TY - JOUR
AU - Metges, C. C.
AB - Abstract The porcine model has been utilized by researchers to study human developmental aspects, surgical procedures, disease impacts, and nutritional interventions for over 60 yr. However, when it comes to investigating how nutrients regulate human gastrointestinal tract (GIT) function and health, and nutritional research evolves from dietary composition to potential nutraceutical applications, the question needs to be asked: is the pig an appropriate translational animal model? Morphologically and functionally, the porcine GIT has more similarity to the human GIT than commonly used models such as rats. Food intake related to BW is much lower in humans than in pigs, which explains the higher and faster growth and earlier maturation in the latter species. In addition, the digestion and metabolism of carbohydrates, prebiotics, lipids, and amino acids produce similar metabolites and subsequent effects on GIT development and health in pigs and humans, making the pig a more comparable model to study human GIT issues than other nonprimate species. However, there are key divergences. Pigs have a functional cecum, larger colon, and the endogenous microbiota of the GIT, while similar at the phylum level is vastly different at the genera level. One notable microbial species that is low to nonexistent in the porcine GIT are the Bifidobacterium, which is a major probiotic species in humans. Several porcine models have been developed to overcome these issues, but they are new and have yet to be fully tested. This paper will provide a comparative analysis between the human and porcine GIT in regards to structure, function, and health and how various nutritional factors influence these characteristics and future developments of the porcine model. INTRODUCTION Human nutrition research investigates how foods we consume influence our growth, development, and health. While some studies can be conducted using human subjects, the majority cannot because of ethical considerations, and so the use of an appropriate translational animal model is required. As a translational model any species used should be closely related to humans in terms of genetics, anatomy, and physiology, so that any results can be directly applicable to humans (Prabhakar, 2012). For more than 60 yr, the domestic pig (Sus scrofa domestica) has been utilized by researchers as a translational model for a wide range of topics including the digestive system and its primary organ the gastrointestinal tract (GIT; Guilloteau et al., 2010; Sangild et al., 2013). Anatomically and physiologically, the porcine GIT is highly similar to that of humans (Guilloteau et al., 2010). Also, porcine GIT functionality, e.g., how nutrients and metabolites produced by digestion or metabolism by resident microbiota are absorbed by the GIT is closer to humans than any other nonprimate model (Brugger et al., 2010). However, there are functional differences between the porcine and human GIT that affect transit time, site of digestion, and nutrient/metabolite production and absorption. Also, large differences exist in growth rate and food intake. The aim of this review was to explore the contemporary scientific literature how carbohydrates, lipids, and amino acids affect GIT development, function, and health in pigs and where comparable literature was available in humans. We conclude with a brief discussion on the limitations of the use of on-farm pig breeds as models for nutrition and health research. Species Comparison Feeding Behavior and Growth. Rats and pigs are 2 of the most commonly utilized animal models for human nutrition research; however, there are differences in feeding behavior and growth that need to be considered when using these 2 model species. At an early age the divergence in these behaviors between species is evident; intake by suckling piglets is ∼0.4 kg colostrum/d (∼1.275 MJ/BWkg0.75) in the first 2 d and 1.1–1.6 kg (∼0.675 MJ/BWkg0.75) milk/d later on, depending on litter size (Quesnel et al., 2015). Milk intake by breastfed human babies aged 3 to 12 mo ranges between 0.4 and 0.9 kg milk/d or 0.45–0.5 MJ/BWkg0.75 (Heinig et al., 1993), while energy intake from solid food increases from 0.39 to 2.18 MJ/d or 0.26–0.29 MJ/BWkg0.75. Weanling pigs (6–15 kg) consume 15–16 meals (0600 to 2400 h) of about 30 g each and eat 0.3–0.5 g/min (Ermer et al., 1994) corresponding to 0.8 to 1.2 MJ/BWkg0.75 (NRC, 1998), while weanling rats consume ∼1 MJ/BWkg0.75 (NRC, 1995). In children between 6 mo and 1 yr, 1–4 yr, and 4–5 yr energy intake increase from 3.6 to 5.9 MJ/d (Deutsche Gesellschaft für Ernährung, 2008). The transition from milk to solid food contrasts in humans vs. commercial piglets and rat pups, which are removed abruptly from the sow/dam between ages 21 to 42 d and in laboratory rats between 18 and 22 d (Puiman and Stoll, 2008; Sangild et al., 2013). Although one has to keep in mind that feeding patterns and dietary intake depend on the social environment, diet type, and diet presentation, from this data it is evident that food intake related to BW is much lower in humans than in pigs, which explains the higher and faster growth and earlier maturation in the latter species. Rats are nocturnal eaters with maximal food intake at the beginning and end of the dark phase and at minimal during the light phase, with food ingestion clustered into bouts (Clifton, 2000). Humans and pigs are meal eaters; with growing pigs (30–90 kg) consuming 8–11 meals of about 270–300 g, each over 1 d where they eat about 20–30 g food/min (Gloaguen et al., 2013; Hyun and Ellis, 2002). In addition, rats practice coprophagy and can ingest over 40% of their fecal output whereby essential nutrients are supplied (Graham and Aman, 1987). Coprophagy can also occur in piglets but to a lesser extent (Soave and Brand, 1991). Fetal pig growth rate was reported to increase from 7.7 to 36 g/d from gestation d 45 to d 110 (McPherson et al., 2004). In the first 4 wk after weaning, piglets increase their BW by 130–250 g/d (Kecman et al., 2014; Mitchell et al., 2012; Montagne et al., 2007), whereas in the first year of life a pig grows with an average rate of approx. 