Infection with the gastrointestinal nematode Ostertagia ostertagi in cattle affects mucus biosynthesis in the abomasum
© Rinaldi et al; licensee BioMed Central Ltd. 2011
Received: 14 January 2011
Accepted: 11 May 2011
Published: 11 May 2011
The mucus layer in the gastrointestinal (GI) tract is considered to be the first line of defense to the external environment. Alteration in mucus components has been reported to occur during intestinal nematode infection in ruminants, but the role of mucus in response to abomasal parasites remains largely unclear. The aim of the current study was to analyze the effects of an Ostertagia ostertagi infection on the abomasal mucus biosynthesis in cattle. Increased gene expression of MUC1, MUC6 and MUC20 was observed, while MUC5AC did not change during infection. Qualitative changes of mucins, related to sugar composition, were also observed. AB-PAS and HID-AB stainings highlighted a decrease in neutral and an increase in acidic mucins, throughout the infection. Several genes involved in mucin core structure synthesis, branching and oligomerization, such as GCNT3, GCNT4, A4GNT and protein disulphide isomerases were found to be upregulated. Increase in mucin fucosylation was observed using the lectin UEA-I and through the evaluation of fucosyltransferases gene expression levels. Finally, transcription levels of 2 trefoil factors, TFF1 and TFF3, which are co-expressed with mucins in the GI tract, were also found to be significantly upregulated in infected animals. Although the alterations in mucus biosynthesis started early during infection, the biggest effects were found when adult worms were present on the surface of the abomasal mucosa and are likely caused by the alterations in mucosal cell populations, characterized by hyperplasia of mucus secreting cells.
The mucus layer in the gastrointestinal (GI) tract forms the first line of defense to the external environment. Mucins (MUC) or mucus glycoproteins are one of the most important components of the mucus barrier and they are classified as membrane bound and secreted mucins depending on their function and location. In cattle, 9 membrane-associated mucins and 8 secreted mucins have been identified . Membrane bound mucins are mostly present on the apical membrane of epithelial cells, where they have been suggested to play a role in cell signaling, while secreted gel forming ones are capable of forming oligomers, crucial in the formation of the visco-elastic mucus gel . Mucins are highly glycosylated proteins, with most of the oligosaccharides attached by O-links to the repetitive sequences rich in threonine and serine. Previous studies have suggested that mucin glycosylation not only protects the mucins from proteolytic enzymes, but it can also be altered in response to mucosal infection and inflammation, probably with protective effects . The carbohydrate structures present on mucins are determined by the expression of specific glycosyltransferases. Thus mucin glycosylation is governed by tissue-specific enzyme expression, host and environmental factors influencing transferase expression .
Trefoil factors (TFF1, TFF2 and TFF3) are also major secretory products of normal mucus-secreting cells in the epithelium in the GI tract. In humans these factors are known to be co-expressed with mucins in mucus producing cells in the stomach and duodenum [5, 6] and they seem to play a major role in wounding responses to maintain mucosal surface integrity, as well as in pathological processes [7, 8]. It has been reported that TFF2, after addition to a mucin solution, significantly increases viscosity and elasticity of the mucus .
Although the precise role of mucus in host defense against nematode infections is not known, previous studies have shown that alteration in mucin production and glycosylation may be related to the capacity of the host to expel GI nematodes. For instance, increased Muc2 production in the intestine during infection with Trichuris muris was observed only in resistant mice and it was correlated with worm expulsion . Moreover levels of Muc4, Muc13 and Muc17 were increased during acute and chronic infections with T. muris, causing thickening of the glycocalix in the intestine . On the other hand, MUC5AC was found to be strongly downregulated in the abomasum of sheep infected with Haemonchus contortus , as well as in experimentally infected sheep selected for resistance to nematode infections . In cattle, an infection with the intestinal nematode Cooperia oncophora also altered the mucus composition, causing induction of MUC2 and a downregulation of MUC5B in the small intestine . Changes in terminal sugars of goblet cells mucins, such as a strong expression of terminal N-acetyl-D-galactosamine, were also observed in rats infected with the intestinal parasite Nippostrongylus brasiliensis around the time of immune-mediated expulsion . Previous studies have also observed important changes in TFF expression during GI nematode infections. Rowe et al.,  have reported an induction of TFF3 accompanied by a progressive loss of TFF2 in the abomasal mucosa of sheep infected with H. contortus. TFF3 was also found to be significantly upregulated in the small intestine of sheep infected with Trichostrongylus columbriformis  and in rats during infection with N. brasiliensis .
The abomasal nematode Ostertagia ostertagi causes a loss in animal production due to depression in food intake and impaired gastrointestinal functions . The presence of larvae and adult worms in the abomasum is accompanied by morphological changes in the host mucosa, such as mucous cell hyperplasia, superficial epithelial damage and loss of acid-producing parietal cells [18–20]. To date little is known about the effects of these cellular changes on the composition of the abomasal mucus layer. The purpose of the current study was to analyze in more detail the effects of an Ostertagia infection in the abomasum on the mucus biosynthesis, i.e. mucins, TFFs, and a set of selected glycogenes and disulphide isomerases involved in the synthesis and oligomerization of mucins.
