![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
The Faculty of Agricultural, Food and Environmental Quality Sciences, Hebrew University of Jerusalem, Rehovot, 76100, Israel
1To whom correspondence should be addressed. E-mail: sklan{at}agri.huji.ac.il.
 |
ABSTRACT |
|---|
 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
KEY WORDS: • chick • mucin • small intestine • starvation • mucus adherent layer
The epithelium of the intestinal tract is covered by a layer of mucus composed predominantly of mucin glycoproteins that are synthesized and secreted by goblet cells. The mucus layer acts as a medium for protection, lubrication, and transport between the luminal contents and the epithelial cells (1).
On the basis of recently deduced amino acid sequences, mucins are now categorized in humans into three distinct families according to the structure of the protein product: gel-forming (MUC2, MUC5AC, MUC5B and MUC6), soluble (MUC7), or membrane-bound (MUC1, MUC3, MUC4 and MUC12) (2). Secretory gel-forming mucins are organized in domains with similar repetitive tandem domains with potential O-glycosylated sites (9–17). The nontandem repetitive region of MUC2, termed D-domains, have a high cysteine content (3,4). These regions exhibit a high level of sequence homology and are positioned similarly in their amino-terminal region to the serum polymeric glycoprotein protein pro-von-Willebrand factor (3,4), which is important in the mucin multimerization process (5). Mucin oligosaccharide chains often terminate with sialic acid or sulfate groups, which accounts for the polyanionic nature of mucins at neutral pH (1). Diversity in the length, composition, branching, and degree of sulfation and acetylation of the oligosaccharide chains attached to the peptide backbone leads to heterogeneity of mucin glycoproteins (6).
The expression of different mucins, defined by differences either in their protein backbones or in their glycosylation patterns, varies both between and within tissues (6,7). Mucin genes are regulated at transcriptional levels by cytokines (8), bacterial products, and growth factors (9). Mucin biosynthesis is also affected by conditions or agents that affect differentiation of precursor cells into mature goblet cells and agents or conditions that uncouple the processes of glycosylation and protein synthesis, or that influence protein synthesis generally such as fasting or malnutrition (9–14).
Mucus is secreted by goblet cells throughout the gastrointestinal tract and forms a gel adherent to the mucosal surface (15). This layer acts as a barrier between the luminal contents and the absorptive system of the intestine and protects the mucosal surface from exogenous or endogenous luminal irritants such as laxatives (16) or bile salts (17). Changes in the properties of this barrier could affect the absorption of both dietary and endogenous macromolecules and ions (18,19).
Little is known about the mechanisms controlling mucin synthesis, secretion, and adherent layer turnover and the response to dietary changes; clarifying these aspects of mucin dynamics constituted the objective of the present study in chickens.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
Male Cobb 28-d-old chicks were blocked into experimental groups on the basis of body weight, equalizing body weight and variance between groups. The control group had free access to water and to a commercial diet (Matmor Feed Mill) (20) formulated to meet or exceed NRC recommendations (21). The starved group had no access to food or water for 72 h. Each dietary treatment was applied to groups of >4 chicks. All procedures were approved by the Animal Care and Welfare Committee of our Institute. At the end of the starvation period, chicks were killed and intestinal segments were removed and gently flushed with 150 mmol/L NaCl to remove the intestinal contents. Samples (
2 cm) were taken from the midpoint of the duodenum, from the midpoint between the point of entry of the bile duct and Meckel’s diverticulum (jejunum), and midway between Meckel’s diverticulum and the ileocecal junction (ileum). Samples for mRNA and protein determination were frozen in liquid N.
Total RNA isolation.
Total RNA was isolated (22) from the intestinal segments using TRI reagent (10 mL/g tissue) according to the manufacturer’s protocol (MRC Molecular Research Center).
Isolation of a chicken intestinal mucin gene fragment.
