Mucus-Pathogen Interactions in the Gastrointestinal Tract of Farmed Animals

Abstract

Gastrointestinal infections cause significant challenges and economic losses in animal husbandry. As pathogens becoming resistant to antibiotics are a growing concern worldwide, alternative strategies to treat infections in farmed animals are necessary in order to decrease the risk to human health and increase animal health and productivity. Mucosal surfaces are the most common route used by pathogens to enter the body. The mucosal surface that lines the gastrointestinal tract is covered by a continuously secreted mucus layer that protects the epithelial surface. The mucus layer is the first barrier the pathogen must overcome for successful colonization, and is mainly composed of densely glycosylated proteins called mucins. The vast array of carbohydrate structures present on the mucins provide an important setting for host-pathogen interactions. This review summarizes the current knowledge on gastrointestinal mucins and their role during infections in farmed animals. We examine the interactions between mucins and animal pathogens, with a focus on how pathogenic bacteria can modify the mucin environment in the gut, and how this in turn affects pathogen adhesion and growth. Finally, we discuss analytical challenges and complexities of the mucus-based defense, as well as its potential to control infections in farmed animals.

1. Introduction

1.1. Animal Production and Infection Challenges

Gastrointestinal (GI) pathogens affect livestock, poultry, and aquaculture production systems worldwide, decreasing overall production parameters and causing considerable economic losses. Besides their effect in increased morbidity, mortality, and reduced feed conversion rates in food animals, GI pathogens can also decrease the safety of food products due to their zoonotic potential.

Intensive food animal production systems require the implementation of biosecurity, prophylactic, and therapeutic measures to prevent and control disease. High animal density in farms in conjunction with stress factors facilitate pathogen transmission. In aquaculture, for example, stocking density and stress are important factors that can increase susceptibility to infections such as those caused by Aeromonas salmonicida and Vibrio spp. [1,2]. Important bacterial pathogens in poultry include Salmonella spp., Escherichia coli, and Clostridium spp. [3,4,5]. Clostridium perfringens is a causative agent of necrotic enteritis (NE), a highly prevalent disease in poultry [6]. Additionally, parasitic coccidial infections by Eimeria spp. are common and affect the poultry industry [7]. Enteric infections due to E. coli, C. perfringens, Brachyspira hyodysenteriae, and Lawsonia intracellularis can reduce productivity and welfare in pigs [8]. Helicobacter suis infection in the pig stomach is associated with gastritis and decreased daily weight gain [9]. The GI parasites Ostertagia ostertagi and Cooperia oncophora have a significant impact in cattle production, mainly affecting the abomasum and intestines, respectively [10,11].

Antimicrobials are extensively used in intensive livestock, poultry, and aquaculture production for diverse purposes including therapeutic (i.e, treatment of diseases) and/or non-therapeutic purposes (i.e., disease control, prophylaxis and growth promotion) [12,13]. However, the use of antimicrobials in food animals has come under scrutiny due its association with the emergence of antimicrobial resistant strains, particularly those that can infect humans [14,15]. Studies have linked the use of antimicrobials for growth promotion, where they are continuously administered in sub-therapeutic doses via feed or water, and antimicrobial resistant bacteria of animal origin [16,17,18]. Pathogens becoming resistant to antimicrobials are a public health issue, and the concerns raised regarding the use of antimicrobials for non-therapeutic purposes in food animals has led to their complete ban in the European Union [19]. As a result, the incidence of several animal diseases such as NE in poultry and diarrhea due to E. coli, L. intracellularis, and Brachyspira spp. in pigs, which were previously under control by in-feed antimicrobials, has increased [20,21,22,23]. Resistance to antimicrobials used in the control and treatment of nematodes has been reported in livestock [24,25]. Consequently, alternative strategies to prevent and treat infections in farmed animals are needed to decrease the risk to human health and increase animal health and productivity.

1.2. Mucus, Mucins, and Mucin Glycosylation

GI mucus and mucins from primates and rodents have been previously reviewed elsewhere [26,27], therefore, we only present a brief summary as a general introduction to mucus in farmed animals.

1.2.1. Mucus

The gastrointestinal mucosa continuously secretes mucus, which consists of 95% water and the remaining 5% is mainly composed of gel-forming highly glycosylated mucin glycoproteins, responsible for the viscous properties of mucus [28,29]. Mucus also contains salts, lipids, and proteins with protective functions, such as lysozyme, immunoglobulins, defensins, growth factors, and trefoil factors [30,31,32,33]. Mucins provide a matrix keeping other secreted defensive molecules in a strategic location [26,34]. Together these components create a slimy organized layer, which is the first barrier pathogens encounter [26,34]. Mucins form large netlike structures, which expand and replenish the mucus from underneath [26,34]. The thickness of the mucus layer is approximately 250 μm in the rat stomach, 150–400 μm in the rat small intestine, and reaching 800–900 μm in the rat colon [35]. Dietary fiber and microbiota have been demonstrated to be important for the generation of a mucus layer during normal conditions in mice [36,37].

1.2.2. Mucins

Mucins are highly glycosylated glycoproteins that generally are large, with the secreted multimeric mucins having a molecular weight in the order of 40 MDa [38]. Mucins consist of a protein core with regions rich in serine and threonine, where O-linked carbohydrate chains are added in the Golgi apparatus during biosynthesis [39]. The two major types of mucins are transmembrane (cell-surface) and secretory mucins. Transmembrane mucins are located on the apical surface of mucosal epithelial cells, and they participate in mucosal defense translating external stimuli to cellular responses [40,41]. In the human gastrointestinal tract, cell-surface mucins include mucin-1 (MUC1), MUC3, MUC4, MUC12, MUC13, MUC15, MUC16, and MUC17 [42].

Secretory mucins can be gel-forming (MUC2, MUC5AC, MUC5B, MUC6, and MUC19) or non-gel-forming (MUC7). In the healthy human stomach, MUC5AC and MUC6 are the major gel-forming mucins located in the surface and glandular region, respectively [43], whereas in the intestine the main secreted mucin is MUC2 [44]. In the GI tract, secretory mucins are produced and secreted by mucous cells of glandular tissues and specialized epithelial cells called goblet cells [45].

1.2.3. Mucin Glycosylation

Post-translational modifications of proteins by glycosylation can occur in N-linked and O-linked form, where O-glycosylation is the main modification of mucins. Mucin-type O-glycosylation takes place in the Golgi complex and is initiated by the addition of α-N-acetylgalactosamine (GalNAc) to the hydroxyl group of Ser/Thr side chains [46,47]. The complex oligosaccharides on proteins have three regions: core region (core 1–core 8), backbone region (type 1 and type 2), and peripheral region. This latter region can be terminated by fucose, galactose, GalNAc, or sialic acid residues, forming histo-blood-group antigens such as A, B, H, Lewis a (Le^a), Lewis b (Le^b), Lewis x (Le^x), Lewis y (Le^y), as well as sialyl-Le^a and sialyl-Le^x structures [48]. The glycans can be further modified by sulfation. The structure of carbohydrates depends on glycosyltransferases expressed by the cell: this is in turn governed by the inherited genetics (i.e., blood group antigens) as well as by tissue location [49]. The mucin oligosaccharide terminal structures vary between species, individuals, and tissue locations within each individual [50,51,52,53,54]. Mucin O-glycans contribute up to 80% of the molecular weight of mucins [28]. The high level of glycosylation enables mucins to function as a protective barrier by lubricating the epithelium and preventing degradation of the protein backbone by proteases [55,56]. Moreover, mucin glycans can modulate cell adhesion, serve as ligands for cell surface receptors, take part in host-pathogen interactions and serve as energy sources for both commensals and pathogens [57,58,59,60,61,62].

1.3. Host-Pathogen Interactions in the Mucus Niche

Microbes commonly interact with the glycan structures on the host’s glycocalyx to colonize mucosal surfaces [63]. The high variety of mucin oligosaccharides forms an extensive repertoire of attachment sites for microorganisms [54,64,65,66], which attach to mucosal glycans via adhesins with different carbohydrate specificities [67,68,69]. Both membrane bound and secreted mucins can play protective roles during infection. Mice lacking the membrane bound mucin Muc1 are more susceptible to Helicobacter pylori and Campylobacter jejuni infections [70,71]. MUC1 expression is upregulated in response to infection, and it also acts as a releasable decoy limiting adhesion of the human gastric pathogen H. pylori to the cell surface [40]. In humans, polymorphisms in MUC1 can contribute to the development of gastritis, gastric cancer, and Crohn’s disease [72,73].

The protective role of secreted intestinal mucins has also been demonstrated in mice lacking Muc2 with increased susceptibility to Salmonella enterica and Citrobacter rodentium infections [74,75], while Muc5C deficiency hinders Trichuris muris clearance [76]. Furthermore, H. pylori-infected rhesus monkeys and children secreting mucins with low H. pylori binding capacity develop higher H. pylori density infections and gastritis than individuals with mucins with high H. pylori binding capacity [53,65], suggesting that binding of pathogens to secreted mucins protects the epithelium. Potentially, binding to mucins hinders the pathogen from close contact with the epithelial cells, and aids in pathogen dissemination with gastric emptying. Adhesin dependent binding to mucin glycans can also cause aggregations that inhibit bacterial proliferation, and affect the expression of pathogen virulence genes [57,77].

Bacteria can also utilize mucus to their advantage: H. pylori is mainly located deep in the mucus layer, which protects the bacterium from gastric acid [78]. Mucus provides nutrients for bacterial growth [79,80]. A number of bacterial strains are able to degrade mucins by producing specific bacterial enzymes, including glycosidases, and use the released glycans as an energy source [81,82,83,84,85,86]. Studies comparing germ free and monoassociated mice have revealed that Bacteroides thetaiotaomicron induces the expression of fucose on cell surface glycoconjugates by secreting signaling molecules, and these fucosylated glycans can be utilized by these bacteria as a carbon source [87].

Host mucin glycans can regulate the proliferation and gene expression of pathogens [77,88,89]. In vitro proliferation assays revealed that culturing H. pylori in the presence of purified human gastric mucins can have a stimulating or an inhibitory effect on bacterial growth depending on the mucin type [57,77]. The expression of genes important for H. pylori pathogenicity can also be differentially regulated by mucins [77]. Similarly, C. jejuni exposure to MUC2 upregulates genes involved in pathogenicity and colonization [90].

Bacterial colonization can alter mucin production and glycosylation quantitatively and qualitatively [50]. For example, H. pylori infection can alter the expression of normal gastric mucins [91]. Furthermore, murine infection studies revealed that H. pylori impairs mucin production and turnover rate in the gastric mucosa [92]. H. pylori can express the blood-group binding adhesin BabA and/or the Sialic acid binding adhesin SabA, which bind to Le^b and sialylated Lewis a (SLe^a) and SLe^x antigens on gastric mucins and glycolipids [61,69]. Primate infection with this pathogen caused time-dependent suppression of fucosylated blood group antigens, decreasing Le^b expression, while the expression of SLe^a and SLe^x antigens increased in the gastric mucosa, affecting the ability of the bacterium to adhere to the host epithelium [65].

