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PHYSIOLOGY, ENDOCRINOLOGY, AND REPRODUCTION |
* Discipline of Agricultural and Animal Science, School of Agriculture, Food and Wine, University of Adelaide, Roseworthy, 5371, South Australia; and South Australian Research and Development Institute, Pig and Poultry Production Institute, Nutrition Research Laboratory, Roseworthy, 5371, South Australia
2 Corresponding author: rebecca.forder{at}adelaide.edu.au
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: host-microbial interactions • development • chick • goblet cell • mucin
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The overlying mucus-gel layer is the first line of defense that foreign bacteria and other pathogens encounter when attempting to traverse the intestinal mucosa. Formation of the mucus gel is through goblet cell secretion of polymeric mucin glycoprotein (Forstner and Forstner, 1994; Klinken et al., 1995). These glycoproteins compete with bacteria for adherence via heterogenous oligosaccharide chains (Belley et al., 1999), thereby preventing noxious agents from coming into contact with the underlying epithelial cells. However, simultaneously, mucin provides a desirable environment for proliferation of specific microflora due to their high carbohydrate content (Deplancke and Gaskins, 2001). Thus, the chemical composition of mucus is essential for establishment of the intestinal barrier.
Histologically, mucins can be separated into 2 broad categories: neutral and acidic, with the latter further subdivided into sulfated and sialylated mucin types (Kiernan, 1990; Forstner and Forstner, 1994). These terms are derived from the chemical nature of the oligosaccharide sugar moieties, and histological techniques have been applied to detect whether particular mucins attribute their acidity or neutrality to the presence of these sugar groups (Kiernan, 1990).
Numerous rodent studies have compared germ-free (GF) and conventionally raised (CR) animals showing distinct changes in mucosal morphology and mucus composition associated with the presence of intestinal microflora. Compared with CR rodents, GF rodents exhibited a decrease in goblet cell size and number (Kandori et al., 1996), with a consequent reduction in mucus layer thickness, indicating a reduction in mucus production (Abrams et al., 1963b; Szentkuti et al., 1990). Compared with CR rodents, GF animals displayed less neutral mucin and sulfated mucin but greater amounts of sialylated mucin in small intestinal mucin fractions (Meslin et al., 1999; Sharma and Schumacher, 2001).
High levels of sulfated and sialylated mucins reportedly coincide with maturation of intestinal barrier function (Fontaine et al., 1996) in newborn rats (Shub et al., 1983) and pigs (Turck et al., 1993). Their presence during early development may be of particular importance as an innate barrier, because the acquired immune system is not fully functional in the neonatal intestine, rendering it more susceptible to infection (Cebra, 1999; Deplancke and Gaskins, 2001).
Currently, little information is available describing the effects of bacterial colonization on the secretory pattern of small intestinal mucins during early development of chicks. Reference to similar numbers of goblet cells containing acidic mucins compared with neutral mucins in CR poultry has been reported; however, ratios of acidic subtypes have not been described (Uni et al., 2003). Thus the aim of the current study was to investigate the effects of bacterial colonization on mucin production in ileal and jejunal goblet cells during early posthatch development of chickens. This study further sought to develop a new in vivo model system to study bacterial-intestinal interactions.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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After hatching, chicks were separated into 2 groups. Group 1 (n = 21) was transported to a brooding pen and raised under conventional conditions (CR; on litter, temperature, 32°C). Group 2 (n = 21) was transferred into a smaller sterile brooding isolator set to 32°C and raised as low bacterial load (LBL) chicks. Both groups received gamma-irradiated (25 kilograys) commercial high-energy broiler starter crumble (Ridley AgriProducts, Murray Bridge, South Australia). The LBL chicks received sterilized water. The CR chicks received normal drinking water dispensed via gravity into a large commercial drinker. Feed and water were available ad libitum during the trial, and all chicks were given free access to feed for 24 h before sampling commenced. The isolators were monitored microbiologically for contamination via daily surface swab analysis. Isolator size restricted the number of chicks that could be used in the trial; therefore, the experiment was replicated to achieve the desired animal numbers. All experimental work was approved by the Animal Ethics Committees of the University of Adelaide and the Department of Primary Industries and Resources of South Australia.