Four different published sequences of the intestinal mucin genes from different sources were compared, i.e., Homo sapiens intestinal/tracheal mucin MUC2 (GI 4505284), Homo sapiens mucin MUC 5AC (GI 3123851), Rattus norvegicus mucin MUC2 (GI 506641) and Mus musculus mucin MUC2 (GI 5532222); two primers, chosen from conserved regions were designed: 5'-GTC TGC AGC ACC TGG GGC AAC TTC C-3' (forward); 5'-GCT CTG CAG GCC GTT GTA GTC CCC G-3' (reverse), which correspond to nucleotides 103–127 and 504–528 of Rat MUC2. Total RNA was amplified using the Promega Access RT-PCR System (Promega Corporation). The program was as follows: 2 min at 94°C, 30 s at 60°C, 2 min at 68°C for 30 cycles followed by 7 min at 68°C. The RT-PCR products were examined on a 1.5% agarose gel, visualized by staining with ethidium bromide, excised from the gel and purified from a gel with High Pure PCR Product Purification kit (Roche Diagnostics GmbH). The chicken intestinal mucin cDNA fragment was subjected to automated sequencing using an Applied Biosystem 373A DNA sequencer. Nucleic acid sequences were analyzed and homology between chicken and other mucin sequences was calculated using DNAMAN version 4 (Lynnon Biosoft).
mRNA analysis.
First strand cDNAs were synthesized from 5 µg of total RNA of each chick using oligo(dT)18 as primers in the presence of MLV reverse transcriptase (Fermentas), for 1 h at 42°C. cDNA were purified from the PCR mix by using High Pure PCR Product Purification kit (Roche Diagnostics GmbH). PCR was carried out with primers chosen from the fragment of the chicken intestinal mucin gene (forward: 5'-TCT TCC GCT ACC CTG GGC TCT GTA A-3'; reverse: 5'-CTC ATG CAG TTC TAG CAA GAT ACT-3') and the housekeeping gene, ß-actin, with primers from the Gallus gallus cytoplasmic ß-actin mRNA (GI 211236) (forward: 5'-CCC TGT ATG CCT CTG GTC GT-3', reverse: 5'-ATC TCG TCT TGT TTT ATG CG-3'). Determination of the linear phase of the amplification PCR was performed with Pfu DNA polymerase (Promega) with pooled cDNA aliquots removed at 10, 15, 20, 25, 30, 35, 40, and 45 cycles. Amplification of the chicken intestinal mucin gene was performed for 41 cycles, which consisted of denaturation (95°C, 30 s), annealing (54°C, 1 min), and extension (72°C, 1 min); ß-actin was amplified at 23 cycles under the same conditions in a different tube. ß-Actin (760 bp) and chicken intestinal mucin (317 bp) PCR products were separated by electrophoresis on 1.5% agarose gel, stained with ethidium bromide, and quantified using a Gel-Pro Analayzer version 3.0 (Media Cybernetics, LP).
Western blot analysis.
Intestinal tissues were homogenized in lysis buffer 10 mmol/L Tris, pH 7.4, containing 150 mmol/L NaCl, 50 mmol/L EDTA, 1% (v:v) Triton, 5 mmol/L dithiothreitol, 4 mmol/L phenylmethylsulfonyl fluoride, and 10 mmol/L NaF, with a tissue:buffer ratio of 1:10 (wt:v) and centrifuged at 12,000 x g at 4°C. Protein concentration in the supernatant was determined by DC Protein Assay (Bio-Rad Laboratories), with bovine serum albumin (BSA) as a standard. Samples were applied to a Sephadex G-150–120 (Sigma Chemical) column and the void volume fraction was collected and subjected to electrophoresis on SDS-polyacrylamide gels [4% (wt:v) acrylamide in the stacking and 5% (wt:v) acrylamide in the running gel]. Gels were subsequently transferred onto nitrocellulose (Schleicher and Schuell). Detection of the mucin glycoprotein was performed after blocking the membrane with 20 g/L BSA in 50 mmol/L Tris-buffered saline, pH 7.4, with 0.05% (v:v) Tween for 3 h. The primary antibody, MUC5AC (Zymed Laboratories) was diluted 1:1000 in 50 mmol/L Tris buffered saline, pH 7.4, with 3 g/L BSA and incubated with the nitrocellulose membrane overnight at 4°C. The membrane was washed and incubated with peroxidase-conjugated donkey anti-mouse IgG (H+L) antibody (Jackson ImmunoResearch Laboratories) for 2 h at room temperature. Immunoblots were developed with Western blotting luminol reagent (Santa Cruz Biotechnology) as recommended by the manufacturer. The density of the positive bands was quantified using a Gel-Pro Analayzer version 3.0 (Media Cybernetics).