400–500 g/d. This contrasts greatly with humans where BW gain from birth to 3 mo was ∼ 28 g/d and decreased to 8 g/d between 12 and 15 mo, whereas the average daily gain over the first year amounts to ∼16 g/d (Heinig et al., 1993). GIT Morphology and Function. Rats and pigs are single stomached (Table 1), hindgut fermenters, and feed on plants and animal alike but at different proportions. Rats are born largely immature, and the gut matures gradually during suckling but more rapid during the weaning phase (Puiman and Stoll, 2008; Sangild et al., 2013). In pigs the proportion of the GIT to total fetal weight increases during the last 2/3 of the gestational period from 2.5% to 6.2% indicating that growth of GIT accelerated during late gestation (McPherson et al., 2004). In piglets small and large intestine weight increase by 50% to 70% in the first 2 wk after weaning (Montagne et al., 2007). Age of maturity depends on the speed of the transition from liquid to solid food. The GIT matures in contemporary pigs shortly after weaning while in humans at the age of 6–12 mo (Guilloteau et al., 2010; Sangild et al., 2013) allowing the use of neonate and adolescent pigs as models for infant and adult human GIT and thus avoid to work with adult-size pigs (200–250 kg BW). Table 1. Geometry and pH of the gastrointestinal tract (GIT) of adult pigs, rats, and humans1   Pig   Rat   Human     Geometry  pH  Geometry  pH2  Geometry  pH  Stomach3  4  3–5  0.002  3.1–5.1  1  1.5–3.5  Duodenum  0.8  6–8  0.1  6.9  0.38  6  Jejunum  19  6–8  1.35  7.4  2.5  7–9  Ileum  0.5  6–8  0.004  7.4  4  5.6–6.8  Cecum  0.24  6–7  0.007  6.4-6.84  0.1–0.34  5.65  Colon  4  6–8  0.011  6.8  1.5  6–7    Pig   Rat   Human     Geometry  pH  Geometry  pH2  Geometry  pH  Stomach3  4  3–5  0.002  3.1–5.1  1  1.5–3.5  Duodenum  0.8  6–8  0.1  6.9  0.38  6  Jejunum  19  6–8  1.35  7.4  2.5  7–9  Ileum  0.5  6–8  0.004  7.4  4  5.6–6.8  Cecum  0.24  6–7  0.007  6.4-6.84  0.1–0.34  5.65  Colon  4  6–8  0.011  6.8  1.5  6–7  1Images of the GIT used with permission from Stevens and Hume (1998). 2 Ward and Coates (1987). 3Stomach geometry is presented as volume (L), while the geometry of the remaining sections of the GIT is presented as length (m). 4 Kararli (1995) 5 Cummings et al. (1987) View Large Table 1. Geometry and pH of the gastrointestinal tract (GIT) of adult pigs, rats, and humans1   Pig   Rat   Human     Geometry  pH  Geometry  pH2  Geometry  pH  Stomach3  4  3–5  0.002  3.1–5.1  1  1.5–3.5  Duodenum  0.8  6–8  0.1  6.9  0.38  6  Jejunum  19  6–8  1.35  7.4  2.5  7–9  Ileum  0.5  6–8  0.004  7.4  4  5.6–6.8  Cecum  0.24  6–7  0.007  6.4-6.84  0.1–0.34  5.65  Colon  4  6–8  0.011  6.8  1.5  6–7    Pig   Rat   Human     Geometry  pH  Geometry  pH2  Geometry  pH  Stomach3  4  3–5  0.002  3.1–5.1  1  1.5–3.5  Duodenum  0.8  6–8  0.1  6.9  0.38  6  Jejunum  19  6–8  1.35  7.4  2.5  7–9  Ileum  0.5  6–8  0.004  7.4  4  5.6–6.8  Cecum  0.24  6–7  0.007  6.4-6.84  0.1–0.34  5.65  Colon  4  6–8  0.011  6.8  1.5  6–7  1Images of the GIT used with permission from Stevens and Hume (1998). 2 Ward and Coates (1987). 3Stomach geometry is presented as volume (L), while the geometry of the remaining sections of the GIT is presented as length (m). 4 Kararli (1995) 5 Cummings et al. (1987) View Large The proximal rat stomach entry area from the esophagus (fundus) is a reservoir for food and site of microbial digestion separated from the glandular part by the limiting ridge (DeSesso and Jacobson, 2001). In the pig the esophageal region of the stomach is nonglandular and an extension of the esophagus, whereas the human stomach is entirely secretory. The small intestine of the pig (fully grown: 16–21 m; 5% duodenum, 90% jejunum, 5% ileum) is about 11–14 times its body length, which is much longer than in rats and humans (Yen, 2001). In adult humans small intestinal lengths range between 4 and 7 m, whereas the large intestine measures between 1.1 to 1.8 m (DeSesso and Jacobson, 2001). In rats compared to humans the proportional small intestinal length (ca. 80% of total GIT) as well as that of the large intestine (18% of total GIT) is similar (DeSesso and Jacobson, 2001). Almost 90% of the length of the small intestine of the rat is jejunum, whereas it is only 38% in humans (DeSesso and Jacobson, 2001). Also, the small intestine of humans and pigs possess plica circularis (Kerckring's folds), which increases the absorptive area, but rats do not (DeSesso and Jacobson, 2001). Thus, the human and pig GIT are capable of absorbing materials faster than that of the rat (DeSesso and Jacobson, 2001). Cecum and varying lengths of the colon are haustrated in pigs and humans but not in rats. The upper digestive tract anatomy from mouth to ileum is similar between rats and humans, but there are major differences in size and architecture of the hindgut (Deglaire and Moughan, 2012; DeSesso and Jacobson, 2001). In the porcine large intestine the cecum, ascending and transverse colon, and proximal portion of the descending colon are arranged in a series of centrifugal and centripetal coils that are structurally different from those of humans (Meurens et al., 2012). In humans, the cecum is poorly defined as compared to rats and pigs and is continued into the colon (Table 