Analysis of potential host-colonization factors in bifidum S17

Dissertation

zur Erlangung des Doktorgrades Dr. rer. nat.

der Fakultät für Naturwissenschaften

der Universität Ulm

vorgelegt von

Christina Westermann

aus Ulm

2015

Amtierender Dekan: Prof. Dr. Joachim Ankerhold

Erstgutachter: PD Dr. Christian Riedel

Zweitgutachter: Prof. Dr. Bernhard Eikmanns

Tag der mündlichen Prüfung: 15.10.2015

Die vorliegende Arbeit wurde im Institut für Mikrobiologie und Biotechnologie an der Universität Ulm angefertigt.

Die Arbeit wurde durch ein Stipendium nach dem Landesgraduiertenförderungsgesetz Baden-Württemberg finanziert.

TABLE OF CONTENTS

1. Introduction ...... 8

1.1. General features of the genus Bifidobacterium ...... 8

1.2. Bifidobacteria as a part of the human microbiota ...... 9

1.3. Bifidobacteria as ...... 10

1.3.1. Definition of probiotics ...... 10

1.3.2. bifidobacteria and their health-promoting effect on human host ...... 11

1.4. Potential adhesive structures of the human gut...... 12

1.4.1. Mucus ...... 12

1.4.2. Intestinal epithelial cells and the glycocalyx ...... 13

1.4.3. Extracellular matrix proteins ...... 14

1.5. Bacterial cell surface architecture – potential adhesion mediating structures ...... 15

1.5.1. Exopolysaccharides ...... 17

1.5.2. Cell envelope proteases ...... 18

1.5.3. Pili ...... 19

1.5.4. Identified single adhesins ...... 20

1.5.4.1. Translocation and cell wall anchoring of cell surface proteins ...... 21

1.6. The aim of this study ...... 23

2. Materials and Methods ...... 24

2.1. Bacterial strains ...... 24

2.1.1. Bifidobacteria ...... 24

2.1.2. Escherichia coli ...... 25

2.2. Plasmids ...... 26

2.3. Eukaryotic Caco-2 cell line ...... 27

2.3.1. Subcultivation of Caco-2 cells...... 27

2.3.2. Cryoconservation of Caco-2 cells ...... 28

2.3.3. Thawing of cryo-stocks ...... 28

2.3.4. Determination of the cell number ...... 28

2.4. Adhesion assays ...... 29

2.4.1. Adhesion assay with Caco-2 cells ...... 29

2.4.2. Competitive adhesion assay ...... 29

2.4.3. Adhesion to extracellular matrix proteins...... 30

2.5. Working with nucleic acids ...... 31

2.5.1. Chromosomal DNA extraction ...... 31

2.5.1.1. Preparation and cell lysis ...... 31

2.5.1.2. Phenol/chloroform/isoamylalcohol extraction ...... 32

2.5.1.3. Ethanol precipitation ...... 32

2.5.2. Isolation of plasmid DNA ...... 32

2.5.3. DNA purification from agarose gels or PCR mixtures ...... 33

2.5.4. Agarose gel electrophoresis ...... 33

2.5.5. Quantification of dsDNA ...... 33

2.5.6. Oligonucleotides ...... 34

2.5.7. Amplification of DNA by polymerase chain reaction ...... 34

2.5.7.1. Temperature gradient PCR ...... 35

2.5.8. Screening colonies by PCR ...... 36

2.5.9. DNA digestion with restriction ...... 37

2.5.10. Klenow fill-in of DNA ends with 5`-overhang ...... 37

2.5.11. Ligation ...... 37

2.5.12. Sequencing of plasmid constructs ...... 38

2.5.13. RNA extraction ...... 38

2.5.14. Reverse Transcription PCR ...... 39

2.6. Transformation on E. coli ...... 40

2.6.1. Preparation of electrocompetent E. coli cells ...... 40

2.6.2. Transformation of electrocompetent E. coli ...... 41

2.6.3. Plasmid artificial modification ...... 41

2.7. Transformation of Bifidobacterium sp. strains...... 42

2.7.1. Preparation of electrocompetent bifidobacteria ...... 42

2.7.2. Transformation of electrocompetent bifidobacteria ...... 42

2.8. Working with proteins...... 43

2.8.1. Preparation of bacterial cell fractions ...... 43

2.8.2. Quantification of proteins ...... 44

2.8.3. Sodium dodecyl sulfate polyacrylamide gel electrophoresis ...... 44

2.8.4. Staining of SDS gels ...... 45

2.8.5. Western Blot ...... 46

2.9. Microscopy ...... 47

2.9.1. Ruthenium red staining ...... 47

2.9.2. Sample embedding and ultrathin sectioning ...... 47

2.9.3. Transmission electron microscopy ...... 48

2.9.4. Scanning electron microscopy ...... 48

2.9.5. Fixing and staining of cells for fluorescence microscopy ...... 48

2.9.6. Fluorescence microscopy ...... 48

2.10. Bioinformatic analysis ...... 49

2.11. Statistical analysis ...... 49

3. Results ...... 50

3.1. Bioinformatic analysis ...... 50

3.1.1. EPS ...... 50

3.1.2. Pili ...... 52

3.1.3. Cell surface proteins ...... 53

3.1.4. like protease ...... 57

3.2. Evidence for presence of adhesion structures ...... 60

3.2.1. EPS ...... 60

3.2.2. Expression analysis ...... 61

3.3. Functional characterization ...... 64

3.3.1. Cloning the putative adhesins ...... 64

3.3.2. Adhesion of the putative adhesins overexpression strains ...... 66

3.3.3. Creating fluorescent B. bifidum S17 expressing the putative adhesin bbif_0295 ..... 69

3.3.4. Competitive adhesion with fluorescent B. bifidum S17 ...... 70

3.3.5. Cloning of bifidocepin ...... 71

3.3.6. Identification of the cellular localization of BBIF_1681 ...... 73

3.3.7. Adhesion to extracellular matrix proteins...... 73

4. Discussion ...... 75

4.1. EPS…………...... 75

4.2. Putative pili ...... 77

4.3. Cell surface proteins ...... 80

4.4. Bifidocepin ...... 84

5. Summary ...... 88

6. Zusammenfassung ...... 89

7. List of Literature ...... 90

8. Addendum ...... 115

8.1. Used enzymes ...... 115

8.2. List of oligonucleotides ...... 116

8.3. Chemicals and manufacturers ...... 124

9. Abbreviations ...... 128

10. Acknowledgment ...... 132

Introduction

1. Introduction

1.1. General features of the genus Bifidobacterium

Bifidobacteria belong to the family and the phylum , which are one of the largest taxonomic units in the domain (Ventura et al., 2007). The members of this genus are anaerobic Gram-positive, that do not form spores and are non-motile (Sgorbati et al., 1995). The first Bifidobacterium was isolated by Henri Tissier in 1899, who termed these bacteria Bacillus bifidus and described them as rod-shaped with bifid morphology (Tissier, 1900) due to their typical Y-form (lat. bifidus: split in two). In 1973, bifidobacteria were reclassified in a separate taxon and Poupard et al. (1973) designated the genus Bifidobacterium.

During the last three decades bifidobacteria became a focus of research because they are an essential part of the human microbiota of the (GIT; Turroni et al., 2012). Bifidobacteria are found to stably colonize the GIT of , birds and insects (Lee and O`Sullivan, 2010; Kopečný et al., 2010; Vásquez et al., 2012). Additionally, they are isolated from a variety of ecological niches, such as human milk, dental caries, raw milk and sewage (Crociani et al., 1996; Klijn et al., 2005; Martín et al., 2009). To date, the genus Bifidobacterium includes 61 species with various subspecies (http://www.dsmz.de/?id=763) and there are hints that the number of species (sp.) will further expand in future (Bottacini et al., 2014).

The genomes of bifidobacteria are characterized by a high G+C content ranging from 55 % to 67 % (Scardovi, 1924; Ventura et al., 2004; Klijn et al., 2005). Bifidobacteria employed a specific pathway called “bifidus shunt” to ferment simple and complex (de Vries and Stouthamer, 1967). The key of this metabolic route is fructose 6- phosphate phosphoketolase (Scardovi, 1924) and the encoding gene is used as genetic marker for bifidobacteria (de Vries and Stouthamer, 1967). Fermentation of 1 mol glucose via the bifidus shunt produces acetate (1.5 Mol) and lactate (1.0 Mol) and theoretically yields 2.5 ATP molecules.

In 2011 the first completely annotated genome sequence of a B. bifidum strain was published by Zhurina et al.. The genomic information of the strain B. bifidum S17 is on a single circular chromosome with a size of 2.2 Mbp and a G+C content of 62 %. B. bifidum S17 was isolated in 2002 from the feces of a breast fed infant (Staudt, 2002). First studies with this strain have shown that this bacterium is strongly adherent to human

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Introduction intestinal epithelial cells (IECs) and possess potent anti-inflammatory capacity both in vitro and in vivo in murine models of colitis (Riedel et al., 2006a; Preising et al., 2010; Philippe et al., 2011). Since then, this strain has been the subject of various other studies. For example, B. bifidum S17 was shown to produce a species-specific lipoprotein BopA, which acts as moonlighting protein and plays a role in adhesion to host tissue (Gleinser et al., 2012). Also, the presence of genes encoding for other potential host-colonization factors were analyzed (Westermann et al., 2012a). However, the mechanisms contributing to the beneficial effects of this strain are still unknown due to the lack of appropriate tools for genetic modification (Brancaccio et al., 2013a).

1.2. Bifidobacteria as a part of the human microbiota

The human GIT is the habitat for a variety of that outnumbers the cells of the human body by a factor of 10 (Backhed et al., 2005). This makes the GIT one of the most densely populated habitats on earth (Whitman et al., 1998; Backhed et al., 2005). The number and complexity of microbial populations gradually increase from the stomach to the colon (Riedel et al., 2014). Most of the members of the are engaged in commensal or mutualistic interactions with the host. The gut microbiota as a whole was shown to be an important asset to the GIT, which has a crucial impact on both in health and disease (Young, 2012). It is involved in the development of the immune system, exclusion of pathogens, production of vitamins and the metabolism of fatty acids, glucose, and bile acids (Deguchi et al., 1985; Baquero and Nombela, 2012).

Dysbiosis in the composition of the microbiota was shown to be related to diverse functional disorders of the gut including inflammatory bowel disease (Tamboli et al., 2004), irritable bowel syndrome (Kassinen et al., 2007), stomach cancer (Parsonnet et al., 1991), mucosa-associated lymphoid tissue lymphoma (Mesnard et al., 2012), obesity (Turnbaugh et al., 2006; Delzenne et al., 2011) and necrotizing enterocolitis (de la Cochetière et al., 2004).

The composition of the microbiota changes during the development to an adult but is reported to be stable in healthy individuals over time once adult’s microbiota is established (Tap et al., 2009; Claesson et al., 2011). The development of the (gut) microbiota starts with the colonization of the GIT at the really beginning of life and is influenced by numerous factors, including the birth process (Musilova et al., 2015), infant diet (breast or

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Introduction formula feeding), nutrition, antibiotic treatment and hygiene conditions (Fanaro et al., 2003; Cheng et al., 2013). Until recently it was believed that the GIT of the newborn is sterile. However, a recent study suggests bacteria are present at low levels in meconium and umbilical cord blood (Cheng et al., 2013). In adults, the dominant phyla of the gut microbiota are Bacteroidetes and Firmicutes, with a significant portion of the Actinobacteria class (Eckburg et al., 2005).

Bifidobacteria are one of the first colonizers of the GIT of newborns (Reuter, 2001; Favier et al., 2002) and are the dominating bacterial group in the colon of breast-fed infants (Kurokawa et al., 2007). The origin of infant’s bacterial community is discussed to be obtained during pregnancy from maternal placenta and during transit through the birth canal being in contact with maternal vaginal and gut bacteria, from parental skin or the environment (Adlerberth and Wold, 2009; Biasucci et al., 2010). However, recent studies showed that one further source of bifidobacteria is human milk and the transfer to newborns is done by breast-feeding (Martín et al., 2009; Fernández et al., 2013). In breast- fed neonates, B. breve, B. bifidum and B. longum dominate the intestinal microbiota (Turroni et al., 2012; Milani et al., 2013), whereas B. adolescentis, B. breve, B. longum and B. infantis are members of the adult microbiota (He et al., 2001a; He et al., 2001b). Formula-fed newborns develop a more complex flora with a more adult profile of Bifidobacterium sp., members of enterobacteria, Streptococcus sp., sp. and Clostridium sp. (Gritz and Bhandari, 2015). After weaning, the numbers of bifidobacteria are decreasing (Gritz and Bhandari, 2015). Studies have estimated that the presence of bifidobacteria in the adult GIT is around 4.3 ± 4.4 % of fecal microbes (Eckburg et al., 2005; Mueller et al., 2006). Thus, they remain a numerically and functionally important group of the microbiota throughout the life span of human being (Reuter, 2001).

1.3. Bifidobacteria as probiotics

1.3.1. Definition of probiotics

The use of live microorganisms as food supplements goes back thousands of years. One of the first food containing viable bacteria were fermented milks and they are consumed in numerous forms until today. Élie Metchnikoff proposed that the administration of fermented milk products have a positive effect on the microbiota of the human gut and therefore on human health (Metchnikoff and Chalmers, 1908; Fuller, 1992). Since that

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Introduction time, harmless bacteria were used as food additives as preventive approach to maintain the balance of the intestinal microbiota and to enhance human well-being (Reuter, 1997; Saarela et al., 2000). Today, probiotics are most widely defined as “live microorganisms which when administered in adequate amounts confer a health benefit on the host” (Food and Agriculture Organization and World Health Organization). However, this definition may change in the future, especially since it has been shown that also inactivated probiotics or preparations of cell structures have beneficial effects on human health (Lee and Salminen, 2009; Kolling et al., 2015).

1.3.2. Probiotic bifidobacteria and their health-promoting effect on human host

Numerous health promoting effects have been associated to bifidobacteria and therefore different strains are extensively used as probiotics by the food and dairy industry worldwide (Masco et al., 2005; Granato et al., 2010). Probiotic bifidobacteria were shown to produce compounds that inhibit the growth of gut pathogens and compete with their adhesion sites and therefore prevent the colonization of pathogens such as Salmonella enerica serovar Typhimurium, Escherichia coli, Listeria monocytogenes, Enterobacter sakazakii, and Clostridium difficle (Collado et al., 2005; Collado et al., 2006). Furthermore, they stimulate the immune response (Gill, 1998; You and Yaqoob, 2012) and display preventive effects in diarrheal diseases (Saavedra et al., 1994; Allen et al., 2013) and colon cancer (Wollowski et al., 2001; Roller et al., 2004; Rafter et al., 2007;). Probiotic bifidobacteria were also shown to improve hypercholesterolemia (Bordoni et al., 2013; Guardamagna et al., 2014), lactose intolerance (Jiang et al., 1996) and stabilize the gut mucosal barrier (Kailasapathy and Chin, 2000; Ewaschuk et al., 2008). Additionally, there are studies showing that bifidobacteria are able to reduce the pro-inflammatory state of patients with inflammatory disorders of the GIT (O'Mahony et al., 2005; Whorwell et al., 2006; Groeger et al., 2013). However, beneficial effects of bifidobacteria and other probiotics on human health are found to be, in most cases, species- and even strain-specific features (Ramos et al., 2013).

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Introduction

1.4. Potential adhesive structures of the human gut

In order to exert their health-promoting effects, bacteria must be able to reach the GIT in viable form and persist for at least a certain period of time in the gut (Serafini et al., 2013). Beside physiological challenges such as gastric acid and bile salt, adhesion of bifidobacteria to human mucus, intestinal epithelial cells, and/or components of the extracellular matrix is an important factor and has frequently been used as a criterion for selection of probiotic bacteria (Dunne et al., 2001).

1.4.1. Mucus

The intestinal epithelium is largely covered with mucus (Figure 1). Mucus is a protective gel on the surface of the mucosal epithelium that prevents access of many pathogens, toxins, and other damaging agents to IECs (Florey, 1955; Macfarlane et al., 2005). Mucus is produced and secreted by goblet cells that are interspersed along the crypt-vilus axis of the mucosa. It is a complex matrix that consists of mucins, water electrolytes, detached cells, enzymes, antibodies, nucleic acid, and also bacteria and bacterial products (Hotta, 2000; Macfarlane et al., 2005).

The major components are mucins that are complex glycoproteins in which the central peptide core are closely packed with side chains (Hayashida et al., 2001; Ikezawa et al., 2004). Mucin monomers are joined by disulfide bridges to form high- molecular-weight polymers. MUC2 is the predominant mucin of the intestinal mucus (Wilson, 2009). The secreted mucus consists of the inner mucus layer that is sterile and the loosely attached outer mucus layer, which contains bacteria (Figure 1; Johansson et al., 2008; McGuckin et al., 2011).

The core protein as well as the carbohydrate side chains of mucin are possible adhesion sites for bacteria. In several lactobacilli species mucin-binding proteins were found (Rojas et al., 2002, Macías-Rodríguez et al., 2009; van Tassel and Miller, 2011). Adhesion to mucus is a multifaceted process, including non-specific adhesion by electrostatic or hydrophobic forces and specific binding mediated by particular molecules (Muñoz- Provencio et al., 2009). Additionally, mucus has been reported to be a source of nutrients for bacteria (Derrien et al., 2004; Ruas-Madiedo et al., 2008).

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Introduction

Figure 1: The intestinal epithelium with the mucus layer and extracellular matrix. Molecules of the ECM are shed into the mucus layer. Damaged IECs expose ECM structures.

1.4.2. Intestinal epithelial cells and the glycocalyx

IECs are largely covered with a mucus layer. In this context, bacteria have only limited access to IECs (Corazziari, 2009). However, it is not excluded that some bacteria reach the epithelial surface, e.g. by degradation of mucus (McGuckin et al., 2011). Moreover, in several diseased states, the mucus layer is disturbed, markedly reduced, or completely absent (Swidsinski et al., 2007; McGuckin et al., 2011; Hoskins, 2012).

The epithelial surface is a single layer of epithelial cells organized into vili, that are projections into the lumen and crypts, which builds tubular invaginations besides the vili. At the base of the crypts reside pluripotent intestinal epithelial stem cells, which continually renew this surface. Absorptive enterocytes, which possesses metabolic and digestive function, borders the epithelial surface. Secretory IECs, including enteroendocrine cells, mucin producing goblet cells and antimicrobial compound producing Paneth cells exhibit digestive or barrier functions of the epithelium (Peterson and Artis, 2014). Epithelial cells are covered with a thin layer called glycocalyx (Figure 1). The main components of the glycocalyx are apical membrane bound mucin proteins. The mucins are densely packed and form long and extended rods described as bottle brushes (Sansonetti, 2004; Johansson et al., 2011).

Possible adhesive sites of IECs are best studied for structures of gastrointestinal pathogens (Lu and Walker, 2001; Lebeer et al., 2010). Interestingly, there are more studies directing adhesion structures compared than host receptors binding to these bacterial

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Introduction structures. The most common receptors of IECs are integrins, cadherins and members of the immunoglobulin-related cell adhesion molecule families, which are frequently recognized by specific pathogenic surface structures (Hauck et al., 2006). Furthermore, pattern recognition receptors detect microorganism-associated molecular patterns, which are widespread and conserved among microorganisms. Toll-like receptors are the best studied pattern recognition receptors, which are transmembrane proteins present at the cell surface of IECs (Lebeer et al., 2010). Nucleotide-binding oligomerization domain-like receptors and C-type lectin receptors are also able to sense pathogenic and commensal bacteria and therefore are possible binding sites (Lebeer et al., 2010; Nishio and Honda, 2012). Probiotic bacterial binding sites on the epithelial cell surface are widely uncharacterized. However, selected probiotic bacteria were shown to adhere to non-mucin producing human cell lines and are able to exclude pathogens and this may occur by competing for the same receptors and binding sites (Servin, 2004; Buffie and Pamer, 2013).

1.4.3. Extracellular matrix proteins

If the epithelial layer is damaged, bacteria gain access to the underlying connective tissue, i.e. the extracellular matrix (ECM). The ECM is a structural support network building a stable macromolecular structure underlying epithelial cells and surrounds connective tissue cells (Figure 1). The ECM consists mainly of collagen, laminin, and fibronectin (Westerlund and Korhonen, 1993).

Collagens are the predominant form of structural proteins found in ECM. In vertebrates system 30 different collagens have been described, which differ in size, function, and tissue distribution (Heino, 2007; Sato et al. 2002; Rozario and de Simone, 2010). Collagen type I is the dominant form found extensively in almost all tissues, type IV and VIII can be found in epithelial cells, but nevertheless there are also other collagen types which are fiber components of most body tissues (Gelse et al. 2003). Collagen is arranged into fibrils, which imparts tensile strength to connective tissues (Badylak et al., 2009).

Beside collagen, laminin is a ubiquitously glycoprotein present as a part of the ECM. Laminins are multifunctional heterotrimeric molecules, whereas 16 types were identified in vertebrates on the basis of different combinations of three subchains. The chains interconnected with each other and form a “crucifix”-shape (Singh et al., 2012).

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Introduction

Fibronectin is also a key compound of the ECM. It is arranged into a mesh of fibrils and linked to cell surface receptors called integrins. Fibronectin is expressed by various cell types. The smallest unit of Fibronectin can be divided into three different types that are repeated within one subunit which form dimers (Mao and Schwarzbauer, 2005).

Fibrinogen is normally found as a major plasma glycoprotein playing a key role in inflammation and in reactions (Upadhyay et al., 2012). However, fibrinogen has been identified as adhesion mediating factor of pathogens and probiotics (Bouchara et al., 1995; Collins et al., 2012). Thus, it is speculated that this structure is a possible for bacteria.

ECM structures have been described in the literature as common bacterial binding substrates. Molecules of the ECM can be shed into the mucus layer from the epithelium. Furthermore, damaged epithelial cells expose the ECM and allow microbial contact (Figure 1; Vélez et al. 2007). The importance of adherence to ECM has been shown for various pathogens and their mechanisms to establish infection (Westerlund and Korhonen, 1993). Pathogenic adhesion to ECM is achieved by various microbial surface exposed proteins with ECM domains (Atzingen et al., 2009; Hendrickx et al., 2009). Probiotics are able to competitive exclude pathogens, speculating that they use the same receptors and binding sites (Servin, 2004; Buffie and Pamer, 2013).

1.5. Bacterial cell surface architecture – potential adhesion mediating structures

Adhesion of commensal to IECs is a complex and multifactorial process. Interaction of bacteria with the host is influenced by the properties of the outer bacterial envelope such as hydrophobicity, roughness or local charge (Pérez et al., 1998; Del Re et al., 2000; Pan et al., 2006). These properties are provided by proteins and other components of the bacterial cell surface (Ventura et al., 2012). By mediating contact between bacteria and host structures bacterial surface contribute to colonization of the GIT and trigger host responses (Kelly et al., 1999).

As Gram-positive organisms, bifidobacteria possess a cell envelope with a complex architecture (Figure 2; Navarre and Schneewind, 1999). Bacterial surface components like proteins, carbohydrates and lipids are naturally the first structures engaged in mediating adhesion to host cells (Ventura et al., 2012; Gonzáles-Rodríguez et al., 2013; Gueimonde

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Introduction et al., 2007). Several studies showed the involvement of extracellular proteins in adhesion of bacteria to IECs (Lebeer et al., 2010; Sánchez et al., 2008; Kleerebezem et al., 2010).

Exopolysaccharides (EPS, section 1.5.1.), which are carbohydrate polymers present on the surface of many bacteria, are one group of molecules involved in adhesion to IECs (López et al., 2012; Fanning et al., 2012b).

Cell envelope proteases (CEPs, section 1.5.2.) were shown to promote bacterial survival and colonization of epithelial surfaces (Schmidtchen et al., 2002; Serruto et al., 2003; Hidalgo-Grass et al., 2006; Karlsson et al., 2007;).

Some recent studies have demonstrated that cell surface pilus structures of bifidobacteria are essential and conserved host-colonization factors (Foroni et al., 2011; O'Connell Motherway et al., 2011; section 1.5.3.).