Measurement of the mucus adherent layer thickness.
The thickness of the mucus adherent layer was estimated by a modification of Corne’s method (23–25). Briefly, 1-cm pieces of intestinal tissue were removed and placed in 10 g/L Alcian blue dye solution in buffer containing 160 mmol/L sucrose and 50 mmol/L sodium acetate, pH 5.8. After 2 h of incubation, excess dye was extracted with 250 mmol/L sucrose. The absorbed dye was extracted from the tissue by incubation in 10 g/L docusate sodium salt solution overnight at room temperature. Samples were cleared by centrifugation at 700 x g and optical density was measured at 620 nm using Alcian Blue solution as a standard. The amount of absorbed dye is reported as µg Alcian Blue/cm2 of intestinal tissue.
Morphological examination.
Intestinal segments were fixed in 4% (v:v) buffered formaldehyde, dehydrated, cleared, and embedded in paraffin. Serial sections were cut at 3 µm, deparaffinized in xylene, rehydrated, and stained with hematoxylin and eosin. Sections were examined by light microscopy.
Mucin staining.
Determination of neutral mucin was by staining 5-µm sections with periodic acid-Schiff reagent (PAS) (26,27). After deparaffinization and rehydration, slides were incubated in 5 g/L periodic acid for 15 min, then washed and incubated with Schiff’s reagent (Sigma Chemical) for 30 min. After being washed in warm water, slides were dehydrated and mounted. The number of PAS-positive cells along the villi was determined by light microscopy. Determination of acid mucin was by staining 5-µm sections with Alcian Blue pH 2.5 (27,28). After deparaffinization and rehydration, slides were incubated in 0.5 mol/L acetic acid for 3 min and then in Alcian Blue solution (10 g/L in 0.5mol/L acetic acid pH 2.5). After being washed in water, the slides were dehydrated and mounted. The number of Alcian Blue positive cells along the villi was counted by light microscopy.
Morphometric measurements.
The area of the goblet cell was calculated from the length and width of goblet cell "cup" in cross sections of the villi. The long diameter of the cup was defined as the distance between the luminal opening and the site of constriction of the cell, and the short diameter was determined at the mid-point of the long diameter and the elliptic area calculated. The density of goblet cells was calculated as the number of goblet cells per unit of surface area (mm2). All measurements were performed with an Olympus light microscope using EPIX XCAP software.
Statistical analysis.
Data were analyzed by 2-way ANOVA with intestinal segment and treatment (control or starved) as main effects using the General Linear Models procedures of SAS (29). Differences between means were tested using Tukey’s test. Differences were considered significant at P < 0.05 unless otherwise stated.
|
RESULTS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
A 426-bp fragment of the chicken intestinal mucin gene isolated by RT-PCR was sequenced and was 60% homologous to Homo sapiens MUC5AC, 48% to Rattus norvegicus MUC2, and 47% to Homo sapiens MUC2 genes. The cDNA sequence of the chicken intestinal mucin was entered into the EMBL nucleotide Sequence Database under accession number AJ487010. The predicted amino acid sequence of this fragment resulted in a predicted translation product of 123 amino acids (Fig. 1). This amino acid sequence was 67% homologous to Homo sapiens MUC5AC, 54% homologous to Homo sapiens MUC2, and 49% homologous to Rattus norvegicus MUC2. Searching the conserved domain database resulted in identification of the von Willebrand factor type D domain in this fragment of the chicken intestinal mucin protein.
|
Chicken intestinal mucin exhibited a band with a molecular weight of >250 kDa after acrylamide gel electrophoresis. This band reacted with anti-human MUC5AC monoclonal antibody, but no reaction was observed with a similar molecular weight fraction of proteins prepared from muscle and liver tissue (Fig. 2). The highest concentrations of the chicken intestinal mucin glycoprotein were found in the proventriculus, and along the small intestine, concentrations increased distally (data not shown).
|
The thickness of the mucus adherent layer ranged from 68 to 76 µg Alcian Blue/cm2 of intestinal tissue. The thickness of the mucus adherent layer did not differ among the small intestinal segments.
Effect of starvation.