1). Humans have smaller colons; as a percentage of the GIT, the colon is > 45% in pigs and only 20% in humans (Miller and Ullrey, 1987). Thus, relative large-intestinal contents in humans show the lowest values compared to rats and pigs (0.3 vs. 2 vs. 5% wet weight/kg BW; Cummings et al., 1987; Rechkemmer et al., 1988). This is somewhat reflected in the estimated contribution of short-chain fatty acid (SCFA) to basal metabolic requirements which ranges from 30% to 76% in pigs, whereas in rats and humans it amounts to 5%–9% (Bugaut, 1987). Pigs and humans are predominantly colon fermenters with cecum fermentation also prevalent but presumably to a lower extent than in the colon because the pig and human cecum makes only 10%–20% of the colon length (Kararli, 1995; Yen, 2001). In contrast, the rat is a cecum fermenter with a cecum length corresponding to 25%–39% of the colon length, and size of cecum differs depending on the diet (DeSesso and Jacobson, 2001; Xiao et al., 2015). This is in line with the observation that the concentration of SCFA in caecal fluids amounts to 170 mmol/L in rats as compared to 90 mmol/L in pigs (Rechkemmer et al., 1988). Thus, the pig stands between rodents and humans in terms of the importance and contribution of cecal fermentation to nutrient utilization. Interestingly, concentrations of total SCFA in jejunum, cecum, and colon (human vs. pig: 1 vs. 4–5; 131 vs. 89–114; 80–123 vs. 96–115 mmol/kg wet weight) are comparable between pigs and humans (Cummings et al., 1987; Loh et al., 2006). Also comparable SCFA patterns in portal and hepatic vein are reported for pigs and humans, albeit with a seemingly greater absolute concentrations in pigs (Cummings et al., 1987; Ingerslev et al., 2014). The digesta transit time is quite comparable between humans and pigs and is altered in a similar manner in response to dietary fiber content, namely passage rate is reduced when fiber content is high. In humans, similar to pigs and rats, chyme traverses the small intestine in 3–4 h (Davis et al., 2001; DeSesso and Jacobson, 2001). The passage time through the large intestine is much longer in all 3 species but varies considerably among species depending on the type of food. Thus, passage through the large intestine was reported to range between 0.5 to 3 d. The average large intestinal transit time in humans was found to be between 28 and 43.5 h, whereas in pigs mean colonic transit time was observed to be 23.5 and 56 h (Hendriks et al., 2012). GIT Microbiota. At birth the GIT is sterile and microbial colonization occurs during and immediately after birth, as the vaginal, maternal, and environmental microbiota are taken up by the neonate (Benno and Mitsuoka, 1986). In humans, the bifidobacteria are the predominant bacterial group found in infant feces within the first 2 wk, followed by clostridia, enterobacteria, streptococci, and staphylococci, while in pig's lactobacilli, streptococci, enterobacteria, bacteroides, and clostridia become the dominant fecal microorganisms (Benno and Mitsuoka, 1986; Heinritz et al., 2013; Wang and Donovan, 2015). The nascent microbial population then remains relatively stable during the suckling period, as milk is the sole nutrient source and a complex evolutionary relationship that has developed between the microbiota and milk oligosaccharides, which are digestible by the microbiota but not the host (German et al., 2008). There are significant differences between human and porcine milk oligosaccharide content (German et al., 2008; Tao et al., 2010) (Table 2) which contribute to the maintenance of the divergent GIT microbial populations between the species as shown for fecal gut microbiota of human and porcine neonates (Wang and Donovan, 2015). However, at the phyla level the GIT microbiota of pigs, rats, and humans is broadly similar, consisting mainly of the Firmicutes and Bacteroidetes (Miller and Ullrey, 1987; Tomas et al., 2012). One of the most fundamental contributions that members of the Firmicutes and Bacteroidetes phyla makes to support GIT development and health is the production of SCFA. In humans and pigs 10% to 30% of expended energy is contributed by the oxidation of SCFA (Bergman, 1990). Chief among the released SCFA are acetate, propionate, and butyrate that act as the main energy sources for host epithelial cells of the large intestine and terminal ileum and play potential roles in regulating intestinal health (Govers et al., 1999). However, pigs, rats, and humans have distinct GIT genera within each phylum that can lead to differences in SCFA production, not accounted for by diet and GIT morphological differences (Furet et al., 2009; Heinritz et al., 2013; Tomas et al., 2012). In addition, the human probiotic Bifidobacterium spp. are scarce or not detectable in the GIT of pigs and rats, whereas in humans they constitute about 4% of the GIT microbiota (Heinritz et al., 2013; Loh et al., 2006). To align the porcine and rat GIT models closer to that of the human, researchers have investigated the use of gnotobiotic human-microbiota-associated (HMA) animals (Pang et al., 2007; Tomas et al., 2012; Zhang et al., 2013a) and cloned pigs. Molecular profiling has shown that HMA piglets and rats have GIT microbial populations similar to those of the donor, and established Bifidobacterium spp. populations, indicating an advantage over traditional models (Pang et al., 2007; Wang and Donovan, 2015). Thus, in terms of digestive tract anatomy, microbiota content, digesta transit time, relative absorptive area, and nutrient digestibility the pig is a more comparable model to study human GIT issues than rat. Table 2. Colostrum and milk yield and composition in rats, pigs and humans (weight %)     Yield1  Water  