Beside the more complex pili structures, there are also single proteins shown to mediate adherence to host structures (Candela et al., 2007; Guglielmetti et al., 2008; Candela et al., 2009; Candela et al., 2010; Gleinser et al., 2012; section 1.5.4.)

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Introduction

Figure 2: Schematic overview of the bifidobacterial surface decoration. Schematic representations of different single adhesins, exoposysaccharide capsule, extracellular proteases and pili with subunits. 1 Source for image of pili pictures: O'Connell Motherway et al., 2011.

1.5.1. Exopolysaccharides

Many bacteria produce a protective coating of carbohydrate polymers known as exopolysaccharides (EPS). These structures can also be found on different strains of Lactobacillus and Bifidobacterium (Ruas-Madiedo et al., 2006) and are either bound to the cell wall, loosely attached to the surface or released into the surrounding environment (de Vuyst et al., 2001; Badel et al., 2011). Bacterial EPS consist of repeating mono- or oligosaccharide subunits, e.g. D-glucose, D-galactose and L-rhamnose, which are connected to each other by varying glycosidic linkages. The resulting homo- or heteropolymers are structurally very diverse and hence differs in their chemical and physical properties (Ruas-Madiedo et al., 2002; Hidalgo-Cantabrana et al., 2014).

EPS seem to possess a number of physiological roles in bacterial ecology. Diversity, chemical characteristics, amount and functionalities of EPS were investigated in detail in lactobacilli and other lactic acid bacteria (Tallon et al., 2003; Mozzi et al., 2006; Nikolic et

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Introduction al., 2012). In bifidobacteria their function is incompletely understood (Fanning et al., 2012a). The EPS of B. breve UCC2003 were shown to be involved in host colonization and protection against Citrobacter rodentium infection and reduce immunogenicity of the strain (Fanning et al., 2012a). EPS of different bifidobacteria increase tolerance to bile and low pH and thus might impact on the GIT transit and persistence in the gut (Ruas-Madiedo et al., 2009a; Alp and Aslim, 2010). Additionally, bifidobacterial EPS was shown to modulate the intestinal microbiota by providing fermentable substrates to other microorganisms (Salazar et al. 2008, Salazar et al., 2009). It was shown that bifidobacteria of human origin synthesize EPS with higher contents of rhamnose than strains of food origin (Hidalgo-Cantabrana et al., 2012). In line with the structural and physicochemical variability of bifidobacterial EPS molecules, in silico analysis revealed a considerable inter-species variability in bifidobacterial gene clusters for EPS synthesis (Bottacini et al., 2010). The G+C content of the EPS clusters differs compared to that of the whole genome, which indicates that these genes were acquired by horizontal gene transfer (Hidalgo- Cantabrana et al., 2014).

1.5.2. Cell envelope proteases

Cell envelope proteases (CEPs) are a group of surface-associated enzymes which are of special interest because they might be involved in interaction with the host due to their localization outside the cell (Navarre and Schneewind, 1999; Mitsuma et al., 2008; González-Rodríguez et al., 2012). CEPs are enzymes which degrade substrates, i.e. from milk and convert them into metabolic active compounds that might also have immunological properties (Yamamoto et al., 1998; Seppo et al., 2003; Fernández-Esplá et al., 2000). Furthermore, CEPs allow the transient binding to IECs due to catalytic breakdown of proteins into peptides or amino acids and adhere at least during the time of cleavage (Sánchez et al., 2008). Some CEPs were shown to cleave and inactivate antimicrobial peptides, which were reported to be a common strategy to promote bacterial survival and colonization of epithelial surfaces (Schmidtchen et al., 2002; Serruto et al., 2003; Hidalgo-Grass et al., 2006; Karlsson et al., 2007). Additionally, the cleavage of possible matrix proteins opens the possibility for bacteria to adhere to subjacent structures.

CEPs possess a C-terminal LPXTG motif for covalent attachment to the cell wall. The LPXTG motif is recognized by a housekeeping sortase, a surface enzyme that catalyses the

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Introduction cleavage and subsequently links the protein covalently to the peptidoglycan network (Navarre and Schneewind, 1999).

1.5.3. Pili

In Gram-negative bacteria, proteinaceous surface appendages such as pili and fimbriae are involved in bacterium-host interactions (Lynch and Crease, 1990). Genes encoding both type IV tight adherence (Tad) pili and sortase dependent Fim pili (-associated adhesion) have identified in bifidobacteria with varying numbers of gene clusters in different species. Pilus-like structures have been described for various Lactobacillus and Bifidobacterium sp. (Kankainen et al., 2009; Foroni et al., 2011; O'Connell Motherway et al., 2011; Turroni et al., 2013).

The tad genes encoding the tight adherence pili have a typical organization with genes for the pilus assembly complex (tadZABC) and those for the pilus subunits (flp and tadE) being located in two adjacent gene clusters and separately located peptidase (tadV) (O'Connell Motherway et al., 2011). The pilus assembly complex consists of the peripheral cytoplasmic membrane located ATPase TadA, which provides the energy for pilus assembly. TadB and TadC in association with TadA form homo- or hetero-oligomeric integral membrane complexes in the membrane for translocation of the pilus subunits. The function of TadZ still remains unclear but it might be responsible for the localization of the pilus assembly complex to the cell poles. Flp is the major structural component of Tad pili and TadE represents the pseudopilin due its pilin-like features but this protein is not assembled into the Flp pilus and only decorates the pilus structure. Flp and TadE are synthesized as prepilins with a hydrophobic leader peptide that is processed by the TadV peptidase. Tad pili are synthesized as bundles of long, thin fibrils consisting of the Flp subunits proteins and their diameter can range between 40 Å and 80 Å (Tomich et al., 2007; O'Connell Motherway et al., 2011). However, precise determinations of the length and diameter of Tad pili of bifidobacteria have not been performed so far.

Typical fim gene clusters for sortase dependent pili encompasses genes encoding for the major pilin subunit, one or two ancillary minor pilin subunits, and the specific class C sortase. The subunits contain the pilin-like motif (TVXXK), a Sec-dependent secretion signal, and a cell wall anchor motif (LPXTG). The major pilin subunit builds the pilus shaft, whereas the minor pilin subunit represents the tip protein. After synthesis of the

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Introduction subunits in the cytoplasm, the proteins are secreted by the Sec secretion system, while signal peptidases cleave off the signal peptides thereby producing precursors of the pilus subunits. During polymerization of pili, these precursor subunits are cross-linked by the specific class C sortases (Hendrickx et al., 2011; Foroni et al., 2011; Turroni et al., 2014).

In B. bifidum S17 one putative tad gene cluster and three clusters for potential sortase- dependent fim pili were identified by in silico analysis (Westermann et al., 2012a). However, these gene clusters and the corresponding pili have not been characterized so far.

1.5.4. Identified single adhesins

Over last several decades, surface proteins of pathogens and their role in host colonization, cell invasion, and immune response modulation have been studied extensively (Patti et al., 1994; Beckmann et al., 2002; Selbach and Backert, 2005; Lilic et al., 2006; Luck et al., 2006; Timmer et al., 2006; Gillen et al., 2008). However, colonization and communication with the host are not limited to pathogens. Similar to pathogens, commensal bacteria colonizing the human GIT interact with each other and the host albeit with different outcomes. It should thus be not surprising that these processes are also mediated by surface proteins (Kolenbrander et al., 2006; Sansonetti and Medzhitov, 2009; Sansonetti, 2011). Interestingly, many of the identified bacterial adhesins are so called moonlighting proteins. Despite their known or assumed intracellular functions, these proteins were shown to mediate adhesion to host structures (Huberts and van der Klei, 2010).

S-layer proteins form the outermost interacting surface of different bacterial species and were shown to act as adhesion factor to the intestinal epithelium (Åvall-Jääskeläinen et al., 2003; Kos et al., 2003). Furthermore, the cell and mucus-binding protein A (CmbA) and mucus adhesion promoting protein (MapA) were described in different Lactobacillus reuteri strains as cell surface proteins playing a role in binding to IECs and mucus (Miyoshi et al., 2006; Jensen et al., 2014). Another group of proteins playing a role in adhesion process of commensal lactobacilli were cleaved by sortase A and anchored to the cell wall (van Pijkeren et al., 2006; Muñoz-Provencio et al., 2012). Recently, glyceraldehydes-3-phosphate dehydrogenase (GAPDH) of Lactobacillus reuteri ZJ617 was shown to act as an adhesion component that plays a role in binding of this strain to IECs (Zhang et al., 2015). The role of collagen binding protein as well as auto- aggregation-promoting protein (AggLb) were proven as crucial surface proteins

20

Introduction contributing adhesion of various Lactobacillus strains to collagen (Yadav et al., 2013; Miljkovic et al., 2015).

In bifidobacteria, the first identified single adhesion protein BopA of B. bifidum MIMBb75 was shown to be involved in adhesion to Caco-2 cells (Guglielmetti et al., 2008). Furthermore, lipoprotein BopA was identified in another B. bifidum and B. longum strain and its role in adhesion process to IECs were confirmed (Gleinser et al., 2012). Moreover, bifidobacterial chaperon DnaK, glutamine synthetase, enolase, bile salt , and phosphoglycerate mutase were shown to be involved in adhesion process (Candela et al., 2007; Candela et al., 2009; Candela et al., 2010).

1.5.4.1. Translocation and cell wall anchoring of cell surface proteins

Extracellular proteins contain sorting signal sequences for protein secretion and correct localization to the target site. In general, there are different secretion mechanisms in Gram- positive bacteria. The main system to secrete proteins into the extracellular environment are the twin arginine translocation (Tat), the hole forming (holin), the WXG100-secretion (WSS), and the secretory pathway (Sec) systems (Desvaux et al., 2009). In Gram-positive bacteria, proteins that are targeted for localization in the cell envelope are mainly secreted via the Sec system (Figure 3; Pallen et al., 2003).

The Sec translocation system consists of a set of integral membrane proteins and is conserved among bacteria, archaea, and eukaryotes (Pohlschröder et al., 1997). The translocon channel is an integral membrane complex composed of SecY, the largest subunit of the protein , SecE, a protein essential for translocation, and SecG. The associated SecA is an ATPase that couples the energy of ATP binding and hydrolysis to the stepwise translocation of proteins (Schiebel et al., 1991; van der Wolk et al., 1993; van der Wolk et al., 1995). Translocated proteins are synthesized as precursors with a cleavable signal peptide (SP). SPs can be located at the N- or C-terminus of a polypeptide. N-terminal SPs contain a stretch of several positively charged lysine and arginine that is followed by a hydrophobic region, which tend to build α-helices when in contact with membrane lipids. C-terminal SPs are rich in hydrophilic amino acids and contains a SP cleavage site (often Ala-X-Ala). After synthesis of the preprotein, two possible translocation routes are possible.

21

Introduction

During the co-translation translocation the protein synthesis machinery is in contact with the translocation channel and the protein is translocated directly without any intermediate states. Alternatively, preproteins are first completely synthesized and then translocated in unfolded conformation. In this case, cytoplasmic chaperons such as GroEl or DnaK are required to prevent folding or aggregation (Beissinger and Buchner, 1998; Fink, 1999). This mechanism is termed post-translation. After crossing the membrane, maturation of the precursor protein to its final folded conformation via sortases takes place (Navarre and Schneewind, 1999; van Wely et al., 2001; Sánchez et al., 2008; Desvaux et al., 2009).

Membrane proteins and lipoproteins are translocated through the bacterial membrane by the integral membrane protein YidC. YidC is associated with the SecYEG complex (Scotti et al., 2000), but is also able to act autonomously for some membrane anchored proteins (Samuelson et al., 2000). YidC interact with the transmembrane segments of these proteins and releases transmembrane helices either one by one or as bundles into the lipid bilayer. Furthermore, YidC is proposed to act as a chaperone during passage of the transmembrane segments (Xie and Dalbey, 2008).

In B. bifidum S17 genes encoding the Sec dependent secretion machinery were found by in silico analysis but the Tat system seems to be missing. The Sec components of In B. bifidum S17 are: SecY (BBIF_1484); SecE (BBIF_0279); SecG (BBIF_0980); SecA (BBIF_1223); YidC (BBIF_1782). It is thus likely that the Sec system is the most relevant system for transport of proteins across the bacterial membrane (Oßwald et al., 2015).

22

Introduction

Figure 3: Schematic representation of different types of surface proteins and their translocation system in Gram-positive bacteria. Proteins containing transmembrane domains anchored to the cytoplasmic membrane (CM). Cell wall-associated proteins are translocated by the Sec system and attached to the cell wall by either (i) covalent linkage via a LPXTG motif or by (ii) non- covalent interactions via e.g. Ig-like or cell wall binding domain(s) (CWBDs). (SP: signal peptide, EC: extracellular, EPS: exopolysaccharides, PG: peptidoglycan, CM: cell membrane, CP: cytoplasm).

1.6. The aim of this study

In this work, possible factors involved in host-colonization of B. bifidum S17 were investigated. B. bifidum S17 is a promising probiotic strain with anti-inflammatory capacity (Preising et al., 2010; Riedel et al., 2006a) and was isolated from the feces of a breast-fed infant (Staudt, 2002). For this purpose, structures displayed on the cell surface were analyzed in detail through bioinformatic approaches, microscopy, expression studies, adhesion assays and generation of overexpression mutants. The identification of structures mediating adhesion to host tissue is of pivotal importance because adhesion to host tissue supports microbe-host-interactions and thus the beneficial health effects of probiotic bifidobacteria.

23

Materials and Methods

2. Materials and Methods

All chemicals used in this study are listed in Table 16 (Addendum) with their corresponding manufacturers.

2.1. Bacterial strains

All bacterial strains used in this study are listed in Table 1. The strains were stored as glycerol stocks at -80 °C. To prepare glycerol stocks, bacteria were grown under strain specific optimal conditions (see sections 2.1.1. and 2.1.2.) over night (o/N) in 10 ml of the appropriate medium and harvested by centrifugation (10 min, 4,000 x g). The bacterial pellet was resuspended in an adequate amount of fresh medium containing 30 % glycerol (v/v, 99 % pure), transferred to a cryotube, vortexed and stored at -80 °C.

2.1.1. Bifidobacteria

Bifidobacteria were routinely cultivated in de Man-Rogosa-Sharp medium (MRS, 55 g/l lactobacilli browth powder; de Man et al. 1960) supplemented with 0.5 g/l L-cysteine hydrochloride monohydrate (MRSc). Bacterial cultures were grown for 16 h at 37 °C to stationary growth phase by incubation statically in sealed jars under anaerobic conditions using AnaeroGenTM sachets (Thermo Scientific, Rockfort, USA). AnaeroGenTM reduces oxygen levels in sealed jars below 1 % within 30 min and simultaneously generates carbon dioxide (CO2) levels between 9 % and 13 %. For the cultivation of strains carrying plasmids the appropriate amount of antibiotic were added to MRSc. Unless stated otherwise, concentrations of 100 µg/ml spectinomycin, 1 µg/ml ampicillin, 5 µg/ml chloramphenicol, or 1 µg/ml erythromycin were used.

24

Materials and Methods

Table 1: Bacterial strains used in this study.

Strain Characteristics Reference

Escherichia coli

E. coli DH10B F- endA1 recA1 galE15 galK16 nupG rpsL InvitrogenTM, ΔlacX74 Φ80lacZ ΔM15 araD139 Darmstadt, Δ(ara,leu)7697 mcrA Δ(mrr-hsdRMS-mcrBC) Germany λ-

E. coli Genomic integration of p16S_ParaB_bbif0710 Brancaccio, 2013b R R ET12567::p16S_ParaB_bbif0710 into 16S-rDNA, CM , Erm , bbif_0710 encodes for a methyltransferase of B. bifidum S17

Bifidobacterium

B. bifidum S17 Intestinal isolate of a breast-fed infant Staudt, 2002

B. longum E18 Intestinal isolate of an adult Staudt, 2002

Rhodobacter

Rhodobacter capsulatus Slime-producing bacteria Drepper, RC Jülich1

Rhodobacter capsulatus RC- Tn5 disrupted genes necessary for Drepper, RC K1 exopolysaccharide synthesis Jülich1 1 Dr. Thomas Drepper, Institute for Molecular Enzyme-Technology, Heinrich-Heine-University of Düsseldorf, Research Center Jülich, Jülich, Germany

2.1.2. Escherichia coli

Lysogeny Broth (LB; 10 g/l tryptone, 5 g/l yeast extract, 10 g/l sodium chloride; Bertani G. 1951) was used for routine cultivation of Escherichia coli (E. coli) strains. Bacteria were grown o/N to stationary growth phase under aerobic conditions at 37 °C with agitation on a rotary shaker (120 rpm; B. Braun Biotech International GmbH, Melsungen, Germany).

E. coli ET12567::p16S_ParaB_bbif0710 was grown in 2 x TY medium (16 g/l tryptone, 10 g/l yeast extract, 5 g/l sodium chloride). Antibiotics were added to the medium as appropriate for the cultured strains at the following concentrations: ampicillin and spectinomycin 100 µg/ml, chloramphenicol 15 µg/ml and erythromycin 300 µg/ml.

25

Materials and Methods

2.2. Plasmids

All plasmids used in this study are listed in Table 2. Maps of plasmids, operons and genes were created using Clone Manager Suite 7 software (Scientific & Educational Software, Cary NC, USA).

Table 2: continuedPlasmids. used in this study.

Plasmid Relevant characteristics Reference pMDY23_Pgap pMDY23 E. coli-Bifidobacterium shuttle vector, Grimm et al., 2014 expression of gusA driven by the promoter of glyceraldehydes-3-phosphate dehydrogenase gene R bbif_0612 (Pgap) of B. bifidum S17, Spc pVG-GFP pMDY23, expression of the green fluorescent protein Grimm et al., 2014 (GFP) under control of constitutive Pgap promoter, SpcR pVG-mCherry pMDY23, expression of the fluorescent protein Grimm et al., 2014 mCherry under control of constitutive Pgap promoter, SpcR pCW pMDY23, containing the constitutive expressed Pgap this work promoter, SpcR pCW_17 pMDY23, expression of the single adhesin bbif_0017 this work R under control of constitutive Pgap promoter, Spc pCW_295 pMDY23, expression of the single adhesin bbif_0295 this work R under control of constitutive Pgap promoter, Spc pCW_450 pMDY23, expression of the single adhesin bbif_0450 this work R under control of constitutive Pgap promoter, Spc pCW_540 pMDY23, expression of the single adhesin bbif_0540 this work R under control of constitutive Pgap promoter, Spc pCW_665 pMDY23, expression of the single adhesin bbif_0665 this work R under control of constitutive Pgap promoter, Spc pCW_1426 pMDY23, expression of the single adhesin bbif_1426 this work R under control of constitutive Pgap promoter, Spc

26

Materials and Methods

Table 2: continued.

Plasmid Relevant characteristics Reference pCW_1769 pMDY23, expression of the single adhesin bbif_1769 this work R under control of constitutive Pgap promoter, Spc pCW_1681 pMDY23, expression of the serine-like protease this work bbif_1681 under control of constitutive Pgap promoter, SpcR pCW_mCherry_295 pMDY23, expression of the single adhesin bbif_0295 this work under control of synthetic constitutive Phyper promoter, expression of the fluorescent protein R mCherry under the constitutive Pgap promoter, Spc

2.3. Eukaryotic Caco-2 cell line

2.3.1. Subcultivation of Caco-2 cells

The human colon adenocarcinoma cell line Caco-2 (Fogh et al., 1977) was used for all experiments in this work and obtained from the American Type Culture Collection (Cat.#: ATCC® HTB-37™). Caco-2 cells were routinely cultivated in Dulbecco´s Modified Eagle´s Medium (DMEM) supplemented with 10 % (v/v) fetal calf serum (FCS), 1 % (v/v) non-essential amino acids (NEAA), 1 % (v/v) penicillin-streptomycin solution, and 2 mM L-glutamine. Before use, FCS was heat-inactivated at 56 °C for 30 min.

Caco-2 cells were handled according manufactures instructions in tissue culture flasks of appropriate size (tissue culture treated, Falcon® Becton Dickinson Labware Europe, Le Pont de Claix, France) and incubated in CB 150 Binder incubator (Binder GmbH, Tuttlingen, Germany). Upon reaching 80 - 90 % confluence, Caco-2 cells were split and subcultured according to the supplier´s recommendation at a ratio of 1:5. For this purpose, the base growth medium was removed and the cells were washed with pre-warmed cell culture phosphate buffered saline (PBS). 3 ml -ethylendiaminetetraacetic acid

(EDTA) solution was added and incubated at 37 °C and 5 % CO2 for 5 – 15 min until the adherent Caco-2 cells were detached. Following detachment 7 ml of complete growth medium was added and the cells were pelleted in a Falcon tube by centrifugation at RT for

27

Materials and Methods

2 min at 300 xg. After removing the supernatant, the cell pellet was resuspended in base growth medium and seeded into fresh tissue culture flasks.

2.3.2. Cryoconservation of Caco-2 cells

Cells were stored at liquid nitrogen at a concentration of 2 x 106 cells/ml in freezing medium (DMEM containing 50 % (v/v) FCS, 10 % (v/v) DMSO (99.8 % pure), 1 % (v/v) NEAA) in cryo tubes. Before long time storage at -196 °C, temperature was ramped down slowly by incubating cells in cryo tubes at -80 C on isopropanol (99 % pure) for 12 h.

2.3.3. Thawing of cryo-stocks

Cells were thawed on ice and then resuspended in 20 ml of the pre-warmed base medium. DMSO of the freezing medium was removed by centrifugation at room temperature (RT) for 2 min and 300 xg. Caco-2 cells were resuspended in 6 ml DMEM medium containing 10 % (v/v) FCS, 1 % (v/v) NEAA and 1 % (v/v) penicillin-streptomycin. Cells were 2 transferred to 25 cm tissue culture flasks, incubated at 37 °C with 5 % CO2 and routinely cultured as described above.

2.3.4. Determination of the cell number

For seeding and storage of Caco-2 cells, the cell number was determined. After detachment of adherent cells with trypsin-EDTA, as described above and an aliquot of 10 µl was mixed with 90 µl trypan blue. 10 µl of stained cells were counted in a hemocytometer (ASSISTEN, Glaswarenfabrik, Karl Hecht GmbH & Co KG, Sondheim v. d. Rhön, Germany) placed under the microscope (Primo Vert, Carl Zeiss AG, Oberkochen, Germany). The numbers of cells in four large squares were counted and cell concentration was calculated using the following formula 1:

푁 ∙ 퐷퐹 ∙ 퐾 푐 = 푐 푁푆

Formula 1: Calculation of the cell concentration. With c: cell concentration (cell/ml), Nc: number 4 -1 of counted cells in all squares, DF: dilution factor, K: chamber factor (1 x 10 µl ), and NS number of counted squares.

28

Materials and Methods

2.4. Adhesion assays

2.4.1. Adhesion assay with Caco-2 cells

Adhesion assay were performed as described in Gleinser et al. (2012) with minor modifications. Briefly, Caco-2 cells were seeded at a density of 2 x 105/well in 24 well plates (tissue culture treated, CELLSTAR®, Greiner Bio-One GmbH, Germany,

Frickenhausen) and incubated at 37 °C with 5 % CO2 for 18 - 21 days to fully differentiated monolayers. At this stage about 1 x 106 cells were counted per well. Medium was changed every two to three days. For adhesion experiments, o/N cultures of 7 bifidobacteria were adjusted to an OD600 of 1 (i.e. approx. 5 x 10 colony forming units per milliliter (cfu/ml)) in DMEM (1 % NEAA). 1 ml bacterial suspension were added to each well resulting in a bacteria:cell ratio of 50:1. Under these conditions, viability of bacteria and Caco-2 cells is not affected for at least 4 h (Riedel et al., 2006a). After 1 h of incubation at 37 °C with 5 % CO2 unbound bacteria were removed by washing the Caco-2 cells three times with PBS. After adding of 1 ml DMEM medium the cell monolayer was scraped off and serial 10-fold dilutions in PBS were plated in spots of 10 µl on MRSc agar plates for determination of colony forming units (CFU) of adherent bacteria. The wells containing the reference strains were not washed and Caco-2 cells with adherent bacteria were directly removed after adhesion incubation time for plating the dilutions on MRSc agar plates for enumeration of CFU of initially added bacteria. After two days of anaerobic incubation at 37 °C the colonies were counted. Adhesion of bifidobacteria to Caco-2 cells was then calculated as percentage of the adherent CFU relative to CFU initially added to Caco-2 cells. All adhesion assays were performed in one technical triplicate and with Caco-2 cells of three independent passages and three independent bacterial cultures (biological replicates). Statistical analysis was performed using ANOVA.

