Protein source and nutrient density in the diets of male broilers from 8 to 21 d of age: Effects on small intestine morphology

Abstract

INTRODUCTION

The jejunum of the gastrointestinal tract is the major site of feed digestion and nutrient absorption. During their migration from crypt base to villus tip, pluripotent columnar cells can differentiate into digestive, absorptive, or mucin-producing roles (Moog, 1950; Cheng and Leblond, 1974a,b). Although intestine organogenesis is primarily programmed by in vivo genetic information, intestine development may be influenced by in vitro nutritional manipulation. Longer villi, shorter crypts, and larger goblet cells have been found in the intestines of broiler chicks fed beneficial additives (Xu et al., 2003; Xia et al., 2004; Smirnov et al., 2006; Zhang et al., 2005; Baurhoo et al., 2007; Salim et al., 2013). An examination of the morphology of these cells may help us elucidate the capacity of the small intestine to utilize nutrients.

Moderate amounts of dietary fiber improve digestive organ development, enzyme production, and nutrient digestibility in birds (Abdelsamie et al., 1983; Gonzalez-Alvarado et al., 2007). Those improvements are mostly due to increasing gastro-duodenal refluxes that enhance the contact between digestive enzymes and nutrients (Duke, 1992). As a plant protein source, dried distiller grains with solubles (DDGS) contain higher amounts of fiber than animal protein sources, such as meat and bone meal (MBM). However, high fiber diets may decrease feed retention time in the digestive tract. Rochell et al. (2012) reported that a DDGS diet has a shorter passage time in the intestine than does an MBM diet. Shorter feed retention time may decrease the duration of contact between chyme and absorptive cells (Washburn, 1991). Nevertheless, avian digestive tracts may have differential responses to DDGS diets and MBM diets.

Nutrient density is another factor that may affect animal intestine development. Higher nutrient densities improve the expression of digestive enzymes (Nitsan et al., 1991) and transporters (Chen et al., 2005; Mott et al., 2008). Structural adjustment is a direct way to control enzyme and transporter levels. Jejunum epithelial numbers have been shown to decrease in chickens fed an energy-restricted diet (Palo et al., 1995). Research on piglets has shown that jejunum villus height increased in response to the feeding of high protein diets (Gu and Li, 2004).

Small intestines of chickens develop rapidly during the first 5 d after birth (Noy and Sklan, 1998), whereas chickens fed a higher nutrient density diet grow faster throughout all growing phases (Saleh et al., 2004; Nahashon et al., 2005; Zhai et al., 2013). Intestinal structures may also be further modified to adapt to nutrition manipulation during latter grow-out phases. Previous research in our lab has shown that high amino acid (AA) or high apparent metabolizable energy (AME) densities in the diets fed to broilers from 8 to 21 d of age improved their feed conversion ratio (FCR). The objectives of this research were to investigate effects of dietary protein source and nutrient density from 8 to 21 d on broiler small intestine morphology at 21 d. The relationship between growth performance (BW gain and FCR) and intestine structure was also studied.

MATERIALS AND METHODS

Birds and Diets

Detailed descriptions of bird management (water, feed, light program, and pen environment) and the arrangement of the treatment groups in the broiler facility were presented in a companion study by Wang et al. (2014). Briefly, a total of 1,120 Ross × Ross 708 male broiler chicks were randomly distributed among 80 floor pens so that 14 birds occupied each pen. The 80 pens were divided into 10 blocks that were distributed throughout the environmentally controlled facility. Birds were fed the same starter diet from 1 to 7 d and one of 8 experimental treatment diets from 8 to 21 d. The 8 treatment diets (2 × 2 × 2 factorial) were randomly assigned to 8 pens within each of the 10 blocks. Each treatment diet contained either a high inclusion of distiller grains with solubles (hDDGS) or a high inclusion of meat and bone meal (hMBM), a moderate or high AA density, and a moderate or high AME density. Ingredients and nutrient contents of experimental diets shown in Table 1 were also provided by Wang et al. (2014). Bird husbandry, handling, and sampling procedures were approved by the Institutional Animal Care and Use Committee of Mississippi State University.