2. Pig

2.1. Pig Gastric Mucins

Pigs are monogastric animals with a gastric mucosa that can be divided into two main parts: a glandular part (containing cardiac gland zone, fundic gland zone, and antrum with pyloric glands) and a nonglandular part called pars esophagea that is covered by a stratified squamous epithelium [93]. The nonglandular region and the cardiac gland zone have a pH between 5 and 7 due to the presence of saliva and cardiac gland bicarbonate secretions [94], while the fundic glands provide a lower pH in the distal part of the stomach [95]. The glandular part of the stomach produces mucins, and the apoprotein content and length of the glycosylated domains differ between mucins isolated from the surface epithelium and the glands, which may represent the pig equivalents of human Muc5ac and Muc6 mucins, respectively [96]. Proteomic analysis of pig gastric mucins identified mainly pig Muc5ac, but also indicated that Muc6 and Muc5b may be part of the mucin repertoire in the healthy pig stomach [97]. Similar to the human tissue localization, pig Muc5ac also localized to the surface epithelium [97]. For an overview on pig gastric mucins and glycosylation refer to Table 1.

[ Table omitted. See PDF. ]

The porcine mucins show inter-individual differences regarding density and glycan profile [97]. Mass spectrometric analysis identified 109 O-glycan structures in purified gastric mucins from three healthy pigs, out of which only 14 were present in all examined samples [97]. Similar to human gastric mucins, the oligosaccharides in pig mucins are mainly extended core 1 and core 2 glycans, but extended core 3 and core 4 structures are also present [97,98]. Pig gastric mucin glycans are mainly terminated by galactose and have a low degree of sialylation. Unlike human gastric mucins from healthy individuals [64], pig gastric mucins are highly sulphated [97,99].

The high acidity of the pig stomach provides a challenging environment for bacteria, thus relatively low numbers of acid-tolerant bacteria colonize this part of the gut [110,111]. Some of these microbes have been described to have beneficial functions for the host [112,113]. For example, lactic acid bacteria, such as Lactobacillus spp., can prevent the colonization of the gut by pathogens and enhance mucosal immunity [114,115,116]. Another advantageous function of some gut bacteria is the contribution to the availability of nutrients that can be used by the host [117]. Sequencing analysis of bacterial DNA revealed the presence of Herbiconiux and Brevundimonas genera in the pig stomach [111], which take part in the degradation of plant-derived food [118,119]. Spiral-shaped bacteria frequently colonize the pig stomach [120,121,122], and their presence has been associated with the development of chronic gastritis and decreased daily weight gain in pigs [9,123,124]. The main Helicobacter species in the pig stomach is H. suis [93], colonizing 60–95% of pigs at slaughter [125]. Fluorescence in situ hybridization performed on gastric tissue sections from H. suis infected pigs demonstrated that it colonizes the mucus layer lining the surface epithelium and the gastric pits of the pig stomach [97]. H. suis can be often found in close contact with parietal cells affecting acid production [97,126,127,128]. H. suis adhesion to glycans present in the pig stomach occurs via two modes: to Galβ3GlcNAcβ3Galβ4Glcβ1 at both neutral and acidic pH, and to negatively charged structures at acidic pH [97]. Furthermore, binding occurs to both mucins and glycolipids (Table 2) [97]. Binding to glycolipids confers close adherence to host epithelial cells, whereas mucins can act as decoys and hinder access to the epithelial cell surface, thus the balance between adhesion to mucins versus glycolipids is probably important for the colonization process and the outcome of the disease.

[ Table omitted. See PDF. ]

2.2. Pig Intestinal Mucins

The colonic mucus layer in healthy pigs, as in mice and humans, is organized in striations parallel to the mucosal surface and is mainly composed of Muc2 [44,101,141,142]. Glucosamine, galactosamine, galactose, fucose, and sialic acid are the main carbohydrates found on mucin glycoproteins in the pig colon [52,143]. Lectin staining of sections from small intestine suggests that the blood group AO genotype affects the glycomic profile of the mucosa [144]. A detailed characterization of the mucin glycosylation profile in the colon of healthy pigs identified high inter-individual variation of O-glycan structures, and equal distribution of N-acetylneuraminic acid (NeuAc) and N-glycolylneuraminic acid (NeuGc) containing structures [52]. Mucin-type cores 1, 2, 3, and 4 where detected with a predominance of core 4 glycans (Table 1) [52].

Pathogen-mucin interaction studies in the pig colon have shown that S. enterica, Brachyspira hampsonii, B. hyodysenteriae, and Trichuris suis increased the levels of Muc5ac (Table 2) [100,101,132,133]. The host’s response to the colonic pathogen B. hyodysenteriae, commonly associated with swine dysentery (SD), has been thoroughly studied from a mucin perspective. B. hyodysenteriae infection results in profound structural changes to the mucus layer, massively inducing mucus production with increased expression of Muc2 and de novo synthesis of Muc5ac [100,101]. Further studies determined that neutrophil elastase and interleukin (IL) 17 expressed in the pig colon during infection synergistically with B. hyodysenteriae induced mucin production via mitogen-activated protein kinase 3 [145]. Glycosylation changes in intestinal mucins can occur as an adaptation to new dietary constituents (i.e., during weaning), physical environment, as well as commensal and pathogenic bacteria [146,147,148,149,150,151,152]. GI bacterial pathogens can induce mucosal glycosylation changes, which in turn can affect bacterial growth and adhesion [57,65,77]. Challenges with the intestinal nematode Trichinella spiralis increased the amount of mucin in goblet cells, and caused changes in sulfation and sialylation in pigs, as determined by histochemical staining [134]. B. hyodysenteriae can also regulate mucin glycosylation synthesis in the pig colon [52]. Structural analysis of the mucins induced during B. hyodysenteriae infection compared to healthy controls showed loss of interindividual variation, shorter glycan chains, and higher abundance of neutral, core 2 and NeuGc containing structures [52]. Additionally, lower abundance of fucosylated and sulfated O-glycans was observed [52]. The increased abundance of shorter mucin O-glycans suggests a faster mucin turnover due to the increased mucin secretion during infection [52]. Furthermore, the mRNA levels of C1GalT, C3GnT, and C2/4GnT-M glycosyltranferases were upregulated in the colon tissue of B. hyodysenteriae infected pigs, linking differences in core glycan abundance to enzyme levels.

Motility and chemotaxis are important virulence factors that can facilitate intestinal colonization. B. hyodysenteriae has chemotactic behavior towards mucin, fucose, and serine [153,154,155]. Studies on E. coli and H. pylori indicate that adhesion to mucins favors the host by limiting colonization and access to the epithelial surface [70,156,157,158]. E. coli uses fimbrial adhesion factors to bind to the mucus layer and colonize the small intestine [129,130,131]. Meijerink et al. showed that a DNA polymorphism influences an alpha(1,2)fucosyltransferase activity related to blood group variants, affecting the mucin phenotype and thus the adhesion of enterotoxigenic E. coli F18 [159].

E. coli K99 fimbriae has affinity for sialic acid residues on mucin glycopeptides and glycolipids found in the pig small intestine [129,130,131]. B. hyodysenteriae also binds to mucin carbohydrates in the mucus layer, with increased adhesion ability to mucins from infected pigs [101]. However, it is yet to be determined the specific glycan structures that B. hyodysenteriae interacts with and whether the binding to colonic mucins is beneficial for the host or the pathogen. Studies have shown that pig intestinal bacteria can use mucin glycoproteins or its components as a carbon source [160,161,162]. Clostridium and Bacteroides isolated from pig colon mucosa are able to hydrolyze pig colonic and gastric mucin carbohydrates [160]. Similarly, several bacterial isolates belonging to the order Clostridiales produced butyrate and grew on mucin [161]. Recently, a novel commensal mucin degrading and butyrate producing bacterium, Cloacibacillus porcorum, was isolated from the pig intestinal mucosa [162].

3. Chicken

Avian species are monogastric animals with a glandular stomach termed proventriculus that secretes hydrochloric acid (HCl) and pepsinogen. Unlike non-avian animals, chickens have in addition a gizzard to breakdown food, a crop for food storage, two ceca pouches, and a cloaca that serves as a common cavity and opening for the GI, urinary and reproductive tracts [163].

Lang et al. [164] described that Muc4, Muc13, and Muc16 transmembrane mucins, and Muc6, Muc2, Muc5ac, and Muc5b gel forming mucins are encoded in the chicken genome and are homologous to the human counterparts. Forder et al. [102] found detectable levels of Muc2 and Muc13, but not Muc1, Muc4, Muc5b, and Muc16 mRNA in the jejunum of healthy chickens (Table 1). The Muc2 expression increases from anterior to posterior through the GI tract, and the Muc2 proline, threonine, and serine (PTS) rich domain diverges from the human motif in amino acid composition and chain length [165].

Struwe et al. [103] characterized the mucin glycosylation profile in healthy chicken caecum, small intestine, and large intestine, detecting predominance of core 3 and core 4 O-glycans in all sections, and an increasing glycan diversity from the caecum towards the small and large intestine. Similarities in the distribution of neutral, sialylated, and sulfated O-glycans were observed in the small and large intestine (Table 1) [103]. The large intestine is the only site with NeuAc/NeuGc containing structures, whereas fucosylated mucin carbohydrates are found throughout the chicken small and large intestine as blood group H, B, and Lewis^a/x like structures [103].

The host’s defense against infection can result in differences in mucin expression, as is the case in necrotic enteritis infection in the small intestinal mucosa. Broiler chickens co-challenged with the coccidian parasite Eimeria spp. and C. perfringens resulted in a 54% increase in Muc5ac mRNA levels compared to unchallenged controls, while Muc2 and Muc13 expression levels decreased (Table 1) [102]. Furthermore, inoculation with Eimeria spp. increases mucus secretion, which may be the reason for increased growth and colonization of the mucolytic C. perfringens in the intestine of Eimeria spp. infected chicks [166]. Eimeria tenella binds to mucins isolated from the duodenum, which inhibits epithelial cell invasion in vitro [167]. Several studies related to bacterial interactions with mucins in chickens refer to C. jejuni colonization. Although C. jejuni is not pathogenic in chickens, it is an important enteric pathogen in humans, and chickens are the main source of infection. In chickens, C. jejuni is localized in the mucus layer and does not colonize epithelial cells [168]. The differential mucus composition in the GI tract of chicken and humans has been suggested to play a role in determining C. jejuni pathogenicity [169,170]. In vitro studies have shown that chicken intestinal mucins, predominantly those from large intestine, decreased C. jejuni binding to human HCT-8 cells [169]. The aforementioned study implied a role for mucin glycosylation in bacterial inhibition, a premise further explored by Struwe et al. [103] who determined that the increased glycan diversity observed in the large intestine corresponded with the high inhibitory effect of mucins from this region on C. jejuni and lower bacterial density. Studies have shown that C. jejuni is chemo attracted to pig gastric and bovine gallbladder mucin, L-aspartate, L-cysteine, L-glutamate, and L-serine amino acids, and L-fucose structures [171]. C. jejuni can recognize glycan structures such as terminal mannose, NeuAc, galactose and fucose, an interaction that is affected by temperature and oxygen [172]. Furthermore, L-fucose upregulates the C. jejuni cj0480c-cj0490 genomic island which allows the bacterium to use fucose as a carbon source for growth [173].