2.4.2. Competitive adhesion assay

Competitive adhesion of bifidobacteria was performed using the fluorescent strains B. bfidium S17/pVG-GFP and B. bifidum S17/pCW_mCherry_295. Adhesion to Caco-2 cells was performed as described above (see section 2.4.1.). However, the bacterial

29

Materials and Methods

6 inoculum was adjusted to OD600 0.1 (i.e. 5 x 10 cfu/ml). For competitive adhesion, 500 µl of B. bifidum S17/pVG-GFP were mixed with 500 µl B. bifidum S17/pCW_mCherry_295 and the mixture was added to Caco-2 cells. Bacterial cells were allowed to adhere during an incubation time of 1 h at 37 °C with 5 % CO2. All unbound bacteria were then removed by washing the Caco-2 cells three times with PBS. Adherent bacteria were fixed (see section 4.5.) and fixed specimens were analyzed by fluorescence microscopy (see section 4.6.). Pixel values of three images were analyzed using ImageJ 1.48v software.

2.4.3. Adhesion to extracellular matrix proteins

Adhesion of bifidobacteria to extracellular matrix proteins were tested as described previous (Muñoz-Provencio et al., 2009) with minor modifications. 96-well microtiter plates were coated with 100 µl/well fibronectin (2 mg/ml) or fibrinogen (2 mg/ml, from bovine plasma) in 50 mM carbonate/bicarbonate buffer (pH 9.6, 1.5 mM di-sodium carbonate, 35 mM sodiumhydrogencarbonate) or 100 µl/well collagen (100 µg/ml, rat tail) in PBS (pH 5.5), by incubation o/N at 4 °C. For fibronectin and fibrinogen 96-well Nunc Polysorp® plates (Thermo Fisher Scientific Inc., Rockford, USA) and for collagen 96-well Nunc Maxisorp® plates (Thermo Fisher Scientific Inc., Rockford, USA) were used. After immobilization of the extracellular matrix proteins, wells were washed three times with 100 µl PBS. The wells were then blocked for 1 h with PBS supplemented with 1 % (v/v) Tween 20. For adhesion assays, o/N bacterial cultures were harvested by centrifugation at

4,000 xg for 10 min. Bacterial pellets were resuspended in 1 ml PBS and OD600 was adjusted to 1 (i.e. 5 x 107 cfu/ml) in PBS. 100 µl aliquots of the cultures were added to each well and plates were incubated o/N at 4 °C. Non-adhered cells were removed by washing the wells three times with 200 µl PBS supplemented with 0.05 % Tween 20. Plates were dried at 55 °C and adhered cells were stained with 100 µl/well crystal violet (1 mg/ml) for 45 min. Wells were washed six times with 100 µl PBS and the stain was released by adding 100µl 50 mM citrate buffer (0.1 M citric acid monohydrate (solution A); 0.1 M trisodium citrate dehydrate (solution B); 59 ml solution A + 41 ml solution B, freshly prepared; pH 4.0). Absorbance was measured at 595 nm using an Infinite® M200 multimode reader (Tecan Group Ltd., Männedorf, Switzerland). Background values (wells with bound substrates and without bacterial cells) were substracted from measured values. Wells coated with 2 mg/ml bovine serum albumin (BSA) in PBS were run as controls for non-specific adhesion. All adhesions were performed in one technical triplicate with three

30

Materials and Methods independent bacterial cultures (biological replicates). The obtained results were confirmed on a qualitative basis by microscopy (40X objective, Zeiss Axio Observer.Z1 microscope, Carl Zeiss AG, Oberkochen, Germany).

2.5. Working with nucleic acids

2.5.1. Chromosomal DNA extraction

Chromosomal DNA was extracted as described by Pospiech and Neumann (1995) with slight modifications. In brief, after the enzymatic cell lysis (see section 2.5.1.1.) protein contaminations were removed by phenol/chloroform/isoamylalcohol extraction (see section 2.5.1.2.) and genomic DNA was precipitated by ethanol (see section 2.5.1.3.).

2.5.1.1. Preparation and cell lysis

For extraction of bacterial chromosomal DNA, 10 ml of a bifidobacterial o/N culture were centrifuged (10 min, 12,900 x g) and the bacterial pellet was washed once in 1 ml TE buffer (10 mM Tris-HCl (pH 7.6), 5 mM EDTA). After washing, the bacterial cell pellet was resuspended in 300 µl TES buffer (50 mM Tris-HCl, 5 mM EDTA, 10 mM NaCl, pH 8.0) and 100 µl lysozyme solution (5 mg/ml in TES buffer) and 40 µl mutanolysin solution (5 U/µl in TES buffer) were added. The reaction mixture was incubated for 2 h at 37° C.

To lyse bacterial cells, 22 µl 10 % (w/v) SDS and 15 µl proteinase K (20 mg/ml in H2O) were added and the mix was further incubated for 1 h at 55 °C with occasional shaking. Addition of 100 µl of saturated NaCl solution (5 M) and gentle inversion of the reaction tubes resulted in precipitation of proteins, visible as white film. The insoluble protein fraction was precipitated by centrifugation at 5,300 xg for 15 min at RT. Further purification of DNA from the supernatant was achieved by phenol/chloroform/isoamylalcohol extraction (section 2.5.1.2.).

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Materials and Methods

2.5.1.2. Phenol/chloroform/isoamylalcohol extraction

To remove residual protein contaminations from DNA solution one volume of phenol/chloroform/isoamylalcohol (25:24:1 (v/v/v)) was added to the solution followed by gentle mixing. After centrifugation (5 min, 16,000 xg) proteins reside in the phenol and interphase, while the aqueous phase contains chromosomal DNA. The phenol-treatment was repeated on the aqueous phase to eliminate protein contaminations totally. The remaining phenol in the aqueous phase was removed by two washing steps with one volume of chloroform/isoamylalcohol (24:1 (v/v)). After thoroughly mixing the two phases, the solution was centrifuged for 5 min at 16,000 xg.

2.5.1.3. Ethanol precipitation

Chromosomal DNA was precipitated by mixing the aqueous phase with 2.5 volumes of ethanol (99 %, -20 °C; Green and Sambrook, 2012). Following an incubation o/N at - 20 °C, the DNA was pelleted by centrifugation (10 min, 14,000 xg). The pellet containing chromosomal DNA was washed with 70 % ethanol (99 % pure). This washing step was repeated with pure ethanol (99 %) and the DNA pellet was air dried. To resuspend dry

DNA, the pellet was solved in 100 µl ddH2O and incubated at 50 °C for 1 h for complete solubilisation.

2.5.2. Isolation of plasmid DNA

Plasmid DNA was isolated using the E.Z.N.A Plasmid Mini Kit I (Omega Bio-Tec, Inc., Norcross, USA) according to manufacturer´s instruction. For isolation of plasmids from bifidobacteria, the bacterial pellet as additionally washed twice in citrate sucrose buffer (0.5 M sucrose, 1 mM citric acid monohydrate, pH 5.8), the bacterial pellet were resuspended in solution I supplemented with 40 mg/ml lysozyme, and 10 µl mutanolysin (5 U/µl) and lysis reaction were performed at 37 °C for 1 h. Isolated plasmid DNA was checked by agarose gel electrophoresis (0.8 % (w/v), see section 2.5.4.). Plasmids, isolated from bifidobacteria, were additionally checked by PCR (see section 2.5.7.).

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Materials and Methods

2.5.3. DNA purification from agarose gels or PCR mixtures

For purification of DNA from agarose gels as well as purification of PCR products and digested plasmids, the NucleoSpin® Extract II kit (Macherey-Nagel GmbH & Co. KG, Düren, Germany) was used according to manufacturer’s instructions.

2.5.4. Agarose gel electrophoresis

DNA and RNA fragments were separated according their size by agarose gel electrophoresis (Sharp et al., 1973). Longer fragments move slower through the agarose matrix compared to shorter fragments. For preparation of agarose gels, 0.8 % (w/v) agarose was dissolved by boiling in 1X TAE buffer (40 mM Tris-HCl, 1 mM EDTA, 10 mM acetic acid, pH 8.0). After cooling down to approx. 50 °C melted agarose was poured into a gel stand (PerfectBlueTM Gelsystem Mini S/M, PEQLAB Biotechnologie GmbH, Erlangen, Germany) and fitted with a comb. After polymerization, the electrophoresis unit was filled with 1X TAE buffer until the gel was completely covered with buffer. The comb was removed directly before sample loading. Samples were mixed with loading dye (0.25 % (w/v) bromphenol blue in 40 % (v/v) glycerol) at a ratio of 5:1 and loaded into individual slots of the gel. GeneRulerTM 1 kb DNA ladder (Thermo Scientifc, Rockfort, USA) was used as molecular weight marker. Electrophoresis was run at 90 V for approx. 40 min. Nucleic acid bands were visualized by staining the gel for 10 min in ethidium bromide staining solution (1 µg/ml in H2O). To remove excess ethidium bromide gels were shortly destained in ddH2O. Agarose gels were exposed to UV light (302 nm) and the fluorescence of nucleic acids stained by intercalating ethidium bromide was imaged in a photodocumentation system (MWG Biotech GmbH, Ebersberg, Germany).

2.5.5. Quantification of dsDNA

Concentration of DNA was measured using the Qubit® dsDNA HS (High Sensitivity) Assay Kit (InvitrogenTM, Darmstadt, Germany) according to the recommendations of the manufacturer. The fluorescence of the dsDNA intercalating dye was measured with an Infinite® M200 multimode reader (Tecan Group Ltd., Männedorf, Switzerland) at an excitation wavelength of 480 nm and an emission wavelength at 530 nm.

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Materials and Methods

2.5.6. Oligonucleotides

Oligonucleotides were designed according to the recommendations of Green and Sambrook (2012) using the Clone Manager Suite 7 software. All oligonucleotides were purchased from Eurofins MWG Operon (Ebersberg, Germany). Table 15 in the Addendum lists all oligonucleotides used in this study.

2.5.7. Amplification of DNA by polymerase chain reaction

Specific DNA fragments were amplified by polymerase chain reaction (PCR; Saiki et al. 1985). During a PCR, a thermostable DNA polymerase is used to synthesize a second DNA strand complementary to a single stranded DNA template molecule in 5´ → 3´ direction. For this reaction, denaturation of dsDNA is required. Specific oligonucleotides are then able to anneal to their priming sequence on the template strand and form the start point for the synthesis of the new DNA strand. For cloning applications, Phusion® High- Fidelity (Thermo Scientific, Rockford, USA) polymerase with the proof-reading activity was used. Control experiments were performed using the non-proof-reading Genaxxon PCR system (Genaxxon, BioScience, Ulm, Germany). Table 4 shows the composition of typical PCR reaction mixtures. All PCRs were performed on a FlexCycler (Analytik Jena AG, Jena, Germany). PCR programs used for amplification are shown in Table 3. PCR products were analyzed by agarose gel electrophoresis (0.8 % (w/v)).

Table 3: PCR program used for amplification with the Phusion or Genaxxon system.

Cycle step Temperature Time Cycles Initial Denaturation 98 °C 30 s 1 Denaturation 98 °C 10 s Annealing 55-60 °C 30s 35 Extension 72 °C 30 s/kb Final extension 72 °C 10 min 1

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Materials and Methods

Table 4: Composition for PCR reaction mix using Phusion or Genaxxon system.

Phusion system Genaxxon system Component Final Volume Final concentration Volume concentration ddH2O - Ad to 20 µl - Ad to 25 µl

Reaction buffer 1X1 4.0 µl 1X2 2.5 µl dNTPs 400 µm each 0.4 µl 200 µm each 2.5 µl (10 mM3/2mM4)

Forward primer 0.2 µM 0.4 µl 0.2 µM 0.5 µl (10 pmol/µl)

Reverse primer 0.2 µM 0.4 µl 0.2 µM 0.5 µl (10 pmol/µl)

Template DNA 25-125 ng 0.5 µl 10-100 ng 1.0 µl

DMSO (100 %)5 3 % (v/v) 0.6 µl 5 % (v/v) 1.25 µl

DNA Polymerase 0.02 U/µl 0.2 µl 1 U/µl 1.0 µl 1 5X Phusion GC Buffer, recommended for GC-rich templates (Thermo Scientific, Rockford, USA) 2 10X Buffer containing 15 mM MgCl2 (Genaxxon, BioScience, Ulm, Germany) 3 dNTPs (10 mM) from Phusion® High-Fidelity Kit (Thermo Scientific, Rockford, USA) 4 dNTPs (2 mM) from Life Technologies, Darmstadt, Germany 5 DMSO (99.8 % pure, Carl Roth GmbH & Co., Karlsruhe, Germany)

2.5.7.1. Temperature gradient PCR

In order to optimize reaction conditions, a temperature gradient PCR was performed. This method enables the establishment of an annealing temperature optimal for each oligonucleotide mix and/or template. This prevents formation of undesired byproducts and allows robust synthesis of the desired target DNA sequence. The Phusion PCR reaction mixture shown in Table 3 and the temperature gradient PCR program shown in Table 5 were used to determine the optimal annealing temperature of difficult target DNA fragments. Temperature gradient PCRs were performed using the FlexCycler (Analytik Jena AG, Jena, Germany). To verify the PCR, products were analyzed by agarose gel electrophoresis (0.8 % (w/v), see section 2.5.4.).

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Materials and Methods

Table 5: PCR program with temperature gradient using the Phusion system.

Cycle step Temperature Time Cycles

Initial Denaturation 95 °C 30 s 1

Denaturation 95 °C 10 s

Annealing 45-75 °C 30 s 35

Extension 72 °C 30 s/kb

Final extension 72 °C 10 min 1

2.5.8. Screening colonies by PCR

After transformation of bacteria (see sections 2.6. and 2.7.), a number of clones were tested for presence of the correct plasmid/insert by colony PCR (Güssow and Clackson, 1989; Sandhu et al., 1989). To proof the ligation of the gene of interest into the vector, a forward primer binding in the vector region and a reversed primer binding in the insert sequence (see Table 15, Addendum) was designed. To perform colony PCR, parts of a bacterial colony were picked and resuspended into 15 µl ddH2O. For disruption of bacterial cells, the mixture was incubated at 95 °C for 10 min with shaking at 1,000 rpm. The lysate was then centrifuged for 5 min at 15,000 xg to pellet cell wall fragments. 1 µl of the supernatant was used as template in a total volume of 12.5 µl using Genaxxon Taq polymerase (Genaxxon, BioScience, Ulm, Germany). For colony PCR, half volume of the Genaxxon system listed in Table 4 was used. The PCR program used in this method is shown in Table 6. PCR products were analyzed by agarose gel electrophoresis (0.8 % (v/v), see section 2.5.4.).

Table 6: PCR grogram for colony PCR.

Cycle step Temperature Time Cycles Initial Denaturation 95 °C 2 min 1 Denaturation 95 °C 20 s Annealing 56 °C 30 s 25 Extension 72 °C 1 min Final extension 72 °C 7 min 1

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Materials and Methods

2.5.9. DNA digestion with restriction enzymes

Restriction endonucleases (see Table 14, Addendum) were used to create defined sticky ends of dsDNA products (Green and Sambrook, 2012). DNA digestions were performed according to the recommendations of the manufacturer or each enzyme or enzyme mixes individually. A typical restriction digest contained 0.5 – 1 µg DNA in a total volume of

5 µl, 1 µl restriction enzyme, 2 µl of the recommended buffer and 12 µl ddH2O to a give final volume of 20 µl. After incubation in a Thermomixer comfort (Eppendorf AG, Hamburg, Germany) for 2 h at restriction enzyme´s optimal reaction temperature (all enzymes in this study 37 °C). Digested DNA was verified by agarose gel electrophoresis (0.8 % (w/v), see section 2.5.4.).

2.5.10. Klenow fill-in of DNA ends with 5`-overhang

The Klenow enzyme exhibits 5`-3`polymerase activity and is thus able to generate blunt DNA ends by fill-in of 5`-overhangs or under specific conditions by degradation of 3`- overhangs. The Klenow enzyme is the larger of two protein fragments of DNA polymerase I of E. coli obtained after cleavage with the protease subtilisin (Klenow and Henningsen, 1970). While it retains 5`-3`polymerase and 3`-5`exonuclease activity it lacks 5`-3`exonuclease activity of the DNA polymerase I holoenzyme.

Klenow fill-in of DNA ends was performed according to manufacturer’s instructions (Life Technologies, Darmstadt, Germany). Briefly, 40 µl of purified DNA (approx. 1 µg), 6 µl of 10x Klenow buffer (500 mM Tris-HCl (pH 8.0 at 25 °C), 50 mM MgCl2, 10 mM DTT) 1.5 µl dNTP mix (2 mM), 0.6 µl Klenow Fragment (10 U/µl) and 11.9 µl nuclease-free water to give a total volume of 60 µl. Reactions were incubated on a Thermomixer comfort for 10 min at 37 °C. Inactivation of Klenow enzyme was performed for 10 min at 75 °C.

2.5.11. Ligation

For plasmid construction, a target DNA fragment (insert) was inserted into the vector with compatible DNA termini using T4 DNA (Thermo Fisher Scientific Inc., Rockfort, USA) according to manufacturer’s instructions. During a ligation, the formation of phosphodiester bonds between juxtaposed 5`-phosphate and 3`-hydroxyl termini of duplex

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Materials and Methods

DNA were catalyzed by the T4 DNA Ligase enzyme (Lehman, 1974). Linear vector and insert DNA were mixed in a molar ratio of 1:5. For ligation, 3.0 µl vector DNA and 5.0 µl insert DNA were added to 2.0 µl T4 DNA Ligase buffer (10X) and 0.5 µl T4 DNA Ligase

(1 U). To give a total volume of 20 µl, 9.5 µl nuclease-free H2O were added. The reaction mixture was incubated o/N at RT. All constructed plasmids were listed in Table 2.

2.5.12. Sequencing of plasmid constructs

The sequence of all inserts of generated plasmids were sent for Sanger sequencing by a commercial service provider (GATC Biotech AG, Köln, Germany). The obtained sequences were analyzed using the alignment function of Clone Manager Suite 7 software. Only mutation free constructs were used further.

2.5.13. RNA extraction

RNA was extracted as described previously (Westermann et al., 2012a). In general, 10 ml MRSc medium were inoculated with B. bifidum S17 and cultures were grown under anaerobic conditions o/N. For isolation of RNA, the bacterial culture were mixed with 40 ml of Quenching buffer (60 % methanol, 66.7 mM HEPES, pH 6.5). The mixture was centrifuged at 4,000 xg for 20 min and 4 °C. The bacterial pellet was then resuspended in

200 µl ice cold ddH2O. After the transfer of the suspension to a cryo-tube filled with 250 mg of glass beads (0.1 mm diameter), 400 µl of acidic phenol, 100 µl of chloroform/isoamylalcohol (24:1 (v/v)), 30 µl of 10 % (w/v) SDS and 30 µl of sodium acetate (3 M, pH 5.2) were added. Bacterial cells were disrupted in a Ribolyser (Precellys® 24, Peqlab Biotechnologie GmbH, Erlangen, Germany) during 3 cycles of 40 s and full speed, with cooling of the samples on ice for 2 min between cycles. By centrifugation of the samples at 18,000 xg for 15 min, cell debris, glass beads and phenol/chloroform were removed. The RNA containing aqueous phase was transferred to a fresh Eppendorf tube® and mixed with 1.5 volume of 100 % ethanol (99 % pure). The mixture was stored o/N at - 20 °C. The precipitated RNA was pelleted by centrifugation at 15,000 xg for 30 min at 4 °C. After washing the pellet once with 1 ml of 70 % (v/v) ice cold ethanol (99 % pure) RNA was dried under airflow in a cleanbench for approx. 20 min. The dry pellet was then resuspended in 200 µl of RNase-free water. Residual DNA was removed by DNaseI

38

Materials and Methods treatment. For this purpose, 30 µl of 10X reaction buffer, 20 µl DNaseI (1U/µl, RNase- free) and 44 µl H2O were added to 200 µl RNA sample and incubated at 37 °C in a Thermomixer comfort for 30 min. Further 10 µl DNaseI were added followed by incubation for another 15 min at 37 °C. DNaseI was then inactivated by adding 30 µl EDTA (25 mM) and incubation at 65 °C for 10 min. DNaseI treatment was repeated until no DNA contamination could be detected by a standard PCR targeting the 16S rRNA gene. DNA-free RNA samples were purified using the RNeasy Mini Kit (Quiagen GmbH, Hilden, Germany) according to the recommendations of the manufacturer. Elution of RNA from columns of the kit was performed by applying two times 50 µl RNase-free H2O to the columns with on-column incubation for 10 min before centrifugation. RNA concentration was quantified by measuring the absorbance at 260 nm using the Infinite® M200 multimode reader (Tecan Group Ltd., Männedorf, Switzerland). Using the Lambert-Beer- law (formula 2) RNA concentrations of the samples were calculated.

퐴260 = 휀 ∙ 푐 ∙ 푑

Formula 2: Lambert-Beer-law, for calculation of the RNA concentration. With A260: absorbance at 260 nm, ε: extinction coefficient (for RNA 40 mg x cm x l-1), c: RNA concentration [µg/ml], d: distance (i.e. path length: 0.5 mm).

2.5.14. Reverse Transcription PCR

Reverse transcription PCR (RT-PCR) was performed according a standard protocol as described previously (Westermann et al., 2012a). The expressions of genes of interest were assessed by mRNA levels using RT-PCR. For this purpose, cDNA were synthesized using RevertAidTM Premium First Strand cDNA Synthesis Kit (Thermo Fisher Scientific Inc., Rockfort, USA) according to manufacturer’s instructions. This kit is based on the use of an engineered version of the Moloney Murine Leukemia Virus (M-MuLV) reverse transcriptase, which is a RNA- and DNA-dependent DNA polymerase deficient in RNaseH activity. For cDNA synthesis 1 µg of total RNA, 1 µl random hexamer primers (0.2 µg/µl) and 1 µl dNTP mix (10 mM) were mixed and RNase-free water was added to a final volume of 15 µl. The reaction mixture was mixed gently, centrifuge briefly, and incubated at 65 °C for 5 min. This step is required to denaturate potential secondary structures of GC

39

Materials and Methods rich RNA templates. After incubation, the mixture was chilled on ice, spun down briefly and placed back on ice. Then, 4 µl of RT buffer (5X) and 1 µl RevertAidTM Enzyme Mix were added and the RT reaction was performed with the following incubations steps (Table 7):

Table 7: PCR program used for sqRT-PCR.

Step Temperature Time

Annealing 25 °C 10 min

Reverse Transcription 60 °C 30 min

Inactivation 85 °C 5 sec

Storage of cDNA occurred at -20 °C for a maximum of one week until use in target specific PCR. For semi-quantification, grey-scale images of agarose gels were analyzed using ImageJ 1.48v software. In principal, quantification was done on the base of individual pixels of the agarose gel image, which correlate to the amount of PCR product. Individual pixels are weighted according to the intensity level on a black-to-white scale. Signal intensities were integrated over a defined area of the agarose gel image around the band with background substraction.

2.6. Transformation on E. coli

2.6.1. Preparation of electrocompetent E. coli cells

Electrocompetent E. coli cells are able to take up foreign DNA upon electroporation. Electrocompetent E. coli cells were prepared according to the protocol of Sheng et al. (1995). 500 ml LB media were inoculated with 500 µl of an o/N culture of the E. coli strain of choice and cultures were grown aerobically at 37 °C with agitation to midexponential growth phase of the culture, i.e. an OD600 of 0.5. The culture was then chilled on ice for 15 min before harvesting bacteria at 4,000 xg for 15 min at 4 °C. Bacteria were washed twice in 200 ml ice cold ddH2O followed by two washing steps in 25 ml ice cold 10 % (v/v) glycerol (99 % pure) at 4,000 xg for 15 min and 4 °C. Then, the bacterial

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Materials and Methods cells were centrifuged at 4,000 xg for 15 min at 4 °C and the bacterial pellet was resuspended in 1,000 µl ice cold 10 % (v/v) glycerol (99 % pure). Aliquots of 50 µl were snap-frozen in liquid nitrogen and stored at -80 °C.