Intestinal Sample Collection
Three chicks were removed from both the brooding pen and the isolator and killed by cervical dislocation at 1, 4, and 7 d posthatch. Rectal swabs were collected to detect bacterial contamination. Segments (1 cm) of jejunum (adjacent to Meckel’s diverticulum) and ileum (adjacent to cecal tonsils) were dissected, flushed with cold sterile saline solution, opened longitudinally, and placed, mucosa side up, onto small pieces of blotting paper. The intestinal specimen was then fixed in 10% buffered formalin. This process was performed for each chick using sterile instruments for each dissection. Fixed samples were dehydrated, cleared, and embedded in paraffin wax for subsequent histological analysis. Consecutive longitudinal sections (7 µm) were placed individually onto poly-L-Lys-coated slides. Sections were then deparaffinized in Histolene (Fronine Laboratory Supplies Pty. Ltd., Riverstone, New South Wales, Australia) and rehydrated in preparation for staining.
Neutral and Acidic Mucin Staining
Because individual goblet cells can potentially produce all 3 types of mucin concurrently, the determination of mucin type required a series of alternative staining techniques. For neutral mucins, sections were subjected to mild acid hydrolysis to eliminate the contribution of sialic acid residues before periodic acid-Schiff (PAS) staining. After rinsing with both tap and distilled water, sections were immersed in periodic acid solution (Sigma, St. Louis, MO) for 20 min, washed, and immersed in Schiff’s Reagent (Sigma) for a further 20 min. Sections were rinsed in tap water for 10 min, dehydrated, and mounted in Entellan (ProSciTech, Kirwan, Queensland, Australia). Staining of acidic mucins required a technique that enabled distinct differentiation between sulfated and sialylated mucins. For this purpose, high iron diamine-alcian blue (HID-AB) pH 2.5 staining was used. Sections were treated in HID solution for 16 h at room temperature, rinsed, and immersed in alcian blue pH 2.5 for 5 min. Sections were then rinsed, dehydrated, mounted in Entellan (ProSciTech), and examined by light microscopy (Olympus BX60 microscope, Olympus, Tokyo, Japan) using a 20x objective and digital color images [super high quality (3,072 x 2,304 pixels)] captured using an Olympus Camedia C-7070 wide-zoom (5.7 to 22.9 mm; 1:2.8 to 4.8), 7.1 megapixels, 4x optical zoom camera.
Morphometry
Jejunal and ileal sections from each bird were stained with PAS and HID-AB pH 2.5. Image analysis programs ImageJ 1.33o (Rasband, 1997–2004) and VideoPro (Version 6.210, Leading Edge Pty Ltd., Adelaide, South Australia) were used in conjunction to measure a variety of parameters for each of the stained sections in which 10 villi/section were measured. ImageJ was used to calculate the number of goblet cells per unit of epithelial area (mm2) and individual goblet cell areas (µm2). For HID-AB pH 2.5, individual counts were obtained for goblet cells that stained either blue or brown. Goblet cells staining both brown and blue were counted separately and termed intermediate. The summed values provided a total count of goblet cells in HID-AB pH 2.5 sections. VideoPro was used to compute measurements of total villus area (µm2); epithelial area (µm2), which was the lamina propria area subtracted from the total villus area; villus length and breadth (µm); crypt depth (µm); and total goblet cell area expressed as a proportion of epithelial area (µm2).
Data Analysis
Statistical analyses were performed using the SPSS software package V11.5 (SPSS Inc., Chicago, IL). Group (LBL vs. CR) x age effects were analyzed using a 2-way AN-OVA fitted with a Bonferroni adjustment. Student t-tests were used to compare acidic and neutral goblet cell numbers. Significance was determined as P 
0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). In CR and LBL birds, jejunal goblet cells exhibited a marked decrease in acidic mucins from d 1 to 4 posthatch (P 0.05) and then by d 7 returned to values comparable to d 1. In the ileum, a similar trend was also observed, occurring only in CR birds (P 0.05). Overall, there were greater numbers of goblet cells containing acidic mucins in the ileum compared with the jejunum on d 4 (jejunum, 1.4 ± 0.1; ileum, 2.1 ± 0.2, P 0.01) and d 7 (jejunum, 2.4 ± 0.2; ileum, 3.4 ± 0.3, P 0.01).
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). Although not statistically significant at d 7 (P = 0.108), the CR group tended to have fewer goblet cells containing sulfated mucins than the LBL group and a greater number of sialylated mucins with an increase in cell number from d 4 (Table 1, Figure 1). Intermediate cells, with staining for both sulfated and sialylated mucin increased in number from d 1 to 4 in LBL and CR birds that were maintained at d 7. By d 7, intermediate goblet cell numbers were greater in CR birds (Table 1).
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0.05) followed by an increase similar to d 1 values at d 7 posthatch (3.2 ± 0.1; P 0.001). When the number of goblet cell-containing neutral mucins (PAS-stained sections) was compared with numbers of cells containing acidic mucins (HID-AB pH 2.5 stained sections) from both LBL and CR birds, there were no significant differences observed (Table 2).
