analysis of potential host-colonization factors in

Analysis of potential host-colonization factors in Bifidobacterium 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 probiotics .............................................................................................. 10

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

1.3.2. Probiotic 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 enzymes .............................................................. 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. Subtilisin 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

8

1. Introduction

1.1. General features of the genus Bifidobacterium

Bifidobacteria belong to the family Bifidobacteriaceae and the phylum Actinobacteria,

which are one of the largest taxonomic units in the domain Bacteria (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 gastrointestinal tract (GIT; Turroni et al.,

2012). Bifidobacteria are found to stably colonize the GIT of mammals, 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 carbohydrates (de

Vries and Stouthamer, 1967). The key enzyme 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

Introduction

9

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 microorganisms 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 gut microbiota 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

Introduction

10

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., Bacteroides 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

Introduction

11

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).

Introduction

12

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 carbohydrate 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).

Introduction

13

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

microorganism adhesion structures compared than host receptors binding to these bacterial

Introduction

14

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).

Introduction

15

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 coagulation 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 binding

site 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

Introduction

16

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.)

Introduction

17

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

Introduction

18

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. casein

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

Introduction

19

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 (fimbria-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

Introduction

20

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

Introduction

21

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 hydrolase, 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 translocase, 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.

Introduction

22

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).

Introduction

23

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.

Materials and Methods

24

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.


25

Table 1: Bacterial strains used in this study.

Strain Characteristics Reference

Escherichia coli

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

InvitrogenTM, Darmstadt, Germany

E. coli ET12567::p16S_ParaB_bbif0710

Genomic integration of p16S_ParaB_bbif0710 into 16S-rDNA, CMR, ErmR, bbif_0710 encodes for a methyltransferase of B. bifidum S17

Brancaccio, 2013b

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-K1

Tn5 disrupted genes necessary for exopolysaccharide synthesis

Drepper, RC 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.


26

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: continued.

Plasmid Relevant characteristics Reference

pMDY23_Pgap pMDY23 E. coli-Bifidobacterium shuttle vector, expression of gusA driven by the promoter of glyceraldehydes-3-phosphate dehydrogenase gene bbif_0612 (Pgap) of B. bifidum S17, SpcR

Grimm et al., 2014

pVG-GFP pMDY23, expression of the green fluorescent protein (GFP) under control of constitutive Pgap promoter, SpcR

Grimm et al., 2014

pVG-mCherry pMDY23, expression of the fluorescent protein mCherry under control of constitutive Pgap promoter, SpcR

Grimm et al., 2014

pCW pMDY23, containing the constitutive expressed Pgap

promoter, SpcR this work

pCW_17 pMDY23, expression of the single adhesin bbif_0017 under control of constitutive Pgap promoter, SpcR

this work


this work


this work


this work


this work


this work

Plasmids used in this study.


27

Table 2: continued.

Plasmid Relevant characteristics Reference


this work

pCW_1681 pMDY23, expression of the serine-like protease bbif_1681 under control of constitutive Pgap promoter, SpcR

this work

pCW_mCherry_295 pMDY23, expression of the single adhesin bbif_0295 under control of synthetic constitutive Phyper promoter, expression of the fluorescent protein mCherry under the constitutive Pgap promoter, SpcR

this work

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 trypsin-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


28

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

transferred to 25 cm2 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:

𝑐 = 𝑁𝑐 ∙ 𝐷𝐹 ∙ 𝐾

𝑁𝑆

Formula 1: Calculation of the cell concentration. With c: cell concentration (cell/ml), Nc: number of counted cells in all squares, DF: dilution factor, K: chamber factor (1 x 104 µl-1), and NS number of counted squares.


29

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

bifidobacteria were adjusted to an OD600 of 1 (i.e. approx. 5 x 107 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


30

inoculum was adjusted to OD600 0.1 (i.e. 5 x 106 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


31

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.).


32

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.).


33

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.


34

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

35 Annealing 55-60 °C 30s

Extension 72 °C 30 s/kb

Final extension 72 °C 10 min 1


35

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

Component

Phusion system Genaxxon system

Final concentration

Volume Final concentration Volume

ddH2O - Ad to 20 µl - Ad to 25 µl

Reaction buffer 1X1 4.0 µl 1X2 2.5 µl

dNTPs (10 mM3/2mM4)

400 µm each 0.4 µl 200 µm each 2.5 µl

Forward primer (10 pmol/µl)

0.2 µM 0.4 µl 0.2 µM 0.5 µl

Reverse primer (10 pmol/µl)

0.2 µM 0.4 µl 0.2 µM 0.5 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.).


36

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


Initial Denaturation 95 °C 30 s 1


35 Annealing 45-75 °C 30 s

Extension 72 °C 30 s/kb


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.


Initial Denaturation 95 °C 2 min 1


25 Annealing 56 °C 30 s

Extension 72 °C 1 min



37

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 Ligase (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


38

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


39

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.

𝐴260 = 𝜀 ∙ 𝑐 ∙ 𝑑

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


40

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


41

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).


42

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.


43

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


44

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


45

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.

Components Amount

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


46

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.


47

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


48

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


49

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 amino acid 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.

Results

50

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).

Results

51

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

EPS

gen

e cl

ust

er I

0049 glycosyltransferase

0050 hypothetical protein

0051 dTDP-glucose 4,6-dehydratase


0053 dTDP-4-dehydrorhamnose 3,5-epimerase/reductase

0054 Transposase


0056 Transposase

0057 Transposase

0058 Glucose-1-phosphat thymidyl transferase

0059 sugar ABC transporter permease

0060 sugar ABC transporter ATP-binding protein


0062 lipopolysaccharide biosynthesis protein

0063 glycosyltransferase

EPS

gen

e

clu

ste

r II

0393 UDP Galactosephosphotransferase

0394 transporter protein



Results

52

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.

Results

53

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

Results

54

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.

Results

55

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

BBIF_

SignalP 3.0 SignalP 4.1

D-score1 Signal

peptide

predicted

cleavage

site2

D-score1 Signal

peptide

predicted

cleavage

site2

0017 0.45 No

0.231 No

0295 0.45 No

0.215 No

0450 0.383 Yes aa 35-36:

VIA*LV 0.351 No

0540 0.253 No

0.150 No

0567 0.158 No

0.234 No

0665 0.398 Yes aa 28-29:

AVA*VV 0.277 No

1426 0.669 Yes aa 39-40:

ALA*GS 0.633 Yes aa 25-26:

AAA*AV

1538 0.605 Yes aa 39-40:

HFA*GM 0.471 Yes aa 45-46:

AYA*VE

1769 0.623 Yes aa 36-37:

GVA*AR 0.540 Yes aa 36-37:

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.

Results

56

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

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

BBIF_ pSORT v2.0 pSORTb v3.0.2

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

1538 0.000 0.308 0.000 Bacterial membrane 0.00 0.12 5.62 4.26 Unknown (possible

multiple localization sites)

1769 0.000 0.363 0.000 Bacterial outside 0.00 3.33 3.33 3.33 Unknown

Results

57

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,

Results

58

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 catalytic triad (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.

Structure Gene (putative) function Reference

EPS

bbif_0049 protection,

immunomodulation,

colonization

Ruas-Madiedo et al., 2010;

Hidalgo-Cantabrana et al.,

2014;

Fanning et al., 2012a;

Alp and Aslim, 2010

bbif_0050

bbif_0051

bbif_0052

bbif_0053

bbif_0054

bbif_0055

bbif_0056

bbif_0057

bbif_0058

Overview of bifidobacterial structures potentially involved in adhesion to host structures.

Results

59


Structure Gene (putative) function Reference

EPS bbif_0059

bbif_0060

bbif_0061

bbif_0062

bbif_0063

bbif_0393

tad pili tadV (bbif_0835) adhesion,

immunomodulation

O'Connell Motherway et al.,

2011 tadZ (bbif_1703)

tadA (bbif_1702)

tadB (bbif_1701)

tadC (bbif_1700)

flp (bbif_1699)

tadE (bbif_1698)

fim1 pili bbif_301 adhesion,

immunomodulation

Foroni et al., 2011;

Turroni et al., 2013 bbif_302

bbif_303


immunomodulation



bbif_1650


immunomodulation



bbif_1762

cell surface

proteins

bbif_0295 adhesion Liu et al., 2015;

O'Flaherty and

Klaenhammer, 2010;

Bateman et al., 2005;

Gagic et al., 2013;

Henderson et al., 2011;

Katerov et al., 2000

bbif_0450

bbif_0540

bbif_0665

bbif_1426

bbif_1538

bbif_1769

subtilisin like

protease

bbif_1681 immunomodulation,

adhesion, ECM

interaction

von Schillde et al., 2012

Williams et al., 2002

Results

60

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.

Results

61

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

Results

62

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).

Results

63

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

Results

64

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

Results

65

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.

Results

66

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

Results

67

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

Results

68

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.

Results

69

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

Results

70

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).

Results

71

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

(B)

mC

herr

y

GF

P

0

1

2

3

4

5

pe

rc

en

t a

re

a [

%]

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).

(A)

Results

72

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.

Results

73

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. BBIF_1681 was detected through anti-His6 antibody. Marker: PageRulerTM 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

Results

74

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).

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.

S17 wt

S17/pCW_1681

(A) (B)

Discussion

75

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

Discussion

76

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.

Discussion

77

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

Discussion

78

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

Discussion

79

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.

Discussion

80

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

Discussion

81

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

Discussion

82

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 protein family 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

Discussion

83

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

Discussion

84

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).

Discussion

85

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

serine protease 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

Discussion

86

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.,

Discussion

87

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.

Summary

88

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.

Zusammenfassung

89

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.

List of Literature

90

7. List of Literature

Adlerberth, I., & Wold, A. E. (2009). Establishment of the gut microbiota in Western infants. Acta

Paediatrica, 98(2), 229-238. doi: 10.1111/j.1651-2227.2008.01060.x.

Allen, S., Wareham, K., Wang, D., Bradley, C., Sewell, B., Hutchings, H., Harris, W., Dhar, A.,

Brown, H., & Foden, A. (2013). A high-dose preparation of lactobacilli and bifidobacteria in the

prevention of antibiotic-associated and Clostridium difficile diarrhoea in older people admitted

to hospital: a multicentre, randomised, double-blind, placebo-controlled, parallel arm trial

(PLACIDE). Health Technology Assessment, 17(57), 1-140. doi: 10.3310/hta17570.

Alp, G., & Aslim, B. (2010). Relationship between the resistance to bile salts and low pH with

exopolysaccharide (EPS) production of Bifidobacterium spp. isolated from infants feces and

breast milk. Anaerobe, 16(2), 101-105. doi: 10.1016/j.anaerobe.2009.06.006.

Anantharaman, V., & Aravind, L. (2003). Evolutionary history, structural features and biochemical

diversity of the NlpC/P60 superfamily of enzymes. Genome Biology, 4(2), R11. doi:10.1186/gb-

2003-4-2-r11.

Apostolou, E., Kirjavainen, P. V., Saxelin, M., Rautelin, H., Valtonen, V., Salminen, S. J., &

Ouwehand, A. C. (2001). Good adhesion properties of probiotics: a potential risk for

bacteremia? FEMS Immunology & Medical Microbiology, 31(1), 35-39.

doi: http://dx.doi.org/10.1111/j.1574-695X.2001.tb01583.x.

Atzingen, M. V., Gómez, R. M., Schattner, M., Pretre, G., Gonçales, A. P., de Morais, Z. M.,

Vasconcellos, S. A., & Nascimento, A. L. (2009). Lp95, a novel leptospiral protein that binds

extracellular matrix components and activates e-selectin on endothelial cells. Journal of

Infection, 59(4), 264-276. doi: 10.1016/j.jinf.2009.07.010.

Åvall-Jääskeläinen, S., Lindholm, A., & Palva, A. (2003). Surface display of the receptor-binding

region of the Lactobacillus brevis S-layer protein in Lactococcus lactis provides nonadhesive

lactococci with the ability to adhere to intestinal epithelial cells. Applied and Environmental

Microbiology, 69(4), 2230-2236. doi: 10.1128/AEM.69.4.2230-2236.2003.

Ayache, J., Beaunier, L., Boumendil, J., Ehret, G., & Laub, D. (2010). Sample Preparation

Handbook for Transmission Electron Microscopy: Techniques. Volume 2, Springer Science &

Business Media. ISBN 978-1-4419-6087-0.

Backhed, F., Ley, R. E., Sonnenburg, J. L., Peterson, D. A., & Gordon, J. I. (2005). Host-bacterial

mutualism in the human intestine. Science, 307(5717), 1915-1920. doi: 307/5717/1915.

Badel, S., Bernardi, T., & Michaud, P. (2011). New perspectives for Lactobacilli

exopolysaccharides. Biotechnology Advances, 29(1), 54-66.

doi: 10.1016/j.biotechadv.2010.08.011.

Badylak, S. F., Freytes, D. O., & Gilbert, T. W. (2009). Extracellular matrix as a biological scaffold

material: structure and function. Acta Biomaterialia, 5(1), 1-13.

doi: 10.1016/j.actbio.2008.09.013.

Balakirev, E. S., & Ayala, F. J. (2003). Pseudogenes: are they “junk” or functional DNA? Annual

Review of Genetics, 37(1), 123-151. doi: 10.1146/annurev.genet.37.040103.103949.

Balestrino, D., Hamon, M. A., Dortet, L., Nahori, M. A., Pizarro-Cerda, J., Alignani, D.,

Dussurget, O., Cossart, P., & Toledo-Arana, A. (2010). Single-cell techniques using

List of Literature

91

chromosomally tagged fluorescent bacteria to study Listeria monocytogenes infection processes.

Applied and Environmental Microbiology, 76(11), 3625-3636. doi: 10.1128/AEM.02612-09.

Baquero, F., & Nombela, C. (2012). The microbiome as a human organ. Clinical Microbiology and

Infection, 18(4), 2-4. doi: 10.1111/j.1469-0691.2012.03916.x.

Bar-Nun, S., & Glickman, M. H. (2012). Proteasomal AAA-ATPases: structure and function.

Biochimica Et Biophysica Acta (BBA)-Molecular Cell Research, 1823(1), 67-82.

doi: 10.1016/j.bbamcr.2011.07.009.

Bateman, A., Birney, E., Durbin, R., Eddy, S. R., Finn, R. D., & Sonnhammer, E. L. L. (1999).

Pfam 3.1: 1313 multiple alignments and profile HMMs match the majority of proteins. Nucleic

Acids Research, 27(1), 260-262. doi: 10.1093/nar/27.1.260.

Bateman, A., Holden, M. T., & Yeats, C. (2005). The G5 domain: a potential N-acetylglucosamine

recognition domain involved in biofilm formation. Bioinformatics, 21(8), 1301-1303.

doi: 10.1093/bioinformatics/bti206.

Beckmann, C., Waggoner, J. D., Harris, T. O., Tamura, G. S., & Rubens, C. E. (2002).

Identification of novel adhesins from Group B streptococci by use of phage display reveals that

C5a peptidase mediates fibronectin binding. Infection and Immunity, 70(6), 2869-2876.

doi: 10.1128/IAI.70.6.2869-2876.2002.

