 (n=7). 3 G = Genotype. 4 A = Age.

#: Rows with different letters, within a microbial group, are significantly different (p<0.05). A,B Values with different superscripts within a column are significantly different (p<0.05). a,b Values with different superscripts within a row are significantly different (p<0.05).

< : Indicates that at least one of the observations used to calculate the least square mean was below detection

level. Numbers in brackets indicate the number of samples below detection level.


! 21

Table 3 Short-chain fatty acid concentrations (mmol/kg sample) in faeces from piglets at 5, 7, 14, 21, 28

and 34 days of age1

Genotype2 p-value Item SENS RES # G3 A4 G*A Acetic acid 0.44 <.0001 0.23

5 42.3 (32.0-52.7) 39.5 (29.5-49.6) a 7 38.1 (29.0-47.1) 35.3 (26.0-44.6) a 14 32.7 (22.1-43.4) 29.9 (20.4-39.4) a 21 38.0 (28.7-47.4) 35.2 (26.2-44.2) a 28 46.5 (37.1-55.9) 43.7 (34.7-52.7) ab 34 58.9 (49.9-67.9) 56.1 (47.2-65.0) b

Propionic acid 0.19 <.0001 0.13 5 12.4 (8.1-16.7) 10.4 (6.2-14.6) a 7 10.9 (7.2-14.6) 8.9 (5.1-12.8) a 14 12.3 (7.8-18.8) 10.3 (6.4-14.2) a 21 12.6 (8.8-16.4) 10.6 (6.9-14.3) a 28 12.0 (8.2-15.9) 10.0 (6.4-13.7) a 34 22.4 (18.7-26.0) 20.4 (16.7-24.0) b

Butyric acid 0.09 0.09 0.48 5 6.6 (3.5-10.7) 4.6 (2.2-8.0) 7 5.3 (2.9-8.4) 3.5 (1.6-6.2) 14 3.0 (1.0-6.1) 1.7 (0.5-3.8) 21 2.5 (0.9-4.8) 1.3 (0.3-3.1) 28 4.0 (1.9-6.9) 2.5 (1.0-4.7) 34 5.3 (3.0-8.4) 3.6 (1.7-6.1)

A+P+B5 0.23 <.0002 0.14 5 61.7 (46.5-77.0) 55.4 (40.6-70.1) a 7 56.0 (43.0-69.0) 49.7 (36.1-63.2) a 14 49.4 (33.5-65.2) 43.0 (29.2-56.8) a 21 54.3 (40.8-67.8) 48.0 (35.0-61.0) a 28 63.8 (50.3-77.3) 57.4 (44.4-70.4) ab 34 87.1 (74.3-100.0) 80.7 (67.9-93.5) b

1 Values are presented as least square means and 95% confidence intervals (in parentheses). 2 SENS = M307GA; RES = M307AA. Number of piglets: SENS=10, except, day 5 (n=8), day 7 (n=9) and day

14 (n=7); RES=6, except day 5 (n=4), day 7 (n=5) and day 14 (n=7). 3 G = Genotype. 4 A = Age. 5 A+P+B = acetic + propionic + butyric acid.

#: Rows with different letters, within a short-chain fatty acid group, are significantly different (p<0.05).


! 22

Table 4 Counts (log cfu/g sample) of selected microbial groups in digesta from the gastrointestinal tract of

34 days old piglets (one week post-weaning)1

Genotype2 p-value Item SENS RES G3 S4 G*S Enterobacteriaceae 0.002 <.0001 0.004

Stomach 5.6 (4.8-6.5) 5.9 (5.0-6.7) Distal small intestine 8.8a (7.9-9.7) 7.5b (6.6-8.3) Caecum 8.8a (8.0-9.7) 7.8b (7.0-8.7) Mid colon 8.9a (8.0-9.8) 7.9b (7.1-8.8)

Haemolytic bacteria 0.02 <.0001 0.28 Stomach 5.2 (4.5-6.0) <4.4 (1) (3.8-5.1) Distal small intestine 7.9 (6.8-9.2) <6.8 (4) (5.8-7.9) Caecum 8.6 (7.4-10) <7.3 (4) (6.3-8.5) Mid colon 8.6 (7.4-10) <7.3 (3) (6.3-8.5)

Clostridium perfringens 0.93 0.63 0.24 Stomach <3.2 (1) (2.6-3.7) 3.2 (2.6-3.7) Distal small intestine <2.8 (5) (2.3-3.4) <2.8 (4) (2.3-3.4) Caecum <3.0 (4) (2.5-3.6) <3.0 (4) (2.5-3.6) Mid colon <3.0 (4) (2.4-3.5) <3.0 (4) (2.4-3.5)

Lactic acid bacteria 0.29 <.0001 0.40 Stomach 9.0 (8.4-9.6) 8.7 (8.1-9.3) Distal small intestine 8.3 (7.8-8.9) 8.0 (7.3-8.6) Caecum 8.6 (8.0-9.2) 8.3 (7.6-8.8) Mid colon 8.6 (8.0-9.2) 8.3 (7.7-8.9)

Total anaerobic bacteria 0.02 <.0001 0.01 Stomach 9.0 (8.7-9.4) 9.3 (9.0-9.6) Distal small intestine 8.9a (8.6-9.3) 8.3b (8.0-8.6) Caecum 9.3 (9.0-9.7) 9.2 (8.9-9.5) Mid colon 9.4 (9.1-9.7) 9.4 (9.1-9.7)

1 Samples from the stomach, distal small intestine, caecum and mid colon were analysed. Values are

presented as least square means and 95% confidence intervals (in parantheses). 2 SENS = M307GA; RES = M307AA . Number of piglets: SENS n=10; RES n=7. 3 G = Genotype. 4 S = Intestinal segment. a,b Values with different superscripts within a row are significantly different (p<0.05).

< : Indicates that at least one of the observations used to calculate the least square mean was below detection

level. Numbers in brackets indicate the number of samples below detection levels.

!


! 23

Table 5 Organic acid concentrations in digesta (mmol/kg wet sample) from the gastrointestinal tract of 34

days old piglets (one week post-weaning)1

Genotype2 p-value Item SENS RES G3 S4 G*S Lactic acid5 0.99 <.0001 0.91

Stomach 19.6 (5.3-62.7) 19.5 (6.9-50.2) Distal small intestine 5.6 (0.6-20.4) 5.5 (1.1-16.2)

Acetic acid 0.01 <.0001 0.01

Stomach 21.1 (4.2-37.9) 26.8 (11.6-41.9) Distal small intestine 4.2 (0.0-21.0) 5.4 (0.0-20.6) Caecum 47.6 (30.7-64.4) 39.0 (24.1-53.9) Mid colon 54.5a (37.7-71.4) 39.4b (24.4-54.4)

Propionic acid6 0.02 0.0001 0.01

Stomach 8.1 (2.7-13.6) 11.7 (6.9-16.5) Caecum 16.7 (11.3-22.2) 13.2 (8.1-18.3) Mid colon 16.6 (11.1-22.0) 11.9 (6.9-16.9)

Butyric acid7 0.90 <.0001 0.83 Stomach 8.3 (1.4-48.0) 8.1 (0.5-41.2) Caecum 2.3 (6.1-30.3) 2.2 (3.8-24.0) Mid colon 1.9 (6.8-29.0) 1.8 (4.6-23.6)


presented as least square means and 95% confidence interval (in parentheses). 2 SENS = M307GA; RES = M307AA. Number of piglets: SENS n=10, except the distal small intestine (n=9);

RES n=7, except the mid colon (n=6) and caecum (n=5). 3 G = Genotype. 4 S = Intestinal segment. 5 Samples from the caecum and mid colon had values below detection level. 6 Samples from the distal small intestine had values below detection level. No superscripts as the G*S

interaction was non-significant after pairwise comparisons using Bonferroni correction. 7 Samples from the distal small intestine had values below detection level. a,b Values with different superscripts within a row are significantly different (p<0.05).


! 24

S1 Table Dry matter content (%) and pH of digesta from the gastrointestinal tract of 34 days old piglets (one

week post-weaning)1

Genotype2 p-value Item SENS RES G3 S4 G*S Dry matter 0.33 <.0001 0.34

Stomach 25.1 (20.8-29.5) 24.0 (20.1-27.9) Distal small intestine 6.0 (1.7-10.3) 4.9 (1.0-8.7) Caecum 7.1 (3.2-10.9) 5.9 (2.2-9.6) Mid colon 7.8 (3.6-11.9) 6.6 (2.9-10.4)

pH 0.86 <.0001 0.55 Stomach 3.0 (2.7-3.2) 3.0 (2.7-3.2) Distal small intestine 7.3 (7.0-7.6) 7.3 (7.0-7.5) Caecum 6.7 (6.4-6.9) 6.7 (6.4-6.9) Mid colon 6.9 (6.7-7.2) 6.9 (6.7-7.2)


presented as least square means and 95% confidence intervals (in parentheses). 2 SENS = M307GA; RES = M307AA. Number of piglets: SENS=10; RES=7, except: pH SENS distal small

intestine (n=9); dry matter SENS piglets caecum and mid colon (n=5) and RES caecum (n=6). 3 G = Genotype. 4 S = Intestinal segment.

!


! 25

Figures:

Fig. 1. Heatmap showing relative abundances (%) of the (A) nine most abundant phyla (relative

abundance > 1%) and (B) 15 most abundant families in faecal samples from ETEC F18-sensitive

(SENS = M307GA) and resistant (RES = M307AA) piglets at 5, 7, 14, 21, 28 and 34 days of age.

Colours represent relative abundances. Number of samples: SENS = 35; RES=30.

Fig. 2. Principal Component Analysis of square root transformed OTU abundances displaying PC1

and PC2. Points are colored for genotype and grouped according to age. Sensitive = M307GA

(SENS); resistant = M307AA (RES). Number of samples: SENS = 35; RES=30.

Fig. 3. Principal Component Analysis of square root transformed OTU abundances displaying PC1

and PC3. Points colored for age. Number of samples: Sensitive = 35; resistant =30.

Fig. 4. Boxplots showing the 25 most significantly changed OTUs when comparing the microbial

community structure of (A) 5 and 28 days old piglets and (B) 28 and 34 days old piglets. Number of

samples: Sensitive = 35; resistant =30.

Fig. S1. Boxplots showing (A) the observed and (B) estimated species richness and (C) the

Shannon diversity index for sensitive and resistant piglets at 5, 7, 14, 21, 28 and 34 days of age.

Number of samples: Sensitive = 35; resistant =30.

Fig. S2. Heatmap showing the relative abundances (%) of the 25 most abundant genera in faecal

samples from ETEC F18-sensitive (SENS = M307GA) and resistant (RES = M307AA) piglets at 5, 7,

14, 21, 28 and 34 days of age. Colours represent relative abundances. Number of samples: SENS =

35; RES=30.


5 7 14 21 28 34

0.4

0

0

1.1

0

0

0.7

92.8

5

0.6

0

0

0

14.1

0

0.8

0.7

83.8

1

35.4

3.3

0

58.5

0

1.2

0

0.3

0.3

0

27.1

70

0.9

0

1.6

0

0

0.1

43.8

51.3

0.4

0.6

0.6

0

1.9

0

37.4

0.3

60.2

0

0.5

0.6

0

0

0.6

72.6

0.8

0

25.2

0.2

0

0.7

0

0.2

2.3

60

0

0

36.6

0.2

0.4

0

0.3

0

0.6

0

0

92.9

0.3

0.1

5.7

0.2

0

0

12.6

0.9

3.5

0.1

82.3

0.3

0.1

1.4

81.1

5

1.1

0.2

9.1

0.7

0.1

0.1

0

5.5

0.3

0

0

84.4

7.2

0.4

1.7

k__Unassigned_OTU_188

Spirochaetes

WPS−2

k__Unassigned_OTU_57

Actinobacteria

Fusobacteria

Proteobacteria

Bacteroidetes

Firmicutes

5 RES 5 SENS 7 RES 7 SENS 14 RES14 SENS 21 RES21 SENS 28 RES28 SENS 34 RES34 SENS

0.1

1.0

10.0

% ReadAbundance

Day 5 Day 7 Day 14 Day 21 Day 28 Day 34

OTU 57

OTU 188

RES SENS RES SENS RES SENS RES SENS RES SENS RES SENS

5 7 14 21 28 34

0.7

31.4

8.6

1

28.5

0.9

0.2

0.5

2.7

2.5

0

0.7

0

17.4

0.4

4.3

23.1

26.1

2.6

0.2

4.9

0.1

0

2.1

0.1

3.4

3

22.8

0.1

0.7

6.9

0.8

8

14.6

4.9

9.6

0.1

2.2

1.2

9.6

1.1

22.6

2.8

4.7

0.1

2.5

0.7

36.5

1.5

5.4

0.1

0.9

0.3

1.1

1.8

19.5

0.5

1.7

19.3

0.2

3.9

2.2

6.8

3.7

7.7

2.1

15.8

1.2

14.8

15.1

1.4

0.6

2.7

1.3

0.8

0.1

4.4

2.3

1.1

1.7

19.9

2

0.7

19.1

1

0.2

8.8

0.4

20.8

0.3

0.2

13.3

0.7

1.1

4.1

5.1

0.6

21.5

0.5

16.8

8.1

2.8

0.2

8

0.2

0.1

0.3

6.9

3.8

1

0.2

0.8

0.8

22.3

2.3

14.1

0.2

29.5

5.7

0.1

0.1

0.1

0.9

0.4

1.2

28.6

3.1

2.2

1.4

0.3

40.3

0.2

1.2

0.1

0.2

1

1.3

0.2

0.3

26

4.3

3.2

1.3

9.3

24.2

0.2

6.1

4.6

1.1

2.9

1.5

6.3

1

27.7

17.1

0.5

0.2

0.2

0.7

0

0

7.2

11.2

1.1

0.3

3.5

0.3

0.2

17.3

3.6

0

5.9

16

0.1

0

30.9

0.6

0

0.5

5.3

Bacteroidetes; S24−7

Bacteroidetes; [Paraprevotellaceae]

Proteobacteria; Enterobacteriaceae

Bacteroidetes; Rikenellaceae

Bacteroidetes; [Odoribacteraceae]

Bacteroidetes; Bacteroidaceae

Firmicutes; Erysipelotrichaceae

Firmicutes; Veillonellaceae

Firmicutes; o__Clostridiales_OTU_17

Bacteroidetes; o__Bacteroidales_OTU_2

Firmicutes; Lachnospiraceae

Bacteroidetes; Prevotellaceae

Firmicutes; Lactobacillaceae

Firmicutes; Clostridiaceae

Firmicutes; Ruminococcaceae

5 RES 5 SENS 7 RES 7 SENS 14 RES14 SENS 21 RES21 SENS 28 RES28 SENS 34 RES34 SENS

0.1

1.0

10.0

% ReadAbundance


RES SENS RES SENS RES SENS RES SENS RES SENS RES SENS

Bacteroidetes; order Bacteroidales OTU 2

Firmicutes; order Clostridiales OTU 17

(a)

(b)

Figure 1


−2.5

0.0

2.5

5.0

−5.0 −2.5 0.0 2.5 5.0PC1 [15%]

PC3

[9.2

%]

age

d_05

d_07

d_14

d_21

d_28

d_34

genotype

resistant

sensitive

PC

3 - v

aria

tion

expl

aine

d 9.

2%

PC1 - variation explained 15%

−2.5

0.0

2.5

5.0

−5.0 −2.5 0.0 2.5 5.0PC1 [15%]

PC3

[9.2

%]

age

d_05

d_07

d_14

d_21

d_28

d_34

genotype

resistant

sensitive

5

7 days

age (days)

714

21

28

34

5 7 14 21 28 34

−6

−3

0

3

−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0PC1 [15%]

PC2

[10%

] genotype

RES

SENS



PC

2 - v

aria

tion

expl

aine

d 10

%

5 7 14 21 28 34

−6

−3

0

3

−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0PC1 [15%]

PC2

[10%

] genotype

RES

SENS

5 7 14 21 28 34

−6

−3

0

3

−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0PC1 [15%]

PC2

[10%

] genotype

RES

SENS



PC

2 - v

aria

tion

expl

aine

d 10

%

5 7 14 21 28 34

−6

−3

0

3

−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0−5.0 −2.5 0.0 2.5 5.0PC1 [15%]

PC2

[10%

] genotype

RES

SENS

Figure 2

Figure 3




Actinobacteria; Corynebacteriaceae; OTU_333

Firmicutes; Ruminococcaceae; OTU_214

Firmicutes; Clostridiaceae; OTU_172

Bacteroidetes; [Paraprevotellaceae]; OTU_110




Firmicutes; Christensenellaceae; OTU_562

Bacteroidetes; o__Bacteroidales_OTU_95; OTU_95





Firmicutes; [Acidaminobacteraceae]; OTU_2665






Firmicutes; Veillonellaceae; OTU_46

Firmicutes; Lachnospiraceae; OTU_2737



0.01 0.10 1.00 10.00Read Abundance (%)

age

28

34


























0.01 0.10 1.00 10.00Read Abundance (%)

age

28

34

age (days)


Read abundance (%)









Bacteroidetes; order Bacteroidales OTU_95




Firmicutes; Peptostreptococcaceae; OTU_793


Firmicutes; Streptococcaceae; OTU_106

Actinobacteria; o__Actinomycetales_OTU_276; OTU_276

Actinobacteria; Coriobacteriaceae; OTU_376



Firmicutes; Lactobacillaceae; OTU_23




Actinobacteria; Actinomycetaceae; OTU_223











0.01 0.10 1.00 10.00Read Abundance (%)

age

5

28


























0.01 0.10 1.00 10.00Read Abundance (%)

age

5

28

age (days)

Read abundance (%)






Actinobacteria; order Actinomycetales OTU_276

(a)

(b)


























0.01 0.10 1.00 10.00Read Abundance (%)

age

28

34


























0.01 0.10 1.00 10.00Read Abundance (%)

age

28

34

age (days)


Read abundance (%)



































0.01 0.10 1.00 10.00Read Abundance (%)

age

5

28


























0.01 0.10 1.00 10.00Read Abundance (%)

age

5

28

age (days)

Read abundance (%)







(a)

(b)


























0.01 0.10 1.00 10.00Read Abundance (%)

age

28

34


























0.01 0.10 1.00 10.00Read Abundance (%)

age

28

34

age (days)


Read abundance (%)



































0.01 0.10 1.00 10.00Read Abundance (%)

age

5

28


























0.01 0.10 1.00 10.00Read Abundance (%)

age

5

28

age (days)

Read abundance (%)







(a)

(b)

Figure 4


5 7 14 21 28 34

2.00

2.25

2.50

2.75

3.00

3.25

3.50

3.75

4.00

4.25

4.50

4.75

5.00

5.25

resistant

sensitiveresistant

sensitiveresistant

sensitiveresistant

sensitiveresistant

sensitiveresistant

sensitive

Genotype

Dive

rsity

inde

x


Genotype

Div

ersi

ty in

dex

5 7 14 21 28 34

375

500

625

750

875

1000

1125

1250

1375

1500

1625

1750

resistantsensitive

resistantsensitive

resistantsensitive

resistantsensitive

resistantsensitive

resistantsensitive

Genotype

Estim

ated

num

ber o

f OTU

s


Genotype

Est

imat

ed n

umbe

r of O

TUs

5 7 14 21 28 34

250

375

500

625

750

875

1000

1125

1250

1375

resistantsensitive

resistantsensitive

resistantsensitive

resistantsensitive

resistantsensitive

resistantsensitive

Genotype

Obs

erve

d nu

mbe

r of O

TUs


Genotype

Obs

erve

d nu

mbe

r of O

TUs

(c)