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), with the increase in cell number contributing to the increase in total goblet cell area. In the jejunum, there was no difference in total PAS-stained goblet cell area between CR and LBL chicks, with only a decrease in goblet cell area in CR chicks from d 1 to 4, which was maintained at d 7 (P 0.05). In the ileum, there were no changes in PAS-stained goblet cell area over the 7-d period in either LBL or CR birds. However, when compared with LBL chicks, CR birds had a significant decrease in goblet cell area at d 4 posthatch (Figure 3). Further analysis of average individual goblet cell area (µm2) at d 4 revealed that CR chicks possessed smaller goblet cells compared with LBL chicks (LBL, 157 ± 3 µm2; CR, 109 ± 5 µm2, P 0.01) and that this was not dependent on villus area (r = 0.009, P >0.05).
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). There were no differences in villus breadth between CR and LBL birds in the ileum. However, in the jejunum, villus breadth was greater in CR chicks at d 4 (Table 3).
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). Statistically significant differences between CR and LBL chicks were observed only in the jejunum, with a greater villus length in CR birds as opposed to LBL at both d 4 and 7 posthatch (Table 3).
Villus Area
Jejunal and ileal total villus area and epithelial area were increased in both CR and LBL chicks from d 1 to 4, with no further increase observed at d 7 (Table 3). Differences between CR and LBL were observed at d 4, with the CR chicks exhibiting a greater villus area, which was more prominent in the jejunum (Table 3). The jejunum villus and epithelial areas of CR birds tended to remain greater at d 7 compared with LBL villi, although statistical significance was not attained (epithelial area, P = 0.058; total villus area, P = 0.067).
Crypt Depth
The villus-crypt axis of poultry is not developed until d 5 posthatch (Uni et al., 2000); hence, crypt depth measurements were only conducted in d 7 birds. In the jejunum, it was found that crypt depth did not differ significantly between LBL and CR birds (LBL, 122 ±27 µm; CR, 161 ±27 µm, P = 0.18). In contrast, ileal crypt depth in LBL birds was significantly lower than in CR birds (LBL, 99± 4 µm; CR, 149 ± 7 µm, P 0.05).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The preservation of sulfated mucins, evident throughout the first week of development in LBL chickens, was consistent with other developmental studies (Hill et al., 1990). A high degree of sulfation is characteristic of immature goblet cells (Turck et al., 1993). Because the level of intestinal microflora was low, the retention of sulfated mucin during posthatch development may be indicative of an immature gut, outlining the influence of bacteria on mucin production and overall gut maturity. The reasons for this are still unclear; however, some bacteria possess mucin-specific glycosidases and proteases, which are able to degrade mucus and facilitate colonization of the epithelial surface (Corfield et al., 1992; Deplancke and Gaskins, 2001). Bacteria such as Helicobacter pylori secrete glycosulfatases, which can cleave sulfate from its linkage to mucin sugars (Roberton and Wright, 1997). The switch from predominately sulfated mucins to acetylated sialylated mucin in neonatal animals could represent a defense strategy. The hydroxyl groups of sialic acids are highly substituted by acetyl esters, which serve as added protection as they block against further glycosidic degradation, with reports that 2 or more acetyl groups inhibit enteric bacterial sialidases (Corfield et al., 1992; Belley et al., 1999). Both sulfate and sialic acid groups have protective properties (Corfield et al., 1992; Roberton and Wright, 1997). As colonization becomes greater, the need for greater protection against mucus degradation is increased, which would explain the observed increase in sialomucin production. Moreover, in the current study, the greater number of goblet cells containing acidic mucin in the ileum compared with the jejunum would suggest the distal ileum may be a preferred region for bacterial colonization. This is consistent with other findings using chicks, which have demonstrated a distal increase in the density of goblet cells along the duodenalileal axis (Uni et al., 2003).
Throughout the current study, intestinal crypts showed a predominant HID-positive staining for sulfated mucins in both CR and LBL birds, with the sialylated stained goblet cells tending to be located from mid to villus tip. Migration rate of goblet cells from crypt to villus tip has been reported to take approximately 2 to 3 d in poultry (Imondi and Bird, 1966; Uni et al., 2000), with CR birds having a greater rate of migration than GF birds (Cook and Bird, 1973). Considering these differences in migration rate and the absence of sialylated mucin in intestinal crypts, the change in mucin composition in CR birds along the villus may have been due to differences in the luminal environment and not to the migration of goblet cells during posthatch development.