Beissinger, M., & Buchner, J. (1998). How chaperones fold proteins. Biological Chemistry, 379(3),

245-259. PMID: 9563819.

Bendtsen, J. D., Nielsen, H., von Heijne, G., & Brunak, S. (2004). Improved prediction of signal

peptides: SignalP 3.0. Journal of Molecular Biology, 340(4), 783-795. doi:

10.1016/j.jmb.2004.05.028.

Bernard, C. S., Bordi, C., Termine, E., Filloux, A., & de Bentzmann, S. (2009). Organization and

PprB-dependent control of the Pseudomonas aeruginosa tad Locus, involved in Flp pilus

biology. Journal of Bacteriology, 191(6), 1961-1973. doi: 10.1128/JB.01330-08.

Bernet, M. F., Brassart, D., Neeser, J. R., & Servin, A. L. (1993). Adhesion of human

bifidobacterial strains to cultured human intestinal epithelial cells and inhibition of

enteropathogen-cell interactions. Applied and Environmental Microbiology, 59(12), 4121-4128.

doi: 0099-2240/93/124121-08$02.00/0.

Bertani, G. (1951). Studies on lysogenesis. I. The mode of phage liberation by lysogenic

Escherichia coli. Journal of Bacteriology, 62(3), 293-300. PMID: 14888646.

Biasucci, G., Rubini, M., Riboni, S., Morelli, L., Bessi, E., & Retetangos, C. (2010). Mode of

delivery affects the bacterial community in the newborn gut. Early Human Development, 86(1),

13-15. doi: 10.1016/j.earlhumdev.2010.01.004.

Bode, L., & Jantscher-Krenn, E. (2012). Structure-function relationships of human milk

oligosaccharides. Advances in Nutrition, 3(3), 383-391. doi: 10.3945/an.111.001404.

Bordoni, A., Amaretti, A., Leonardi, A., Boschetti, E., Danesi, F., Matteuzzi, D., Roncaglia, L.,

Raimondi, S., & Rossi, M. (2013). Cholesterol-lowering probiotics: in vitro selection and in

vivo testing of bifidobacteria. Applied Microbiology and Biotechnology, 97(18), 8273-8281.

doi: 10.1007/s00253-013-5088-2.

Bottacini, F., Medini, D., Pavesi, A., Turroni, F., Foroni, E., Riley, D., Giubellini, V., Tettelin, H.,

van Sinderen, D., & Ventura, M. (2010). Comparative genomics of the genus Bifidobacterium.

Microbiology, 156(11), 3243-3254. doi: 10.1099/mic.0.039545-0.

List of Literature

92

Bouchara, J., Tronchin, G., Larcher, G., & Chabasse, D. (1995). The search for virulence

determinants in Aspergillus fumigatus. Trends in Microbiology, 3(8), 327-330.

doi: 10.1016/S0966-842X(00)88965-9.

Brancaccio, V. F., Zhurina, D. S., & Riedel, C. U. (2013a). Tough nuts to crack: site-directed

mutagenesis of bifidobacteria remains a challenge. Bioengineered, 4(4), 197-202.

doi: 10.4161/bioe.23381.

Brancaccio, V. F. (2013b). Characterization of Host Colonization Factors of Bifidobacterium

bifidum S17 and Development of Molecular Tools. Doctoral Thesis. University of Ulm, Ulm,

Germany.

Buffie, C. G., & Pamer, E. G. (2013). Microbiota-mediated colonization resistance against

intestinal pathogens. Nature Reviews Immunology, 13(11), 790-801. doi: 10.1038/nri3535.

Candela, M., Bergmann, S., Vici, M., Vitali, B., Turroni, S., Eikmanns, B. J., Hammerschmidt, S.,

& Brigidi, P. (2007). Binding of human plasminogen to Bifidobacterium. Journal of

Bacteriology, 189(16), 5929-5936. doi: 10.1128/JB.00159-07.

Candela, M., Biagi, E., Centanni, M., Turroni, S., Vici, M., Musiani, F., Vitali, B., Bergmann, S.,

Hammerschmidt, S., & Brigidi, P. (2009). Bifidobacterial enolase, a cell surface receptor for

human plasminogen involved in the interaction with the host. Microbiology, 155(10), 3294-

3303. doi: 10.1099/mic.0.028795-0.

Candela, M., Centanni, M., Fiori, J., Biagi, E., Turroni, S., Orrico, C., Bergmann, S.,

Hammerschmidt, S., & Brigidi, P. (2010). DnaK from Bifidobacterium animalis subsp. lactis is

a surface-exposed human plasminogen receptor upregulated in response to bile salts.

Microbiology, 156(6), 1609-1618. doi: 10.1099/mic.0.038307-0.

Candela, M., Perna, F., Carnevali, P., Vitali, B., Ciati, R., Gionchetti, P., Rizzello, F., Campieri,

M., & Brigidi, P. (2008). Interaction of probiotic Lactobacillus and Bifidobacterium strains with

human intestinal epithelial cells: adhesion properties, competition against enteropathogens and

modulation of IL-8 production. International Journal of Food Microbiology, 125(3), 286-292.

doi: 10.1016/j.ijfoodmicro.2008.04.012.

Cheng, J., Palva, A. M., de Vos, W. M., & Satokari, R. (2013). Contribution of the intestinal

microbiota to human health: from birth to 100 years of age. Current Topics in Microbiology and

Immunology, 358(1), 323-346. doi: 10.1007/82_2011_189.

Claesson, M. J., Cusack, S., O'Sullivan, O., Greene-Diniz, R., de Weerd, H., Flannery, E.,

Marchesi, J. R., Falush, D., Dinan, T., Fitzgerald, G., Stanton, C., van Sinderen, D., O'Connor,

M., Harnedy, N., O'Connor, K., Henry, C., O'Mahony, D., Fitzgerald, A. P., Shanahan, F.,

Twomey, C., Hill, C., Ross, R. P., & O'Toole, P. W. (2011). Composition, variability, and

temporal stability of the intestinal microbiota of the elderly. Proceedings of the National

Academy of Sciences of the United States of America, 108(1), 4586-4591.

doi: 10.1073/pnas.1000097107.

Collado, M. C., Gueimonde, M., Hernandez, M., Sanz, Y., & Salminen, S. (2005). Adhesion of

selected Bifidobacterium strains to human intestinal mucus and the role of adhesion in

enteropathogen exclusion. Journal of Food Protection, 68(12), 2672-2678. PMID: 16355841.

Collado, M. C., Gueimonde, M., Sanz, Y., & Salminen, S. (2006). Adhesion properties and

competitive pathogen exclusion ability of bifidobacteria with acquired acid resistance. Journal

of Food Protection, 69(7), 1675-1679. PMID: 16865903.

List of Literature

93

Collins, J., van Pijkeren, J., Svensson, L., Claesson, M. J., Sturme, M., Li, Y., Cooney, J. C., van

Sinderen, D., Walker, A. W., & Parkhill, J. (2012). Fibrinogen‐binding and platelet‐aggregation

activities of a Lactobacillus salivarius septicaemia isolate are mediated by a novel

fibrinogen‐binding protein. Molecular Microbiology, 85(5), 862-877. doi: 10.1111/j.1365-

2958.2012.08148.x.

Colombatti, A., Bonaldo, P., & Doliana, R. (1993). Type A modules: interacting domains found in

several non-fibrillar collagens and in other extracellular matrix proteins. Matrix, 13(4), 297-306.

doi: 10.1016/S0934-8832(11)80025-9.

Corazziari, E. S. (2009). Intestinal mucus barrier in normal and inflamed colon. Journal of

Pediatric Gastroenterology and Nutrition, 48(2), 54-55. doi: 10.1097/MPG.0b013e3181a117ea.

Corvey, C., Stein, T., Düsterhus, S., Karas, M., & Entian, K. (2003). Activation of subtilin

precursors by Bacillus subtilis extracellular serine proteases subtilisin (AprE), WprA, and Vpr.

Biochemical and Biophysical Research Communications, 304(1), 48-54. doi: 10.1016/S0006-

291X(03)00529-1.

Crociani, F., Biavati, B., Alessandrini, A., Chiarini, C., & Scardovi, V. (1996). Bifidobacterium

inopinatum sp. nov. and Bifidobacterium denticolens sp. nov., two new species isolated from

human dental caries. International Journal of Systematic Bacteriology, 46(2), 564-571.

doi: 10.1099/00207713-46-2-564.

de la Cochetière, M. F., Piloquet, H., des Robert, C., Darmaun, D., Galmiche, J. P., & Roze, J. C.

(2004). Early intestinal bacterial colonization and necrotizing enterocolitis in premature infants:

the putative role of Clostridium. Pediatric Research, 56(3), 366-370. doi:

10.1203/01.PDR.0000134251.45878.D5.

de Man, J. C., Rogosa, M., & Sharpe, E. M. (1960). A medium for the cultivation of lactobacilli.

Journal of Applied Bacteriology, 23(1), 130-135. doi: 10.1111/j.1365-2672.1960.tb00188.x.

de Vries, W., & Stouthamer, A. H. (1967). Pathway of glucose fermentation in relation to the

taxonomy of bifidobacteria. Journal of Bacteriology, 93(2), 574-576. PMID: 6020562

de Vuyst, L., de Vin, F., Vaningelgem, F., & Degeest, B. (2001). Recent developments in the

biosynthesis and applications of heteropolysaccharides from lactic acid bacteria. International

Dairy Journal, 11(9), 687-707. doi: 10.1016/S0958-6946(01)00114-5.

Deguchia, Y., Morishitaa, T., & Mutaia, M. (1985). Comparative studies on synthesis of water-

soluble vitamins among human species of bifidobacteria. Agricultural and Biological

Chemistry, 49(1), 13-19. doi: 10.1080/00021369.1985.10866683.

del Re, B., Sgorbati, B., Miglioli, M., & Palenzona, D. (2000). Adhesion, autoaggregation and

hydrophobicity of 13 strains of Bifidobacterium longum. Letters in Applied Microbiology,

31(6), 438-442. doi: 10.1046/j.1365-2672.2000.00845.x.

Delzenne, N. M., Neyrinck, A. M., Backhed, F., & Cani, P. D. (2011). Targeting gut microbiota in

obesity: effects of prebiotics and probiotics. Nature Reviews. Endocrinology, 7(11), 639-646.

doi: 10.1038/nrendo.2011.126.

Denou, E., Pridmore, R. D., Berger, B., Panoff, J. M., Arigoni, F., & Brussow, H. (2008).

Identification of genes associated with the long-gut-persistence phenotype of the probiotic

Lactobacillus johnsonii strain NCC533 using a combination of genomics and transcriptome

analysis. Journal of Bacteriology, 190(9), 3161-3168. doi: 10.1128/JB.01637-07.

List of Literature

94

Derrien, M., Vaughan, E. E., Plugge, C. M., & de Vos, W. M. (2004). Akkermansia muciniphila

gen. nov., sp. nov., a human intestinal mucin-degrading bacterium. International Journal of

Systematic and Evolutionary Microbiology, 54(Pt 5), 1469-1476. doi: 10.1099/ijs.0.02873-0.

Dertli, E., Mayer, M. J., & Narbad, A. (2015). Impact of the exopolysaccharide layer on biofilms,

adhesion and resistance to stress in Lactobacillus johnsonii FI9785. BMC Microbiology, 15(1),

8. doi: 10.1186/s12866-015-0347-2.

Desvaux, M., Hebraud, M., Talon, R., & Henderson, I. R. (2009). Secretion and subcellular

localizations of bacterial proteins: a semantic awareness issue. Trends in Microbiology, 17(4),

139-145. doi: 10.1016/j.tim.2009.01.004.

Dower, W. J., Miller, J. F., & Ragsdale, C. W. (1988). High efficiency transformation of E. coli by

high voltage electroporation. Nucleic Acids Research, 16(13), 6127-6145.

doi: 10.1093/nar/16.13.6127.

Dunne, C., O'Mahony, L., Murphy, L., Thornton, G., Morrissey, D., O'Halloran, S., Feeney, M.,

Flynn, S., Fitzgerald, G., Daly, C., Kiely, B., O'Sullivan, G. C., Shanahan, F., & Collins, J. K.

(2001). In vitro selection criteria for probiotic bacteria of human origin: correlation with in vivo

findings. The American Journal of Clinical Nutrition, 73(2), 386-392. PMID: 11157346.

Eckburg, P. B., Bik, E. M., Bernstein, C. N., Purdom, E., Dethlefsen, L., Sargent, M., Gill, S. R.,

Nelson, K. E., & Relman, D. A. (2005). Diversity of the human intestinal microbial flora.

Science, 308(5728), 1635-1638. doi: 10.1126/science.1110591.

Ewaschuk, J. B., Diaz, H., Meddings, L., Diederichs, B., Dmytrash, A., Backer, J., Looijer-van

Langen, M., & Madsen, K. L. (2008). Secreted bioactive factors from Bifidobacterium infantis

enhance epithelial cell barrier function. American Journal of Physiology.Gastrointestinal and

Liver Physiology, 295(5), 1025-1034. doi: 10.1152/ajpgi.90227.2008.

Fanaro, S., Vigi, V., Chierici, R., & Boehm, G. (2003). Fecal flora measurements of breastfed

infants using an integrated transport and culturing system. Acta Paediatrica, 92(5), 634-635.

doi: 10.1111/j.1651-2227.2003.tb02521.x

Fanning, S., Hall, L. J., & van Sinderen, D. (2012). Bifidobacterium breve UCC2003 surface

exopolysaccharide production is a beneficial trait mediating commensal-host interaction through

immune modulation and pathogen protection. Gut Microbes, 3(5), 420-425.

doi: 10.4161/gmic.20630.

Fanning, S., Hall, L. J., Cronin, M., Zomer, A., MacSharry, J., Goulding, D., Motherway, M. O.,

Shanahan, F., Nally, K., Dougan, G., & van Sinderen, D. (2012). Bifidobacterial surface-

exopolysaccharide facilitates commensal-host interaction through immune modulation and

pathogen protection. Proceedings of the National Academy of Sciences of the United States of

America, 109(6), 2108-2113. doi: 10.1073/pnas.1115621109.

Fassel, T. A., Schaller, M. J., & Remsen, C. C. (1992). Comparison of alcian blue and ruthenium

red effects on preservation of outer envelope ultrastructure in methanotrophic bacteria.

Microscopy Research and Technique, 20(1), 87-94. doi: 10.1002/jemt.1070200109.

Favier, C. F., Vaughan, E. E., de Vos, W. M., & Akkermans, A. D. (2002). Molecular monitoring

of succession of bacterial communities in human neonates. Applied and Environmental

Microbiology, 68(1), 219-226. doi: 10.1128/AEM.68.1.219-226.2002.

Fernández, L., Langa, S., Martín, V., Maldonado, A., Jimenez, E., Martín, R., & Rodríguez, J. M.

(2013). The human milk microbiota: origin and potential roles in health and disease.

Pharmacological Researc, 69(1), 1-10. doi: 10.1016/j.phrs.2012.09.001.