(a)

(b)

Figure S1


5 7

1421

2834

0.7

2.80 0.8

0.1

0.2 016.2 0 06.3

0.7 00.7

0.2

0.1

0.6 00.5

13.4

2.7

2.5

0.6031.4

0.7

1.3

3.8

0.50 0 0.5 00.3

0.2

2.1

1.1

16.7

5.9 023.1 0 0.3

1.8

0.1

1.9

10.9

2.6

0.63

0.3

2.1

2.6

0.1

6.5

0.8

4.4

0.4

2.2

0.7

0.8

3.1 00.7

0.18 1.1

14.6

0.9

0.2

4.4

4.7

3.3

2.3

0.8

0.5

0.5

2.8

1.3

2.4

0.5

0.6

0.1

1.3 00.9

0.3

0.3

19.5

0.1

0.3

0.1

2.6

1.8

36.5

0.3

3.8

1.7

2.5

0.4

0.5 20.5

1.1

0.7

0.2

0.8

0.8

7.7

1.5

1.4

0.9

1.4

6.8

0.62 2.4

14.8

1.7

3.1

2.8

0.5

2.7

2.1

0.5

02 10.4

1.9

0.3

2.3

0.2

0.1

0.9

0.7

4.4

2.1

1.4

1.5

8.8

2.2

7.3

5.6

1.3

1.3

1.7

0.5

20.8

0.2

0.2

0.2

1.1

4.5

0.4

0.5

0.33 0.6

0.15 0.7

2.8

13.3

2.5

0.4

1.3

8.1

4.58 1.1

1.6

0.9

2.1

0.2

0.1

0.9

0.1

0.5

0.7

0.6

0.2

0.1

0.1

0.7

0.2

6.9

8.5

0.6

2.5

3.2

0.5

29.5

1.4

2.3

5.7

0.8

0.5

1.3

0.8

00.1

0.2

0.4

0.5

1.2 2 11.6

1.4 1 0.1

1.1

0.2

1.7

1.9

1.8

3.1

1.1

5.7

10.4

0.9

17.1

2.6

1.2

0.6

0.6

0.3

0.4

0.2

0.3

0.8

4.6

1.4

2.5

3.9

1.3

1.1

8.1

1.3

0.6

2.3 11.8

0.2

0.3

3.1

6.1

8.6

3.8

0.5

1.1

0.201 2.8

1.3

5.9

0.6

7.2

2.7

0.7

2.8

0.2

0.3

1.1 0 1.7

0.2

0.7

6.3 00.8 40.5

5.9

2.1

2.80 4.700.3

0.5

22.5

0.8

2.2 0 0.4

0.4

3.1

2.2

0.4

0.4 01.8

0.4

0.5

3.6 00.8

o__B

acte

roid

ales

_OTU

_74;

o__

Bact

eroi

dale

s_OT

U_74

Rum

inoc

occa

ceae

; f__

Rum

inoc

occa

ceae

_OTU

_18

[Par

apre

vote

llace

ae];

[Pre

vote

lla]

Lach

nosp

irace

ae; f

__La

chno

spira

ceae

_OTU

_78

Rum

inoc

occa

ceae

; f__

Rum

inoc

occa

ceae

_OTU

_35

Lach

nosp

irace

ae; C

opro

cocc

us

Ente

roba

cter

iace

ae; f

__En

tero

bact

eria

ceae

_OTU

_41

Rum

inoc

occa

ceae

; Osc

illosp

ira

[Odo

ribac

tera

ceae

]; Bu

tyric

imon

as

Rum

inoc

occa

ceae

; f__

Rum

inoc

occa

ceae

_OTU

_15

Clos

tridi

acea

e; f_

_Clo

strid

iace

ae_O

TU_1

3

Erys

ipel

otric

hace

ae; p−7

5−a5

Clos

tridi

acea

e; f_

_Clo

strid

iace

ae_O

TU_3

32

Bact

eroi

dace

ae; B

acte

roid

es

Rum

inoc

occa

ceae

; f__

Rum

inoc

occa

ceae

_OTU

_5

Rum

inoc

occa

ceae

; f__

Rum

inoc

occa

ceae

_OTU

_175

5

Rum

inoc

occa

ceae

; f__

Rum

inoc

occa

ceae

_OTU

_12

o__C

lost

ridia

les_

OTU_

17; o

__Cl

ostri

dial

es_O

TU_1

7

Rum

inoc

occa

ceae

; Rum

inoc

occu

s

o__B

acte

roid

ales

_OTU

_2; o

__Ba

cter

oida

les_

OTU_

2

Clos

tridi

acea

e; C

lost

ridiu

m

Clos

tridi

acea

e; f_

_Clo

strid

iace

ae_O

TU_6

Clos

tridi

acea

e; f_

_Clo

strid

iace

ae_O

TU_3

Prev

otel

lace

ae; P

revo

tella

Lact

obac

illace

ae; L

acto

bacil

lus

5 R

ES 5

SEN

S 7

RES

7 S

ENS

14 R

ES14

SEN

S21

RES

21 S

ENS

28 R

ES28

SEN

S34

RES

34 S

ENS

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

RE

S

SE

NS

RE

S

SE

NS

RE

S

SE

NS

RE

S

SE

NS

RE

S

SE

NS

RE

S

SE

NS

Ord

er B

acte

roid

ales

; OTU

2

Fam

ily C

lost

ridia

ceae

; OTU

3

Fam

ily C

lost

ridia

ceae

; OTU

6

Ord

er C

lost

ridia

les;

OTU

17

Fam

ily R

umin

ococ

cace

ae; O

TU 1

2

Fam

ily R

umin

ococ

cace

ae; O

TU 1

755

Fam

ily R

umin

ococ

cace

ae; O

TU 5

Fam

ily C

lost

ridia

ceae

; OTU

332

Fam

ily C

lost

ridia

ceae

; OTU

13

Fam

ily R

umin

ococ

cace

ae; O

TU 1

5

Fam

ily E

nter

obac

teria

ceae

; OTU

41

Fam

ily R

umin

ococ

cace

ae; O

TU 3

5

Fam

ily L

achn

ospi

race

ae; O

TU 7

8

Fam

ily R

umin

ococ

cace

ae; O

TU 1

8

Ord

er B

acte

roid

ales

; OTU

74

Day

5D

ay 7

Day

14

Day

21

Day

28

Day

34

Figu

re S

2



Chapter 6

Manuscript 3

Influence of Bacillus spp. spores and gentamicin on the gut microbiotaof suckling and newly weaned piglets

Ann-Sofie Riis Poulsen, Nadieh de Jonge, Jeppe Lund Nielsen, Ole Højberg, Char-lotte Lauridsen, Nuria Canibe.

In preparation.



ControlNo pro- or antibiotics

Faeces Day 7, 28 and 42

Digesta Stomach, ileum, caecum

and mid colonDay 3, 28 and 42

AntibioticPiglets administered

gentamicin day 4, 5 and 6


ProbioticSows and piglets administered

Bacillus spp. spores


Probiotic + antibioticSows administered Bacillus spp. spores, and piglets administered Bacillus spp.

spores and gentamicin

Faeces Day 7, 28, and 42

6 sows6 sows 6 sows 6 sows







1. 16S rRNA gene sequencing• V1-V3 region• Illumina MiSeq

2. Classical culture3. Organic acid analysis4. Biogenic amine analysis5. Gene expression analysis

• Inflammatory markers• Tight junction proteins

Intestinal tissue Ileum




24 sows12 piglets from each litter included

! 1

Influence of Bacillus spp. spores and gentamicin on the gut microbiota of suckling and newly

weaned piglets

Ann-Sofie Riis Poulsena, Nadieh de Jongeb, Jeppe Lund Nielsenb, Ole Højberga, Charlotte

Lauridsena, Nuria Canibea*

a Aarhus University, Faculty of Science and Technology, Department of Animal Science, Blichers

Allé 20, P.O. Box 50, DK-8830, Tjele, Denmark

b Center for Microbial Communities, Department of Chemistry and Bioscience, Aalborg University,

Aalborg, Denmark

* Corresponding author.

Aarhus University,


Blichers Allé 20, P.O. Box 50,


Tel.: +45 87 15 80 19

Fax.: +45 87 15 42 49



! 2

Abstract

Diarrhea is highly prevalent in neonatal piglets and is often treated with antibiotics like gentamicin.

Administrating antibiotics to newborn piglets may have short- and long-term consequences on gut

microbiota and immune system development. We hypothesise that these consequences may be

alleviated by concurrent probiotic administration. The objective was therefore to investigate the

effect of administrating gentamicin and a mixture of Bacillus licheniformis, B. subtilis and B.

amyloliquefaeceans spores on the gut microbiota of piglets pre- and post-weaning. Twenty-four

sows and their litters were randomly allocated to four treatment groups receiving; a) Bacillus spore

mixture to sows and piglets (PRO); b) gentamicin to piglets day 4, 5 and 6 of age (AB); c) Bacillus

spore mixture to sows and piglets, and gentamicin to piglets (PRO+AB); or d) no administration of

probiotics or antibiotics (CTRL). The study included 12 piglets from each litter. Faecal samples

were collected day 7, 14, 21, 28, 35 and 42. Piglets were sacrificed for intestinal digesta and tissue

day 3, 28 and 42. Selected samples were subjected to amplicon sequencing of the 16S rRNA gene,

culture counts, and organic acid, biogenic amine and tissue gene expression analysis (TNF-α, IL-

10, COX-2, ZO-1, OCLN, CLDN-4 and CLDN-2). Treatment had a significant effect on the

composition of the faecal microbial community on day 28 (padonis=0.003) and 42 (padonis=0.008), and

the colonic community on day 28 (padonis=0.014). Faecal species richness and diversity (p≤0.001),

and colonic richness (p=0.001), were higher in AB compared to PRO piglets on day 28, and were

not significantly different from day 42. Species richness and diversity were numerically lower in

CTRL piglets compared to AB piglets, and numerically higher compared to PRO piglets.

Significant differences in low abundant OTUs were observed between treatment groups. PRO

piglets had the highest faecal concentration of iso-butyric acid on day 7 (p=0.04) and a higher

butyric acid concentration compared to CTRL piglets (p=0.02); otherwise were no significant

effects observed in organic acid and biogenic amine concentrations, gene expression or bacterial

counts. The results show that administrating oral gentamicin to piglets shortly after birth may affect

gut microbial composition, and that it is counteracted by concurrent administration of Bacillus

spores, suggestively due to spores competitively excluding early colonisers. We conclude that both

gentamicin and Bacillus spores influence the gut microbial diversity of piglets, although

administration of the tested antibiotic did not result in severe dysbiosis. The importance of the

microbiological findings in relation to gut and animal health needs further investigation.


! 3

Introduction

The newborn young has an immature immune system and is hence highly susceptible to infections.

Despite vaccination programs, newborn piglets are in high risk of developing diarrhea in their first

week of life and the condition often leads to antibiotic treatment (Kongsted, 2013). The etiology is

commonly unknown prior to therapeutic intervention, why the empirically chosen antibiotics often

are of broad-spectrum. However, administering antibiotics to newborn piglets may have short- and

long-term consequences on the developing gastrointestinal microbiota and the local immune

system, which may ultimately result in increased disease susceptibility later in life.

Microbial gut colonisation begins during birth when the newborn comes into contact with

bacteria from the mother and the surrounding environment. This initial colonisation leads to an

array of complex processes responsible for establishing the gut microbiota (Mackie et al., 1999).

The gut microbiota plays a pivotal role in maturing the gut and local immunological functions

(Sommer and Baeckhed, 2013). During early life, however, the gut microbiota is instable and highly

susceptible to disturbances by influencing factors, antibiotics being one of them (Nylund et al.,

2014). Administering antibiotics within the first week of life has been shown to cause short- and

long-term changes on the composition of the gut microbiota and on intestinal gene expressions

related to immunological functions (Schokker et al., 2014; Schokker et al., 2015). Human studies

have furthermore reported an association between antibiotic treatments in early life and immune-

related disorders (Alm et al., 2008). Alleviating the negative effect of antibiotics on the immature

gut microbiota might be crucial in improving both intestinal and general health of piglets.

Probiotics are defined as ‘live microbial feed supplements which beneficially affect the host

animal by improving its intestinal microbial balance’ (Fuller, 1989) and are believed to exert their

anti-pathogenic effects by competing with pathogens for intestinal mucosal binding sites and

nutrients, and inhibiting pathogen growth by producing organic acids and antibiotic-like compounds

(Vondruskova et al., 2010). Bacteria commonly used as porcine probiotics belong to the

Lactobacillus, Enterococcus, and Bacillus genera. Unlike most other probiotics, Bacillus spp. is a

spore-forming bacteria, and hence more resistant to unfavorable environmental conditions than are

probiotics found as vegetative cells only (Leser et al., 2008). Bacillus cereus var. Toyoi has been

reported to reduce the intestinal number of enterotoxigenic Escherichia coli, diarrhea incidence and

morbidity in weaned piglets (Taras et al., 2005; Scharek et al., 2007a; Papatsiros et al., 2011). It has

furthermore been reported to have immunological effects by increasing the number of T-

lymphocytes locally in the intestinal tissue and faecal IgA concentration (Scharek et al., 2007a;


! 4

Scharek et al., 2007b). Also B. licheniformis and B. subtilis have been found to decrease morbidity,

mortality and post-weaning diarrhoea (Alexopoulos et al., 2004). Human studies, focusing on the

use of probiotics in relation to antibiotic treatment, have reported Lactobacillus casei to reduce the

incidence of antibiotic-associated diarrhoea and counteract the effect of antibiotics on the diversity

of the gut microbiota (Pirker et al., 2012).

However, detailed information on the effects of early life antibiotic therapy and

concomitantly probiotic administration on the gut microbiota and local immune parameters in

piglets are lacking. The aim of the current study is therefore to investigate the effect of

administrating antibiotics and probiotics on the gut microbiota in piglets. We hypothesise that

administration of gentamicin, a broad-spectrum aminoglycoside, will change the microbial

community of the gut, and that the expected negative effects of antibiotic administration are

alleviated by concomitantly administration of Bacillus spp. spores.


Study design

The present study was conducted according to the license obtained from the Danish Animal


administration. Sows and piglets were housed according to the general guidelines on housing of

pigs before and after farrowing, and after weaning.

A total of 288 piglets (crossbred Danish Landrace x Yorkshire x Duroc; mixed females and

males) from 24 sows (parity varying between one to seven) were included in the study. All sows

originated from Christiansminde Multisite K/S. The study was run in three blocks with eight sows

each. Before farrowing, sows were randomly divided into four treatment groups: (a) Sows fed a

probiotic mixture from ten days pre-partum until 28 days post-partum, and piglets fed the same

probiotic mixture during suckling and in the post-weaning period (PRO); (b) Piglets administered

gentamicin orally at 4, 5, and 6 days of age (AB); (c) Sows fed a probiotic mixture from ten days

pre-partum until 28 days post-partum, and piglets fed a probiotic mixture during lactation and in the

post-weaning period and administered oral gentamicin at 4, 5, and 6 days of age (PRO+AB); (d)

Neither sows nor piglets was fed probiotics or antibiotics (CTRL).

No creep-feed was allowed pre-weaning. Post-weaning feed was free from zinc oxide (Table

1) and no sawdust was allowed in the pens. All piglets were weaned at 28 days of age and housed

litter-wise. The study ended two weeks post-weaning. Clinical conditions and occurrence of


! 5

diarrhoea were recorded daily. Diarrhoea was recorded per litter and scored as 0, 1 or 2. The score 0

was given when no diarrhoea was observed in a litter, score 1 when mild diarrhoea was observed,

and score 2 when severe diarrhoea was observed.

Probiotics

The probiotic mixture consisted of a total of nine Bacillus subtilis, B. amyloliquefaeceans, and B.

licheniformis spore strains. Sows were fed 1x1010 cfu per kg feed two times daily from ten days pre-

partum until the piglets were weaned. Piglets were orally administered 2x109 cfu per day on day 3,

5, 7, 10, 13, 16, 20, 24, and 28 of age; 4x109 cfu per day at 33 days of age, and 8x109 cfu per day at

38 days of age. Piglets not receiving probiotics were administered an equal volume of sterilised

water.

Antibiotics

Piglets were orally administered 5 mg gentamicin (Gentacin Vet.) as a single bolus on day 4, 5, and

6 of age. Piglets not receiving the antibiotic were administered an equal volume of sterilised water.

Sample and data collection

Piglets were weighed weekly. Rectal faecal samples were collected weekly from three piglets from

each litter. Two piglets from each litter were euthanised at 3 days of age and one piglet from each

litter at 28 and 42 days of age. 3-day-old piglets (<5 kg) were euhanised by a blunt trauma to the

head, and 28 and 42 days old piglets were euthanized using a captive bolt gun followed by bleeding.

The abdomen was incised and the gastrointestinal tract removed. Luminal content (digesta) from the

stomach, small intestine (proximal and distal segments), caecum and colon (proximal, middle and

distal segment) was collected immediately after killing. Collected digesta from the two 3-day-old

piglets was pooled. pH was measured and the digesta was weighed and taken to the laboratory for

further analysis. Bacterial enumeration by culture was performed on digesta samples from the

stomach, distal small intestine, caecum, and mid colon. Faecal and digesta subsamples were stored

at -20°C for organic acid and biogenic amine analyses. Other faecal and digesta subsamples were

snap-frozen in liquid nitrogen and stored at -80°C for 16S rRNA gene amplicon sequencing. An

intestinal tissue sample from 50% of the length of the distal small intestine was carefully rinsed

with PBS, cut into two pieces of 1 cm2 and stored in Dulbecco’s modified Eagle’s medium

(DMEM) on ice until LPS-stimulation.


! 6

DNA extraction

Samples for DNA extraction included 214 faecal samples (day 7, 28 and 42; two day 7 samples

from the PRO group were missing) from 72 piglets and 144 digesta samples (distal small intestine

and mid colon from day 3, 28 and 42) from 72 piglets. DNA was extracted with the E.Z.N.A. Stool

DNA Kit (Omega Bio-Tek, inc., VWR international) following a standard protocol with the

following exceptions: In step 2, 450 µl SLX-Mlus buffer was added followed by 2x20 s bead

beating (FastPrep FP120; Bio 101 Savant/MP Biomedicals, USA). In step 3, 50 µl DS Buffer and

Proteinase K solution was added and followed by bead beating for 20 s. Step 4 was followed by

centrifugation at 2,000 g for 30 s. In step 5, 170 µl SP2 buffer was added and followed by bead

beating for 20 s. If no supernatant was present after step 8, another 180 µl SLX-Mlus, 20 µl DS

buffer and 67 µl SP2-buffer were added and the samples vortexed for 30 s., put on ice for 3 min and

centrifuged at maximum speed for 5 min. DNA extract purity was evaluated with Nanodrop

ND1000 (Thermo Scientific, USA) and quantified fluorometrically with Qubit 3.0 HS dsDNA assay

(Life Technologies, Thermo Fisher Scientific, USA). DNA concentrations were normalized to 5

ng/µl.

16S rRNA amplicon sequencing

Amplicon libraries were generated by targeted amplification of the V1-V3 hypervariable regions of

the bacterial 16S rRNA gene. The PCR reaction (25 µl) contained 10 ng template DNA, Platinum®

High Fidelity buffer (x1), dNTP (400 uM of each), MgSO4 (1.5 mM) and Platinum® Taq DNA

polymerase High Fidelity (1U) and barcoded library adapters (400 nM). V1-V3 primers: 27F

AGAGTTTGATCCTGGCTCAG and 534R ATTACCGCGGCTGCTGG. Thermocycler settings:

Initial denaturation at 95˚C for 2 min, 30 cycles of 95˚C for 20 s, 56˚C for 30 s, 72˚C for 60 s and

final elongation at 72˚C for 5 min. PCR reactions were run in duplicate for each sample and pooled

before purification. Purification of the amplicon libraries was performed using the Agencourt

AMPure XP bead protocol (Beckman Coulter, USA) and eluted in 23 µL nuclease-free water.