Small Intestine Sampling

At 21 d, 2 birds per pen (20 chicks/dietary treatment) were dissected for determination of duodenum, jejunum, and ileum lengths. Birds were weighed, euthanized by CO₂ asphyxiation, and dissected. Subsequently, 3 intestinal segments (duodenum, jejunum, and ileum) were excised, and their lengths were individually measured. The duodenum section extended from the pylorus of the gizzard to the end of the duodenal loop, the jejunum section extended from the end of the duodenal loop to Meckel’s diverticulum, and the ileum section extended from Meckel’s diverticulum to the ileocecal junction. Tissue samples (1.5 cm) were obtained from the midpoint of the jejunum of each chick and were placed in 10% buffered formalin phosphate (Fisher Scientific, Fair Lawn, NJ) for subsequent morphological examination.

Jejunum samples were dehydrated, cleared, and embedded in paraffin. Serial sections (5 μm) of each sample were cut, mounted on glass slides, and stained with periodic acid-schiff (PAS) and Alcian blue (ALB) stains. Neutral mucin-producing goblet cells were detected by staining with PAS reagent (McManus, 1948), and acid mucin–producing goblet cells were detected by staining with ALB reagent (Lev and Spicer, 1964).

Jejunum Morphological Examination

Villi and goblet cells were photographed under a light microscope (Micromaster, Fisher Scientific, Pittsburgh, PA) using the method presented by Fasina et al. (2010). Morphometric parameters of jejunum villi were performed at a magnification of 40×, and measurements of goblet cells were performed at a magnification of 400×. All measurements were analyzed using ImageJ software (National Institutes of Health, Bethesda, MD).

Morphometric parameters recorded included total villus height (from the tip to the bottom of each villus), mid-point villus width, villus area (calculated by multiplying villus height by mid-point width), crypt depth (from the base to its opening), and villus to crypt ratio (V:C; calculated by dividing villus height by crypt depth) according to the procedure of Fasina et al. (2010). Also, the muscle thickness of the jejunum was measured from the submucosal and muscular layer boundary to the muscular layer and peritoneum boundary. Goblet cell densities were expressed as the number of goblet cells per unit of villus height.

Statistics Analysis

A randomized complete block design (pen location as block) with 10 replications (block as a replication unit) was used to test for effects of dietary treatment on small intestine length, small intestine segment length, goblet cell density, and each villus parameter. The treatments were set in a 2 × 2 × 2 factorial arrangement. All parameters were analyzed using SAS (SAS Institute, 2010). A 3-way ANOVA using the PROC MIXED procedure was used to determine the significance of responses to protein source, AA, and AME levels, as well as their interactions. When significant global effects were observed, comparisons of least squares means (Tukey-Kramer) were used to detect significant differences among treatment means. The protein source, AA, and AME levels were designated as fixed effects and block as a random effect. Linear correlations between the lengths of the whole small intestine and its segments (duodenum, jejunum, and ileum), each morphological parameter (muscle thickness, crypt depth, villus height, and villus width) at 21 d, and BW gain, as well as FCR from 8 to 21 d were analyzed using the PROC CORR procedure. Body weight gain and FCR were reported in a published companion paper (Wang et al., 2014). Global effects, differences among least squares means, and correlations were considered significant at P ≤ 0.05.

RESULTS

Small Intestine Length

As compared to hMBM in diets, high inclusion of distiller grains with solubles (hDDGS) increased small intestine length by 3.00 cm (2.5%) (P = 0.045, Table 2). An interaction between protein source and AA level existed for broiler jejunum length (P = 0.017). In chicks fed diets containing a high AA density, chicks fed hDDGS diets exhibited longer jejuna than those fed hMBM diets.

Morphological Characteristics of the Jejunum Villus

Morphological examination showed that dietary treatment did not affect jejunum villus height, width, areas, crypt depth, villus to crypt ration (V:C), or goblet cell size and density (Table 3). A 3-way interaction among dietary protein source, AA density, and high AME density was found for muscle thickness (P = 0.027). Birds fed an hDDGS diet with moderate AA and high AME densities exhibited thicker intestinal muscle layers than those fed either hDDGS diets with high AA and high AME densities, hMBM diets with high AA and high AME densities, hMBM diets with high AA and moderate AME densities, or hMBM diets with moderate AA and high AME densities.

Correlation of Intestine Morphology with Growth Performance

Lengths of the small intestine, duodenum, jejunum, and ileum, as well as muscle thickness, villus height, and width were not correlated to BW gain or FCR (P > 0.05, data not shown). However, crypt depth was negatively correlated to BW gain (P = 0.042) and positively correlated to FCR (P = 0.029).