List of Literature

95

Fernández-Esplá, M. D., Garault, P., Monnet, V., & Rul, F. (2000). Streptococcus thermophilus

cell wall-anchored proteinase: release, purification, and biochemical and genetic

characterization. Applied and Environmental Microbiology, 66(11), 4772-4778. doi:

10.1128/AEM.66.11.4772-4778.2000.

Fink, A. L. (1999). Chaperone-mediated protein folding. Physiological Reviews, 79(2), 425-449.

PMID: 10221986.

Finn, R. D., Bateman, A., Clements, J., Coggill, P., Eberhardt, R. Y., Eddy, S. R., Heger, A.,

Hetherington, K., Holm, L., Mistry, J., Sonnhammer, E. L. L., Tate, J., & Punta, M. (2014).

Pfam: the protein families database. Nucleic Acids Research, 42(1), 222-230. doi:

10.1093/nar/gkt1223.

Florey, H. (1955). Mucin and the protection of the body. Proceedings of the Royal Society of

London. Series B, Biological Sciences, 143(911), 147-158. PMID:14371602.

Fogh, J., Fogh, J. M., & Orfeo, T. (1977). One hundred and twenty-seven cultured human tumor

cell lines producing tumors in nude mice. Journal of the National Cancer Institute, 59(1), 221-

226. PMID: 327080.

Foroni, E., Serafini, F., Amidani, D., Turroni, F., He, F., Bottacini, F., O'Connell Motherway, M.,

Viappiani, A., Zhang, Z., Rivetti, C., van Sinderen, D., & Ventura, M. (2011). Genetic analysis

and morphological identification of pilus-like structures in members of the genus

Bifidobacterium. Microbial Cell Factories, 10(1), 1-13. doi: 10.1186/1475-2859-10-S1-S16.

Fraser, J. S., Maxwell, K. L., & Davidson, A. R. (2007). Immunoglobulin-like domains on

bacteriophage: weapons of modest damage? Current Opinion in Microbiology, 10(4), 382-387.

doi: S1369-5274(07)00087-2.

Fuller, R. (1992). History and development of probiotics. In: Fuller, R., Probiotics. The scientific

basis. pp 1-8. Springer Netherlands, ISBN: 978-94-010-5043-2.

Gagic, D., Wen, W., Collett, M. A., & Rakonjac, J. (2013). Unique secreted-surface protein

complex of Lactobacillus rhamnosus, identified by phage display. Microbiology open, 2(1), 1-

17. doi: 10.1002/mbo3.53.

Gardy, J. L., Laird, M. R., Chen, F., Rey, S., Walsh, C. J., Ester, M., & Brinkman, F. S. (2005).

PSORTb v.2.0: expanded prediction of bacterial protein subcellular localization and insights

gained from comparative proteome analysis. Bioinformatics, 21(5), 617-623.

doi: 10.1093/bioinformatics/bti057

Gelse, K., Pöschl, E., & Aigner, T. (2003). Collagens—structure, function, and biosynthesis.

Advanced Drug Delivery Reviews, 55(12), 1531-1546. doi: 10.1016/j.addr.2003.08.002.

Gill, H. (1998). Stimulation of the immune system by lactic cultures. International Dairy Journal,

8(5-6), 535-544. doi: 10.1016/S0958-6946(98)00074-0.

Gillen, C. M., Courtney, H. S., Schulze, K., Rohde, M., Wilson, M. R., Timmer, A. M., Guzman,

C. A., Nizet, V., Chhatwal, G. S., & Walker, M. J. (2008). Opacity factor activity and epithelial

cell binding by the serum opacity factor protein of Streptococcus pyogenes are functionally

discrete. The Journal of Biological Chemistry, 283(10), 6359-6366.

doi: 10.1074/jbc.M706739200.

Gleinser M, Grimm V, Zhurina D, Yuan J, Riedel CU. (2012). Improved adhesive properties of

recombinant bifidobacteria expressing the Bifidobacterium bifidum-specific lipoprotein BopA.

Microbial Cell Factories, 11(80), 1-14. doi: 10.1186/1475-2859-11-80.

List of Literature

96

Glenting, J., Beck, H. C., Vrang, A., Riemann, H., Ravn, P., Hansen, A. M., Antonsson, M., Ahrne,

S., Israelsen, H., & Madsen, S. (2013). Anchorless surface associated glycolytic enzymes from

Lactobacillus plantarum 299v bind to epithelial cells and extracellular matrix proteins.

Microbiological Research, 168(5), 245-253. doi: 10.1016/j.micres.2013.01.003.

González-Rodríguez, I., Ruiz, L., Gueimonde, M., Margolles, A., & Sánchez, B. (2013). Factors

involved in the colonization and survival of bifidobacteria in the gastrointestinal tract. FEMS

Microbiology Letters, 340(1), 1-10. doi: http://dx.doi.org/10.1111/1574-6968.12056.

González-Rodríguez, I., Sánchez, B., Ruiz, L., Turroni, F., Ventura, M., Ruas-Madiedo, P.,

Gueimonde, M., & Margolles, A. (2012). Role of extracellular transaldolase from

Bifidobacterium bifidum in mucin adhesion and aggregation. Applied and Environmental

Microbiology, 78(11), 3992-3998. doi: 10.1128/AEM.08024-11.

Granato, D., Branco, G. F., Cruz, A. G., Faria, J. A. F., & Shah, N. P. (2010). Probiotic Dairy

products as functional foods. Comprehensive Reviews in Food Science and Food Safety, 9(1),

455-470. doi: 10.1111/j.1541-4337.2010.00.

Green, M. R., & Sambrook, J. (2012). Molecular cloning: a laboratory manual. Cold Spring

Harbor Laboratory Press. ISBN: 978-1-936113-42-2.

Grimm, V., Gleinser, M., Neu, C., Zhurina, D., & Riedel, C. U. (2014). Expression of fluorescent

proteins in bifidobacteria for analysis of host-microbe interactions. Applied and Environmental

Microbiology, 80(9), 2842-2850. doi: 10.1128/AEM.04261-13.

Gritz, E. C., & Bhandari, V. (2015). The human neonatal gut microbiome: a brief review. Frontiers

in Pediatrics, 3(17), 1-12, doi: 10.3389/fped.2015.00017.

Groeger, D., O'Mahony, L., Murphy, E. F., Bourke, J. F., Dinan, T. G., Kiely, B., Shanahan, F., &

Quigley, E. M. (2013). Bifidobacterium infantis 35624 modulates host inflammatory processes

beyond the gut. Gut Microbes, 4(4), 325-339. doi: 10.4161/gmic.25487.

Guardamagna, O., Amaretti, A., Puddu, P. E., Raimondi, S., Abello, F., Cagliero, P., & Rossi, M.

(2014). Bifidobacteria supplementation: effects on plasma lipid profiles in dyslipidemic

children. Nutrition, 30(7-8), 831-836. doi: 10.1016/j.nut.2014.01.014.

Gueimonde, M., Margolles, A., Clara, G., & Salminen, S. (2007). Competitive exclusion of

enteropathogens from human intestinal mucus by Bifidobacterium strains with acquired

resistance to bile—A preliminary study. International Journal of Food Microbiology, 113(2),

228-232. doi: 10.1016/j.ijfoodmicro.2006.05.017.

Guglielmetti S, Tamagnini I, Mora D, Minuzzo M, Scarafoni A, Arioli S, Hellman J, Karp M,

Parini C. (2008). Implication of an outer surface lipoprotein in adhesion of Bifidobacterium

bifidum to Caco-2 cells. Applied and Environmental Microbiology, 74(15), 4695-4702. doi:

10.1128/AEM.00124-08.

Hamburger, Z. A., Brown, M. S., Isberg, R. R., & Bjorkman, P. J. (1999). Crystal structure of

invasin: a bacterial integrin-binding protein. Science, 286(5438), 291-295.

doi: 10.1126/science.286.5438.291.

Hammerschmidt, S., Wolff, S., Hocke, A., Rosseau, S., Muller, E., & Rohde, M. (2005).

Illustration of pneumococcal polysaccharide capsule during adherence and invasion of epithelial

cells. Infection and Immunity, 73(8), 4653-4667. doi: 10.1128/IAI.73.8.4653-4667.2005.

List of Literature

97

Hauck, C. R., Agerer, F., Muenzner, P., & Schmitter, T. (2006). Cellular adhesion molecules as

targets for bacterial infection. European Journal of Cell Biology, 85(3), 235-242.

doi: 10.1016/j.ejcb.2005.08.002.

Hautefort, I., Thompson, A., Eriksson‐Ygberg, S., Parker, M., Lucchini, S., Danino, V., Bongaerts,

R., Ahmad, N., Rhen, M., & Hinton, J. (2008). During infection of epithelial cells Salmonella

enterica serovar Typhimurium undergoes a time‐dependent transcriptional adaptation that

results in simultaneous expression of three type 3 secretion systems. Cellular Microbiology,

10(4), 958-984. doi: 10.1111/j.1462-5822.2007.01099.x.

Hayashida, H., Ishihara, K., Ichikawa, T., Okayasu, I., Kurihara, M., Saigenji, K., & Hotta, K.

(2001). Expression of a specific mucin type recognized by monoclonal antibodies in the rat

gastric mucosa regenerating from acetic acid-induced ulcer. Scandinavian Journal of

Gastroenterology, 36(5), 467-473. doi: 10.1080/00365520116855.

He, F., Ouwehan, A. C., Hashimoto, H., Isolauri, E., Benno, Y., & Salminen, S. (2001). Adhesion

of Bifidobacterium spp. to human intestinal mucus. Microbiology and Immunology, 45(3), 259-

262. doi: 10.1111/j.1348-0421.2001.tb02615.x.

He, F., Ouwehand, A. C., Isolauri, E., Hashimoto, H., Benno, Y., & Salminen, S. (2001).

Comparison of mucosal adhesion and species identification of bifidobacteria isolated from

healthy and allergic infants. FEMS Immunology and Medical Microbiology, 30(1), 43-47. doi:

S0928-8244(00)00232-7.

Heino, J. (2007). The collagen family members as cell adhesion proteins. Bioessays, 29(10), 1001-

1010. doi: 10.1002/bies.20636.

Henderson, B., Nair, S., Pallas, J., & Williams, M. A. (2011). Fibronectin: a multidomain host

adhesin targeted by bacterial fibronectin-binding proteins. FEMS Microbiology Reviews, 35(1),

147-200. doi: 10.1111/j.1574-6976.2010.00243.x.

Hendrickx, A. P., Willems, R. J., Bonten, M. J., & van Schaik, W. (2009). LPxTG surface proteins

of enterococci. Trends in Microbiology, 17(9), 423-430. doi: 10.1016/j.tim.2009.06.004.

Hendrickx, A. P., Budzik, J. M., Oh, S., & Schneewind, O. (2011). Architects at the bacterial

surface—sortases and the assembly of pili with isopeptide bonds. Nature Reviews Microbiology,

9(3), 166-176. doi: 10.1038/nrmicro2520.

Hidalgo-Cantabrana, C., López, P., Gueimonde, M., Clara, G., Suárez, A., Margolles, A., & Ruas-

Madiedo, P. (2012). Immune modulation capability of exopolysaccharides synthesised by lactic

acid bacteria and bifidobacteria. Probiotics and Antimicrobial Proteins, 4(4), 227-237. doi:

10.1007/s2602-012-9110-2.

Hidalgo-Cantabrana, C., Sánchez, B., Milani, C., Ventura, M., Margolles, A., & Ruas-Madiedo, P.

(2014). Genomic overview and biological functions of exopolysaccharide biosynthesis in

Bifidobacterium spp.. Applied and Environmental Microbiology, 80(1), 9-18.

doi:10.1128/AEM.02977-13.

Hidalgo-Grass, C., Mishalian, I., Dan-Goor, M., Belotserkovsky, I., Eran, Y., Nizet, V., Peled, A.,

& Hanski, E. (2006). A streptococcal protease that degrades CXC chemokines and impairs

bacterial clearance from infected tissues. The EMBO Journal, 25(19), 4628-4637.

doi: 10.1038/sj.emboj.7601327.

Hörmannsperger, G., von Schillde, M., & Haller, D. (2013). Lactocepin as a protective microbial

structure in the context of IBD. Gut Microbes, 4(2), 152-157. doi: 10.4161/gmic.23444.

List of Literature

98

Horn, N., Wegmann, U., Dertli, E., Mulholland, F., Collins, S. R., Waldron, K. W., Bongaerts, R.

J., Mayer, M. J., & Narbad, A. (2013). Spontaneous mutation reveals influence of

exopolysaccharide on Lactobacillus johnsonii surface characteristics. PloS One, 8(3), e59957.

doi: 10.1371/journal.pone.0059957.

Hoskins, L. C. (2012). Degradation of mucus glycoproteins in the gastrointestinal tract. In:

Horowitz, M. (Ed.), The Glycoconjugates V2: Mammalian Glycoproteins and Glycolipids and

Proteoglycans. pp 235-253. Elsevier. ISBN: 9780323148832.

Hotta, K. (2000). "Gastric mucus", a mysterious and interesting substance. Trends in Glycoscience

and Glycotechnology, 12(63), 59-68. doi: 10.1111/j.1365-2982.2004.00580.x

Huberts, D. H., & van der Klei, Ida J. (2010). Moonlighting proteins: an intriguing mode of

multitasking. Biochimica Et Biophysica Acta (BBA)-Molecular Cell Research, 1803(4), 520-

525. doi: 10.1016/j.bbamcr.2010.01.022.

Hultgfien, S. J., Jaco, B., Bu, F., & Branden, C. (1993). PapD and superfamily of periplasmic

immunoglobulin-like pilus chaperones. Volume 44. In: Meurant, G., Advances in Protein

Chemistry. pp 99-124. Academic Press. ISBN: 9780080582177.

Ikezawa, T., Goso, Y., Ichikawa, T., Hayashida, H., Kurihara, M., Okayasu, I., Saigenji, K., &

Ishihara, K. (2004). Appearance of specific mucins recognized by monoclonal antibodies in rat

gastric mucosa healing from HCl-induced gastric mucosal damage. Journal of

Gastroenterology, 39(2), 113-119. doi: 10.1007/s00535-003-1261-1.

Jensen, H., Roos, S., Jonsson, H., Rud, I., Grimmer, S., van Pijkeren, J. P., Britton, R. A., &

Axelsson, L. (2014). Role of Lactobacillus reuteri cell and mucus-binding protein A (CmbA) in

adhesion to intestinal epithelial cells and mucus in vitro. Microbiology, 160(4), 671-681. doi:

10.1099/mic.0.073551-0.

Jensen, P. R., & Hammer, K. (1998). The sequence of spacers between the consensus sequences

modulates the strength of prokaryotic promoters. Applied and Environmental Microbiology,

64(1), 82-87. PMCID: PMC124675.

Jiang, T., Mustapha, A., & Savaiano, D. A. (1996). Improvement of lactose digestion in humans by

ingestion of unfermented milk containing Bifidobacterium longum. Journal of Dairy Science,

79(5), 750-757. doi: 10.3168/jds.S0022-0302(96)76422-6.

