Download 24 Gastrointestinal protein and amino acid metabolism in

24 Gastrointestinal protein and amino acid
metabolism in growing animals1
D. G. Burrin
USDA/ARS Children’s Nutrition Research Center,
Department of Pediatrics, Baylor College of Medicine,
1100 Bates Street, Houston, Texas 77030
The tissues of the gastrointestinal tract play a major role in the metabolism
of protein and amino acids in growing animals. Intestinal tissue has a high
rate of protein metabolism, which is directly linked to the high rates of proliferation, protein secretion, cell death and desquamation of various epithelial and lymphoid cells within the mucosa. Because the small intestine is the
first tissue exposed to the diet, it has a key regulatory role in the digestion,
absorption, metabolism and availability of dietary protein and amino acids
for growth. The major oxidative fuels for the intestine are glutamate, glutamine, aspartate and glucose; however, some essential amino acids are also
oxidized, such as lysine, leucine and phenylalanine. Among the essential
amino acids, threonine utilization is particularly high, and may be critical
for normal intestinal function. Intestinal protein and amino acid metabolism
is primarily regulated by the modality (enteral versus parenteral) and quantity of nutrient intake, and to a lesser extent by the composition of nutrients.
1
This work is a publication of the USDA=ARS Children’s Nutrition Research Center, Department of
Pediatrics, Baylor College of Medicine and Texas Children’s Hospital, Houston, TX. The work was supported in part by federal funds from the U.S. Department of Agriculture Agricultural Research Service,
Cooperative Agreement No. 58-6258-6001, by the National Institutes of Health R01 HD33920. The contents of this publication do not necessarily reflect the views or policies of the U.S. Department of Agriculture, nor does mention of trade names, commercial products, or organizations imply endorsement by
the U.S. Government.
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Biology of the Intestine in Growing Animals
R. Zabielski, P.C. Gregory and B. Weström (Eds.)
# 2002 Elsevier Science. B.V. All rights reserved.
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D. G. Burrin
Intestinal metabolism has a significant influence on the quantity and pattern
of amino acids required for growth, particularly arginine and threonine.
1. INTRODUCTION
During the last 20 years, research studies established that the tissues of the
gastrointestinal tract have a substantial impact on whole-body protein and
amino acid metabolism in growing animals. The portal-drained visceral
(PDV) tissues, comprised largely of gastrointestinal tissues, contribute
between 3 and 6% of body weight, but they account for 20 to 35% of
whole-body protein turnover and energy expenditure (Burrin et al., 1989;
Yen et al., 1989; Nieto and Lobley, 1999; Stoll et al., 1999a). The disproportionate impact of gastrointestinal tissues on whole-body metabolism is
a function of their relatively high fractional rates of protein synthesis and
oxygen consumption. Studies with domestic animals have demonstrated that
the fractional synthesis rate of protein in the intestinal tissues is several-fold
higher than that in peripheral tissues, such as muscle (Lobley et al., 1980,
1992; Attaix et al., 1986; Burrin et al., 1992; Davis et al., 1996). Although
we know much about the heterogeneous phenotypes and characteristic functions of the cells that comprise the intestinal mucosa, the quantitative influence of these different cell types on mucosal protein metabolism as a whole
has yet to be fully delineated. Moreover, we have a limited knowledge of
how the spatial localization of cells within the mucosa determines the
source of substrate, its supply, and rate of metabolism.
Two critically important physiological functions of the intestine that are
necessary for maximal growth of the organism are: (1) to digest and
absorb dietary nutrients, and (2) to serve as a biological barrier in order
to prevent or minimize effects of environmental pathogens, toxins and
other antigenic molecules. Studies during the past decade have suggested
that ensuring an adequate nutrient supply to the intestine, mainly via the
oral or enteral route, is essential to maintain these functions. Currently,
efforts are increasing to quantify the extent to which the intestine uses
nutrients for its own purposes, and modifies the rate and pattern of amino
acid supply for peripheral tissue growth (Reeds et al., 1993; Reeds et al.,
1999; Burrin et al., 2001). Given the significance of the intestinal tissues
to whole-body metabolism, the investigation of those factors that substantially alter intestinal mass has become increasingly relevant to the study of
whole-animal growth.
