Todd J. Applegate
Kristi Thompson
Purdue University
Department of Animal Science
West Lafayette, IN 47907
U.S.A.
Introduction
The gastro-intestinal tract (GIT) in the turkey is a very dynamic organ system. Knowledge of its functionality and relationship with different nutrients in the diet can greatly increase our ability as nutritionists to formulate more precise diets. As such, these more precise diets can modify nutrient retention, growth, nutrient partitioning, and overall health of our birds.
Embryonic & early post-hatching intestinal development
During the early posthatch growth and development period, there is a tremendous energetic allocation to GIT growth at the expense of most other body parts (Konarzawski et al., 1990). Intestinal development has been the subject of several reviews, for a more detailed discussion on digestive tract development in poultry, one should review Moran (1985), Dibner et al. (1996), Sell (1996), Noy and Sklan (1997), and Jin et al. (1998).
Within the embryonic chick intestine, the brush border begins to develop as a network of microfilaments, which aggregate to form uniform projections as early as 7 to 11 days of incubation (Chambers and Grey, 1979). The microfilaments elongate to form rootlets between 11 and 15 days and further form a terminal web and microvilli between 19 days of incubation and 5 days of age. This provides both structure and support to the brush border epithelium (Chambers and Grey, 1979). Other researchers have also noted villi containing short microvilli as early as 16 days of incubation (Overton and Shoup, 1964; Lim and Low, 1977). At hatch, however, portions of the lower small intestine still contain epithelial discontinuities (Bayer et al., 1975). In addition, formation of goblet cell pores and extensive convolution of intestinal villi does not occur until approximately 1 week of age or later (Bayer et al., 1975). Not only is the small intestine morphologically immature at hatch, it is also functionally immature in terms of digestive and absorptive capabilities (Holdworth and Hastings-Wilson, 1967; Sell et al., 1991; Obst and Diamond, 1992).
During the first week after hatching, the poult's small intestine increases in weight 9-fold and doubles in length (Sell et al., 1991). As a proportion of yolk-free BW, this represents an increase in relative weight of the small intestine from 1.6% at hatch to 6.9% by 8 days of age (Sell et al., 1991). In chicks, the small intestine will increase in length over the first 5 days after hatch, even during an imposed fast (Baranyiova, 1972). Increases in the relative weight of the small intestine, intestinal diameter, and villus length only occur after feeding, however (Baranyiova, 1972; Baranyiova and Holman, 1976). Energy for support of rapid GIT growth during the acclimation period is at the expense of all other tissues and organs (Konarzewski et al., 1990). Several authors have described the intestine as a relatively "flexible system" which can undergo morphological and functional adjustments depending on the demands of the animal (Moran, 1985; Obst and Diamond, 1992; Starck, 1996). For example, Uni et al. (1995) noted a positive correlation of jejunal and ileal villus volumes, and enterocyte density with feed intake in chicks during the first 2 weeks of age (r=0.77, 0.67, and 0.62, respectively).
Functional maturation of the small intestine is both a physical and physiological process and is one of the main constraints to early growth after hatching (Konarzewski et al., 1989, 1990; Ricklefs et al., 1998). Physiological maturation (in terms of digestive and absorptive functionality) of the digestive tract involves the increased production of pancreatic and intestinal enzymes (Nitsan et al., 1991a,b; Sell et al., 1991; Pinchasov and Noy, 1994) and changes in nutrient transporters (Holdsworth and Hastings-Wilson; 1967; Shehata et al., 1984; Obst and Diamond, 1992). However, the physical development of the gastrointestinal tract, namely the increase in surface area of the small intestine, can be a more limiting factor to early growth than changes in digestion or nutrient transporters (Uni et al., 1995, 1996).
In the chick, villus length throughout the small intestine increases over two-fold during the first 5 days after hatch (Dibner et al., 1996), villus volume (mm3/cm3) increases 3- to 4- fold from hatch to 10 days of age (Uni et al., 1995), and the apical surface area of jejunal enterocytes increases from 300 to 410 μm2 from 1 to 14 days of age (Ferrer et al., 1995). Other morphologic changes include changes in the rates of intestinal crypt cell proliferation and enterocyte turnover (Starck, 1998). Enterocyte turnover rates in the small intestine were originally reported to take 2 days in the chick (Imondi and Bird, 1966) but has been more also reported to vary from 2.8 to 2.9 days in the hatchling to 8.9 to 10.6 days in adult Japanese quail (Starck, 1996).