Individual libraries were quantified with Quant-iT HS dsDNA assay (Life Technologies, USA) and

quality checked on a Tapestation 2200 (Agilent, USA). Libraries were pooled in equimolar

concentrations, and diluted to 4 nM. The library pool was sequenced using an Illumina MiSeq

(Illumina, USA) and MiSeq reagent kit v3 (2x300 PE).


! 7

Amplicon bioinformatic processing and analysis

The obtained raw sequencing reads were quality filtered and trimmed using trimmomatic (v0.32)

(Bolger et al., 2014), only keeping reads with a minimum length of 275 bp. The trimmed reads were

merged using FLASH v. 1.2.7 (Magoc and Salzberg, 2011) and read pairs between 425 and 525 bp

in length were formatted for use with the UPARSE workflow (Edgar, 2013). Reads were

dereplicated and clustered into Operational Taxonomical Units (OTUs) using USEARCH7 at 97%

sequence similarity. Taxonomy was assigned using the RDP-classifier as implemented in QIIME

(Caporaso et al., 2010) with a minimum confidence of 0.8 and Greengenes (version 08-2013) as a

reference database. Results were analysed in R studio (version 0.99.489 for Mac) using the Ampvis

package (Albertsen et al., 2015).

Organic acid and biogenic amine analysis

The concentrations of short-chain fatty acids and lactic acid in faeces and digesta samples were

measured as previously described by Canibe et al. (2007). Biogenic amine concentrations

(agmatine, putrescine, cadaverine and tyramine) were measured according to the following

procedure. Samples (including a blank sample) were diluted 10-fold with an internal standard

solution and homogenized in a Smasher paddle blender (bioMérieux Industry, USA) for 2 min.

Proteins were precipitated by mixing 20 µl sample or standard mix with 780 µl 0.1 M hypochloric

acid and 240 µl 2 M perchloric acid, followed by incubation at room temperature for 5 min and

centrifugation at 13,000 rpm for 5 min. Derivates were produced by adding 320 µl 0.5 M sodium

bicarbonate to 200 µl supernatant from the protein precipitation step, followed by the addition of

600 µl 5 mM Fmoc-solution and mixing. The samples were subsequently heated at 40°C for 10

min, followed by mixing with 40 µl concentrated hypochloric acid. Samples were centrifuged at

13,000 rpm for 5 minutes. 600 µl supernatant was transferred to a vial and analysed by high-

performance liquid chromatography (HPLC).

Microbiological analysis of digesta samples

For microbial plating, digesta samples (approximately 5 g) were transferred to bottles containing 50

ml of pre-reduced salt medium (Holdeman et al., 1977). The bottle content was then transferred to a

CO2 flushed bag and homogenized in a Smasher paddle blender for 2 min. One ml of the digesta



! 8

fold dilutions were prepared using the technique previously described by Miller and Wolin (1974).

The samples were plated on selective (and indicative) and non-selective agar plates.


incubation for 1 day. Yeasts were enumerated on Malt, Chlortetracycline and -Amphenicol agar

(Merck 1.03753 (yeast extract), 1.05397 (malt extract), 1.07224 (bacto-pepton), 1.08337 (glucose),

1.01614 (agar-agar) and Oxoid Sr0177E) after aerobic incubation for 2 days. Hemolytic bacteria

were enumerated on blood agar (Oxoid Pb5039A) after aerobic incubation for 1 day. Clostridium

perfringens were enumerated using the pour-plate technique on Tryptose Sulfit Cycloserine agar

(Merck 1.11972, 1.00888) after anaerobic incubation for 1 day. Lactic acid bacteria were

enumerated on de Man, Rogosa and Sharpe agar (Merck 1.10660) after anaerobic incubation for 2

days. Total anaerobic bacteria were enumerated in roll tubes containing pig colon fluid-glucose-

cellobiose agar (Holdeman et al., 1977) and incubated for 7 days. Bacillus spores were enumerated

on Casein soya bean digest broth agar (Oxoid CM0129) after aerobic incubation for 1 day. Prior to

Bacillus spore enumeration, the sample was incubated in a water bath at 80˚C for 20 min. All plates

and roll-tubes were incubated at 37˚C.

LPS-stimulation of ileal tissue

One 1 cm2 tissue sample was transferred to 1 ml DMEM and 10 µl PBS (control) and the second 1

cm2 tissue sample was transferred to 1 ml DMEM and 10 µl LPS (1 µg/µl; Sigma L4391). The two

test tubes were incubated at 37˚C for 120 min during constant gently shaking. After ended

incubation, the tubes were stored on ice for 10 min and then transferred to 700 µl RNAlater (Sigma-

Aldrich), stored at 5˚C for one day and then stored at -20˚C until gene expression analysis.

Gene expression analysis

The intestinal tissue sample was added to a 2 ml Eppendorf tube together with a 5 mm Stainless

Steel Bead and homogenized on the TissueLyser system (Qiagen). Total mRNA for gene expression

analysis was extracted using the NucleoSpin RNA isolation kit (Macherey-Nagel, Germany)

following the manufacturers protocol. The Nanodrop ND1000 was used for measuring RNA

quantity and assessing RNA quality (Thermo Scientific, USA). All samples were diluted to 100

ng/µl and converted to cDNA using the High-Capacity cDNA Reverse Transcription kit (Applied

Biosystems, USA) according to the manufacturers protocol. Real Time PCR reactions contained 2

µl cDNA and 8 µl mix containing primers, probe (Table 2) and 2X TaqMan mastermix, and run in


! 9

triplicate on the Applied Biosystems ViiA 7 Real-Time PCR system (Life Technologies) with the

following thermocycler settings: 50˚C for 2 min, 95˚C for 10 min and 40 cycles of 95˚C for 15 s

and 60˚C for 60 s. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as

housekeeping gene. The Applied Biosystem ViiA 7 software was used for determining the gene

expression cycle threshold (Ct) values. Relative quantification (comparative threshold method) was

used for evaluating the gene expression level using non-LPS stimulated ileal samples from 28 days

old CTRL pigs as reference samples. For each sample the Ct value of the GAPDH gene was

subtracted from the Ct value of the target gene (ΔCt). The average ΔCt value of the reference

animals was then subtracted from the ΔCt value of all samples (ΔΔCt). The expression of the target

gene was given as fold change calculated by 2-ΔΔCt.

Statistical analysis

Principal Component Analysis was performed on square root transformed OTU abundances.

Significance of treatment was tested on the first two principal components (PCs) using the envfit

parametric test and on the Bray-Curtis dissimilarity matrix using the Adonis test (Oksanen, 2015).

The parametric Wald-test (Love et al., 2014) was used to test for significant OTU abundance

differences between the four treatment groups. OTUs with an adjusted p<0.001 were considered

significantly different between the respective treatments.

The impact of treatment and age on bacterial, organic acid and biogenic amine parameters,

microbial richness, diversity index, gene expression levels and body weight were investigated by

fitting the data to a linear mixed model using the lmer function from the lme4 package (Bates, 2014)

using R studio (Version 0.99.489 for Mac). Diet, age, and intestinal segment (the latter included

when analysing samples obtained from slaughter) were included as fixed effects, while piglet and

sow were included as random effects (by including random intercept terms) to account for multiple

observations from the same litter and the same pig. When analysing the body weight variable, the

piglet body weight at the day of birth was included as a co-variate. The fixed effects were tested

using an F-test with Kenward-Roger approximation, where the reduced model was tested against

the full model. This was done using the KRmodcomp function in the pbkrtest package (Halekoh,

2014). When a fixed effect was found to be significant, a post-hoc test was performed using the

multcomp package and Bonferroni adjustment to correct for multiple comparisons (Hothorn, 2008).

Effects were considered significant when p<0.05.


! 10

Results

General observations

Weekly body weights of piglets are shown in Table 3. Body weight was not affected by treatment

(p=0.89) or gender (p=0.72) (data not shown). During the suckling period, PRO and PRO+AB

piglets had a higher frequency of grade 1 and 2 diarrhoea, while after weaning there was a higher

frequency of grade 1 diarrhoea in PRO+AB piglets (Table 4).

Gut microbiota – culture results

Treatment had a significant effect on the number of Bacillus spores, with the highest numbers in

PRO and PRO+AB piglets compared to AB and CTRL piglets (p<0.001). In addition, age and

segment had an interacting effect. The number of Bacillus spp. spores in the caecum was higher on

day 3 and 28 than on day 42 (p≤0.04) and in the mid colon, it was higher on day 28 (p≤0.003) than

on day 3 and 42 (Figure 1A).

Treatment did not have an effect on the number of Enterobacteriaceae, haemolytic bacteria,

C. perfringens, total anaerobic bacteria or lactic acid bacteria in any of the investigated gut

segments (Figure 1B-F). The number of Enterobacteriaceae in the stomach was highest on day 3

compared to day 28 (p=0.01), in the ileum the number was higher day 3 compared to day 28 and 42

(p<0.02), and in the caecum and mid colon, the number was highest on day 3 (p<0.001). In all

segments, the number of haemolytic bacteria was highest on day 3 (p<0.001). The numbers of lactic

acid bacteria in the stomach and ileum were lowest on day 3 (p≤0.04); and in the mid colon, the

number was lowest on day 28 (p<0.001) (Figure 1F). In the stomach, ileum, and caecum, C.

perfringens numbers decreased with age, being lowest on day 42 (p<0.001), and in the mid colon,

the number was highest on day 3 compared to day 28 (p=0.03), and lowest on day 42 compared to

day 3 and 28 (p<0.001). In the stomach, the number of total anaerobic bacteria was lowest on day 3

(p≤0.02); in the caecum, the counts were highest on day 3 compared to day 42 (p<0.001); and in the

mid colon, the number was highest on day 3 compared to day 28 and day 42 (p≤0.01). Treatment

had no effect on pH in any of the investigated segments (data not shown).

Bacterial metabolites

Age and intestinal segment were found to have an interacting effect on the ileal and mid colonic

digesta concentrations of the investigated biogenic amines (cadaverine and putrescine) and organic

acids, while treatment did not have a significant effect (Table 5 and 6). Agmatin and tyramin were


! 11

not included in the analysis as the majority of the observations were zero/below detection level.

Ileal and colonic concentrations of cadaverine and putrescine were consistently lower (numerically)

in AB piglets compared to the other three groups. In the ileal digesta, concentrations of cadaverine

and putrescine were highest on day 42 (p<0.001; p≤0.13), while in the mid colon, the highest

concentration of putrescine was found on day 3 (p<0.001), and the concentration of cadaverine was

higher on day 3 compared to day 28 (p<0.001). The ileal concentration of acetic acid was higher on

day 42 compared to day 3 (p=0.006) and the lactic acid concentration was highest on day 42

(p<0.001). In the colonic digesta, the concentrations of acetic and butyric acid were highest on day

42 (p<0.001), while the concentrations of propionic and valeric acid were highest on day 42

(p<0.001; p≤0.04) and lowest on day 3 (p≤0.01; p≤0.003). The concentration of iso-valeric acid was

higher on day 42 compared to day 3 (p=0.03) in both ileal and colonic digesta. The concentration of

iso-butyric acid was similar between day 3, 28 and 42 regardless of intestinal segment. Colonic

concentrations of propionic acid on day 28 and 42 were unexpectedly higher than the acetic acid

concentrations.

Organic acid concentrations in faecal samples from day 7, 28, and 42 are shown in Table 7.

PRO piglets had the highest faecal concentration of iso-butyric acid on day 7 (p=0.04) and a higher

concentration of butyric acid compared to CTRL piglets also on day 7 (p=0.02). Treatment had no

other significant effect on faecal organic acid concentrations. The concentrations of iso-butyric and

iso-valeric acid were higher on day 7 compared to day 42 in PRO piglets (p=0.008; p=0.006) and

the concentration of propionic acid was highest on day 42 (p<0.001) for PRO+AB, CTRL and AB

piglets (p<0.001), and higher on day 42 compared to day 3 in PRO piglets (p=0.003). The faecal

concentration of butyric acid was higher in PRO piglets compared to CTRL piglets on day 7

(p=0.02) and highest on day 42 in AB (p≤0.009) and PRO+AB (p≤0.02) piglets, and higher on day

42 compared to day 7 (p<0.001) in CTRL piglets. The faecal acetic acid concentration was highest

on day 42 (p<0.001) and lowest on day 7 (p≤0.02). Neither treatment nor age had an effect on the

faecal concentration of valeric acid.

Gut microbiota – 16S rRNA gene amplicons

Sequencing of 356 samples yielded a total of 10,638,599 reads. A sequencing depth of 10,000 reads

was considered appropriate from rarefaction curves (data not shown) and excluded 23 samples from

further analysis. Recovered reads were clustered into 3400 OTUs and classified into 20 phyla, 100


! 12

families and 191 genera. Three of the four detected Bacillus species were classified to species level

and belonged to B. safensis, B. coagulans and B. cereus.

Faeces

Figure 2 shows the relative abundances of the ten most abundant phyla and 20 most abundant

genera in faecal samples from day 7, 28 and 42. Firmicutes and Bacteroidetes, followed by

Fusobacteria, were the most abundant phyla on day 7 and 28 irrespective of treatment. Firmicutes,

followed by Bacteroidetes, were the most abundant phyla on day 42, whereas Fusobacteria was

detected at very low levels (Figure 2). At genus level, Bacteroides, Lactobacillus, Prevotella and

Fusobacterium were the most abundant genera on day 7; Prevotella and Fusobacterium most

abundant on day 28; and Lactobacillus and Prevotella most abundant on day 42 irrespective of

treatment. 12 of the most abundant genera belonged to Firmicutes, six to Bacteroidetes, one to

Fusobacteria and one to Proteobacteria. The observed microbial community richness and diversity

index (Shannon index) was lower in PRO piglets compared to AB piglets (p=0.0001; p<0.001) and

PRO+AB piglets (p=0.03; p=0.046) on day 28, while the estimated richness was lower in PRO

piglets compared to AB piglets (p=0.0004) (Figure 3). Age had a significant effect on microbial

richness and diversity, with the observed and estimated species richness and Shannon index lowest

on day 7 (p<0.001), irrespective of treatment. In addition, for treatments CTRL, PRO and PRO+AB

the microbial richness and diversity was highest on day 42 (p<0.02) (figure 3).

There was a significant effect of treatment on the overall community composition of the four

treatment groups on day 28 (padonis=0.003) and 42 (padonis=0.008) (Figure 4). Performing a

redundancy analysis on day 28 and 42 revealed four groupings with slight overlap, indicating that

the four different treatments tended to result in four different communities (Figure 4C and 4E). Age

(padonis=0.001) (Figure S1) and litter (padonis=0.002) had a significant effect on the faecal microbial

composition. Table 8 shows the OTUs that were found in significantly different read abundances

between treatment groups. Generally were the significantly different OTUs present in relatively low

read abundances.

Digesta

Figure 5A and 5B show the relative abundances of the eight most prevalent phyla and 20 most

prevalent genera in ileal digesta from day 3, 28 and 42 of age. Firmicutes was clearly the most

abundant phylum on all three days investigated, irrespective of treatment. On day 3, Fusobacteria


! 13

was measured to be between 2.1 and 12.1 % in all treatments. Lactobacillus was the most abundant

genus irrespective of age and treatment. On day 3, SMB53, Clostridium, Sarcina, Streptococcus and

Fusobacterium followed Lactobacillus, with the relative abundance of Sarcina ranging between 0.7

% (AB group) and 20.8 % (PRO+AB group). On day 28, SMB53, 02d06, Sarcina and Mitsuokella

followed Lactobacillus, with the relative abundance of Mitsoukella ranging between 0.1 % (CTRL

group) and 13.8 % (PRO group). On day 42, 02d06, SMB53, and Streptococcus followed

Lactobacillus.

Figure 5C and 5D show the relative abundances of the eight most prevalent phyla and 20

most prevalent genera in digesta from mid-colon on day 3, 28, and 42 of age. Firmicutes,

Bacteroidetes, and Fusobacteria were the most abundant phyla on day 3 irrespective of treatment,

while Firmicutes and Bacteroidetes were the most abundant phyla on day 28 and 42 irrespective of

treatment. Fusobacteria was also abundant on day 28 in PRO and PRO+AB piglets. At genus level,

Fusobacterium, Bacteroides, Lactobacillus and Clostridium were the most abundant genera on day

3 in all treatment groups. Prevotella, Lactobacillus, and Fusobacterium dominated on day 28, with

Fusobacterium being most abundant in PRO and PRO+AB piglets, and Prevotella most abundant in

PRO+AB piglets. Lactobacillus and Prevotella were the dominating genera on day 42, with the

highest abundance of Lactobacillus in CTRL piglets. The observed richness of colonic digesta on

day 28 was higher in AB piglets compared to PRO (p=0.001) and PRO+AB (p=0.02) piglets, and

the estimated richness was higher in AB compared to PRO (p=0.001) piglets (Figure 6). Age had a

significant effect on the observed and estimated species richness in colon contents. The observed

species richness was lowest on day 3 in all treatments groups (p≤0.02), except PRO+AB, and

highest on day 42 in PRO (p=0.01) and PRO+AB piglets (p=0.001). The estimated species richness

was lowest on day 3 in AB and CTRL piglets (p<0.001), and lowest on day 3 (p≤0.04) and highest

on day 42 (p≤0.05) in PRO and PRO+AB piglets. The Shannon diversity index was lowest on day 3

in AB piglets (p≤0.03), higher on day 28 compared to day 3 (p=0.006) in CTRL piglets, and higher

day 42 compared to day 3 in PRO+AB piglets (p<0.001) (figure 6).

Treatment had an effect on the overall microbial community composition in the mid colon

on day 28 (padonis=0.013) (Figure 7). Otherwise had treatment no significant effect on the microbial

communities (data not shown). Age had an effect on the overall microbial communities in the ileum

and mid-colon (p=0.001) (Figure S2). Samples from the ileum and mid colon clustered closer

together on day 3 compared to day 28 and 42. Litter had no effect on the microbial communities of

the ileum and mid colon (data not shown). Table 8 shows the OTUs that were found in


! 14

significantly different read abundance between treatment groups. Generally the significantly

different OTUs were present in relatively low read abundances.

Ileal gene-expression

Comparative expression of TNF-α, COX-2, IL-10, ZO-1, OCLN, CLDN-2, and CLDN-4 genes are

seen in Figure 8. LPS stimulation of ileal tissue increased the expression of TNF-α (p=0.05) and

COX-2 (p=0.04), but had no effect on IL-10, ZO-1, OCLN, CLDN-2 and CLDN-4 gene expression.

Treatment had no effect on the gene expression level of any of the investigated genes. TNF-α

(p<0.001), CLD-4 (p=0.05) and ZO-1 (p=0.01) genes were found to be expressed at higher levels

on day 28 compared to day 42

Discussion

The potential of using probiotics has been intensively investigated in both humans and animals. In

pigs, several probiotics have been tested and some have been found to have beneficial effects on the

gut microbial community and intestinal immunological parameters. However, knowledge on the

combined effects of pro- and antibiotics on the gut microbiota and intestinal health of pigs is

lacking. To our knowledge, this study is the first to investigate the combined effect of Bacillus spp.

spores and gentamicin on the gut microbiota and gut health parameters in piglets.