Food deprivation for 72 h decreased body weight by 30% (data not shown, P < 0.01) ; in the small intestine, the villous surface area was decreased 27% in the duodenum, 36% in the jejunum, and 26% in the ileum (P < 0.05 for all segments) (Fig. 3A). Goblet cells as a percentage of the total cells (Fig. 3B) and goblet cell density (Fig. 3C) were unaffected by starvation. Starvation did not affect the proportion of goblet cells stained for acidic and neutral mucins (data not shown). However, the size of the goblet cells in the starved chicks was dramatically greater than that in the control group (Fig. 4A). The area of the goblet cell "cup" containing the mucin granules was increased by starvation in all intestinal segments: by 33% in the duodenum, 100% in the jejunum, and 66% in the ileum (Fig. 4B, P < 0.05 in all segments). The expression of chicken intestinal mucin mRNA was examined by semiquantitative, two-step RT-PCR. Expression of chicken intestinal mucin mRNA was increased 47% by starvation in the duodenum, 480% in the jejunum, and was not affected in the ileum (Fig. 5). Starvation caused a 400% increase in total mucin glycoprotein concentration in the duodenum and a 216% increase in the jejunum, with no effect in the ileum (Fig. 6). The increase in mucin glycoprotein concentration in starved chicks did not result in a thicker mucus adherent layer. On the contrary, this layer was reduced 22% in duodenum, 26% in jejunum, and 45% in ileum of the starved chicks (Fig. 7).
|
|
|
|
|
|
DISCUSSION |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
A fragment of the chicken intestinal mucin gene was isolated and sequenced in this study. The predicted amino acid sequence of the gene fragment isolated had the highest homology (67%) to the human intestinal/tracheal mucin gene MUC 5AC (30) and lower homology to rat intestinal mucin MUC2 and human mucin MUC2 genes (4,31). The differences between chicken intestinal mucin and mammalian mucin genes might be due to evolutionary divergence. However, despite the differences in homology, the chicken mucin protein included the location of the predicted domain that contained a sequence corresponding to the sequence of the von Willebrand factor type D domain. This domain is required for the multimerization process (5); thus the chicken mucin protein apparently maintains the polymerizing ability.
The capacity for synthesis of mucin was evaluated in this study by quantifying mucin mRNA expression and the amount of mucin determined by measuring its concentration. To separate mucin glycoproteins, high-molecular-weight proteins were separated by gel filtration and a single band was found by SDS-PAGE electrophoresis with a MW > 250 kDa. Western blot analysis of chicken intestinal mucin glycoprotein revealed that mucin protein content increased distally along the small intestine. A similar pattern of increasing mucin concentration along the small intestine was described in other species (31–33).
Once mucin is synthesized in the goblet cells and secreted to the intestinal surface, it forms a layer that undergoes continuous degradation and renewal (1). This appears to constitute a diffusion barrier between luminal nutrients and digestive and transport sites on the brush border membrane of the enterocytes (34). Thus, changes in the thickness of the mucus adherent layer would be expected to influence nutrient digestion processes. The thickness of the adherent gastrointestinal mucus layer is the result of the balance between the rate of secretion of mucin and its degradation through enzymatic digestion and mechanical shear (35). The mucus adherent layer was of similar thickness throughout the chicken small intestine. In a previous study in rats, the thickness of the mucus adherent layer was similar in the duodenum and jejunum, and thicker in the ileum (36). In chicks, the mucin protective properties in the small intestine may not differ among intestinal segments.
Mucins, which form a surface network above the villi, are comprised of both neutral and acidic molecules. The ability of the mucus layer to protect the epithelium from different pathogens and its participation in nutrient absorption are often attributed to the presence of charged groups in the mucin molecules (47,48). Some nutritional manipulations in chicks, such as feeding maize-based diets and xylanase supplements (13), were found to change the profile of mucin types in the small intestine. In this study, starvation, a simple nutritional manipulation, was used to examine processes of mucin dynamics. In 4-wk-old chicks, the proportion of goblet cells (of total epithelial cells) and the proportion of goblet cells containing acidic and neutral mucins were unaffected by starvation. However, starvation decreased body weight, intestinal villous surface area, and caused a thinner mucus adherent layer. The presence of nutrients in the intestine is important for maintenance of normal mucosal function. It was shown that in White Leghorn hens starved for 36 h, starvation decreased villous height throughout the small intestine (37). Sakamoto et al. (38) showed villous atrophy and a reduction in the absolute quantity of mucus gel in rat ileum associated with total parenteral nutrition. It should be noted that in this study, chicks were deprived of both feed and water, making it impossible to separate the effects of these two variables.