2.6.2. Transformation of electrocompetent E. coli

Transformation of E. coli cells was performed as described in Dower et al. (1988) with slight modifications. An aliquot of 50 µl electrocompetent cells was thawed on ice and then mixed with 5 µl plasmid DNA in ice cold cuvettes (1 mm gap, Bio-Rad, Munich, Germany). After 5 min incubation on ice, bacteria were electroporated with a single pulse at 25 µF, 200 ohms, and 2 kV cm-1 in a Gene Pulser® II (Bio-Rad, Munich, Germany). Immediately after the pulse, 1 ml pre-warmed (37 °C) SOC medium (5 g/l yeast extract, 2 g/l tryptone, 10 mM sodium chloride, 2.5 mM potassium chloride, 10 mM magnesium chloride, 10 mM magnesium sulfate) was added and bacteria were incubated at 37 °C for 1 h for regeneration. Transformed bacteria were then plated on LB-agar containing the appropriate antibiotic and incubated o/N at 37 °C.

2.6.3. Plasmid artificial modification

The plasmid artificial modification (PAM) is a method to improve bacterial transformation efficiency. This method aims at overcoming restriction-modification systems that protect target bacteria against foreign DNA (Makarova et al., 2013). For transformation of plasmids into B. bifidum S17, the PAM system was used. For this purpose, the methyltransferase negative E. coli ET12567 strain was used. In a previous study

(Brancaccio, 2013b), p16S_ParaB_bbif0710 carrying the gene encoding for one of the methyltransferases of B. bifidum S17 was integrated into the genome of E. coli ET12567.

The resulting strain ET12567::p16S_ParaB_bbif0710 expresses the methyltransferase of B. bifidum S17 following induction with arabinose. Thus, plasmids transformed into E. coli

ET12567::p16S_ParaB_bbif0710 for amplification can be methylated in a manner specific for B. bifidum S17. After transformation of these plasmids into B. bifidum S17, the DNA is recognized as host DNA and prevented for degradation through restriction endonucleases thereby increasing transformation efficiency (Brancaccio, 2013b).

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Materials and Methods

2.7. Transformation of Bifidobacterium sp. strains

2.7.1. Preparation of electrocompetent bifidobacteria

Bifidobacteria were transformed as described previously (Kim et al., 2010) with minor modifications. For preparation of electrocompetent bifidobacteria, 10 ml MRSc supplemented with 0.2 M sucrose were inoculated with 200 µl of a fresh o/N culture in MRSc and incubated anaerobically at 37 °C for approx. 8 h. 100 µl of this culture were used to inoculate 10 ml MRSc supplemented with 0.2 M sucrose and incubated under anaerobic conditions at 37 °C o/N. 50 ml modified Rogosa (composition see table 8) supplemented with 16 % (w/v) fructo-oligosaccharide (Actilight®; in case for B. longum E18 1 % (w/v) glucose) were inoculated with 2 ml of the o/N culture and incubated anaerobically at 37 °C until bacteria reached an OD600 of 0.6 - 0.8. At this stage, bacteria were harvested by centrifugation at 4,500 rpm for 10 min followed by three washing steps in 30 ml ice cold wash buffer (1 mM ammonium citrate (pH 6), 0.5 M sucrose). After the final washing step, the bacterial cell pellet was resuspended in adequate volume of ice cold wash buffer and immediately used for electroporation.

2.7.2. Transformation of electrocompetent bifidobacteria

For transformation, 5 µl plasmid DNA were mixed with 50 µl prepared competent bifidobacteria cells in ice cold cuvettes (1 mm gap, Bio-Rad, Munich, Germany) as described (Kim et al., 2010). Transformation was carried out with a single pulse of 12.5 kV cm-1, 25 µF and 200 ohms in Gene Pulser® II (Bio-Rad, Munich, Germany). Following electroporation, bacteria were resuspended in 800 µl pre-warmed (37 °C) RCM broth supplemented with 0.05 % (w/v) L-cysteine hydrochloride and 1 % (w/v) fructo- oligosaccharide (Actilight®, Syral – Beghin Meiji, Marckolsheim, France) and were incubated anaerobically at 37 °C for 2.5 h. Then, transformed bifidobacteria were plated on MRSc agar containing the appropriate antibiotic and incubated under anaerobic conditions at 37 °C for 2 days.

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Materials and Methods

Table 8: Composition of the modified Rogosa medium.

Components Amount (g/l)

Trypticase peptone 10

Yeast extract 2.5

Tryptose 3

Dipotassium phosphate dibasic (K2HPO4) 3

Potassium phosphate monobasic (KH2PO4) 3

Ammonium acetate (C2H4O2 NH4) 1.86

Ammonium chloride (NH4Cl) 0.44

Pyruvic acid (CH3COCOOH) 0.2

L-Cysteine hydrochloride monohydrate 0.5

Tween 80 1 ml

Magnesium Sulfate Heptahydrate (MgSO4 x 7 H2O) 0.0575

Manganese(II) Sulfate Hydrate (MnSO4 x H2O) 0.015

Iron(II) Sulfate Heptahydrate (FeSO4 x 7 H2O) 0.0034 ddH2O Ad to 1 l

2.8. Working with proteins

2.8.1. Preparation of bacterial cell fractions

Bacterial cell fractions were prepared as described previously (Kotani et al., 1975) with minor changes. An o/N culture of 100 ml was harvested by centrifugation at 4,000 xg for 20 min. Supernatants containing secreted proteins were stored at -20 °C until further investigations. For separation of the cell envelope and cytoplasm, the volume of the bacterial pellet was estimated and the pellet was resuspended in three volumes of 50 mM Tris-HCl (pH 7.6). Bacterial cells were then lysed in a French Press high pressure cell disrupter (SML Aminco, SelectScience Ltd., Corston, Bath, USA) at a pressure of 500 psi. As control 1 ml of the crude cell extract was taken and stored at -20 °C. The cell lysate was centrifuged at 45,000 xg for 1.5 h at 4 °C in a CU-L8-60M ultracentrifuge (rotor TFT 65.13, Beckman® Coulter GmbH, Krefeld, Germany). The resulting supernatant containing

43

Materials and Methods the cytoplasmic fraction were stored at -20 °C. The bacterial cell pellet representing the cell envelope fraction was resuspend in 700 µl Tris-HCl (pH 7.6) and containing cell envelope proteins were dissolved by three cycles of ultrasound (1 min) in a Sonorex Super RK 255 (Bandelin electronic, Berlin, Germany) with cooling of samples on ice for 1 min between cycles. To avoid protein aggregations 0.02 % (v/v) Triton X-100 (Sigma Aldrich Chemie GmbH, Steinheim, Germany) was added. The cell envelope fraction was stored o/N at 4 °C. The protein concentrations in the different fractions were quantified as described below (see section 3.2.).

2.8.2. Quantification of proteins

The quantification of protein concentrations was performed using the BCATM Protein Assay Kit (Pierce Biotechnology, Thermo Scientific, Rockford, USA) according to manufacturer’s instructions. This assay is based on the selective colorimetric detection of cuprous cations by bicinchoninic acid (BCA). Cuprous cations are the result of the chelat reaction of copper with proteins in an alkaline environment, commonly known as biuret reaction. Two molecules BCA react with one cuprous ion and generate product with an intense purple color. The BCA/copper complex has a characteristic absorbance at 562 nm, which is directly proportional to increasing protein concentrations. For calculation of the concentration of protein samples, a standard curve of different concentrations of bovine serum albumin (BSA) was used. All experiments were performed in duplicate. The colorimetric detection were done using an Infinite® M200 multimode reader (Tecan Group Ltd., Männedorf, Switzerland).

2.8.3. Sodium dodecyl sulfate polyacrylamide gel electrophoresis

Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) is a method for separating proteins based on their ability to move within an electrical field depending on their molecular weights. SDS-PAGE was performed according a standard protocol (Green and Sambrook, 2012). The anionic SDS detergent denatures secondary and tertiary protein structures, thus the proteins are present in their linear form as polypeptide chains. Furthermore, SDS coats the proteins and their innate electrical charge, resulting in a negative electrical charge proportional to their molecular mass. The gel system used is

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Materials and Methods based on tris-glycine gels which consist of a stacking gel and the resolving gel component. The stacking gel has large pores and a lower pH than the electrophoresis buffer (see Table 9). This helps to focus the proteins into sharp bands at the beginning of the separation. Then, polypeptides migrate through the small pores of the resolving gel and the speed of migration depends on their molecular mass. The composition of a 12.5 % polyacrylamide gel is listed in Table 9 below.

Table 9: Composition of stacking and separation gels for SDS-PAGE.

Amount Components Stacking gel (5 %) Resolving gel (12.5 %) ddH2O 2,15 ml 0.9 ml 40 % Rotiphorese®1 1.5 ml 0.188 ml 1.5 M Tris HCl2 1.25 ml (pH 8.8) 0.188 ml (pH 6.8) 10 % (w/v) SDS3 0.05 ml 0.015 ml 10 % (w/v) APS3 0.05 ml 0.015 ml TEMED3 0.002 ml 0.0015 ml 1 Rotiphorese®, BioRad Laboratories GmbH, Munich, Germany 2 Sigma Aldrich Chemie GmbH, Steinheim, Germany 3 Carl Roth GmbH & Co., Karlsruhe, Germany

SDS-PAGE was performed using the Mini-PROTEAN® Tetra System (BioRad Laboratories GmbH, Munich, Germany) in 1X SDS electrophoresis buffer (25 mM Tris, 250 mM glycine, 0.1 % (w/v) SDS). Before loading the samples, 25 µg of protein were mixed with 20 µl of 1X Laemmli buffer (50 mM Tris-HCl, pH 6.8, 2 % (w/v) SDS, 0.1 % (v/v) bromphenol blue, 10 % (v/v) glycerol, 10 % (v/v) β-mercaptoethanol) and denatured at 95 °C for 10 min (Laemmli, 1970). Electrophoresis was run at 25 mA per gel for approx. 1 h. PageRulerTM Prestained Protein Ladder (Thermo Scientific, Rockfort, USA) was used as molecular weight marker.

2.8.4. Staining of SDS gels

Protein bands on SDS gels were visualized using InstantBlue (Expedeon, Cambridgeshire, United Kingdom) solution. InstantBlue is a Coomassie based stain, which allows the ultrafast detection of low amounts of proteins (5 ng) in a single step. The SDS gel was

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Materials and Methods stained in 30 ml InstantBlue for approx. 30 min until the protein bands were visible. Excess staining solution was removed by washing the gel in 50 ml H2O for approx. 10 min.

2.8.5. Western Blot

Western Blot is the transfer of proteins resolved by SDS-PAGE onto a support material and the subsequent specific identification of proteins using antibodies. In this study, all proteins analyzed by Western Blot were labeled with a His6-tag and were detected using a monoclonal anti-His6 antibody. This specific antibody was then detected with a secondary antibody conjugated with horseradish peroxidase (HRP), an enzyme which results in chemiluminescence by reaction with the Pierce ECL Western Blotting Substrate (Thermo Scientific, Rockfort, USA).

To perform Western Blots a standard procedure was carried out (Green and Sambrook, 20112). After separating the proteins as described above, the polyacrylamide gel was incubated for 5 min in blotting buffer (48 mM Tris-base, 39 mM glycine, 20 % (v/v) methanol; pH 9.2). Prior to transfer, a positively charged nitrocellulose membrane (Macherey-Nagel GmbH & Co. KG, Düren, Germany) was incubated for 5 min in blotting buffer. To assemble the blotting apparatus (Trans-Blot® TurboTM Transfer System, BioRad Laboratories GmbH, Munich, Germany) two layers of Whatman® gel blotting paper (7x10 cm, grade GB003, GE Healthcare Europe GmbH, Freiburg, Germany) saturated with blotting buffer were placed on the bottom of the cassette. After that the nitrocellulose membrane was laid onto the Whatman® gel blotting paper stack. Then the SDS-gel was carefully placed on the membrane and covered with three further layers of Whatman® gel blotting paper soaked in blotting buffer. Air bubbles between the layers were removed using a roll. The cassette was then closed and locked with the lid. Blotting was performed for 30 min at 25 V and 1 A. Successful protein transfer was confirmed by presence of the prestained bands of the molecular weight marker on the nitrocellulose membrane and their absence in the polyacrylamide gel.

After the blot, the membrane was washed in TBS-T buffer (50 mM Tris-base, 150 mM NaCl, 0.1 % (v/v) Tween 20, pH 7.6) for 5 min. Unspecific binding sites were blocked by incubating the membrane into TBS-T containing 5 % (w/v) BSA for 1 h at RT. Following blocking, the membrane was incubated with the primary mouse ant-His6 IgG1 monoclonal antibody (Catalog # 11922416001, dilution 1:400 in TBS-T + 5 % (w/v) BSA) o/N at 4 °C.

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Materials and Methods

After three washing steps with TBS-T for 10 min, the membrane was incubated with the secondary rabbit anti-mouse polyclonal antibody coupled with HRP (Catalog # P0260, dilution 1:1000 in TBS-T + 5 % (w/v) BSA) for 1 h at RT. The membrane was washed again three times for 10 min in TBS-T at RT. Pierce ECL Western Blotting Substrate was used according to manufacturer’s instructions as substrate for horseradish peroxidase. The resulting chemiluminescence was detected on X-ray films (Agfa HealthCare GmbH, Bonn, Germany) with an adequate exposure time.

2.9. Microscopy

2.9.1. Ruthenium red staining

Polysaccharide capsules of bifidobacteria were visualized by lysine based ruthenium red staining method (Fassel et al, 1992) and subsequent electron microscopy. Bifidobacteria were grown to stationary growth phase and harvested at 4,000 xg for 10 min at RT. Bacterial pellet was then resuspended in pre-fixation solution (0.075 % ruthenium red, 75 mM lysine, 2.5 % (v/v) glutaraldehyde, solved in 0.1 M cacodylate buffer, pH 7.0) and incubated for 20 min at RT. After centrifugation (4,000 xg, 10 min, RT) bacteria were fixed in fixation solution (0.075 % ruthenium red, 2.5 % (v/v) glutaraldehyde in 0.1 M cacodylate buffer, pH 7.0) for 2 h. After washing three times with 0.1 M cacodylat buffer, samples were dehydrated by immersion and incubation in increasing concentrations of ethanol (10 %, 25 %, 50 %, 70 %, 95 %, and 100 % (v/v) in H2O).

2.9.2. Sample embedding and ultrathin sectioning

Fixed and stained samples were embedded and sectioned as described elsewhere (Ayache et al., 2010) with slight modifications. Shortly, after sample dehydration, embedding started with 33 % (v/v) Epon (Fluka AG, Buchs, Switzerland) for 15 min, then 50 % (v/v) Epon for 30 min, 66 % (v/v) Epon for 1 h and finally 100 % Epon o/N. Dilutions of Epon were made in 1-propanol. The next day, fresh Epon was added and polymerization was performed at 60 °C for 48 h. For TEM analysis, samples were prepared by ultrathin sectioning (approx. 60 to 80 nm) on a Leica Ultracut UCT ultramicrotome using a diamond

47

Materials and Methods

knife. Ultrathin sections were collected in a hutch filled with ddH2O, transferred to a bare copper grid and air dried.

2.9.3. Transmission electron microscopy

For transmission electron microscopy (TEM), ultrathin sections were accessory contrasted with lead citrate for one minute (Reynolds, 1963). Afterwards, bacteria were analyzed in an EM10 transmission electron microscope (Carl Zeiss AG, Oberkochen, Germany). Images were taken at an acceleration voltage of 80 kV.

2.9.4. Scanning electron microscopy

For scanning electron microscopy (SEM), ruthenium red stained cells were critical point dried using CO2 with the Critical Point Dryer COD 030 (Bal-Tec, Principality of Lichtenstein). After drying, the samples were rotary coated with 3 nm of platinum-carbon by electron beam evaporation using Baf 300 device (Bal-Tec, Principality of Lichtenstein). The specimens were then visualized using a Hitachi S-5200 scanning electron microscopy (Hitachi, Tokyo, Japan) at an accelerating voltage of 4 kV.

2.9.5. Fixing and staining of cells for fluorescence microscopy

The procedure of sample preparation and fluorescence microscopy was described previously (Grimm et al., 2014) with slight variations. For fixation of Caco-2 cells with adherent fluorescent bifidobacteria, cells were incubated with 4 % (w/v) paraformaldehyde in PBS for 30 min at RT. After three washes with PBS, cell nuclei were stained with Hoechst diluted 1:10,000 in PBS for 10 min at RT. Cells were washed with PBS and the surface was covered with Mowiol®.

2.9.6. Fluorescence microscopy

Fluorescence microscopy was performed on Zeiss Axio Observer.Z1 microscope (Carl Zeiss AG, Oberkochen, Germany) using a 40X objective (Grimm et al., 2014). The appropriate filter sets to detect fluorescence of Hoechst (excitation at 365 ± 10 nm, emission at 445 ± 50 nm), GFP (excitation at 470 ± 40 nm, emission at 525 ± 50 nm) and

48

Materials and Methods mCherry (excitation at 545 ± 30 nm, emission at 610 ± 75 nm) were used. Images were obtained with the ZEN 2012 software (Carl Zeiss AG, Oberkochen, Germany). For quantification, images were analyzed using ImageJ 1.48v software. In principal, quantification is based on individual red and green pixels of the fluorescence image independent of the fluorescent intensity. Individual pixels are weighted according to the amount on a black-to-white scale with background substraction.

2.10. Bioinformatic analysis

Bioinformatic analysis was performed as described previously (Westermann et al., 2012a). Proteins with domains known or suspected to be involved in microbe-host interactions were identified using a previously published list of domains (Kankainen et al., 2009). Domains were obtained from the publically available protein families (Pfam; Finn et al., 2014) and the institute for genomic research (TIGR) databases. Using the HMMER package (Wheeler and Eddy, 2013), deduced sequences of all open reading frames identified in the genome of B. bifidum S17 were searched for these domains (Bateman et al., 1999). Signal peptides were identified by using the SignalP 3.1 and SignalP 4.0 software (Bendtsen et al., 2004; Petersen et al., 2011). The subcellular localization of proteins was predicted using pSORTb v2.0 and pSORTb v3.0.2 (Gardy et al., 2005) and transmembrane helices were identified with Transmembrane prediction using Hidden Markov Model (TMHMM; Krogh et al., 2001). Cell-wall associated domains and LPXTG motifs were predicted with Hidden Markov Model (HMM) from Pfam (Sonnhammer et al., 1998; Finn et al., 2014). Protein domain structures were analyzed in more detail using InterProScan (Zdobnov and Apweiler, 2001).

2.11. Statistical analysis

Experiments were performed in technical and biological triplicates as indicated. The results are shown as mean ± standard deviation of biological replicates. Statistical analysis was performed by ANOVA or pair-wise comparisons by Student t test as indicated in the figure captions. Levels of statistical significance are indicated as follows: * p ≤ 0.05; ** p ≤ 0.01; *** p ≤ 0.001; **** p ≤ 0.0001.

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Results

3. Results

The first direct contact between bacteria and their environment is usually mediated by structures decorating the cell surface of bacteria. There are different structures known to be present at the cell wall and potentially involved in this process. For a range of bacteria, proteins, teichoic acids and polysaccharides are known to interact with host structures (Lebeer et al., 2010). For bifidobabacteria, cell surface associated EPS (Fanning et al., 2012a), type IV tight adherence, sortase dependent pili (Turroni et al., 2013, Foroni et al., 2011), and other surface proteins (Sànchez et al., 2010) were shown to play a role in host colonization and attachment to host tissues. Interestingly, many of the identified bacterial adhesins are so called moonlighting proteins. Lipoprotein BopA was shown by us and others to play a role in adhesion process to IECs (Guglielmetti et al., 2008; Gleinser et al., 2012). Moreover, bifidobacterial chaperon DnaK, glutamine synthetase, enolase, bile salt hydrolase, and phosphoglycerate mutase are further examples for moonlighting proteins with function in bacterial adhesion (Candela et al., 2007; Candela et al., 2009; Candela et al., 2010). However, the present study focused on specific surface structures of B. bifidum S17 potentially involved in host colonization. Bioinformatic analysis (section 3.1.) allowed the identification of different structures potentially involved in this process. The evidence (section 3.2.) of some of these structures was investigated and some functional characterizations (section 3.3) were done.

3.1. Bioinformatic analysis

3.1.1. EPS

Many bacteria including bifidobacteria, were shown to produce EPS, which are carbohydrate polymers present as an extracellular layer on the surface of microorganisms (Suresh Kumar et al., 2007; Ruas-Madiedo et al., 2009b). The EPS of probiotic bacteria are of special interest, because it has been shown that EPS has the capability to modulate the host immune response and help bacteria to overcome gastrointestinal challenges (Denou et al., 2008; Lòpez et al., 2011; Lebeer et al., 2012; Fanning et al., 2012a; Nikolic et al., 2012).

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Results

Phenotypic observation that could be linked to differences in EPS structure and composition is that, in liquid cultures, B. longum E18 grows equally distributed throughout the culture and is difficult to precipitate by centrifugation. By contrast, B. bifidum S17 grows at the bottom of the growth vessel and forms tightly packed pellets after centrifugation. For this reason, B. longum E18 was used as control. Two putative EPS biosynthesis gene cluster (blong_0398-0417 and blong_2008-2021) were found in the genome of B. longum E18 (Zhurina et al., 2013). In silico analysis of a previous work (Oehlke, 2013) revealed the presence of two gene clusters (bbif_0049-0063 and bbif_0393- 0396) encoding the EPS assembly machinery in B. bifidum S17. Table 10 shows the identified genes with the corresponding predicted function.

Table 10: Predicted genes of B. bifidum S17 involved in EPS production.

Gene bbif_ Predicted function 0049 glycosyltransferase 0050 hypothetical protein 0051 dTDP-glucose 4,6-dehydratase 0052 hypothetical protein 0053 dTDP-4-dehydrorhamnose 3,5-epimerase/reductase

0054 Transposase 0055 hypothetical protein 0056 Transposase 0057 Transposase 0058 Glucose-1-phosphat thymidyl

EPS gene cluster I cluster EPS gene 0059 sugar ABC transporter permease 0060 sugar ABC transporter ATP-binding protein 0061 hypothetical protein 0062 lipopolysaccharide biosynthesis protein 0063 glycosyltransferase

0393 UDP Galactosephosphotransferase 0394 transporter protein 0395 hypothetical protein

cluster II cluster EPS gene EPS gene 0396 hypothetical protein

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3.1.2. Pili

In silico analysis of the complete genome sequence of B. bifidum S17 led to identification of four gene clusters potentially encoding for pili (Westermann, 2012b). These clusters include one for putative type IV Tad (tight adherence) pili (Figure 4) and three for potential sortase-dependent pili (Figure 5).

The genes of the tad cluster are typical organized in two adjacent operons and a separately located gene and show good homology to the tad genes of B. breve UCC2003 (Westermann, 2012b). One operon contains the tadZABC genes encoding for the tad pilus assembly complex (Figure 4). A second adjacent operon harbors the genes encoding the structural pilus subunits flp and tadE. flp encodes the precursor of the major pilin and the tadE gene product is the minor pilin subunit. The corresponding peptidase TadV is encoded on a separate locus.

Figure 4: Genetic organization of the putative tad pilus cluster identified in the genome of B. bifidum S17. Genes and the putative function of their products are indicated by colored arrows. Blue bars indicate the amplified region in RT-PCR. aNote (p. 114).

The gene clusters for potential sortase-dependent pili were designated fim1 (bbif_0301- 0303), fim2 (bbif_1648-1650), and fim3 (bbif_1760-1762) and are shown in Figure 5. Sortase-dependent pilus clusters are typically organized as operons consisting of a gene encoding the major pilin subunit, a gene encoding a minor pilin subunit and a dedicated sortase.