DISCUSSION

In the current trial, total small intestine lengths increased in broilers fed high inclusion of distiller grains with solubles (hDDGS) diets. This result is consistent with that of Barekatain et al. (2013), who reported that feeding sorghum DDGS to broilers increased their gastrointestinal tract size. Crude fiber concentration in the hDDGS diets was higher than that in the hMBM diets (Table 1). Previous studies have shown that the use of dietary ingredients with high fiber contents increased intestinal length (Abdelsamie et al., 1983; Jorgensen et al., 1996; Sklan et al., 2003; Gonzalez-Alvarado et al., 2007). In addition, soluble fiber increases digesta viscosity (Dikeman and Fahey, 2006), which subsequently increases intestinal laxation and results in longer intestinal lengths (Smits et al., 1997). A longer small intestinal length allows for a greater digestive and absorptive area. Broilers fed hDDGS diets exhibited longer small intestines, but their growth rate through the end of the feeding trial was similar to those fed hMBM diets (Wang et al., 2014). It has been reported the retention time of DDGS in the gastrointestinal tract of broilers fed semi-purified diets is shorter than that of broilers fed MBM (Rochell et al., 2012). On the other hand, increasing the levels of supplementary poultry fat in hDDGS diets (Table 1) may increase feed retention time (Mateos and Sell, 1981; Mateos et al., 1982). However, the retention time of the experimental diets in the birds of the current trial was not investigated. Studies concerning the effects of the dietary inclusion of DDGS and MBM on nutrient utilization in broilers should include tests on feed passage rate.

Intestinal crypts, also known as the precursors of intestinal epithelial cells, contain pluripotent stem cells at their base and functional enterocytes at their periphery. In the present trial, jejunum crypt depth was found to be negatively correlated to BW gain and positively correlated to FCR. These results indicate that chicks with shorter crypts exhibit better growth performance. It is possible that more crypt cells differentiated into functional epithelial cells in the fast growing broilers. In addition, shorter crypt depths are indicative of a longer time needed for cell regeneration (Holt et al., 1985; Smith et al., 1990; Pagan et al., 1999; Gao et al., 2008). The turnover of intestinal epithelial cells is normally fast and requires higher amounts of energy and protein to maintain the rapid production of new cells (Creamer et al., 1961; Imondi and Bird, 1996). Shorter crypt depths may reduce intestinal maintenance, thus sparing energy and protein for use in muscle deposition (Xu et al., 2003; Markovicva et al., 2009).

Intestinal length relative to body length is shorter in broilers than in other livestock animals. Furthermore, small intestines in broilers that are short and smooth have poorer digestive and absorptive capabilities. In the current trial, when chicks were fed diets with moderate AA and high AME densities, chicks fed hDDGS diets exhibited thicker jejunum muscle layers than did those fed hMBM diets. It was reported that high fiber diets increased intestinal size (Sklan et al., 2003; Gonzalez-Alvarado et al., 2007). Previous studies have also shown that dietary fiber activated intestinal peristalsis (Esonu et al., 2001, 2004). In addition, muscle thickness in the duodenum is positively related to increased numbers of fiber-consuming microbial products in the ceca (Maisonnier et al., 2003). Therefore, the high fiber content in hDDGS diets may have stimulated intestinal muscle growth. Here specifically, dietary fiber in DDGS may have increased bacterial populations and muscle layer growth in the jejuna.

Goblet cells secret mucin in the digestive tract to protect the intestinal membrane from digestive enzyme degradation and pathogen invasion. Nevertheless, although shorter jejunum crypt depths were found to be associated with improved growth performance, goblet cell size and density was not affected by dietary treatment in the present trial. However, mucin production may be affected by dietary treatment. It has been reported that mucin production is greatly influenced by dietary threonine (Horn et al., 2009) and fiber (Smirnov et al., 2006) levels. However, mucin concentration in the broilers fed the experimental diets was not evaluated in the current trial. Future studies should be conducted to further elucidate how dietary treatments affect feed digestion and nutrient absorption, microbial activity, brush border enzyme activity, and glucose and AA transporter expression levels.

In the companion study, but the feed conversion of broiler chicks was affected only by dietary nutrient density, not by protein source (Wang et al., 2014). However, in the current study, protein source affected intestinal structure. In conclusion, high nutrient density diets may improve broiler performance without affecting their intestinal structure. In addition, an hDDGS diet may facilitate small intestine longitudinal growth in broilers.

We thank Ms. Donna Morgan of the Mississippi State University, Department of Poultry Science, for her technical assistance during this research.

REFERENCES