The most remarkable effect of starvation was the enlargement of the goblet cells in the small intestine. A similar increase in the size of goblet cells was observed in hatchlings with delayed access to feed after hatch (39). This accumulation of mucin in goblet cells might be explained by changes in the interrelated processes of mucin synthesis, secretion, and degradation. Mucin synthesis depends on mRNA expression, followed by translation. Starvation increased the amounts of both mRNA and mucin glycoprotein in two of the three small intestinal segments examined. This phenomenon of enhanced mRNA expression after starvation was described in previous studies that showed upregulation of intestinal transporters such as sodium glucose transporter-1 in chickens, (20) and PepT1 in rats (40), and enzymes such as aminopeptidase in chickens and rats (20,41). This may be explained by enhanced translation to provide digestive ability for "upcoming" nutrients. Another possibility is suggested by recent studies in rat glioma cells, which showed that increased cationic amino acid transporter protein concentration during amino acid deprivation is due to both enhanced translation of mRNA and increased mRNA levels due to increased mRNA stability (42). Yaman et al. (42) proposed that increased mRNA stability is an adaptive response of cells to nutritional deprivation, mediated by "Human R" embryonic lethal abnormal visual-like RNA binding protein, a member of the embryonic lethal abnormal vision family of RNA binding proteins that protects mRNAs from degradation (43). An additional explanation may be that mRNA stability was influenced by the eukaryotic initiation factor 5A, which affects turnover of mRNA when protein synthesis decreases during nutrient deprivation (44).
The thickness of the mucus adherent layer is affected by the rate of mucin secretion and by the rate of mucin layer degradation; in this study, starvation caused the thickness of the mucin layer to decrease, whereas mucin protein concentrations increased. Because regulation of mucus secretion in the gastrointestinal tract is cholinergic, acetyl- and butyrylcholinesterase, which are responsible for the breakdown of acetylcholine after release, may be important (45). Several studies indicated that starvation influences cholinesterase activity. Leparoux et al. (46) reported increased acetylcholinesterase-specific activity in the jejunum of starved rats and Shambaugh et al. (47) found greater acetylcholinesterase activity in brain cells of rat fetuses from starved dams. In this study in chicks, starvation did not change the composition of goblet cell mucin as indicated by the staining used here. However, a mucus layer of decreased thickness in starved chicks may decrease the barrier properties of this layer, which could lead to increased exposure of the epithelium to luminal harmful agents.
The mucin layer at the mucosal surface is a nutrient source for many intestinal organisms; thus, some of the diverse bacterial species may utilize polysaccharide derived from mucin (48). It was shown that mucolytic bacteria have a competitive advantage during nutrient deprivation such as during malnutrition or total parenteral nutrition (49). Deplancke et al. (50) showed that the lack of enteral nutrients in TPN might encourage commensal or pathogenic bacteria, leading to increased mucus-associated bacteria using mucus as a substrate; for example, Clostridium perfringens, an opportunistic pathogen, was specifically enriched in the total parenteral nutrition. Thus, changes in intestinal microflora profile might contribute to the decrease of the mucus adherent layer during starvation; however, this requires further investigation.
This study described methods for analysis of chicken mucin dynamics and indicated that starvation enhances synthesis of the mucus glycoproteins and appears to perturb processes of secretion and degradation, which lead to the accumulation of mucin within the goblet cells.
Manuscript received 18 November 2003. Initial review completed 16 December 2003. Revision accepted 21 January 2004.
|
LITERATURE CITED |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
1. Forstner, J. F., Oliver, M. G. & Sylvester, F. A. (1995) Production, structure and biologic relevance of gastrointestinal mucins. Blaser, M. J. Smith, P. D. Ravdin, J. I. Greenberg, H. B. Guerrant, R. L. eds. Infections of the Gastrointestinal Tract 1995:71-88 Raven Press New York, NY. .