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Figure 5: Genetic organization of the potential sortase-dependent pili cluster fim1 (bbif_0301- 0303), fim2 (bbif_1648-1650), and fim3 (bbif_1760-1762) of B. bifidum S17. Genes are indicated as arrows: sortase genes (light blue), genes for major pilin subunits (yellow), and minor pilin subunits (black). Red triangles indicate LPXTG motifs detected in silico. Blue bars indicate the amplified region in RT-PCR. aNote (p. 114).

3.1.3. Cell surface proteins

Adhesion to cultured intestinal epithelial cell in vitro is partially mediated by the surface lipoprotein BopA (Gleinser et al., 2012). However, these experiments indicated that there are additional mechanisms that might be involved in adhesion of B. bifidum S17 to IECs. To prognosticate proteins potentially involved in adhesion, the predicted proteome of B. bifidum S17 was screened in silico (Westermann et al., 2012a) for proteins carrying domains known or predicted to be involved in the adhesion to host tissue using InterProScan, Pfam (Protein families), and TIGR (The Institute for Genomic Research) databases as described in a previous publication (Kankainen et al., 2009). Additionally, the theoretical proteome was screened for all proteins containing signal sequences (SignalP, pSORT) targeting the cell envelope such as sortase recognition motifs. Transmembrane spanning regions were identified through TMHMM (Transmembrane prediction using Hidden Markov Model) analysis. Sugar binding proteins with predicted enzymatic function were removed from the list of potential proteins adhesins. This yielded a list of nine potential adhesion proteins besides BopA. Identified proteins with adhesion associated domains were designated as putative adhesins. The sizes of the putative adhesins range between 1992 amino acids (aa) (BBIF_0450) to only 175 aa (BBIF_0295; Figure 6). The

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Results existing analysis (Westermann et al., 2012a) was updated using current versions of the software packages (Tables 11 and 12). This revealed that the initial analysis with SignalP 3.0 predicted signal peptides for BBIF_0450 and BBIF_0665 could not be confirmed by the new analysis (SiganlP 4.1).

Furthermore, different cleavage sites were predicted for two putative adhesins (BBIF_1426, BBIF_1538). Only for BBIF_1769 the result of the previous in silico analysis of a predicted signal peptide was confirmed. Also, most of the predictions of subcellular localization with pSORTb v3.02 did not confirm the earlier results obtained with pSORT v2.0. Only BBIF_0665 was predicted as a membrane protein by both versions of pSORT. BBIF_0295 (W-score 9.97, i.e. 99.7 % probability for cell wall localization) and BBIF_0450 (W-score 8.90) were predicted to be cell wall proteins and BBIF_1426 is predicted to be localized extracellular (E-score 9.73). The subcellular localization of BBIF_0017, BBIF_0540, BBIF_1538, and BBIF_1769 was not predicted with any reasonable probability. BBIF_0017 has neither a predicted signal peptide nor a transmembrane spanning region. Additionally, the predicted LPXTG motif cannot be confirmed with newer software versions. Interestingly, BBIF_0567 was predicted as cytoplasmic protein with a C-score of 7.5 (i.e. 75 % probability for localization in the cytoplasm) and no signal peptide was identified. For this reason, BBIF_0017 and BBIF_0567 were removed from the list of single adhesins.

Figure 6 shows the remaining putative adhesins with their according domains. One protein containing a collagen associated domain, one protein with fibronectin associated domain, one protein with Ig-like domain, one protein with lectin associated domain, one protein with von Willebrand factor type A domain, one protein with G5 domain, and one protein with NlpC/P60 domain were identified in theoretical proteome of B. bifidum S17.

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Table 11: Potential surface adhesin proteins with predicted signal peptide with SignalP 3.0 and SignalP 4.1.

SignalP 3.0 SignalP 4.1 predicted predicted BBIF_ Signal Signal D-score1 cleavage D-score1 cleavage peptide peptide site2 site2

0017 0.45 No 0.231 No

0295 0.45 No 0.215 No

aa 35-36: 0450 0.383 Yes 0.351 No VIA*LV

0540 0.253 No 0.150 No

0567 0.158 No 0.234 No

aa 28-29: 0665 0.398 Yes 0.277 No AVA*VV

aa 39-40: aa 25-26: 1426 0.669 Yes 0.633 Yes ALA*GS AAA*AV

aa 39-40: aa 45-46: 1538 0.605 Yes 0.471 Yes HFA*GM AYA*VE

aa 36-37: aa 36-37: 1769 0.623 Yes 0.540 Yes GVA*AR GVA*AR

1 D-score, discrimination score, calculated by SignalP 2 amino acid (aa) position, sequence and exact cleavage site (*) of the predicted signal peptides. For amino acids, the one letter code was used.

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Table 12: Putative surface adhesin proteins and their predicted subcellular localization sites. Scores for subcellular localization were calculated by pSORT.

pSORT v2.0 pSORTb v3.0.2 BBIF_ C-score1 M-score2 E-score3 Final prediction C-score1 M-score2 W-score4 E-score3 Final prediction

0017 0.000 0.501 0.000 Bacterial membrane 2.50 2.50 2.50 2.50 Unknown

0295 0.000 0.380 0.000 Bacterial membrane 0.01 0.01 9.97 0.01 Cellwall

0450 0.000 0.554 0.000 Bacterial membrane 0.01 0.53 8.90 0.56 Cellwall

0540 0.000 0.153 0.000 Bacterial membrane 2.50 2.50 2.50 2.50 Unknown

0567 0.000 0.423 0.000 Bacterial membrane 7.50 1.15 0.62 0.73 Cytoplasmic

0665 0.000 0.552 0.000 Bacterial membrane 0.00 10.00 0.00 0.00 Cytoplasmic Membrane

1426 0.000 0.308 0.000 Bacterial membrane 0.00 0.09 0.18 9.73 Extracellular Unknown (possible 1538 0.000 0.308 0.000 Bacterial membrane 0.00 0.12 5.62 4.26 multiple localization sites)

1769 0.000 0.363 0.000 Bacterial outside 0.00 3.33 3.33 3.33 Unknown

1 C-score, cytoplasm score 2 M-score, cell membrane score 3 E-score, extracellular score 4 W-score, cell wall score

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Figure 6: Domain composition of B. bifidum S17 proteins predicted to be associated with the bacterial cell envelope and a putative function in the interaction of bacteria with host structures. Domains are indicated in colored boxes. Regions targeted by RT-PCR are indicated by blue bars. aNote (p. 114).

3.1.4. Subtilisin like protease

In a previous work, a putative protease BBIF_1681 belonging to the subtilases S8 family of proteins was identified by genome analysis of B. bifidum S17 (Westermann, 2012b). BBIF_1681 displays sequence similarities to lactocepin, an immunomodulatory protease of Lactobacillus casei. However, the similarities are mostly limited to the catalytic domain,

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Results whereas potential substrate recognition sites differ. With a size of 4068 bp, bbif_1681 belongs to one of the largest genes in the genome of B. bifidum S17. By in silico analysis a subtilase S8 domain, two potential sugar-binding domains (FIVAR domains), a C-terminal LPXTG motif, two transmembrane spanning helices, and a secretion signal were identified (Figure 7). The gene bbif_1681 was shown to be expressed under normal culture condition in stationary growth phase (Westermann, 2012b). Furthermore, BBIF_1681 was predicted to be a bacterial membrane protein and thus belongs potentially to CEPs. For a number of extracellular proteases of other microorganisms’ important functions in interaction with the host (Navarre and Schneewind, 1999; Mitsuma et al., 2008; González-Rodríguez et al., 2012) and immunomodulatory function (von Schillde et al., 2012) were shown. Based on these findings BBIF_1681 was named bifidocepin and further analyzed.

Figure 7: BBIF_1681 with the signal peptide (yellow rectangle), the (His, Asp, Ser), the subtilase S8 domain (blue rectangle), the potential sugar binding sites (red sphere, FIVAR, found in various architecture), and the transmembrane domain (TM, grey rectangle).

Table 13 shows an overview of bioinformatic identified structures and the corresponding genes with putative functions. Further analyses of these structures were performed.

Table 13: continued.Overview of bifidobacterial structures potentially involved in adhesion to host structures. Structure Gene (putative) function Reference EPS bbif_0049 protection, Ruas-Madiedo et al., 2010; bbif_0050 immunomodulation, Hidalgo-Cantabrana et al., bbif_0051 colonization 2014; bbif_0052 Fanning et al., 2012a; bbif_0053 Alp and Aslim, 2010 bbif_0054 bbif_0055 bbif_0056 bbif_0057 bbif_0058

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Table 13: continued.

Structure Gene (putative) function Reference EPS bbif_0059 bbif_0060 bbif_0061 bbif_0062 bbif_0063 bbif_0393 tad pili tadV (bbif_0835) adhesion, O'Connell Motherway et al., tadZ (bbif_1703) immunomodulation 2011 tadA (bbif_1702) tadB (bbif_1701) tadC (bbif_1700) flp (bbif_1699) tadE (bbif_1698) fim1 pili bbif_301 adhesion, Foroni et al., 2011; bbif_302 immunomodulation Turroni et al., 2013 bbif_303 fim2 pili bbif_1648 adhesion, Foroni et al., 2011; bbif_1649 immunomodulation Turroni et al., 2013 bbif_1650 fim3 pili bbif_1760 adhesion, Foroni et al., 2011; bbif_1761 immunomodulation Turroni et al., 2013 bbif_1762 cell surface bbif_0295 adhesion Liu et al., 2015; proteins bbif_0450 O'Flaherty and bbif_0540 Klaenhammer, 2010; bbif_0665 Bateman et al., 2005; bbif_1426 Gagic et al., 2013; bbif_1538 Henderson et al., 2011; bbif_1769 Katerov et al., 2000 subtilisin like bbif_1681 immunomodulation, von Schillde et al., 2012 protease adhesion, ECM Williams et al., 2002 interaction

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3.2. Evidence for presence of adhesion structures

3.2.1. EPS

The cell surface layers of B. bifidum S17 and B. longum E18 were analyzed in stationary growth phase by transmission electron microcopy (TEM) and scanning electron microscopy (SEM; Figure 8). As controls Rhodobacter capsulatus wildtype (wt) and a non-capsulated, isogenic mutant were included.

Presence and absence of an EPS capsule could be clearly demonstrated for Rhodobacter capsulatus wt and its non-capsulated mutant, respectively. Bifidobacteria display different amounts and structures of EPS. B. longum E18 is covered with a thick but rather fuzzy layer of EPS. B. bifidum S17 shows fewer and cloudy EPS structures, which seems to be attached more loosely to the cells compared to the compact structures of the other bifidobacterial strain.

Figure 8: Visualization of the EPS capsules of Rhodobacter capsulatus1 wt and non-capsulated mutant, B. bifidum S17 and B. longum E18 in stationary growth phase by TEM and SEM1. Bacteria and EPS were fixed and stained by lysine-based ruthenium red staining. The scale bars indicate a size of 200 µm (TEM) or 2 µm (SEM). 1Images were kindly provided by Dr. Daria Zhurina.

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3.2.2. Expression analysis

Expression of the tad and fim genes was analyzed by semi-quantitative RT-PCR on RNA isolated from bacteria harvested in exponential and stationary growth phase (primer sequences see Table 15, Addendum). Expressions of the genes were normalized against 16S rRNA encoding gene expression in exponential and stationary growth phase.

Most of the genes encoding the tad locus are expressed higher when in bacteria harvested in exponential growth phase (Figure 9). Only the peptidase gene tadV showed higher expression in stationary growth phase. Equal gene expression in both growth phases was shown for the tadE gene, encoding a structural pilus subunit. The genes tadZA are expressed at low levels in stationary growth phase (Figure 9B). The highest expression could be observed for the genes tadZB in exponential growth phase. tadA is the weakest expressed gene in both growth phases. By contrast, no signal could be detected for flp encoding the putative major pilin precursor under tested conditions (Figure 9).

Figure 9: Semi-quantitative assessment of expression in exponential (black bars) and stationary (green bars) growth phase by densitometrical analysis of gel images. For each gene signal intensities of RT-PCR reactions with RNA samples of three independent cultures were quantified and normalized to signal intensities of 16S rRNA. RT-PCR on 16S rRNA was run as a control for equal RNA quantities. Sections of gel images with RT-PCR products are shown in boxes (left: exponential growth phase, right: stationary growth phase).

Transcripts of all genes of the fim2 and fim3 were detected in both growth phases with somewhat higher expression of most genes in exponential growth phase (Figure 10 B-C). All genes of fim2 cluster showed a comparable high adhesion in exponential growth phase, whereas the genes of the fim3 cluster are lower expressed. Additionally, the intergenic

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Results regions in the fim2 and fim3 gene clusters were also expressed (Figure 10 B-C). Of the fim1 cluster, only the gene bbif_0302 is expressed under the tested conditions and no signal could be detected for the genes bbif_0301 and bbif_0303 (Figure 10 A). Only the intergenic region between bbif_302 and bbif_303 is expressed. Like bbif_0302, the intergenic region showed higher expression in exponential growth phase.

Figure 10: Semi-quantitative assessment of expression in exponential (black bars) and stationary (green bars) growth phase by densitometrical analysis of gel images. For each gene signal intensities of RT-PCR reactions with RNA samples of three independent cultures were quantified and normalized to signal intensities of 16S rRNA. RT-PCR on 16S rRNA was run as a control for equal RNA quantities. Sections of gel images with RT-PCR products are shown in boxes (left: exponential growth phase, right: stationary growth phase). Sections of gel images with RT-PCR products are shown in boxes (left: exponential growth phase, right: stationary growth phase).

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Expression of the genes bbif_0017, bbif_0295, bbif_0450, bbif_0540, bbif_0567, bbif_0665, bbif_1426, bbif_1538, and bbif_1769 was analyzed by semi quantitative RT- PCR on RNA samples isolated during exponential and stationary growth (primer sequences see Table 15, Addendum). Transcripts for all of the potential single adhesion protein encoding genes except bbif_1426 could be detected in both growth phases (Figure 11). For bbif_0295, bbif_0450, bbif_0665, and bbif_1538 expression levels were higher in exponential growth phase and this effect was most evident for bbif_0295 (Figure 11). By contrast, bbif_0540 shows higher expression in stationary growth phase (Figure 11). No growth phase dependent expression was observed for bbif_1769 (Figure 11), encoding a protein with a G5 and collagen triple helix repeat domain. Expression levels of bbif_0295, bbif_0450, bbif_0567, and bbif_0665 in exponential growth phase are in the same range. Lowest expression in exponential growth phase was observed for bbif_0540.

Figure 11: Expression of the identified genes encoding the putative adhesins was analyzed by RT-PCR on RNA samples isolated in exponential (left band) and stationary (right band) growth phase. Results of RNA samples from one representative of three independent cultures are shown. Semi-quantitative assessment of expression in exponential (black bars) and stationary (green bars) growth phase by densitometrical analysis of gel images. For each gene signal intensities of RT-PCR reactions with RNA samples of three independent cultures were quantified and normalized to signal intensities of 16S rRNA. RT-PCR on 16S rRNA was run as a control for equal RNA quantities.

As indicated by semi quantitative RT-PCR most of the putative adhesins are expressed at higher levels in exponential growth phase. Further experiments with Caco-2 cell revealed that adhesion of B. bifidum S17 was about 2-fold higher when bacteria were harvested in

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Results exponential growth phase compared to the adhesion of bacteria in stationary growth phase (Figure 12).

Figure 12: Adhesion of B. bifidum S17 grown to exponential or stationary growth phase to Caco- 2 cells. Values are mean ± standard deviation of triplicate measurements. Results from one representative of three independent experiments are shown. Statistical analysis was performed using Student’s t-test (*p<0.05). aNote (p. 114).

3.3. Functional characterization

3.3.1. Cloning the putative adhesins

To further investigate the role of the identified putative adhesins in adhesion to host tissue, the corresponding genes were cloned into an E. coli – Bifidobacterium shuttle vector pMDY23 under control of the Pgap promoter for constitutive high level expression in bifidobacteria (Grimm et al., 2014, Sun et al., 2014a). For this purpose, the reporter gene gusA after Pgap promoter was removed with the restriction enzymes XhoI and HindIII (Table 14, Addendum) and the vector backbone pMDY23 was isolated from a 0.8 % (w/v) agarose gel after size dependent separation. The predicted cell surface adhesins encoding genes were amplified by PCR introducing restriction sites and a C-terminal His6-tag using the corresponding primer pairs shown in Table 15 (Addendum). Following digestion with the endonucleases XhoI and HindIII (Tabel 14, Addendum), the PCR product was ligated to vector backbone pMDY23 as a transcriptional fusion to Pgap. As control, a plasmid containing the promoter Pgap without a gene was constructed. For this purpose, the reporter gene gusA was removed from pMDY23 by restriction with XhoI and HindIII. The vector

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Results backbone pMDY23 after isolation from 0.8 % agarose gel and DNA ends were blunted by Klenow fill-in reaction. Then the vector backbone was religated yielding pCW (Figure 13). The resulting plasmids pCW, pCW_295, pCW_450, pCW_540, pCW_665, pCW_1426, and pCW_1769 (Figure 13) were transformed into E. coli DH10B and positive clones were selected on agar plates containing spectinomycin. Following confirmation by restriction analysis, plasmids were subcloned into E. coli ET12567::p16S_ParaB_bbif0710 for amplification. This E. coli strain lacks own DNA methylases and expresses the type II methyltransferase BBIF_0710 of B. bifidum S17 and can thus used for PAM cloning into B. bifidum S17. After transformation, colonies were screened for the presence of the plasmid by PCR using primers pVG_cont_f and corresponding reversed primer (Table 15, Addendum). The plasmid of a positive clone was isolated and the sequence was checked by Sanger sequencing using specific primers (Table 15, Addendum). Confirmed potential cell surface adhesin plasmids were transformed into B. bifidum S17 and B. longum E18. After transformation, bacteria were cultivated on agar plates containing spectinomycin. Positive clones were checked for presence of plasmid by PCR using primers pVG_cont_f and corresponding reversed primer (Table 15, Addendum).

All constructs were successfully transformed into B. bifidum S17 and B. longum E18 except the plasmid containing bbif_1538. Despite successful amplification of a mutation- free PCR product, ligation to XhoI/HindIII-cut pMDY23 backbone, and transformation into E. coli DH10B and ET12567::p16S_ParaB_bbif0710 neither recombinant B. bifidum S17 nor B. longum E18 strains could be obtained after several transformations.

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Figure 13: Schematic representation of plasmids generated for expression of putative adhesins. PCR products of the coding genes (green arrows) were cloned as a transcriptional fusion downstream of Pgap promotor into the backbone of pMDY23 following excision of gusA by restriction with XhoI and HindIII. Plasmid features: Pgap promoter (Pgap), E. coli origin of replication (rep) for E. coli, pMDY23 origin of replication (repAB) for bifiodbacteria, spectinomycin resistance gene (spc).

3.3.2. Adhesion of the putative adhesins overexpression strains

Following successful generation of recombinant B. bifidum S17 and B. longum E18 strains, adhesion to Caco-2 cells was tested (Figure 14 and 15). As controls wt strains and strains carrying the plasmid pCW were used. As observed in previous studies (Riedel et al., 2006b; Preising et al., 2010; Gleinser et al., 2012), B. bifidum S17 wt shows a high level of adhesion to Caco-2 cells. Adhesion of the strain harboring the plasmid pCW was indistinguishable from that of the wt strain. However, adhesion of B. bifidum S17 could be significantly increased by expression of bbif_0295, bbif_0450, bbif_0665, bbif_1426, and

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Results bbif_1769. The increase in adhesion was most prominent for strains B. bifidum S17/pCW_295 B. bifidum S17pCW_450, B. bifidum S17/pCW_1426, and B. bifidum S17/pCW_1769. A minor significant effect could be observed for B. bifidum S17/pCW_665. Expression of bbif_0540 did not alter adhesion of B. bifidum S17.

Figure 14: Adhesion of B. bifidum S17, recombinant derivatives expressing single adhesins to Caco-2 cells. As an empty vector control, B. bifidum S17/pCW was included. Adherent bacteria were determined by spot plating on MRSC agar plates and expressed as percentage of the initially added bacteria (i.e. 5 x 107 bacteria/well). Values are mean ± standard deviation of three independent experiments each performed in triplicate. Statistical analysis was performed using ANOVA with Bonferroni post-test analysis for multiple comparisons. Levels of statistical significance are indicated by asterisks (*p<0.05; ** p<0.01, *** p<0.001, **** p<0.0001).

In contrast to the results obtained with B. bifidum S17 strains, expression of most genes for single adhesions did not result in altered adhesion of B. longum E18 (Figure 15). Similar to

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Results previous studies (Preising et al., 2010; Gleinser et al., 2012), B. longum E18 wt showed only a marginal adhesion (0.02-0.06 %) to Caco-2 cells. As observed for B. bifidum S17, presence of the empty vector pCW had no effect on adhesion of B. longum E18. Similarly, expression of most genes did not improve the adhesive capacity of B. longum E18. The only strain that actually did show improved adhesion to Caco-2 cells was B. longum E18/pCW_665 with 1.9 fold higher adhesion than the wt strain.

Figure 15: Adhesion of B. longum E18, recombinant derivatives expressing single adhesins to Caco-2 cells. As an empty vector control, B. longum E18/pCW was included. Adherent bacteria were determined by spot plating on MRSC agar plates and expressed as percentage of the initially added bacteria (i.e. 5 x 107 bacteria/well). Values are mean ± standard deviation of three independent experiments each performed in triplicate. Statistical analysis was performed using ANOVA with Bonferroni post-test analysis for multiple comparisons. There are no significant differences.

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3.3.3. Creating fluorescent B. bifidum S17 expressing the putative adhesin bbif_0295

Expression of the gene encoding BBIF_0295 resulted in the most prominent improvement in adhesion of B. bifidum S17 to Caco-2 cells. Thus, it was attempted to confirm this effect in a second independent approach. For this purpose, the corresponding gene was cloned into the plasmid pVG_mCherry. This plasmid contains the gene encoding for the fluorescent protein mCherry under the control of the promoter Pgap and allows detection of bifidobacteria by fluorescent microscopy.

To construct this plasmid, the bbif_0295 gene was cloned into the E. coli –

Bifidobacterium shuttle vector pVG_mCherry under the control of the constitutive Phyper promoter. For this purpose, the plasmid pVG_mCherry was opened with the restriction enzyme NdeI and BglII. The gene bbif_0295 was amplified using the primer pair BBIF295_XhoI_fw and BBIF295_BglII_rev (Table 15, Addendum) introducing restriction sites XhoI and BglII. The Phyper promoter was similarly amplified using primers

Phyper_NdeI_fw and Phyper_XhoI_rev introducing restriction sites NdeI and XhoI during amplification. This synthetic promoter was described to have high transcriptional activity in a number of Gram-positive bacteria (Jensen et al. 1998; Riedel et al., 2007; Balestrino et al. 2010) including bifidobacteria (Sun, unpublished results). Following digestion with the endonuclease XhoI, the promoter Phyper was ligated to the PCR product of bbif_0295. A second amplification with the primer Phyper_NdeI_fw and BBIF295_BglII_rev was performed. After digestion with the endonucleases NdeI and BglII, the PCR construct was ligated with the plasmid pVG_mCherry. The resulting plasmid pCW_mCherry_295 (Figure 16) was transformed into E. coli DH10B and positive clones were selected on agar plates containing spectinomycin. Following confirmation by restriction analysis, pCW_mCherry_295 was subcloned into E. coli ET12567::p16S_ParaB_bbif0710 for amplification. This E. coli strain lacks own DNA methylases and expresses the type II methyltransferase BBIF_0710 of B. bifidum S17 and can thus used for PAM cloning. After transformation, colonies were screened for the presence of the plasmid by PCR using primers pVG_cont_fw and BBIF0295_cont_rev (Table 15, Addendum). The plasmid of a positive clone was isolated and the sequence was checked by Sanger sequencing using specific primers (Table 15, Addendum). Confirmed pCW_mCherry_295 was transformed into B. bifidum S17. After transformation, bacteria were cultivated on agar plates

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Results containing spectinomycin. Positive clones were tested for fluorescence under the microscope.