Materials and methods
Experimental design and tissue collection
Nematode-free Holstein calves, aged 6 to 8 months, were randomly selected and divided in experimental groups. The animals were kept indoors to prevent accidental infection with nematode parasites. The calves were fed hay and commercial pellets, and given ad libitum access to water. For gene transcription analyses, a trial including 4 groups of animals (n = 4 in each group) was performed (Trial 1). Animals sacrified at day 0 were used as negative controls, the remaining animals were orally infected with 100 000 O. ostertagi L3 larvae/animal and killed at 6, 9, and 24 days post infection (dpi). These time points corresponded with the presence of late L3/early L4, L4 and adult worm stages in the abomasum, respectively, as observed at slaughtering. For histological analyses, a second trial including three groups of animals (n = 3 in each group) was performed (Trial 2). Animals sacrified at day 0 were used as negative controls, the remaining animals were orally infected with 100 000 O. ostertagi L3 larvae/animal and killed at 14 and 21 dpi. One additional group of four animals was maintained for 60 days on pasture to acquire a natural O. ostertagi infection before euthanasia was performed at 60 days post exposure (dpe). Samples from these animals were collected for both gene transcription and histological analyses. For gene transcription analyses, tissue samples were collected from the fundic region of the abomasum, snap frozen in liquid nitrogen and stored at -80°C until RNA was extracted. For histological analyses abomasal tissues were stored in Carnoy's solution (60% ethanol, 30% chloroform, 10% glacial acetic acid) and in 10% formaldehyde in phosphate buffered saline (PBS), both freshly made. All experiments had been approved by the ethical committee of the Faculty of Veterinary Medicine at Ghent University.
RNA extraction and cDNA synthesis
Total RNA was extracted from tissue samples using Trizol (Invitrogen) and further purified using the RNeasy Mini kit (Qiagen). To remove contaminating genomic DNA (gDNA), on-column DNase digestion was performed using the RNase-free DNase set (Qiagen) according to the manufacturer's instructions. RNA quality was verified using an Experion™Automated Electophoresis System (Bio-Rad), and concentrations were determined using a NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies). Genomic DNA contamination was checked by the SuperScript One-Step RT PCR (Invitrogen) using intron-spanning primers for gapdh (Additional file 1).
Quantitative Real-time PCR
Quantitative Real-time PCR (qRT-PCR) was performed for the following genes (Additional file 1 Table S1): mucins (MUC1, MUC2, MUC5AC, MUC5B, MUC6, MUC20); trefoil factors (TFF1, TFF2, TFF3); glycosyltransferases [alpha-1, 4-N-acetylglucosaminyltransferase (A4GNT); betaGal beta-1,3-N-acetylglucosaminyltransferase 3 (B3GNT3); core 1 synthase, glycoprotein-N-acetylgalactosamine 3-beta-galactosyltransferase 1 (C1GALT1); glucosaminyl (N-acetyl) transferase 2, I-branching enzyme (I blood group) (GCNT2); glucosaminyl (N-acetyl) transferase 3, mucin type (GCNT3); glucosaminnyl (N-acetyl) transferase 4, core 2 (GCNT4)]; sulfotransferases [heparan sulfate (glucosamine) 3-O-sulfotransferase 1 (HS3ST1); heparan sulfate (glucosamine) 3-O-sulfotransferase 2 (HS3ST2); galactose-3-O-sulfotransferase 1 (GAL3ST1); sialyltransferases [ST3 beta-galactoside alpha-2,3-sialyltransferase 4 (ST3GAL4)]; fucosyltransferases (FUT1, FUT2, FUT4, FUT10); and disulphide isomerases [anterior gradient homolog 2 (AGR2); protein disulfide isomerase family A, member 3 (PDIA3); protein disulfide isomerase family A, member 4 (PDIA4)].
One μg of total RNA was converted to cDNA using the iScript cDNA synthesis kit (Bio-Rad), following the manufacturer's instructions. Real-time-PCR analyses was carried out with the SYBR Green Master Mix (Applied Biosystems) using 400 nM of each amplification primer and 2 μL of single-stranded cDNA (10 ng of the input total RNA equivalent) in a 20 μL reaction volume. Amplification cycles were performed on a StepOnePlus Real-Time PCR System (Applied Biosystems) under the following conditions: 95°C for 20 s followed by 35 cycles of 95°C for 5 s and optimal annealing temperature (Ta) for 30 s (Additional file 1). The primer sets used to amplify the different genes were designed using the Primer3 software http://frodo.wi.mit.edu/primer3/ and are listed in Additional file 1. Reaction efficiencies were measured based on a standard curve using dilution series of pooled cDNA from all the samples. A melting curve analysis was performed at the end of the reaction to ensure specificity of the primers. In addition, PCR products were cloned in the pGEM-T vector according to the manufacturer's instruction (Promega) and sequenced. Every assay included cDNA samples in duplicate and a non-template control. Ct values were transformed in relative quantity using the delta Ct method applying the formula: Q = E (min Ct - sample Ct), with Q = sample quantity relative to sample with highest expression, E = amplification efficiency, and min Ct = lowest Ct value = Ct value of sample with the highest expression level. To minimize technical mistakes and therefore misinterpretation of results, qRT-PCRs for each gene were carried out on the same plate with all samples.