The intent of this review is to characterize gastrointestinal protein
and amino acid metabolism by examining (1) the underlying influence of
anatomy and cellular function, (2) the major metabolic fates of amino acids,
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and (3) the relative significance of nutrition, ontogeny, infection and inflammation.
2. ANATOMICAL AND MORPHOLOGICAL BASIS
OF GUT METABOLISM
2.1. Regional metabolism within the gastrointestinal tract
There are notable differences in the rates of metabolism within the tissues of
the gastrointestinal tract. Studies in developing animals indicate that the
rates of protein synthesis are higher in the small intestine than in the stomach and large intestine (Attaix and Arnal, 1987; Attaix et al., 1992; Burrin
et al., 1999a). Within the small intestine, the protein synthesis rate is highest
in the duodenum, and declines longitudinally (fig. 1) (Attaix et al., 1992;
Stoll et al., 2000a). There are numerous estimates of non-essential amino
acid metabolism (mainly glutamine, arginine, citrulline and ornithine) in
primary isolated enterocytes (Darcy-Vrillon et al., 1994; Quan et al.,
1998; Wu, 1998), colonocytes (Mouille et al., 1999), and colon carcinoma
cell-lines (Selamnia et al., 1998). One study comparing isolated cells
from the jejunum, caecum and colon found no difference in the glutamine
Fig. 1.
Fractional rates of protein synthesis is small and large intestinal setments of milkfed and weaned lambs. (Adapted from Attaix et al., 1992).
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oxidation rate, despite regional differences in glucose and short-chain fatty
acid oxidation (Fleming et al., 1991).
2.2. Cellular basis of mucosal metabolism
The high rates of protein metabolism in the gut are directly linked to the
high rates of proliferation, protein secretion, cell death and desquamation
of various epithelial and lymphoid cells within the mucosa. In vivo labeling
studies in young pigs show that the epithelial cells have a life span of about
3 to 10 days, depending on the intestinal region and stage of development
(Smith and Jarvis, 1978; Fan et al., 2001). Within the small intestinal epithelium, the proliferative crypt compartment contains pluripotent stem cells,
which undergo mitosis and differentiation into four cell lineages: absorptive
enterocytes, mucin-producing goblet cells, antibacterial peptide- producing
Paneth cells, and enteroendocrine cells. The differentiated phenotype of
these cells defines their main functions, which are to digest and absorb dietary nutrients and provide an innate mucosal barrier to the luminal environment. Lymphoid cells are the other major class of cells located mainly in the
lamina propria region beneath the epithelial monolayer. These cells include
T- and B-lymphocytes, macrophages, neutrophils, mast cells, dendritic cells
and fibroblasts that are diffusely organized around a highly vascular and
neuronal network. Studies in vitro show that these epithelial (Higashiguchi
et al., 1993; Wu 1998) and lymphoid (Szondy and Newsholme, 1990;
Dugan et al., 1994) cell types exhibit high rates of protein synthesis and
glutamine metabolism. The protein secretory capacity of goblet cells is also
substantial, and mucin protein represents a significant component of endogenous secretions that pass undigested to the terminal ileum (Lien et al.,
1997; Montagne et al., 2000).
2.3. Luminal vs arterial substrate input
Another important anatomical consideration with respect to the metabolism
of epithelial cells is that they derive nutrients from two separate sources: the
intestinal lumen, via the apical=brush-border membrane, and the blood, via
the basolateral membrane. Studies in piglets with isotopically labeled substrates have shown that there is significant utilization of both luminal and
arterial amino acids. It is of interest to note that, in pigs fed a milk-based
formula, the input of amino acids from the luminal route (i.e. diet) is far
greater than that from the arterial circulation (fig. 2). With respect to the
uptake of amino acids, particularly glutamate and glutamine, there is markedly greater uptake from the luminal (67–96%) than the arterial (11–21%)
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Fig. 2. Contribution of dietary and arterial amino acid input into the portal-drained visceral
tissue in young pigs fed cow’s milk formula. (Adapted from Van Goudoever et al., 2000;
Stoll et al., 1999b; Stoll et al., 1998; Burrin et al., 1999b).