Johansson, M. E., Ambort, D., Pelaseyed, T., Schütte, A., Gustafsson, J. K., Ermund, A.,

Subramani, D. B., Holmén-Larsson, J. M., Thomsson, K. A., & Bergström, J. H. (2011).

Composition and functional role of the mucus layers in the intestine. Cellular and Molecular

Life Sciences, 68(22), 3635-3641. doi: 10.1007/s00018-011-0822-3.

Johansson, M. E., Phillipson, M., Petersson, J., Velcich, A., Holm, L., & Hansson, G. C. (2008).

The inner of the two Muc2 mucin-dependent mucus layers in colon is devoid of bacteria.

Proceedings of the National Academy of Sciences of the United States of America, 105(39),

15064-15069. doi: 10.1073/pnas.0803124105.

Jubair, M., Atanasova, K. R., Rahman, M., Klose, K. E., Yasmin, M., Yilmaz, Ö., Morris Jr, J. G.,

& Ali, A. (2014). Vibrio cholerae persisted in microcosm for 700 days inhibits motility but

promotes biofilm formation in nutrient-poor lake water microcosms. PloS One, 9(3), e92883.


Juhas, M., Wiehlmann, L., Salunkhe, P., Lauber, J., Buer, J., & Tümmler, B. (2005). GeneChip

expression analysis of the VqsR regulon of Pseudomonas aeruginosa TB. FEMS Microbiology

Letters, 242(2), 287-295. doi: http://dx.doi.org/10.1016/j.femsle.2004.11.020.

List of Literature

99

Kachlany, S. C., Planet, P. J., Bhattacharjee, M. K., Kollia, E., DeSalle, R., Fine, D. H., & Figurski,

D. H. (2000). Nonspecific adherence by Actinobacillus actinomycetemcomitans requires genes

widespread in bacteria and archaea. Journal of Bacteriology, 182(21), 6169-6176.

doi: 10.1128/JB.182.21.6169-6176.2000.

Kailasapathy, K., & Chin, J. (2000). Survival and therapeutic potential of probiotic organisms with

reference to Lactobacillus acidophilus and Bifidobacterium spp.. Immunology and Cell Biology,

78(1), 80-88. doi: 10.1046/j.1440-1711.2000.00886.x.

Kankainen, M., Paulin, L., Tynkkynen, S., von Ossowski, I., Reunanen, J., Partanen, P., Satokari,

R., Vesterlund, S., Hendrickx, A. P., Lebeer, S., De Keersmaecker, S. C., Vanderleyden, J.,

Hamalainen, T., Laukkanen, S., Salovuori, N., Ritari, J., Alatalo, E., Korpela, R., Mattila-

Sandholm, T., Lassig, A., Hatakka, K., Kinnunen, K. T., Karjalainen, H., Saxelin, M., Laakso,

K., Surakka, A., Palva, A., Salusjarvi, T., Auvinen, P., & de Vos, W. M. (2009). Comparative

genomic analysis of Lactobacillus rhamnosus GG reveals pili containing a human- mucus

binding protein. Proceedings of the National Academy of Sciences of the United States of

America, 106(40), 17193-17198. doi: 10.1073/pnas.0908876106.

Karlsson, C., Andersson, M. L., Collin, M., Schmidtchen, A., Bjorck, L., & Frick, I. M. (2007).

SufA--a novel subtilisin-like serine proteinase of Finegoldia magna. Microbiology, 153(12),

4208-4218. doi: 10.1099/mic.0.2007/010322-0.

Kassinen, A., Krogius-Kurikka, L., Makivuokko, H., Rinttila, T., Paulin, L., Corander, J., Malinen,

E., Apajalahti, J., & Palva, A. (2007). The fecal microbiota of irritable bowel syndrome patients

differs significantly from that of healthy subjects. Gastroenterology, 133(1), 24-33. doi: S0016-

5085(07)00734-2.

Katerov, V., Lindgren, P. E., Totolian, A. A., & Schalen, C. (2000). Streptococcal opacity factor: a

family of bifunctional proteins with lipoproteinase and fibronectin-binding activities. Current

Microbiology, 40(3), 149-156. doi: 10.1007/s002849910031.

Kelly, G., Prasannan, S., Daniell, S., Fleming, K., Frankel, G., Dougan, G., Connerton, I., &

Matthews, S. (1999). Structure of the cell-adhesion fragment of intimin from enteropathogenic

Escherichia coli. Nature Structural Biology, 6(4), 313-318. doi: 10.1038/7545.

Kil, K. S., Cunningham, M. W., & Barnett, L. A. (1994). Cloning and sequence analysis of a gene

encoding a 67-kilodalton myosin-cross-reactive antigen of Streptococcus pyogenes reveals its

similarity with class II major histocompatibility antigens. Infection and Immunity, 62(6), 2440-

2449. doi: 0019-9567/94/$04.00+0.

Kim, J. Y., Wang, Y., Park, M. S., & Ji, G. E. (2010). Improvement of transformation efficiency

through in vitro methylation and SacII site mutation of plasmid vector in Bifidobacterium

longum MG1. Journal of Microbiology and Biotechnology, 20(6), 1022-1026.

doi: 10.4014/jmb.1003.03014.

Kinoshita, H., Uchida, H., Kawai, Y., Kawasaki, T., Wakahara, N., Matsuo, H., Watanabe, M.,

Kitazawa, H., Ohnuma, S., & Miura, K. (2008). Cell surface Lactobacillus plantarum LA 318

glyceraldehyde‐3‐phosphate dehydrogenase (GAPDH) adheres to human colonic mucin.

Journal of Applied Microbiology, 104(6), 1667-1674. doi: 10.1111/j.1365-2672.2007.03679.x.

Kleerebezem, M., Hols, P., Bernard, E., Rolain, T., Zhou, M., Siezen, R. J., & Bron, P. A. (2010).

The extracellular biology of the lactobacilli. FEMS Microbiology Reviews, 34(2), 199-230.

doi: 10.1111/j.1574-6976.2010.00208.x.

List of Literature

100

Klenow, H., & Henningsen, I. (1970). Selective elimination of the exonuclease activity of the

deoxyribonucleic acid polymerase from Escherichia coli B by limited proteolysis. Proceedings

of the National Academy of Sciences of the United States of America, 65(1), 168-175.

PMID: 4905667.

Klijn, A., Mercenier, A., & Arigoni, F. (2005). Lessons from the genomes of bifidobacteria. FEMS

Microbiology Reviews, 29(3), 491-509. doi: S0168-6445(05)00034-3.

Kolenbrander, P. E., Palmer, R. J.,Jr, Rickard, A. H., Jakubovics, N. S., Chalmers, N. I., & Diaz, P.

I. (2006). Bacterial interactions and successions during plaque development. Periodontology

2000, 42, 47-79. doi: 10.1111/j.1600-0757.2006.00187.x.

Kolling, Y., Salva, S., Villena, J., Marranzino, G., & Alvarez, S. (2015). Non-viable immunobiotic

Lactobacillus rhamnosus CRL1505 and its peptidoglycan improve systemic and respiratory

innate immune response during recovery of immunocompromised-malnourished mice.

International Immunopharmacology, 25(2), 474-484. doi: S1567-5769(15)00052-1.

Kopečný, J., Mrázek, J., & Killer, J. (2010). The presence of bifidobacteria in social insects, fish

and reptiles. Folia Microbiologica, 55(4), 336-339. doi: 10.1007/s12223-010-0053-2.

Kos, B., Suskovic, J., Vukovic, S., Simpraga, M., Frece, J., & Matosic, S. (2003). Adhesion and

aggregation ability of probiotic strain Lactobacillus acidophilus M92. Journal of Applied

Microbiology, 94(6), 981-987. doi: 10.1046/j.1365-2672.2003.01915.x.

Kotani, S., Watanabe, Y., Shimono, T., Narita, T., & Kato, K. (1975). Immunoadjuvant activities

of cell walls, their water-soluble fractions and peptidoglycan subunits, prepared from various

Gram-positive bacteria, and of synthetic n-acetylmuramyl peptides. Zeitschrift Für

Immunitätsforschung, Experimentelle Und Klinische Immunologie, 149(2-4), 302-319. PMID:

126564.

Krogh, A., Larsson, B., von Heijne, G., & Sonnhammer, E. L. (2001). Predicting transmembrane

protein topology with a hidden Markov model: application to complete genomes. Journal of

Molecular Biology, 305(3), 567-580. doi: 10.1006/jmbi.2000.4315.

Kurokawa, K., Itoh, T., Kuwahara, T., Oshima, K., Toh, H., Toyoda, A., Takami, H., Morita, H.,

Sharma, V. K., Srivastava, T. P., Taylor, T. D., Noguchi, H., Mori, H., Ogura, Y., Ehrlich, D.

S., Itoh, K., Takagi, T., Sakaki, Y., Hayashi, T., & Hattori, M. (2007). Comparative

metagenomics revealed commonly enriched gene sets in human gut microbiomes. DNA

Research, 14(4), 169-181. doi: 10.1093/dnares/dsm018.

Laakso K, Koskenniemi K, Koponen J, Kankainen M, Surakka A, Salusjarvi T, Auvinen P,

Savijoki K, Nyman TA, Kalkkinen N, Tynkkynen S, Varmanen P. (2011). Growth phase-

associated changes in the proteome and transcriptome of Lactobacillus rhamnosus GG in

industrial-type whey medium. Microbial Biotechnology, 4(6), 746-766. doi: 10.1111/j.1751-

7915.2011.00275.x.

Lebeer S, Claes I, Tytgat HL, Verhoeven TL, Marien E, von Ossowski I, Reunanen J, Palva A, Vos

WM, Keersmaecker SC, Vanderleyden J. (2012). Functional analysis of Lactobacillus

rhamnosus GG pili in relation to adhesion and immunomodulatory interactions with intestinal

epithelial cells. Applied and Environmental Microbiology, 78(1), 185-193. doi:

10.1128/AEM.06192-11.

Lebeer, S., Vanderleyden, J., & de Keersmaecker, S. C. (2010). Host interactions of probiotic

bacterial surface molecules: comparison with commensals and pathogens. Nature Reviews

Microbiology, 8(3), 171-184. doi: 10.1038/nrmicro2297.

List of Literature

101

Lee, Y. K., & Salminen, S. (2009). Handbook of probiotics and probiotics. John Wiley & Sons.

ISBN: 9780470432617.

Lee, J. H., & O'Sullivan, D. J. (2010). Genomic insights into bifidobacteria. Microbiology and

Molecular Biology Reviews, 74(3), 378-416. doi: 10.1128/MMBR.00004-10.

Lehman, I. R. (1974). DNA ligase: structure, mechanism, and function. Science, 186(4166), 790-

797. doi: 10.1126/science.186.4166.790.

Lilic, M., Vujanac, M., & Stebbins, C. E. (2006). A common structural motif in the binding of

virulence factors to bacterial secretion chaperones. Molecular Cell, 21(5), 653-664. doi: S1097-

2765(06)00048-7.

Liu, S., Rich, J. O., & Anderson, A. (2015). Antibacterial activity of a cell wall hydrolase from

Lactobacillus paracasei NRRL B-50314 produced by recombinant Bacillus megaterium.

Journal of Industrial Microbiology & Biotechnology, 42(2), 229-235. doi: 10.1007/s10295-014-

1557-6.

López, P., González-Rodríguez, I., Gueimonde, M., Margolles, A., & Suárez, A. (2011). Immune

response to Bifidobacterium bifidum strains support Treg/Th17 plasticity. PLoS One, 6(9),

e24776. doi: 10.1371/journal.pone.0024776.

López, P., Monteserin, D. C., Gueimonde, M., Clara, G., Margolles, A., Suarez, A., & Ruas-

Madiedo, P. (2012). Exopolysaccharide-producing Bifidobacterium strains elicit different in

vitro responses upon interaction with human cells. Food Research International, 46(1), 99-107.

doi: 10.1016/j.foodres.2011.11.020.

Lu, L., & Walker, W. A. (2001). Pathologic and physiologic interactions of bacteria with the

gastrointestinal epithelium. The American Journal of Clinical Nutrition, 73(6), 1124-1130.

PMID: 11393190.

Luck, S. N., Badea, L., Bennett-Wood, V., Robins-Browne, R., & Hartland, E. L. (2006).

Contribution of FliC to epithelial cell invasion by enterohemorrhagic Escherichia coli

O113:H21. Infection and Immunity, 74(12), 6999-7004. doi: 10.1128/IAI.00435-06.

Luo, Y., Frey, E. A., Pfuetzner, R. A., Creagh, A. L., Knoechel, D. G., Haynes, C. A., Finlay, B.

B., & Strynadka, N. C. (2000). Crystal structure of enteropathogenic Escherichia coli intimin-

receptor complex. Nature, 405(6790), 1073-1077. doi: 10.1038/35016618.

Lynch, M., & Crease, T. J. (1990). The analysis of population survey data on DNA sequence

variation. Molecular Biology and Evolution, 7(4), 377-394. doi: 0737-030/90/0704-008$02.00.

Macfarlane, S., Woodmansey, E. J., & Macfarlane, G. T. (2005). Colonization of mucin by human

intestinal bacteria and establishment of biofilm communities in a two-stage continuous culture

system. Applied and Environmental Microbiology, 71(11), 7483-7492.

doi: 10.1128/AEM.71.11.7483-7492.2005.

Macías‐Rodríguez, M., Zagorec, M., Ascencio, F., Vázquez‐Juárez, R., & Rojas, M. (2009).

Lactobacillus fermentum BCS87 expresses mucus‐and mucin‐binding proteins on the cell

surface. Journal of Applied Microbiology, 107(6), 1866-1874. doi: 10.1111/j.1365-

2672.2009.04368.x.

Makarova, K. S., Wolf, Y. I., & Koonin, E. V. (2013). Comparative genomics of defense systems

in archaea and bacteria. Nucleic Acids Research, 41(8), 4360-4377. doi: 10.1093/nar/gkt157.

Mao, Y., & Schwarzbauer, J. E. (2005). Fibronectin fibrillogenesis, a cell-mediated matrix

assembly process. Matrix Biology, 24(6), 389-399. doi: 10.1016/j.matbio.2005.06.008.

List of Literature

102

Marco, M. L., Bongers, R. S., de Vos, W. M., & Kleerebezem, M. (2007). Spatial and temporal

expression of Lactobacillus plantarum genes in the gastrointestinal tracts of mice. Applied and

Environmental Microbiology, 73(1), 124-132. doi: 10.1128/AEM.01475-06.

Marcobal, A., & Sonnenburg, J. (2012). Human milk oligosaccharide consumption by intestinal

microbiota. Clinical Microbiology and Infection, 18(s4), 12-15. doi: 10.1111/j.1469-

0691.2012.03863.x.

Mariat, D., Firmesse, O., Levenez, F., Guimaraes, V., Sokol, H., Dore, J., Corthier, G., & Furet, J.

P. (2009). The Firmicutes/Bacteroidetes ratio of the human microbiota changes with age. BMC

Microbiology, 9(123), 1-6. doi: 10.1186/1471-2180-9-123.