Statistical analysis was carried out using GraphPad Prism software. The Nonparametric Mann Whitney U test was used to determine differences between infected and control groups. A P-value of ≤ 0,05 was considered significant.
Immediately after collection, tissue samples were placed in Carnoy's solution for 24 h at room temperature (RT) and then in 70% ethanol at RT. The samples were then dehydrated through a graded ethanol, cleared in xylene, and embedded in paraffin. For histochemical examination, serial paraffin sections were cut at 8 μm thickness. Tissue sections were deparaffinized in xylene and isopropanol followed by rehydration through a graded ethanol series and stained with periodic acid Schiff (PAS) , Alcian blue (AB)-PAS (pH 2.5) [22, 23] and High iron diamine (HID)-AB stainings as previously described  with slight modifications. Briefly, for PAS staining samples were incubated with periodic acid for 10 min, washed, and covered with Schiff's solution (Merck) for 20 min. A wash in sulfite water (12 mL Na2S2O5 + 10 mL HCl + 200 mL distilled water) for 15 min was followed by counterstain with hematoxylin. For the AB-PAS staining, samples were incubated in AB solution for 15 min, rinsed in water and incubated with Schiff's solution for 15 min. Following a 15 min wash in sulfite water they were counterstained with hematoxylin. Finally for the HID-AB staining, sections were incubated in diamine solution for 16 h and then counterstained with 1% AB in 3% acetic acid for 10 min. All the sections were then dehydrated in ethanol, cleared in xylene and mounted in synthetic medium for observation using light microscopy. Sections from control (0 dpi; n = 3) and infected animals (14 and 21 dpi, 60 dpe; n = 3) were used. Three pictures for each section were taken and analyzed.
Immunohistochemistry for TFF3
Immediately after collection, tissue samples were placed in 10% formaldehyde for 24 h at RT followed by distilled water for 1 h at RT and then 70% ethanol at RT. The samples were then dehydrated through a graded ethanol, cleared in xylene, and embedded in paraffin. Serial paraffin sections were cut at 5 μm thickness and mounted onto APES-coated glass slides. Tissue sections were deparaffinized in xylene and isopropanol, followed by rehydration through a graded ethanol series and immunostained with anti-hTFF3 (rabbit polyclonal, 1:100, Santa Cruz Biotechnology). Antigen retrieval was performed by incubating the slides in citrate buffer, microwave heating, and cooling for 30 min at 4°C. Non specific staining was blocked with 1% of BSA and 0.3% Triton X-100 pH 7.5 (Sigma) in PBS for 15 min at RT, followed by incubation with 1% of BSA in PBS for 30 min at RT. Sections were subsequentely incubated overnight at 4°C with the primary antibody diluted in 1% BSA in PBS. Slides were covered with Alexa Fluor® 488 goat anti-rabbit IgG (H+L) (Invitrogen) for 1 h at RT. Counterstaining was performed with DAPI (4',6-diamidino-2-phenylindole, dilactate; 1:1000 in PBS; Invitrogen) for 5 min at RT. Then sections were dehydrated in ethanol, cleared in xylene and mounted in synthetic medium for observation using fluorescent microscopy. Negative control stainings were carried out using the same procedure except that the primary antibody was omitted and replaced with buffer. Sections from control (0 dpi; n = 4) and infected animals (60 dpe; n = 4) were used. Three pictures for each section were taken and analyzed.
The origins and sugar binding properties of the lectins used in this study.
Inhibitor b (mM)c
RCA 120 (1)
Lectin staining, subjectively described from ++ (very strong) to - (absent) during a primary infection with O.ostertagi.