Energy2  Protein  Fat  Lactose  OS3  References  Rat  Colostrum  –  –  –  8.9  14.7  2.5  –  (Keen et al., 1981)    Milk  0.05–0.08  58.4–75  –  7.8–13  8–17  1.7–3.7  –  (Keen et al., 1981; Kliewer and Rasmussen, 1987; Wattez et al., 2015; Young and Rasmussen, 1985)  Pig  Colostrum  3.5  73–79  5.7–6.7  7.7–16.9  5.4–8  2.0–4.1  –  (Beyer et al., 2007; Declerck et al., 2015; Quesnel et al., 2015; Rehfeldt et al., 2011; Hurley, 2015)    Milk  9.4  78–82  4.4–6.5  5–7.5  7–10.1  4.3–5.6  –  (Beyer et al., 2007; Le et al., 1998; Quesnel et al., 2015)  Human  Colostrum  0.6  –  –  2.3  2.1  4.2  1.8–3.5  (Kunz et al., 1999)    Milk  0.8  87.6  2.6–3.3  0.8-1.3  2–3.5  7.5–8  0.9–2.7  (Heinig et al., 1993; Jans et al., 2015; Kunz et al., 1999; Stam et al., 2013; Wojcik et al., 2009)      Yield1  Water  Energy2  Protein  Fat  Lactose  OS3  References  Rat  Colostrum  –  –  –  8.9  14.7  2.5  –  (Keen et al., 1981)    Milk  0.05–0.08  58.4–75  –  7.8–13  8–17  1.7–3.7  –  (Keen et al., 1981; Kliewer and Rasmussen, 1987; Wattez et al., 2015; Young and Rasmussen, 1985)  Pig  Colostrum  3.5  73–79  5.7–6.7  7.7–16.9  5.4–8  2.0–4.1  –  (Beyer et al., 2007; Declerck et al., 2015; Quesnel et al., 2015; Rehfeldt et al., 2011; Hurley, 2015)    Milk  9.4  78–82  4.4–6.5  5–7.5  7–10.1  4.3–5.6  –  (Beyer et al., 2007; Le et al., 1998; Quesnel et al., 2015)  Human  Colostrum  0.6  –  –  2.3  2.1  4.2  1.8–3.5  (Kunz et al., 1999)    Milk  0.8  87.6  2.6–3.3  0.8-1.3  2–3.5  7.5–8  0.9–2.7  (Heinig et al., 1993; Jans et al., 2015; Kunz et al., 1999; Stam et al., 2013; Wojcik et al., 2009)  1kg/d. 2kJ/g; Gross energy; Pig: lactation d 2–29 (lower values during the last 2 wk of lactation); Human: lactation mo 3 to 12. 3Oligosaccharides (non-Lactose). View Large Table 2. Colostrum and milk yield and composition in rats, pigs and humans (weight %)     Yield1  Water  Energy2  Protein  Fat  Lactose  OS3  References  Rat  Colostrum  –  –  –  8.9  14.7  2.5  –  (Keen et al., 1981)    Milk  0.05–0.08  58.4–75  –  7.8–13  8–17  1.7–3.7  –  (Keen et al., 1981; Kliewer and Rasmussen, 1987; Wattez et al., 2015; Young and Rasmussen, 1985)  Pig  Colostrum  3.5  73–79  5.7–6.7  7.7–16.9  5.4–8  2.0–4.1  –  (Beyer et al., 2007; Declerck et al., 2015; Quesnel et al., 2015; Rehfeldt et al., 2011; Hurley, 2015)    Milk  9.4  78–82  4.4–6.5  5–7.5  7–10.1  4.3–5.6  –  (Beyer et al., 2007; Le et al., 1998; Quesnel et al., 2015)  Human  Colostrum  0.6  –  –  2.3  2.1  4.2  1.8–3.5  (Kunz et al., 1999)    Milk  0.8  87.6  2.6–3.3  0.8-1.3  2–3.5  7.5–8  0.9–2.7  (Heinig et al., 1993; Jans et al., 2015; Kunz et al., 1999; Stam et al., 2013; Wojcik et al., 2009)      Yield1  Water  Energy2  Protein  Fat  Lactose  OS3  References  Rat  Colostrum  –  –  –  8.9  14.7  2.5  –  (Keen et al., 1981)    Milk  0.05–0.08  58.4–75  –  7.8–13  8–17  1.7–3.7  –  (Keen et al., 1981; Kliewer and Rasmussen, 1987; Wattez et al., 2015; Young and Rasmussen, 1985)  Pig  Colostrum  3.5  73–79  5.7–6.7  7.7–16.9  5.4–8  2.0–4.1  –  (Beyer et al., 2007; Declerck et al., 2015; Quesnel et al., 2015; Rehfeldt et al., 2011; Hurley, 2015)    Milk  9.4  78–82  4.4–6.5  5–7.5  7–10.1  4.3–5.6  –  (Beyer et al., 2007; Le et al., 1998; Quesnel et al., 2015)  Human  Colostrum  0.6  –  –  2.3  2.1  4.2  1.8–3.5  (Kunz et al., 1999)    Milk  0.8  87.6  2.6–3.3  0.8-1.3  2–3.5  7.5–8  0.9–2.7  (Heinig et al., 1993; Jans et al., 2015; Kunz et al., 1999; Stam et al., 2013; Wojcik et al., 2009)  1kg/d. 2kJ/g; Gross energy; Pig: lactation d 2–29 (lower values during the last 2 wk of lactation); Human: lactation mo 3 to 12. 3Oligosaccharides (non-Lactose). View Large Nutrients: Interaction and Regulation of GIT Development and Health Carbohydrates. The most well-studied group of carbohydrates in human and porcine nutrition studies are the nondigestible carbohydrates (NDC) as results indicate that they can directly and/or indirectly modify GIT health and function (Anguita et al., 2007). Direct effects include modification of GIT viscosity, water retention capacity, and the rate of digesta passage (Anguita et al., 2007). However, the majority of studies have focused on the indirect roles of NDC, which are mediated by the fermentation of NDC by resident microbiota leading to SCFA production (Aumiller et al., 2015). In humans, NDC fermentation occurs primarily in the colon (inulin and oligofructose), while in pigs this occurs precaecal starting already in the stomach (Branner et al., 2004; Eberhard et al., 2007). However, changes in dietary NDC composition (Bird et al., 2007) can lead to shifts in the site of digestion in both species (Govers et al., 1999; Jonathan et al., 2013). This leads to changes in the resident microbiota, either by increasing the number or diversity of bacteria responsible for the production of SCFA (Bird et al., 2007) and in some cases reducing the numbers of potentially pathogenic microorganisms (Haenen et al., 2013; Petkevicius et al., 2007). The mechanisms behind the reduction in pathogenic microorganisms are not clear, but it may be a function of altered pH, antibacterial or antiparasitic effects of SCFA, or the potential bacterial binding properties of some NDC (Hopwood et al., 2004; Petkevicius et al., 2004). As the number and diversity of SCFA producing bacteria increases so do the levels of SCFA measured in different regions of the colon (Haenen et al., 2013) and in feces (Muir et al., 2004). As SCFA are the primary fuel of colonic epithelial cells, it is proposed that increased SCFA concentrations are also responsible for the observed increase in colon length via stimulation of epithelial cell growth (Bird et al., 2007). In addition, increased butyrate produced in response to NDC has been associated