Martín, R., Jimenez, E., Heilig, H., Fernández, L., Marin, M. L., Zoetendal, E. G., & Rodríguez, J.

M. (2009). Isolation of bifidobacteria from breast milk and assessment of the bifidobacterial

population by PCR-denaturing gradient gel electrophoresis and quantitative real-time PCR.


Masco, L., Huys, G., De Brandt, E., Temmerman, R., & Swings, J. (2005). Culture-dependent and

culture-independent qualitative analysis of probiotic products claimed to contain bifidobacteria.

International Journal of Food Microbiology, 102(2), 221-230. doi: S0168-1605(05)00059-0.

McGuckin, M. A., Lindén, S. K., Sutton, P., & Florin, T. H. (2011). Mucin dynamics and enteric

pathogens. Nature Reviews Microbiology, 9(4), 265-278. doi: 10.1038/nrmicro2538.

Mei, S., Zhang, J., Zhang, X., & Tu, X. (2015). Solution structure of a bacterial immunoglobulin-

like domain of the outer membrane protein (LigB) from Leptospira. Proteins, 83(1), 195-200.

doi: 10.1002/prot.24723.

Mesnard, B., De Vroey, B., Maunoury, V., & Lecuit, M. (2012). Immunoproliferative small

intestinal disease associated with Campylobacter jejuni. Digestive and Liver Disease, 44(9),

799-800. doi: 10.1016/j.dld.2012.03.020

Metchnikoff, E., & Chalmers, M. P. (1908). The prolongation of life; optimistic studies, (Ed.)

Springer Publishing Company. ISBN: 9780826118776.

Meyer, H., Bug, M., & Bremer, S. (2012). Emerging functions of the VCP/p97 AAA-ATPase in

the ubiquitin system. Nature Cell Biology, 14(2), 117-123. doi: 10.1038/ncb2407.

Milani, C., Hevia, A., Foroni, E., Duranti, S., Turroni, F., Lugli, G. A., Sánchez, B., Martín, R.,

Gueimonde, M., van Sinderen, D., Margolles, A., & Ventura, M. (2013). Assessing the fecal

microbiota: an optimized ion torrent 16S rRNA gene-based analysis protocol. PloS One, 8(7),

e68739. doi: 10.1371/journal.pone.0068739.

Miljkovic, M., Strahinic, I., Tolinacki, M., Zivkovic, M., Kojic, S., Golic, N., & Kojic, M. (2015).

AggLb Is the Largest Cell-Aggregation Factor from Lactobacillus paracasei subsp. paracasei

BGNJ1-64, Functions in Collagen Adhesion, and Pathogen Exclusion In Vitro. PLoS One,

10(5), e0126387. doi: 10.1371/journal.pone.0126387.

Mitsuma, T., Odajima, H., Momiyama, Z., Watanabe, K., Masuguchi, M., Sekine, T., Shidara, S.,

& Hirano, S. (2008). Enhancement of gene expression by a peptide p (CHWPR) produced by

Bifidobacterium lactis BB‐12. Microbiology and Immunology, 52(3), 144-155.

doi: 10.1111/j.1348-0421.2008.00022.x.

Miyoshi, Y., Okada, S., Uchimura, T., & Satoh, E. (2006). A mucus adhesion promoting protein,

MapA, mediates the adhesion of Lactobacillus reuteri to Caco-2 human intestinal epithelial

cells. Bioscience, Biotechnology, and Biochemistry, 70(7), 1622-1628. doi: 10.1271/bbb.50688.

List of Literature

103

Molmenti, E. P., Ziambaras, T., & Perlmutter, D. H. (1993). Evidence for an acute phase response

in human intestinal epithelial cells. The Journal of Biological Chemistry, 268(19), 14116-14124.

PMID: 7686149.

Mozzi, F., Vaningelgem, F., Hebert, E. M., Van der Meulen, R., Foulquie Moreno, M. R., Font de

Valdez, G., & de Vuyst, L. (2006). Diversity of heteropolysaccharide-producing lactic acid

bacterium strains and their biopolymers. Applied and Environmental Microbiology, 72(6), 4431-

4435. doi: 10.1128/AEM.02780-05.

Mueller, S., Saunier, K., Hanisch, C., Norin, E., Alm, L., Midtvedt, T., Cresci, A., Silvi, S.,

Orpianesi, C., Verdenelli, M. C., Clavel, T., Koebnick, C., Zunft, H. J., Dore, J., & Blaut, M.

(2006). Differences in fecal microbiota in different European study populations in relation to

age, gender, and country: a cross-sectional study. Applied and Environmental Microbiology,

72(2), 1027-1033. doi: 10.1128/AEM.72.2.1027-1033.2006.

Muñoz-Provencio, D., Llopis, M., Antolin, M., de Torres, I., Guarner, F., Pérez-Martínez, G., &

Monedero, V. (2009). Adhesion properties of Lactobacillus casei strains to resected intestinal

fragments and components of the extracellular matrix. Archives of Microbiology, 191(2), 153-

161. doi: 10.1007/s00203-008-0436-9.

Muñoz-Provencio, D., Rodríguez-Diaz, J., Collado, M. C., Langella, P., Bermudez-Humaran, L.

G., & Monedero, V. (2012). Functional analysis of the Lactobacillus casei BL23 sortases.


Musilova, S., Rada, V., Vlkova, E., Bunesova, V., & Nevoral, J. (2015). Colonisation of the gut by

bifidobacteria is much more common in vaginal deliveries than Caesarean sections. Acta

Paediatrica, 104(4), 184-186. doi: 10.1111/apa.12931.

Navarre, W. W., & Schneewind, O. (1999). Surface proteins of gram-positive bacteria and

mechanisms of their targeting to the cell wall envelope. Microbiology and Molecular Biology

Reviews : MMBR, 63(1), 174-229. doi: 1092-2172/99/$04.00+0.

Nikolic, M., López, P., Strahinic, I., Suárez, A., Kojic, M., Fernández-García, M., Topisirovic, L.,

Golic, N., & Ruas-Madiedo, P. (2012). Characterisation of the exopolysaccharide (EPS)-

producing Lactobacillus paraplantarum BGCG11 and its non-EPS producing derivative strains

as potential probiotics. International Journal of Food Microbiology, 158(2), 155-162.

doi: 10.1016/j.ijfoodmicro.2012.07.015.

Nishio, J., & Honda, K. (2012). Immunoregulation by the gut microbiota. Cellular and Molecular

Life Sciences, 69(21), 3635-3650. doi: 10.1007/s00018-012-0993-6.

O'Connell Motherway, M., Zomer, A., Leahy, S. C., Reunanen, J., Bottacini, F., Claesson, M. J.,

O'Brien, F., Flynn, K., Casey, P. G., Munoz, J. A., Kearney, B., Houston, A. M., O'Mahony, C.,

Higgins, D. G., Shanahan, F., Palva, A., de Vos, W. M., Fitzgerald, G. F., Ventura, M., O'Toole,

P. W., & van Sinderen, D. (2011). Functional genome analysis of Bifidobacterium breve

UCC2003 reveals type IVb tight adherence (Tad) pili as an essential and conserved host-

colonization factor. Proceedings of the National Academy of Sciences of the United States of

America, 108(27), 11217-11222. doi: 10.1073/pnas.1105380108.

Oehlke, S. (2013). Investigation of capsular polysaccharides of bifidobacteria using bioinformatic

and genetic approaches. Master Thesis. University of Ulm, Ulm, Germany.

O'Flaherty, S. J., & Klaenhammer, T. R. (2010). Functional and phenotypic characterization of a

protein from Lactobacillus acidophilus involved in cell morphology, stress tolerance and

adherence to intestinal cells. Microbiology, 156(11), 3360-3367. doi: 10.1099/mic.0.043158-0.

List of Literature

104

O'Mahony, L., McCarthy, J., Kelly, P., Hurley, G., Luo, F., Chen, K., O'Sullivan, G. C., Kiely, B.,

Collins, J. K., Shanahan, F., & Quigley, E. M. (2005). Lactobacillus and bifidobacterium in

irritable bowel syndrome: symptom responses and relationship to cytokine profiles.

Gastroenterology, 128(3), 541-551. doi: S0016508504021559.

Osswald, A., Westermann, C., Sun, Z., & Riedel, C. U. (2015). A Phytase-Based Reporter System

for Identification of Functional Secretion Signals in Bifidobacteria. PloS One, 10(6), e0128802.


Pallen, M. J., Chaudhuri, R. R., & Henderson, I. R. (2003). Genomic analysis of secretion systems.

Current Opinion in Microbiology, 6(5), 519-527. doi: 10.1016/j.mib.2003.09.005.

Pan W-H, Li P-L, Liu Z. (2006). The correlation between surface hydrophobicity and adherence of

Bifidobacterium strains from centenarians’ faeces. Anaerobe, 12(3), 148-152.

doi: 10.1016/j.anaerobe.2006.03.001.

Parsonnet, J., Friedman, G. D., Vandersteen, D. P., Chang, Y., Vogelman, J. H., Orentreich, N., &

Sibley, R. K. (1991). Helicobacter pylori infection and the risk of gastric carcinoma. The New

England Journal of Medicine, 325(16), 1127-1131. doi: 10.1056/NEJM199110173251603.

Patti, J. M., Allen, B. L., McGavin, M. J., & Hook, M. (1994). MSCRAMM-mediated adherence of

microorganisms to host tissues. Annual Review of Microbiology, 48(1), 585-617.

doi: 10.1146/annurev.mi.48.100194.003101.

Pearson, J. P., Ward, R., Allen, A., Roberts, N. B., & Taylor, W. H. (1986). Mucus degradation by

pepsin: comparison of mucolytic activity of human pepsin 1 and pepsin 3: implications in peptic

ulceration. Gut, 27(3), 243-248. PMCID: PMC1433406.

Pelzer, A., Schwarz, C., Knapp, A., Wirtz, A., Wilhelm, S., Smits, S., Schmitt, L., Funken, H., &

Jaeger, K. (2015). Functional expression, purification, and biochemical properties of subtilase

SprP from Pseudomonas aeruginosa. MicrobiologyOpen, 4(3),1-10. doi: 10.1002/mbo3.275.

Pérez PF, Minnaard Y, Disalvo EA, De Antoni GL. (1998). Surface properties of bifidobacterial

strains of human origin. Applied and Environmental Microbiology, 64(1), 21-26. doi: 0099-

2240/98/$04.00+0.

Petersen, T. N., Brunak, S., von Heijne, G., & Nielsen, H. (2011). SignalP 4.0: discriminating

signal peptides from transmembrane regions. Nature Methods, 8(10), 785-786. doi:

10.1038/nmeth.1701.

Peterson, L. W., & Artis, D. (2014). Intestinal epithelial cells: regulators of barrier function and

immune homeostasis. Nature Reviews Immunology, 14(3), 141-153. doi: 10.1038/nri3608.

Philippe, D., Heupel, E., Blum-Sperisen, S., & Riedel, C. U. (2011). Treatment with

Bifidobacterium bifidum 17 partially protects mice from Th1-driven inflammation in a

chemically induced model of colitis. International Journal of Food Microbiology, 149(1), 45-

49. doi: 10.1016/j.ijfoodmicro.2010.12.020.

Pohlschröder, M., Prinz, W. A., Hartmann, E., & Beckwith, J. (1997). Protein translocation in the

three domains of life: variations on a theme. Cell, 91(5), 563-566. doi: S0092-8674(00)80443-2.

Pospiech, A., & Neumann, B. (1995). A versatile quick-prep of genomic DNA from Gram-positive

bacteria. Trends in Genetics, 11(6), 217-218. doi: S0168-9525(00)89052-6.

Poupard, J. A., Husain, I., & Norris, R. F. (1973). Biology of the bifidobacteria. Bacteriological

Reviews, 37(2), 136-165. PMID: 4578756.

List of Literature

105

Preising, J., Philippe, D., Gleinser, M., Wie, H., Blum, S., Eikmanns, B. J., Niess, J.-H., Riedel, C.

U.. (2010). Selection of Bifidobacteria based on adhesion and anti-inflammatory capacity in

vitro for amelioration of murine colitis. Applied Environmental Microbiology, 76(9), 3048-

3051. doi: 10.1128/AEM.03127-09.

Rafter, J., Bennett, M., Caderni, G., Clune, Y., Hughes, R., Karlsson, P. C., Klinder, A., O'Riordan,

M., O'Sullivan, G. C., Pool-Zobel, B., Rechkemmer, G., Roller, M., Rowland, I., Salvadori, M.,

Thijs, H., Van Loo, J., Watzl, B., & Collins, J. K. (2007). Dietary synbiotics reduce cancer risk

factors in polypectomized and colon cancer patients. The American Journal of Clinical

Nutrition, 85(2), 488-496. PMID: 17284748.

Ramos, C. L., Thorsen, L., Schwan, R. F., & Jespersen, L. (2013). Strain-specific probiotics

properties of Lactobacillus fermentum, Lactobacillus plantarum and Lactobacillus brevis

isolates from Brazilian food products. Food Microbiology, 36(1), 22-29.

doi: 10.1016/j.fm.2013.03.010.

Reuter, G. (1997). Present and Future of Probiotics in Germany and in Central Europe. Bioscience

and Microflora, 16(2), 43-51. doi: 10.12938/bifidus1996.16.43.

Reuter, G. (2001). The Lactobacillus and Bifidobacterium microflora of the human intestine:

composition and succession. Current Issues in Intestinal Microbiology, 2(2), 43-53.

PMID: 11721280.

Reynolds, E. S. (1963). The use of lead citrate at high pH as an electron-opaque stain in electron

microscopy. The Journal of Cell Biology, 17, 208-212. PMID: 13986422.

Riedel, C. U., Foata, F., Philippe, D., Adolfsson, O., Eikmanns, B. J., & Blum, S. (2006a). Anti-

inflammatory effects of bifidobacteria by inhibition of LPS-induced NF-kappaB activation.

World Journal of Gastroenterology, 12(23), 3729-3735. doi: 10.3748/wjg.v12.i23.3729.

Riedel, C. U., Foata, F., Goldstein, D. R., Blum, S., Eikmanns, B. J. (2006b). Interaction of

bifidobacteria with Caco-2 cells-adhesion and impact on expression profiles. International

Journal of Food Microbiology, 110(1), 62-68. doi: 10.1016/j.ijfoodmicro.2006.01.040.

Riedel, C. U., Monk, I. R., Casey, P. G., Morrissey, D., O'Sullivan, G. C., Tangney, M., Hill, C., &

Gahan, C. G. (2007). Improved luciferase tagging system for Listeria monocytogenes allows

real-time monitoring in vivo and in vitro. Applied and Environmental Microbiology, 73(9),

3091-3094. doi: 10.1128/AEM.02940-06.

Riedel, C. U., Schwiertz, A., Egert, M., & Marchesi, J. (2014). The stomach and small and large

intestinal microbiomes. In: Marchesi, J. R., The human microbiota and microbiome. pp 1-19.

CABI. ISBN: 9781780640495.