During the first 7 days after hatching in the poult, villus growth also increases over two-fold, while enterocyte migration remains constant at approximately 2μm/hour (Applegate et al., 1999). As such, enterocytes take approximately 4 to 5 days to migrate from the crypt region to be sloughed from the villus tip. Between hatching and 7 days of age, cells along the entire length of the crypt-villus axis stain positive for a compound indicating they are actively dividing. Therefore, the increased functionality of the villus in the poult after hatching appears to be affected more by the net accumulation of cells, rather than the turnover of the mucosal epithelial surface (as has been previously suggested).
Changes in the structural components of the intestine
Delayed access to feed greatly affects intestinal morphology and growth of the bird after feeding. For example, Corless and Sell noted that holding time of poults can have prolonged effects on overall growth to 28 days of age. Delayed access to feed is followed by microvilli clumping (within 24 hours) and delayed jejunal mucosal development and crypt structure up to 9 days after hatching (Uni et al., 1998). Baranyiova and Holman (1976) reported that when chicks were fasted between hatching and 5 days of age, villus height and intestinal diameter did not change. Even after chicks are given access to feed, jejunal and ileal villus volume is reduced, crypt depth is reduced and crypt morphology remains abnormal. Short-term fasting after hatching has also been reported to slow or stop GIT epithelium engaged in cell division (Cameron et al., 1964). This may also be reflected in morphological changes associated with fasting.
Research from our laboratory also suggests prolonged effects on intestinal development when birds undergo a delay in placement after hatching. For example, Potturi et al. (2005) noted that poults that underwent a 48 hours delay in placement had 50 micron shorter and 6.8 micron narrower villi with 5.9 micron shallower crypts up at 5 days after placement. Also of note was a lower number of cells actively dividing coinciding with a greater number undergoing programmed cell death in those same birds that were delayed up to 48 hours prior to placement. From a microbial exposure standpoint, there was also a greater amount of aerobes associated with the lower small intestine 1-day post-placement in the delayed placement birds, but not at 5 days post-placement.
Short-term changes in villus morphology may not, however, adversely affect subsequent digestion and absorption of nutrients upon refeeding. Transient reductions in feed intake up to 75% of ad libitum intake in 6-weeks -old chickens for example can substantially reduce villus height but does not influence nutrient utilization upon refeeding (Michael and Hodges, 1973). Other morphological changes in villus height have been reportedly influenced by: feed restriction, competition with normal GIT microflora, coccidial infections, dietary fiber content, and changes in growth or productive needs (Moran, 1985).
Compounds that can potentiate intestinal maturation
The yolk stalk can provide a passageway for transport of yolk material into the small intestine up to 72 hours after hatching and this yolk material could be detected as proximal as the duodenum (Esteban et al., 1991; Noy et al., 1996). Jeurissen et al. (1991) reported that colloidal carbon injected into the yolk sac of chicken eggs at 10 days of incubation was subsequently absorbed from the yolk sac into the lymphoid tissue around the yolk stalk and was subsequently transported to leukocytes and mononuclear phagocytes. This transport may be a means for physical passage of yolk-derived antigens, which could then aid in early immunological maturation of the intestine. Insulin and IGF-I have been found to be deposited into the yolk by the hen (Scavo et al., 1989) and both have been demonstrated to significantly influence metabolic, growth, and differentiation processes during early embryonic development (de Pablo et al., 1990). In domestic farm animals, the stimulatory effects of maternal colostrum and milk on intestinal development via various growth factors (insulin, EGF, IGF-I and IGF-II) have received considerable attention (Odle et al., 1996 for review). If such growth factors are transported into the intestinal lumen from the yolk sac, they could potentially stimulate intestinal maturation.
Uni et al. (1998) noted that yolk sac ablation of chicks at hatching caused a transient decrease in villus volume and crypt depth throughout the small intestine. Intubation with yolk contents after yolk sac ablation, however, increased body weight, enhanced pancreatic and liver development, and restored relative pancreatic lipase activity when compared with ablated chicks (Nitsan et al., 1995). These indirect results, therefore, suggest that the yolk does contain substances that aid in the functional development of the small intestine.