Analysis of 16S rRNA gene sequences confirmed an effect of treatment on the overall

colonic and faecal microbial communities at the day of weaning (day 28), where AB piglets,

compared to PRO piglets, consistently had the richest and most diverse microbiota. Contrary to the

other treatment groups, the microbial richness and diversity of AB piglets were the same on day 28

as on day 42 (lower on day 28 for the other treatment groups). These observations could indicate an

accelerated microbial gut colonisation in the distal gut segments of AB piglets, perhaps caused by

gentamicin inhibiting bacterial groups that would colonise the gut under normal (non-treated)

circumstances, thereby allowing a more diverse community to colonise faster. In accordance with

these results, Kim et al. (2012) suggested an accelerated maturation of the faecal microbiota in

tylosin-supplemented pigs which the non-treated pigs, however, eventually catched up on. Schokker

et al. (2015) suggested that the effects of intrinsic factors, e.g. early impact of an antibiotic, are most

apparent at times when the gut microbiota has been more or less stabilized, as seen just prior to

weaning and several weeks post-weaning, and our results are generally supportive of this.


! 15

Interestingly, though, the ileal microbiota was the most stable community across treatments without

any significant effects on the overall community, species richness or diversity on day 28 and 42 of

age. Well aware of the varying mode of action among antibiotics, subcutaneous tulathromycin

injection at four days of age have been shown to have short- and long-term effects on the jenunal

microbiota of piglets by decreasing the microbial diversity (Schokker et al., 2014; Schokker et al.,

2015). The short-term effect was seen when the piglets were 8 days of age, the long-term effect was

seen at 176 days of age, while no effect was seen at 55 days of age. So, if the ileal digesta had been

collected from piglets a few days after antibiotic administration in the current study, we would

perhaps have found gentamicin to have a more profound effect on the ileal microbiota. An

explanation might also be, that the spectrum of activity of gentamicin merely was incompatible with

the community found in the ileum. Only faecal samples were collected in the immediate period

following antibiotic administration and it could be speculated that the antibiotic effect was not large

enough to be reflected in these samples.

Enumerating specific bacterial groups by culture did not show any differences between

treatment groups. As gentamicin is especially active against gram-negative infections (Adams,

2001), we did expect a reduction in the number of Enterobacteriaceae cultured from intestinal

digesta. A possible explanation is that the digesta samples were collected 22 and 36 days after the

last gentamicin dose was administered, thus allowing enough time for the piglets to be re-colonised

with gram-negatives from the environment. Daily administration of subtherapeutic doses of

virginiamycin has been shown to have a decreasing effect on anaerobic bacterial and lactoballi

counts and an increasing effect on coliform counts (Jensen, 1988).

The authors found the effect of gentamicin to be unexpectedly low. Oral antibiotics are

generally thought to cause intestinal dysbiosis, as seen in e.g. humans where diarrhoea is a common

complication to antibiotic therapy (Breves et al., 2000; Pirker et al., 2012). However, there exists

many different classes of antibiotics that target different spectra of bacteria, and the development of

antibiotic-associated diarrhea has been shown to be dependent on the type of antibiotic used (Anand

et al., 1994). The relatively low effect of gentamicin may have impeded the detection of potential

effect(s) of Bacillus spores on the gut microbial community in the current study.

Bacillus spores were included to assess whether they were able to counteract the effect of

early gentamicin administration. To allow piglets to be exposed to the spores already from birth,

they were included in the sow feed from ten days before expected farrowing. Plate cultures of

Bacillus spores showed that piglets in the PRO and PRO+AB groups had the highest concentration


! 16

of spores along the length of their gastrointestinal tracts on day 3 post-farrowing compared to

piglets in the CTRL and AB groups. Sampling on day 3 was done prior to probiotic administration,

demonstrating that the spores must have been vertically transferred from the sow to the piglets,

which is in accordance with Taras et al. (2005). Our results suggest that probiotics administered to

the dam with the feed during late gestation are transmitted to the offspring.

Piglets given the Bacillus spores (PRO and PRO+AB groups) had higher spore counts than

the other groups. However, the applied method quantified only the number of spores and not

vegetative cells, why we were not able to account for the precise degree of spore germination.

Nevertheless, earlier studies report that B. subtilis and B. licheniformis are able to germinate in the

gut of grower pigs, though only showing limited ability to grow (Leser et al., 2008). Due to the

resilient spore structure (Cutting, 2011), 16S rRNA sequencing were expected to detect germinated

cells only. The detected abundances of Bacillus genera in the ileum, colon and faeces were,

however, extremely low regardless of sample type, age and previous spore administration. If only a

small percentage of the spores germinated, the detection threshold of the sequencing method may

have been too high compared to more traditional methods as suggested by Lagier et al. (2012).

Despite the apparent lack of germination, the oral administration of Bacillus spores did seem

to have an in vivo effect. The consistently lower richness of the communities of the distal colon of

PRO piglets suggests that the normal colonisation pattern during nursing was hindered by the

spores. This was especially evident as the increased colonic and faecal richness and diversity in AB

piglets seemed to be counteracted by spore supplementing. Bacillus spores have previously been

reported to be able to adhere to intestinal cells (Tam et al., 2006), and it could thus be speculated

that the spores were able to competitively halt the colonisation pattern seen after gentamicin

administration. Previous reports on the effect of different probiotics on the microbial diversity are

contradicting. In agreement with our findings, Wang et al. (2012) found Lactobacillus acidophilus

and Pediococcus acidilactici to decrease species richness and diversity of colonic digesta from pigs

in the post-weaning period. The action of decreasing the number of colonising species may have a

health promoting effect when the infection pressure is high. In the current study, piglets were

challenged only during the natural process of weaning, and under these circumstances did spore

administration not improve piglet health. In fact, piglets administered Bacillus spores tended to

have a higher frequency of diarrhoea before and after weaning. As there are no general guidelines

on probiotic use in pigs, the dose and interval may have been sub-optimal for use in piglets.

Furthermore, an increased intestinal permeability has been associated with the use of Bacillus


! 17

cereus var. Toyoi (Altmeyer et al., 2014), which would be expected to have a compromising effect

on intestinal health.

Weaning is a stressful event for piglets, where numerous factors contribute to physiological

and microbial changes in the gut (Lalles et al., 2007). Intrinsic and extrinsic factors as litter, nursing

mother and genotype are found to have the biggest influence on the gut microbiota when the piglets

still suckle, while this influence decreases after weaning (Bian et al., 2016). As a consequence, the

reduced effect of gentamicin and spore administration two weeks after weaning was expected, and

the event of introducing solid foods, the stress of being separated from the sow and transported to a

new stable environment, all seemed to have a stronger influence on the gut microbiota.

The establishment of a gut microbiota is a prerequisite for an optimal development of the

intestinal immune system during early life (Sommer and Baeckhed, 2013), and administering

antibiotics to piglets shortly after birth, has been shown to alter the expression of genes related to

immunological functions (Benis et al., 2015; Schokker et al., 2015). The genes investigated in the

current experiment encoded pro- and inflammatory cytokines and cell-to-cell adhesion proteins,

however, neither early gentamicin nor continuous spore administration was able to alter the

expression of these genes. Hence, treatment apparently did not have any effect on the intestinal

barrier or inflammatory status. These results may have been different, it the piglets had been

exposed to an infection challenge.

Age is an important driver of gut microbiota maturation and is an important influencing

factor (Slifierz et al., 2015). In the present study, sequencing data from day 3 and 28 clearly showed

the effect of age. The microbial communities on day 42 was furthermore clearly different from

those on day 3 and 28, though the event of weaning on day 28 causing changes in diet, environment

and stress complicates the interpretation of the gut microbial changes from 28 to 42 days of age.

Microbial richness and diversity of the colonic and faecal communities increased up until day 42,

while it was more or less constant in the ileum. Thus the ileal community is quickly colonised by

the adult-like number and proportion of species, albeit the colonising species change with age.

Conclusion

In conclusion, administering gentamicin on three consecutive days, from 4 days post-partum,

caused an increased diversity and richness of the colonic and faecal microbiota, which was evident

3 weeks later. Administration of Bacillus spores, on the other hand, decreased species richness and

diversity and hence counteracted the effect of gentamicin. We were not able to show that spore


! 18

administration beneficially affected the health of newly weaned piglets. Yet, the importance of

these findings in relation to the clinical status of the animals needs further investigation. Such an

investigation should include a long-term study following the pigs into their adult life and include a

challenge trial. The study furthermore underlines the importance of distinguishing between different

classes of antibiotics when discussing dysbiosis in relation to antibiotic treatment.

Acknowledgements

The authors would like to thank Trine Poulsen, Helle Handll, Lene Rosborg Dal, Hanne Purup,

Inger Marie Jepsen and Thomas Rebsdorf for their technical assistance during the experiment, and

subsequent extensive sample processing and analysis. Also thanks to the staff of SB for taking good

care of the sows and piglets from the beginning to the end of the experiment.


! 19

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! 22

Tables:

Table 1

Item %

Dry matter 88.4

Protein 22.2

Fat 7.4

Ash 4.9

Chemical analyses were performed by Eurofins Steins Laboratory A/S, Vejen, Denmark.

Table 2

Information on primers. probes and genes for real-time PCR

Target gene Primer sequences Accession number GAPDH Forward: 5’-GTCGGAGTGAACGGATTTGG-3’

Reverse: 5’-CAATGTCCACTTTGCCAGAGTTAA-3’ Probe: 5’-CGCCTGGTCACCAGGGCTGCT-3’

AF017079

TNF-α Forward: 5’-AACCCTCTGGCCCAAGGA-3’ Reverse: 5’-GGCGACGGGCTTATCTGA-3’ Probe: FAM-TCAGATCATCGTCTCAAAC-MGB

X57321

COX-2 Forward: 5’-GGGACGATGAACGGCTGTT-3’ Reverse: 5’-CACAATCTTAATCGTTTCTCCTATCAGT-3’ Probe: 5’-AGACGAGCAGGCTGA-3’

NM_214321

IL-10 Forward: 5’-GAGGAGGTGAAGAGTGCCTTTA-3’ Reverse: 5’-CTCACCCATGGCTTTGTAGACA-3’ Probe: FAM-CCTCTCTTGGAGCTTGC-MGB

L20001

ZO-1 TaqMan® Gene Expression Assay. catalogue no. 4351370. assay ID ss03373514_m1 (Applied Biosystems. Thermo Fisher Scientific)

OCLN TaqMan® Gene Expression Assay. catalogue no. 4351370. assay ID ss03377507_u1 (Applied Biosystems. Thermo Fisher Scientific)

CLDN-2 TaqMan® Gene Expression Assay. catalogue no. 4351370. assay ID ss03375002_u1 (Applied Biosystems. Thermo Fisher Scientific)

CLDN-4 TaqMan® Gene Expression Assay. catalogue no. 4351370. assay ID ss03375006_u1 (Applied Biosystems. Thermo Fisher Scientific)

GAPDH = glyderaldehyde-3-phosphate dehydrogenase; TNF-α = tumour necrosis factor-α; COX-2 =

cyclooxygenase-2; IL-10 = interleukin-10; ZO-1 = zonula occludens-1; OCLN = occludin; CLDN-2 =

claudin-2; CLDN-4 = claudin-4


! 23

Table 3

Piglet body weight (kg) at birth and 7, 14, 21, 28, 35 and 42 days of age1

Treatment group2 p-value Days of age CTRL AB PRO PRO+AB

T3 A4 T*A

0 1.5 (1.4-1.6) 1.5 (1.4-1.6) 1.5 (1.4-1.6) 1.5 (1.4-1.6) 0.89 <0.001 <0.001

7 2.7 (2.6-2.9) 2.8 (2.6-2.9) 2.8 (2.6-3.9) 2.5 (2.4-2.7)

14 5.0 (4.7-5.4) 4.9 (4.6-5.3) 4.9 (4.5-5.2) 4.7 (4.4-5.0)

21 7.5 (7.0-8.0) 7.0 (6.5-7.5) 6.9 (6.4-7.4) 7.0 (6.5-7.5)

28 9.7 (9.0-10.4) 9.2 (8.5-9.8) 8.6 (8.0-9.2) 9.4 (8.8-10.1)

35 9.9 (9.3-10.7) 9.5 (8.9-10.2) 9.1 (8.5-9.8) 9.6 (9.0-10.3)

42 11.3 (10.6-12.2) 10.8 (10.0-11.6) 10.3 (9.6-11.0) 11.1 (10.4-11.9)

1 Values are presented as least square means and 95% confidence intervals (in parentheses). 2 CTRL = control; AB = antibiotic group; PRO = probiotic group; PRO+AB = probiotic+antibiotic group.

Number of observations: CTRL = 414; AB = 395; PRO = 414; PRO+AB = 386. 3 T = treatment group 4 A = age.

! ! ! ! !Table 4

Diarrhoea frequencies during suckling and after weaning1

Treatment group2

Diarrhoea score CTRL AB PRO PRO+AB

Suckling

Grade 1 0.17 0.15 0.28 0.2

Grade 2 0 0.01 0.09 0.07

Weaned

Grade 1 0.49 0.33 0.43 0.62

Grade 2 0.08 0.07 0.08 0.12

1 Values are presented as mean frequencies. 2 CTRL = control; AB = antibiotic group; PRO = probiotic group; PRO+AB = probiotic+antibiotic group.


!24

Tab

le 5

Con

cent

ratio

n of

bio

geni

c am

ines

cad

aver

ine

and

putre

scin

e (m

g/kg

sam

ple)

in il

eal a

nd m

id c

olon

ic d

iges

ta fr

om p

igle

ts a

t 3, 2

8, a

nd 4

2

days

of a

ge 1

Trea

tmen

t gro

up2

p-

valu

e

C

TRL

AB

PR

O

PRO

+AB

#

T3 S*

A4

Cad

aver

ine

0.

39

<0.0

01

Day

3

Ile

um

5.4

(0.1

-18.

3)

2.4

(0-1

1.4)

9.

0 (1

.5-2

7.0)

4.

6 (0

-16.

3)

a

M

id c

olon

20

5.7

(83.

0-50

2.5)

13

6.6

(54.

3-33

5.6)

28

7.0

(116

.4-7

00.1

) 18

6.3

(74.

9-45

5.4)

A

D

ay 2

8

Ileum

9.

6 (2

.4-2

5.5)

5.

2 (0

.4-1

6.2)

14

.8

(4.6

-37.

5)

8.4

(1.8

-22.

8)

a

M

id c

olon

24

.2

(9.1

-57.

1)

15.0

(4

.9-3

7.3)

35

.0

(14.

0-80

.9)

21.6

(7

.9-5

1.4)

B

D

ay 4

2

Ileum

11

5.5

(51.

0-25

6.1)

76

.1

(32.

9-17

0.6)

16

1.8

(72.

0-35

7.8)

10

4.4

(45.

9-23

1.9)

b

Mid

col

on

70.3

(3

0.2-

157.

5)

45.8

(1

9.0-

104.

6)

99.0

(4

3.3-

220.

7)

63.4

(2

7.1-

142.

5)

AB

Putre

scin

e

0.28

<0

.001

D

ay 3

Ileum

19

.2

(7.6

-43.

3)

9.6

(2.9

-23.

2)

22.3

(9

.0-4

9.6)

15

.7

(5.9

-36.

0)

a

M

id c

olon

52

6.9

(253

.8-1

089.

9)

302.

3 (1

44.9

-626

.5)

598.

2 (2

88.1

-123

8.5)

44

4.7

(213

.9-9

20.4

) A

D

ay 2

8

!!

!D

ay 2

8!Ile

um

21.1

(9

.4-4

3.4)

10

.7

(4.0

-23.

3)

24.4

(1

1.0-

50.4

) 17

.3

(7.4

-36.

2)

a

M

id c

olon

18

.9

(8.3

-39.

1)

9.5

(3.4

-20.

8)

21.9

(9

.9-4

4.9)

15

.5

(6.5

-32.

5)

B

Day

42

!

!!

Day

42!

Ileum

60

.1

(30.

0-11

7.4)

33

.2

(15.

9-66

) 68

.7

(34.

4-13

4.0)

50

.3

(24.

8-98

.7)

b

M

id c

olon

44

.8

(21.

9-88

.2)

24.4

(1

1.2-

49.1

) 51

.2

(25.

3-10

0.7)

37

.3

(18.

0-74

.0)

B


!25

1 Val

ues a

re p

rese

nted

as l

east

squa

re m

eans

and

95%

con

fiden

ce in

terv

als (

in p

aren

thes

es).

2 C

TRL

= co

ntro

l; A

B =

ant

ibio

tic g

roup

; PR

O =

pro

biot

ic g

roup

; PR

O+A

B =

pro

biot

ic+a

ntib

iotic

gro

up.

Num

ber o

f pig

lets

: CTR

L=32

; AB

=32;

PR

O=3

1; P

RO

+AB

=32.

3 T

= tre

atm

ent g

roup

4 S*

A =

inte

ract

ion

betw

een

inte

stin

al se

gmen

t and

age

.

a, b

: Ile

al sa

mpl

es. R

ows w

ith d

iffer

ent l

ette

rs, w

ithin

a b

ioge

nic

amin

e gr

oup,

are

sign

ifica

ntly

diff

eren

t (p<

0.05

).

A, B

: Mid

col

on sa

mpl

es. R

ows w

ith d

iffer

ent l

ette

rs, w

ithin

a b

ioge

nic

amin

e gr

oup,

are

sign

ifica

ntly

diff

eren

t (p<

0.05

).

Maj

ority

of a

gmat

ine

and

tyra

min

e m

easu

rem

ents

wer

e ze

ro/b

elow

det

ectio

n le

vel.


!26

Tab

le 6

Con

cent

ratio

n of

org

anic

aci

ds (m

mol

/kg

sam

ple)

in il

eal a

nd c

olon

ic d

iges

ta fr

om p

igle

ts 3

, 28,

and

42

days

of a

ge1

Trea

tmen

t gro

up2

p-

valu

e

C

TRL

AB

PR

O

PRO

+AB

#

T3 S*

A4

Ace

tic a

cid

0.

55

<0.0

01

Day

3

Ile

um

0.9

(0.6

-3.9

) 1.

0 (0

.6-4

.4)

0.8

(0.5

-3.4

) 0.

8 (0

.5-3

.0)

a

M

id c

olon

4.

1 (2

.6-2

1.8)

4.

5 (2

.9-2

4.2)

3.

7 (2

.4-1

9.7)

3.

4 (2

.2-1

7.8)

A

D

ay 2

8

Ileum

1.

5 (1

.0-6

.9)

1.7

(1.1

-7.7

) 1.

4 (0

.9-6

.2)

1.3

(0.8

-5.5

) ab

M

id c

olon

5.

7 (3

.9-2

9.7)

6.

4 (4

.3-3

3.0)

5.

2 (3

.6-2

7.0)

4.

8 (3

.2-2

4.5)

A

D

ay 4

2

Ileum

1.

9 (1

.3-8

.8)

2.1

(1.4

-9.9

) 1.

7 (1

.2-7

.9)

1.5

(1.1

-7.1

) b

Mid

col

on

20.6

(1

4.0-

110.

1)

22.8

(1

5.5-

121.

9)

18.8

(1

2.7-

100.

3)

17.1

(1

1.6-

91.1

) B

Pr

opio

nic

acid

0.56

<0

.001

D

ay 3

Ileum

0.

4 (0

.4-0

.7)

0.4

(0.4

-0.9

) 0.

4 (0

.4-0

.7)

0.4

(0.4

-0.5

)

Mid

col

on

3.3

(2.0

-5.2

) 3.

9 (2

.4-6

.2)

3.5

(2.2

-5.6

) 2.