with reduced colonic epithelial apoptosis (Mentschel and Claus, 2003) and elevated expression of the SCFA transporter monocarboxylate transporter 1 in the cecum (Haenen et al., 2013). However, elevated levels of SCFA are not necessarily an indicator of improved intestinal health, as the SCFA produced by the resident microbiota need to be absorbed and utilized by the epithelial cells. Martin et al. (2000) have shown that the butyrate produced in response to different NDC is absorbed at different efficiencies by the colonic mucosa, thus providing potential health benefits. Overall, these changes mimic those observed in humans (Muir et al., 2004) and are proposed to aid in the prevention of colon cancer, as the distal colonic regions are where tumors most commonly occur (Flisikowska et al., 2012). A porcine model of colon cancer has been produced through gene-targeted mutations in the adenomatous polyposis coli gene that produces all the phenotypic characteristics observed in humans with familial adenomatous polyposis (Flisikowska et al., 2012). However, to date, no published studies investigating the effect of NDC or SCFA supplementations using this model are available. Prebiotics. Prebiotics are subset of NDC specifically utilized to stimulate the growth and metabolism of the Bifidobacteria, Lactobacilli, and Eubacteria spp. in the large intestine (Rastall and Gibson, 2015). This leads to changes in the profile of SCFA produced, decreased pH, changes in the digestion site of other nutrients, and the production of gases and other metabolites (ammonia, succinate, polyamines; Bruzzese et al., 2006; Lamichhane et al., 2014). These factors can be highly variable as they are inextricably linked to the host's microbiome, health status and the structure of the prebiotic. To date, the most-studied prebiotics are the fructo-oligosaccharides (FOS) and inulin, galacto-oligosaccharides (GOS), polydextrose (PDX), and human milk oligosaccharide (HMO). A large body of research on the use of prebiotics has focused on the neonatal and weaning periods (Jacobi and Odle, 2012). The use of prebiotics in neonatal studies focuses on replicating the protective effects associated with the oligosaccharide content of maternal milk. The oligosaccharide content of human milk is the highest of all mammals and is the third largest solid component after lactose and fat (Boehm and Stahl, 2007); however, the oligosaccharide content in pigs is not available quantitatively (Tao et al., 2010; Table 2). Infant formulas have a very low oligosaccharide content which has been linked to the observed differences between the microbiome and health indicators of formula- and milk-fed infants (Boehm and Stahl, 2007). Research in the use of supplemental prebiotics in neonatal piglets indicates that their presence can produce beneficial effects. Healthy piglets fed formula supplemented with PDX have increased ileal Lactobacilli counts, propionate and lactate concentrations, and decreased pH (Herfel et al., 2011). In addition, PDX supplementation reduces the expression of the pro-inflammatory cytokines TNFα, IL-1β, and IL-1 (Herfel et al., 2011). In disease models, HMO, short-chain GOS, and long-chain FOS have all been shown to reduce the duration of rotavirus-induced diarrhea in milk-fed piglets, but through potentially different mechanisms. Human milk oligosaccharides increased pH, the abundance of butyrate producing Lachospiraceae and modulated the immune response by enhancing the abundance of IFNγ and IL-10 mRNA, while the only observed effect of short-chain GOS and long-chain FOS was to promote the RV-specific immunoglobulin M response (Li et al., 2014). Preterm infants are at high risk of developing necrotizing enterocolitis, a potentially fatal disease where the epithelium of the intestine necrotizes (Barnes et al., 2012). After the removal of necrotic tissue, the remaining intestinal tissue often has inadequate surface area for digestion and absorption of nutrients, a state known as intestinal failure (IF). In a piglet model of IF, prebiotic treatment improved intestinal function by increasing ileal mucosa weight, ileal protein, and ileal villus length through increased epithelial cell proliferation. In addition, prebiotic supplementation upregulated intestinal peptide and glutamine transport (Barnes et al., 2012). Weaning from a highly digestible liquid to less digestible, more-complex solid diet is a stressful period for the weanling and can result in the occurrence of diarrheal diseases, which represent a serious health problem in humans and pigs alike. Increased susceptibility is associated with the loss of maternal protection factors (in maternal milk-fed neonates) and profound changes in the intestinal structure, metabolism, and microbiome. The addition of FOS to a transitional piglet diet reduced the number of coliform bacteria and increased the number of Bifidobacteria in the colon (Shim et al., 2005). Diets supplemented with FOS have also been shown to increase the prevalence of 2 beneficial Lactobacilli species in the ileum and colon and improve intestinal health and function by reducing intestinal pH and jejunum villus height when combined with lactose (Pierce et al., 2006). Oligosaccharide-supplemented diets of newly weaned pigs were shown to increase yeast concentration in the small and large intestines (Mikkelsen et al., 2003), butyric acid concentration in the large intestine, and improve growth, nutrient digestibility, and small intestinal morphology (Liu et al., 2008a). Pigs with acute cholera toxin initiated secretory diarrhea had more Lactobacilli in the small intestine and cecum, whereas densities of Enterobacteriaceae in the mucosa of the small intestine, cecum, and