A practical approach to applying such compounds has been investigated by researchers at North Carolina State University by studying the effects of in ovo administration of peptide YY. Peptide YY has demonstrated effects in other specie of inhibiting gut motility (Savage et al., 1987) and stimulating small intestinal absorption of glucose (Bird et al., 1996). Application of peptide YY in ovo at transfer demonstrated benefits of improving BW from hatch up to 3 days of age in the poult (Coles et al., 1999a) and BW and feed efficiency up to 7 days of age in the chick (Coles et al., 1999b).
Other practical approaches to enhancing intestinal development include early feeding and in ovo feeding. For example, Sklan et al. (2000) demonstrated that weight loss in poults emerging from the shell could be minimized if a starter diet was placed within the hatching trays. Without the early access to feed, poults lost between 0.14 and 0.17 g/hour; whereas if poults were given early access to feed, they emerged from the hatcher 3.9 g (if hatched during 0-22 hours of hatching window) or 0.8 g heavier (if hatched during the 22-48 hours of hatching window). Uni and Ferket (2003; 2004) have utilized this information to potentiate the metabolic shift and intestinal development in the chick and the poult by in ovo feeding. Published research with chicks (Tako et al., 2004; Uni and Ferket, 2004) has reported that administration of maltose, sucrose, dextrin, and β-hydroxy-β-methylbutyrate to the amniotic fluid of chicken eggs at 18 days of incubation increased intestinal surface and mucosal carbohydrase activity up to 3 days after hatching and improved body weights to at least 14 days of age.
In considering early shifts in metabolism, the newly hatched poult is in a gluconeogenic state (deriving energy from body reserves) and will remain so until it consumes exogenous nutrients (Romanoff, 1960). Researches noted above as well as other researchers in the past have tried to facilitate –or- expedite this shift in metabolism through administration of glucose or metabolic derivatives at the hatchery. Administration of these compounds will temporarily shift the poult from a gluconeogenic and ketogenic metabolic condition towards to a glycolytic state. This shift, however, is exacerbated if feed and water are not immediately available after the initial glucose surge (Moran, 1989; 1990). In other words, if the poult doesn't have feed and water after the initial shift in metabolism, it has a tendency to crash.
Impact of Intestinal Development on Growth
Turkey poults and Pekin ducklings hatch from similar sized eggs, have similar lengths of incubation, and BW at hatch. Yet, the male Pekin duckling attains a mature market BW of approximately 3.2 kg in 6 weeks or less whereas the male turkey only weighs approximately 2.3 kg at the same age. Despite having similar BW at hatching, the BW of ducklings is nearly double that of turkey poults by 7 days of age (Applegate and Lilburn, 1998). Much of this phenomenal growth difference can be attributed to the rapid development of the small intestine. By 7 days of age, the duckling's lower small intestine is 3.7X heavier, 1.6X longer, and 2.3X more dense (g/cm). Histological sections of the distal jejunum reveal that the duck also has a more rapid villus growth from hatch to 3 days of age (as depicted in Figure 1), thereby allowing the duck to achieve this phenomenal growth (vs the poult) during this critical 7 days growth period. Better understanding of these differences may allow us to adopt strategies to try to upregulate or accentuate intestinal functionality in the developing poult.
Role and Composition of GIT Microbiota
The avian gastrointestinal tract (GIT) accommodates an extensive and diverse indigenous microbiota that is crucial to proper intestinal function and defense. Generally, the most beneficial and perhaps important role of the microbiota is colonization resistance, or the ability of the microbiota to inhibit the colonization of the gut by exogenous pathogenic bacteria (Berg, 1996). However, other research has indicated that the intestinal microbiota may also contribute to small intestinal development (Hooper, 2004).