9 (1

.7-4

.6)

A

Day

28

Ile

um

0.5

(0.4

-1.0

) 0.

4 (0

.4-0

.9)

0.6

(0.4

-1.1

) 0.

4 (0

.4-0

.8)

M

id c

olon

6.

5 (4

.4-9

.6)

7.7

(5.2

-11.

2)

7.0

(4.7

-10.

2)

5.8

(3.9

-8.5

) B

D

ay 4

2

Ileum

0.

4 (0

.4-0

.5)

0.4

(0.4

-0.9

) 0.

4 (0

.4-0

.5)

0.4

(0.4

-0.4

)

Mid

col

on

28.5

(1

9.8-

40.8

) 33

.3

(23.

2-47

.7)

30.2

(2

1-43

.4)

25.3

(1

7.6-

36.4

) C

B

utyr

ic a

cid

0.

93

- M

id c

olon

Day

3

2.7

(0.3

-5.1

) 2.

0 (0

-4.4

) 2.

0 (0

-4.4

) 2.

1 (0

-4.4

) A

D

ay 2

8 4.

3 (2

.2-6

.4)

3.6

(1.5

-5.7

) 3.

6 (1

.4-5

.7)

3.6

(1.5

-5.8

) A

D

ay 4

2 11

.2

(9.1

-13.

3)

10.5

(8

.4-1

2.6)

10

.5

(8.4

-12.

6)

10.6

(8

.4-1

2.7)

B

0.87

-


!27

Val

eric

aci

d M

id c

olon

Day

3

0.9

(0.1

-1.7

) 0.

5 (0

-1.3

) 0.

6 (0

-1.4

) 0.

6 (0

-1.3

) A

D

ay 2

8 2.

0 (1

.3-2

.8)

1.7

(1.0

-2.4

) 1.

8 (1

.1-2

.5)

1.7

(1.0

-2.5

) B

D

ay 4

2 2.

8 (2

.1-3

.5)

2.5

(1.7

-3.2

) 2.

5 (1

.8-3

.3)

2.5

(1.8

-3.2

) C

Is

o-bu

tyric

aci

d

0.90

-

Mid

col

on

D

ay 3

0.

8 (0

.4-1

.3)

0.7

(0.3

-1.2

) 0.

9 (0

.4-1

.3)

0.7

(0.3

-1.2

)

Day

28

1.2

(0.8

-1.6

) 1.

2 (0

.8-1

.6)

1.3

(0.9

-1.7

) 1.

1 (0

.7-1

.5)

D

ay 4

2 1.

3 (0

.9-1

.7)

1.3

(0.9

-1.7

) 1.

4 (1

.0-1

.8)

1.2

(0.8

-1.6

)

Iso-

vale

ric a

cid

0.

91

- M

id c

olon

Day

3

0.7

(0.7

-0.7

) 0.

7 (0

.7-0

.8)

0.7

(0.7

-0.7

) 0.

7 (0

.7-0

.7)

A

Day

28

0.7

(0.7

-1.2

) 0.

8 (0

.7-1

.3)

0.7

(0.7

-1.2

) 0.

7 (0

.7-1

.1)

AB

D

ay 4

2 0.

8 (0

.7-1

.2)

0.9

(0.7

-1.4

) 0.

8 (0

.7-1

.3)

0.7

(0.7

-1.1

) B

La

ctic

aci

d

0.41

-

Ileum

Day

3

10.9

(7

.0-1

6.3)

10

.7

(6.9

-16.

0)

7.7

(4.7

-11.

8)

9.1

(5.7

-13.

7)

A

Day

28

10.1

(6

.8-1

4.6)

9.

9 (6

.6-1

4.3)

7.

0 (4

.4-1

0.6)

8.

3 (5

.5-1

2.2)

A

D

ay 4

2 38

.4

(27.

7-52

.7)

37.7

(2

7.2-

51.8

) 28

.6

(20.

4-39

.7)

32.8

(2

3.6-

45.2

) B

1 V

alue

s are

pre

sent

ed a

s lea

st sq

uare

mea

ns a

nd 9

5% c

onfid

ence

inte

rval

s (in

par

enth

eses

).

2 CTR

L =

cont

rol;

AB

= a

ntib

iotic

gro

up; P

RO

= p

robi

otic

gro

up; P

RO

+AB

= p

robi

otic

+ant

ibio

tic g

roup

.

Num

ber o

f pig

lets

: CTR

L=32

; AB

=32;

PR

O=3

1; P

RO

+AB

=32.

3 T

= tre

atm

ent g

roup

. 4 S*

A =

inte

ract

ion

betw

een

inte

stin

al se

gmen

t and

age

.

a, b

: Ile

al sa

mpl

es. R

ows w

ith d

iffer

ent l

ette

rs, w

ithin

an

acid

gro

up, a

re si

gnifi

cant

ly d

iffer

ent (p<

0.05

).

A, B

, C: M

id c

olon

sam

ples

. Row

s with

diff

eren

t let

ters

, with

in a

n ac

id g

roup

, are

sign

ifica

ntly

diff

eren

t (p<

0.05

).


!28

Tab

le 7

Con

cent

ratio

n of

shor

t-cha

in fa

tty a

cids

(mm

ol/ k

g sa

mpl

e) in

faec

al m

atte

r fro

m p

igle

ts 7

, 28,

and

42

days

of a

ge1

Tr

eatm

ent g

roup

2

p-va

lues

A

ge

CTR

L A

B

PRO

PR

O+A

B

# T3

A4

T*A

A

cetic

aci

d

0.32

<0

.001

0.

42

7 28

.8

(23.

8-33

.7)

24.6

(1

9.7-

29.5

) 29

.0

(24.

2-33

.9)

29.0

(2

4.2-

33.8

) a

28

34

.9

(30.

2-39

.6)

30.7

(2

6.0-

35.3

) 35

.2

(30.

4-39

.9)

35.1

(3

0.4-

39.7

) b

42

59

.9

(55.

3-64

.6)

55.7

(5

1.1-

60.4

) 60

.2

(55.

5-64

.9)

60.1

(5

5.4-

64.8

) c

Pr

opio

nic

acid

0.76

<0

.001

0.

03

7 8.

6A

(4.7

-12.

4)

7.8A

(4

.1-1

1.5)

11

.6A

(8

.0-1

5.2)

7.

8A

(4.3

-11.

2)

28

14.2

A

(10.

9-17

.5)

10.8

A

(7.5

-14.

1)

14.9

AB

(1

1.4-

18.4

) 12

.9A

(9

.6-1

6.2)

42

24

.8B

(2

1.5-

28.0

) 24

.5B

(2

1.2-

27.8

) 19

.5B

(1

6.2-

22.8

) 25

.4B

(2

2.0-

28.8

)

B

utyr

ic a

cid

0.

84

<0.0

01

0.00

4 7

3.7A

b (0

.6-6

.9)

5.4A

ab

(2.4

-8.5

) 10

.2a

(7.3

-13.

2)

5.2A

ab

(2.4

-8.1

)

28

8.

2AB

(5

.4-1

0.9)

5.

2A

(2.4

-7.9

) 7.

6 (4

.7-1

0.4)

6.

6A

(3.8

-9.3

)

42

12

.1B

(9

.4-1

4.9)

11

.2B

(8.4

-13.

9)

8.0

(5.3

-10.

7)

11.7

B

(8.9

-14.

5)

Val

eric

aci

d

0.52

0.

11

0.09

7

2.2

(1.6

-2.8

) 1.

9 (1

.3-2

.5)

2.4

(1.8

-3.0

) 2.

1 (1

.5-2

.7)

28

2.7

(2.1

-3.3

) 2.

4 (1

.8-2

.9)

2.9

(2.3

-3.4

) 2.

6 (2

.0-3

.1)

42

2.7

(2.2

-3.3

) 2.

4 (1

.9-3

.0)

2.9

(2.3

-3.5

) 2.

6 (2

.0-3

.2)

Iso-

buty

ric a

cid

0.

63

0.91

0.

01

7 1.

6a (0

.9-2

.4)

1.6a

(0.9

-2.4

) 3.

1Ab

(2.4

-3.8

) 1.

7a (1

.0-2

.4)

28

2.3

(1.6

-3.0

) 1.

8 (1

.1-2

.4)

2.3A

B

(1.6

-3.0

) 2.

0 (1

.3-2

.7)

42

2.1

(1.5

-2.8

) 2.

4 (1

.8-3

.1)

1.8B

(1

.1-2

.4)

2.2

(1.6

-2.9

)

Is

o-va

leric

aci

d

0.68

0.

29

0.03

7

1.3

(0.6

-2.0

) 1.

4 (0

.8-2

.1)

2.6A

(1

.9-3

.3)

1.4

(0.8

-2.0

)

28

2.

2 (1

.5-2

.8)

1.7

(1.0

-2.3

) 2.

1AB

(1.5

-2.8

) 1.

8 (1

.2-2

.4)

42

1.6

(1.0

-2.2

) 1.

9 (1

.3-2

.5)

1.3B

(0

.7-1

.9)

1.7

(1.1

-2.4

)


!29

1 Val

ues a

re p

rese

nted

as l

east

squa

re m

eans

and

95%

con

fiden

ce in

terv

als (

in p

aren

thes

es).

2 C

TRL

= co

ntro

l; A

B =

ant

ibio

tic g

roup

; PR

O =

pro

biot

ic g

roup

; PR

O+A

B =

pro

biot

ic+a

ntib

iotic

gro

up.

Num

ber o

f pig

lets

: CTR

L=49

; AB

=50;

PR

O=4

9; P

RO

+AB

=51.

3 T

= tre

atm

ent g

roup

4 A

= a

ge.

a, b

, c: R

ows w

ith d

iffer

ent l

ette

rs a

re si

gnifi

cant

ly d

iffer

ent (p<

0.05

). a,

b Val

ues w

ith d

iffer

ent s

uper

scrip

ts w

ithin

a ro

w a

re si

gnifi

cant

ly d

iffer

ent.

A,B

: Val

ues w

ith d

iffer

ent s

uper

scrip

ts w

ithin

a c

olum

n ar

e si

gnifi

cant

ly d

iffer

ent.


! 30

Table 8 List of the OTUs that were significantly different between treatment groups1

Age Treatments

compared Total OTUs

Significantly different OTU(s)

OTU classification Log2 fold change

Ileum 3 PRO+AB vs. PRO 182 OTU_143 Bacteroides 7.2 28 PRO+AB vs. CTRL 189 OTU_4 Streptococcus 4.3 42 AB vs. PRO 95 OTU_362 Clostridium 5.9 Mid colon

3 AB vs. PRO 259 OTU_22 Fusobacterium 9.3 PRO+AB vs. AB 266 OTU_22 Fusobacterium 7.9 PRO+AB vs. PRO 230 OTU_872 Order Clostridiales -5.4 OTU_387 Streptococcus -6.4 AB vs. PRO 257 OTU_22 Fusobacterium 9.2 OTU_43 Prevotella -7.7 28 AB vs. PRO 734 OTU_13

9 more.. Bacteroides 7.7

AB vs. CTRL 719 OTU_273 OTU_219 OTU_262

Campylobacter Family Ruminococcaceae Order Tremblayales

9.5 6.4 6.5

PRO vs. CTRL 629 OTU_13 Bacteroides 9.2 PRO+AB vs. AB 694 OTU_386

OTU_13 Anaerovibrio Bacteroides

8.2 5.7

AB vs. PRO OTU_13 Bacteroides 7.7 OTU_2428 Lachnospiraceae 9.8 OTU_179 Rikellaceae 10 OTU_82 Coriobacteriaceae 8.2 OTU_2 Fusobacterium 6.9 OTU_666 Bacteroides 8.1 OTU_297 Bacteroides 9.9 OTU_36 Oscillospira 5.8 OTU_359 Veillonellaceae 8.7 42 AB vs. CTRL 924 OTU_86 Order RF39 7.6 PRO+AB vs. AB 914 OTU_86 Order RF39 6.3 PRO+AB vs. CTRL 901 OTU_33

OTU_9 Lactobacillus Lactobacillus

-5.3 -5.2

Faeces

7 AB vs. CTRL 451 OTU_3118 Family Ruminococcaceae 6.5 PRO+AB vs. AB 407 OTU_340

OTU_46 OTU_84 OTU_299 OTU_1880

Bacteroides Bacteroides Bacteroides Dorea Bacillus

6.7 -4.2 4.7 3.3 5.6


! 31

OTU_545 Family Lachnospiraceae 3.3 PRO+AB vs. PRO 406 OTU_24 Fusobacterium -6.2 PRO+AB vs. CTRL 440 OTU_24

OTU_79 Fusobacterium Order Bacteroidales

-4.6 -4.5

AB vs. PRO 423 OTU_3118 Family Ruminococcaceae 4.4 28 AB vs. CTRL 1195 OTU_50

OTU_2333 Megasphaera Oscillospira

3.2 -2.8

PRO vs. CTRL 1071 OTU_129 20 more…

Bacteroides -6.2

PRO+AB vs. AB 1194 OTU_762 OTU_166

Ruminococcus Anaerovibrio

-6.2 4.8

PRO+AB vs. PRO 1099 OTU_135 178 more…

Bacteroides 6.4

PRO+AB vs. CTRL 1175 OTU_268 OTU_891 OTU_402 OTU_2333

Bacteroides Family S24-7 Bacteroides Oscillospira

3.2 4.5 3.5 3.8

AB vs. PRO 1114 OTU_404 Family Mogibacteriaceae -6.0 182 more… 42 AB vs. CTRL 1264 OTU_391 Order Bacteroidales 4.7 PRO vs. CTRL 1252 OTU_691

OTU_5 YRC22 Lactobacillus

2.1 -3.2

PRO+AB vs. AB 1306 OTU_865 OTU_391 OTU_121

Ruminococcus Order Bacteroidales Family Christensenellaceae

-6.0 4.2 -5.0

PRO+AB vs. PRO 1277 OTU_348 Family Veillonellaceae 5.3 AB vs. PRO 1332 OTU_391 Order Bacteroidales 6.1 OTU_31 p-75-a5 -3.3 OTU_256 Faecalibacterium 2.7 OTU_689 Lachnospiraceae -6.0 OTU_62 Coriobacteriaceae -2.8 OTU_720 Anaerovibrio -5.2 OTU_126 Peptococcus -2.5 OTU_273 Campylobacter 2.8

1 CTRL = control; AB = antibiotic group; PRO = probiotic group; PRO+AB = probiotic+antibiotic group.


! 32

Figures:

Fig. 1. Enumeration of selected bacterial groups (A-F) in digesta (log cfu/g) from the stomach,

ileum, caecum and mid colon sampled day 3, 28 and 42 from piglets administered gentamicin (AB;

n=71); piglets administered Bacillus spores (PRO; n=68); piglets administered both gentamicin and

Bacillus spores (PRO+AB; n=71) and control piglets not receiving gentamicin or Bacillus spores

(CON; n=72). Values are presented as least-square means and the 95% confidence intervals

presented as vertical bars. Bars embraced by horisontal brackets market by * (0.01≤p<0.05), **

(0.001≤p<0.01) or *** (p<0.001) are significantly different.

Fig. 2. Heatmaps of faecal samples collected at 7, 28 and 42 days of age from piglets administered

gentamicin (AB; n=53), Bacillus spores (PRO; n=50), both gentamicin and Bacillus spores

(PRO+AB; n=53) and control piglets not receiving gentamicin or Bacillus spores (CTRL; n=51).

Heatmaps show the relative abundances (%) of (A) the ten most abundant phyla and (B) the 20

most abundant genera in faecal samples. Colours represent the relative abundances.

Fig. 3. Observed and estimated species richness and Shannon diversity index of faecal samples

collected at 7, 28 and 42 days of age from piglets administered gentamicin (AB; n=53), Bacillus

spores (PRO; n=50), both gentamicin and Bacillus spores (PRO+AB; n=53) and control piglets not

receiving gentamicin or Bacillus spores (CTRL; n=51). Boxplots show the (A) observed and (B)

estimated species richness and (C) Shannon diversity index.

Fig. 4. Principal Component Analysis (PCA) of square root transformed OTU abundances

originating from faecal samples (n=207) displaying PC1 and PC2. (A) Day 7 samples; (B) Day 28

samples; (C) Constrained PCA on day 28 samples; (D) Day 42 samples; (E) Constrained PCA on

day 42 samples. Points are coloured according to treatment. AB: Piglets administered gentamicin;

PRO: Piglets administered Bacillus spores; PRO+AB: Both administered gentamicin and Bacillus

spores; CONTROL: Control piglets not receiving gentamicin or Bacillus spores.

Fig. 5. Heatmaps of ileal and colonic digesta samples collected at 3, 28 and 42 days of age from

piglets administered gentamicin (AB; n=33), Bacillus spores (PRO; n=33), both gentamicin and

Bacillus spores (PRO+AB; n=31) and control piglets not receiving gentamicin or Bacillus spores

(CTRL; n=29). Heatmaps show the relative abundances (%) of (A) the eight most abundant phyla in


! 33

ileal digesta, (B) the 20 most abundant genera in ileal digesta, (C) the eight most abundant phyla in

colonic digesta and (D) the 20 most abundant genera in colonic digesta. Colours represent the

relative abundances.

Fig. 6. Observed and estimated species richness and Shannon diversity index of ileal and colonic

digesta samples collected at 3, 28 and 42 days of age from piglets administered gentamicin (AB;

n=33), Bacillus spores (PRO; n=33), both gentamicin and Bacillus spores (PRO+AB; n=31) and

control piglets not receiving gentamicin or Bacillus spores (CONTORL; n=29). Boxplots show the

(A) observed and (B) estimated species richness and (C) Shannon diversity index.

Fig. 7. (A) Principal Component Analysis (PCA) of square root transformed OTU abundances in

colonic digesta (n=65) displaying PC1 and PC2. Points are coloured according to treatment and

grouped according to age. (B) Constrained PCA of square root transformed OTU abundances in

colonic digesta day 28 (n=17). Points are coloured for treatment. AB: Piglets administered

gentamicin; PRO: Piglets administered Bacillus spores; PRO+AB: Both administered gentamicin

and Bacillus spores; CONTORL: Control piglets not receiving gentamicin or Bacillus spores.

Fig. 8. Gene expression levels of (A) TNF-α, (B) IL-10, (C) Cyclo-oxygenase-2, (D) ZO-1, (E)

OCLN, (F) CLDN-4 and (G) CLDN-2 in ileal tissue collected day 28 (n=22) and 42 (n=34) from

piglets administered gentamicin (AB); piglets administered Bacillus spores (PRO); piglets

administered both gentamicin and Bacillus spores (PRO+AB) and control piglets not receiving

gentamicin or Bacillus spores (CONTROL), that has either been left untreated or stimulated with

LPS. Values are presented as least-square means and the 95% confidence intervals presented as

vertical bars. Bars embraced by horisontal brackets market by * (0.01≤p<0.05), ** (0.001≤p<0.01)

or *** (p<0.001) are significantly different.

Fig. S1. Principal Component Analysis of square root transformed OTU abundances in faeces

(n=207) displaying PC1 and PC2. Points are coloured according to age.

Fig. S2. Principal Component Analysis of square root transformed OTU abundances in ileal (n=61)

and colonic (n=65) digesta displaying PC1 and PC2. Points are coloured according to age and

shaped according to segment.