2. Moniax, N., Escande, F., Porchet, N., Aubert, J.-P. & Batra, S. K. (2001) Structural organization and classification of the human mucin genes. Front. Biosci. 6:D1192-D1206.[Medline]
3. Gum, J. R., Hicks, J. W., Toribara, N. W., Rothe, E. M., Lagace, R. E. & Kim, Y. S. (1992) The human MUC2 intestinal mucin has cysteine-rich domains located both upstream and downstream of its central repetitive region. J. Biol. Chem. 267:21375-21383.
4. Gum, J. R., Hicks, J. W., Toribara, N. W., Siddiki, B. & Kim, Y. S. (1994) Molecular cloning of human intestinal mucin (MUC2) cDNA. J. Biol. Chem. 269:2440-2446.
5. Titani, K. & Walsh, K. A. (1988) Human von Willebrand factor: the molecular glue of platelet plugs. Trends Biochem. Sci. 13:94-97.[Medline]
6. Neutra, M. R. & Forstner, J. F. (1987) Gastrointestinal mucus: synthesis, secretion and function. Johnson, L. R. eds. Physiology of the Gastrointestinal Tract 2nd ed. 1987:975-1009 Raven Press New York, NY. .
7. Gendler, S. J. & Spicer, A. P. (1995) Epithelial mucin genes. Annu. Rev. Physiol. 57:607-634.[Medline]
8. Temann, U. A., Prasad, B., Gallup, M. W., Basbaum, C., Ho, S. B., Flavell, R. A. & Rankin, J. A. (1997) A novel role for murine IL-4 in vivo: induction of MUC5AC gene expression and mucus hypersecretion. Am. J. Respir. Cell Mol. Biol. 16:471-478.[Abstract]
9. Arnold, J. W., Klimpel, G. R. & Niesel, D. W. (1993) Tumor necrosis factor (TNF
) regulates intestinal mucus production during salmonellosis. Cell. Immunol. 151:336-344.[Medline]
10. De Ritis, G., Falchuk, Z. M. & Trier, J. S. (1975) Differentiation and maturation of cultured fetal rat jejunum. Dev. Biol. 45:304-317.[Medline]
11. O’Doherty, P. J. A. & Kuksis, A. (1975) Effect of puromycin in vitro on protein and glycerolipid biosynthesis in isolated epithelial cells of rat intestine. Int. J. Biochem. 6:435-441.
12. Sherman, P., Forstner, J. F., Roomi, N., Kharti, I. & Forstner, G. G. (1985) Mucin depletion in the intestine of malnourished rats. Am. J. Physiol. 248:G418-G423.
13. Sharma, R., Fernandez, F., Hinton, M. & Schumacher, U. (1997) The influence of diet on the mucin carbohydrates in the chick intestinal tract. Cell. Mol. Life Sci. 53:935-942.[Medline]
14. Langhout, D. J., Schutte, J. B., Van Leeuwen, P. V., Wiebenga, J. & Tamminga, S. (1999) Effect of dietary high- and low-methylated citrus pectin on the activity of the ileal microflora and morphology of the small intestinal wall of broiler chicks. Br. Poult. Sci. 40:340-347.[Medline]
15. Forstner, J. F. & Forstner, G. G. (1994) Gastrointestinal mucus. Johnson Leonard, R. eds. Physiology of the Gastrointestinal Tract 3rd ed. 1994:1255-1284 Raven Press New York, NY. .
16. Yagi, T., Miyawaki, Y., Nishikawa, A., Horiyama, S., Yamuchi, K. & Kuwano, S. (1990) Prostaglandin E2-mediated stimulation of mucus synthesis and secretion by rhenin anthone, the active metabolite of sennosides A and B in mouse colon. J. Pharm. Pharmacol. 42:542-545.[Medline]
17. Teem, M. V. & Phillips, T. E. (1972) Perfusion of the hamster jejunum with conjugated and unconjugated bile acids: inhibition of water absorption and effects on morphology. Gastroenterology 62:261-267.[Medline]
18. Satchithanandam, S., Vargofcak-Apker, M., Calvert, R. J., Leeds, A. R. & Cassidy, M. M. (1990) Alteration of gastrointestinal mucin by fiber feeding in rats. J. Nutr. 120:1179-1184.
19. Quarterman, J. (1987) Metal absorption and the intestinal mucus layer. Digestion 37:1-9.
20. Gal-Garber, O., Mabjeesh, S. J., Sklan, D. & Uni, Z. (2000) Partial sequence and expression of the gene for and activity of the sodium glucose transporter in the small intestine of fed, starved and refed chickens. J. Nutr. 130:2174-2179.