At hatch, the intestinal tract of the poult is generally sterile. However, within hours of hatch, bacteria from the poults' environment begin to colonize the intestinal tract. During the days following hatch, bacterial species compete with one another in an attempt to become established within the intestine. For example, lactobacilli appear by the fourth day and large populations of obligate anaerobes appear soon after (Mead and Adams, 1975). This process, known as ecological succession (Wilson et al., 1986), continues through stages until a stable climax bacterial population that closely resembles that of adult birds develops within the small intestine (Smith, 1965; Rolfe, 1997). At this point, researchers are uncertain as to when an adult population is established. Research by Lee et al. (2002) has indicated that populations of indigenous microbiota vary significantly with age up to 49 days. However, no research has been conducted past this point.
The indigenous microflora of the mammalian adult intestinal tract is composed of 400 to 500 known bacterial species, 99.9% of which are obligate anaerobes (Moore and Holdeman, 1974). Of the more than 400 species inhabiting the GIT, however, 99% of the indigenous microflora is composed of 30 to 40 bacterial species (Moore and Holdeman, 1974), and these bacteria, along with the remaining 1% of other species, occupy all available niches or habitats and establish a stable ecosystem within the GIT (Berg, 1996). It is likely that the turkey intestinal tract harbors similar numbers of bacterial species.
The distribution of indigenous microflora within the GIT is not random, but organized qualitatively and quantitatively along vertical and horizontal regions in the GIT (Berg, 1996). The vertical distribution refers to the distribution of bacteria from the oral cavity to the colon, and concentrations of bacteria are vastly differently among different sections of the GIT. Furthermore, bacteria are distributed horizontally along the GIT as well, and occupy the intestinal lumen, the loose and tightly adherent mucus lining, crypt spaces, and adhere to the epithelial cells (Rozee et al., 1982). Thus, each segment and horizontal layer of the GIT harbors its own specific bacterial community (van der Wielen et al., 2002).
In birds, the largest concentrations of bacterial populations are found in the distal ileum, ceca, and colon, although bacterial populations are also found in the crop, proventriculus, and gizzard as well. Bacterial populations within the crop are dominated by the lactobacilli species, and may contain low concentrations of other organisms such as Clostridium perfringens, micrococci, staphylococci, and yeast (Sarra et al., 1985; Mead, 1997). The proventriculus and gizzard are relatively inhospitable to bacteria due to a low pH and rapid transit of feed and thus only a few species of bacteria are located within these organs (Mead, 1997). Among those bacteria reported to be found in the gizzard and proventriculus are Lactobacillus as well as low populations of E. coli, enterococci, and yeasts (Smith, 1965).
The duodenal and jejunal sections of the intestine contain relatively low microbial densities which increase considerably at the distal ileum, ceca, and colon to densities of 1011 bacteria per gram (wet weight) of cecal content (Savage, 1977; Mead, 1997). Analysis of the ileum of 6-week-old broiler chickens yielded an average count of 108 to 109 bacterial cells per gram of ileal digesta and 1011 bacterial cells per gram of ileal mucosa, with 95% of the bacterial cells identified as gram positive (Gong et al., 2002). In this study, Lactobacilli, E. cecorum, and butyrate-producing bacteria were the three major bacterial groups that were detected and identified in the ileum and ceca. However, it appears that the ileum and ceca contain different populations, as Zhu et al. (2002) analyzed broiler chicken cecal content and mucosa and determined that only four phylogenetic groups, Clostridium leptum, Sporomusa sp., Clostridium coccoides, and the enteric bacteria group, compromised 89% of all bacterial sequences isolated from the ceca. The remaining 11% of bacterial species were represented by the Atopobium group (3.6%), the Bacillus-Lactobacillus-Streptococcus subdivision (1.5%), Bacteroides group (1.9%), Actinomyces and relatives (1.3%), and Pseudomonas and relatives (0.7%).
GIT microbiota benefits to the bird
The resident intestinal microbiota provide a wide range of benefits to the host. Generally, the most beneficial and perhaps important role of the microbiota is colonization resistance, or the ability of the microbiota to inhibit the colonization of the gut by exogenous pathogenic bacteria (Berg, 1996). Inhibition of potential pathogens by the indigenous microbiota is achieved by numerous mechanisms, including competition for epithelial attachment sites and nutrients, and the production of bacteriocins, short chain fatty acids, and modified bile acids that are inhibitory to pathogens (Rolfe, 1997).