Figure 16: Schematic representation of plasmid pCW_mCherry_295 generated for expression of bbif_0295 and the fluorescent protein mCherry. PCR product of the coding gene bbif_0295 (green arrows) under control of Phyper promoter was cloned upstream of Pgap promotor into the plasmid pVG_mCherry by restriction with NdeI and BglII. Plasmid features: Phyper promoter (Phyper), Pgap promoter (Pgap), E. coli pMB1 origin of replication (rep) for E. coli, pMDY23 origin of replication (repAB) for bifiodbacteria, spectinomycin resistance gene (spc).

3.3.4. Competitive adhesion with fluorescent B. bifidum S17

To confirm the effect of BBIF_0295, competitive adhesion assay was performed with the strains B. bifidum S17/pVG_GFP and pCW_mCherry_295. After adhesion, unbound bacteria were washed off and cells were fixed with paraformaldehyde. Nuclei from Caco-2 cells were stained with Hoechst. Fluorescent bifidobacteria and cell nuclei were imaged by fluorescence microscopy (Figure 17 A). This revealed that a proportion of bacteria bound to Caco-2 cells were labeled by red fluorescence indicating that they were B. bifidum S17/pCW_mCherry_295. Further quantitative pixel analysis of green and red fluorescent signals independent of the fluorescence intensities using ImageJ software 1.48v confirmed that the proportion of red fluorescent bacteria Caco-2-bound was higher than the proportion of green fluorescent bacteria (Figure 17 B).

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(A) (B)

5 ]

% 4

[

a

e 3

r

a

t

n 2

e

c r

e 1 p 0

y r P r F e G h C m

Figure 17: (A) Fluorescence microscopy of the competitive adhesion of B. bifidum S17/pVG- GFP (green) and B. bifidum S17/pCW_mCherry_295 (red) to Caco-2 cells. Nuclei of Caco-2 cells were stained with Hoechst (blue). Image was acquired using Zeiss Axio Observer.Z1 microscope and a 40X objective. The scale bar indicates a size of 20 µm. (B) Pixel values displayed in percent of red and green areas of three fluorescence microscopy images. The images were analyzed using ImageJ software (1.48v).

3.3.5. Cloning of bifidocepin

Due to sequence similarities to lactocepin (von Schillde et al., 2012), bifidocepin could act as immunomodulatory protease or is able to interact with host structures. As shown for Staphylocoddus epidermidis NCTC11047, proteins containing FIVAR domains are able to bind fibronectin (Williams et al., 2002). To further analyze the function of bifidocepin, overexpression strains were produced and adhesion assay to fibronectin was performed to test either protease or adhesion activity.

The bbif_1681 gene encoding bifidocepin was cloned into the E. coli – Bifidobacterium shuttle vector pMDY23 under the control of the constitutive Pgap promoter. For this purpose, the reporter gene gusA after Pgap promoter was removed with the restriction enzymes XhoI and HindIII (Table 14, Addendum) and the vector backbone pMDY23 was isolated from a 0.8 % (w/v) agarose gel after size dependent separation. The gene bbif_1681 was amplified by PCR introducing restriction sites and a C-terminal His6-tag using the primer pair BBIF1681_XhoI_fw and BBIF1681_HindIII_rev (Table 15; Addendum). Following digestion with the endonucleases XhoI and HindIII (Tabel 14, Addendum), the PCR product was ligated to vector backbone pMDY23 as a transcriptional fusion to Pgap. The resulting plasmid pCW_1681 (Figure 18) was transformed into E. coli

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DH10B and positive clones were selected on agar plates containing spectinomycin. Following confirmation by restriction analysis, pCW_1681 was subcloned into E. coli

ET12567::p16S_ParaB_bbif0710 for amplification. This E. coli strain lacks own DNA methylases and expresses the type II methyltransferase BBIF_0710 of B. bifidum S17 and can thus used for PAM cloning. After transformation, colonies were screened for the presence of the plasmid by PCR using primers pVG_cont_f and BBIF1681_cont_rev (Table 15, Addendum). The plasmid of a positive clone was isolated and the sequence was checked by Sanger sequencing using specific primers (Table 15, Addendum). Confirmed pCW_1681 was transform into B. bifidum S17, B. breve S27, and B. longum E18. After transformation, bacteria were cultivated on agar plates containing spectinomycin. Positive clones were checked for presence of pCW_1682 by PCR using primers pVG_cont_f and BBIF1681_cont_rev (Table 15, Addendum).

Figure 18: E. coli - Bifidobacterium shuttle vector pCW_1681 for expression of bifidocepin (BBIF_1681) with a C-terminal His6-tag using the constitutive Pgap promoter. Cleavage sites of relevant restriction enzymes are indicated. Plasmid features: E. coli origin of replication (rep (pMB1)), Bifidobacterium origin of replication (repAB), spectinomycin resistance gene (spc), promoter of glyceraldehydes-3-phosphate dehydrogenase gene (Pgap), bbif_1681 gene encoding bifidocepin.

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3.3.6. Identification of the cellular localization of BBIF_1681

In order to check the subcellular localization of bifidocepin, crude extracts as well as cytoplasmic, cell envelope and secreted protein fractions of B. bifidum S17/pCW_1681, B. breve S27/pCW_1681, and B. longum E18/pCW_1681 were prepared and analyzed by

Western Blot using a mouse anti-His6 antibody and HRP labeled anti-mouse antibody for detection of His-tagged bifidocepin (Figure 19). A signal was at approx. 140 kDa, i.e. the expected size of His-tagged bifidocepin, in the cell envelope fractions of all strains tested. No signal could be detected for the cytoplasm and supernatant fraction. Bifidocepin could also be detected in crude cell extracts of B. breve S27 and B. longum E18.

Figure 19: Western Blot on cell fractions ([1] crude cell extract, [2] cytoplasm, [3] cell envelope, [4] supernatant of (A) B. bifidum S17/pCW_1681, (B) B. breve S27/pCW_1681, and (C) B. longum E18/pCW_1681. Samples were loaded with a protein concentration of 0.5 mg. TM BBIF_1681 was detected through anti-His6 antibody. Marker: PageRuler Prestained Protein Ladder.

3.3.7. Adhesion to extracellular matrix proteins

To further investigate the role of bifidocepin, the adhesion to extracellular matrix proteins (fibronectin, collagen, and fibrinogen) were tested. For this purpose, 96-well plates were covered with these compounds of the extracellular matrix and adhesion of bacteria was tested with a crystal violet assay. Neither wt strains of B. bifidum S17, B. breve S27, and B. longum E18 nor the recombinant derivatives harboring for pCW_1681 expression of bifidocepin showed adhesion to fibrinogen or collagen (data not shown). B. bifidum S17 (Figure 20 C) but not B. breve S27 and B. longum E18 (data not shown) showed adhesion

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Results to fibronectin. However, adhesion of B. bifidum S17/pCW_1681 to fibronectin was decreased (Figure 20). None of the strains tested showed adhesion to BSA ruling out non- specific interactions.

BBIF_0450 contains a fibronectin binding domain and BBIF_1769 harbors domains described to be involved in adhesion to collagen and fibronectin. Also, B. bifidum S17 was shown to adhere to fibronectin (Figure 20). Thus, further experiments were performed testing adhesion of recombinant bifidobacteria expressing bbif_0450 and bbif_1769 to fibronectin. However, none of the genes were able to improve adhesion of B. bifidum S17 (Figure 20) or B. longum E18 to this ECM protein (data not shown).

(A) (B)

S17 wt

S17/pCW_1681

Figure 20: Adhesion of B. bifidum S17 and the corresponding recombinant strains harboring pCW_0, pCW_450, and pCW_1681 for expression of potential cell surface adhesion protein BBIF_0450 and bifidocepin to fibronectin. As controls wells were coated with BSA. Adhesion was analyzed by crystal violet assay (A) or microscopy (B). Background subtraction was done using values of coated wells without bacteria. Microscopic images (B) were acquired with a Zeiss Axio Observer.Z1 microscope using a 10x objective. The scale bar (red) indicates a size of 20 µM. Statistical analysis was performed using ANOVA with Bonferroni post-test analysis for multiple comparisons. Levels of statistical significance are indicated by asterisks (* p<0.05; ** p<0.01; *** p<0.001). The pairs (wt, pCW), (wt, pCW_450), and (pCW, pCW_450) have no significant differences.

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4. Discussion

Bifidobacteria are one of the major genera in the gastrointestinal tract of humans (Kurokawa et al., 2007; Mariat et al., 2009; Yatsunenko et al., 2012). They are one of the most frequently commercialized probiotics, due to their reported beneficial effects to human health. In order to exert a positive effect on their host, bifidobacteria must be able to survive the human intestine passage and persist at least for some time in the gut (González-Rodríguez et al., 2013). Adherence of bacteria to host structures is discussed as one of the key features important for colonization and interaction with the host. Thus, one of the selection criteria for potential probiotic strains is their ability to adhere to IECs (Lee and Salminen, 2009). As a consequence, investigations of the molecular mechanisms promoting adhesion of probiotic bacteria to host tissue are of pivotal importance.

In the presented study, cell surface structures of B. bifidum S17 with potential role in adhesion to host structures were investigated. Furthermore, the role in adhesion of some predicted single adhesion proteins was investigated in functional assays.

4.1. EPS

Microbial EPS are produced of a vast variety of bacteria and seems to possess protective function against environmental stress, immunomodulatory activity, and adherence to host structures (Ruas-Madiedo et al., 2009a; O'Connell Motherway et al., 2011). Recently, the EPS of Lactobacillus rhamnosus KF5 was reported to stimulate the immune system through increasing splenocytes proliferation in vitro (Shao et al., 2014). Wang and co- authors showed that EPS extracted from Lactobacillus plantarum 70810 is able to inhibit the proliferation of HT-29 tumor cells (Wang et al., 2014). The ability of purified EPS from 13 strains belonging to the three Bifidobacterium species B. animalis, B. longum, and B. pseudocatenulatum to induce T cell differentiation was shown to be highly dependent on the type of polymer (Lòpez et al., 2012). Surface associated EPS from B. breve UCC2003 was shown to provide stress tolerance and have a positive impact during in vivo persistence (Fanning et al., 2012a). However, using a model of an EPS-producing B. breve UCC2003 strain and EPS-deficient derivative strain, the absence of EPS increased the level and activity of adaptive immune response. EPS positive strains were able to reduce colonization levels of the gut pathogen Citrobacter rodentium (Fanning et al., 2012b). Whether EPS extracted from Lactobacillus rhamnosus GG had no effect on the adhesion

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Discussion of its host and B. animalis IPLA-R1 to human intestinal mucus, EPS from B. animalis IPLA-R1 and B. longum NB667 was shown to reduce the adhesion of Lactobacillus rhamnosus GG and B. animalis IPLA-R1 significantly (Ruas-Madiedo et al., 2006). Thus, EPS can possess different sometimes oppositional effects which seem to be highly strain specific.

However, the structural characteristics of EPS responsible these differences have not been identified so far (Hidalgo-Cantabrana et al., 2014). Genome analysis of B. bifidum S17 revealed the presence of putative EPS biosynthesis genes (Table 10; Oehlke, 2013). The existence of EPS on the surface of this strain grown in vitro was analyzed using TEM and SEM with lysine and ruthenium red stained polysaccharide structures (Hammerschmidt et al., 2005; Jubair et al., 2014).

In line with the genomic data, EPS structures were visible around the cells of B. bifidum S17 and B. longum E18. The EPS structures of B. bifidum S17 and B. longum E18 appeared to be completely different in thickness and composition. While B. longum E18 had continuous, thick yet somewhat fuzzy layer of EPS, the EPS of B. bifidum S17 seems to be more compact and present only as small patches or blebs in distinct. Phenotypic observation that could be linked to differences in EPS structure and composition is that, in liquid cultures, B. longum E18 grows equally distributed throughout the culture and is difficult to precipitate by centrifugation. By contrast, B. bifidum S17 grows at the bottom of the growth vessel and forms tightly packed pellets after centrifugation. Similar observations have been described for Lactobacillus johnsonii FI9785, for which the authors reported higher autoaggregation of mutants with reduced EPS (Dertli et al., 2015). Although the discontinuous, patchy EPS structure of B. bifidum S17 might be an artifact of dehydration during fixation and staining, this points towards different polysaccharides composition in the EPS of the two strains as reported for other bifidobacteria (Salazar et al., 2009, Ruas-Madiedo et al., 2010).

Differences in extent and composition of EPS might also provide an explanation for the differences in adhesion to Caco-2 cells (8-11 % for B. bifidum S17 vs. 0.02-0.06 % for B. longum E18). Lactobacillus mutants with reduced EPS production adhere at significantly higher levels to HT29 cell line than the wt strain producing large amounts of EPS (Horn et al., 2013). A possible explanation could be the thickness and density of the EPS surrounding the cell, which maybe interfere with structures that mediate adhesion.

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There are a number of studies that reveal that EPS structures possess immunomodulatory effects, facilitate pathogen exclusion, play a role in bacterium-host interaction, prevent bacteria for pH stress, and mediate bile resistance (Fanning et al., 2012a, Fanning et al., 2012b, Wu et al., 2010, Alp and Aslim, 2010, Ruas-Madiedo et al., 2007). In order to investigate if the EPS structures of B. bifidum S17 and B. longum E18 are indeed EPS with similar properties, EPS of these strains will have to be purified or mutants deficient in EPS of the respective strains have to generate.

4.2. Putative pili

Further in silico analysis of the genome of B. bifidum S17 for structures potentially involved in colonization was performed and one cluster of genes for type IV tight adherence (tad) pili and three gene clusters for sortase dependent pili (fim1, fim2, fim3) were identified. For Gram-positive bacteria, Tad pili were described mainly to be found in pathogens, like Yersinia pestis, Vibrio cholera, Mycobacterium tuberculosis, Pseudomonas aeruginosa, Bordetella pertusis, to name a few (Tomich et al., 2007). In pathogens, this kind of pili was described to be a virulence factor and mediate adhesion to host tissue (Tomich et al., 2007; Bernard et al., 2009). Bifidobacterial Tad pili were shown previously to be highly expressed under in vivo conditions in the murine GIT and play a role in colonization from B. breve UCC2003 of the murine intestine (O'Connell Motherway et al. 2011).

Expression of the tad pili cluster of B. bifidum S17 was tested during in vitro growth in exponential and stationary phase by RT-PCR. Most of the genes encompassing the tad locus are expressed with higher expression in exponential growth phase. Absence of an RT-PCR signal for the putative precursor of the major pilin subunit encoding gene flp under laboratory growth conditions suggests that no functional Tad pili are build during in vitro growth. For B. breve UCC2003 it was shown that the expression of tad pili encoding genes was up-regulated in vivo (O'Connell Motherway et al., 2011). This might indicate that the genes encoding for tad pili of B. bifidum S17 are only expressed when bacteria are in contact with their host. For some tad genes, expression regulation by quorum sensing or transcriptional regulators was shown (Schuster et al., 2003; Vallet et al., 2004; Juhas et al., 2005). There are also some reports of other genes of Bifidobacterium and Lactobacillus strains which expressions are induced when bacteria in contact with the murine gut (Marco

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Discussion et al., 2007; O'Connell Motherway et al., 2011). So far, host factors responsible for this expression up-regulation are still unknown. However, the presence of genes encoding tad pili indicate that at least in vivo this could be one possible adhesion mediating factor of B. bifidum S17.

Sortase-dependent pili were found in different members of the genus Bifidobacterium and Lactobacillus (Foroni et al., 2011; Lebeer et al., 2012; Turroni et al., 2014). The pivotal role of sortase-dependent pili in efficient adherence and immunomodulatory interactions with human gut cells were shown for Lactobacillus rhamnosus GG (Lebeer et al., 2012). In bifidobacteria, the role of sortase-dependent pili in adhesion to ECM proteins and IECs was shown by expression of the corresponding genes of B. bifidum PRL2010 in non piliated Lactococcus lactis. Fibronectin deglycosylation of O-linked and N-linked oligosaccharides are described to be putative receptors of Pil3 pili, whereas mannose and fucose are potential receptors for Pil2. Furthermore, immunomodulatroy effects for Pil3 pili was shown by enhancement of expression levels of TNF-α and inhibition of interleukin 10 response in the murine GIT in vivo (Turroni et al., 2013). Similarly to Tad pili, expression of two of the three gene clusters for sortase dependent pili of B. bifidum PRL2010 are up-regulated under in vivo conditions. Pili gene expression is also altered in response to conditions that reflect the GIT and the co-cultivation with other commensal bacteria (Turroni et al. 2014). Expression of bifidobacterial sortase dependent pili is influenced when bacteria exposed to thermal, osmotic, and acidity stress. Turroni and coworker demonstrated also that the expression level of pil3 locus of B. bifidum PRL2010 is enhanced upon co-cultivation with Lactobacillus paracasei and B. breve 12L (Turroni et al. 2014). These results indicate that pili not only support the interaction with host cells but also provide species independent microorganism-microorganism interaction.

Expression of the sortase dependent pili cluster of B. bifidum S17 was tested during in vitro growth in exponential and stationary phase by RT-PCR. Transcripts of all genes of fim2 and fim3 gene cluster were detected with higher expression of most genes in exponential growth phase. Growth phase dependent expression was also shown for genes encoding pili and other proteins potentially involved in persistence and survival of Lactobacillus rhamnosus GG with higher levels of expression in exponential growth phase. Thus, it was speculated that higher expression in exponential phase of pili encoding genes provides a competitive advantage for L. rhamnosus GG by promoting its survival and persistence in the GIT (Laakso et al. 2011). RT-PCR signals for intergenic regions of fim2

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Discussion and fim3 gene clusters indicate transcription as polycistronic operons under the tested conditions. This contradicts the results of Foroni and co-authors who suggested a differential regulation of the pilin genes and the corresponding sortases (Foroni et al. 2011). Further expression analysis using the primers specified in the study by Foroni et al. (2011) on the same cDNA yielded no PCR product while DNA was used as positive control (data not shown). Although we cannot rule out that there is a single out of the limit of detection, this result showed that the primers used by Foroni et al. for amplification of intergenic regions of fim2 and fim3 clusters are not optimal.

Interestingly, the transcript of bbif_1649 (i.e. the minor pilin subunit encoding gene of the fim2 cluster) contains a guanine homopolymer sequence at the start of the coding sequence (data not shown). Genes containing nucleotide polymorphism sides are described as pseudogenes with nonfunctional sequences, and therefore the corresponding mRNA is possibly not transcribed to a functional protein (Balakirev and Ayala, 2003). Similar frameshift-causing sequences were observed in pilus clusters of different bifidobacteria. The fimB (bbpr_1822, minor pilin subunit) of the pil1 gene cluster of B. bifidum PRL2010 and two genes (bbr_0113, bbr_1889) encompassing two of three pili encoding gene cluster in B. breve UCC2003 are described as pseudogenes due to a frameshift within a stretch of nine, 11 or 10 guanine nucleotides (Foroni et al. 2011, O'Connell Motherway et al. 2011). The authors suggested that, under certain environmental conditions the frameshift along the guanine residues is suppressed resulting in functional proteins (Foroni et al. 2011). In Salmonella enteric serovar Typhimurium, analysis of gene expression with microarrays showed, that pseudogenes are expressed during infection of epithelial cells (Hautefort et al., 2008). Therefore, it could be speculated that bbif_1649 is expressed to a functional protein in vivo during colonization. However, to prove this hypothesis more experiments are necessary.

Two genes of fim1 cluster are not expressed under the tested conditions. It could be suggest, that intact fim1 pili are only produced when the gene expression is induced by special stimuli. However, through the results of the present study it can be hypothesized that at least one pili encoding gene cluster is expressed to functional pili on the cell surface of B. bifidum S17 under tested conditions.

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4.3. Cell surface proteins

For different Lactobacillus strains mucin-, fibronectin-binding proteins as well as S-layer proteins, pilus structures and moonlighting proteins were described to contribute to adhesion (Rojas et al. 2002, Macías-Rodríguez et al. 2009, Kos et al. 2003, Weiss and Jespersen 2010, von Ossowski et al. 2010, Kinoshita et al. 2008). A few studies have also investigated bifidobacterial adhesion to IECs. Adhesion of B. breve strain 4 was shown to be mediated by proteinaceous components (Bernet et al. 1993). Candela et al. (2007) showed that four bifidobacterial strains belonging to three different species (B. lactis, B. bifidum, B. longum) are able to bind plasminogen, a glycoprotein found amongst others tissue in the intestine (Zhang et al., 2002). In B. animalis subsp. lactis BI07 DnaK was identified as protein with a plasminogen-binding motif on their cell surface (Candela et al., 2010). Furthermore, the cell surface lipoprotein BopA was shown to mediate adhesion to Caco-2 cells in B. bifidum strains (Guglielmetti et al. 2008, Gleinser et al., 2012). Further proteins including the enzyme transaldolase of B. bifidum A8 (Gonzáles-Rodríguez et al. 2013) have been described to be involved in adhesion to host tissue.

Kankainen and co-authors published a list with protein domains known to be involved in adhesion (Kankainen et al., 2009). In Lactobacillus rhamnosus GG they identified nine proteins with collagen associated domain, nine proteins with fibronectin associated domain, one protein with Immunoglobulin (Ig) associated domain, two proteins with invasion associated domain, four proteins with mucus associated domain, seven proteins with peptidoglycan associated domain and two pili proteins. All together, they found 33 proteins with adhesion related domains in total in Lactobacillus rhamnosus GG. However, comparative genome analysis of 14 Lactobacillus strains revealed the presence of six till 37 proteins with adhesion related domains in total.

Nine predicted cell surface proteins with domains known or suspected to interact with host structures were identified through in silico analysis of the predicted proteome of B. bifidum S17 (Westermann et al., 2012a). For updating the data, the analysis with actual versions of software were performed and seven putative adhesins remained for further analysis. For B. bifidum S17, one protein with collagen associated domain, one protein with fibronectin associated domain, one protein with Ig associated domain, one protein with lectin associated domain, one protein with general adhesion associated domain, two proteins with epithelium associated domains were identified. Compared to the 33 identified proteins with adhesion related domains of Lactobacillus rhamnosus GG, only seven proteins with

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Discussion adhesion related domains could be identified. However, there are no hints that the number of identified proteins with adhesion related domains correlate with the adhesion ability of the corresponding strain. Lactobacillus delbrueckii ATCC 11842 with six identified proteins with adhesion related domains (no protein with mucus associated domain) showed higher adhesion to intestinal mucus compared to Lactobacillus rhamnosus GG with 33 identified proteins with adhesion related domains (four proteins with mucus associated domain; Apostolou et al., 2001; Kankainen et al., 2009). Whereas no protein with epithelium associated domain could be found in L. rhamnosus GG, two proteins with epithelium associated domain were identified in B. bifidum S17. There is also no protein with lectin and general adhesion associated domain identified in L. rhamnosus GG, whereas one could be found in B. bifidum S17 respectively. Four proteins with mucus associated domain, two proteins with invasin associated domain and seven proteins with peptidoglycan associated domain were identified in L. rhamnosus GG and no proteins with any of these domains could be found in the theoretical proteome of B. bifidum S17. However, detailed investigations are necessary to elicit the role in adhesion of each identified protein with adhesion related domain in B. bifidum S17.

BBIF_0295 and BBIF_1538 were identified as proteins containing one or more bacterial Ig-like domains (defined by Pfam domain database: PF07523, PF02368). Other surface proteins contain bacterial Ig-like domains and were described to be involved in adhesion and binding to extracellular structures (Hultgfien et al., 1993; Takai and Ono, 2001; Fraser et al., 2007; Gagic et al., 2013). Furthermore, this domain was found in surface proteins of pathogenic bacteria including Leptospira interrogans, enteropathogenic E. coli, Yersinia enterocolitica, Yersinia pseudotuberculosis, and Streptococcus pneumonia, which are described as virulence factors involved in adhesion to host cells (Hamburger et al., 1999; Kelly et al., 1999; Luo et al., 2000; Wang et al., 2013; Mei et al., 2015;).