colon were significantly lower in FOS-supplemented animals compared with the nonsupplemented pigs (Oli et al., 1998). Similar results have also been reported in pigs with parasitic nematode infections (Petkevicius et al., 1997; Petkevicius et al., 2007). Inulin-supplemented diets have also been shown to lower the establishment of Oesophagostomum dentatum (Petkevicius et al., 1997), retard the growth (Thomsen et al., 2005), egg excretion, and female worm fecundity of Trichuris suis (Petkevicius et al., 2007). Lipids. Dietary lipids are essential for human and porcine growth and development as they are crucial components of biological membranes, involved in cell signaling, and act as substrates for various enzymes (Black and Rohwer-Nutter, 1991). Lipids contain fatty acids (saturated, monounsaturated, and polyunsaturated), and their derivatives, mono-, di-, and triglycerides (TG), phospholipids, and cholesterol. Their uptake by enterocytes is both passive and active (Iqbal and Hussain, 2009). Medium-chain triglycerides (MCT) and fatty acids (MCFA) have been the focus of numerous studies because porcine and human milk both have a low MCT/MCFA content, and they are small enough to passively diffuse into enterocytes, thus require no energy expenditure for uptake (Odle, 1997). Weanling piglets fed MCFA have been shown to have enhanced nutrient absorption, immune function compared with unsupplemented piglets (Dierick et al., 2004). Medium-chain fatty acids have also been shown to have antibacterial properties (Dierick et al., 2002). Increased release of MCFA into the GIT has been shown to reduce total microbial counts, Lactobacilli and E. coli (Dierick et al., 2002) and lactic acid bacteria counts in the stomach, cecum, and small intestine (Lai et al., 2015). Long-chain fatty acids have also been investigated for their ability to improve intestinal function and health. The most widely researched LCFA group are the omega 3-fatty acids; α-Linolenic acid (ALA), docosahexaenoic acid (DHA), and eicosapentaenoic acid (EPA), which are polyunsaturated fatty acids (PUFA) that have also been associated with a decreased risk of coronary heart disease, compared to saturated fatty acids. The long chain fatty acid ALA is found in plant oils while DHA and EPA are primarily found in marine oils. Supplementation of the maternal porcine diet during pregnancy and lactation, with linseed oil (rich in ALA), significantly changes the fatty acid composition, structure, and permeability factors of the neonatal ileum (Boudry et al., 2009; Hess et al., 2008). In a similar model investigating the effect of DHA-rich fish oil, researchers were able to show significant changes in the fatty acid profile of the weanling piglet jejunum that may support enhanced glucose uptake (Gabler et al., 2008). In contrast, neonatal piglets not exposed to PUFA in utero fed a milk-based formula with ALA or EPA had negligible effects on gross villus/crypt morphology, but still showed increased enrichment of the PUFA, ALA, and EPA in the jejunum and ileum, with the percentage ALA enrichment higher than EPA (Hess et al., 2008). Dietary PUFA intake has also been shown to influence intestinal endotoxin transport (Mani et al., 2013). Intestinal endotoxin is considered a predisposing factor for diseases such as obesity and diabetes. Diets rich in the PUFA, DHA, and EPA were shown to reduce endotoxin transport, while those high in saturated fatty acids increased endotoxin transport. The researchers suggest dietary fat intake may be associated with fatty acid regulation of intestinal membrane lipid raft mediated permeability (Mani et al., 2013). Alternative sources of PUFA have also been investigated for their role in supporting intestinal function and health. A study using malnourished neonatal piglets fed an adapted milk formula containing PUFA obtained and purified from pig brains, showed the intestine of supplemented piglets recovered more completely from the histologic lesions and biochemical alterations caused by malnutrition than piglets fed a control formula (Lopez-Pedrosa et al., 1999). The researchers observed increased intestinal cell growth, normalized jejunum lipid and fatty acid composition, and reduced histologic lesions caused by malnutrition. Elevated blood cholesterol (hypercholesterolemia) has been implicated in a wide range of human diseases. A pig model of hypercholesterolemia has shown that lupin protein isolate can lower cholesterol absorption by modifying the expression of intestinal genes required for uptake (Radtke et al., 2014). Amino Acids. Amino acids are not just precursors of proteins and other nitrogenous metabolites, they also regulate cell signaling pathways that control nutrient uptake and metabolism, DNA synthesis, immune function, tissue remodeling, hormone/mitogen secretion, and a vast array of additional functions in all tissues (Wu, 2014). The arginine family of AA have traditionally been overlooked in studies because of their relative abundance in the diet and being dispensable according to traditional AA classification (Wu, 2014). An extensively studied member of this family with regards to its effects on the GIT is Gln. Glutamine functions as a key respiratory fuel for all cells of the small intestine, surpassing glucose and fatty acids; contributes significantly to intestinal glutathione synthesis; and is an essential precursor of nucleic acids, nucleotides, amino sugars, and amino acids (Windmueller and Spaeth, 1978). Yet, Gln plays a significant role beyond the support of metabolic processes within the GIT. Experimental evidence shows that Gln supplementation prevents jejunal atrophy in postweaning piglets (Wu et al., 1996), reduces the abundance of mucosal stress