The presence of neutral mucins in ileal and jejunal goblet cells of day-old chicks was consistent with previous studies in poultry (Uni et al., 2003) but differed from mammalian models. It has been reported that little to no neutral mucin was detected in the lower small intestine and colon of neonatal rodents and pigs but did increase with age (Hill et al., 1990; Turck et al., 1993; Deplancke and Gaskins, 2001). Germ-free studies have used rodent and pig models, which during the first few weeks after birth are dependent on maternal milk resources. Chicks, however, must have the capacity to digest complex carbohydrates immediately after hatch (Sklan, 2001). Thus, the gut of a day-old chick requires advanced intestinal development compared with a day-old rodent or pig, in which the intestine is comparable to chicks at d 18 of incubation (Uni et al., 2003). In the current study, whether the presence of neutral mucins in the ileum and jejunum was due to bacterial colonization or dietary components, or both, is yet to be determined. Bacterial species, mainly type-1 fimbriated, have been demonstrated to possess receptors for mannose residues in vitro (Firon et al., 1984; Marc et al., 1998; Vimal et al., 2000). As the animals age, there is increased bacterial adhesion to mannose and, consequently, reduced susceptibility to infection (Nagy et al., 1992). The production of neutral mucins could therefore serve as a protective mechanism against invasion by pathogenic bacteria (Runnels et al., 1980; Dean-Nystrom and Samuel, 1994). In this study, because neutral mucins in both the LBL and CR chicks displayed similar patterns, their presence at this time point was likely the result of increased intestinal maturity to facilitate the breakdown of complex carbohydrates. Future studies could measure neutral mucin content, particularly that of mannose residues, at a later stage of development to determine the extent to which bacteria may influence the production of this mucin type.
The differences in villus length, breadth, and area between the ileum and jejunum were consistent with previous findings conducted in poultry, with the jejunum displaying an increase in all 3 parameters compared with the ileum (Iji et al., 2001), supporting its importance as a site for nutrient digestion (Iji et al., 2001). It has been well documented that villus length and crypt depth increase with age (Uni et al., 1995; Iji et al., 2001), which was observed in both CR and LBL birds. The increased mucosal development observed in CR birds compared with LBL birds may have been due to bacterial-diet interactions and the need for greater absorptive area to accommodate the by-products associated with microbial fermentation. The increased villus length and crypt depth evident in CR birds was consistent with previous studies (Abrams et al., 1963a; Cook and Bird, 1973; Kleessen et al., 2003). However, this was not true of other reports in which crypt depth was found to be greater in CR animals, but villus length was significantly decreased (Shirkey et al., 2006). These studies were conducting using pigs; thus, species differences in overall intestinal morphology and also contributing factors such as microbial colonization and digestive enzyme function may have contributed to the contrasting results.
In the current study, the dramatic changes in mucin composition in CR birds at d 4 posthatch could have coincided with an increase in immune system development. With the depletion of the yolk sac, and subsequent maternal antibody resources, the stimulation of goblet cells to alter mucin glycosylation may have functioned to defend against pathogenic infection at this stage in development. At d 4 posthatch, it has been reported that an upregulation of mRNA expression of proteins involved in immune function such as antimicrobial peptides and proinflammatory cytokines was greatly increased in the gut-associated lymphoid tissue (Bar-Shira et al., 2003). Cytokines have been reported to increase the extent of mucin production and goblet cell proliferation (Blanchard et al., 2004) and also to produce changes in the glycosylation of mucins (Beum et al., 2005). Bacterial endotoxins, such as lipopolysaccharides, are the major outer surface membrane component of gram-negative bacteria and have been found to upregulate mRNA expression and secretion of cytokine IL-8 and mucin genes MUC5AC and MUC5B (Smirnova et al., 2003). Relative expression of proinflammatory cytokines IL-1β and IL-6 has been reported to be highest in pigs inoculated with adult porcine feces and nonpathogenic Escherichia coli compared with GF animals (Shirkey et al., 2006), both of which have been reported to trigger mucin release and upregulate MUC gene expression (Enss et al., 2000; Deplancke and Gaskins, 2001).
The development of gut immunity and its interactions with mucin dynamics and bacterial colonization warrants further investigation. The protective properties of mucin glycoprotein could be utilized for the development of novel therapies, such as administration of probiotics and prebiotics for the treatment and prevention of infection in younger animals. Findings of the current study provide insight into the influence of intestinal microflora on goblet cell and mucosal cytoarchitecture during posthatch development.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Received for publication June 4, 2007. Accepted for publication July 16, 2007.
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REFERENCES |
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