Rojas, M., Ascencio, F., & Conway, P. L. (2002). Purification and characterization of a surface

protein from Lactobacillus fermentum 104R that binds to porcine small intestinal mucus and

gastric mucin. Applied and Environmental Microbiology, 68(5), 2330-2336.

doi: 10.1128/AEM.68.5.2330-2336.2002.

Roller, M., Pietro Femia, A., Caderni, G., Rechkemmer, G., & Watzl, B. (2004). Intestinal

immunity of rats with colon cancer is modulated by oligofructose-enriched inulin combined

with Lactobacillus rhamnosus and Bifidobacterium lactis. The British Journal of Nutrition,

92(6), 931-938. doi: S0007114504002569.

Rosberg-Cody E, Liavonchanka A, Gobel C, Ross RP, O'Sullivan O, Fitzgerald GF, Feussner I,

Stanton C. (2011). Myosin-cross-reactive antigen (MCRA) protein from Bifidobacterium breve

List of Literature

106

is a FAD-dependent fatty acid hydratase which has a function in stress protection. BMC

Biochemistry, 12(9), 1-12. doi: 10.1186/1471-2091-12-9.

Rozario, T., & de Simone, D. W. (2010). The extracellular matrix in development and

morphogenesis: a dynamic view. Developmental Biology, 341(1), 126-140. doi:

10.1016/j.ydbio.2009.10.026.

Ruas-Madiedo, P., Hugenholtz, J., & Zoon, P. (2002). An overview of the functionality of

exopolysaccharides produced by lactic acid bacteria. International Dairy Journal, 12(2), 163-

171. doi: http://dx.doi.org/10.3168/jds.S0022-0302(03)73619-4.

Ruas-Madiedo, P., Gueimonde, M., Margolles, A., de los Reyes-Gavilan, C. G., & Salminen, S.

(2006). Exopolysaccharides produced by probiotic strains modify the adhesion of probiotics and

enteropathogens to human intestinal mucus. Journal of Food Protection, 69(8), 2011-2015.

PMID: 16924934.

Ruas-Madiedo, P., Moreno, J. A., Salazar, N., Delgado, S., Mayo, B., Margolles, A., & de Los

Reyes-Gavilan, C. G. (2007). Screening of exopolysaccharide-producing Lactobacillus and

Bifidobacterium strains isolated from the human intestinal microbiota. Applied and


Ruas-Madiedo, P., Gueimonde, M., Fernández-Garcia, M., de los Reyes-Gavilan, C. G., &

Margolles, A. (2008). Mucin degradation by Bifidobacterium strains isolated from the human

intestinal microbiota. Applied and Environmental Microbiology, 74(6), 1936-1940.

doi: 10.1128/AEM.02509-07.

Ruas-Madiedo, P., Gueimonde, M., Arigoni, F., de los Reyes-Gavilan, C. G., & Margolles, A.

(2009a). Bile affects the synthesis of exopolysaccharides by Bifidobacterium animalis. Applied

and Environmental Microbiology, 75(4), 1204-1207. doi: 10.1128/AEM.00908-08.

Ruas-Madiedo, P., Salazar, N., & Clara, G. (2009b). Biosynthesis and Chemical Composition of

Exopolysaccharides. In: Ullrich, M., Bacterial Polysaccharides: Current Innovations and

Future Trends. pp. 279-310. Horizon Scientific Press. ISBN. 9781904455455.

Ruas-Madiedo, P., Medrano, M., Salazar, N., De Los Reyes-Gavilan, C. G., Pérez, P. F., &

Abraham, A. G. (2010). Exopolysaccharides produced by Lactobacillus and Bifidobacterium

strains abrogate in vitro the cytotoxic effect of bacterial toxins on eukaryotic cells. Journal of

Applied Microbiology, 109(6), 2079-2086. doi: 10.1111/j.1365-2672.2010.04839.x.

Ruggiero, A., Tizzano, B., Pedone, E., Pedone, C., Wilmanns, M., & Berisio, R. (2009). Crystal

structure of the resuscitation-promoting factor ΔDUF RpfB from M. tuberculosis. Journal of

Molecular Biology, 385(1), 153-162. doi: 10.1016/j.jmb.2008.10.042.

Russell, T. M., Delorey, M. J., & Johnson, B. J. (2013). Borrelia burgdorferi BbHtrA degrades

host ECM proteins and stimulates release of inflammatory cytokines in vitro. Molecular

Microbiology, 90(2), 241-251. doi: 10.1111/mmi.12377.

Saarela, M., Mogensen, G., Fonden, R., Matto, J., & Mattila-Sandholm, T. (2000). Probiotic

bacteria: safety, functional and technological properties. Journal of Biotechnology, 84(3), 197-

215. doi: S0168165600003758.

Saavedra, J. M., Bauman, N. A., Oung, I., Perman, J. A., & Yolken, R. H. (1994). Feeding of

Bifidobacterium bifidum and Streptococcus thermophilus to infants in hospital for prevention of

diarrhoea and shedding of rotavirus. Lancet, 344(8929), 1046-1049. doi: S0140-6736(94)91708-

6.

List of Literature

107

Saiki, R. K., Scharf, S., Faloona, F., Mullis, K. B., Horn, G. T., Erlich, H. A., & Arnheim, N.

(1985). Enzymatic amplification of beta-globin genomic sequences and restriction site analysis

for diagnosis of sickle cell anemia. Science, 230(4732), 1350-1354. PMID: 2999980.

Salazar, N., Gueimonde, M., Hernandez-Barranco, A. M., Ruas-Madiedo, P., & de los Reyes-

Gavilan, C. G. (2008). Exopolysaccharides produced by intestinal Bifidobacterium strains act as

fermentable substrates for human intestinal bacteria. Applied and Environmental Microbiology,

74(15), 4737-4745. doi: 10.1128/AEM.00325-08.

Salazar, N., Prieto, A., Leal, J. A., Mayo, B., Bada-Gancedo, J. C., de los Reyes-Gavilan, C. G., &

Ruas-Madiedo, P. (2009). Production of exopolysaccharides by Lactobacillus and

Bifidobacterium strains of human origin, and metabolic activity of the producing bacteria in

milk. Journal of Dairy Science, 92(9), 4158-4168. doi: 10.3168/jds.2009-2126.

Salminen, S., Ouwehand, A., Benno, Y., & Lee, Y. K. (1999). Probiotics: how should they be

defined?. Trends in Food Science & Technology, 10(13), 107-110. doi: 10.1016/S0924-

2244(99)00027-8.

Samuelson, J. C., Chen, M., Jiang, F., Moller, I., Wiedmann, M., Kuhn, A., Phillips, G. J., &

Dalbey, R. E. (2000). YidC mediates membrane protein insertion in bacteria. Nature,

406(6796), 637-641. doi: 10.1038/35020586.

Sánchez, B., Bressollier, P., & Urdaci, M. C. (2008). Exported proteins in probiotic bacteria:

adhesion to intestinal surfaces, host immunomodulation and molecular cross-talking with the

host. FEMS Immunology and Medical Microbiology, 54(1), 1-17. doi: 10.1111/j.1574-

695X.2008.00454.x.

Sánchez, B., Urdaci, M. C., & Margolles, A. (2010). Extracellular proteins secreted by probiotic

bacteria as mediators of effects that promote mucosa-bacteria interactions. Microbiology,

156(11), 3232-3242. doi: 10.1099/mic.0.044057-0.

Sandhu, G. S., Precup, J. W., & Kline, B. C. (1989). Rapid one-step characterization of

recombinant vectors by direct analysis of transformed Escherichia coli colonies. Biotechniques,

7(7), 689-690. PMID: 2517212.

Sansonetti, P. J. (2004). War and peace at mucosal surfaces. Nature Reviews Immunology, 4(12),

953-964. doi: 10.1038/nri1499.

Sansonetti, P. J., & Medzhitov, R. (2009). Learning tolerance while fighting ignorance. Cell,

138(3), 416-420. doi: 10.1016/j.cell.2009.07.024.

Sansonetti, P. J. (2011). To be or not to be a pathogen: that is the mucosally relevant question.

Mucosal Immunology, 4(1), 8-14. doi: 10.1038/mi.2010.77.

Sato, K., Yomogida, K., Wada, T., Yorihuzi, T., Nishimune, Y., Hosokawa, N., & Nagata, K.

(2002). Type XXVI collagen, a new member of the collagen family, is specifically expressed in

the testis and ovary. The Journal of Biological Chemistry, 277(40), 37678-37684.

doi: 10.1074/jbc.M205347200.

Scardovi, V. (1924). The genus Bifidobacterium Orla-jensen. Volume 5: The Actinobacteria, In:

Whitman, W., Goodfellow, M., Kämpfer, P., Busse, H.-J., Trujillo, M., Ludwig, W. & Suzuki,

K.-I., Bergey´s manual of systematic bacteriology. pp. 1418-1434. Springer Science & Business

Media. ISBN: 9780387682334.

List of Literature

108

Schiebel, E., Driessen, A. J., Hartl, F., & Wickner, W. (1991). Δμ H and ATP function at different

steps of the catalytic cycle of preprotein translocase. Cell, 64(5), 927-939.

doi: http://dx.doi.org/10.1016/0092-8674(91)90317-R.

Schmidtchen, A., Frick, I., Andersson, E., Tapper, H., & Björck, L. (2002). Proteinases of common

pathogenic bacteria degrade and inactivate the antibacterial peptide LL‐37. Molecular


Schuster, M., Lostroh, C. P., Ogi, T., & Greenberg, E. P. (2003). Identification, timing, and signal

specificity of Pseudomonas aeruginosa quorum-controlled genes: a transcriptome analysis.

Journal of Bacteriology, 185(7), 2066-2079. doi: 10.1128/JB.185.7.2066-2079.2003.

Scotti, P. A., Urbanus, M. L., Brunner, J., de Gier, J. W., von Heijne, G., van der Does, C.,

Driessen, A. J., Oudega, B., & Luirink, J. (2000). YidC, the Escherichia coli homologue of

mitochondrial Oxa1p, is a component of the Sec translocase. The EMBO Journal, 19(4), 542-

549. doi: 10.1093/emboj/19.4.542.

Selbach, M., & Backert, S. (2005). Cortactin: an Achilles' heel of the actin cytoskeleton targeted by

pathogens. Trends in Microbiology, 13(4), 181-189. doi: S0966-842X(05)00053-3.

Seppo, L., Jauhiainen, T., Poussa, T., & Korpela, R. (2003). A fermented milk high in bioactive

peptides has a blood pressure-lowering effect in hypertensive subjects. The American Journal of

Clinical Nutrition, 77(2), 326-330. PMID: 12540390.

Serafini, F., Strati, F., Ruas-Madiedo, P., Turroni, F., Foroni, E., Duranti, S., Milano, F., Perotti,

A., Viappiani, A., Guglielmetti, S., Buschini, A., Margolles, A., van Sinderen, D., & Ventura,

M. (2013). Evaluation of adhesion properties and antibacterial activities of the infant gut

commensal Bifidobacterium bifidum PRL2010. Anaerobe, 21, 9-17.

doi: 10.1016/j.anaerobe.2013.03.003.

Serruto, D., Adu‐Bobie, J., Scarselli, M., Veggi, D., Pizza, M., Rappuoli, R., & Aricò, B. (2003).

Neisseria meningitidis App, a new adhesin with autocatalytic serine protease activity. Molecular


Servin, A. L. (2004). Antagonistic activities of lactobacilli and bifidobacteria against microbial

pathogens. FEMS Microbiology Reviews, 28(4), 405-440. doi: 10.1016/j.femsre.2004.01.003.

Sgorbati, B., Biavati, B., and Palenzona, D. (1995). The genus Bifidobacterium. In Wood, B. J. B.

and Holzapfel, W. H., The genera of Lactic Acid Bacteria. pp. 279-306. Springer US. ISBN:

978-1-4613-7666-8.

Shao, L., Wu, Z., Zhang, H., Chen, W., Ai, L., & Guo, B. (2014). Partial characterization and

immunostimulatory activity of exopolysaccharides from Lactobacillus rhamnosus KF5.

Carbohydrate Polymers, 107, 51-56. doi: 10.1016/j.carbpol.2014.02.037.

Sharp, P. A., Sugden, B., & Sambrook, J. (1973). Detection of two restriction endonuclease

activities in Haemophilus parainfluenzae using analytical agarose--ethidium bromide

electrophoresis. Biochemistry, 12(16), 3055-3063. doi: 10.1021/bi00740a018.

Sheng, Y., Mancino, V., & Birren, B. (1995). Transformation of Escherichia coli with large DNA

molecules by electroporation. Nucleic Acids Research, 23(11), 1990-1996. doi:

10.1093/nar/23.11.1990.

Singh, B., Fleury, C., Jalalvand, F., & Riesbeck, K. (2012). Human pathogens utilize host

extracellular matrix proteins laminin and collagen for adhesion and invasion of the host. FEMS

Microbiology Reviews, 36(6), 1122-1180. doi: 10.1111/j.1574-6976.2012.00340.x.

List of Literature

109

Sonnhammer, E. L., Eddy, S. R., Birney, E., Bateman, A., & Durbin, R. (1998). Pfam: multiple

sequence alignments and HMM-profiles of protein domains. Nucleic Acids Research, 26(1),

320-322. doi: 10.1093/nar/26.1.320.

Staudt, C. (2002). Wechselwirkungen von Bifidobakterien mit Darmepithelzellen und mit

extrazellulären Matrix – und Plasmaproteinen. Doctoral Thesis, University of Ulm, Ulm,

Germany.

Sun, Z., Westermann, C., Yuan, J., & Riedel, C. U. (2014). Experimental determination and

characterization of the gap promoter of Bifidobacterium bifidum S17. Bioengineered, 5(6), 371-

377. doi: 10.4161/bioe.34423.

Suresh Kumar, A., Mody, K., & Jha, B. (2007). Bacterial exopolysaccharides–a perception.

Journal of Basic Microbiology, 47(2), 103-117. doi: 10.1002/jobm.200610203.

Swidsinski, A., Loening-Baucke, V., Theissig, F., Engelhardt, H., Bengmark, S., Koch, S., Lochs,

H., & Dorffel, Y. (2007). Comparative study of the intestinal mucus barrier in normal and

inflamed colon. Gut, 56(3), 343-350. doi: 10.1136/gut.2006.098160.

Takai, T., & Ono, M. (2001). Activating and inhibitory nature of the murine paired

immunoglobulin-like receptor family. Immunological Reviews, 181, 215-222. doi:

10.1034/j.1600-065X.2001.1810118.

Tallon, R., Bressollier, P., & Urdaci, M. C. (2003). Isolation and characterization of two

exopolysaccharides produced by Lactobacillus plantarum EP56. Research in Microbiology,

154(10), 705-712. doi: 10.1016/j.resmic.2003.09.006.

Tamboli, C. P., Neut, C., Desreumaux, P., & Colombel, J. F. (2004). Dysbiosis in inflammatory

bowel disease. Gut, 53(1), 1-4. doi: 10.1136/gut.53.1.1.