1 3 5 7

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

4.5

5.0

5.5

6.0

6.5

7.0

7.5

8.0

8.5

9.0

9.5

10.0

3 28 42 3 28 42 3 28 42 3 28 42Age (days)

Log

cfu/

g di

gest

a Treatment

AB

CON

PRO

PRO+AB

Total anaerobic bacteria

***

*** ** **

Log

cfu/

g di

gest

a

Age (days)

Total anaerobic bacteriaStomach Ileum Caecum Mid colon

1 3 5 7

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

4.5

5.0

5.5

6.0

6.5

7.0

7.5

8.0

8.5

9.0

9.5

10.0

3 28 42 3 28 42 3 28 42 3 28 42Age (days)

Log

cfu/

g di

gest

a Treatment

AB

CON

PRO

PRO+AB

EnterobacteriaceaeEnterobacteriaceaeStomach Ileum Caecum Mid colon

*

**

*** ****** ***

Age (days)Lo

g cf

u/g

dige

sta

1 3 5 7

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

4.5

5.0

5.5

6.0

6.5

7.0

7.5

8.0

8.5

9.0

9.5

10.0

3 28 42 3 28 42 3 28 42 3 28 42Age (days)

Log

cfu/

g di

gest

a Treatment

AB

CON

PRO

PRO+AB

Haemolytic bacteriaHaemolytic bacteriaStomach Ileum Caecum Mid colon

Log

cfu/

g di

gest

a

Age (days)

******

******

****** *** ***

1 3 5 7

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

4.5

5.0

5.5

6.0

6.5

7.0

7.5

8.0

8.5

9.0

9.5

10.0

3 28 42 3 28 42 3 28 42 3 28 42Age (days)

Log

cfu/

g di

gest

a Treatment

AB

CON

PRO

PRO+AB

Lactic acid bacteria

Log

cfu/

g di

gest

a

Age (days)

Stomach Ileum Caecum Mid colon

Lactic acid bacteria

*** * * ****

***

1 3 5 7

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

4.5

5.0

5.5

6.0

6.5

7.0

7.5

8.0

3 28 42 3 28 42 3 28 42 3 28 42Age (days)

Log

cfu/

g di

gest

a Treatment

AB

CON

PRO

PRO+AB

Clostridium perfringensClostridium perfringensStomach Ileum Caecum Mid colon

Age (days)

Log

cfu/

g di

gest

a

*

******

***

***

***

***

***

***

1 3 5 7

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

4.5

5.0

5.5

6.0

6.5

3 28 42 3 28 42 3 28 42 3 28 42Age (days)

Log

cfu/

g di

gest

a Treatment

AB

CON

PRO

PRO+AB

BacillusBacillus spp. sporesStomach Ileum Caecum Mid colon

Age (days)

Log

cfu/

g di

gest

a

*********

***

(A) (B)

(D)(C)

(E) (F)

Figure 1


7 28 42

0.1

0

0.1

0.1

0.9

0

14.3

33.7

0

50.2

0.4

0

19.9

43.1

0

0

0.6

0

35.8

0

40.4

0.2

0

0.1

0

15.1

0

0

1.4

42.6

0

45.8

0.2

0

0

0

1

0

16.6

35.9 54.1

0.3

30

2

0.7

1.1

2.8

1.4

0.7

5.3

44.7

0.5

1.2

10

0.1

37.7

0.7

2.3

1.4

0.7

17.2

38.1

39

0.6

1.1

0.5

0.5

0.2

0.3

1.8

0.6

7.8

48.3

0.4

36

0.6

0.8

0.4

2

1.2 1.5

0

0.2

64

0.4

29.2

1.5

0.7

0

1.8

1.9

63.5

0.6

1

0

0

0.6

30.6

1.2

0

1.1

4.7

0.1

59.4

32.1

0.5

0

0

0.2

1.2

0

1.3

2.5

0

0.2

61.2

0.4

32.2

0.6

1

Lentisphaerae

Cyanobacteria

Synergistetes

Tenericutes

Actinobacteria

Spirochaetes

Proteobacteria

Fusobacteria

Bacteroidetes

Firmicutes

7 A

B

7 C

ONT

ROL

7 P

RO

7 P

RO+A

B

28 A

B

28 C

ONT

ROL

28 P

RO

28 P

RO+A

B

42 A

B

42 C

ONT

ROL

42 P

RO

42 P

RO+A

B

0.1

1.0

10.0

% ReadAbundance

Day 7 Day 28 Day 42

AB CTRL PRO PRO+AB AB CTRL PRO PRO+AB AB CTRL PRO PRO+AB

7 28 42

0.2

18.8

0

1.5

0

0.6

0.4

9.6

2.11.2

0

0

35

0

0

14.3

0.3

0.9

0

1.3

0.2

17.5

0

0.5

1.21.6

28.1

0

0.1

1.6

19.9

0.4

8

2.2

0

0

0.7

0.1

0.3

0.4

0

0

0.8

27.4

0

22.8

2.7

15.1

0.4

8.9

2.6

1.1

0

1.3

0

0

0.2

0.2

0

0.1

32.4

0.3

2.1

0

8.7

1.1

0.4

1.4

0

20.2

0

0.4

16.6

0.2

0.1

0.4

1.7

0.9

0

0

3.5

1.6

1.7

4.3

2.4

3.6

0.2

0.1

0.3

0.3

1.6

6.2

0.90.5

4.6

0.1

0.1

5.3

1.8

3.5

7.8

1.7

2.6

4.7

0.2

0.1

1.10.3

0.6

3.5

0.6

0.2

0.1

10

1

12.5

2.96

0.3

1.3

2.72.6

0.4

0.70.9

2.7

6

0.6

6

0.1

17.2

0.1

17

4

0.9

5.8

0.1

0.4

0.4

0.7

1.6

2.1

8.5

2.7

0.1

0.6

10.8

3.8

0.2

1.8

1.7

3.4

0.4

3.4

0.2

0.2

7.8

0.2

0.9

1.3

2.2

0.5

10.3

0.7

0.6

1.3

1.1

0.8

3.1

12.2

2.8

0.3

1.2

3.7

2.2

0.6

2.9

2

0.7

0.2

4.1

0.9

0.1

0.7

3.5

13.2

1.6

1.8

15.7

0.61.7

0.8

0.4

2.4

3.3

0.2

0

1.92.8

1.2

1.70.25.7

1.8

1.3

0.7

0.61.7

0.7

0.5

1.7

6.5

0.3

0.114.1

1.9

0.4

2.6

3.3

3.31.6

2.5

0.4

0.2

0.4

3.9

0.2

0.2

1.7

0.6

16.4

1.9

1.1

0.8

0.8

2.8

2.2

13.3

3.2

1.5Firmicutes; Anaerovibrio

Proteobacteria; CampylobacterBacteroidetes; f__p−2534−18B5_OTU_58

Firmicutes; SMB53Firmicutes; Clostridium

Firmicutes; MegasphaeraFirmicutes; Streptococcus

Firmicutes; RoseburiaBacteroidetes; Parabacteroides

Firmicutes; p−75−a5Firmicutes; Coprococcus

Firmicutes; 02d06Bacteroidetes; o__Bacteroidales_OTU_17

Bacteroidetes; [Prevotella]Firmicutes; Ruminococcus

Firmicutes; OscillospiraFusobacteria; Fusobacterium

Bacteroidetes; PrevotellaFirmicutes; Lactobacillus

Bacteroidetes; Bacteroides

7 A

B

7 C

ONT

ROL

7 P

RO

7 P

RO+A

B

28 A

B

28 C

ONT

ROL

28 P

RO

28 P

RO+A

B

42 A

B

42 C

ONT

ROL

42 P

RO

42 P

RO+A

B

0.1

1.0

10.0

% ReadAbundance

Day 7 Day 28 Day 42


Bacteroidetes; order Bacteroidales OTU 17

Bacteroidetes; family p-2534-18B5 OTU 58

(A)

(B)

Figure 2


7 28 42

125

250

375

500

625

750

875

1000

AB

CONTROLPRO

PRO+AB AB

CONTROLPRO

PRO+AB AB

CONTROLPRO

PRO+AB

Treatment

Obs

erve

d nu

mbe

r of O

TUs

Obs

erve

d nu

mbe

r of O

TUs


Treatment

Day 7 Day 28 Day 42

7 28 42

125

250

375

500

625

750

875

1000

1125

1250

AB

CONTROLPRO

PRO+AB AB

CONTROLPRO

PRO+AB AB

CONTROLPRO

PRO+AB

Treatment

Estim

ated

num

ber o

f OTU

sE

stim

ated

num

ber o

f OTU

s


Treatment

Day 7 Day 28 Day 42

7 28 42

1.50

1.75

2.00

2.25

2.50

2.75

3.00

3.25

3.50

3.75

4.00

4.25

4.50

4.75

5.00

5.25

AB

CONTROLPRO

PRO+AB AB

CONTROLPRO

PRO+AB AB

CONTROLPRO

PRO+AB

Treatment

Dive

rsity

inde

xD

iver

sity

inde

x


Treatment

Day 7 Day 28 Day 42

(B)

(A)

(C)

Figure 3


−6

−3

0

3

6

−3 0 3 6PC1 [13.9%]

PC2

[10.

8%]

Treatment

AB

CONTROL

PRO

PRO+AB

Faeces day 7

PC

2 - v

aria

tion

expl

aine

d 10

.8%


PCA - faeces day 7

−4

0

4

−5.0 −2.5 0.0 2.5 5.0PC1 [18.2%]

PC2

[11.

7%]

Treatment

AB

CONTROL

PRO

PRO+AB

Faeces day 28

PC

2 - v

aria

tion

expl

aine

d 11

.7%


PCA - faeces day 28

−10

−5

0

5

−4 0 4 8RDA1 [8.2%]

RDA2

[2.1

%]

Treatment

AB

CONTROL

PRO

PRO+AB

Faeces day 28

RD

A2

- var

iatio

n ex

plai

ned

2.1%

RDA1 - variation explained 8.2%

RDA - faeces day 28

−3

0

3

6

−3 0 3PC1 [16.4%]

PC2

[9.1

%]

Treatment

AB

CONTROL

PRO

PRO+AB

Faeces day 42

PC

2 - v

aria

tion

expl

aine

d 9.

1%


PCA - faeces day 42

−5

0

5

−5 0 5 10RDA1 [4%]

RDA2

[2.2

%]

Treatment

AB

CONTROL

PRO

PRO+AB

Faeces day 42

RD

A2

- var

iatio

n ex

plai

ned

2.2%

RDA1 - variation explained 4%

RDA - faeces day 42

(A)

(E)(D)

(C)(B) −6

−3

0

3

6

−3 0 3 6PC1 [13.9%]

PC2

[10.

8%]

Treatment

AB

CONTROL

PRO

PRO+AB

Faeces day 7

PC

2 - v

aria

tion

expl

aine

d 10

.8%


PCA - faeces day 7

−4

0

4

−5.0 −2.5 0.0 2.5 5.0PC1 [18.2%]

PC2

[11.

7%]

Treatment

AB

CONTROL

PRO

PRO+AB

Faeces day 28

PC

2 - v

aria

tion

expl

aine

d 11

.7%


PCA - faeces day 28

−10

−5

0

5

−4 0 4 8RDA1 [8.2%]

RDA2

[2.1

%]

Treatment

AB

CONTROL

PRO

PRO+AB

Faeces day 28

RD

A2

- var

iatio

n ex

plai

ned

2.1%


RDA - faeces day 28

−3

0

3

6

−3 0 3PC1 [16.4%]

PC2

[9.1

%]

Treatment

AB

CONTROL

PRO

PRO+AB

Faeces day 42

PC

2 - v

aria

tion

expl

aine

d 9.

1%


PCA - faeces day 42

−5

0

5

−5 0 5 10RDA1 [4%]

RDA2

[2.2

%]

Treatment

AB

CONTROL

PRO

PRO+AB

Faeces day 42

RD

A2

- var

iatio

n ex

plai

ned

2.2%

RDA1 - variation explained 4%

RDA - faeces day 42

(A)

(E)(D)

(C)(B)

Figure 4


328

42

0.5

2.2

0.2 01.1

95.1 00.8

0.3

0.5 0 0086.5

0.7

12.1

0.5

1.2 089.4 6 3 00

7.8

0.5

1.1

88.8 0 01.7

0.1

0.3

0.4

0.5

0.2 0 098.3

0.1

0.1

0.2 00 098.4

0.3

0.8

096.3

0.1

0.8

1.2

1.6 00

0.3

0.2

3.1

0.7

95.6 0 00

0.2 00.3

0.20 0099.3

0.2

0.2 099.3 0 0.2 0 0

99.1 00 0.1

0.4

0.3 00

01.6

0.2 00.6

0.2

97.3 0

Tene

ricut

es

Cyan

obac

teria

TM7

Actin

obac

teria

Bact

eroi

dete

s

Prot

eoba

cter

ia

Fuso

bact

eria

Firm

icute

s

AB 3

CONTROL 3

PRO 3

PRO+AB 3

AB 28

CONTROL 28

PRO 28

PRO+AB 28

AB 42

CONTROL 42

PRO 42

PRO+AB 42

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Ileum

Day

3D

ay 2

8D

ay 4

2Ile

um

AB

C

TRL

PR

O

PR

O+A

B

AB

C

TRL

PR

O

PR

O+A

B

AB

C

TRL

PR

O

PR

O+A

B

328

42

0.2

0.70 057.6

0.1

0.7

10.4 00.7

1.9

0.2

0.1

0.7 00.1

2.1

1.2

4.8

12.9

0.1

0.2057.2

6.8 00.1

3.7 12 0.3

0.1

0.2

0.1

0.1

0.3

12.1 10.31 2.3

1.1

0.6

37.7 00.3

8.5

0.2

3.7

0.20 0.7

0.5

27.7

2.2 05.5 0 1.2

0.3

5.9

0.3

7.7

1.1

2.5

0.3

0.3

42.1

0.1

14.2 00.2

0.1

1.3

20.8 0 0 0.2

0.8

4.5

0.1

0.2

1.3

33.6

0.4

0.1

3.2 07.7

3.5

4.7 0 0.1

0.5

3.79 0.2

26.3 00.1

2.2

6.8 1 0.1 0 0.1

0.8

0.1

1.1

53.7

0.2

0.1

1.2

0.1

2.6

28.5

0.1 00.6

0.3

1.5

0.5

3.6 0 0.1

2.6

53.2

0.3

0.5

1.4

0.4

0.7

16.8

0.4

0.9

13.8 01.5 000.1

0.1

3.3

0.2

61.3 00.2

1.4

0.81 02.6

1.1

4.1 00.3

1.1

2.5

8.3

7.9

0.1

064.1 0 00 0.2

0.5

0.1 00.1 00.6

0.1

3.6 0 00021.3 8

34.6

4.1 00.3

50.1 001.9

0.1 00.1 0 0.2

0.2 00.3 06.9 00

000.8 031 2.5

0.3 0 00.8

0.1

0.5

47.9 00.1

4.4

9.5

0.1 00

55 0026.1

0.1 00.5

1.4 00.2

0.3 02.8

0.5

0.2

10.3 0 0.60 0

Prot

eoba

cter

ia; A

ctin

obac

illus

Prot

eoba

cter

ia; S

utte

rella

Actin

obac

teria

; Cor

yneb

acte

rium

Firm

icute

s; G

emel

laBa

cter

oide

tes;

Bac

tero

ides

Firm

icute

s; A

cidam

inoc

occu

sBa

cter

oide

tes;

Pre

vote

llaPr

oteo

bact

eria

; f__

Ente

roba

cter

iace

ae_O

TU_1

36Fi

rmicu

tes;

o__

Clos

tridi

ales

_OTU

_872

Firm

icute

s; V

eillo

nella

Firm

icute

s; M

egas

phae

raFi

rmicu

tes;

Tur

iciba

cter

Firm

icute

s; M

itsuo

kella

Fuso

bact

eria

; Fus

obac

teriu

mFi

rmicu

tes;

Stre

ptoc

occu

sFi

rmicu

tes;

Sar

cina

Firm

icute

s; C

lost

ridiu

mFi

rmicu

tes;

SM

B53

Firm

icute

s; 0

2d06

Firm

icute

s; L

acto

bacil

lus

AB 3

CONTROL 3

PRO 3

PRO+AB 3

AB 28

CONTROL 28

PRO 28

PRO+AB 28

AB 42

CONTROL 42

PRO 42

PRO+AB 42

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Ileum

Day

28

Day

3D

ay 2

8D

ay 4

2Ile

um

AB

C

TRL

P

RO

PR

O+A

B

AB

C

TRL

P

RO

PR

O+A

B

AB

C

TRL

P

RO

PR

O+A

B

Firm

icut

es; o

rder

Clo

strid

iale

s O

TU 8

72

Pro

teob

acte

ria; f

amily

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0.2

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0.7

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4.6

0.3

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63.3

0.4

21.4

0.3

72.8

0.5

1.7

1.8

0.8

64.4

0.3

0.6

3.3

28.6

0.4

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14.6

0.4

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29.7

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1.2

Cyan

obac

teria

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s

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AB 28

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AB 42

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PRO 42

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0.1

1.0

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ead

Abu

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3.4

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3.5

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0.10 0.6

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0.6

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Bact

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eaFi

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p−7

5−a5

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naer

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s; 0

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s; L

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ia; F

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AB 3

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AB 28

CONTROL 28

PRO 28

PRO+AB 28

AB 42

CONTROL 42

PRO 42

PRO+AB 42

0.1

1.0

10.0

% R

ead

Abu

ndan

ce

Colo

nD

ay 3

Day

28

Day

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Col

on

AB

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RO

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RO

+AB

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PR

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(A)

(B)

(C)

(D)

Figu

re 5


Ileum Mid colon

125

250

375

500

625

750

875

1000

3 28 42 3 28 42Age (days)

Obs

erve

d nu

mbe

r of O

TUs

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Obs

erve

d nu

mbe

r of O

TUs

Age (days)

Ileum Mid colon

125

250

375

500

625

750

875

1000

1125

1250

3 28 42 3 28 42Age (days)

Estim

ated

num

ber o

f OTU

s

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Est

imat

ed n

umbe

r of O

TUs

Age (days)

Ileum Mid colon

0.50

0.75

1.00

1.25

1.50

1.75

2.00

2.25

2.50

2.75

3.00

3.25

3.50

3.75

4.00

4.25

4.50

4.75

5.00

5.25

5.50

5.75

6.00

3 28 42 3 28 42Age (days)

Dive

rsity

inde

x

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Div

ersi

ty in

dex

Age (days)

(A)

(B)

(C)

Ileum Mid colon

125

250

375

500

625

750

875

1000

3 28 42 3 28 42Age (days)

Obs

erve

d nu

mbe

r of O

TUs

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colonO

bser

ved

num

ber o

f OTU

s

Age (days)

Ileum Mid colon

125

250

375

500

625

750

875

1000

1125

1250

3 28 42 3 28 42Age (days)

Estim

ated

num

ber o

f OTU

s

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Estim

ated

num

ber o

f OTU

s

Age (days)

Ileum Mid colon

0.50

0.75

1.00

1.25

1.50

1.75

2.00

2.25

2.50

2.75

3.00

3.25

3.50

3.75

4.00

4.25

4.50

4.75

5.00

5.25

5.50

5.75

6.00

3 28 42 3 28 42Age (days)

Dive

rsity

inde

x

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Div

ersi

ty in

dex

Age (days)

(A)

(B)

(C)

Ileum Mid colon

125

250

375

500

625

750

875

1000

3 28 42 3 28 42Age (days)

Obs

erve

d nu

mbe

r of O

TUs

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Obs

erve

d nu

mbe

r of O

TUs

Age (days)

Ileum Mid colon

125

250

375

500

625

750

875

1000

1125

1250

3 28 42 3 28 42Age (days)

Estim

ated

num

ber o

f OTU

s

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Estim

ated

num

ber o

f OTU

s

Age (days)

Ileum Mid colon

0.50

0.75

1.00

1.25

1.50

1.75

2.00

2.25

2.50

2.75

3.00

3.25

3.50

3.75

4.00

4.25

4.50

4.75

5.00

5.25

5.50

5.75

6.00

3 28 42 3 28 42Age (days)

Dive

rsity

inde

x

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Div

ersi

ty in

dex

Age (days)

(A)

(B)

(C)

Ileum Mid colon

125

250

375

500

625

750

875

1000

3 28 42 3 28 42Age (days)

Obs

erve

d nu

mbe

r of O

TUs

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Obs

erve

d nu

mbe

r of O

TUs

Age (days)

Ileum Mid colon

125

250

375

500

625

750

875

1000

1125

1250

3 28 42 3 28 42Age (days)

Estim

ated

num

ber o

f OTU

s

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Estim

ated

num

ber o

f OTU

s

Age (days)

Ileum Mid colon

0.50

0.75

1.00

1.25

1.50

1.75

2.00

2.25

2.50

2.75

3.00

3.25

3.50

3.75

4.00

4.25

4.50

4.75

5.00

5.25

5.50

5.75

6.00

3 28 42 3 28 42Age (days)

Dive

rsity

inde

x

Treatment

AB

CONTROL

PRO

PRO+AB

Ileum Mid colon

Div

ersi

ty in

dex

Age (days)

(A)

(B)

(C)

Figure 6


3 28 42

0

5

−4 −2 0 2 −4 −2 0 2 −4 −2 0 2PC1 [26.4%]

PC2

[8.4

%]

Treatment

AB

CONTROL

PRO

PRO+AB

Day 3 Day 28 Day 42

PC

2 - v

aria

tion

expl

aine

d 8.