Alteration of mucus biosynthesis during Ostertagia infection
Transcription profile of genes during a primary infection with O.ostertagi
2.07 ± 0.3*
2.93 ± 0.3*
3.29 ± 0.3*
2.75 ± 0.2*
2.78 ± 0.5*
1.50 ± 0.2*
3.99 ± 0.6*
3.28 ± 0.6*
Secreted gel forming
4.08 ± 1.1*
2.25 ± 0.5*
1.50 ± 0.2*
1.56 ± 0.7*
3.09 ± 1.0*
2.80 ± 0.4*
4.39 ± 1.4*
4.52 ± 0.5*
3.89 ± 1.5*
3.32 ± 0.8*
2.57 ± 0.2*
3.90 ± 0.5*
2.50 ± 1.1*
4.12 ± 2.4*
2.22 ± 0.8*
3.81 ± 1.4*
2.59 ± 0.8*
18.57 ± 11.7*
58.57 ± 13.2*
49.28 ± 16.7*
2.93 ± 0.6*
3.73 ± 0.3*
5.90 ± 1.0*
4.29 ± 0.6*
2.92 ± 0.3*
2.33 ± 0.2*
3.02 ± 0.2*
3.14 ± 0.3*
3.06 ± 0.6*
2.09 ± 0.2*
2.22 ± 0.5*
2.35 ± 0.2*
Backbone and terminal
2.87 ± 0.8*
1.60 ± 0.2
12.44 ± 4.2*
6.74 ± 1.0*
3.32 ± 0.3*
2.03 ± 0.3
3.12 ± 0.8*
2.75 ± 0.1*
4.51 ± 1.1*
3.11 ± 0.6*
2.57 ± 0.8*
4.61 ± 0.4*
1.14 ± 0.2*
2.10 ± 0.6*
4.52 ± 2.6*
2.35 ± 0.5*
1.60 ± 0.1*
1.20 ± 0.2*
1.89 ± 0.3*
1.15 ± 0.1*
1.94 ± 0.5*
1.27 ± 0.2*
0.77 ± 0.1*
1.33 ± 0.1*
2.35 ± .4*
2.56 ± 0.8*
3.13 ± 0.4*
3.15 ± 0.2*
2.77 ± 0.5*
2.89 ± 1.0*
3.09 ± 0.5*
3.13 ± 0.6*
1.65 ± 0.2*
1.47 ± 0.2*
1.46 ± 0.2*
1.58 ± 0.1*
3.41 ± 0.6*
3.44 ± 0.4*
5.82 ± 0.9*
5.24 ± 0.8*
3.16 ± 0.3*
2.60 ± 0.4*
4.12 ± 1.0*
4.46 ± 0.6*
2.58 ± 0.2*
3.51 ± 0.8*
6.95 ± 0.9*
5.37 ± 0.9*
Alteration of neutral and acidic mucins during Ostertagia infection
Immunohistochemistry for TFF3
Alteration in the mucin saccharide residues during Ostertagia infection
The importance of mucus as a host defensive mechanism against intestinal nematode infections has been reported in several animal species, including sheep, cattle and rodents [13, 15, 25, 26], but the role of mucus in the response to abomasal parasites remains largely unclear. Apart from the recently reported changes in TFF3, GCNT3 and gastrokine 2 transcription levels during an O. ostertagi infection , little is known about the effect of this parasite on abomasal mucus composition and synthesis.
In the current study, increased gene expression of mucins, which are normally present in the abomasum, was observed for MUC1, MUC6 and MUC20. Although the transcriptional upregulation started early during infection, the highest changes were found when adult worms were present on the surface of the abomasal mucosa, after emergence from the infected glands. Transcription level of MUC5AC, another mucin normally present in abomasal tissues, did not change during infection. Previous studies in humans have shown that the gastric mucus layer is able to react to invading pathogens, such as Helicobacter pylori, through the modification of the expression profile of MUC1, MUC5AC and MUC6, but the role of these changes still remains unknown [28–30].
Qualitative changes of mucins, related to sugar composition, were also observed. Histochemistry of neutral and acidic (sialylated and sulfated) mucins in uninfected animals was found to be similar to what has been described in previous studies in abomasum of sheep . Consistent with previous observations [19, 32], the abomasal mucosa was found to be thickened at 24 dpi and in animals exposed for 60 days to a natural infection, compared to uninfected control animals. The length of the fundic glands was increased, due to the hyperplasia of the mucus secreting epithelium, as shown by PAS staining. The hyperplasia of these mucus-secreting cells may be the reason for the detected increase in mucin gene transcription levels during the infection. At 14 dpi, AB-PAS and HID-AB stainings showed that the abomasal mucosa was characterized by depletion of both neutral and acidic mucins, confined to the area invaded by the parasite larvae, indicating a strictly localized response to the parasite. The limited modification observed at this stage of the infection may be an attempt of the parasite to create an optimal environment for itself through the production of enzymes that break down mucin protein and carbohydrate structures, as previously suggested . In the rest of the mucosa, during the infection, the alteration in mucin composition, in particular in the hyperplastic glands, was characterized by a decrease of neutral and an increase in acidic mucins produced by the MNCs in the neck region of the gland. These alterations in mucin composition are very similar to what has been described in sheep during a primary infection with H. contortus . Since the modification of the glycosylation status of mucins has been reported to relate with alteration in the viscoelastic properties of mucus [34, 35] and the possible attachment of the parasite [16, 31], these alterations may be an attempt of the host to eliminate the parasite.