4. Cow

Unlike pigs and chickens, which are monogastric, bovines are polygastric animals with a four-compartment stomach composed of rumen, reticulum, omasum, and abomasum, followed by the small and large intestines. The abomasal compartment is functionally similar to the stomach of monogastric animals including humans, as it secretes HCl and pepsinogen. In bovines, genome wide analysis has allowed the identification of several secreted (Muc2, Muc5ac, Muc5b, Muc6, Muc7, and Muc19) and membrane bound mucins (Muc1, Muc3a, Muc4, Muc12, Muc13, Muc15, Muc16, Muc20, and Muc21), the majority of which are transcribed in the GI tract of adult healthy cows (Table 1) [104]. Gel-forming mucins are expressed from the abomasum onwards and membrane-associated mucins have a more general distribution throughout the GI tract [104]. In the abomasum are the main secreted mucin genes expressed are Muc5ac and Muc6, while Muc2 and Muc5b are predominantly expressed in the intestine [104]. Calf small intestinal mucin isolated by density gradient centrifugation has a glycan composition rich in galactose, followed by N-acetylglucosamine (GlcNAc), GalNAc, fucose, mannose, and NeuAc [105].

Few studies have characterized the effect of GI infections on mucus composition in cattle, and mostly focus has been given in response to parasitic infections. Nematode infections in the GI tract of cows can affect mucus composition and mucin glycan biosynthesis. This is the case for C. oncophora and O. ostertagi infections in the small intestine and abomasum respectively (Table 2) [135,136]. Both parasites induce a Th2 like immune response with upregulation of IL-4 and eosinophilia [174]. Infection with the nematode C. oncophora results in increased mRNA and protein levels of Muc2 in the small intestine [136]. O. ostertagi infection results in thickening of the abomasal mucosa and hyperplasia of mucus secreting cells [135], accompanied by increased transcription levels of Muc1, Muc6, Muc20, and fucosyltrasferases in the abomasum [135]. Despite of Muc5ac being commonly found in the abomasum, its expression remained unchanged during infection [135]. Soon after infection of the bovine GI tract, both C. oncophora and O. ostertagi induced expression of the mucin glycan biosynthesis enzyme gene Gcnt3, in addition to Gcnt4 and A4gnt induced by O. ostertagi [135,136], further supporting that changes in the mucus niche occur during infection.

Mucins can play a protective role limiting pathogen colonization [158]. Purified bovine Muc1 inhibited binding of E. coli, S. enterica, Staphylococcus aureus, and Bacillus subtilis to a human intestinal cell line in vitro in a dose dependent manner [157]. Sialic acid, l-fucose, and d-mannose were among the Muc1 carbohydrates involved in both E. coli and S. enterica binding inhibition [157]. Enterotoxigenic E. coli (ETEC) is a common cause of diarrhea in calves [175]. ETEC adheres to calve intestinal mucus through the interaction of bacterial pili and mucus glycoproteins in a pH dependent manner (Table 2) [137]. Different fimbriae recognized different receptors, K99 pili adhered to sialic acid and galactose, while F41 pili bound to desialylated glycans [137]. Unlike ETEC, enterohemorrhagic E. coli (EHEC) O157:H7 is not pathogenic in cattle. However, the GI of cattle is an important source of this bacterium that can cause disease in humans. Aperce et al. showed that E. coli O157:H7 grows in the presence of bovine crude intestinal mucus [176] and E. coli O157:H7 can use mucin carbohydrates found in the cow small intestinal mucus layer as a carbon source for growth [177]. E. coli O157:H7 utilizes fucose, galactose, GalNAc, GlcNAc, mannose, and NeuAc as the sole carbon sources when grown on minimal medium, with GlcNAc or sialic acid having the highest growth yield compared to the glucose control [177]. Furthermore, in vitro bacterial competitive assays demonstrated that bovine mucus-derived mannose, GlcNAc, NeuAc, and galactose, give EHEC a growth advantage due to a higher expression of genes involved in carbohydrate catabolism and faster utilization of glycans than commensal bacteria [177]. This phenomenon has also been observed in other organisms, such as the utilization of L-fucose by C. jejuni carrying the putative fucose permease (fucP) gene which is upregulated in the presence of fucose [178]. Similarly, induction of nanH, dctp, nanA, and nanK gene expression in Vibrio cholerae allows it to utilize sialic acid as a carbon source for growth [179].

5. Fish

The fish gastrointestinal tract anatomy differs between species; some have a stomach whereas others do not. Some fish have an intestine with small outpocketings in the anterior region called pyloric caeca to increase the surface area of the digestive epithelium, whereas other species may have a spiral intestine for the same purpose. The Atlantic salmon, Salmo salar, a prominent species in aquaculture, has a stomach and pyloric caeca in the anterior part of the intestine [180].

Five orthologues of the human and murine MUC1–MUC20 mucins have been identified in the genome of pufferfish, Fugu rubripes [181], and 13 in the zebrafish, Danio rerio [182]. Among cultured species, Sveen et al. [107] recently identified seven putative mucin genes of the Muc2 and Muc5 families in Atlantic salmon using annotation, transcription, and domain structure approaches. Pérez-Sánchez et al. [106] identified I-Muc, Muc13, Muc18, Muc2, Muc2-like and Muc19 in gilthead sea bream intestine (Table 1). Generally, studies involved in teleost mucin gene analyses rely on homologies with mammalian or zebrafish mucin genes (i.e., a study predating the aforementioned publication by Sveen et al. reported partial mucin sequences with homology to human MUC2 and MUC5 mucins in salmon skin [183]).

There is limited knowledge on fish GI mucin glycosylation (Table 1), except for Atlantic salmon where a total of 91 structures were identified on mucins isolated from pyloric caeca, proximal intestine, and distal intestine using mass spectrometry [51]. The structures present differed between regions: pyloric caecal mucins carried 56 structures, proximal intestine 65 structures, and distal intestine 75 structures [51]. However, within each tissue the glycan profile was remarkably similar; salmon mucin glycans showed much lower inter-individual variation in structure repertoire [51], compared with for example human or pig mucin glycans [52,64]. The major structures found were core 5 and extended core 5, with one NeuAc linked to the reducing end GalNAc in a α2-6 linkage, and most glycans were sialylated (NeuAc, 83–92%), Table 1 [51]. Core 1 and 2 glycans were detected in low abundance [51]. In addition, mucin glycosylation has been studied indirectly in carp by histochemistry and lectin staining of goblet cells [108,109]. β-GlcNAc, α1-6 fucose, α-GalNAc, β-Gal, and sialic acid containing structures were found in common nase, Chondrostoma nasus, with higher level of sialyation in the posterior part than in the anterior [109]. In chub, Squalius cephalus, the majority of neutral glycans in anterior intestine were fucosylated while β-Gal and β-GalNAc were found in the posterior intestine [109]. Similar to common nase, the highest level of sialylation was found in the posterior intestine [109]. Terminal α-fucose and β-GlcNAc were prominent in the anterior intestine of the Grass carp, Ctenopharyngodon idella, while β-Gal and α-GalNAc were more dominant in the posterior intestine, with sialylated and sulfated glycans present in both regions [109]. In the common carp, Cyprinus carpio, the major terminal glycan was β-GalNAc; β-GlcNAc was also present, and among acidic structures sulfated glycans were predominant [108,109]. The aforementioned studies show that terminal GalNAc and GlcNAc are often found in fish mucins, and these can be found linked to sialic acids.

The microbiota, infection, mucin expression, and glycosylation appear linked also in fish. Several studies demonstrate mucus related changes during infection (Table2): i.e., the number of mucous cells in the striped trumpeter, Latris lineata, increased near the attachment sites of the parasite Chondracanthus goldsmidi [184]. Edwardsiella ictaluri infection in channel catfish, Ictalurus punctatus, altered the expression of genes with similarities to the zebrafish Muc2 and Mucb mucins in the gut [139], and infection with the intestinal parasite Enteromyxum leei decreased the mRNA expression of I-Muc, Muc13, Muc2 and Muc2 like mucin genes in gilthead sea bream [106]. An altered mucin glycosylation, such as the appearance of fucose on carp gut mucins after endotoxin treatment, has been suggested to be a microbial clearance mechanism by enhancing bacterial adherence [185], and glycosylation changes that lead to a decrease in sialic acids may give access to cryptic glycan structures that bacteria can utilize [58]. Thirty percent of the studied gut microbiota in turbot, Scophtalmus maximus, inhibited the growth of Vibrio anguillarum, A. salmonicida, and Aeromonas hydrophila [186], therefore mucus related changes that affect the microbiota composition has the potential to indirectly affect pathogen density in the gut.

An increased mucus secretion can aid in the removal of pathogens from the epithelial surface [142]. In zebrafish, LPS has been shown to induce mucus secretion using in vivo imaging [187], changes in the intestinal mucin content of common carp occur in response to LPS [185], and V. cholerae infection in zebrafish increases the quantity of mucus expelled into the surrounding water [188]. The adhesion capacity and interactions of bacteria in mucus varies with species and host specificity, Table 2. Vibrio alginolyticus adheres to crude gilthead sea bream, Sparus aurata, skin, gill and intestinal mucus, and is able to use mucus as an energy source [140]. Pathogenic vibrios, namely V. alginolyticus, V. anguillarum, V. harveyi, and V. tubiashi, are also chemotactic towards skin, gill, and intestinal mucins of gilthead sea bream [189]. Galactose containing epitopes could be of importance for E. ictaluri adherence to channel catfish olfactory mucosa, as binding decreased after preincubation with galactose [190]. A. salmonicida binds to NeuAc on mucin O-glycans, the elimination of which reduces the binding ability of this pathogen to Atlantic salmon mucins [138]. A. salmonicida showed higher binding to gut mucins compared to skin mucins, with gut mucins having a higher level of sialylation (mono-, di-, and trisialylated O-glycan structures), longer glycan chains, and higher glycan diversity than skin mucins [138]. Thus, it is likely that binding occurs not only to NeuAc, but also rather to glycan structures including NeuAc. Despite being a target for adhesion, A. salmonicida is not able to hydrolyze NeuAc from mucins [58]. Atlantic salmon intestinal mucins enhance A. salmonicida growth and enzymatic removal of NeuAc increases growth further by exposing GlcNAc [58]. Calcium ion levels and pH also play a role in the adhesion and growth mechanism of A. salmonicida in contact with salmon mucins, underlining the importance of environmental factors in fish host-pathogen interactions [58].