The indigenous microbiota also provide other benefits to the intestine. Specific short chain fatty acids produced by the fermentation processes of the microbiota serve as a major energy source to enterocytes in the colon (Guarner and Malagelada, 2003) and as stimulators of epithelial cell proliferation in the small and large intestine (Frankel et al., 1994.), suggesting a role in maintaining the intestinal integrity that is so critical to preventing disease. Additionally, the indigenous microbiota is capable of converting dietary precarcinogens and carcinogens into noncarcinogenic compounds (Berg, 1996). Lastly, indigenous microbiota are known to synthesize vitamins and to increase the absorption of calcium, magnesium, and iron (Hill, 1997; Miyazawa et al., 1996).
However, the role of bacteria in intestinal development should not be overlooked. The intestinal microbiota take part in bacteria-host cross-talk that allows for increased colonization of "beneficial" bacteria such as Bacteroides thetaiotamicron in the developing intestine. Briefly, B. thetaiotamicron induces the production of host-derived fucosylated glycans by secreting a bacterial signal to the host cell which prompts it to produce the fucosylated glycans that B. thetaiotamicron thrives on (Bry et al., 1996). It was noted that prior to weaning, the intestinal glycans of mice terminated predominantly in sialic acid, but after weaning and the subsequent introduction of B. thetaiotamicron, most glycans terminated in fucose. Hence, a mutually beneficial relationship is established for both the bacteria and the developing intestinal tract of the host; the B. thetaiotamicron flourishes while the epithelial attachment site of the host is protected by the presence of an indigenous organism as opposed to a pathogenic organism (Bry et al., 1996; Hooper et al., 2002). Interestingly, germfree mice do not develop fucosylated glycans and hence do not receive the same benefits that their conventional counterparts enjoy (Bry et al., 1996).
Intestinal microbiota also contribute to intestinal angiogenesis, or the vascular development, of developing intestinal tracts. In 2002, Stappenbeck and coworkers noted that the development of the mouse's intestinal vascular network corresponded with the development of a complex microbiota within the GIT. Further research revealed that germ-free mice had arrested intestinal villi capillary development when compared with their conventional counterparts, and that a 10 day colonization of the germ-free mice with B. thetaiotamicron significantly improved the capillary development within the intestine. The data suggests that microbes are regulating vascular development via bacteria-host cross-talk with the Paneth cells of the host. The benefits of this particular relationship are apparent; improved vascularization increases the intestine's absorptive capabilities and thus intestinal development, which may translate into improved growth rates.
Interestingly, bacteria also stimulate immune development of the intestine. Rhee et al. (2004) determined that proper development of the intestinal immune development is dependent upon the presence of intestinal microbiota. Colonization of the intestine increases circulating levels of antibodies such as IgA (Cebra, 1999), which aids the intestinal immune system in defending against pathogenic bacteria such as Salmonella or E. coli.
Barrier functions of the GIT
As previously indicated, intestinal microbiota contribute significantly to intestinal development and health, but must also frequently interact with the immune system as well as the gut barrier. In that respect, the intestine is an organ that must meet two seemingly incompatible goals: to maximize nutrient uptake and to minimize antigenic insult while tolerating the presence of indigenous microbiota and other antigens introduced by the presence of feed within the intestinal tract. Both of these functions require a number of multifaceted interactions between numerous physiological systems such as the physical gut barrier and the gut immune system and the indigenous microbiota. The synchronization of these systems is crucial to maintaining nutrient uptake, utilization, and thus animal growth as well as defending against constantly adapting bacteria and other antigenic insults.
Among the first line of non-immune defenses associated with gut barrier function is the mucus layer above the epithelial cells, which contains two sections; a loosely attached and a tightly adherent layer of mucus. Both layers are known to accumulate bacteriocidal and bacteriostatic compounds, and secretory immunoglobulin A, compounds which are capable of neutralizing or killing bacteria. The loosely attached mucus layer traps and therefore neutralizes potential pathogens and then removes the pathogens as the mucus layer is flushed from the intestinal tract (Nuetra and Forstner, 1987). The tightly adherent layer serves to lubricate and protect the epithelial tissue and constitutes a diffusion barrier as well (Farhadi et al., 2003).