Genes involved in mucin core structure synthesis and branching were observed to be altered in infected animals compared to controls. Biosynthesis of mucin O-linked glycan is a complicated process, which starts with the formation of N-acetylgalactosamine (GalNAC), followed by the synthesis of four main core structures (core 1, 2, 3 and 4) that can be branched to form a high variety of glycans. GCNT enzymes are involved in these processes and among the genes analyzed, GCNT3 and GCNT4 were found to be upregulated. Interestingly, in a previous study on cattle infected with C. oncophora, strong upregulation of GCNT3 in intestinal goblet cells and in columnar epithelial cell was noticed throughout the infection. As observed in the current study, the increased transcription level started early during the infection (6 dpi). GCNT3 catalyzes a key rate-limiting step in mucins biosynthesis. An early upregulation during nematode infections suggests an enhancement in mucin secretion and an early capacity of the host to respond to the presence of the parasite, before the major alterations and damages in the mucosa appear, independently from the species of the invading parasite. During infection with O. ostertagi some of the sugar residues of the mucins in the abomasal mucosa were also found to be altered. Staining with the UEA-I lectin, which binds to α-L-Fucosyl residues, was increased during infection, in particular in the secreted mucus and in SMCs, compared to control animals. This observation is in contrast with the results of Hoang et al.  where a reduced UEA binding was observed in the abomasal fundus during infection with T. circumcincta. Almost all fucose in mucus is found on mucins where it has an important effect on the viscosity of the mucus [34, 37]. The increased levels of fucose in the tissue during infection are consistent with the observed increase in transcription levels of FUT2 and FUT4, coding for enzymes transfering Fuc α-1,2 and Fuc α-1,3 respectively, on mucins. Similarly in the small intestine of rats infected with N. brasiliensis, an induced fucosylation, due to an upregulation of Fut2 gene expression, has been reported . Disulphide isomerases are a family of enzymes that play an important role in the process of disulphide bond formation of gel forming mucins [39, 40]. They are expressed in several tissues, including the stomach and the intestine where they are produced by mucus secreting cells [40, 41]. Increased transcription levels of 3 disulphide isomerases (AGR2, PDIA3 and PDIA4) were observed after an O. ostertagi infection. Since the polymerization of mucin monomers is crucial in the formation of viscoelastic mucus, an increase of disulphide isomerases level may increase the gastric mucus viscosity. Although further studies analyzing the rheological properties of mucus during infection need to be done, it is possible that all the modification observed in mucus may be related to an attempt of the host to eliminate the invading pathogens.
Trefoil factor peptides are normally synthesized and secreted in human gastric and intestinal mucosa . In humans, TFF1 is predominantly located in the foveolar cells of the gastric mucosa, TFF2 in the MNCs and deep in the pyloric glands, while TFF3, also called intestinal trefoil factor, is expressed mainly by goblet cells of the large and small intestine [43, 44]. TFF3 has recently also been localized in the human gastric cardia . In the current study, a strong upregulation of TFF3 was observed from 9 dpi onward. Immunofluorescence confirmed the increased expression of TFF3 in the mucosa of infected animals and showed that SMCs produce this peptide in the bovine fundus of uninfected calves, similar to what has been described in man . In infected animals the hyperplastic mucus secreting cells appear to be the ones producing TFF3. The TFF3 upregulation is consistent with a recent study of Li et al.  that have showed an increase of TFF3 mRNA levels in primary and repeated infections with O. ostertagi. TFF3 upregulation has also been observed during H. contortus and T. colubriformis infections in sheep . TFF1 was also observed to be upregulated at 24 dpi. To our knowledge this is the first time that TFF1 upregulation is observed during a GI nematode infection. In man, gastric pit cells (SMCs) in the cardia are able to synthesize TFF1, TFF3 and MUC5AC . It is possible that the hyperplastic TFF3-expressing cells in the abomasum of infected calves also produce TFF1. The potential role of TFF3 during nematode infections has been related to mucosal defense and tissue restitution [15, 27] but it still remains unknown if TFF1 may also contribute to tissue repair.
In conclusion, this study has shown that the abomasal mucin, TFFs and glycogenes transcription levels, as well as mucin glycosylation patterns are significantly altered during an Ostertagia infection in cattle. These changes are likely caused by the alterations in mucosal cell populations, characterized by hyperplasia of mucus secreting cells. The effect of these changes on the protective mucus barrier is still unclear. Studies on host immune response against O. ostertagi have highlighted that expulsion of adult worms during primary infection is uncommon [46, 47], therefore alteration in mucin sugar composition and mucus viscosity does not seem to be an efficient method to eliminate this parasite from the abomasum in this stage of the infection. Further studies comparing the mucosal changes in primary infected animals versus animals with an acquired immunity against O. ostertagi, will be helpful in clarifying if mucus has a protective role against this parasite.