21. National Research Council (1994) Nutrient Requirements of Poultry 9th ed. 1994 National Academy of Science Washington, DC.
22. Chomczynski, P. & Sacchi, N. (1987) Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 162:156-159.[Medline]
23. Corne, S. J., Morrissey, S. M. & Woods, R. J. (1974) Proceedings: a method for the quantitative estimation of gastric barrier mucus. J. Physiol. 242:116P-117P.
24. Kitagawa, H., Takeda, F. & Kohei, H. (1986) A simple method for estimation of gastric mucus and effects of antiulcerogenic agents on the decrease in mucus during water-immersion stress in rats. Arzneim.-Forsch. 36:1240-1244.[Medline]
25. Parmar, N. S. & Desai, J. K. (1993) A review of the current methodology for the evaluation of gastric and duodenal anti-ulcer agents. Indian J. Pharmacol. 25:120-135.
26. McManus, J.F.A. (1948) Histological and histochemical uses of periodic acid. Stain Technol. 23:99.
27. American Forces Institute of Pathology (1992) Laboratory Methods in Histotechnology 1992 American Registry of Pathology Washington, DC.
28. Lev, R. & Spicer, S. (1964) Specific staining of sulfate groups with alcian blue at low pH. J. Histochem. Cytochem. 12:309.[Medline]
29. SAS Institute Inc. (1986) SAS User’s Guide. Version 6 Edition 1986 SAS Institute Cary, NC.
30. Guyonnet Duperat, V., Audie, J. P., Debailleul, V., Laine, A., Buisine, M. P., Galiegue-Zouitina, S., Pigny, P., Degand, P., Aubert, J. P. & Porchet, N. (1995) Characterization of the human mucin gene MUC5AC: a consensus cysteine-rich domain for 11p15 mucin genes. Biochem. J. 305:211-219.
31. Ohmori, H., Dohrman, A. F., Gallup, M., Tsuda, T., Kai, H., Gum, J. R., Jr., Kim, Y. S. & Basbaum, C. B. (1994) Molecular cloning of the amino-terminal region of a rat MUC 2 mucin gene homologue. Evidence for expression in both intestine and airway. J. Biol. Chem. 269:17833-17840.
32. Gum, J. R., Byrd, J. W., Hicks, J. W., Toribara, N. W., Lamport, D. T. & Kim, Y. S. (1989) Molecular cloning of human intestinal mucin cDNAs. Sequence analysis and evidence for genetic polymorphism. J. Biol. Chem. 264:6480-6487.
33. Van Klinken, B. J., Einerhand, A. W., Duits, L. A., Makkink, M. K., Tytgat, K. M., Renes, I. B., Verburg, M., Buller, H. A. & Dekker, J. (1999) Gastrointestinal expression and partial cDNA cloning of murine Muc2. Am. J. Physiol. 276:G115-G124.
34. Smithson, K. W., Millar, D. B., Jacobs, L. R. & Gray, G. M. (1981) Intestinal diffusion barrier: unstirred water layer or membrane surface mucous coat?. Science (Washington, DC) 214:1241-1244.
35. Allen, A. (1989) Gastrointestinal mucus. Handbook of Physiology. The Gastrointestinal System. Salivary, Gastric, Pancreatic, and Hepatobiliary Secretion 1989:359-382 American Physiology Society Bethesda, MD.
36. Atuma, C., Strugala, V., Allen, A. & Holm, L. (2001) The adherent gastrointestinal mucus gel layer: thickness and physical state in vivo. Am. J. Physiol. 280:G922-G929.
37. Yamauchi, K., Kamisoyama, H. & Isshiki, Y. (1996) Effects of fasting and refeeding on structures of the intestinal villi and epithelial cells in White Leghorn hens. Br. Poult. Sci. 37:909-921.[Medline]
38. Sakamoto, K., Hirose, H., Onizuka, A., Hayashi, M., Futamura, N., Kawamura, Y. & Ezaki, T. (2000) Quantitative study of changes in intestinal morphology and mucus gel on total parenteral nutrition in rats. J. Surg. Res. 94:99-106.[Medline]
39. Uni, Z., Smirnov, A. & Sklan, D. (2003) Pre- and posthatch development of goblet cells in the broiler small intestine: effect of delayed access to feed. Poult. Sci. 82:320-327.