6. Methodological Considerations for Mucus/Mucin Studies

The interest in mucus and mucins has expanded notably during recent years, and for researchers moving into this field, it is important to be aware of factors that result in commonly used tools to investigate changes after challenge being less suitable for mucin studies compared to other molecules. Firstly, the secreted mucins are very large and do not typically enter standard gels used for sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE), instead, what is found on such gels are either small (i.e., membrane bound) mucins or fragments of the large mucins, indicating the need for special gels. Secondly, since mucins are to 85% carbohydrate and subject to major post-translational regulation, mucin mRNA levels do not always reflect the amount of mucin/mucus being produced [142]. Histology on its own, without for example metabolic labelling can also be deceptive because important mucus functions are mainly carried out after secretion. Mucin production and secretion rate are important parameters in this context, secreted mucus is lost during processing using most histology methods and mainly the amount of mucus inside the goblet cells is evaluated. A decreased secretion could lead to an increased amount of mucus in the cells, and conversely, an increased secretion could lead to a decreased amount of mucus in the cells, none of which reflects the amount of mucus produced. Furthermore, these methods do not take into account the complexity of mucus, mucins, and mucin glycosylation. Therefore, it is necessary to combine several methods, such as glycan analysis, metabolic labeling of mucins, and histological evaluation to get a more complete understanding of mucus related events.

7. Complexities and Potential

Several studies suggest that mucus and mucins are important for regulating growth, composition, and behavior of the microflora (including pathogens), and that these in turn regulate mucus and mucins. For example, the commensal flora can improve the mucus barrier and pathogens can affect mucus via the inflammatory response, thus these factors are intertwined. However, the mucus-pathogen interactions are complex and depend on microbe and host species; therefore, in order to achieve such advancements, more knowledge is needed on this topic. One can envision many aspects where these systems can be utilized to prevent/treat disease, such as via feeds that enhance the mucus barrier and shape a beneficial microflora, potentially decreasing the amount and severity of infections, or via feeding a beneficial microflora directly. Furthermore, breeds may differ in mucus related parameters such as mucin glycosylation, and selective breeding could take mucin mutations and glycosylation into account once an “ideal” mucin repertoire has been identified. There is also potential for temporary “mucus boosting” treatment, for example with the purpose of “flushing out” a pathogen or to prophylactically enhance the barrier to prevent infections from spreading within a site.

Acknowledgments

This work was supported by the Swedish research council Formas (221-2013-590), the Swedish Research Council (2016-05154_3), and the Engkvists and WM Lundgren foundations. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Conflicts of Interest

The authors declare no conflicts of interest.

References

1. Kent, M.L. Marine netpen farming leads to infections with some unusual parasites. Int. J. Parasitol. 2000, 30, 321–326.

2. Lafferty, K.D.; Harvell, C.D.; Conrad, J.M.; Friedman, C.S.; Kent, M.L.; Kuris, A.M.; Powell, E.N.; Rondeau, D.; Saksida, S.M. Infectious diseases affect marine fisheries and aquaculture economics. Ann. Rev. Mar. Sci. 2015, 7, 471–496.

3. Gast, R.K.; Shivaprasad, H.L.; Barrow, P.A. Salmonella infections. In Diseases of Poultry, 13th ed.; Swayne, D.E., Ed. John Wiley & Sons, Ltd.: Hoboken, NJ, USA, 2013; pp. 675–736.

4. Nolan, L.K.; Barnes, H.J.; Vaillancourt, J.-P.; Abdul-Aziz, T.; Logue, C.M. Colibacillosis. In Diseases of Poultry, 13th ed.; Swayne, D.E., Ed.; John Wiley & Sons, Ltd.: Hoboken, NJ, USA, 2013; pp. 751–805.

5. Logue, C.M.; Songer, J.G.; Uzal, F.; Opengart, K. Clostridial diseases. In Diseases of Poultry, 13th ed.; Swayne, D.E., Ed.; John Wiley & Sons, Ltd.: Hoboken, NJ, USA, 2013; pp. 943–970.

6. Van Immerseel, F.; Rood, J.I.; Moore, R.J.; Titball, R.W. Rethinking our understanding of the pathogenesis of necrotic enteritis in chickens. Trends Microbiol. 2009, 17, 32–36.

7. Haug, A.; Gjevre, A.G.; Skjerve, E.; Kaldhusdal, M. A survey of the economic impact of subclinical eimeria infections in broiler chickens in norway. Avian Pathol. J. WVPA 2008, 37, 333–341.

8. Economou, V.; Gousia, P. Agriculture and food animals as a source of antimicrobial-resistant bacteria. Infect. Drug Resist. 2015, 8, 49–61.

9. De Bruyne, E.; Flahou, B.; Chiers, K.; Meyns, T.; Kumar, S.; Vermoote, M.; Pasmans, F.; Millet, S.; Dewulf, J.; Haesebrouck, F.; et al. An experimental Helicobacter suis infection causes gastritis and reduced daily weight gain in pigs. Vet. Microbiol. 2012, 160, 449–454.

10. Nilsson, O.; Sorelius, L. trichostrongyle infections of cattle in sweden. Nord. Vet. Med. 1973, 25, 65–78.

11. Fox, M.T. Pathophysiology of infection with gastrointestinal nematodes in domestic ruminants: Recent developments. Vet. Parasitol. 1997, 72, 285–297.

12. Cabello, F.C. Heavy use of prophylactic antibiotics in aquaculture: A growing problem for human and animal health and for the environment. Environ. Microbiol. 2006, 8, 1137–1144.

13. Marshall, B.M.; Levy, S.B. Food animals and antimicrobials: Impacts on human health. Clin. Microbiol. Rev. 2011, 24, 718–733.

14. Levy, S.B.; Marshall, B. Antibacterial resistance worldwide: Causes, challenges and responses. Nat. Med. 2004, 10, S122–S129.

15. European Food Safety Authority; European Centre for Disease Prevention and Control. The european union summary report on antimicrobial resistance in zoonotic and indicator bacteria from humans, animals and food in 2015. EFSA J. 2017, 15, 4694.

16. Alexander, T.W.; Yanke, L.J.; Topp, E.; Olson, M.E.; Read, R.R.; Morck, D.W.; McAllister, T.A. Effect of subtherapeutic administration of antibiotics on the prevalence of antibiotic-resistant Escherichia coli bacteria in feedlot cattle. Appl. Environ. Microbiol. 2008, 74, 4405–4416.

17. Diarra, M.S.; Silversides, F.G.; Diarrassouba, F.; Pritchard, J.; Masson, L.; Brousseau, R.; Bonnet, C.; Delaquis, P.; Bach, S.; Skura, B.J.; et al. Impact of feed supplementation with antimicrobial agents on growth performance of broiler chickens, Clostridium perfringens and enterococcus counts, and antibiotic resistance phenotypes and distribution of antimicrobial resistance determinants in Escherichia coli isolates. Appl. Environ. Microbiol. 2007, 73, 6566–6576.

18. Varga, C.; Rajic, A.; McFall, M.E.; Reid-Smith, R.J.; McEwen, S.A. Associations among antimicrobial use and antimicrobial resistance of Salmonella spp. Isolates from 60 alberta finishing swine farms. Foodborne Pathog. Dis. 2009, 6, 23–31.

19. EPC. Ban on Antibiotics as Growth Promoters in Animal Feed Enters into Effect. Available online: http://europa.eu/rapid/press-release_IP-05-1687_en.htm (accessed on 9 May 2018).

20. Grave, K.; Kaldhusdal, M.C.; Kruse, H.; Harr, L.M.; Flatlandsmo, K. What has happened in norway after the ban of avoparcin? Consumption of antimicrobials by poultry. Prev. Vet. Med. 2004, 62, 59–72.

21. Alvarez-Ordonez, A.; Martinez-Lobo, F.J.; Arguello, H.; Carvajal, A.; Rubio, P. Swine dysentery: Aetiology, pathogenicity, determinants of transmission and the fight against the disease. Int. J. Environ. Res. Public Health 2013, 10, 1927–1947.

22. Casewell, M.; Friis, C.; Marco, E.; McMullin, P.; Phillips, I. The european ban on growth-promoting antibiotics and emerging consequences for human and animal health. J. Antimicrob. Chemother. 2003, 52, 159–161.

23. Dibner, J.J.; Richards, J.D. Antibiotic growth promoters in agriculture: History and mode of action. Poult. Sci. 2005, 84, 634–643.

24. Gasbarre, L.C.; Smith, L.L.; Lichtenfels, J.R.; Pilitt, P.A. The identification of cattle nematode parasites resistant to multiple classes of anthelmintics in a commercial cattle population in the us. Vet. Parasitol. 2009, 166, 281–285.

25. Sutherland, I.A.; Leathwick, D.M. Anthelmintic resistance in nematode parasites of cattle: A global issue? Trends Parasitol. 2011, 27, 176–181.

26. Linden, S.K.; Sutton, P.; Karlsson, N.G.; Korolik, V.; McGuckin, M.A. Mucins in the mucosal barrier to infection. Mucosal Immunol. 2008, 1, 183–197.

27. McGuckin, M.A.; Linden, S.K.; Sutton, P.; Florin, T.H. Mucin dynamics and enteric pathogens. Nat. Rev. Microbiol. 2011, 9, 265–278.

28. Bansil, R.; Turner, B.S. Mucin structure, aggregation, physiological functions and biomedical applications. Curr. Opin. Colloid Interface Sci. 2006, 11, 164–170.

29. Sellers, L.A.; Allen, A.; Morris, E.R.; Ross-Murphy, S.B. Mucus glycoprotein gels. Role of glycoprotein polymeric structure and carbohydrate side-chains in gel-formation. Carbohydr. Res. 1988, 178, 93–110.

30. Schumacher, G.F.B.; Kim, M.H.; Hosseinian, A.H.; Dupon, C. Immunoglobulins, proteinase inhibitors, albumin, and lysozyme in human cervical mucus. Am. J. Obstet. Gynecol. 1977, 129, 629–636.

31. Jones, D.E.; Bevins, C.L. Paneth cells of the human small intestine express an antimicrobial peptide gene. J. Biol. Chem. 1992, 267, 23216–23225.

32. Mukherjee, S.; Zheng, H.; Derebe, M.G.; Callenberg, K.M.; Partch, C.L.; Rollins, D.; Propheter, D.C.; Rizo, J.; Grabe, M.; Jiang, Q.X.; et al. Antibacterial membrane attack by a pore-forming intestinal c-type lectin. Nature 2014, 505, 103–107.

33. Allen, A.; Bell, A.; Mantle, M.; Pearson, J.P. The structure and physiology of gastrointestinal mucus. In Mucus in Health and Disease—II; Chantler, E.N., Elder, J.B., Elstein, M., Eds.; Springer: Boston, MA, USA, 1982; pp. 115–133.