input. Interestingly, the uptake of luminal glucose, both on a relative and
absolute basis, is considerably less than that of amino acids ( 6% of
input). It is possible that this combined phenomenon is a function of the
inherent amino acid and glucose transport capacities of the apical (luminal)
and basolateral (arterial) surfaces of intestinal enterocytes. However, in
absolute molar terms, the uptake of some amino acids from the arterial
and luminal routes is nearly equal, because the rate of arterial amino acid
input is more than five times greater than that from the diet. Thus, some
amino acids used by the gut are derived equally from the diet and arterial
supply. In contrast, intestinal glucose is supplied largely by the arterial circulation (85% total), with only a small contribution from the diet (15%).
The relatively high rates of first-pass metabolism of dietary amino acids
may be a phenomenon that is largely confined to the small intestine rather
than the stomach or large intestine. That is because when intact protein is
consumed, the level of free amino acids is normally low in the stomach
as well as the large intestine due to bacterial fermentation. If the digestive
environment in the stomach and large intestine limits the luminal amino
acid availability, then the arterial circulation must meet the amino acid
needs of these tissues. This idea is supported by the evidence that during
total parenteral nutrition, atrophy occurs primarily in the proximal region
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of the small intestine, while the mass of the distal small intestine, stomach
and large intestine is preserved (Goldstein et al., 1985; Morgan et al., 1987;
Ganessunker et al., 1999).
2.4. Crypt vs villus cell metabolism
The extent to which epithelial cells derive their nutrients from the luminal
or vascular input is further influenced by their stage of differentiation and
physical location along the crypt-villus axis. Early studies (Alpers, 1972)
showed that crypt cells are more highly labeled with isotopic tracers derived
from the blood, whereas villus cells are more highly labeled with tracers
given luminally (fig. 3). Although these findings seem logical, in a general
sense, we still know very little about how the stage of differentiation and
amino acid transporter expression on the apical and basolateral surface of
enterocytes affect the rate and pattern of nutrient utilization. Studies with
cultured enterocytes indicate that the mitogenic effect of glutamine appears
to be dependent on its metabolism, but interestingly, it cannot be reproduced
with glutamate. This apparent specificity of glutamine as a crypt cell mitogen may be due to the fact that the crypt cell lines used in these studies had a
limited capacity to transport glutamate across the cell membrane. Yet, this
contradicts the findings of extensive intestinal glutamate metabolism documented in vivo (Windmueller and Spaeth, 1975; Reeds et al., 1996). On the
Fig. 3. Relative incorporation of luminally (-3H-leucine) and intravenously (*-14Cleucine) administered leucine into enterocytes isolated from the crypt to villus axis of the
intestinal mucosa (adapted from Alpers, 1972).
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other hand, it may be that crypt cells are capable of basolateral transport of
blood-borne glutamine (but not glutamate) for proliferative processes, but as
the cells differentiate and mature, they acquire the ability to transport luminal glutamate for use as a metabolic fuel and as a biosynthetic precursor
(fig. 4). This idea is supported by evidence that shows that glutamine transport is high in undifferentiated Caco-2 cells, but in confluent differentiated
cells glutamine transport is decreased, whereas glutamate transport is substantially increased (Mordrelle et al., 2000).
3. METABOLIC FATE OF AMINO ACIDS
Once taken up by the intestinal tissues, amino acids can be utilized for three
major metabolic purposes: (1) incorporation into protein; (2) conversion via
Fig. 4. Model of mucosal glutamate, glutamine, and aspartate metabolism derived from
dietary intake and the arterial circulation.