40. Ihara, T., Tsujikawa, T., Fujiyama, Y. & Bamba, T. (2000) Regulation of PepT1 peptide transporter expression in the rat small intestine under malnourished conditions. Digestion 61:59-67.[Medline]
41. Ihara, T., Tsujikawa, T., Fujiyama, Y., Ueyama, H., Ohkubo, I. & Bamba, T. (2000) Enhancement of brush border membrane peptidase activity in rat jejunum induced by starvation. Pfleug. Arch. 440:75-83.
42. Yaman, I., Fernandez, J., Sarkar, B., Schneider, R. J., Snider, M. D., Nagy, L. E. & Hatzoglou, M. (2002) Nutritional control of mRNA stability is mediated by a conserved AU-rich element that binds the cytoplasmic shuttling protein HuR. J. Biol. Chem. 277:41539-41546.
43. Brennan, C. M. & Steitz, J. A. (2001) HuR and mRNA stability. Cell. Mol. Life Sci. 58:266-277.[Medline]
44. Veress, I., Haghighi, S., Pulkka, A. & Pajunen, A. (2000) Changes in gene expression in response to polyamine depletion indicates selective stabilization of mRNAs. Biochem. J. 346:185-191.
45. Specian, R. D. & Neutra, M. R. (1980) Mechanism of rapid mucus secretion in goblet cells stimulated by acetylcholine. J. Cell Biol. 85:626-640.
46. Leparoux, S., Sine, J. P., Ferrand, R. & Colas, B. (1992) Behaviour of butyrylcholinesterase in the intestinal epithelial cells of starved and refed rats. Int. J. Biochem. 24:263-266.[Medline]
47. Shambaugh, G. E., 3rd., Koehler, R. R. & Radosevich, J. A. (1988) Nutrition and fetal brain maturation. II. Impact of maternal starvation on changing levels of acetylcholinesterase and enolase in vitro. Exp. Cell Res. 175:344-353.[Medline]
48. Rowland, I. R. & Malett, A. K. (1990) The influence of dietary fiber on microbial enzyme activity in the gut. Kritchevsky, D. Bonfield, C. Anderson, J. eds. Dietary Fiber: Chemistry, Physiology and Health Effects 1990:195-206 Plenum Press New York, NY. .
49. Miller, R. & Hoskins, L. (1981) Mucin degradation in human colon ecosystems. Fecal population densities of mucin-degrading bacteria estimated by the "most probable number" method. Gastroenterology 81:759-765.[Medline]
50. Deplancke, B., Vidal, O., Ganessunker, D., Donovan, S. M., Mackie, R. I. & Gaskins, H. R. (2002) Selective growth of mucolytic bacteria including Clostridium perfringens in a neonatal piglet model of total parenteral nutrition. Am. J. Clin. Nutr. 76:1117-1125.
This article has been cited by other articles:
|
|
 |
|
  T. J. Applegate, G. Schatzmayr, K. Pricket, C. Troche, and Z. Jiang Effect of aflatoxin culture on intestinal function and nutrient loss in laying hens Poult. Sci., June 1, 2009; 88(6): 1235 - 1241. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. R. Mott, P. B. Siegel, K. E. Webb Jr., and E. A. Wong Gene Expression of Nutrient Transporters in the Small Intestine of Chickens from Lines Divergently Selected for High or Low Juvenile Body Weight Poult. Sci., November 1, 2008; 87(11): 2215 - 2224. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. M. Byrne, M. Clyne, and B. Bourke Campylobacter jejuni adhere to and invade chicken intestinal epithelial cells in vitro Microbiology, February 1, 2007; 153(2): 561 - 569. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. L. Thompson and T. J. Applegate Feed withdrawal alters small-intestinal morphology and mucus of broilers. Poult. Sci., September 1, 2006; 85(9): 1535 - 1540. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. Smirnov, R. Perez, E. Amit-Romach, D. Sklan, and Z. Uni Mucin Dynamics and Microbial Populations in Chicken Small Intestine Are Changed by Dietary Probiotic and Antibiotic Growth Promoter Supplementation J. Nutr., February 1, 2005; 135(2): 187 - 192. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