marker corticotropin-releasing factor, and prevents weaning-induced decreases in tight junction proteins, markers of intestinal barrier permeability (Wang et al., 2015). Glutamine supplementation also improves GIT oxidative-defense capacity and promotes small intestine growth (Wang et al., 2008). In mature sow milk, Gln is the most abundant free amino acid (Table 3), whereas among protein-bound amino acids, Gln plus Glu form the highest group (Wu and Knabe, 1994). This finding coincides with the fact that maturation of the pig neonatal intestine requires high amounts of Gln as fuel and for the development of the gut-associated lymphoid tissue as well as a number of other physiological functions (Wu et al., 2011). It has been shown that dietary supplementation with Gln can improve whole body and intestinal growth and prevent intestinal oxidative injury and inflammatory disease in piglets (Haynes et al., 2009; Rezaei et al., 2013). Dysfunction of tight junction integrity, resulting in increased GIT permeability, is associated with poor recovery from surgery, serious medical conditions, and gastrointestinal diseases in humans and piglets (De-Souza and Greene, 2005). Research shows that oral Gln supplementation can protect the GIT of weaned piglets from E. coli-induced damage (Yi et al., 2005), suppress destructive inflammatory and regulatory cytokine responses, and decrease damage to tight junction proteins and intestinal electrolyte movement (Ewaschuk et al., 2011). Table 3. Amino acid composition of porcine and human milk, values are arranged from lowest to highest concentration1 Milk Free Amino Acids (µmol/L)  Milk Protein-Bound Amino Acids (mmol/L)  Human  Porcine  Human  Porcine  Amino Acid  Concentration  Amino Acid  Concentration  Amino Acid  Concentration  Amino Acid  Concentration  Methionine  10  Tryptophan  18  Methionine  Trace  Histidine  4  Isoleucine  14  Isoleucine  21  Histidine  2  Lysine  4  Phenylalanine  16  Methionine  23  Arginine  3  Arginine  6  Histidine  22  Phenyalanine  36  Phenylalanine  3  Glycine  7  Tyrosine  22  Leucine  45  Tyrosine  4  Phenyalanine  7  Arginine  30  Lysine  63  Glycine  4  Tyrosine  8  Cysteine  31  Arginine  65  Threonine  5  Isoleucine  9  Proline  49  Tyrosine  71  Alanine  6  Threonine  9  Leucine  55  Proline  111  Valine  6  Alanine  11  Aspartate  55  Valine  133  Serine  7  Valine  11  Valine  57  Cystine  382  Lysine  7  Serine  13  Lysine  58  Serine  427  Isoleucine  8  Asparagine+Aspartate  14  Threonine  79  Threonine  438  Asparagine+Aspartate  8  Leucine  19  Glycine  84  Aspartate  479  Proline  10  Methionine  24  Serine  100  Histidine  492  Leucine  11  Proline  25  Glutamine  135  Alanine  643  Glutamine+Glutamate  14  Glutamine+Glutamate  37  Alanine  200  Glutamate  908          Taurine  287  Taurine  1268          Glutamate  1175  Glycine  1352              Glutamine  3445          Milk Free Amino Acids (µmol/L)  Milk Protein-Bound Amino Acids (mmol/L)  Human  Porcine  Human  Porcine  Amino Acid  Concentration  Amino Acid  Concentration  Amino Acid  Concentration  Amino Acid  Concentration  Methionine  10  Tryptophan  18  Methionine  Trace  Histidine  4  Isoleucine  14  Isoleucine  21  Histidine  2  Lysine  4  Phenylalanine  16  Methionine  23  Arginine  3  Arginine  6  Histidine  22  Phenyalanine  36  Phenylalanine  3  Glycine  7  Tyrosine  22  Leucine  45  Tyrosine  4  Phenyalanine  7  Arginine  30  Lysine  63  Glycine  4  Tyrosine  8  Cysteine  31  Arginine  65  Threonine  5  Isoleucine  9  Proline  49  Tyrosine  71  Alanine  6  Threonine  9  Leucine  55  Proline  111  Valine  6  Alanine  11  Aspartate  55  Valine  133  Serine  7  Valine  11  Valine  57  Cystine  382  Lysine  7  Serine  13  Lysine  58  Serine  427  Isoleucine  8  Asparagine+Aspartate  14  Threonine  79  Threonine  438  Asparagine+Aspartate  8  Leucine  19  Glycine  84  Aspartate  479  Proline  10  Methionine  24  Serine  100  Histidine  492  Leucine  11  Proline  25  Glutamine  135  Alanine  643  Glutamine+Glutamate  14  Glutamine+Glutamate  37  Alanine  200  Glutamate  908          Taurine  287  Taurine  1268          Glutamate  1175  Glycine  1352              Glutamine  3445          1Porcine: all values (29 d of lactation) obtained from Wu and Knabe (1994), except Taurine from Zhang et al. (2015). Human: free amino acid values (21–60 d of lactation) obtained from Zhang et al. (2013b). Protein-bound amino acid values from Lemons et al. (1983). View Large Table 3. Amino acid composition of porcine and human milk, values are arranged from lowest to highest concentration1 Milk Free Amino Acids (µmol/L)  Milk Protein-Bound Amino Acids (mmol/L)  Human  Porcine  Human  Porcine  Amino Acid  Concentration  Amino Acid  Concentration  Amino Acid  Concentration  Amino Acid  Concentration  Methionine  10  Tryptophan  18  Methionine  Trace  Histidine  4  Isoleucine  14  Isoleucine  21  Histidine  2  Lysine  4  Phenylalanine  16  Methionine  23  Arginine  3  Arginine  6  Histidine  22  Phenyalanine  36  Phenylalanine  3  Glycine  7  Tyrosine  22  Leucine  45  Tyrosine  4  Phenyalanine  7  Arginine  30  Lysine  63  Glycine  4  Tyrosine  8  Cysteine  31  Arginine  65  Threonine  5  Isoleucine  9  Proline  49  Tyrosine  71  Alanine  6  Threonine  9  Leucine  55  Proline  111  Valine  6  Alanine  11  Aspartate  55  Valine  133  Serine  7  Valine  11  Valine  57  Cystine  382  Lysine  7  Serine  13  Lysine  58  Serine  427  Isoleucine  8  Asparagine+Aspartate  14  Threonine  79  Threonine  438  Asparagine+Aspartate  8  Leucine  19  Glycine  84  Aspartate  479  Proline  10  Methionine  24  Serine  100  Histidine  492  Leucine  11  Proline  25  Glutamine  135  Alanine  643  Glutamine+Glutamate  14  Glutamine+Glutamate  37  