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transamination into other amino acids, metabolic substrates and biosynthetic
intermediates; and (3) complete oxidation to CO2. In the first two pathways,
amino acids can be deposited and recycled by the body for purposes of
growth or other biological functions. In the case of some amino acids,
namely threonine and cysteine, incorporation into endogenous secretions
that are fermented in the large intestine represents a net nutritional
loss. Likewise, if the amino acids are completely oxidized to CO2 by the
mucosal cells, this is also a nutritional loss, especially in the case of essential amino acids.
3.1. Protein synthesis and degradation
The major metabolic fate of amino acids in the gut is considered to be their
incorporation into cellular proteins. Although this is true of essential amino
acids, there is extensive metabolism of several non-essential amino acids.
Numerous studies have measured the rates of protein synthesis in various
tissues of the gastrointestinal tract. In general, these studies show that the
fractional rates of protein synthesis are relatively high compared to other
major tissues such as muscle, due to the high rates of cell turnover. Moreover, gut protein synthesis rates are modulated by several nutritional and
hormonal factors, the details of which are described below. However, there
is a lack of fundamental knowledge about the cellular and molecular
mechanisms that regulate protein synthesis in gut tissues in comparison with
our knowledge of other tissues, such as liver and muscle. Recent evidence
has shown that the stimulation of muscle protein synthesis by food intake,
insulin and amino acids is mediated by activation of intracellular signal
pathways which include IRS-1, PI-3 kinase, MAP-kinase, mTOR and
p70-S6-kinase (Kimball et al., 1998; Davis et al., 2000; Kimball et al.,
2000; Anthony et al., 2001). These signaling pathways play a critical role
in the proliferation and differentiation of intestinal epithelial cells (Taupin
and Podolsky, 1999; Wang et al., 2001) and can be activated by amino acids
(Rhoads et al., 2000); yet, their role in the regulation of protein synthesis is
largely unknown. We know even less about the regulation of protein degradation in gut tissues, whether at the molecular and cellular level, or with
respect to nutritional and hormonal regulation (Baracos et al., 2000).
3.2. Nonessential amino acid metabolism
Of the amino acids ingested in the diet, the intestinal metabolism of glutamine and glutamate has been studied most extensively (fig. 5). It has
been known for more than 25 years that there is substantial utilization of
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Fig. 5. Synthesis and interconversion of glutamate and glutamine. (1) Glutamate dehydrogenase; (2) Alanine aminotransferase, aspartate aminotransferase, or branched-chain
aminotransferase; (3) Glutaminase; (4) Glutamine synthetase.
glutamine and glutamate by the intestine, and much of the glutamine is
derived from the arterial circulation (Windmueller and Spaeth, 1975;
1980). More important, however, is the fact that in situ studies (Windmueller and Spaeth, 1980) have shown that at least 50% of the dietary glutamate
and glutamine taken up by the gut is completely oxidized to CO2. These
results were recently confirmed by in vivo studies of 13C-labeled glutamate
and glutamine metabolism by the PDV tissues in piglets indicating that as
much as 95% of enteral glutamate is metabolized by the gut, largely to
CO2 (Stoll et al., 1999a) (fig. 6). Consistent with this finding, studies in premature infants and adult humans, based on estimates of whole-body kinetics
of enterally fed 13C- and 15N-labeled glutamate and glutamine, indicate that
approximately 50 to 95% of the dietary glutamine and glutamate is
extracted by splanchnic tissues, and most of this is oxidized to CO2 (Battezzati et al., 1995; Darmaun et al., 1997; Haisch et al., 2000).