4%


−10

−5

0

5

10

−8 −4 0 4RDA1 [24.1%]

RDA2

[8.6

%]

Treatment

AB

CONTROL

PRO

PRO+AB

RDA - colon day 28

RD

A2

- var

iatio

n ex

plai

ned

8.6%



(A)

(B)

Figure 7


2842

0.0

0.5

1.0

1.5

Cont

rol

LPS

Cont

rol

LPS

Tiss

ue

Fold change

Treatment

AB CONT

ROL

PRO

PRO

+AB

TNF−

alfa

TNF-⍺

Tiss

ue

Fold change*

****

Con

trol

LPS

-stim

ulat

edC

ontro

lLP

S-s

timul

ated

2842

0.0

0.5

1.0

Cont

rol

LPS

Cont

rol

LPS

Tiss

ue

Fold change

Treatment

AB CONT

ROL

PRO

PRO

+AB

IL10

IL-10

Fold change

Tiss

ueC

ontro

lLP

S-s

timul

ated

Con

trol

LPS

-stim

ulat

ed

2842

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

4.5

5.0

5.5

6.0

6.5

7.0

7.5

8.0

8.5

9.0

9.5

10.0

Cont

rol

LPS

Cont

rol

LPS

Tiss

ue

Fold change

Treatment

AB CONT

ROL

PRO

PRO

+AB

COX2

COX-2

Fold change

Tiss

ue

*

*

Con

trol

LPS

-stim

ulat

edC

ontro

lLP

S-s

timul

ated

2842

0.0

0.5

1.0

1.5

Cont

rol

LPS

Cont

rol

LPS

Tiss

ue

Fold change

Treatment

AB CONT

ROL

PRO

PRO

+AB

ZO1

**

Fold change

Tiss

ue

ZO-1

Con

trol

LPS

-stim

ulat

edC

ontro

lLP

S-s

timul

ated

(A)

(B)

(D)

(C) Fi

gure

8 (1/2)


2842

0.0

0.5

1.0

1.5

Cont

rol

LPS

Cont

rol

LPS

Tiss

ue

Fold change

Treatment

AB CONT

ROL

PRO

PRO

+AB

OCL

NOCLN

Fold change

Tiss

ueC

ontro

lLP

S-s

timul

ated

Con

trol

LPS

-stim

ulat

ed

2842

0.0

0.5

1.0

1.5

2.0

Cont

rol

LPS

Cont

rol

LPS

Tiss

ue

Fold change

Treatment

AB CONT

ROL

PRO

PRO

+AB

CLDN

4

Fold change

Tiss

ue

CLDN-4

Con

trol

LPS

-stim

ulat

edC

ontro

lLP

S-s

timul

ated

2842

0.0

0.5

1.0

Cont

rol

LPS

Cont

rol

LPS

Tiss

ue

Fold change

Treatment

AB CONT

ROL

PRO

PRO

+AB

CLDN

2CLDN-2

Fold change

Tiss

ueC

ontro

lLP

S-s

timul

ated

Con

trol

LPS

-stim

ulat

ed

(E)

(F)

(G) Fi

gure

8 (2/2)


−4

−2

0

2

4

−5.0 −2.5 0.0 2.5PC1 [19.6%]

PC2

[15.

8%]

10

20

30

40Age

Sample_Type

Ileum

Mid colon

PC

2 - v

aria

tion

expl

aine

d 15

.8%


−4

−2

0

2

−2 0 2 4PC1 [21.6%]

PC2

[9.6

%]

10

20

30

40

Age

PC

2 - v

aria

tion

expl

aine

d 9.

6%

PC1 - variation explained 21.6%Figure S1

Figure S2


Chapter 7. General discussion 153

Chapter 7

General discussion

Intensive pig production systems, optimised for taking advantage of the pigs fullreproductive potential, demand adult pigs and piglets to cope with unnatural con-ditions as extremely large litter sizes and early weaning. Piglets experience a majorhurdle after weaning, where intestinal disorders and diarrhoea are highly prevalent,leading to decreased animal welfare and frequent use of antibiotics (DANMAP,2014). The gut microbiota, and the process of its development, is appreciated asan essential part of the mammalian gut, serving numerous functions in relation toboth animal and human health. The dynamic nature of the gut microbial com-munity allows it to be manipulated by a range of external and internal factors,many of which can potentially be taken advantage of in the search for measurescapable of improving gut health. Hence, a deeper understanding of how, when,and to what extent the piglet gut microbiota can be manipulated is pivotal, if suchmanipulating actions are to be further developed, and, with time, find general ap-plications in enhancing gut robustness. Thus, the overall aim of the present thesiswas to investigate the influencing effects of external and host-related factors onthe development of the piglet gut microbiota.

The following discussion highlights the major findings from the animal experi-ments presented in chapter 4, 5 and 6, and discusses these in relation to the currentscientific knowledge.

7.1 Manipulating the gut microbiota

The major objective of the current thesis was to characterise if, and how, BC,FUT1 genotype, gentamicin, and Bacillus spp. spores influenced the microbialcolonisation of the piglet gut. This point is discussed in the following sections,

154 Chapter 7. General discussion

including relevant experimental issues.

7.1.1 Milk-based dietary intervention

The aim of manuscript 1 (chapter 4) was to investigate the isolated effects of eachof the included milk-based diets, and required piglets from the same litter to beassigned to three different dietary groups. Piglets were fed no other feed thanmilk, and one week was therefore considered to be enough time for the milk-dietto exert its potential effects on the gut microbiota (e.g. Heinritz et al., 2016). Toallow sufficient milk intake, sow milk-fed piglets were housed together with the sowduring the experimental period. Thus, continuous contact between the sow andsow milk-fed piglets, different housing environments and the stress of weaning wereexpected to cause variation between piglets suckling the sow and piglets in the twoexperimental dietary groups. The sow milk-fed piglets were therefore included asa reference group.

In accordance with previous dietary studies performed on suckling piglets (Caoet al., 2016; Frese et al., 2015; Yeruva et al., 2016), manuscript 1 illustrates thatdifferent diets, in this case different milk types/sources, clearly change the compo-sition of the gut microbiota. This observation is in agreement with human studies,where breast-fed and formula-fed children have been found to have different gutmicrobiotas (e.g. Bokulich et al., 2016). As mother’s own milk is preferred, butnot always available, milk replacers are indispensable feeding supplements, andformula manufacturers are constantly trying to optimise their products (Martinet al., 2016).

The rise of interest for BC stems in particular from its high content of bioactivecomponents as growth hormones and immunomodulators (Pandey et al., 2011). Inour study, the finding of a change in genera belonging to lactic acid bacteria inBC-fed piglets, and a rise in the number of potentially pathogenic Enterobacteri-aceae in MR-fed piglets confirms, that BC provides components to the gut thatare in fact able to promote a gut microbiota harbouring host-beneficial bacteria.This observation agrees with previous studies in piglets, where MR-feeding is ac-companied by a rise in Enterobacteria (Yeruva et al., 2016). The lower incidence ofdiarrhoea in BC-fed piglets, compared to those fed MR, suggests that some of thebioactive components in BC are indeed able to provide protection against intesti-nal pathogens. In accordance with the general perception of mother’s own milkbeing the preferred nutrition for suckling piglets, it was obvious that sow milk wassuperior to the investigated experimental diets. However, when mother’s milk is


not available, the loss of important compounds may be counteracted by the rangeof bioactive components found in BC. Such a potential can be taken advantageof when piglets are weaned. Weaning is commonly associated with decreased feedintake, weight loss or growth check, and diarrhoea caused by ETEC (Fairbrotheret al., 2005; Lalles et al., 2007). Previous studies on BC and its use in piglets post-weaning have shown, that BC has a beneficial effect on both weight gain and feedintake (Boudry et al., 2008; Huguet et al., 2012). Due to the experimental set-upof our study, we are not able to make any conclusions on the effect of BC on pigletbody weight under normal weaning conditions. As enteric colibacillose is highlyprevalent after weaning, the apparent ability of BC to reduce the number of Enter-obacteriaceae in digesta suggests that BC has the potential to reduce colonisationof ETEC, and hence reduce the number of piglets with diarrhoea and in need ofantibiotics. In support of this, results published by Sugiharto et al. (2015), thatoriginate from the same animal experiment as the one presented in manuscript1, furthermore confirm that the number of mucosa-associated Enterobacteriaceaeand haemolytic bacteria indeed was reduced in the jejunum and ileum of BC-fedpiglets (Sugiharto et al., 2015). Hence, provision of BC seems to be a promisingstrategy to enhance gut health of (weak) piglets.

7.1.2 ETEC F18 susceptibility

Earlier studies have reported pig breed, and hence genotype, to influence the gutmicrobial composition (Bian et al., 2016; Pajarillo et al., 2015). The rationalebehind manuscript 2 varied from those of manuscript 1 and 3, as the objective wasto investigate whether a genetic mutation causing resistance to ETEC F18 alsocaused an unintentional change in the microbial colonisation of the gut. FUT1expression is age-dependent (Bao et al., 2012), and piglets were studied from birthuntil 34 days of age. This period included weaning at day 28, an event wheresusceptible piglets were highly likely to become infected by ETEC F18.

As the FUT1 enzyme is expressed in the gut (Bao et al., 2012) and acts byfucosylating glycoproteins and -lipids (Meijerink et al., 2000), lack of a functionalenzyme may impose changes to other glycoproteins found in close relation to theintestinal mucosa. From the results of manuscript 2, we were not able to showthat piglets with different FUT1 genotypes (307A/G and 307G/G) also experiencedsignificant differences in their early gut colonisation (i.e. during suckling and thefirst week after weaning). Piglets were partly confirmed to have different suscepti-bilities to ETEC F18, as the number of luminal intestinal haemolytic bacteria was


significantly different between piglets with different genotypes. As the haemolyticcolonies unfortunately were not serotyped, we do not know with certainty whetherthe colonies indeed did represent ETEC F18.

The study only included 17 piglets originating from two different sows, andwas an obvious limitation of the study. Bian et al. (2016) show that genotypeis a weaker influencing factor than weaning and age, suggesting that potentialeffects of genotype are harder to detect, which, combined with the low number ofanimals, potentially can be the reason for the observed results. From the currentexperiment, we could not provide evidence that ETEC F18 resistant piglets werein risk of developing a gut microbiota that could potentially compromise animalhealth later in life.

7.1.3 Bacillus spores and gentamicin

Antibiotics are generally believed to have a big influence on the composition of thegut microbiota. As the colonisation process in young animals is pivotal to properintestinal immune education (Sommer and Baeckhed, 2013), perturbations in earlylife may have detrimental long-term effects as suggested by e.g. the associationbetween early antibiotic administration and allergic conditions in children (Almet al., 2008; Goksor et al., 2013). Gentamicin is a commonly used drug for treatingsystemic infections in newborn infants (Chirico et al., 2009) and neonatal diarrhoeain piglets. Hence, gentamicin was believed to be the most relevant antibiotic toinvestigate in newborn piglets. In agreement with previous studies investigatingthe influential effects of antibiotics on the gut microbial community in pigs (Looftet al., 2014b; Schokker et al., 2014), we found gentamicin to affect the colonic andfaecal microbial communities. Surprisingly, this effect was most evident severalweeks after the piglets had been treated with the antibiotic. The effect of gen-tamicin was lower than expected, and gentamicin was not associated with signs ofdysbiosis (e.g. diarrhoea). Actually, our results showed that gentamicin increasedthe microbial diversity (Shannon index), which was in accordance with Schokkeret al. (2014). When compared to gentamicin, administering Bacillus spores hadthe opposite effect on the microbial richness and diversity. Such a counteract-ing effect was previously reported in humans administered Lactobacillus casei forthree days after an antibiotic treatment had been completed (Pirker et al., 2013).Whereas the combined effect of an antibiotic and a probiotic reduced the num-ber of people developing antibiotic-associated diarrhoea in the human study, ourstudy did not provide evidence that the combined effect of gentamicin and Bacil-


lus spores resulted in clinically healthier piglets. On the contrary, litters receivingBacillus spores had a higher frequency of diarrhoea than those only administeredgentamicin, especially during suckling. Li et al. (2012) reported an increased diar-rhoea incidence when the probiotic dose of Lactobacillus rhamnosus was increasedfrom 1x1010 to 1x1012 cfu, though both doses were able to reduce diarrhoea inci-dence after ETEC F4 challenge. Hence, the amount of Bacillus spores provided inthe current study may have been too high, but may have shown protective effectsduring an infection trial. The lack of uniformity between probiotic studies in pigs,as mentioned in the introduction, leaves us without solid theoretical knowledgeupon which we can base our choice of dose on. Thus, future studies on the use ofprobiotics for improving piglet gut health are needed.

7.2 Age-related succession of the gut microbiota

Manuscript 2 and 3 (presented in chapter 5 and 6) include longitudinal studies ofthe gut microbiota spanning a minimum of four weeks. The two studies clearlydemonstrate the importance of age in shaping the gut microbial communities ofpiglets. In both studies, a gradual change in the overall microbial community wasobserved during the suckling period. Additionally, there was a clear differencebetween the microbial communities on the day of weaning and 1-2 weeks post-weaning. Besides the impact of age, changes in microbial community profiles afterweaning are also caused by the dietary shift (from sow milk to solid feed) and themere impact of being weaned (stress and change of environment).

The earliest samples were collected three days after birth (manuscript 3), wherea colonisation process in obvious progress was evident. This is in agreement witholder (Jensen, 1998; Swords et al., 1993) and newer literature (Frese et al., 2015;Petri and Hill, 2010) stating that a gut microbiota is established within the first24 hours after birth. From birth until weaning, the microbiota became graduallyricher in species, and species diversity increased, peaking around 21 days of agein manuscript 2. In manuscript 3, the piglets were sampled until 42 days of age,and in this case, species richness and diversity increased from 28 to 42 days of age(however not seen when administered antibiotics). In agreement with our results,Frese et al. (2015) reported an increase in faecal diversity until two weeks afterweaning, and Slifierz et al. (2015) an increase in faecal species richness until oneweek after weaning. Differences between results obtained in manuscript 2 and 3might be caused by the fact that different procedures for 16S rRNA gene analysis


were used for sample and bioinformatic processing.On phylum level, the faecal community was consistently dominated by Firmi-

cutes and Bacteroidetes irrespective of age and experimental intervention. Further-more, in accordance with Frese et al. (2015) and Slifierz et al. (2015), Firmicutessteadily increased its dominance as the piglets aged. On genus level the picturewas slightly more inconsistent, and the dominating genera- and age-related differ-ences differed between experiments. These observations highlight the variabilitybetween studies and the difficulty of comparing experiments.

7.3 Sampling the gut microbial communities

The gut microbiota is a composed community consisting of several microhabitats.It is widely known that different compartments along the length and width of thegut vary considerably between each other (Looft et al., 2014a). The results frommanuscript 1, 2, and 3 expectedly agreed with these previous reports. It was alsoevident from these data that in the search of the effects of an influencing factor, thesite investigated may be of major importance. In manuscript 1, the effect of dietwas observed in digesta, whereas manuscript 3 revealed an effect of the antibioticand probiotic in the mid colon and faeces. The mucosa-associated community wasnot investigated in any of the animal studies.

In retrospect, to pursue the objective of manuscript 2, subjecting intestinal di-gesta or, perhaps even better, mucosa from the small intestine to high-throughput16S rRNA gene sequencing would have been useful. As the potentially changedglycoproteins are expected to be found in the mucus layer or on cell surfaces (Hes-selager et al., 2016), it would have been highly relevant to include this niche aswell. Also, sampling of intestinal digesta at an early age shortly after antibiotictherapy would have added more information to the data presented in manuscript3. Nonetheless, sampling of luminal contents from the length of the gut requiresanimals to be sacrificed. Sacrificing too many animals early in the experiment maybe a problem if the number of animals beforehand is low, which was the case inmanuscript 2.

Faecal samples were consistently included to be able to evaluate the effect oftime. This approach has been applied by several papers (e.g. Frese et al., 2015;Slifierz et al., 2015) and obviously allows the same animal to be sampled multipletimes. However, when faecal samples are deemed insufficient, intestinal digestafrom different piglets belonging to the same litter has to be sampled multiple


times. Such an approach inevitably introduces variation due to natural variabilitybetween individual piglets.

7.4 High-throughput sequencing versus microbial

culture

Applying high-throughput sequencing of the 16S rRNA gene, as done in all ex-periments of the present thesis, provides an in-depth insight into the diversity ofthe gut microbiota, joined by a thorough knowledge on the taxonomy of presentbacteria. This method has been increasingly applied in the study of complex mi-crobial environments during the last decade. This molecular method, however,potentially introduces biases during sample preparation and sequencing, and doesnot allow absolute quantification. In an effort to circumvent this problem, we in-cluded microbial plate culture as well. From the very beginning we were awarethat traditional culture would be insufficient in regards to the detailed characteri-sation of the complex microbial communities in the gut, and the presented studiesalso confirm these limitations. Manuscript 1 and 2, however, underline the impor-tance of including culture of potentially pathogenic bacteria, here the family ofEnterobacteriaceae and haemolytic bacteria. Furthermore, some pathogens maybe found in numbers too low to be detected by current sequencing platforms, andhence require use of other methods for their detection (Lagier et al., 2015).

The application of two different platforms turned out to complicate the overallinterpretation of the obtained results. Due to the need of additional extensivequality filtering and trimming, the sequences from the Ion Torrent platform wererelatively short (166 base-pairs). The faecal samples appeared abnormally rich,which could be assigned to the short read lengths (Engelbrektson et al., 2010).Also, only ten of the 25 most abundant species were assigned to genus level, andnumerous OTUs were not taxonomically assigned or assigned to order level only.These results demonstrate the importance of platform specifications as e.g. readlength, when assigning taxonomy to 16S rRNA gene fragments.