Alanine  200  Glutamate  908          Taurine  287  Taurine  1268          Glutamate  1175  Glycine  1352              Glutamine  3445          Milk Free Amino Acids (µmol/L)  Milk Protein-Bound Amino Acids (mmol/L)  Human  Porcine  Human  Porcine  Amino Acid  Concentration  Amino Acid  Concentration  Amino Acid  Concentration  Amino Acid  Concentration  Methionine  10  Tryptophan  18  Methionine  Trace  Histidine  4  Isoleucine  14  Isoleucine  21  Histidine  2  Lysine  4  Phenylalanine  16  Methionine  23  Arginine  3  Arginine  6  Histidine  22  Phenyalanine  36  Phenylalanine  3  Glycine  7  Tyrosine  22  Leucine  45  Tyrosine  4  Phenyalanine  7  Arginine  30  Lysine  63  Glycine  4  Tyrosine  8  Cysteine  31  Arginine  65  Threonine  5  Isoleucine  9  Proline  49  Tyrosine  71  Alanine  6  Threonine  9  Leucine  55  Proline  111  Valine  6  Alanine  11  Aspartate  55  Valine  133  Serine  7  Valine  11  Valine  57  Cystine  382  Lysine  7  Serine  13  Lysine  58  Serine  427  Isoleucine  8  Asparagine+Aspartate  14  Threonine  79  Threonine  438  Asparagine+Aspartate  8  Leucine  19  Glycine  84  Aspartate  479  Proline  10  Methionine  24  Serine  100  Histidine  492  Leucine  11  Proline  25  Glutamine  135  Alanine  643  Glutamine+Glutamate  14  Glutamine+Glutamate  37  Alanine  200  Glutamate  908          Taurine  287  Taurine  1268          Glutamate  1175  Glycine  1352              Glutamine  3445          1Porcine: all values (29 d of lactation) obtained from Wu and Knabe (1994), except Taurine from Zhang et al. (2015). Human: free amino acid values (21–60 d of lactation) obtained from Zhang et al. (2013b). Protein-bound amino acid values from Lemons et al. (1983). View Large In the early postnatal period, Arg is an essential dietary AA because during this period the sole diet of maternal milk is a relatively poor source of Arg (Table 3), and the Arg precursors Pro and Cit (Davis et al., 1994); suckling piglet performance benefits from additional Arg supplementation (Mateo et al., 2008; Wu et al., 2004). Furthermore, Arg supplementation protects and enhances intestinal mucosal immune barrier function and maintains intestinal integrity in weaned pigs (Zhu et al., 2013). Insufficient Arg has been linked to the onset of necrotizing enterocolitis in premature infants (Wu et al., 2004). To account for the observed Arg deficiency in the perinatal period, researchers have investigated the use of supplemental Pro, Orn, and Cit as precursors of Arg synthesis to neonatal piglets. Urschel et al. (2006) show that neonatal piglets fed an Arg-deficient diet supplemented with Cit are able to increase Arg to levels similar to supplementation with Arg alone, whereas Pro and Orn cannot not. The authors suggest this is due to the low abundance of the Orn transporter ORNT1 and the inefficient conversion of Pro to Arg in enterocytes of preweaning pigs. Postweaning, Arg becomes a conditionally essential AA as endogenous production becomes a multiorgan process, and the diet contains sufficient Arg for supporting metabolic needs; only during times of disease or metabolic stress does Arg become essential. A study of E. coli LPS-induced damage in weaned piglets shows that Arg supplementation can also protect the GIT by modulating the intestinal inflammatory response through increased numbers of intraepithelial lymphocytes, CD4+ T cells, CD8+ T cells, and IgA-secreting cells, and decreasing Peyer's patch cell apoptosis and mast cell number in the intestinal mucosa (Zhu et al., 2013). Metabolomic analysis of weaned piglets shows supplementation with Arg leads to alterations in the metabolites produced by resident microbiota of the small and large intestine. The researchers suggest this is due to the production of nitric oxide (NO) by cells of the lumen (He et al., 2009). This observation is in agreement with other studies which show that the beneficial effect of Arg is not direct, but mediated by Arg metabolites produced in cells of the GIT, in particular, NO. Two studies (Liu et al., 2008a; Liu et al., 2008b) show that the production of NO in response to oral Arg supplementation protects the GIT of weaned piglets from E. coli LPS-induced damage. The use of Arg to improve GIT function and health is a two-edged sword, low doses have no appreciable effect, while doses that are too high adversely affect microvascular development (Zhan et al., 2008). Like Arg, Pro, and Cit, porcine and human milk are both deficient (Table 3) in protein-bound Gly (Davis et al., 1994). Glycine supplementation of milk fed piglets has been shown to enhance small-intestinal villus height, antioxidative capacity, and stimulate Gly transporter abundance (Wang et al., 2014), while a therapeutic study using Cys to treat piglets with an induced model of inflammatory bowel disease showed Cys ameliorated intestinal inflammation and permeability (Kim et al., 2009). CONCLUSION Identifying and understanding how nutritional components interact with and regulate the function and health of the human GIT requires the use of appropriate animal models. Although rats and mice are commonly used models for studying fundamental GIT processes, the pig with its greater size, similar eating patterns, and GIT physiology, is a closer translational model for testing nutritional interventions. 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TI - REVIEW: The pig as a model for humans: Effects of nutritional factors on intestinal function and health
JF - Journal of Animal Science
DO - 10.2527/jas.2015-9788
DA - 2016-09-01
UR - https://www.deepdyve.com/lp/oxford-university-press/review-the-pig-as-a-model-for-humans-effects-of-nutritional-factors-on-0yQqU0Lz4b
SP - 441
EP - 452
VL - 94
IS - suppl_3
DP - DeepDyve
ER -