34. Hansson, G.C.; Johansson, M.E. The inner of the two MUC2 mucin-dependent mucus layers in colon is devoid of bacteria. Gut Microbes 2010, 1, 51–54.

35. Atuma, C.; Strugala, V.; Allen, A.; Holm, L. The adherent gastrointestinal mucus gel layer: Thickness and physical state in vivo. Am. J. Physiol. Gastrointest. Liver Physiol. 2001, 280, G922–G929.

36. Schroeder, B.O.; Birchenough, G.M.H.; Stahlman, M.; Arike, L.; Johansson, M.E.V.; Hansson, G.C.; Backhed, F. Bifidobacteria or fiber protects against diet-induced microbiota-mediated colonic mucus deterioration. Cell Host Microbe 2018, 23, 27–40.

37. Rodriguez-Pineiro, A.M.; Johansson, M.E. The colonic mucus protection depends on the microbiota. Gut Microbes 2015, 6, 326–330.

38. Sheehan, J.K.; Brazeau, C.; Kutay, S.; Pigeon, H.; Kirkham, S.; Howard, M.; Thornton, D.J. Physical characterization of the MUC5AC mucin: A highly oligomeric glycoprotein whether isolated from cell culture or in vivo from respiratory mucous secretions. Biochem. J. 2000, 347 Pt 1, 37–44.

39. Strous, G.J.; Dekker, J. Mucin-type glycoproteins. Crit. Rev. Biochem. Mol. Biol. 1992, 27, 57–92.

40. Linden, S.K.; Florin, T.H.J.; McGuckin, M.A. Mucin dynamics in intestinal bacterial infection. PLoS ONE 2008, 3.

41. Singh, P.K.; Hollingsworth, M.A. Cell surface-associated mucins in signal transduction. Trends Cell Biol. 2006, 16, 467–476.

42. Hattrup, C.L.; Gendler, S.J. Structure and function of the cell surface (tethered) mucins. Annu. Rev. Physiol. 2008, 70, 431–457.

43. Ho, S.B.; Roberton, A.M.; Shekels, L.L.; Lyftogt, C.T.; Niehans, G.A.; Toribara, N.W. Expression cloning of gastric mucin complementary DNA and localization of mucin gene expression. Gastroenterology 1995, 109, 735–747.

44. Herrmann, A.; Davies, J.R.; Lindell, G.; Martensson, S.; Packer, N.H.; Swallow, D.M.; Carlstedt, I. Studies on the “insoluble” glycoprotein complex from human colon. Identification of reduction-insensitive Muc2 oligomers and C-terminal cleavage. J. Biol. Chem. 1999, 274, 15828–15836.

45. Verdugo, P. Goblet cells secretion and mucogenesis. Annu. Rev. Physiol. 1990, 52, 157–176.

46. Hanisch, F.G. O-glycosylation of the mucin type. Biol. Chem. 2001, 382, 143–149.

47. Bennett, E.P.; Mandel, U.; Clausen, H.; Gerken, T.A.; Fritz, T.A.; Tabak, L.A. Control of mucin-type O-glycosylation: A classification of the polypeptide galnac-transferase gene family. Glycobiology 2012, 22, 736–756.

48. Green, C. The abo, lewis and related blood group antigens; a review of structure and biosynthesis. FEMS Microbiol. Immunol. 1989, 1, 321–330.

49. Rini, J.; Esko, J.; Varki, A. Glycosyltransferases and glycan-processing enzymes. In Essentials of Glycobiology, 2nd ed.; Varki, A., Cummings, R.D., Esko, J.D., Freeze, H.H., Stanley, P., Bertozzi, C.R., Hart, G.W., Etzler, M.E., Eds.; Cold Spring Harbor Laboratory: Cold Spring Harbor, NY, USA, 2009.

50. Freitas, M.; Axelsson, L.G.; Cayuela, C.; Midtvedt, T.; Trugnan, G. Microbial-host interactions specifically control the glycosylation pattern in intestinal mouse mucosa. Histochem. Cell Biol. 2002, 118, 149–161.

51. Jin, C.; Padra, J.T.; Sundell, K.; Sundh, H.; Karlsson, N.G.; Lindén, S.K. Atlantic salmon carries a range of novel O-glycan structures differentially localized on skin and intestinal mucins. J. Proteome Res. 2015, 14, 3239–3251.

52. Venkatakrishnan, V.; Quintana-Hayashi, M.P.; Mahu, M.; Haesebrouck, F.; Pasmans, F.; Linden, S.K. Brachyspira hyodysenteriae infection regulates mucin glycosylation synthesis inducing an increased expression of core-2 O-glycans in porcine colon. J. Proteome Res. 2017, 16, 1728–1742.

53. Linden, S.; Semino-Mora, C.; Liu, H.; Rick, J.; Dubois, A. Role of mucin lewis status in resistance to Helicobacter pylori infection in pediatric patients. Helicobacter 2010, 15, 251–258.

54. Robbe, C.; Capon, C.; Coddeville, B.; Michalski, J.C. Structural diversity and specific distribution of O-glycans in normal human mucins along the intestinal tract. Biochem. J. 2004, 384, 307–316.

55. Garner, B.; Merry, A.H.; Royle, L.; Harvey, D.J.; Rudd, P.M.; Thillet, J. Structural elucidation of the N- and O-glycans of human apolipoprotein(a): Role of O-glycans in conferring protease resistance. J. Biol. Chem. 2001, 276, 22200–22208.

56. Crouzier, T.; Boettcher, K.; Geonnotti, A.R.; Kavanaugh, N.L.; Hirsch, J.B.; Ribbeck, K.; Lieleg, O. Modulating mucin hydration and lubrication by deglycosylation and polyethylene glycol binding. Adv. Mater. Interfaces 2015, 2, 1500308.

57. Skoog, E.C.; Padra, M.; Aberg, A.; Gideonsson, P.; Obi, I.; Quintana-Hayashi, M.P.; Arnqvist, A.; Linden, S.K. Baba dependent binding of Helicobacter pylori to human gastric mucins cause aggregation that inhibits proliferation and is regulated via arss. Sci. Rep. 2017, 7, 40656.

58. Padra, J.T.; Sundh, H.; Sundell, K.; Venkatakrishnan, V.; Jin, C.; Samuelsson, T.; Karlsson, N.G.; Linden, S.K. Aeromonas salmonicida growth in response to atlantic salmon mucins differs between epithelial sites, is governed by sialylated and N-acetylhexosamine-containing O-glycans, and is affected by Ca2+. Infect. Immun. 2017, 85, e00189-17.

59. Fukuda, M. Roles of mucin-type O-glycans in cell adhesion. Biochim. Biophys. Acta 2002, 1573, 394–405.

60. Yeaman, C.; Le Gall, A.H.; Baldwin, A.N.; Monlauzeur, L.; Le Bivic, A.; Rodriguez-Boulan, E. The O-glycosylated stalk domain is required for apical sorting of neurotrophin receptors in polarized mdck cells. J. Cell Biol. 1997, 139, 929–940.

61. Linden, S.; Nordman, H.; Hedenbro, J.; Hurtig, M.; Boren, T.; Carlstedt, I. Strain- and blood group-dependent binding of Helicobacter pylori to human gastric MUC5AC glycoforms. Gastroenterology 2002, 123, 1923–1930.

62. Linden, S.K.; Bierne, H.; Sabet, C.; Png, C.W.; Florin, T.H.; McGuckin, M.A.; Cossart, P. Listeria monocytogenes internalins bind to the human intestinal mucin MUC2. Arch. Microbiol. 2008, 190, 101–104.

63. Hooper, L.V.; Gordon, J.I. Glycans as legislators of host-microbial interactions: Spanning the spectrum from symbiosis to pathogenicity. Glycobiology 2001, 11, 1R–10R.

64. Jin, C.; Kenny, D.T.; Skoog, E.C.; Padra, M.; Adamczyk, B.; Vitizeva, V.; Thorell, A.; Venkatakrishnan, V.; Linden, S.K.; Karlsson, N.G. Structural diversity of human gastric mucin glycans. Mol. Cell. Proteom. 2017, 16, 743–758.

65. Linden, S.; Mahdavi, J.; Semino-Mora, C.; Olsen, C.; Carlstedt, I.; Boren, T.; Dubois, A. Role of abo secretor status in mucosal innate immunity and h. Pylori infection. PLoS Pathog. 2008, 4, e2.

66. Linden, S.; Mahdavi, J.; Hedenbro, J.; Boren, T.; Carlstedt, I. Effects of pH on Helicobacter pylori binding to human gastric mucins: Identification of binding to non-MUC5AC mucins. Biochem. J. 2004, 384, 263–270.

67. Linden, S.K.; Wickstrom, C.; Lindell, G.; Gilshenan, K.; Carlstedt, I. Four modes of adhesion are used during Helicobacter pylori binding to human mucins in the oral and gastric niches. Helicobacter 2008, 13, 81–93.

68. Borén, T.; Falk, P.; Roth, K.A.; Larson, G.; Normark, S. Attachment of Helicobacter pylori to human gastric epithelium mediated by blood group antigens. Science 1993, 262, 1892–1895.

69. Aspholm-Hurtig, M.; Dailide, G.; Lahmann, M.; Kalia, A.; Ilver, D.; Roche, N.; Vikstrom, S.; Sjostrom, R.; Linden, S.; Backstrom, A.; et al. Functional adaptation of baba, the H. Pylori ABO blood group antigen binding adhesin. Science 2004, 305, 519–522.

70. McGuckin, M.A.; Every, A.L.; Skene, C.D.; Linden, S.K.; Chionh, Y.T.; Swierczak, A.; McAuley, J.; Harbour, S.; Kaparakis, M.; Ferrero, R.; et al. MUC1 mucin limits both Helicobacter pylori colonization of the murine gastric mucosa and associated gastritis. Gastroenterology 2007, 133, 1210–1218.

71. McAuley, J.L.; Linden, S.K.; Png, C.W.; King, R.M.; Pennington, H.L.; Gendler, S.J.; Florin, T.H.; Hill, G.R.; Korolik, V.; McGuckin, M.A. MUC1 cell surface mucin is a critical element of the mucosal barrier to infection. J. Clin. Investig. 2007, 117, 2313–2324.

72. Carvalho, F.; Seruca, R.; David, L.; Amorim, A.; Seixas, M.; Bennett, E.; Clausen, H.; SobrinhoSimoes, M. Muc1 gene polymorphism and gastric cancer—An epidemiological study. Glycoconj. J. 1997, 14, 107–111.

73. Franke, A.; McGovern, D.P.B.; Barrett, J.C.; Wang, K.; Radford-Smith, G.L.; Ahmad, T.; Lees, C.W.; Balschun, T.; Lee, J.; Roberts, R.; et al. Genome-wide meta-analysis increases to 71 the number of confirmed crohn's disease susceptibility loci. Nat. Genet. 2010, 42, 1118–1125.