In addition to the role of glutamate as metabolic fuel, studies in rats
(Horvath et al., 1996) have shown that adding 4% glutamate to an elemental
diet deficient in both glutamine and glutamate significantly increases intestinal growth, crypt cell mitosis, and lactase activity, without affecting body
weight gain. These results serve to challenge the long-standing notion that
glutamine is the primary oxidative fuel for the gut, and suggest that glutamate and perhaps aspartate ingested in the diet are equally important intestinal fuels. Moreover, studies with short-term cultures of isolated enterocytes
have shown that, along with glutamine, glucose is a major intestinal oxidative fuel (Darcy-Vrillon et al., 1994; Kight and Fleming, 1995; Wu et al.,
1995). Although our results indicate that significant quantities of glucose
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Fig. 6. Relative rates of arterial and dietary substrate uptake and oxidation by the portaldrained visceral tissues in young pigs fed cow’s milk formula (GLN, glutamine; GLU,
glutamate; GLUC, glucose; ASP, aspartate; Adapted from stoll et al., 1999a).
are indeed oxidized by the gut, they demonstrate a preferential utilization of
arterial rather than dietary glucose. It is important to note that, although
glucose represents a major oxidative fuel (29%) in terms of total PDV
CO2 production, the proportion of glucose oxidized completely to CO2 is
substantially less than that of either glutamate or glutamine. This finding
is entirely consistent with results from cultured enterocytes showing that
glutamine can potently suppress glucose oxidation, whereas glucose has little impact on glutamine oxidation (Kight and Fleming, 1995; Wu et al.,
1995). The implication is that glutamine and glutamate are preferentially
channeled towards mitochondrial oxidation, while most of the glucose is
utilized for other metabolic or biosynthetic purposes. A small percentage
of the portal glucose utilization is metabolized to alanine (7.5%) and lactate
(5.9%), which leaves approximately 60% unaccounted for. The biosynthetic
products potentially derived from gut glucose utilization are numerous;
however, incorporation into mucin glycoproteins, fatty acids and lipids
presents a variety of intriguing possibilities.
The importance of glutamine and glutamate to gut metabolism goes
beyond their roles as oxidative fuel for rapidly proliferating cells, such as
enterocytes and lymphocytes; they also function as important precursors
Gastrointestinal protein and amino acid metabolism in growing animals
705
of nucleic acids, nucleotides, amino sugars, amino acids, and glutathione
(Smith, 1990; Souba, et al., 1990; Rouse et al., 1995; Klimberg, 1996; Gate
et al., 1999). A number of studies with adult rats, neonatal pigs and porcine
enterocytes have demonstrated that glutamine=glutamate are precursors
for the synthesis of several non-essential amino acids, including alanine,
aspartate, proline, citrulline and arginine (Windmueller and Spaeth, 1980;
Blaicher et al., 1993; Wu et al., 1995; Wu and Knabe, 1995; Stoll et al.,
1999a). In addition, Reeds et al. (1997) reported that luminal glutamate,
rather than glutamine-derived glutamate, is the preferential source of glutathione synthesis in the intestinal mucosa of the infant pig, suggesting a
highly compartmentalized catabolism of intestinal glutamate and glutamine.
It is also noteworthy that the enzymes necessary for de novo synthesis of
both glutamate and glutamine are present in gut tissues, and their expression
is up-regulated during the suckling-to-weanling transition (Shenoy et al.,
1996; Madej et al., 1999).
Studies with cultured intestinal epithelial cells have demonstrated that
glutamine has pluripotent actions, including stimulation of cell proliferation, differentiation, ornithine decarboxylase (ODC) and immediate early
gene (c-jun) expression, and polyamine synthesis (Kandil et al., 1995; Wang
et al., 1996). Glutamine also suppresses apoptosis, which suggests that it
may be an important survival factor (Papaconstantinou et al., 1998). In a
recent review, Rhoads (1999) postulated that glutamine stimulates cell proliferation by a signaling mechanism which involves the activation of two
related, but distinct, classes of mitogen-activated protein kinases. The biochemical mechanism whereby glutamine affects intestinal function may
be related to its conversion to glucosamine, which reduces the cellular
NADPH and suppresses nitric oxide synthesis (Wu et al., 2001).
3.3. Essential amino acid metabolism
A number of essential amino acids play a key role in gut function and thus
their metabolism by the gut has a critical influence on essential amino acid
requirements. Dietary threonine and cysteine are utilized by the intestinal
mucosa for mucin and glutathione synthesis. From a nutritional perspective,
although threonine is considered essential, cysteine is not. However,
cysteine can be synthesized from the essential amino acid, methionine.