Tap, J., Mondot, S., Levenez, F., Pelletier, E., Caron, C., Furet, J. P., Ugarte, E., Munoz-Tamayo,

R., Paslier, D. L., Nalin, R., Dore, J., & Leclerc, M. (2009). Towards the human intestinal

microbiota phylogenetic core. Environmental Microbiology, 11(10), 2574-2584. doi:

10.1111/j.1462-2920.2009.01982.x.

Timmer, A. M., Kristian, S. A., Datta, V., Jeng, A., Gillen, C. M., Walker, M. J., Beall, B., &

Nizet, V. (2006). Serum opacity factor promotes group A streptococcal epithelial cell invasion

and virulence. Molecular Microbiology, 62(1), 15-25. doi: 10.1111/j.1365-2958.2006.05337.x.

Tissier, M. H. (1900). Recherches sur la flore intestinale des nourrissons état normal et

pathologique. Doctoral Thesis, University of Paris, Paris, France.

Tomich, M., Planet, P. J., & Figurski, D. H. (2007). The tad locus: postcards from the widespread

colonization island. Nature Reviews.Microbiology, 5(5), 363-375. doi: 10.1038/nrmicro1636.

Tran, S. L., Guillemet, E., Gohar, M., Lereclus, D., & Ramarao, N. (2010). CwpFM (EntFM) is a

Bacillus cereus potential cell wall peptidase implicated in adhesion, biofilm formation, and

virulence. Journal of Bacteriology, 192(10), 2638-2642. doi: 10.1128/JB.01315-09.

Turnbaugh, P. J., Ley, R. E., Mahowald, M. A., Magrini, V., Mardis, E. R., & Gordon, J. I. (2006).

An obesity-associated gut microbiome with increased capacity for energy harvest. Nature,

444(7122), 1027-1031. doi: 10.1038/nature05414.

Turroni, F., Peano, C., Pass, D. A., Foroni, E., Severgnini, M., Claesson, M. J., Kerr, C.,

Hourihane, J., Murray, D., Fuligni, F., Gueimonde, M., Margolles, A., De Bellis, G., O'Toole, P.

W., van Sinderen, D., Marchesi, J. R., & Ventura, M. (2012). Diversity of bifidobacteria within

the infant gut microbiota. PloS One, 7(5), e36957. doi: 10.1371/journal.pone.0036957.

List of Literature

110

Turroni, F., Serafini, F., Foroni, E., Duranti, S., O'Connell Motherway, M., Taverniti, V.,

Mangifesta, M., Milani, C., Viappiani, A., Roversi, T., Sánchez, B., Santoni, A., Gioiosa, L.,

Ferrarini, A., Delledonne, M., Margolles, A., Piazza, L., Palanza, P., Bolchi, A., Guglielmetti,

S., van Sinderen, D., & Ventura, M. (2013). Role of sortase-dependent pili of Bifidobacterium

bifidum PRL2010 in modulating bacterium-host interactions. Proceedings of the National

Academy of Sciences of the United States of America, 110(27), 11151-11156. doi:

10.1073/pnas.1303897110.

Turroni, F., Serafini, F., Mangifesta, M., Arioli, S., Mora, D., van Sinderen, D., & Ventura, M.

(2014). Expression of sortase-dependent pili of Bifidobacterium bifidum PRL2010 in response

to environmental gut conditions. FEMS Microbiology Letters, 357(1), 23-33. doi:

10.1111/1574-6968.12509.

Upadhyay, S. K., Gautam, P., Pandit, H., Singh, Y., Basir, S. F., & Madan, T. (2012). Identification

of fibrinogen-binding proteins of Aspergillus fumigatus using proteomic approach.

Mycopathologia, 173(2-3), 73-82. doi: 10.1007/s11046-011-9465-z.

Vallet, I., Diggle, S. P., Stacey, R. E., Camara, M., Ventre, I., Lory, S., Lazdunski, A., Williams,

P., & Filloux, A. (2004). Biofilm formation in Pseudomonas aeruginosa: fimbrial cup gene

clusters are controlled by the transcriptional regulator MvaT. Journal of Bacteriology, 186(9),

2880-2890. doi: 10.1128/JB.186.9.2880-2890.2004.

van der Wolk, J., Klose, M., Breukink, E., Demel, R. A., de Kruijff, B., Freudl, R., & Driessen, A.

J. (1993). Characterization of a Bacillus subtilis SecA mutant protein deficient in translocation

ATPase and release from the membrane. Molecular Microbiology, 8(1), 31-42. doi:

10.1111/j.1365-2958.1993.tb01200.x.

van der Wolk, J. P., Klose, M., de Wit, J. G., den Blaauwen, T., Freudl, R., & Driessen, A. J.

(1995). Identification of the magnesium-binding domain of the high-affinity ATP-binding site

of the Bacillus subtilis and Escherichia coli SecA protein. The Journal of Biological Chemistry,

270(32), 18975-18982. PMID: 7642557.

van Pijkeren, J. P., Canchaya, C., Ryan, K. A., Li, Y., Claesson, M. J., Sheil, B., Steidler, L.,

O'Mahony, L., Fitzgerald, G. F., van Sinderen, D., & O'Toole, P. W. (2006). Comparative and

functional analysis of sortase-dependent proteins in the predicted secretome of Lactobacillus

salivarius UCC118. Applied and Environmental Microbiology, 72(6), 4143-4153. doi:

10.1128/AEM.03023-05.

van Tassell, M. L., & Miller, M. J. (2011). Lactobacillus adhesion to mucus. Nutrients, 3(5), 613-

636. doi: 10.3390/nu3050613.

van Wely, K. H., Swaving, J., Freudl, R., & Driessen, A. J. (2001). Translocation of proteins across

the cell envelope of Gram-positive bacteria. FEMS Microbiology Reviews, 25(4), 437-454. doi:

http://dx.doi.org/10.1111/j.1574-6976.2001.tb00586.x.

Vásquez, A., Forsgren, E., Fries, I., Paxton, R. J., Flaberg, E., Szekely, L., & Olofsson, T. C.

(2012). Symbionts as major modulators of insect health: lactic acid bacteria and honeybees.

PloS One, 7(3), e33188. doi: 10.1371/journal.pone.0033188.

Vélez, M. P., De Keersmaecker, S. C., & Vanderleyden, J. (2007). Adherence factors of

Lactobacillus in the human gastrointestinal tract. FEMS Microbiology Letters, 276(2), 140-148.

doi: http://dx.doi.org/10.1111/j.1574-6968.2007.00908.x.

List of Literature

111

Ventura, M., Turroni, F., Motherway, M. O., MacSharry, J., & van Sinderen, D. (2012). Host–

microbe interactions that facilitate gut colonization by commensal bifidobacteria. Trends in

Microbiology, 20(10), 467-476. doi: 10.1016/j.tim.2012.07.002.

Ventura, M., Canchaya, C., Tauch, A., Chandra, G., Fitzgerald, G. F., Chater, K. F., & van

Sinderen, D. (2007). Genomics of Actinobacteria: tracing the evolutionary history of an ancient

phylum. Microbiology and Molecular Biology Reviews, 71(3), 495-548. doi:

10.1128/MMBR.00005-07.

Ventura, M., van Sinderen, D., Fitzgerald, G. F., & Zink, R. (2004). Insights into the taxonomy,

genetics and physiology of bifidobacteria. Antonie Van Leeuwenhoek, 86(3), 205-223. doi:

10.1023/B:ANTO.0000047930.11029.ec.

Volkov, A., Liavonchanka, A., Kamneva, O., Fiedler, T., Goebel, C., Kreikemeyer, B., & Feussner,

I. (2010). Myosin cross-reactive antigen of Streptococcus pyogenes M49 encodes a fatty acid

double bond hydratase that plays a role in oleic acid detoxification and bacterial virulence. The

Journal of Biological Chemistry, 285(14), 10353-10361. doi: 10.1074/jbc.M109.081851.

von Ossowski, I., Reunanen, J., Satokari, R., Vesterlund, S., Kankainen, M., Huhtinen, H.,

Tynkkynen, S., Salminen, S., de Vos, W. M., & Palva, A. (2010). Mucosal adhesion properties

of the probiotic Lactobacillus rhamnosus GG SpaCBA and SpaFED pilin subunits. Applied and


von Schillde, M. A., Hormannsperger, G., Weiher, M., Alpert, C. A., Hahne, H., Bauerl, C., van

Huynegem, K., Steidler, L., Hrncir, T., Pérez-Martinez, G., Kuster, B., & Haller, D. (2012).

Lactocepin secreted by Lactobacillus exerts anti-inflammatory effects by selectively degrading

proinflammatory chemokines. Cell Host & Microbe, 11(4), 387-396.

doi: 10.1016/j.chom.2012.02.006.

Wang, K., Li, W., Rui, X., Chen, X., Jiang, M., & Dong, M. (2014). Characterization of a novel

exopolysaccharide with antitumor activity from Lactobacillus plantarum 70810. International

Journal of Biological Macromolecules, 63, 133-139. doi: 10.1016/j.ijbiomac.2013.10.036.

Wang, T., Zhang, J., Zhang, X., Xu, C., & Tu, X. (2013). Solution structure of the Big domain

from Streptococcus pneumoniae reveals a novel Ca2 -binding module. Scientific Reports,

3(1079), doi: 10.1038/srep01079.

Weiss, G., & Jespersen, L. (2010). Transcriptional analysis of genes associated with stress and

adhesion in Lactobacillus acidophilus NCFM during the passage through an in vitro

gastrointestinal tract model. Journal of Molecular Microbiology and Biotechnology, 18(4), 206-

214. doi: 10.1159/000316421.

Westerlund, B., & Korhonen, T. (1993). Bacterial proteins binding to the mammalian extracellular

matrix. Molecular Microbiology, 9(4), 687-694. doi: 10.1111/j.1365-2958.1993.tb01729.x.

Westermann, C., Zhurina, D., Baur, A., Shang, W., Yuan, J., & Riedel, C. U. (2012a). 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.

Westermann, C. (2012b). Analysis of potential host-colonization and probiotic factors of different

bifidobacteria. Master Thesis, University of Ulm, Ulm, Germany.

Wheeler, T. J., & Eddy, S. R. (2013). nhmmer: DNA homology search with profile HMMs.

Bioinformatics, 29(19), 2487-2489. doi: 10.1093/bioinformatics/btt403.

List of Literature

112

Whitman, W. B., Coleman, D. C., & Wiebe, W. J. (1998). Prokaryotes: the unseen majority.

Proceedings of the National Academy of Sciences of the United States of America, 95(12),

6578-6583. PMID: 9618454.

Whorwell, P. J., Altringer, L., Morel, J., Bond, Y., Charbonneau, D., O'Mahony, L., Kiely, B.,

Shanahan, F., & Quigley, E. M. (2006). Efficacy of an encapsulated probiotic Bifidobacterium

infantis 35624 in women with irritable bowel syndrome. The American Journal of

Gastroenterology, 101(7), 1581-1590. doi: 10.1111/j.1572-0241.2006.00734.x.

Wikström, M., & Linde, A. (1986). Ability of oral bacteria to degrade fibronectin. Infection and

Immunity, 51(2), 707-711. doi: 0019-9567/86/020707-05$02.00/0.

Williams, R. J., Henderson, B., Sharp, L. J., & Nair, S. P. (2002). Identification of a fibronectin-

binding protein from Staphylococcus epidermidis. Infection and Immunity, 70(12), 6805-6810.

doi: 10.1128/IAI.70.12.6805-6810.2002.

Wilson, M. (2009). Bacteriology of humans: an ecological perspective. John Wiley & Sons. ISBN:

9781444300383.

Wollowski, I., Rechkemmer, G., & Pool-Zobel, B. L. (2001). Protective role of probiotics and

prebiotics in colon cancer. The American Journal of Clinical Nutrition, 73(2), 451-455.

Woods, D. E., Straus, D. C., Johanson, W. G.,Jr, & Bass, J. A. (1981). Role of fibronectin in the

prevention of adherence of Pseudomonas aeruginosa to buccal cells. The Journal of Infectious

Diseases, 143(6), 784-790. PMID: 11157356.

Wu, M. H., Pan, T. M., Wu, Y. J., Chang, S. J., Chang, M. S., & Hu, C. Y. (2010).

Exopolysaccharide activities from probiotic bifidobacterium: Immunomodulatory effects (on

J774A.1 macrophages) and antimicrobial properties. International Journal of Food

Microbiology, 144(1), 104-110. doi: 10.1016/j.ijfoodmicro.2010.09.003.

Xie, K., & Dalbey, R. E. (2008). Inserting proteins into the bacterial cytoplasmic membrane using

the Sec and YidC translocases. Nature Reviews.Microbiology, 6(3), 234-244. doi:

10.1038/nrmicro1845.

Xu, Q., Chiu, H., Farr, C. L., Jaroszewski, L., Knuth, M. W., Miller, M. D., Lesley, S. A., Godzik,

A., Elsliger, M., & Deacon, A. M. (2014). Structures of a bifunctional cell wall hydrolase CwlT

containing a novel bacterial lysozyme and an NlpC/P60 DL-endopeptidase. Journal of

Molecular Biology, 426(1), 169-184. doi: 10.1016/j.jmb.2013.09.011.

Yadav, A. K., Tyagi, A., Kaushik, J. K., Saklani, A. C., Grover, S., & Batish, V. K. (2013). Role of

surface layer collagen binding protein from indigenous Lactobacillus plantarum 91 in adhesion

and its anti-adhesion potential against gut pathogen. Microbiological Research, 168(10), 639-

645. doi: 10.1016/j.micres.2013.05.003.

Yamamoto, N., Ono, H., Maeno, M., Ueda, Y., Takano, T., & Momose, H. (1998). Classification

of Lactobacillus helveticus strains by immunological differences in extracellular proteinases.

Bioscience, Biotechnology, and Biochemistry, 62(6), 1228-1230.

doi: http://doi.org/10.1271/bbb.62.1228

Yatsunenko, T., Rey, F. E., Manary, M. J., Trehan, I., Dominguez-Bello, M. G., Contreras, M.,

Magris, M., Hidalgo, G., Baldassano, R. N., & Anokhin, A. P. (2012). Human gut microbiome

viewed across age and geography. Nature, 486(7402), 222-227. doi: 10.1038/nature11053.

List of Literature

113

You, J., & Yaqoob, P. (2012). Evidence of immunomodulatory effects of a novel probiotic,

Bifidobacterium longum bv. infantis CCUG 52486. FEMS Immunology and Medical

Microbiology, 66(3), 353-362. doi: 10.1111/j.1574-695X.2012.01014.x.

Young, V. B. (2012). The intestinal microbiota in health and disease. Current Opinion in

Gastroenterology, 28(1), 63-69. doi: 10.1097/MOG.0b013e32834d61e9.

Zdobnov, E. M., & Apweiler, R. (2001). InterProScan--an integration platform for the signature-

recognition methods in InterPro. Bioinformatics, 17(9), 847-848.

doi: 10.1093/bioinformatics/17.9.847.

Zhang, L., Seiffert, D., Fowler, B. J., Jenkins, G. R., Thinnes, T. C., Loskutoff, D. J., Parmer, R. J.,

& Miles, L. A. (2002). Plasminogen Has a Broad Extrahepatic Distribution. Thrombosis

Haemostasis, 87(3), 493-501. PMID: 11916082.