Additionally, fibronecctin type III domains (PF00041) were discovered in Gram-positive bacteria, belonging to microbial surface components recognizing the extracellular matrix component fibronectin (Henderson et al., 2011). The potential surface adhesion protein BBIF_0450 was identified containing a fibronectin type III domain.

The myosin cross-reactiv antigen (PF06100) was first discovered in Streptococcus pyrogenes and was demonstrated to play a role in adherence to keratinocytes (Kil et al., 1994; Volkov et al., 2010). This domain was also identified in a cell wall protein of Lactobacillus acidophilus and seems to mediate adhesion to IEC (O'Flaherty and

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Klaenhammer, 2010). Furthermore, the myosin cross-reactive antigen is conserved among many probiotic bacteria (Volkov et al., 2010). This domain was also shown to play a role in stress tolerance of B. breve (Rosberg-Cody et al. 2011) and Lactobacillus acidophilus (O'Flaherty and Klaenhammer 2010). In B. bifidum S17 the potential surface protein BBIF_0540 contains a streptococcal 67 kDa myosin-cross-reactive antigen like family domain.

In Streptococci the von Willebrand factor domain was found to be involved in adhesion process to extracellular matrix components like fibronectin (Colombatti et al., 1993; Katerov et al., 2000). There are also reports on promotion of unspecific adhesion by proteins with von Willebrand factor domains (Kachlany et al., 2000). A protein with a von Willebrand factor was identified encoded by gene bbif_0665.

NlpC/P60 domain have either a signal peptide or additional a transmembrane spanning region, indicating an extracellular localization of the protein (Anantharaman and Aravind, 2003). For example cell wall lipoproteins containing the NlpC/P60 family domain (PF00877) were shown to be involved in adhesion to epithelial cells (Anantharaman and Aravind, 2003; Tran et al., 2010; Liu et al., 2015). BBIF_1426 with an NlpC/P60 domain was identified through in silico analysis of the theoretical proteome of B. bifidum S17.

The G5 domain (PF07501), which is named after its conserved glycine residues, is widespread among cell surface binding proteins and found in proteins involved in cell wall metabolism (Bateman et al., 2005; Ruggiero et al., 2009). One of these proteins, BBIF_1769, contains a G5 domain described to recognize N-acetylglucosamines, found in human milk oligosaccharides (HMOs, Marcobal and Sonnenburg, 2012). Furthermore, BBIF_1769 contains a lysozyme like domain (NCBI CDD Conserved Protein Domain Family code cl00222), which is described to be involved in the hydrolysis of beta-1,4-linked polysaccharides, as it can be found in HMOs (Bode and Jantscher- Krenn, 2012; Xu et al., 2014). Thus, BBIF_1769 possibly supports colonization of breast- fed infants.

Expression of the genes encoding the identified putative adhesins of B. bifidum S17 was tested during in vitro growth in exponential and stationary phase by RT-PCR. Except bbif_1426, all genes encoding putative adhesins are expressed at both growth phases. Most of the genes encoding putative adhesins showed a higher expression in exponential growth

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Discussion phase. Only the gene bbif_0540 encoding a protein with a myosin cross-reactive antigen domain is higher expressed in stationary growth phase.

Due to the growth phase dependent expression of the putative adhesins, adhesion to Caco-2 cells with bacteria harvested in exponential and stationary growth phase was tested. In line with increased expression of most of the putative adhesins, B. bifidum S17 adheres significantly higher to Caco-2 cells when bacteria in exponential growth phase (Figure 12). To allow a stable bacterial population in the human GIT, bacteria must be in exponential growth phase. Thus, it explains higher expression of proteins involved in adhesion and colonization in exponential growth phase.

To functionally demonstrate a contribution of the putative adhesins, recombinant strains of B. bifidum S17 and B. longum E18 were generated. These strains harbored plasmids for high level constitutive expression of all identified putative adhesins except BBIF_1538 and were used in adhesion assays with Caco-2 cells. Expression of all but one (BBIF_0540) putative adhesins significantly increased adhesion of B. bifidum S17 to Caco-2 cells showing that, in principal, these proteins somehow contribute to adhesion to IECs. BBIF_0540 contains two streptococcal 67 kDa myosin-cross-reactive antigen like family domain and is not able to enhance the adhesion of B. bifidum S17 to IECs. This domain is also involved in stress tolerance (O'Flaherty and Klaenhammer 2010; Rosberg-Cody et al. 2011), which could be explain that BBIF_0540 act not as adhesion protein and possess other functions.

Further experiments are required to determine the receptors of these putative adhesins on Caco-2 cells and to elucidate the mechanism of adhesion.

Expression of BBIF_0295 led to most prominent increase in adhesion of B. bifidum S17. To confirm its role as an adhesin, bbif_0295 was cloned under the control of a synthetic promoter into the plasmid pVG-mCherry carrying a constitutively expressed gene for the fluorescent protein mCherry. A recombinant strain harboring this plasmid (pCW_mCherry_295) and another B. bifidum strain tagged with green fluorescence by pVG-GFP were used in adhesion assays to test adhesion of both strains in competition.

Bacteria expressing bbif_0295 are able to adhere better in comparison to B. bifidum S17/pVG-GFP (Figure 17). This result indicates that BBIF_0295 plays a role in adhesion of this strain and enhance the competitiveness amongst the same strain. Similar to the BopA study, this result demonstrates that the expression platform consisting of the vector

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Discussion

backbone of pMDY23 and the Pgap promoter can be used to generate recombinant bifidobacteria with improved adhesive properties. It has been shown recently that bifidobacteria are able to compete for adhesion sites with pathogenic bacteria (Collado et al. 2006; Candela et al. 2008). Thus, (over)expression of adhesins might improve this probiotic property of bifidobacteria.

Interestingly, expression of none of the putative adhesins led to increased adhesion of B. longum E18 to Caco-2 cells. A plausible explanation why expression of the putative adhesins enhance adhesion of B. bifidum S17 but not B. longum E18 to IECs are differences in EPS. As demonstrated by electron microscopy, B. longum E18 is surrounded by a think continuous layer of EPS whereas B. bifidum S17 produces less EPS which is distributed around cells in small patches. Using TEM pictures, EPS of B. longum E18 has an estimated thickness of about 100 - 150 nm. BBIF_0450, the biggest putative adhesin, has a mass of 208.4 kDa. Comparative analysis (BLAST) of the protein sequence of BBIF_0450 resulted in high homology to ATPase AAA of different bifidobacteria (86- 100 %). ATPases AAA described as conserved proteins, which build hexamer structures, with subunits containing amongst other domains a globular region (Bar-Nun et al., 2012; Meyer et al., 2012). For globular proteins the mass of BBIF_0450 corresponds to a size of approx. 4 nm (Erickson, 2009). It is thus possible that putative adhesins analyzed in this study have too small sizes to penetrate the thick EPS layer of B. longum E18 consequently preventing their functions as adhesins. Whether Tad and sortase-dependent pili, which are structures bigger than 100 nm, act as adhesins to IECs and whether these structures can be expressed to enhance adhesion of bifidobacteria needs to be tested in further experiments.

4.4. Bifidocepin

In a previous study, the subtilisin-like protease BBIF_1681 (bifidocepin) was identified in the predicted proteome of B. bifidum S17. This protease comprises 1355 amino acid residues and is one of the largest proteins encoded by B. bifidum S17. It shows a high degree of similarity to lactocepin of L. paracasei, which has anti-inflammatory activity by selectively degrading pro-inflammatory chemokines of the host (von Schillde et al., 2012; Hörmannsperger et al., 2013).

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Using a His6-tagged version of bifidocepin expressed in different bifidobacteria and detection by Western Blot on different cell fraction clearly demonstrated that the protein is located primary in the cell envelope fraction (Figure 19) and is thus a CEP. This is consistent with the prediction of a signal peptide for protein secretion at the N-terminus and a transmembrane domain at the C-terminus and suggests that the protein has a large extracellular domain and is anchored in the cytoplasmic membrane. The localization of bifidocepin in the cell envelope would allow interactions with the host or host structures. However, we cannot rule out that bifidocepin is posttranslational modified or cleaved autoproteolytic and released in the extracellular milieu. Protease concentrations in the supernatant are maybe below the detection limit of Western Blot, wherefore no signal for presence of protease could be detected in the supernatant fraction (Figure 19). The subtilase S8 domain of bifidocepin is described as Vpr-like (NCBI CDD Conserved Protein Domain Family code cd07474: peptidases S8 subtilisin Vpr-like). The extracellular protease Vpr of Bacillus subtilis ATCC 6633 maturates by posttranslational modifications of the propeptide and proteolytic cleavage of the leader peptide (Corvey et al., 2003). Furthermore, the expression of adhesion and penetration protein of the human pathogen Neisseria meningitides, in E. coli showed that the protein is exported to the surface, processed by an endogenous serine-protease domain, which is the super group of subtilisin-like proteases, and released in the culture supernatant where it mediates adhesion to IECs (Serruto et al., 2003). Recently, another human pathogen Pseudomonas aeruginosa PAO1, causing nosocomial infections, was shown to produce the subtilisin-like SprP which is autocatalytically activated by proteolysis (Pelzer et al., 2015).

A more in-depth sequence analysis revealed that the similarities between lactocepin and bifidocepin are limited to the proteolytic domain, whereas potential substrate recognition sites differ dramatically. Bifidocepin contains two FIVAR (found in various architectures, PF07554) domains, which are mostly present in cell wall associated proteins. FIVAR domains are described as uncharacterized sugar-binding domains. Streptococcus epidermidis strain NCTC11047 was shown to possess proteins with FIVAR domains that bind to fibronectin (Williams et al., 2002). It was thus hypothesized that bifidocepin is a CEP with fibronectin binding capacity. Fibronectin is a component of the ECM, which is part of the connective tissue of the intestinal mucosa underneath the epithelium (Westerlund and Korhonen, 1993; Mao and Schwarzbauer, 2005). Besides fibronectin, ECM contains other proteins including collagen and fibrinogen (Molmenti et al., 1993). In

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Discussion case of a damaged epithelial layer, the ECM is accessible for microbial colonization and infection (Westerlund and Korhonen, 1993) and all ECM proteins were shown to be common targets for bacterial attachment (Vélez et al. 2007). In this context, bifidobacterial binding to ECM may play a role in competition and displacement of pathogens. In this scenario, the FIVAR domain could be the substrate binding domain with the subtilisin domain carrying the enzymatic activity possibly catalyzing proteolytic cleavage of fibronectin. Moreover, one other putative adhesin of B. bifidum S17 contain a domain that potentially interacts with fibronectin (BBIF_0450, PF00041). The gene bbif_0450 is expressed in both growth phases and adhesion to Caco-2 cells with the overexpression strains resulted in an enhanced adhesion (Figure 14).

To test this hypothesis, a range of different extracellular matrix proteins were tested as possible substrates or adhesion sites for bifidocepin (BBIF_1681) and the putative single adhesin BBIF_0450 containing fibronectin binding domain. Neither B. bifidum S17 wt nor recombinant derivatives expressing BBIF_0450 were able to adhere to collagen and fibrinogen under tested conditions (data not shown).

By contrast, B. bifidum S17 showed good adhesion to fibronectin. Surprisingly, expression of BBIF_0450, which contains a fibronectin binding domain, did not increase adhesion to fibronectin. One explanation could be that more than one protein is involved in adhesion to fibronectin as shown for Lactobacillus plantarum 299v that requires two anchorless surface associated glycolytic enzymes for fibronectin binding (Glenting et al. 2013). Another possibility is that experiments were carried out in a way that does not allow for detection of increased adhesion e.g. as a consequence of saturation of binding sites. This is supported by the microscopic examination, which revealed that the amount of added bacteria leads to an almost complete coverage of the fibronectin surface (Figure 20 B). Thus experiments have to be repeated with less bacteria.

Interestingly, overexpression of bifidocepin by B. bifidum S17 resulted in a significant reduction in adhesion to fibronectin. As stated above, bifidocepin is a subtilisin-like protease with a potential fibronectin binding domain. It is thus possible, that fibronectin is indeed a substrate for binding and degradation of fibronectin by bifidocepin, which results in a degradation of the adhesion sites and thus reduced adhesion.

Fibronectinolytic activity was identified as virulence factor in different pathogens, like Pseudomonas aeruginosa or Borrelia burgdorferi (Woods et al., 1981; Russell et al.,

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Discussion

2013). Degradation of fibronectin could be the initiation step for the development of infectious process of the pathogenic bacteria (Wikström and Linde, 1986). However, B. bifidum S17 possess no invasive potential but shows anti-inflammatory activity (Preising et al., 2010). If the epithelial layer is damaged, i.e. during inflammation, pathogenic and commensal bacteria gain access to underlying ECM proteins. Probiotic bacteria compete with pathogens maybe due to the use of same strategies, therefore occupy spaces and prevent invasion of pathogenic bacteria. Additionally, it could be possible that B. bifidum S17 inhibit the inflammatory process concomitant. Our results showed that the CEP bifidocepin plays no role in long-time adhesion of B. bifidum S17 and maybe only mediate temporary adhesion during breakdown of fibronectin structures. Detailed investigations are necessary to proof if this protease is involved in the metabolism by degrading of other substrates that enables B. bifidum S17 to colonize the GIT and maybe produce bioactive compounds through cleavage of fibronectin (Russell et al., 2013). Bifidocepin shows a high degree of sequence similarity to lactocepin of L. paracasei, which was shown to possess anti-inflammatory activity by degrading pro-inflammatory chemokines (von Schillde et al., 2012; Hörmannsperger et al., 2013). Further detailed investigations are necessary to proof the capability of the protease bifidocepin to cleave pro-inflammatory molecules, maybe others than that cleaved by lactocepin, due to different substrate recognition sites.

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Summary

5. Summary

Adhesion of probiotic bacteria plays a role in colonization and persistence of the host GIT. B. bifidum S17 was identified as a strain with high adhesion to IECs (Riedel et al., 2006b; Preising et al. 2010). In this study, possible structures mediating adhesion to IECs are investigated.

Bioinformatic analysis of the genome of B. bifidum S17 revealed the presence of genes encoding EPS biosynthesis, potential proteinaceous surface appendages, seven potential single adhesins, and a subtilisin-like protease.

EPS structures around B. bifidum S17 and B. longum E18 were identified by TEM and SEM. While B. bifidum S17 displayed a compact and patchy EPS layer, B. longum E18, a strain showing only weak adhesion to IECS, has a thick and fuzzy EPS structure.

The predicted tad pili and sortase-dependent pili encoding gene clusters, as well as single adhesin genes were analyzed for expression in exponential and stationary growth by RT- PCR. The results strongly suggest that two sortase-dependent pili encoding gene cluster were organized as operons and are expressed under in vitro conditions. For both clusters, expression levels were higher in exponential growth phase. Additionally, most of the seven single adhesins are higher expressed when bacteria in exponential growth phase. In line with this result, B. bifidum S17 shows higher adhesion to Caco-2 cells in exponential growth phase.

Adhesion assay with recombinant bifidobacteria expressing putative single adhesins were performed to investigate the role of these protein in adhesion to IECs. Expression of all but one adhesin increased adhesion of B. bifidum S17 suggesting that these proteins indeed play a role in adhesion. However, adhesion of B. longum E18 was not increased upon expression of the respective adhesins possibly as a result of the thick EPS layer preventing interaction with the receptors.

A subtilisin-like protease (BBIF_1681, bifidocepin) was identified as cell envelope protease. B. bifidum S17 adheres strongly to fibronectin but adhesion is significantly reduced when bifidocepin was overexpressed suggesting a role in fibronectin binding and degradation.

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Zusammenfassung

6. Zusammenfassung

Die Adhäsion von probiotischen Bakterien spielt eine Rolle in der Beständigkeit und Kolonisierung des gastrointestinalen Trakts des Wirtes. Der kommensale Bakterienstamm B. bifidum S17 wurde als stark adhärenter Stamm an intestinale Epithelzellen identifiziert (Riedel et al., 2006b; Preising et al. 2010). In dieser Studie wurden mögliche Strukturen untersucht, die diesen Effekt bewirken.

Die bioinformatische Analyse des Genoms von B. bifidum S17 zeigten das Vorhandensein von Genen, die für die EPS Biosynthese, potentielle proteinöse Oberflächenanhänge, sieben potentielle Einzel-Adhäsine und eine Subtilisin-ähnliche Protease codieren.

EPS Strukturen, die Zellen von B. bifidum S17 und B. longum E18 umhüllen, wurden durch SEM und TEM identifiziert. Während B. bifidum S17 eine dünne und kompakte EPS Schicht hat, besitzt B. longum E18 schwache Adhäsionseigenschaften an intestinale Epithelzellen und eine dicke und flockige EPS Struktur.

Die Expression der prognostizierten tad Pili und Sortase-abhängige Pili codierenden Gencluster, als auch der sieben Einzel-Adhäsin Genen, wurde während der exponentiellen und stationären Wachstumsphase analysiert. Die Ergebnisse deuten darauf hin, dass zwei Sortase-abhängige Pili codierende Gencluster als Operon organisiert sind und unter in vitro Bedingungen exprimiert werden. Für beide Gencluster war die Expression in der exponentiellen Wachstumsphase höher. Ebenso sind die meisten Gene der sieben Einzel- Adhäsine in der exponentiellen Wachstumsphase stärker exprimiert. In Übereinstimmung mit diesen Ergebnissen konnte eine höhere Adhärenz von B. bifidum S17 an Caco-2 Zellen in der exponentiellen Wachstumsphase gezeigt werden. Indessen konnte die Adhäsion von B. longum E18 durch die Präsenz der Einzel-Adhäsine nicht erhöht werden. Möglicherweise wurde die Interaktion mit Rezeptoren aufgrund der dicken EPS Schicht verhindert.

Die Subtilisin-ähnliche Protease (BBIF_1681, Bifidocepin) wurde als Zellhüllenprotease identifiziert. B. bifidum S17 adhäriert stark an Fibronektin, wobei sich die Adhäsion signifikant reduziert wenn Bifidocepin überexprimiert wird. Dies weist auf eine Rolle in der Bindung und dem Abbau von Fibronektin hin.

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aNote: With kind permission from Springer Science+Business Media: Symbiosis, Exploring the genome sequence of Bifidobacterium bifidum S17 for potential players in host-microbe interactions., 58(1), 2012, 191-200, Westermann, C., Zhurina, D., Baur, A., Shang, W., Yuan, J., & Riedel, C. U., Figure 4 (modified), 5 (modified), 6 (modified), 12 (modified).

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Parts of this dissertation have already been published in:

Westermann, C., Zhurina, D., Baur, A., Shang, W., Yuan, J., & Riedel, C. U. (2012). Exploring the genome sequence of Bifidobacterium bifidum S17 for potential players in host-microbe interactions. Symbiosis, 58(1), 191-200. doi: 10.1007/s13199-012-0205-z.

114

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8. Addendum

8.1. Used enzymes

All enzymes used in this study are listed in the following Table 14.

Table 14: Enzymes and their manufacturers used in this study.

Enzymes Manufacturer

Restriction enzymes XhoI Thermo Fisher Scientific Inc., Rockfort, USA HindIII Thermo Fisher Scientific Inc., Rockfort, USA NdeI Thermo Fisher Scientific Inc., Rockfort, USA BglII Thermo Fisher Scientific Inc., Rockfort, USA

DNA modifying enzymes DNaseI (RNase free) Thermo Fisher Scientific Inc., Rockfort, USA Klenow Fragment MBI Fermentas GmbH, St. Leon-Rot, Germany Phusion hot start polymerase Finnzymes Oy, Espoo, Finland Taq DNA Polymerase Genaxxon BioScience, Ulm, Germany T4-DNA- Ligase Thermo Fisher Scientific Inc., Rockfort, USA

Protein modifying enzymes Lysozyme Roche Diagnostics GmbH, Mannheim, Germany Mutanolysin Sigma Aldrich Chemie GmbH, Steinheim, Germany Proteinase K Roche Diagnostics GmbH, Mannheim, Germany

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8.2. List of oligonucleotides

All oligonucleotides used in this study are listed in the following Table 15.

Table 15: Oligonucleotides used for experiments and cloning work. Restriction sites (italics), stop codon (bold), His-tag sequence (underlined).

Table 15: continued. Name Sequence Amplicon size Purpose [bp] Pili encoding gene cluster BBIF301_fw 5´- GGCACTGCGATTGCGGCCAATG -3´ 510 RT-PCR for bbif_0301 BBIF301_rev 5´- GAAGTAGTTGAAGTAGAATG -3´ 301/302_fw 5´- CGTGACTGGCTTGTTGTATGG-3´ 343 RT-PCR for intergenic region between bbif_0301 and 301/302_rev 5´- AGGTGACCAACGTCAAGTCC-3´ bbif_0302 BBIF302_fw 5´- GAGACGACAGTGGACACTGC -3´ 234 RT-PCR for bbif_0302 BBIF302_rev 5´- CCATCAAATCAGTGGTGCC -3´ 302/303_fw 5´- GTTCACCGTGGCCATCGATG-3´ 700 RT-PCR for intergenic region

between bbif_0302 and 302/303_rev 5´- GCATGATGATGAGCTGTATGC-3´ bbif_0303

cluster 912 BBIF303_fw 5´- GCAGATCCTACCGAAACAGCC -3´ RT-PCR for bbif_0303

im1 BBIF303_rev 5´- GTCATGATGGTGCGCGTGTCATG -3´

f BBIF1648_fw 5´- CGCTGCTGACGCCATCGGCGTCGATC-3´ 529 RT-PCR for bbif_1648 BBIF_1648_rev 5´- CGTGACCGTCGAATACAAGG-3´ 1648/1649_fw 5´- CTTGAACGTGTGTCCAGTCAATTGGTCCGC -3´ RT-PCR for intergenic region cluster 459 between bbif_1648 and 1648/1649_rev 5´- CTCCATCGCAAAGACCAAGGACGACGAG -3´ bbif_1649

fim2 fim2

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Table 15: continued. Name Sequence Amplicon size Purpose [bp]

BBIF1649_fw 5´- GGCTACTGTCATCCTTGTGGAAC -3´ 521 RT-PCR for bbif_1649 BBIF1649_rev 5´- CGTTCCTGAACTTCCTCATCCTG -3´ 1649/1650_fw 5´- CGTTGAGGACGAGATCGTAGATGAGG -3´ RT-PCR for intergenic region

631 between bbif_1649 and 5´- GGTGATGGGCGAGACCTTGGGCTACAAGG -3´ 1649/1650_rev bbif_1650

cluster BBIF1650_fw 5´- GTGGTAGATGGGCAGGTCGACCCCAATC -3´ 424 RT-PCR for bbif_1650 fim2 BBIF1650_rev 5´- GAGCCGTCCTGGGCCTGGAGG -3´ RT_1760fw 5´- GATGTCCACGTGGAATGTGG -3´ 529 RT-PCR for bbif_1760 RT_1760rev 5´- GAGAGACCGACGCTCTTGCC -3´ 1760/1761_fw 5´- CGTCACCGCTCACGGTGGCGGAGAAC -3´ RT-PCR for intergenic region 588 between bbif_1761 and 1760/1761_rev 5´- CCAATGGCAAGGACATCGTGCGTTAC -3´ bbif_1760 BBIF1761_fw 5´- TGAGCGTGTACTCGCCGTTCTTC -3´ 619 RT-PCR for bbif_1761 BBIF1761_rev 5´- CATGTCCGCCTATGACACCTAC -3´ 1761/1762_fw 5´- CGTGCGCAGCATCCAGCACCTTCTGG -3´ RT-PCR for intergenic region