This research was funded by a PhD grant of the 'Institute for the Promotion of Innovation through Science and Technology in Flanders' (IWT-Vlaanderen) (IWT-SB/61028/Hoorens), the 'Fund for Scientific Research Flanders' (F.W.O. 1.5.005.07) and Ghent University Concerted Research Actions.
- Hoorens PR, Rinaldi M, Li RW, Goddeeris B, Claerebout E, Vercruysse J, Geldhof P: Genome wide analysis of the bovine mucin genes and their gastrointestinal transcription profile. BMC Genomics. 2011, 12: 140-10.1186/1471-2164-12-140.PubMed CentralView ArticlePubMedGoogle Scholar
- Corfield AP, Carroll D, Myerscough N, Probert CS: Mucins in the gastrointestinal tract in health and disease. Front Biosci. 2001, 6: D1321-1357. 10.2741/Corfield.View ArticlePubMedGoogle Scholar
- Linden SK, Sutton P, Karlsson NG, Korolik V, McGuckin MA: Mucins in the mucosal barrier to infection. Mucosal Immunol. 2008, 1 (3): 183-197. 10.1038/mi.2008.5.View ArticlePubMedGoogle Scholar
- Jass JR, Walsh MD: Altered mucin expression in the gastrointestinal tract: a review. J Cell Mol Med. 2001, 5 (3): 327-351. 10.1111/j.1582-4934.2001.tb00169.x.View ArticlePubMedGoogle Scholar
- Hoffmann W: Trefoil factor family (TFF) peptides: regulators of mucosal regeneration and repair, and more. Peptides. 2004, 25 (5): 727-730. 10.1016/j.peptides.2004.03.019.View ArticlePubMedGoogle Scholar
- Hoffmann W, Jagla W, Wiede A: Molecular medicine of TFF-peptides: from gut to brain. Histol Histopathol. 2001, 16 (1): 319-334.PubMedGoogle Scholar
- Hoffmann W: Trefoil factors TFF (trefoil factor family) peptide-triggered signals promoting mucosal restitution. Cell Mol Life Sci. 2005, 62 (24): 2932-2938. 10.1007/s00018-005-5481-9.View ArticlePubMedGoogle Scholar
- Thim L, Madsen F, Poulsen SS: Effect of trefoil factors on the viscoelastic properties of mucus gels. Eur J Clin Invest. 2002, 32 (7): 519-527. 10.1046/j.1365-2362.2002.01014.x.View ArticlePubMedGoogle Scholar
- Hasnain SZ, Wang H, Ghia JE, Haq N, Deng Y, Velcich A, Grencis RK, Thornton DJ, Khan WI: Mucin gene deficiency in mice impairs host resistance to an enteric parasitic infection. Gastroenterology. 2010, 138 (5): 1763-1771. 10.1053/j.gastro.2010.01.045.PubMed CentralView ArticlePubMedGoogle Scholar
- Hasnain SZ, Thornton DJ, Grencis RK: Changes in the mucosal barrier during acute and chronic Trichuris muris infection. Parasite Immunol. 2011, 33 (1): 45-55. 10.1111/j.1365-3024.2010.01258.x.PubMed CentralView ArticlePubMedGoogle Scholar
- Rowe A, Gondro C, Emery D, Sangster N: Sequential microarray to identify timing of molecular responses to Haemonchus contortus infection in sheep. Vet Parasitol. 2009, 161 (1-2): 76-87. 10.1016/j.vetpar.2008.12.023.View ArticlePubMedGoogle Scholar
- Ingham A, Reverter A, Windon R, Hunt P, Menzies M: Gastrointestinal nematode challenge induces some conserved gene expression changes in the gut mucosa of genetically resistant sheep. Int J Parasitol. 2008, 38 (3-4): 431-442. 10.1016/j.ijpara.2007.07.012.View ArticlePubMedGoogle Scholar
- Li RW, Li C, Elsasser TH, Liu G, Garrett WM, Gasbarre LC: Mucin biosynthesis in the bovine goblet cell induced by Cooperia oncophora infection. Vet Parasitol. 2009, 165 (3-4): 281-289. 10.1016/j.vetpar.2009.07.008.View ArticlePubMedGoogle Scholar
- Ishikawa N, Horii Y, Nawa Y: Immune-mediated alteration of the terminal sugars of goblet cell mucins in the small intestine of Nippostrongylus brasiliensis-infected rats. Immunology. 1993, 78 (2): 303-307.PubMed CentralPubMedGoogle Scholar
- Menzies M, Reverter A, Andronicos N, Hunt P, Windon R, Ingham A: Nematode challenge induces differential expression of oxidant, antioxidant and mucous genes down the longitudinal axis of the sheep gut. Parasite Immunol. 2010, 32 (1): 36-46. 10.1111/j.1365-3024.2009.01156.x.View ArticlePubMedGoogle Scholar