Therefore, increased metabolism of methionine to meet cysteine needs
may become limiting for growth, and nutritionally significant. The secretory
mucins play a key role in the innate immune defense of the mucosa, and the
core protein of the major intestinal mucins (van Klinkenet al., 1997)
contains a large amount of threonine and cysteine. Likewise, cysteine is a
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component of the tripeptide antioxidant glutathione, which is critical for the
maintenance of the structural integrity and barrier function of the intestinal
mucosa (Martenssonet al., 1991). Given that the rates of mucosal synthesis
and secretion of glutathione and mucins are likely significant, it follows
that their production by the gut could have a quantitatively significant
impact on the animal’s dietary requirement for threonine, cysteine, and perhaps methionine. For threonine, but not cysteine, there is evidence to support this hypothesis. First, based on net portal utilization, as much as 60
to 85% of dietary threonine is utilized by the gut (Stoll et al., 1998; Van
Goudoever et al., 2000). More conclusive, however, was a recent study
(Bertolo et al., 1998) demonstrating that the threonine requirement of piglets maintained by parenteral nutrition was only 40% of that observed in
piglets receiving enteral feedings. Moreover, a subsequent report found
that feeding threonine-deficient diets to piglets significantly reduces intestinal mass and goblet cell numbers, and this suppression of intestinal growth
cannot be fully restored by providing threonine parenterally (Ball et al.,
1999). Another study demonstrated that the methionine requirement of
piglets maintained by parenteral nutrition was 35% lower than that of enterally fed piglets (Shoveller et al., 2000; 2001). This value is lower than
the value of 46% based on our measurements of net portal utilization of
dietary methionine. Nevertheless, both findings suggest that the gut utilizes
substantial amounts of dietary methionine, and raise the question of
whether methionine is being metabolized to cysteine and used for intestinal
mucin synthesis.
Dietary arginine is essential for young piglets. Studies have shown that
the small intestine is an important site of arginine and proline synthesis
(fig. 7) (Murphy et al., 1996; Wu, 1998; Stoll et al., 1999a). Similarly, in
adult rats and weanling pigs (Windmueller and Spaeth, 1980; Dugan et al.,
1995), intestinal citrulline synthesis from glutamine, glutamate and proline
is the main source for circulating citrulline, which plays a critical role in
arginine homeostasis. In many adult animals and humans, arginine is
synthesized by the kidney from intestinally derived citrulline at rates that
are inadequate to support growth in neonatal animals. As the milk of
humans, pigs, rats and many other mammals is deficient in arginine (Davis
et al., 1994), it is considered an essential amino acid for growing animals. In
the healthy suckling pig, the intestinal synthesis of arginine provides only
about half of the animal’s needs for growth; thus, arginine is required in
the diet. Moreover, the net intestinal synthesis of arginine declines substantially during the late suckling period (Wu and Morris, 1998). These observations raise the possibility that both the endogenous (via gut synthesis) and
dietary arginine supply may be limiting for maximal growth of suckling
piglets. Studies with enterocytes from newborn pigs (Blaicher et al.,
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707
Fig. 7. Metabolic pathways and interconversion of arginine, ornithine, glutamate, and proline.
(1) arginase; (2) ornithine aminotransferase; (3) glutamate-g-semi-aldehyde dehydrogenase;
(4) g-glutamyl kinase; (5) g-glutamyl dehydrogenase; (6) spontaneous, non-enzymatic;
(7) pyrroline-5-carboxylase reductase; (8) proline oxidase; (9) ornithine transcarbamoylase;
(10) NO synthase; (11) arginine decarboxylase.