7.5 Linking gut microbial community composition

to gut health and animal robustness

Diarrhoea is frequently used as a clinical measure of gut health and also broaderas an indicator of lack of animal health. In manuscript 1 (chapter 4) the increaseddiarrhoea incidence was associated with an increased detection of Enterobacteri-aceae and haemolytic bacteria, whereas a lower diarrhoea incidence was associatedwith an increased detection of Lactobacillus. This is in accordance with the Enter-obacteriaceae to Lactobacillus ratio often used as a measure of gut health (Castilloet al., 2007). However, whether this measure is consistently associated with anincreased animal health is unknown. Though the number of studied animals werelow, occurrence of diarrhoea was included as a clinical observation of piglet health.In manuscript 3, the included measures of gut health (Enterobacteriaceae, Lacto-bacillus, inflammation and intestinal tight-junction markers) were not significantlydifferent between the experimental groups, and not in accordance with the observeddiarrhoea frequencies. Species richness and diversity, on the other hand, did differbetween treatments. Petersen and Round (2014) suggested that a gut microbiotafree from dysbiosis was characterised by a highly diverse community. However, thegeneral perception of a healthy gut being associated with a microbial communityof high diversity and richness is currently challenged. In humans, a high microbialrichness has been associated with long colonic transit time and increased proteinfermentation (Roager et al., 2016), which generally is not considered beneficial tothe host. Also, as seen in manuscript 1, the piglets with the highest diarrhoea in-cidence were also the ones with the highest microbial richness and diversity. Theseresults stress the complexity of the gut microbiota, and that the definition of ahealthy gut may change according to the investigated circumstances.

Chapter 8. Conclusion 161

Chapter 8

Conclusion

The results of the presented Ph.D.-study contribute to the existing knowledge onhow different factors influence the colonisation and succession of the gut microbiotain piglets. The major findings and conclusions from the three animal experimentsare presented in the following.

• Diet had a major impact on the intestinal luminal microbiota.

• When mother’s own milk was not available, BC was a more suitable substi-tute, compared to a regular milk replacer, in maintaining a health-promotinggut microbiota.

• Especially the ileal community of BC fed piglets had a closer resemblance tothat of sow milk-fed piglets with regard to the abundance of potential entericpathogens.

• Genetic manipulation of the FUT1 gene did not notably affect colonisationof the microbiota, investigated by analysing faecal samples.

• Gentamicin and Bacillus spores changed the colonic and faecal microbiota,which was especially evident just before weaning at 28 days of age.

• Gentamicin increased community species richness and diversity, while Bacil-lus spore administration had the opposite effect.

• No negative effect on the animals general health was observed after gentam-icin administration. On the contrary, piglets administered Bacillus sporestended to have a higher diarrhoea frequency.

162 Chapter 8. Conclusion

• Age had a major effect on the gut microbiota, causing a gradual change ofgut communities as the piglets aged.

In conclusion, the investigated external factors, and especially diet, had themost significant influence on the colonisation of the gut microbiota in piglets. Noeffect was seen when FUT1 genotype was manipulated. BC showed the mostpromising results in shaping the gut microbiota in a direction favouring healthierpiglets.

Chapter 9. Perspectives and future work 163

Chapter 9

Perspectives and future work

The close relationship between the gut microbiota and host health unfolds an arrayof opportunities for potentially influencing piglet health by delicately orchestratingthe complex gut microbiota. The presented results confirm that the gut microbiotaof piglets is manipulative by various factors. Especially the dietary interventionwith BC showed promising results in relation to improving piglet gut health. Thereported results indicate that BC is able to favour a beneficial gut microbiota,which could be useful both during suckling and in relation to weaning. Due to theresemblance in gut structure and the omnivorous dietary habit between humansand pigs, such perspectives might also be of relevance in infants.

Increased litter sizes in the modern pig industry have required many pig farmersto implement the use of milk replacers (De Vos et al., 2014). Substituting regularmilk replacers with BC could potentially enhance the performance of piglets inthe need of sow milk supplementation, increasing survival chances and robustnesslater in life, e.g. after weaning. Also, in relation to weaning, including BC maycontribute to an increased gut health, hence increase animal welfare and reduce theneed for antibiotics. Before such measures can be implemented, additional studiesare needed. Focus should be on finding the proportion of BC needed to obtain thebeneficial effects, how it is best administered, and the long-term effects. The highprotein content might pose problems due to protein fermentation and potentialproduction of harmful by-products. An important aspect is also whether its useis rentable for the pig farmer. If the price is too high, the odds of a successfulimplementation are bad. However, to be able to implement its use on a wide scale,the production has to be able to keep up with the demand. As the lactating cowonly produces colostrum for a short period of time, and the calf needs to be fed it aswell, BC might not be produced in large enough amounts, making it unobtainable

164 Chapter 9. Perspectives and future work

or too expensive.Though we did not find supplementation with Bacillus spores to beneficially

influence gut health, its potential use should not be disregarded. The current ev-idence on probiotics is in some regards indeed contradicting. However, assigningsuch observations to the functionality of the probiotics alone should be done withcare. The variability between studies is substantial and underlines the need foruniformity. Interestingly, the investigated Bacillus spores were shown to counter-act the effect of gentamicin, and hence the concomitant use of these should befurther investigated. Further studies should emphasise investigating the long-termeffects of Bacillus spp. spores and how they influence performance of piglets dur-ing infection challenges. Use of probiotics could find application in the pre- andpost-weaning period. Administrating probiotics to sows might be a way of verti-cally transferring probiotic species to the offspring, and may be a way of increasinghealth of the mother sow.

As genetic manipulation of specific intestinal pathogen receptors is an appliedapproach for coping with ETEC infections after weaning, the lack of evidence ofa changed colonisation of the indigenous gut microbiota favours the continued useof such a procedure. However, additional work is needed to support the presentedfindings. Future studies should include a larger number of animals and analysesof the mucosa-associated bacterial community.

In general, future studies on the gut microbial community should be supportedby extensive functional analyses, and measures of both the piglets clinical and sub-clinical condition. This could provide a deeper insight into the actual significanceof microbial community changes in the gut.

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Appendix A. 181

Appendix A

The table below provides information on the microbial composition in the stom-ach, small intestine and large intestine of piglets immediately after birth, duringsuckling, and during the first two weeks after weaning.

• Frese et al. (2015) used V4 16S rRNA gene sequencing and Illumina MiSeq,and piglets were weaned at 21 days of age.

• Gancarcikova et al. (2008) used culture, and piglets were weaned at 28 daysof age.

• Melin et al. (1997) used culture, and piglets were weaned at 35 days of age.

• Petri and Hill (2010) used sequencing and used an unknown technique; pigletswere not weaned.

• Slifierz et al. (2015) used V4 16S rRNA gene sequencing and Ilumina Miseq,and piglets were weaned at 21 days of age.

• Swords et al. (1993) used culture, and piglets were weaned at 21 days of age.

182 Appendix A.

Age Taxa Sample type Reference

Sequencing (taxa in %)

0.25 d Clostridiaceae (33.8)Enterobacteriaceae (25)Moraxellaceae (8.8)Lactobacillaceae (1.5)Streptococcaceae (1.5)

Peptostreptococcaceae (1.5)

Pooled digesta Petri and Hill (2010)

0.5 d Clostridiaceae (50)Streptococcaceae (19.7)Enterobacteriaceae (13.2)Pasteurellaceae (5.3)


1 d Streptococcaceae (37.7)Clostridiaceae (27.3)Lactobacillaceae (6.5)Enterobacteriaceae (3.9)Pasteurellaceae (3.9)Peptostreptococcaceae (1.3)Lachnospiraceae (1.3)Micrococcaceae (1.3)


2 d Streptococcaceae (29.9)Lactobacillaceae (16.9)Clostridiaceae (7.8)Bacteroidaceae (5.2)Veillonellaceae (5.2)Moraxellaceae (3.9)Peptostreptococcaceae (1.3)Lachnospiraceae (1.3)Pasteurellaceae (1.3)Micrococcaceae (1.3)


3 d Streptococcaceae (22.8)Lactobacillaceae (19.0)Clostridiaceae (12.7)Moraxellaceae (10.1)Peptostreptococcaceae (7.6)Pasteurellaceae (5.1)Veillonellaceae (3.8)Enterobacteriaceae (1.3)Ruminococcaceae (1.3)Lachnospiraceae (1.3)


Continues on the next page

Appendix A. 183


5 d Lactobacillaceae (50.0)Moraxellaceae (6.4)Lachnospiraceae (6.4)Streptococcaceae (3.8)Pasteurellaceae (2.6)Veillonellaceae (2.6)Clostridiaceae (1.3)Enterobacteriaceae (1.3)Peptostreptococcaceae (1.3)Micrococcaceae (1.3)


10 d Lactobacillaceae (46.7)Ruminococcaceae (10.7)Enterobacteriaceae (5.3)Veillonellaceae (5.3)Clostridiaceae (4.0)Moraxellaceae (4.0)Lachnospiraceae (2.7)Pastrurellaceae (1.3)Micrococcaceae (1.3)


20 d Lactobacillaceae (44.6)Ruminococcaceae (9.5)Lachnospiraceae (9.5)Streptococcaceae (5.4)Moraxellaceae (2.7)Clostridiaceae (1.4)Pasteurellaceae (1.4)Micrococcaceae (1.4)


1 d Bacteroidaceae (21.5)Enterobacteriaceae (21.0)Clostridiaceae (15.0)Lachnospiraceae (12.2)Lactobacillaceae (5.0)Streptococcaceae (4.0)Enterococcaceae (2.5)Ruminococcaceae (2.0)Erysipelotrichaceae (1.0)Prevotellaceae (<1.0)

Faeces Frese et al. (2015)

3 d Enterobacteriaceae (27.5)Bacteroidaceae (19.0)Lachnospiraceae (13.5)Lactobacillaceae (8.0)Clostridiaceae (6.0)Ruminococcaceae (4.5)Erysipelotrichaceae (3.0)Enterococcaceae (2.5)Streptococcaceae (2.0)Prevotellaceae (<1.0)



184 Appendix A.


5 d Enterobacteriaceae (19.0)Bacteroidaceae (13.5)Lachnospiraceae (13.0)Lactobacillaceae (10.5)Clostridiaceae (6.5)Ruminococcaceae (4.5)Enterococcaceae (4.0)Erysipelotrichaceae (3.5)Streptococcaceae (3.0)Prevotellaceae (<1.0)


7 d Enterobacteriaceae (17.5)Lachnospiraceae (14.5)Bacteroidaceae (13.5)Lactobacillaceae (10.5)Erysipelotrichaceae (5.0)Ruminococcaceae (5.0)Enterococcaceae (4.0)Clostridiaceae (3.5)Streptococcaceae (2.0)Prevotellaceae (<1.0)


14 d Enterobacteriaceae (25.0)Lachnospiraceae (14.0)Erysipelotrichaceae (7.5)Enterococcaceae (7.5)Lactobacillaceae (7.0)Bacteroidaceae (6.0)Ruminococcaceae (4.0)Clostridiaceae (2.0)Streptococcaceae (1.0)Prevotellaceae (<1.0)


21 d Bacteroidaceae (16.5)Lachnospiraceae (16.0)Enterobacteriaceae (12.0)Erysipelotrichaceae (10.0)Ruminococcaceae (6.0)Lactobacillaceae (5.0)Enterococcaceae (3.0)Clostridiaceae (2.0)Streptococcaceae (<1.0)Prevotellaceae (<1.0)



Appendix A. 185


28 d Ruminococcaceae (16.0)Lachnospiraceae (13.5)Prevotellaceae (12.0)Erysipelotrichaceae (11.0)Lactobacillaceae (10.0)Bacteroidaceae (4.0)Streptococcaceae (3.0)Enterobacteriaceae (2.5)Clostridiaceae (1.5)Enterococcaceae (1.0)


35 d Lactobacillaceae (19.0)Prevotellaceae (17.0)Ruminococcaceae (16.0)Lachnospiraceae (6.0)Erysipelotrichaceae (3.0)Streptococcaceae (1.5)Clostridiaceae (1.0)Enterobacteriaceae (<0.1)Bacteroidaceae (<1.0)Enterococcaceae (<1.0)


42 d Lactobacillaceae (20.0)Prevotellaceae (17.5)Lachnospiraceae (12.0)Ruminococcaceae (10.0)Streptococcaceae (8.5)Erysipelotrichaceae (4.0)Clostridiaceae (1.0)Enterobacteriaceae (<0.1)Bacteroidaceae (<1.0)Enterococcaceae (<1.0)


1-3 d Clostridium (17.9)Escherichia (15.0)Fusobacterium (10.5)Clostridium XIVa (4.3)Lactobacillus (4.2)

Faeces Slifierz et al. (2015)

7-21 d Clostridium (8.8)Escherichia (8.6)Lactobacillus (8.2)Clostridium XIVa (7.4)unclassified Firmicutes (5.2)


28-35 d Megasphaera (14.0)Lactobacillus (12.3)Clostridium (4.6)Succinivibrio (4.1)unclassified Firmicutes (2.2)



186 Appendix A.


42-49 d Megasphaera (21.2)Lactobacillus (12.3)Roseburia (4.2)Unclassified Firmicutes (3.9)Erysipelotrichaceae (3.9)


Culture (taxa in log-transformed cfu numbers)

3 h Total aerobes (3.9)Total anaerobes (2.7)Coliforms (2.6)E. coli (0)Bacteroides spp. (0)Clostridium spp. (0)Lactobacillus spp. (0)

Distal colonic digesta Swords et al. (1993)

6 h Total anaerobes (6.9)Total aerobes (6.7)Clostridium spp. (6.7)Coliforms (6.2)E. coli (5.6)Bacteroides spp. (0)Lactobacillus spp. (0)


9 h Total anaerobes (8.4)Coliforms (8.4)Clostridium spp. (8.2)Total aerobes (8.1)E. coli (7.8)Bacteroides spp. (0)Lactobacillus spp. (0)


12 h Total aerobes (8.8)Clostridium spp. (8.5)Coliforms (8.4)Total anaerobes (7.8)E. coli (7.3)Bacteroides spp. (0)Lactobacillus spp. (0)


1 d Total anaerobes (9.6)Total aerobes (8.3)Clostridium spp. (8.2)Coliforms (7.9)E. coli (6.9)Lactobacillus spp. (5.9)Bacteroides spp. (0)



Appendix A. 187


2 d Total anaerobes (8.1)Coliforms (7.7)Total aerobes (7.4)E. coli (6.6)Clostridium spp. (6.5)Lactobacillus spp. (4.6)Bacteroides spp. (0)


3 d Total anaerobes (9.7)Clostridium spp. (9.1)Total aerobes (9.0)Coliforms (8.7)E. coli (8.7)Lactobacillus spp. (5.1)Bacteroides spp. (0)


4 d Total anaerobes (9.9)Clostridium spp. (9.6)Total aerobes (9.0)Lactobacillus spp. (7.8)Coliforms (7.5)E. coli (6.8)Bacteroides spp. (6.3)


5 d Total anaerobes (9.9)Clostridium spp. (9.8)Total aerobes (8.8)Lactobacillus spp. (7.5)Coliforms (7.3)E. coli (7.0)Bacteroides spp. (<5.0)


6 d Total anaerobes (9.5)Total aerobes (8.2)Bacteroides spp. (7.7)Clostridium spp. (7.6)Coliforms (7.3)E. coli (7.2)Lactobacillus spp. (6.9)


7 d Total anaerobes (9.8)Clostridium spp. (9.7)Total aerobes (8.9)Coliforms (6.9)Bacteroides spp. (6.6)Lactobacillus spp. (6.0)E. coli (5.8)



188 Appendix A.


8 d Total anaerobes (8.4)Clostridium spp. (8.0)Bacteroides spp. (7.4)Lactobacillus spp. (6.7)Total aerobes (6.6)Coliforms (6.6)E. coli (4.2)


9 d Bacteroides spp. (8.9)Clostridium spp. (8.9)Total anaerobes (8.7)Coliforms (7.6)Total aerobes (6.9)Lactobacillus spp. (6.7)E. coli (5.9)


10 d Total anaerobes (9.0)Clostridium spp. (7.6)Coliforms (7.1)Bacteroides spp. (<7.0)Total aerobes (6.0)E. coli (6.0)Lactobacillus spp. (5.9)


15 d Clostridium spp. (9.3)Total anaerobes (8.8)Bacteroides spp. (8.8)Coliforms (7.8)Lactobacillus spp. (7.6)Total aerobes (6.5)E. coli (4.3)


20 d Total anaerobes (9.1)Clostridium spp. (9.1)Bacteroides spp. (8.6)Lactobacillus spp. (7.6)Coliforms (7.0)E. coli (6.5)Total aerobes (6.4)


25 d Total anaerobes (10.6)Clostridium spp. (9.5)Bacteroides spp. (8.4)Lactobacillus spp. (7.7)Coliforms (7.6)E. coli (6.9)Total aerobes (5.9)



Appendix A. 189


30 d Clostridium spp. (10.4)Total anaerobes (8.9)Bacteroides spp. (8.7)Lactobacillus spp. (8.1)Coliforms (7.9)Total aerobes (7.5)E. coli (5.1)


2 d Total aerobes (7.4)Lactobacilli (6.4)Enterococci (6.4)E. coli (4.6)Enterobacteriaceae (4.0)

Jejunal digesta Gancarcikova et al. (2008)



14 d Total aerobes (7.5)Lactobacilli (7.0)Enterococci (5.0)Enterobacteriaceae (4.0)E. coli (3.8)


21 d Total aerobes (7.0)Lactobacilli (6.0)E. coli (5.4)Enterococci (5.0)Enterobacteriaceae (4.8)




35 d Total aerobes (6.4)Lactobacilli (6.0)Enterococci (4.0)E. coli (3.8)Enterobacteriaceae

(not detected)





190 Appendix A.


2 d Total aerobes (9.0)Lactobacilli (7.4)Enterobacteriaceae (6.9)E. coli (6.6)Enterococci (6.0)

Ileal digesta Gancarcikova et al. (2008)

7 d Total aerobes (7.8)Enterobacteriaceae (7.2)E. coli (7.1)Lactobacilli (7.0)Enterococci (4.0)


14 d Lactobacilli (9.2)E. coli (7.7)Total aerobes (7.5)Enterobacteriaceae (7.1)Enterococci (5.0)


21 d Total aerobes (8.8)Lactobacilli (8.0)E. coli (6.9)Enterobacteriaceae (6.6)Enterococci (6.0)




35 d Total aerobes (9.2)Lactobacilli (8.0)Enterobacteriaceae (5.8)E. coli (5.2)Enterococci (5.0)


42 d Lactobacilli (8.6)Enterococci (8.0)Total aerobes (7.0)E. coli (6.2)Enterobacteriaceae (5.2)


2 d Total aerobes (9.4)E. coli (9.0)Enterobacteriaceae (9.0)Lactobacilli (8.9)Enterococci (8.0)

Caecal digesta Gancarcikova et al. (2008)




Appendix A. 191








35 d Lactobacilli (9.8)Total aerobes (9.4)Enterococci (7.2)E. coli (6.8)Enterobacteriaceae (6.0)


42 d Lactobacilli (9.8)Total aerobes (9.1)Enterococci (8.9)E. coli (5.9)Enterobacteriaceae (5.4)


7 d E. coli (8.8)Enterococci (7.7)C. perfringens (4.6)

Faeces Melin et al. (1997)

14 d Enterococci (8.3)E. coli (8.2)C. perfringens (4.0)


21 d E. coli (8.4)Enterococci (7.4)C. perfringens (0)







192 Appendix A.


38 d Enterococci (6.5)E. coli (5.6)C. perfringens (0)




colonisation and succession of the gut microbiota in suckling- and newly weaned piglets · 3....

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