74. Zarepour, M.; Bhullar, K.; Montero, M.; Ma, C.; Huang, T.; Velcich, A.; Xia, L.; Vallance, B.A. The mucin Muc2 limits pathogen burdens and epithelial barrier dysfunction during Salmonella enterica serovar typhimurium colitis. Infect. Immun. 2013, 81, 3672–3683.

75. Bergstrom, K.S.; Kissoon-Singh, V.; Gibson, D.L.; Ma, C.; Montero, M.; Sham, H.P.; Ryz, N.; Huang, T.; Velcich, A.; Finlay, B.B.; et al. Muc2 protects against lethal infectious colitis by disassociating pathogenic and commensal bacteria from the colonic mucosa. PLoS Pathog. 2010, 6, e1000902.

76. Hasnain, S.Z.; Evans, C.M.; Roy, M.; Gallagher, A.L.; Kindrachuk, K.N.; Barron, L.; Dickey, B.F.; Wilson, M.S.; Wynn, T.A.; Grencis, R.K.; et al. MUC5AC: A critical component mediating the rejection of enteric nematodes. J. Exp. Med. 2011, 208, 893–900.

77. Skoog, E.C.; Sjoling, A.; Navabi, N.; Holgersson, J.; Lundin, S.B.; Linden, S.K. Human gastric mucins differently regulate Helicobacter pylori proliferation, gene expression and interactions with host cells. PLoS ONE 2012, 7, e36378.

78. Schreiber, S.; Konradt, M.; Groll, C.; Scheid, P.; Hanauer, G.; Werling, H.O.; Josenhans, C.; Suerbaum, S. The spatial orientation of Helicobacter pylori in the gastric mucus. Proc. Natl. Acad. Sci. USA 2004, 101, 5024–5029.

79. Miller, R.S.; Hoskins, L.C. Mucin degradation in human colon ecosystems. Fecal population densities of mucin-degrading bacteria estimated by a “most probable number” method. Gastroenterology 1981, 81, 759–765.

80. Martens, E.C.; Chiang, H.C.; Gordon, J.I. Mucosal glycan foraging enhances fitness and transmission of a saccharolytic human gut bacterial symbiont. Cell Host Microbe 2008, 4, 447–457.

81. Anderson, K.M.; Ashida, H.; Maskos, K.; Dell, A.; Li, S.C.; Li, Y.T. A clostridial endo-beta-galactosidase that cleaves both blood group a and b glycotopes: The first member of a new glycoside hydrolase family, GH98. J. Biol. Chem. 2005, 280, 7720–7728.

82. Fujita, M.; Tsuchida, A.; Hirata, A.; Kobayashi, N.; Goto, K.; Osumi, K.; Hirose, Y.; Nakayama, J.; Yamanoi, T.; Ashida, H.; et al. Glycoside hydrolase family 89 alpha-n-acetylglucosaminidase from Clostridium perfringens specifically acts on glcnac alpha1,4gal beta1r at the non-reducing terminus of O-glycans in gastric mucin. J. Biol. Chem. 2011, 286, 6479–6489.

83. Hoskins, L.C.; Agustines, M.; McKee, W.B.; Boulding, E.T.; Kriaris, M.; Niedermeyer, G. Mucin degradation in human colon ecosystems. Isolation and properties of fecal strains that degrade abh blood group antigens and oligosaccharides from mucin glycoproteins. J. Clin. Investig. 1985, 75, 944–953.

84. Corfield, A.P.; Wagner, S.A.; O’Donnell, L.J.; Durdey, P.; Mountford, R.A.; Clamp, J.R. The roles of enteric bacterial sialidase, sialate O-acetyl esterase and glycosulfatase in the degradation of human colonic mucin. Glycoconj. J. 1993, 10, 72–81.

85. Png, C.W.; Linden, S.K.; Gilshenan, K.S.; Zoetendal, E.G.; McSweeney, C.S.; Sly, L.I.; McGuckin, M.A.; Florin, T.H. Mucolytic bacteria with increased prevalence in ibd mucosa augment in vitro utilization of mucin by other bacteria. Am. J. Gastroenterol. 2010, 105, 2420–2428.

86. Crost, E.H.; Tailford, L.E.; Le Gall, G.; Fons, M.; Henrissat, B.; Juge, N. Utilisation of mucin glycans by the human gut symbiont ruminococcus gnavus is strain-dependent. PLoS ONE 2013, 8, e76341.

87. Bry, L.; Falk, P.G.; Midtvedt, T.; Gordon, J.I. A model of host-microbial interactions in an open mammalian ecosystem. Science 1996, 273, 1380–1383.

88. Berry, M.; Harris, A.; Lumb, R.; Powell, K. Commensal ocular bacteria degrade mucins. Br. J. Ophthalmol. 2002, 86, 1412–1416.

89. Mothey, D.; Buttaro, B.A.; Piggot, P.J. Mucin can enhance growth, biofilm formation, and survival of streptococcus mutans. FEMS Microbiol. Lett. 2014, 350, 161–167.

90. Tu, Q.V.; McGuckin, M.A.; Mendz, G.L. Campylobacter jejuni response to human mucin MUC2: Modulation of colonization and pathogenicity determinants. J. Med. Microbiol. 2008, 57, 795–802.

91. Byrd, J.C.; Yan, P.; Yunker, C.; Bresalier, R.S. Aberrant expression of gland-type gastric mucin in the surface epithelium of h-pylori-infected patients. Gastroenterology 1997, 113, 455–464.

92. Navabi, N.; Johansson, M.E.; Raghavan, S.; Linden, S.K. Helicobacter pylori infection impairs the mucin production rate and turnover in the murine gastric mucosa. Infect. Immun. 2013, 81, 829–837.

93. Haesebrouck, F.; Pasmans, F.; Flahou, B.; Chiers, K.; Baele, M.; Meyns, T.; Decostere, A.; Ducatelle, R. Gastric helicobacters in domestic animals and nonhuman primates and their significance for human health. Clin. Microbiol. Rev. 2009, 22, 202–223.

94. Yen, J.-T. Swine Nutrition, 2nd ed.; CRC Press Taylor and Francis Group: Boca Raton, FL, USA, 2000.

95. Merchant, H.A.; McConnell, E.L.; Liu, F.; Ramaswamy, C.; Kulkarni, R.P.; Basit, A.W.; Murdan, S. Assessment of gastrointestinal ph, fluid and lymphoid tissue in the guinea pig, rabbit and pig, and implications for their use in drug development. Eur. J. Pharm. Sci. 2011, 42, 3–10.

96. Nordman, H.; Davies, J.R.; Carlstedt, I. Mucus glycoproteins from pig gastric mucosa: Different mucins are produced by the surface epithelium and the glands. Biochem. J. 1998, 331, 687–694.

97. Padra, M.; Adamczyk, B.; Benktander, J.; Flahou, B.; Skoog, E.C.; Padra, J.T.; Smet, A.; Jin, C.; Ducatelle, R.; Samuelsson, T.; et al. Helicobacter suis binding to carbohydrates on human and porcine gastric mucins and glycolipids occurs via two modes. Virulence 2018, 9, 898–918.

98. Karlsson, N.G.; Nordman, H.; Karlsson, H.; Carlstedt, I.; Hansson, G.C. Glycosylation differences between pig gastric mucin populations: A comparative study of the neutral oligosaccharides using mass spectrometry. Biochem. J. 1997, 326 Pt 3, 911–917.

99. Nordman, H.; Davies, J.R.; Herrmann, A.; Karlsson, N.G.; Hansson, G.C.; Carlstedt, I. Mucus glycoproteins from pig gastric mucose: Identification of different mucin populations from the surface epithelium. Biochem. J. 1997, 326, 903–910.

100. Wilberts, B.L.; Arruda, P.H.; Kinyon, J.M.; Madson, D.M.; Frana, T.S.; Burrough, E.R. Comparison of lesion severity, distribution, and colonic mucin expression in pigs with acute swine dysentery following oral inoculation with “Brachyspira hampsonii” or Brachyspira hyodysenteriae. Vet. Pathol. 2014, 51, 1069–1108.

101. Quintana-Hayashi, M.P.; Mahu, M.; De Pauw, N.; Boyen, F.; Pasmans, F.; Martel, A.; Premaratne, P.; Fernandez, H.R.; Teymournejad, O.; Vande Maele, L.; et al. The levels of Brachyspira hyodysenteriae binding to porcine colonic mucins differ between individuals, and binding is increased to mucins from infected pigs with de novo MUC5AC synthesis. Infect. Immun. 2015, 83, 1610–1619.

102. Forder, R.E.; Nattrass, G.S.; Geier, M.S.; Hughes, R.J.; Hynd, P.I. Quantitative analyses of genes associated with mucin synthesis of broiler chickens with induced necrotic enteritis. Poult. Sci. 2012, 91, 1335–1341.

103. Struwe, W.B.; Gough, R.; Gallagher, M.E.; Kenny, D.T.; Carrington, S.D.; Karlsson, N.G.; Rudd, P.M. Identification of O-glycan structures from chicken intestinal mucins provides insight into campylobactor jejuni pathogenicity. Mol. Cell. Proteom. 2015, 14, 1464–1477.

104. Hoorens, P.R.; Rinaldi, M.; Li, R.W.; Goddeeris, B.; Claerebout, E.; Vercruysse, J.; Geldhof, P. Genome wide analysis of the bovine mucin genes and their gastrointestinal transcription profile. BMC Genom. 2011, 12, 140.

105. Montagne, L.; Toullec, R.; Lalles, J.P. Calf intestinal mucin: Isolation, partial characterization, and measurement in ileal digesta with an enzyme-linked immunosorbent assay. J. Dairy Sci. 2000, 83, 507–517.

106. Perez-Sanchez, J.; Estensoro, I.; Redondo, M.J.; Calduch-Giner, J.A.; Kaushik, S.; Sitja-Bobadilla, A. Mucins as diagnostic and prognostic biomarkers in a fish-parasite model: Transcriptional and functional analysis. PLoS ONE 2013, 8, e65457.

107. Sveen, L.R.; Grammes, F.T.; Ytteborg, E.; Takle, H.; Jørgensen, S.M. Genome-wide analysis of atlantic salmon (Salmo salar) mucin genes and their role as biomarkers. PLoS ONE 2017, 12, e0189103.

108. Neuhaus, H.; van der Marel, M.; Caspari, N.; Meyer, W.; Enss, M.L.; Steinhagen, D. Biochemical and histochemical study on the intestinal mucosa of the common carp Cyprinus carpio L., with special consideration of mucin glycoproteins. J. Fish Biol. 2007, 70, 1523–1534.

109. Fiertak, A.; Kilarski, W. Glycoconjugates of the Intestinal Goblet Cells of Four Cyprinids. Cell. Mol. Life Sci. 2002, 59, 1724–1733.

110. Conway, P.L.; Gorbach, S.L.; Goldin, B.R. Survival of lactic acid bacteria in the human stomach and adhesion to intestinal cells. J. Dairy Sci. 1987, 70, 1–12.