The intestinal epithelium itself also contributes to gut barrier function. Apart from being a physical barrier between the lumen of the intestine and the interstitial space, the epithelium is also composed of numerous differentiated cells that migrate to the tip of the intestinal villi and then slough off into the lumen of the intestine. Additionally, in vitro research has indicated that a Salmonella or Shigella infection of cultured human intestinal cells stimulates cell apoptosis, though it is unclear if the same phenomenon occurs in vivo (Kim et al., 1998). Researchers believe that the continual migration, apoptosis, and renewal of these cells maintains a strong and healthy barrier against invasion and destruction (Gaskins, 1997).
One particular set of cells, the Paneth cells, reside within the crypts of the intestine and possess secretory granules containing high concentrations of antimicrobial peptides known as defensins, cryptidins, and angiogenins. When bacterial populations within the crypts become too concentrated, the Paneth cells discharge the granule contents into the lumen of the intestine (Ganz and Lehrer, 1994), thereby reducing bacterial concentrations near the stem cells and shaping the bacterial milieu surrounding the stem cells.
Immune functions of the GIT
Non-immunologic contributions to gut barrier function are very important, yet the immune system cannot be overlooked. Researchers have estimated that the intestinal tract contains greater than 70% of all immune cells found within the body and that nearly one-fourth of the intestinal mucosa is composed of lymphoid tissue (Kagnoff, 1993). Thus, the intestinal immune system has been given its own designation as GALT, or gastrointestinal-associated lymphoid tissue, and contributes greatly to gut barrier function on a day-to-day basis.
One of the primary components of the GALT is the lamina propria of the intestine, which is the connective tissue that underlies the epithelium of the gut and contains a sizeable population of immune cells such as macrophages, lymphocytes, immunoglobulins, and dendritic cells, among others. Highly vascularized and richly innervated by the enteric nervous system, the lamina propria (Gaskins, 1997) interacts with other components of the GALT such as the Peyer's patches, also one of the primary components of GALT, to maintain gut health and function. The Peyer's patches, which are aggregations of lymphoid follicles found primarily in the ileal epithelium, are located within the intestinal epithelium and thus are in contact with the digesta. In 12-weeks-old chickens, there are five or six Peyer's patches, though in adult chickens, only one patch can be consistently found (Schat and Meyers, 1991). The Peyer's patches contain M cells which continuously "sample" the contents of the intestine and transport them to the lamina propria for processing by the GALT.
Immune cells within the Peyer's patch and lamina propria are presented with the sampled bacteria and initiate a chain of events that help the immune system recognize and respond to the bacteria. Among these responses is the production of a secretory immunoglobulin A (IgA), an antibody that is specific to the bacteria it was presented with. Secretory IgA are secreted into the intestinal lumen and provide protection to the intestine by attaching to bacteria, neutralizing the bacteria and preventing the bacteria from adhering to and injuring or destroying epithelial cells. Furthermore, secretory IgA is also capable of killing the bacteria directly, or by presenting the bacteria to immune cells that will destroy the bacteria (Kagnoff, 1993). Large concentrations of secretory IgA exist within the lumen of the intestine and coat many of the bacteria within the lumen, thus contributing to maintaining gut barrier function. Additionally, the production of IgA within the intestine stimulates the production of memory B-cells, or cells that automatically recognize the specific bacteria in subsequent encounters. This recognition is advantageous to the animal because the immune systems is able to recognize and respond to the bacteria without the processing that was necessary during the first exposure, thus decreasing the response time (Kagnoff, 1993).
Maintaining gut barrier function and immunity is crucial to the growth and well being of the turkey. Because the intestine serves as a chief interface between the turkey's external and internal environment, any disruptions of the existing gut barrier function may lead to serious production problems that will ultimately affect growth and mortality rates. Problems that can lead to impaired gut barrier function include viral and bacterial infections, the presence of mycotoxins in feed, parasites, and fat rancidity (Hoerr, 2001). Stress should not be overlooked as a source of gut barrier disruption, either.