- Yamauchi J, Kawai Y, Yamada M, Uchikawa R, Tegoshi T, Arizono N: Altered expression of goblet cell- and mucin glycosylation-related genes in the intestinal epithelium during infection with the nematode Nippostrongylus brasiliensis in rat. APMIS. 2006, 114 (4): 270-278. 10.1111/j.1600-0463.2006.apm_353.x.View ArticlePubMedGoogle Scholar
- Fox MT: Pathophysiology of infection with gastrointestinal nematodes in domestic ruminants: recent developments. Vet Parasitol. 1997, 72 (3-4): 285-297. 10.1016/S0304-4017(97)00102-7. discussion 297-308View ArticlePubMedGoogle Scholar
- Murray M, Jennings FW, Armour J: Bovine ostertagiasis: structure, function and mode of differentiation of the bovine gastric mucosa and kinetics of the worm loss. Res Vet Sci. 1970, 11 (5): 417-427.PubMedGoogle Scholar
- Ross JG, Dow C: The course and development of the abomasal lesions in calves experimentally infected with the nematode parasite Ostertagia ostertagi. Brit Vet J. 1965, 121: 228-233.Google Scholar
- Snider TG, Williams JC, Knox JW, Marbury KS, Crowder CH, Willis ER: Sequential histopathologic changes of type I, pre-type II and type II ostertagiasis in cattle. Vet Parasitol. 1988, 27 (1-2): 169-179. 10.1016/0304-4017(88)90072-6.View ArticlePubMedGoogle Scholar
- McManus JF: Histological demonstration of mucin after periodic acid. Nature. 1946, 158: 202-View ArticlePubMedGoogle Scholar
- Mowry RW: Alcian blue techniques for the histochemical study of acidic carbohydrates. J Histochem Cytochem. 1956, 4: 407-Google Scholar
- Mowry RW: Observations on the use of sulfuric acid in ether for the sulfation of hydroxyl groups in tissue sections. J Histochem Cytochem. 1958, 6 (2): 82-83.PubMedGoogle Scholar
- Spicer SS: Diamine Methods for Differentialing Mucosubstances Histochemically. J Histochem Cytochem. 1965, 13: 211-234. 10.1177/13.3.211.View ArticlePubMedGoogle Scholar
- Hoang VC, Williams MA, Simpson HV: Monosaccharide composition of fundic and duodenal mucins in sheep infected with Haemonchus contortus or Teladorsagia circumcincta. Vet Parasitol. 2010, 170 (3-4): 253-261. 10.1016/j.vetpar.2010.02.014.View ArticlePubMedGoogle Scholar
- Takeda K, Hashimoto K, Uchikawa R, Tegoshi T, Yamada M, Arizono N: Direct effects of IL-4/IL-13 and the nematode Nippostrongylus brasiliensis on intestinal epithelial cells in vitro. Parasite Immunol. 2010, 32 (6): 420-429. 10.1111/j.1365-3024.2010.01200.x.View ArticlePubMedGoogle Scholar
- Li RW, Hou Y, Li C, Gasbarre LC: Localized complement activation in the development of protective immunity against Ostertagia ostertagi infections in cattle. Vet Parasitol. 2010, 174 (3-4): 247-256. 10.1016/j.vetpar.2010.08.037.View ArticlePubMedGoogle Scholar
- Byrd JC, Yunker CK, Xu QS, Sternberg LR, Bresalier RS: Inhibition of gastric mucin synthesis by Helicobacter pylori. Gastroenterology. 2000, 118 (6): 1072-1079. 10.1016/S0016-5085(00)70360-X.View ArticlePubMedGoogle Scholar
- Kang HM, Kim N, Park YS, Hwang JH, Kim JW, Jeong SH, Lee DH, Lee HS, Jung HC, Song IS: Effects of Helicobacter pylori Infection on gastric mucin expression. J Clin Gastroenterol. 2008, 42 (1): 29-35. 10.1097/MCG.0b013e3180653cb7.View ArticlePubMedGoogle Scholar
- Matsuzwa M, Ota H, Hayama M, Zhang MX, Sano K, Honda T, Ueno I, Akamatsu T, Nakayama J: Helicobacter pylori infection up-regulates gland mucous cell-type mucins in gastric pyloric mucosa. Helicobacter. 2003, 8 (6): 594-600. 10.1111/j.1523-5378.2003.00185.x.View ArticlePubMedGoogle Scholar
- Newlands GF, Miller HR, Jackson F: Immune exclusion of Haemonchus contortus larvae in the sheep: effects on gastric mucin of immunization, larval challenge and treatment with dexamethasone. J Comp Pathol. 1990, 102 (4): 433-442. 10.1016/S0021-9975(08)80164-8.View ArticlePubMedGoogle Scholar
- Martin WB, Thomas BAC, Urquhart GM: Chronic diarreha in housed cattle due to atypical parasitic gastritis. Vet Rec. 1957, 736-739.Google Scholar