111. Motta, V.; Trevisi, P.; Bertolini, F.; Ribani, A.; Schiavo, G.; Fontanesi, L.; Bosi, P. Exploring gastric bacterial community in young pigs. PLoS ONE 2017, 12, e0173029.

112. Yang, F.; Hou, C.; Zeng, X.; Qiao, S. The use of lactic acid bacteria as a probiotic in swine diets. Pathogens 2015, 4, 34–45.

113. Alexopoulos, C.; Georgoulakis, I.E.; Tzivara, A.; Kritas, S.K.; Siochu, A.; Kyriakis, S.C. Field evaluation of the efficacy of a probiotic containing bacillus licheniformis and bacillus subtilis spores, on the health status and performance of sows and their litters. J. Anim. Physiol. Anim. Nutr. 2004, 88, 381–392.

114. Yeung, C.Y.; Chiang Chiau, J.S.; Chan, W.T.; Jiang, C.B.; Cheng, M.L.; Liu, H.L.; Lee, H.C. In vitro prevention of Salmonella lipopolysaccharide-induced damages in epithelial barrier function by various lactobacillus strains. Gastroenterol. Res. Pract. 2013, 2013, 973209.

115. Liu, H.; Zhang, J.; Zhang, S.; Yang, F.; Thacker, P.A.; Zhang, G.; Qiao, S.; Ma, X. Oral administration of lactobacillus fermentum I5007 favors intestinal development and alters the intestinal microbiota in formula-fed piglets. J. Agric. Food Chem. 2014, 62, 860–866.

116. Rauch, M.; Lynch, S.V. Probiotic manipulation of the gastrointestinal microbiota. Gut Microbes 2010, 1, 335–338.

117. Stevens, C.E.; Hume, I.D. Contributions of microbes in vertebrate gastrointestinal tract to production and conservation of nutrients. Physiol. Rev. 1998, 78, 393–427.

118. Kim, B.C.; Park, D.S.; Kim, H.; Oh, H.W.; Lee, K.H.; Shin, K.S.; Bae, K.S. Herbiconiux Moechotypicola sp. Nov., a xylanolytic bacterium isolated from the gut of hairy long-horned toad beetles, moechotypa diphysis (pascoe). Int. J. Syst. Evol. Microbiol. 2012, 62, 90–95.

119. Hu, X.; Yu, J.M.; Wang, C.Y.; Chen, H. Cellulolytic bacteria associated with the gut of dendroctonus armandi larvae (Coleoptera: Curculionidae: Scolytinae). Forests 2014, 5, 455–465.

120. Queiroz, D.M.; Rocha, G.A.; Mendes, E.N.; Lage, A.P.; Carvalho, A.C.; Barbosa, A.J. A spiral microorganism in the stomach of pigs. Vet. Microbiol. 1990, 24, 199–204.

121. Roosendaal, R.; Vos, J.H.; Roumen, T.; van Vugt, R.; Cattoli, G.; Bart, A.; Klaasen, H.L.; Kuipers, E.J.; Vandenbroucke-Grauls, C.M.; Kusters, J.G. Slaughter pigs are commonly infected by closely related but distinct gastric ulcerative lesion-inducing gastrospirilla. J. Clin. Microbiol. 2000, 38, 2661–2664.

122. Krakowka, S.; Ringler, S.S.; Flores, J.; Kearns, R.J.; Eaton, K.A.; Ellis, J.A. Isolation and preliminary characterization of a novel helicobacter species from swine. Am. J. Vet. Res. 2005, 66, 938–944.

123. Park, J.H.; Lee, B.J.; Lee, Y.S.; Park, J.H. Association of tightly spiraled bacterial infection and gastritis in pigs. J. Vet. Med. Sci. 2000, 62, 725–729.

124. De Witte, C.; Devriendt, B.; Flahou, B.; Bosschem, I.; Ducatelle, R.; Smet, A.; Haesebrouck, F. Helicobacter suis induces changes in gastric inflammation and acid secretion markers in pigs of different ages. Vet. Res. 2017, 48, 34.

125. Hellemans, A.; Chiers, K.; De Bock, M.; Decostere, A.; Haesebrouck, F.; Ducatelle, R.; Maes, D. Prevalence of ‘candidatus Helicobacter suis’ in pigs of different ages. Vet. Rec. 2007, 161, 189–192.

126. Zhang, G.; Ducatelle, R.; Mihi, B.; Smet, A.; Flahou, B.; Haesebrouck, F. Helicobacter suis affects the health and function of porcine gastric parietal cells. Vet. Res. 2016, 47, 101.

127. Danon, S.J.; Lee, A. Other gastric helicobacters and spiral organisms. In Helicobacter Pylori: Physiology and Genetics; Mobley, H.L.T., Mendz, G.L., Hazell, S.L., Eds.; ASM Press: Washington, DC, USA, 2001.

128. Hellemans, A.; Chiers, K.; Decostere, A.; De Bock, M.; Haesebrouck, F.; Ducatelle, R. Experimental infection of pigs with ‘candidatus Helicobacter suis’. Vet. Res. Commun. 2007, 31, 385–395.

129. Lindahl, M.; Carlstedt, I. Binding of k99 fimbriae of enterotoxigenic Escherichia coli to pig small intestinal mucin glycopeptides. J. Gen. Microbiol. 1990, 136, 1609–1614.

130. Kyogashima, M.; Ginsburg, V.; Krivan, H.C. Escherichia coli K99 binds to N-glycolylsialoparagloboside and N-glycolyl-GM3 found in piglet small intestine. Arch. Biochem. Biophys. 1989, 270, 391–397.

131. Teneberg, S.; Willemsen, P.; de Graaf, F.K.; Karlsson, K.A. Receptor-active glycolipids of epithelial cells of the small intestine of young and adult pigs in relation to susceptibility to infection with Escherichia coli K99. FEBS Lett. 1990, 263, 10–14.

132. Kim, C.H.; Kim, D.; Ha, Y.; Cho, K.D.; Lee, B.H.; Seo, I.W.; Kim, S.H.; Chae, C. Expression of mucins and trefoil factor family protein-1 in the colon of pigs naturally infected with Salmonella typhimurium. J. Comp. Pathol. 2009, 140, 38–42.

133. Kringel, H.; Iburg, T.; Dawson, H.; Aasted, B.; Roepstorff, A. A time course study of immunological responses in Trichuris suis infected pigs demonstrates induction of a local type 2 response associated with worm burden. Int. J. Parasitol. 2006, 36, 915–924.

134. Theodoropoulos, G.; Hicks, S.J.; Corfield, A.P.; Miller, B.G.; Kapel, C.M.; Trivizaki, M.; Balaskas, C.; Petrakos, G.; Carrington, S.D. Trichinella spiralis: Enteric mucin-related response to experimental infection in conventional and spf pigs. Exp. Parasitol. 2005, 109, 63–71.

135. Rinaldi, M.; Dreesen, L.; Hoorens, P.R.; Li, R.W.; Claerebout, E.; Goddeeris, B.; Vercruysse, J.; van Den Broek, W.; Geldhof, P. Infection with the gastrointestinal nematode ostertagia ostertagi in cattle affects mucus biosynthesis in the abomasum. Vet. Res. 2011, 42, 61.

136. Li, R.W.; Li, C.; Elsasser, T.H.; Liu, G.; Garrett, W.M.; Gasbarre, L.C. Mucin biosynthesis in the bovine goblet cell induced by Cooperia oncophora infection. Vet. Parasitol. 2009, 165, 281–289.

137. Mouricout, M.A.; Julien, R.A. Pilus-mediated binding of bovine enterotoxigenic Escherichia coli to calf small intestinal mucins. Infect. Immun. 1987, 55, 1216–1223.

138. Padra, J.T.; Sundh, H.; Jin, C.; Karlsson, N.G.; Sundell, K.; Linden, S.K. Aeromonas salmonicida binds differentially to mucins isolated from skin and intestinal regions of atlantic salmon in an N-acetylneuraminic acid-dependent manner. Infect. Immun. 2014, 82, 5235–5245.

139. Li, C.; Zhang, Y.; Wang, R.; Lu, J.; Nandi, S.; Mohanty, S.; Terhune, J.; Liu, Z.; Peatman, E. Rna-seq analysis of mucosal immune responses reveals signatures of intestinal barrier disruption and pathogen entry following Edwardsiella ictaluri infection in channel catfish, ictalurus punctatus. Fish Shellfish Immunol. 2012, 32, 816–827.

140. Balebona, M.C.; Andreu, M.J.; Bordas, M.A.; Zorrilla, I.; Moriñigo, M.A.; Borrego, J.J. Pathogenicity of Vibrio alginolyticus for cultured gilt-head sea bream (Sparus aurata L.). Appl. Environ. Microbiol. 1998, 64, 4269–4275.

141. Johansson, M.E.; Phillipson, M.; Petersson, J.; Velcich, A.; Holm, L.; Hansson, G.C. The inner of the two Muc2 mucin-dependent mucus layers in colon is devoid of bacteria. Proc. Natl. Acad. Sci. USA 2008, 105, 15064–15069.

142. Gustafsson, J.K.; Navabi, N.; Rodriguez-Pineiro, A.M.; Alomran, A.H.; Premaratne, P.; Fernandez, H.R.; Banerjee, D.; Sjovall, H.; Hansson, G.C.; Linden, S.K. Dynamic changes in mucus thickness and ion secretion during citrobacter rodentium infection and clearance. PLoS ONE 2013, 8, e84430.

143. Marshall, T.; Allen, A. The isolation and characterization of the high-molecular-weight glycoprotein from pig colonic mucus. Biochem. J. 1978, 173, 569–578.

144. Priori, D.; Colombo, M.; Koopmans, S.J.; Jansman, A.J.; van der Meulen, J.; Trevisi, P.; Bosi, P. The A0 blood group genotype modifies the jejunal glycomic binding pattern profile of piglets early associated with a simple or complex microbiota. J. Anim. Sci. 2016, 94, 592–601.

145. Quintana-Hayashi, M.P.; Navabi, N.; Mahu, M.; Venkatakrishnan, V.; Fernandez, H.R.; Haesebrouck, F.; Pasmans, F.; Linden, S.K. Neutrophil elastase and il17 expressed in the pig colon during Brachyspira hyodysenteriae infection synergistically with the pathogen induce increased mucus transport speed and production via MAPK3. Infect. Immun. 2017.

146. Hu, J.; Nie, Y.; Chen, J.; Zhang, Y.; Wang, Z.; Fan, Q.; Yan, X. Gradual changes of gut microbiota in weaned miniature piglets. Front. Microbiol. 2016, 7, 1727.

147. Chen, L. The maturing development of gut microbiota in commercial piglets during the weaning transition. Front. Microbiol. 2017, 8, 1688.