Stress responses within the GIT
Animals in a production setting face many different types of stressors that not only influence both the health and the productivity of a flock but also disrupt gut barrier function and cause hormonal changes within the birds. Upon the introduction of stress, two pathways are initiated in order to help the animal cope with the stress. The first pathway, known as the sympathetic adrenal medullary system (SAMS) is activated immediately after stressful events begin, and causes the release of catecholamines such as epinephrine and norepinephrine. These catecholamines prepare the bird for the classic "fight or flight" response of increased heart rate and blood pressure, while also stimulating the hypothalamus pituitary adrenal axis (HPA), a system that will aid the bird in adapting to long-term stressors by stimulating the release of corticotrophin releasing hormone from the hypothalamus. The corticotrophin releasing hormone acts upon the anterior pituitary gland by stimulating the release of adrenocorticotropin hormone, which in turn stimulates the adrenal cortical cells to secrete corticosterone. Corticosterone orchestrate alternate physiological patterns, mobilize energy reserves, and suppress processes that are not necessary to immediate survival (Henry, 1992).
The ultimate effect of catecholamines and corticosterone is detrimental to bird growth and productivity, as these hormones increases GIT transit time (Tur et al., 1989), and simultaneously cause decreases in growth and body weight (Saadoun et al., 1987) due to increased muscle breakdown and abdominal fat deposition (Hayashi et al. 1994). Adding insult to injury, however, is the fact that corticosterone also suppresses the immune system (Padgett and Glaser, 2003) by decreasing levels of antibodies such as IgA (Gross, 1992) and thus increasing susceptibility to viral infections (Gross et al., 1980) and coccidiosis (Isobe and Lillehoj, 1993). Furthermore, research by Rahman et al. (2000) indicated a ten-fold increase in the growth of Salmonella typhimurium when cultured with the catecholamines norepinephrine and epinephrine. The presence of catecholamines within the intestine also causes a significant increase in gram negative bacteria in the intestines of mice (Lyte and Bailey, 1997), indicating possible disruptions of the indigenous microbiota and gut barrier function of the intestine as well as potential for pathogen colonization.
Many different situations generate the stress response, such as temperature fluctuations (Davis and Siopes, 1987), social interactions (Pesti and Howarth, 1983), feed deprivation (Beuving et al., 1986), transport, and restraint (Freeman and Manning, 1976), and dietary changes. These stressors cause changes within the indigenous microbiota as well as in gut barrier function and ultimately influence growth and well being. Unfortunately, however, it is impossible to avoid all stressors within a production setting. Thus, the goal should be to minimize stressors as much as possible in order to minimize insult to the gastrointestinal tract and thus avoid growth depression.
Studies on nutrient uptake realized in meat yield & GIT health
Different investigators have explored the functionality of different nutrient transporters. One such example is the uptake of betaine by the small intestine (Kettunen et al., 2001). These researchers explored the uptake mechanisms of betaine, and noted that uptake of betaine in the duodenum and jejunum requires both a Na-dependant and Na-independent component. In comparing uptake, body compartmentalization, and usage as a methyl donor of betaine, methionine, and choline, Ketunen et al. (2001) noted that betaine had a more rapid uptake into plasma and was more soluble than that of methionine and choline (which were more closely associated with plasma lipoproteins). Once these methyl donors are delivered to muscle, choline and methionine have a much greater association with the lipid fraction of the tissue, whereas betaine has a much higher association with the water fraction. This association, therefore, can be related to reports of betaine aiding in osmoregulation during coccidial challenges (Augustine et al., 1997; Hess et al., 1998; Matthews and Southern, 1998).
Stangeland et al. (2000), in taking this partitioning information to somewhat of a more practical level, investigated the impact of feeding two pounds/ton of betaine on growth and breast meat yield from 5 to 20 weeks of age.
These researchers noted that there were no significant differences in growth or feed conversion between an NRC fed control group vs. those fed an NRC diet + 2 lb betaine/ton. Toms fed betaine, however, yielded an additional ½ pound of breast meat. This additional 6 to 8 cent investment in feed costs (from 5 to 20 weeks of age), therefore, yielded an additional 86 cents in breast meat.
In conclusion, knowledge of how the intestinal tract develops in concert with the microbiota in the GIT will greatly aid our ability to formulate diets and provide compounds and environments that provide for optimization of a synergistic relationship between them.
References are available on request
From Proceedings of the "Midwest Poultry Federation Convention", St. Paul, Minnesota, U.S.A.