- Young CJ, McKeand JB, Knox DP: Proteinases released in vitro by the parasitic stages of Teladorsagia circumcincta, an ovine abomasal nematode. Parasitology. 1995, 110 (Pt 4): 465-471.View ArticlePubMedGoogle Scholar
- Ishibashi Y, Takayama G, Inouye Y, Taniguchi A: Carbocisteine normalizes the viscous property of mucus through regulation of fucosylated and sialylated sugar chain on airway mucins. Eur J Pharmacol. 2010, 641 (2-3): 226-228. 10.1016/j.ejphar.2010.05.045.View ArticlePubMedGoogle Scholar
- Wiggins R, Hicks SJ, Soothill PW, Millar MR, Corfield AP: Mucinases and sialidases: their role in the pathogenesis of sexually transmitted infections in the female genital tract. Sex Transm Infect. 2001, 77 (6): 402-408. 10.1136/sti.77.6.402.PubMed CentralView ArticlePubMedGoogle Scholar
- Hoang VC, Williams MA, Simpson HV: Effects of weaning and infection with Teladorsagia circumcincta on mucin carbohydrate profiles of early weaned lambs. Vet Parasitol. 2010, 171 (3-4): 354-360. 10.1016/j.vetpar.2010.04.007.View ArticlePubMedGoogle Scholar
- Girod S, Zahm JM, Plotkowski C, Beck G, Puchelle E: Role of the physiochemical properties of mucus in the protection of the respiratory epithelium. Eur Respir J. 1992, 5 (4): 477-487.PubMedGoogle Scholar
- Holmen JM, Olson FJ, Karlsson H, Hansson GC: Two glycosylation alterations of mouse intestinal mucins due to infection caused by the parasite Nippostrongylus brasiliensis. Glycoconj J. 2002, 19 (1): 67-75. 10.1023/A:1022589015687.View ArticlePubMedGoogle Scholar
- Ellgaard L, Ruddock LW: The human protein disulphide isomerase family: substrate interactions and functional properties. EMBO Rep. 2005, 6 (1): 28-32. 10.1038/sj.embor.7400311.PubMed CentralView ArticlePubMedGoogle Scholar
- Park SW, Zhen G, Verhaeghe C, Nakagami Y, Nguyenvu LT, Barczak AJ, Killeen N, Erle DJ: The protein disulfide isomerase AGR2 is essential for production of intestinal mucus. Proc Natl Acad Sci USA. 2009, 106 (17): 6950-6955. 10.1073/pnas.0808722106.PubMed CentralView ArticlePubMedGoogle Scholar
- Iida KI, Miyaishi O, Iwata Y, Kozaki KI, Matsuyama M, Saga S: Distinct distribution of protein disulfide isomerase family proteins in rat tissues. J Histochem Cytochem. 1996, 44 (7): 751-759. 10.1177/44.7.8675996.View ArticlePubMedGoogle Scholar
- Kouznetsova I, Peitz U, Vieth M, Meyer F, Vestergaard EM, Malfertheiner P, Roessner A, Lippert H, Hoffmann W: A gradient of TFF3 (trefoil factor family 3) peptide synthesis within the normal human gastric mucosa. Cell Tissue Res. 2004, 316 (2): 155-165. 10.1007/s00441-004-0854-1.View ArticlePubMedGoogle Scholar
- Podolsky DK, Lynch-Devaney K, Stow JL, Oates P, Murgue B, DeBeaumont M, Sands BE, Mahida YR: Identification of human intestinal trefoil factor. Goblet cell-specific expression of a peptide targeted for apical secretion. J Biol Chem. 1993, 268 (9): 6694-6702.PubMedGoogle Scholar
- Wong WM, Poulsom R, Wright NA: Trefoil peptides. Gut. 1999, 44 (6): 890-895. 10.1136/gut.44.6.890.PubMed CentralView ArticlePubMedGoogle Scholar
- Kouznetsova I, Kalinski T, Peitz U, Monkemuller KE, Kalbacher H, Vieth M, Meyer F, Roessner A, Malfertheiner P, Lippert H, Hoffmann W: Localization of TFF3 peptide in human esophageal submucosal glands and gastric cardia: differentiation of two types of gastric pit cells along the rostro-caudal axis. Cell Tissue Res. 2007, 328 (2): 365-374. 10.1007/s00441-006-0350-x.View ArticlePubMedGoogle Scholar
- Gasbarre LC: Effects of gastrointestinal nematode infection on the ruminant immune system. Vet Parasitol. 1997, 72 (3-4): 327-337. 10.1016/S0304-4017(97)00104-0. discussion 337-343View ArticlePubMedGoogle Scholar
- Michel JF, Lancaster MB, Hong C: Ostertagia ostertagi: protective immunity in calves. The development in calves of a protective immunity to infection with Ostertagia ostertagi. Exp Parasitol. 1973, 33 (1): 179-186. 10.1016/0014-4894(73)90023-4.View ArticlePubMedGoogle Scholar
This article is published under license to BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.