586 between bbif_1761 and 1761/1762_rev 5´- GGGGACTGAAGAACGGCGAGTACACG -3´ bbif_1762

cluster BBIF1762_fw 5´- CGTAGCCTGTGGAAAGATCCTTC -3´ RT-PCR for bbif_1762 500

im3 im3 BBIF1762_rev 5´- GTGAAACCAGCCGTGCCTATTTG -3´ (proximal part)

f TadE_fw 5´- CACCGCCATCCTGTGGCCGAG -3´

161 RT-PCR for bbif_1698 (tadE) TadE_rev 5´- GTCCGTCAACGCCGTGGCGAG -3´ Flp_fw 5´- TCATCCCACACTCAATGACTTC -3´

cluster 290 RT-PCR for bbif_1699 (flp)

ad Flp_rev 5´- ATGAATATCGAGGAGACGATG -3´

t

117

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Table 15: continued. Name Sequence Amplicon size Purpose [bp] BBIF1700_fw 5´- CACACTCAATGCCTTCTTGAC -3´ 282 RT-PCR for bbif_1700 (tadC) BBIF1700_rev 5´- GAATATCGAGGAGACGATG -3´ BBIF1701_fw 5´- GAAATCATGCAACAGCCTGG -3´ 428 RT-PCR for bbif_1701 (tadB) BBIF1701_rev 5´- GATGGTCGTGCTGGCATGGATG -3´ BBIF1702_fw 5´- CTCAACGGTGACGATACGGTC -3´ 467 RT-PCR for bbif_1702 (tadA) BBIF1702_rev 5´- GCACGCGATCCAACGGTCACC -3´

BBIF1703_fw 5´- GATGGCACCGACGACTGCATC -3´ 795 RT-PCR for bbif_1703 (tadZ) BBIF1703_rev 5´- CAAGCCCATGACCATGCCGCGAG -3´

cluster TadV_fw 5´- GATCGAACCTTAGCCACAGCG -3´ 287 RT-PCR for bbif_0835 (tadV) tad TadV_rev 5´- GGATCGCCATCGGGTGCCTGAC -3´

16S_fw 5´- GGAGGTGATCCAGCCGCACCTTC -3´ RT-PCR for bbif_1306 243 (16S ribosomal RNA) 16S_rev 5´- GTAAACGGTGGACGCTGGATGTG -3´

16SrRNA Putative single adhesins Forward primer to control the integration of a single adhesin pCW_cont_fw 5´- GAGTAAACTTGGTCTACAG -3´ - into the vector (binds in the plasmid region), also used for sequencing Reversed primer for sequencing of single adhesin

pCW_cont_rev 5´- GTCTGAGCCGTCGGCGTCCGC-3´ - encoding genes after ligation with the vector

pCW

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Addendum

Table 15: continued. Name Sequence Amplicon size Purpose [bp] Forward primer for Phyp_NdeI_fw 5´- GACTACCATATGTTCTTGAAGACGAAAGGGCC -3 amplification of the promoter Phyper with NdeI restriction site 98 Reversed primer for

Phyp_XhoI_rev 5´- GACCTCGAGTCTCCTTCTACATTTTTAAC -3 amplification of the promoter Phyper with XhoI restriction site

hyper

P BBIF0295_XhoI_fw 5´- GACTACCTCGAGGTGACGTACGCCGCCGATACC -3´ For amplification of bbif_0295 564 with XhoI and HindIII BBIF0295_HindIII_rev 5´- GACAAGCTTTTAATGATGATGATGATGATGGATGTCGCGACG -3´ restriction site, with His6-tag

537 For colony PCR, to control the BBIF0295_cont_rev 5´- CGGACTTCTCCTTGACGGTG -3´ integration of the gene (with pCW_cont_fw) bbif_0295 into the vector

For amplification of bbif_0295 564 with XhoI and BglII restriction BBIF_0295_BglII_rev 5´- GACAGATCTTTAATGATGATGATGATGATGGATGTCGCGACG -3´ (with site, with His6-tag, for cloning BBIF0295_XhoI_fw) into pVG_mCherry

BBIF0295_fw 5´- CGATACCTCGCTTACCGCCAC -3´ RT-PCR for bbif_0295, also 291 used for sequencing of a part BBIF0295_rev 5´- CTTGACGGTGGCCACACGGGT -3´ of the gene

bbif_0295

BBIF0450_XhoI_fw 5´- GACTACCTCGAGATGAGCAAGCAGCCTAACCAG -3´ For amplification of bbif_0450 6018 with XhoI and HindIII BBIF0450_HindIII_rev 5´- GACAAGCTTTCAATGATGATGATGATGATGTGGTCGGTTTGAGGC -3´ restriction site, with His6-tag

bbif_0450

119

Addendum

Table 15: continued. Name Sequence Amplicon size Purpose [bp] For colony PCR, to control the 539 BBIF0450_cont_rev 5´- GCTGGACGATGTCGAAACG -3´ integration of the gene (with pCW_cont_fw) bbif_0450 into the vector

BBIF0450_fw 5´- GTTGACGCAGTTGAGGTCGGAG -3´ 486 RT-PCR for bbif_0450 BBIF0450_rev 5´- GACCAAGCAGCCTAACCAGCG -3´ BBIF0450_seq1 5´- GCATCCATGCGCAAACGGTC -3´ - BBIF0450_seq2 5´- CCGTTGACGCTGCAATACCC -3´ - BBIF0450_seq3 5´- GAACGCACTCGGCAGTTTCC -3´ - BBIF0450_seq4 5´- GCAGAAGCTGCAGTACGACAAG -3´ - BBIF0450_seq5 5´- GCACCGCGAACGAACTGAAG -3´ - Primer for sequencing BBIF0450_seq6 5´- GACGATTCGGCGAAGCTGAC -3´ -

BBIF0450_seq7 5´- CCCGTGACCATCATCTATTC -3´ - BBIF0450_seq8 5´- CTCGGCAACATGCACAACG -3´ -

50 BBIF0450_seq9 5´- GTATCCGCCGACGATTTCAC -3´ - BBIF0450_seq10 5´- CACAAGCGCGTGTATCTG -3´ -

Bbif_04 BBIF0450_seq11 5´- CGATGTCTCGCAGTGGAT -3´ - BBIF0540_XhoI_fw 5´- GACTACCTCGAGATGTATTATTCCAGTGGCAAC -3´ For amplification of bbif_0540 5´- GACAAGCTTTCAATGATGATGATGATGATGGATCGCCCCGTACTCGCG 1917 with XhoI and HindIII BBIF0540_HindIII_rev -3´ restriction site, with His6-tag

For colony PCR, to control the 592 integration of the gene BBIF0540_cont_rev 5´- GTCCTTGTTGAGCCAGTAATAC -3´ (with pCW_cont_fw) bbif_0540 into the vector, bbif_0540 also used for sequencing of a

120

Addendum

Table 15: continued. Name Sequence Amplicon size Purpose [bp] BBIF0540_fw 5´- CAGTGGCAACTATGAAGCGTTC -3´ 536 RT-PCR for bbif_0540 BBIF0540_rev 5´- CAGTACAGCCAGAAGTTGG -3´ BBIF0665_XhoI_fw 5´- GACTACCTCGAGATGAGACCGATGACATTCGC -3´ For amplification of bbif_0665 5´- 1068 with XhoI and HindIII BBIF0665_HindIII_rev GACAAGCTTTCAATGATGATGATGATGATGTAGCATCCTCCGTGACATG - restriction site, with His6-tag 3´ For colony PCR, to control the 821 integration of the gene BBIF0665_cont_rev 5´- CATACAGCACGATGATGTC -3´ bbif_0665 into the vector, (with pCW_cont_fw) also used for sequencing of a part of the gene

BBIF0665_fw 5´- CTGATTGATTTG AGCGACAGC -3´ RT-PCR for bbif_0665 432 BBIF0665_rev 5´- GCTCCAGCGATGAGACGCTGA -3´

bbif_0665

BBIF1426_XhoI_fw 5´- GACTACCTCGAGATGCCTGTGTGCTACAATCATGCC -3´ 846 For amplification of bbif_0665 with XhoI and HindIII BBIF1426_HindIII_rev 5´- GACAAGCTTTTAATGATGATGATGATGATGGATGATTCGGTAGTAC -3´ restriction site, with His6-tag For colony PCR, to control the 505 integration of the gene BBIF1426_cont_rev 5´- CACGTTCAGGGATTCCACAC -3´ bbif_1426 into the vector, (with pCW_cont_fw) also used for sequencing of a

part of the gene BBIF1426_fw 5´- GCTCCTCTTCGGCCTTCTTGG -3´ BBIF1426_rev 5´- CCTGTGTGCTACAATCATGCCTC -3´ 324 RT-PCR for bbif_1426

bbif_1426

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Addendum

Table 15: continued. Name Sequence Amplicon size Purpose [bp] BBIF1538_XhoI_fw 5´-GACTACCTCGAGATGAACCGAAGAATATCTATG -3´ 5289 For amplification of bbif_1538 BBIF1538_HindIII_rev 5´- GACAAGCTTTCAATGATGATGATGATGATGGCGCTTGGCCAGT -3´ with XhoI and HindIII restriction site, with His6-tag

538 For colony PCR, to control the BBIF1538_cont_rev 5´- GACCGTATACCAGATGTCCTC -3´ integration of the gene (with pCW_cont_fw) bbif_1538 into the vector

BBIF1538_fw 5´- GTCGATCAGCGCGTTCGTG -3´ RT-PCR for bbif_1538 377 BBIF1538_rev 5´- CAGACCACCACCATCCCG -3´

bbif_1538 BBIF1769_XhoI_fw 5´- GACTACCTCGAGATGGCCAAGGCGGTGGACACCAG -3´ For amplification of bbif_1769 1572 with XhoI and HindIII BBIF1769_HindIII_rev 5´- GACAAGCTTTCAATGATGATGATGATGATGGTACCAGCCGATTTG -3´ restriction site, with His6-tag For colony PCR, to control the 544 integration of the gene bbif_1769 into the vector, BBIF1769_cont_rev 5´- GCTGATGGTCGTCTGATATGC -3´ (with pCW_cont_fw) also used for sequencing of a part of the gene

BBIF1769_fw 5´- GTGGACACCACGACGTTTTG -3´ RT-PCR for bbif_1769 69 BBIF1769_rev 5´- CTTGTTGAGGCTGATGCCTTTG -3´ 549

bbif_17

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Addendum

Table 15: continued. Name Sequence Amplicon size Purpose [bp] Subtilisin like protease bifidocepin

BBIF1681_XhoI_fw 5´- GACTACCTCGAGATGGCCAATAAGCAATGGCCTCGCTGG -3´ For amplification of bbif_1681 4089 with XhoI and HindIII 5´- TACAAGCTTCTAATGATGATGATGATGATGGCGACGCACCGCGTCACG BBIF1681_HindIII_rev restriction site, with His6-tag -3´ For colony PCR, to control the 2338 integration of the gene BBIF1681_cont_rev 5´- GCTCACCGCGGATCGCAAGACC -3´ bbif_1681 into the vector, (with pCW_cont_fw) also used for sequencing of a part of the gene BBIF1681_seq1 5´- GAAGAAGCCGTTCTTGTGCAGC -3´ - BBIF1681_seq2 5´- GCATCATCCTCCTGGGTGAC -3´ - Primer for sequencing BBIF1681_seq3 5´- GCGTAGTTCACGGAGTACTGGC -3´ -

bbif_1681

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Addendum

8.3. Chemicals and manufacturers

All used chemicals with their corresponding manufacturers are listed in the following Table 16.

Table 16: Chemicals and manufacturers.

Table 16: continued. Chemical/reagent Manufacturer 1 kb DNA ladder GeneRulerTM MBI Fermentas GmbH, St. Leon-Rot, Germany Acetic acid (100 % pure) AppliChem GmbH, Darmstadt, Germany Actilight® Fructo-oligosaccharides Syral – Beghin Meiji, Marckolsheim, France Agar Difco Laboratories, Detroit, USA Agarose NEEO ultra-quality Carl Roth GmbH & Co., Karlsruhe, Germany

Ammonium acetate (NH4CH3CO2) Sigma Aldrich Chemie GmbH, Steinheim, Germany

Ammonium chloride (NH4Cl) Sigma Aldrich Chemie GmbH, Steinheim, Germany

Ammonium citrate dibasic ((NH4)2C6H4O7) Sigma Aldrich Chemie GmbH, Steinheim, Germany

Ammonium sulfate (NH4)2SO4) Sigma Aldrich Chemie GmbH, Steinheim, Germany Ampicillin sodium salt Carl Roth GmbH & Co., Karlsruhe, Germany AnaeroGenTM Thermo Scientific, Rockfort, USA Ammoniumperoxodisulfat (APS) Carl Roth GmbH & Co., Karlsruhe, Germany BactoTM Tryptone Becton Dickinson GmbH, Heidelberg, Germany BactoTM Yeast extract Becton Dickinson GmbH, Heidelberg, Germany Bovine serum albumin (BSA) Thermo Fisher Scientific Inc., Rockford, USA Bromphenol blue Merck KGaA, Darmstadt, Germany Cacodylate buffer (0.2 M, pH 7) Biotang Inc., Lexington, USA

Carbon dioxide (CO2) MTI Industriegase, Neu-Ulm, Germany

Calcium chloride (CaCl2) Merck KGaA, Darmstadt, Germany Chloroform J. T. Baker, Griesheim, Germany chloroform/isoamylalcohol AppliChem GmbH, Darmstadt, Germany Chloramphenicol Merck KgaA, Darmstadt, Germany

Citric acid monohydrate (H2O C6H8O7 x H2O) Sigma Aldrich Chemie GmbH, Steinheim, Germany Crystal violet Fluka AG, Buchs, Switzerland Collagen, rat tail (#11179179001, Version 15) Roche, Basel, Switzerland Desoxy-nucleotides triphosphates (dNTPs) MBI Fermentas GmbH, St. Leon-Rot, Germany Dimethyl sulfoxide (DMSO, 99.8 % pure) Carl Roth GmbH & Co., Karlsruhe, Germany

Dipotassium phosphate (K2HPO4) Merck KGaA, Mannheim, Germany

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Addendum

Table 16: continued. Chemical/reagent Manufacturer Di-sodium ethylendiaminetetraacetate Carl Roth GmbH & Co., Karlsruhe, Germany dehydrate (EDTA) Di-sodium hydrogenphosphate heptahydrogen Merck KGaA, Mannheim, Germany

(Na2HPO4 x 7 H2O) Dithiothreitol (DTT) Carl Roth GmbH & Co., Karlsruhe, Germany dNTPs (10 mM) Life Technologies, Darmstadt, Germany Dulbecco´s Modified Eagle´s Medium (DMEM) Sigma Aldrich Chemie GmbH, Steinheim, Germany Epon 812 Fluka AG, Buchs, Switzerland Erythromycin Sigma Aldrich Chemie GmbH, Steinheim, Germany Ethanol (99 % pure) Sigma Aldrich Chemie GmbH, Steinheim, Germany Ethidium bromide Carl Roth GmbH & Co., Karlsruhe, Germany Fetal calf serum (FCS) Biochrom AG, Berlin, Germany Fibrinogen from bovine plasma (#1001621516) Sigma Aldrich Chemie GmbH, Steinheim, Germany Fibronectin (#356008) BD Difco Laboratories, Detroit, USA Glass beads (0.1 mm) Carl Roth GmbH & Co., Karlsruhe, Germany Glutaraldehyde Agar Scientific Ltd., Stanstead, UK Glycerol (99 % pure) Sigma Aldrich Chemie GmbH, Steinheim, Germany Glycine Sigma Aldrich Chemie GmbH, Steinheim, Germany Hoechst 33342 trihydrochloride trihydrate Life Technologies, Darmstadt, Germany Hydrochloric acid (HCl) 37 % Merck KGaA, Darmstadt, Germany Imidazole Carl Roth GmbH & Co., Karlsruhe, Germany

Iron(II) Sulfate Heptahydrate (FeSO4 x 7 H2O) Sigma Aldrich Chemie GmbH, Steinheim, Germany Isoamyl alcohol (99 % pure) Fluka Chemikalien, Buchs, Switzerland Isopropanol (99 % pure) Sigma Aldrich Chemie GmbH, Steinheim, Germany Lactobacilli de Man-Rogosa-Sharp Medium BD Difco Laboratories, Detroit, USA L-cystein hydrochloride monohydrate Merck KgaA, Darmstadt, Germany Lead citrate Atlanta Chemie, Heidelberg, Germany L-glutamine Sigma Aldrich Chemie GmbH, Steinheim, Germany Lysine Sigma Aldrich Chemie GmbH, Steinheim, Germany

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Addendum

Table 16: continued. Chemical/reagent Manufacturer

Magnesium chloride (MgCl2) Carl Roth GmbH & Co., Karlsruhe, Germany

Magnesium sulfate heptahydrate (MgSO4 x 7 Sigma Aldrich Chemie GmbH, Steinheim,

H2O) Germany

Manganese(II) Sulfate Hydrate (MnSO4 x H2O) Sigma Aldrich Chemie GmbH, Steinheim, Germany β-mercaptoethanol Carl Roth GmbH & Co., Karlsruhe, Germany

Methanol (H3COH) Sigma-Aldrich Chemie GmbH, Steinheim, Germany

Mouse anti-His6 IgG1 monoclonal antibodies Roche Diagnostics GmbH, Mannheim, (Catalog # 11922416001) Germany Mowiol® 4-88 Sigma Aldrich Chemie GmbH, Steinheim, Germany N-2-Hydroxyetylpiperazine-N´-2-Ethanesulfonic Carl Roth GmbH & Co., Karlsruhe, Germany acid (HEPES) Nitrogen (liquid) VWR International GmbH, Darmstadt, Germany Non-essential amino acids (NEEA, 100 x) PAA Laboratories GmbH, Pasching, Austria Page RulerTM Prestained Protein Ladder MBI Fermentas GmbH, St. Leon-Rot, Germany Paraformaldehyde Th. Geyer GmbH & Co. KG, Renningen, Germany Penicillin-streptomycin solution PAA Laboratories GmbH, Pasching, Austria Peptone Becton Dickinson GmbH, Heidelberg, Germany Phosphate bufferd saline (PBS) PAA Laboratories GmbH, Pasching, Austria Platinum-carbon Sigma Aldrich Chemie GmbH, Steinheim, Germany Polyacrylamid (Rothiphorese Gel 30) Carl Roth GmbH & Co., Karlsruhe, Germany Polysorbate (Tween®) 20 Fluka by Carl Roth, Karlsruhe, Germany Potassium chloride (KCl) Sigma Aldrich Chemie GmbH, Germany

Potassium dihydrogen orthophosphate (KH2PO4) Merck KgaA, Darmstadt, Germany

Pyruvic acid (CH3COCOOH, 98 %) Sigma Aldrich Chemie GmbH, Steinheim, Germany Rabbit anti-mouse polyclonal antibodies, horse Dako Deutschland GmbH, Hamburg, Germany radish peroxidase coupled (Catalog # P0260) RevertAidTM Premium First Strand cDNA MBI Fermentas GmbH, St. Leon-Rot, Germany Synthesis Kit

RNase-free water (H2O) Life Technologies, Darmstadt, Germany Roti®-Aqua-Phenol (acidic phenol) Carl Roth GmbH & Co., Karlsruhe, Germany Roti®-Phenol Carl Roth GmbH & Co., Karlsruhe, Germany

Roti®-Phenol/Chloroform/Isoamyl alcohol Carl Roth GmbH & Co., Karlsruhe, Germany

126

Addendum

Table 16: continued. Chemical/reagent Manufacturer (25:24:1) Ruthenium red Sigma Aldrich Chemie GmbH, Steinheim, Germany

Sodium acetate dehydrate (CH3COONa) Fluka by Carl Roth, Karlsruhe, Germany

Sodium bicarbonate (NaHCO3) Sigma Aldrich Chemie GmbH, Steinheim, Germany

Sodium carbonate (Na2CO3) Sigma Aldrich Chemie GmbH, Steinheim, Germany Sodium chloride (NaCl) Fluka by Carl Roth, Karlsruhe, Germany Sodium dodecyl sulfate (SDS) Carl Roth GmbH & Co., Karlsruhe, Germany Sodium hydroxide (NaOH) AppliChem GmbH, Darmstadt, Germany Sodium hydrogenphosphate dihydrogen Carl Roth GmbH & Co., Karlsruhe, Germany

(Na2HPO4 x 2H2O) Spectinomycin Sigma Aldrich Chemie GmbH, Steinheim, Germany Sucrose Fluka AG, Buchs, Switzerland Tetramethylethylenediamine (TEMED) Carl Roth GmbH & Co., Karlsruhe, Germany Tris(hydroxymethyl)-aminomethan (Tris) USB Corporation, Cleveland, USA Tris-(2-Carboxyethyl)phosphine hydrochloride Sigma Aldrich Chemie GmbH, Steinheim, (Tris-HCl) Germany

Tri-sodium citrate dehydrate (C6H5O7Na3 x 2 H2O) AppliChem GmbH, Darmstadt, Germany Triton X-100 Sigma Aldrich Chemie GmbH, Steinheim, Germany Trypsin-EDTA Sigma Aldrich Chemie GmbH, Steinheim, Germany Tryptone BD Difco Laboratories, Detroit, USA Tween 20 Thermo Fisher Scientific Inc., Rockfort, USA Tween 80 Thermo Fisher Scientific Inc., Rockfort, USA Yeast extract BD Difco Laboratories, Detroit, USA

127

Abbreviations

9. Abbreviations

Δ gene deletion

% per cent

°C degree Celsius

µ micro (10-6) xg times gravity (9.81 m/s2)

Å Ångström (1.0 x 10-10 meters)

A ampere aa amino acids approx. approximately

APS Ammoniumperoxodisulfat

ATCC American Type Culture Collection

B. Bifidobacterium

BCA bicinchominic acid bp basepair

BSA bovine serum albumin

CEP cell envelope protease cfu colony forming units

CmR chloramphenicol resistance

CO2 carbon dioxide ds double stranded

DMEM Dulbecco´s Modified Eagle´s Medium

DMSO dimethylsulfoxide

DSMZ German Collection of Microorganisms and Cell Cultures

DNA deoxyribonucleic acid dNTP deoxyribose nucleoside triphosphate

E. Escherichia

128

Abbreviations e.g. Latin `exemplȋ grȃtiȃ´; English: for example et al. Llatin `et alii´; English: and others

EDTA ethylendiaminetetraacetic acid

EmrR Erythromycin resistance

EPS exopolysaccharides (deutsch: Exopolysaccharide)

FCS fetal calf serum

Fig. Figure fim fimbria-associated adhesion g/l gram per liter h hour

HCl hydrochloric acid

HEPES 2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid

His6 Histidine tag with six histidine

HMO human milk oligosaccharide

HRP horseradish peroxidase i.e. Latin `id est´, English: that is

IEC intestinal epithelial cells (deutsch: intestinale Epithelzellen)

Ig immunglobulin k kilo (103) kb Kilo (103) basepair

L. Lactobacillus

LB Lysogeny Broth

Ltd. Limited m milli (10-3)

M molar

Mbp mega (106) base pairs min minute

MRSC Man-Rogosa-Sharpe medium containing 0.5 g/l L-cysteine hydrochloride

129

Abbreviations

NaCl sodium chloride

NCBI Nactional Centre for Biotechnology Information

NEAA non-essential amino acids nm nanometer (10-9)

OD600 optical density at 600 nm o/N over night

PCR polymerase chain reaction

PBS phosphate buffered saline

Pfam Protein families

® registered trademark rpm rounds per minute

RNA ribonucleic acid rRNA ribosomal ribonucleic acid

RT room temperature

RT-PCR reverse transcription - polymerase chain reaction

SDS sodium dodecylsulfate

SDS-PAGE sodium dodecylsulfate polyacrylamide gel electrophoresis

SEM scanning electron microscopy (deutsch: Rasterelektronenmikrsokopie)

SP signal peptide sp. species

SpcR spectinomycin resistance subsp. subspecies tad tight adherence

TM transmembrane

TEM transmission electron microscopy (deutsch: Transmissionselektronenmikroskopie)

TIGR The Institute for Genomic Research

TMHMM Transmembrane prediction using Hidden Markov Model

130

Abbreviations

Tris Tris(hydroxymethyl)-aminomethan

TM `trademark´

U unit

UV ultraviolet

V volt v/v volume per volume wt wildtype w/v weight per volume

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Acknowledgment

10. Acknowledgment

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Acknowledgment

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Curriculum Vitae

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Ich versichere hiermit, dass ich die Arbeit ohne fremde Hilfe und ausschließlich unter Verwendung der angegebenen Quellen angefertigt habe. Alle Ausführungen, die wörtlich oder sinngemäß übernommen wurden sind als solche gekennzeichnet.

Ulm, den 28.07.2015

Christina Westermann

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