148. Kelly, D.; King, T.P. The influence of lactation products on the temporal expression of histo-blood group antigens in the intestines of suckling pigs: Lectin histochemical and immunohistochemical analysis. Histochem. J. 1991, 23, 55–60.

149. Kelly, D.; King, T.P.; McFadyen, M.; Coutts, A.G. Effect of preclosure colostrum intake on the development of the intestinal epithelium of artificially reared piglets. Biol. Neonate 1993, 64, 235–244.

150. Liu, P.; Pieper, R.; Rieger, J.; Vahjen, W.; Davin, R.; Plendl, J.; Meyer, W.; Zentek, J. Effect of dietary zinc oxide on morphological characteristics, mucin composition and gene expression in the colon of weaned piglets. PLoS ONE 2014, 9, e91091.

151. Pestova, M.I.; Clift, R.E.; Vickers, R.J.; Franklin, M.A.; Mathew, A.G. Effect of weaning and dietary galactose supplementation on digesta glycoproteins in pigs. J. Sci. Food Agric. 2000, 80, 1918–1924.

152. King, T.P.; Begbie, R.; Slater, D.; McFadyen, M.; Thom, A.; Kelly, D. Sialylation of intestinal microvillar membranes in newborn, sucking and weaned pigs. Glycobiology 1995, 5, 525–534.

153. Kennedy, M.J.; Yancey, R.J., Jr. Motility and chemotaxis in serpulina hyodysenteriae. Vet. Microbiol. 1996, 49, 21–30.

154. Milner, J.A.; Sellwood, R. Chemotactic response to mucin by serpulina hyodysenteriae and other porcine spirochetes: Potential role in intestinal colonization. Infect. Immun. 1994, 62, 4095–4099.

155. Naresh, R.; Hampson, D.J. Attraction of brachyspira pilosicoli to mucin. Microbiology 2010, 156, 191–197.

156. Sando, L.; Pearson, R.; Gray, C.; Parker, P.; Hawken, R.; Thomson, P.C.; Meadows, J.R.; Kongsuwan, K.; Smith, S.; Tellam, R.L. Bovine MUC1 is a highly polymorphic gene encoding an extensively glycosylated mucin that binds bacteria. J. Dairy Sci. 2009, 92, 5276–5291.

157. Parker, P.; Sando, L.; Pearson, R.; Kongsuwan, K.; Tellam, R.L.; Smith, S. Bovine MUC1 inhibits binding of enteric bacteria to caco-2 cells. Glycoconj. J. 2010, 27, 89–97.

158. Linden, S.K.; Sheng, Y.H.; Every, A.L.; Miles, K.M.; Skoog, E.C.; Florin, T.H.J.; Sutton, P.; McGuckin, M.A. MUC1 limits Helicobacter pylori infection both by steric hindrance and by acting as a releasable decoy. PLoS Pathog. 2009, 5.

159. Meijerink, E.; Neuenschwander, S.; Fries, R.; Dinter, A.; Bertschinger, H.U.; Stranzinger, G.; Vogeli, P. A DNA polymorphism influencing alpha(1,2)fucosyltransferase activity of the pig fut1 enzyme determines susceptibility of small intestinal epithelium to Escherichia coli f18 adhesion. Immunogenetics 2000, 52, 129–136.

160. Stanley, R.A.; Ram, S.P.; Wilkinson, R.K.; Roberton, A.M. Degradation of pig gastric and colonic mucins by bacteria isolated from the pig colon. Appl. Environ. Microbiol. 1986, 51, 1104–1109.

161. Levine, U.Y.; Looft, T.; Allen, H.K.; Stanton, T.B. Butyrate-producing bacteria, including mucin degraders, from the swine intestinal tract. Appl. Environ. Microbiol. 2013, 79, 3879–3881.

162. Looft, T.; Levine, U.Y.; Stanton, T.B. Cloacibacillus porcorum sp. Nov., a mucin-degrading bacterium from the swine intestinal tract and emended description of the genus cloacibacillus. Int. J. Syst Evol. Microbiol. 2013, 63, 1960–1966.

163. Svihus, B. Function of the digestive system1. J. Appl. Poult. Res. 2014, 23, 306–314.

164. Lang, T.; Hansson, G.C.; Samuelsson, T. An inventory of mucin genes in the chicken genome shows that the mucin domain of MUC13 is encoded by multiple exons and that ovomucin is part of a locus of related gel-forming mucins. BMC Genom. 2006, 7, 197.

165. Jiang, Z.; Applegate, T.J.; Lossie, A.C. Cloning, annotation and developmental expression of the chicken intestinal MUC2 gene. PLoS ONE 2013, 8, e53781.

166. Collier, C.T.; Hofacre, C.L.; Payne, A.M.; Anderson, D.B.; Kaiser, P.; Mackie, R.I.; Gaskins, H.R. Coccidia-induced mucogenesis promotes the onset of necrotic enteritis by supporting Clostridium perfringens growth. Vet. Immunol. Immunopathol. 2008, 122, 104–115.

167. Tierney, J.B.; Matthews, E.; Carrington, S.D.; Mulcahy, G. Interaction of eimeria tenella with intestinal mucin in vitro. J. Parasitol. 2007, 93, 634–638.

168. Young, K.T.; Davis, L.M.; Dirita, V.J. Campylobacter jejuni: Molecular biology and pathogenesis. Nat. Rev. Microbiol. 2007, 5, 665–679.

169. Alemka, A.; Whelan, S.; Gough, R.; Clyne, M.; Gallagher, M.E.; Carrington, S.D.; Bourke, B. Purified chicken intestinal mucin attenuates campylobacter jejuni pathogenicity in vitro. J. Med. Microbiol. 2010, 59, 898–903.

170. Byrne, C.M.; Clyne, M.; Bourke, B. Campylobacter jejuni adhere to and invade chicken intestinal epithelial cells in vitro. Microbiology 2007, 153, 561–569.

171. Hugdahl, M.B.; Beery, J.T.; Doyle, M.P. Chemotactic behavior of campylobacter jejuni. Infect. Immun. 1988, 56, 1560–1566.

172. Day, C.J.; Tiralongo, J.; Hartnell, R.D.; Logue, C.A.; Wilson, J.C.; von Itzstein, M.; Korolik, V. Differential carbohydrate recognition by campylobacter jejuni strain 11168: Influences of temperature and growth conditions. PLoS ONE 2009, 4, e4927.

173. Stahl, M.; Friis, L.M.; Nothaft, H.; Liu, X.; Li, J.; Szymanski, C.M.; Stintzi, A. L-fucose utilization provides campylobacter jejuni with a competitive advantage. Proc. Natl. Acad. Sci. USA 2011, 108, 7194–7199.

174. Gasbarre, L.C.; Leighton, E.A.; Sonstegard, T. Role of the bovine immune system and genome in resistance to gastrointestinal nematodes. Vet. Parasitol. 2001, 98, 51–64.

175. Nataro, J.P.; Kaper, J.B. Diarrheagenic Escherichia coli. Clin. Microbiol. Rev. 1998, 11, 142–201.

176. Aperce, C.C.; Heidenreich, J.M.; Drouillard, J.S. Capacity of the bovine intestinal mucus and its components to support growth of Escherichia coli o157:H7. Animal 2014, 8, 731–737.

177. Bertin, Y.; Chaucheyras-Durand, F.; Robbe-Masselot, C.; Durand, A.; de la Foye, A.; Harel, J.; Cohen, P.S.; Conway, T.; Forano, E.; Martin, C. Carbohydrate utilization by enterohaemorrhagic Escherichia coli o157:H7 in bovine intestinal content. Environ. Microbiol. 2013, 15, 610–622.

178. Muraoka, W.T.; Zhang, Q. Phenotypic and genotypic evidence for l-fucose utilization by campylobacter jejuni. J. Bacteriol. 2011, 193, 1065–1075.

179. Almagro-Moreno, S.; Boyd, E.F. Sialic acid catabolism confers a competitive advantage to pathogenic vibrio cholerae in the mouse intestine. Infect. Immun. 2009, 77, 3807–3816.

180. Lokka, G.; Austbo, L.; Falk, K.; Bjerkas, I.; Koppang, E.O. Intestinal morphology of the wild atlantic salmon (Salmo salar). J. Morphol. 2013, 274, 859–876.

181. Lang, T.; Alexandersson, M.; Hansson, G.C.; Samuelsson, T. Bioinformatic identification of polymerizing and transmembrane mucins in the puffer fish fugu rubripes. Glycobiology 2004, 14, 521–527.

182. Lang, T.; Hansson, G.C.; Samuelsson, T. Gel-forming mucins appeared early in metazoan evolution. Proc. Natl. Acad. Sci. USA 2007, 104, 16209–16214.

183. Micallef, G.; Bickerdike, R.; Reiff, C.; Fernandes, J.M.; Bowman, A.S.; Martin, S.A. Exploring the transcriptome of atlantic salmon (Salmo salar) skin, a major defense organ. Mar. Biotechnol. (N. Y.) 2012, 14, 559–569.

184. Andrews, M.; Battaglene, S.; Cobcroft, J.; Adams, M.; Noga, E.; Nowak, B. Host response to the chondracanthid copepod chondracanthus goldsmidi, a gill parasite of the striped trumpeter, latris lineata (Forster), in tasmania. J. Fish Dis. 2010, 33, 211–220.

185. Neuhaus, H.; van der Marel, M.; Caspari, N.; Meyer, W.; Enss, M.L.; Steinhagen, D. Biochemical and histochemical effects of perorally applied endotoxin on intestinal mucin glycoproteins of the common carp Cyprinus carpio. Dis. Aquat. Org. 2007, 77, 17–27.

186. Westerdahl, A.; Olsson, J.C.; Kjelleberg, S.; Conway, P.L. Isolation and characterization of turbot (Scophtalmus maximus)-associated bacteria with inhibitory effects against vibrio anguillarum. Appl. Environ. Microbiol. 1991, 57, 2223–2228.

187. Jevtov, I.; Samuelsson, T.; Yao, G.; Amsterdam, A.; Ribbeck, K. Zebrafish as a model to study live mucus physiology. Sci. Rep. 2014, 4, 6653.

188. Mitchell, K.C.; Breen, P.; Britton, S.; Neely, M.N.; Withey, J.H. Quantifying vibrio cholerae enterotoxicity in a zebrafish infection model. Appl. Environ. Microbiol. 2017, 83, e00783-17.

189. Bordas, M.A.; Balebona, M.C.; Rodriguez-Maroto, J.M.; Borrego, J.J.; Moriñigo, M.A. Chemotaxis of pathogenic vibrio strains towards mucus surfaces of gilt-head sea bream (Sparus aurata L.). Appl. Environ. Microbiol. 1998, 64, 1573–1575.

190. Wolfe, K.G.; Plumb, J.A.; Morrison, E.E. Lectin binding characteristics of the olfactory mucosa of channel catfish: Potential factors in attachment of Edwardsiella ictaluri. J. Aquat. Anim. Health 1998, 10, 348–360.