Zhang, W., Wang, H., Gao, K., Wang, C., Liu, L., & Liu, J. (2015). Lactobacillus reuteri

glyceraldehyde-3-phosphate dehydrogenase functions in adhesion to intestinal epithelial cells.

Canadian Journal of Microbiology, 61(5), 373-380. doi: 10.1139/cjm-2014-0734.

Zhurina, D., Zomer, A., Gleinser, M., Brancaccio, V. F., Auchter, M., Waidmann, M. S.,

Westermann, C., van Sinderen, D., Riedel, C. U. (2011). Complete genome sequence of

Bifidobacterium bifidum S17. Journal of Bacteriology, 193(1), 301-302. doi: 10.1128/JB.01180-

10.

Zhurina, D., Dudnik, A., Waidmann, M. S., Grimm, V., Westermann, C., Breitinger, K. J., Yuan,

J., van Sinderen, D., & Riedel, C. U. (2013). High-Quality Draft Genome Sequence of

Bifidobacterium longum E18, Isolated from a Healthy Adult. Genome Announcements, 1(6), 1-

2. doi: 10.1128/genomeA.01084-13.

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).

List of Literature

114

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.

Addendum

115

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

Addendum

116

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).


Name Sequence Amplicon size [bp]

Purpose

Pili encoding gene cluster

fim

1 c

lust

er

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 bbif_0302 301/302_rev 5´- AGGTGACCAACGTCAAGTCC-3´

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 bbif_0303 302/303_rev 5´- GCATGATGATGAGCTGTATGC-3´

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

BBIF303_rev 5´- GTCATGATGGTGCGCGTGTCATG -3´

fim

2 cl

ust

er

BBIF1648_fw 5´- CGCTGCTGACGCCATCGGCGTCGATC-3´ 529 RT-PCR for bbif_1648

BBIF_1648_rev 5´- CGTGACCGTCGAATACAAGG-3´

1648/1649_fw 5´- CTTGAACGTGTGTCCAGTCAATTGGTCCGC -3´

459 RT-PCR for intergenic region between bbif_1648 and bbif_1649 1648/1649_rev 5´- CTCCATCGCAAAGACCAAGGACGACGAG -3´

Addendum

117



Purpose

BBIF1649_fw 5´- GGCTACTGTCATCCTTGTGGAAC -3´

521 RT-PCR for bbif_1649 BBIF1649_rev 5´- CGTTCCTGAACTTCCTCATCCTG -3´

1649/1650_fw 5´- CGTTGAGGACGAGATCGTAGATGAGG -3´

631 RT-PCR for intergenic region between bbif_1649 and bbif_1650 1649/1650_rev 5´- GGTGATGGGCGAGACCTTGGGCTACAAGG -3´

BBIF1650_fw 5´- GTGGTAGATGGGCAGGTCGACCCCAATC -3´ 424 RT-PCR for bbif_1650

BBIF1650_rev 5´- GAGCCGTCCTGGGCCTGGAGG -3´

fim

3 c

lust

er

RT_1760fw 5´- GATGTCCACGTGGAATGTGG -3´ 529 RT-PCR for bbif_1760

RT_1760rev 5´- GAGAGACCGACGCTCTTGCC -3´

1760/1761_fw 5´- CGTCACCGCTCACGGTGGCGGAGAAC -3´ 588

RT-PCR for intergenic region between bbif_1761 and bbif_1760 1760/1761_rev 5´- CCAATGGCAAGGACATCGTGCGTTAC -3´

BBIF1761_fw 5´- TGAGCGTGTACTCGCCGTTCTTC -3´ 619 RT-PCR for bbif_1761

BBIF1761_rev 5´- CATGTCCGCCTATGACACCTAC -3´

1761/1762_fw 5´- CGTGCGCAGCATCCAGCACCTTCTGG -3´ 586

RT-PCR for intergenic region between bbif_1761 and bbif_1762 1761/1762_rev 5´- GGGGACTGAAGAACGGCGAGTACACG -3´

BBIF1762_fw 5´- CGTAGCCTGTGGAAAGATCCTTC -3´ 500

RT-PCR for bbif_1762 (proximal part) BBIF1762_rev 5´- GTGAAACCAGCCGTGCCTATTTG -3´

tad

clu

ster

TadE_fw 5´- CACCGCCATCCTGTGGCCGAG -3´ 161 RT-PCR for bbif_1698 (tadE)

TadE_rev 5´- GTCCGTCAACGCCGTGGCGAG -3´

Flp_fw 5´- TCATCCCACACTCAATGACTTC -3´ 290 RT-PCR for bbif_1699 (flp)

Flp_rev 5´- ATGAATATCGAGGAGACGATG -3´

fim

2 c

lust

er

Addendum

118



Purpose

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´

TadV_fw 5´- GATCGAACCTTAGCCACAGCG -3´ 287 RT-PCR for bbif_0835 (tadV)

TadV_rev 5´- GGATCGCCATCGGGTGCCTGAC -3´

16

SrR

NA

16S_fw 5´- GGAGGTGATCCAGCCGCACCTTC -3´ 243

RT-PCR for bbif_1306 (16S ribosomal RNA)

16S_rev 5´- GTAAACGGTGGACGCTGGATGTG -3´

Putative single adhesins

pC

W

pCW_cont_fw 5´- GAGTAAACTTGGTCTACAG -3´ -

Forward primer to control the integration of a single adhesin into the vector (binds in the plasmid region), also used for sequencing

pCW_cont_rev 5´- GTCTGAGCCGTCGGCGTCCGC-3´ -

Reversed primer for sequencing of single adhesin encoding genes after ligation with the vector

tad

clu

ster

Addendum

119



Purpose P

hyp

er

Phyp_NdeI_fw 5´- GACTACCATATGTTCTTGAAGACGAAAGGGCC -3

98

Forward primer for amplification of the promoter Phyper with NdeI restriction site

Phyp_XhoI_rev 5´- GACCTCGAGTCTCCTTCTACATTTTTAAC -3

Reversed primer for amplification of the promoter Phyper with XhoI restriction site

bb

if_0

295

BBIF0295_XhoI_fw 5´- GACTACCTCGAGGTGACGTACGCCGCCGATACC -3´ 564

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

BBIF0295_cont_rev 5´- CGGACTTCTCCTTGACGGTG -3´ 537

(with pCW_cont_fw)

For colony PCR, to control the integration of the gene bbif_0295 into the vector

BBIF_0295_BglII_rev 5´- GACAGATCTTTAATGATGATGATGATGATGGATGTCGCGACG -3´ 564

(with BBIF0295_XhoI_fw)

For amplification of bbif_0295 with XhoI and BglII restriction site, with His6-tag, for cloning into pVG_mCherry

BBIF0295_fw 5´- CGATACCTCGCTTACCGCCAC -3´ 291

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

bb

if_0

450

BBIF0450_XhoI_fw 5´- GACTACCTCGAGATGAGCAAGCAGCCTAACCAG -3´

6018

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

Addendum

120



Purpose

BBIF0450_cont_rev 5´- GCTGGACGATGTCGAAACG -3´ 539

(with pCW_cont_fw)


BBIF0450_fw 5´- GTTGACGCAGTTGAGGTCGGAG -3´ 486 RT-PCR for bbif_0450

BBIF0450_rev 5´- GACCAAGCAGCCTAACCAGCG -3´

BBIF0450_seq1 5´- GCATCCATGCGCAAACGGTC -3´ -

Primer for sequencing

BBIF0450_seq2 5´- CCGTTGACGCTGCAATACCC -3´ -

BBIF0450_seq3 5´- GAACGCACTCGGCAGTTTCC -3´ -

BBIF0450_seq4 5´- GCAGAAGCTGCAGTACGACAAG -3´ -

BBIF0450_seq5 5´- GCACCGCGAACGAACTGAAG -3´ -

BBIF0450_seq6 5´- GACGATTCGGCGAAGCTGAC -3´ -

BBIF0450_seq7 5´- CCCGTGACCATCATCTATTC -3´ -

BBIF0450_seq8 5´- CTCGGCAACATGCACAACG -3´ -

BBIF0450_seq9 5´- GTATCCGCCGACGATTTCAC -3´ -

BBIF0450_seq10 5´- CACAAGCGCGTGTATCTG -3´ -

BBIF0450_seq11 5´- CGATGTCTCGCAGTGGAT -3´ -

BBIF0540_XhoI_fw 5´- GACTACCTCGAGATGTATTATTCCAGTGGCAAC -3´

1917 For amplification of bbif_0540 with XhoI and HindIII restriction site, with His6-tag BBIF0540_HindIII_rev

5´- GACAAGCTTTCAATGATGATGATGATGATGGATCGCCCCGTACTCGCG -3´

BBIF0540_cont_rev 5´- GTCCTTGTTGAGCCAGTAATAC -3´ 592

(with pCW_cont_fw)

For colony PCR, to control the integration of the gene bbif_0540 into the vector, also used for sequencing of a b

bif

_054

0

Bb

if_0

45

0

Addendum

121



Purpose

BBIF0540_fw 5´- CAGTGGCAACTATGAAGCGTTC -3´ 536 RT-PCR for bbif_0540

BBIF0540_rev 5´- CAGTACAGCCAGAAGTTGG -3´

bb

if_0

66

5

BBIF0665_XhoI_fw 5´- GACTACCTCGAGATGAGACCGATGACATTCGC -3´


5´- GACAAGCTTTCAATGATGATGATGATGATGTAGCATCCTCCGTGACATG -3´

BBIF0665_cont_rev 5´- CATACAGCACGATGATGTC -3´ 821

(with pCW_cont_fw)

For colony PCR, to control the integration of the gene bbif_0665 into the vector, also used for sequencing of a part of the gene

BBIF0665_fw 5´- CTGATTGATTTG AGCGACAGC -3´

432

RT-PCR for bbif_0665

BBIF0665_rev 5´- GCTCCAGCGATGAGACGCTGA -3´

bb

if_1

426

BBIF1426_XhoI_fw 5´- GACTACCTCGAGATGCCTGTGTGCTACAATCATGCC -3´ 846

For amplification of bbif_0665 with XhoI and HindIII restriction site, with His6-tag BBIF1426_HindIII_rev 5´- GACAAGCTTTTAATGATGATGATGATGATGGATGATTCGGTAGTAC -3´

BBIF1426_cont_rev 5´- CACGTTCAGGGATTCCACAC -3´ 505

(with pCW_cont_fw)


BBIF1426_fw 5´- GCTCCTCTTCGGCCTTCTTGG -3´

324 RT-PCR for bbif_1426 BBIF1426_rev 5´- CCTGTGTGCTACAATCATGCCTC -3´

Addendum

122



Purpose b

bif

_15

38

BBIF1538_XhoI_fw 5´-GACTACCTCGAGATGAACCGAAGAATATCTATG -3´

5289

For amplification of bbif_1538 with XhoI and HindIII restriction site, with His6-tag

BBIF1538_HindIII_rev 5´- GACAAGCTTTCAATGATGATGATGATGATGGCGCTTGGCCAGT -3´

BBIF1538_cont_rev 5´- GACCGTATACCAGATGTCCTC -3´ 538

(with pCW_cont_fw)


BBIF1538_fw 5´- GTCGATCAGCGCGTTCGTG -3´ 377


BBIF1538_rev 5´- CAGACCACCACCATCCCG -3´

bb

if_1

76

9

BBIF1769_XhoI_fw 5´- GACTACCTCGAGATGGCCAAGGCGGTGGACACCAG -3´ 1572

For amplification of bbif_1769 with XhoI and HindIII restriction site, with His6-tag BBIF1769_HindIII_rev 5´- GACAAGCTTTCAATGATGATGATGATGATGGTACCAGCCGATTTG -3´

BBIF1769_cont_rev 5´- GCTGATGGTCGTCTGATATGC -3´

544

(with pCW_cont_fw)


BBIF1769_fw 5´- GTGGACACCACGACGTTTTG -3´

549


BBIF1769_rev 5´- CTTGTTGAGGCTGATGCCTTTG -3´

Addendum

123



Purpose

Subtilisin like protease bifidocepin

bb

if_1

68

1

BBIF1681_XhoI_fw 5´- GACTACCTCGAGATGGCCAATAAGCAATGGCCTCGCTGG -3´


5´- TACAAGCTTCTAATGATGATGATGATGATGGCGACGCACCGCGTCACG -3´

BBIF1681_cont_rev 5´- GCTCACCGCGGATCGCAAGACC -3´ 2338

(with pCW_cont_fw)


BBIF1681_seq1 5´- GAAGAAGCCGTTCTTGTGCAGC -3´ -

Primer for sequencing BBIF1681_seq2 5´- GCATCATCCTCCTGGGTGAC -3´ -

BBIF1681_seq3 5´- GCGTAGTTCACGGAGTACTGGC -3´ -

Addendum

124

8.3. Chemicals and manufacturers

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

Table 16.

Table 16: Chemicals and manufacturers.


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

Addendum

125



Di-sodium ethylendiaminetetraacetate

dehydrate

(EDTA)

Carl Roth GmbH & Co., Karlsruhe, Germany

Di-sodium hydrogenphosphate heptahydrogen

(Na2HPO4 x 7 H2O)

Merck KGaA, Mannheim, Germany

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

Addendum

126



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

Magnesium sulfate heptahydrate (MgSO4 x 7

H2O)

Sigma Aldrich Chemie GmbH, Steinheim,

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

(Catalog # 11922416001)

Roche Diagnostics GmbH, Mannheim,

Germany

Mowiol® 4-88 Sigma Aldrich Chemie GmbH, Steinheim,

Germany

N-2-Hydroxyetylpiperazine-N´-2-Ethanesulfonic

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

radish peroxidase coupled (Catalog # P0260)

Dako Deutschland GmbH, Hamburg, Germany

RevertAidTM Premium First Strand cDNA

Synthesis Kit

MBI Fermentas GmbH, St. Leon-Rot, Germany

RNase-free water (H2O) Life Technologies, Darmstadt, Germany

Roti®-Aqua-Phenol (acidic phenol) Carl Roth GmbH & Co., Karlsruhe, Germany

Roti®-Phenol


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

Addendum

127



(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

(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

(Tris-HCl)

Sigma Aldrich Chemie GmbH, Steinheim,

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

Abbreviations

128

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

Abbreviations

129

e.g. Latin èxemplȋ grȃtiȃ´; English: for example

et al. Llatin èt 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 ìd 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

Abbreviations

130

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

Abbreviations

131

Tris Tris(hydroxymethyl)-aminomethan

TM `trademark´

U unit

UV ultraviolet

V volt

v/v volume per volume

wt wildtype

w/v weight per volume

Acknowledgment

132

10. Acknowledgment

Der Inhalt dieser Seite wurde aus Datenschutzgründen entfernt.

Acknowledgment

133


Curriculum Vitae

134

Curriculum Vitae - Christina Westermann


Curriculum Vitae

135


Curriculum Vitae

136


137

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

analysis of potential host-colonization factors in

Documents