1993; Wu and Knabe, 1995) and developing mouse and rat small intestine
(Hurwitz and Kretchmer, 1986; DeJonge et al., 1998) have shown developmental changes and metabolic zonation along the crypt-villus axis of arginine-metabolizing enzymes. At birth, the enterocytes of the upper villus
(DeJonge et al., 1998) are the major site of arginine synthesis, but gradually
become the major site of net citrulline production as intestinal arginase
expression increases via a glucocorticoid-dependent mechanism. This transition is compensated by the gradually increasing capacity of the kidney to
use citrulline for arginine synthesis. Thus, following the transition from
suckling to weanling, the intestine appears to become a site of arginine
degradation rather than synthesis (see review, Wu and Morris, 1998).
The limited arginine degradation by enterocytes from newborn pigs
ensures a maximum output of arginine (synthesized from glutamine or
derived from milk) into the portal circulation for utilization by extraintestinal tissues. Type II arginase, a mitochondrial enzyme that is expressed at
lower levels in kidney, brain, small intestine, mammary gland, and macrophages but not in liver, is distinct from type I arginase, a cytosolic enzyme
that is highly expressed in liver as a component of the urea cycle. The
induction of type II arginase in enterocytes after weaning possibly regulates
the availability of arginine for the synthesis of nitric oxide (NO), ornithine
and thus, polyamines, proline, and glutamate, as well as ureagenesis in the
small intestinal mucosa (fig. 7). lthough considerably less than in the liver,
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gut ureagenesis may be a first-line defense in detoxification of ammonia
derived from tissue metabolism and luminal microorganisms (Wu, 1995).
However, the major end-products of arginine metabolism by intestinal
enterocytes from weaned pigs are proline and ornithine. Proline, required
for collagen synthesis, is one of the most abundant amino acids in human
milk (Davis et al., 1994). Arginine-derived proline is not detectable in enterocytes of newborn or suckling pigs. However, providing proline in the diet
can ameliorate the hyperammonemia associated with dietary arginine deficiency in neonatal pigs (Brunton et al., 1999). Thus, while proline is not
considered an absolute dietary essential nutrient, it may be conditionally
essential for maintaining arginine synthesis in neonates. It is apparent from
a number of studies that a normally functioning gut is important for maintenance of whole-body arginine and proline status, especially in neonates.
Furthermore, these amino acids become conditionally essential under conditions that markedly reduce gut mass or compromise function, such as
massive small bowel resection (Wakabayashi et al., 1995).
Recent studies in cultured intestinal cells (Blaicher et al., 1995) have
shown that ornithine derived from arginine metabolism is converted to polyamines (fig. 8). Polyamines (putrescine, spermidine, spermine, cadaverine)
Fig. 8. Polyamine synthesis. (1) ornithine decarboxylase; (2) spermidine synthetase;
(3) spermidine synthase; (4) polyamine oxidase; (5) methionine adenosyl transferase;
(6) S-adenosymethionine decarboxylase.
Gastrointestinal protein and amino acid metabolism in growing animals
709
are ubiquitous cationic amines involved in cell proliferation and differentiation in many tissues, including the gastrointestinal tract. Ornithine decarboxylase (ODC) and S-adenosyl-methionine decarboxylase (SAMDC),
converting ornithine to putrescine and putrescine to spermine, respectively,
are the rate-limiting enzymes in polyamine synthesis. The synthesis of
polyamines from arginine is negligible in enterocytes of newborn and suckling animals (Blaicher et al., 1991; Blaicher et al., 1992), but increases in
enterocytes of postweaning animals, concurrent with the induction of both
arginase and ODC (Wu and Morris, 1998). Luminal administration of polyamines increases intestinal growth in adult rats (Seidel et al., 1985) and has
been shown to enhance intestinal maturation (Grant et al., 1990) and cell
proliferation in developing rats (Dufour et al., 1988). Polyamines are present in human milk in micromolar concentrations for up to 4 months of lactation (Pollack et al., 1992; Buts et al., 1995). Results of a recent study in
cultured intestinal cells suggested that both human and rat milk, but neither
bovine milk nor infant formula, contain sufficient amounts of polyamines to
sustain cell growth during inhibition of polyamine synthesis with difluoromethylornithine (DFMO), a specific and irreversible inhibitor of ODC
(Capano et al., 1998). Thus, when the ingestion of milkborne polyamines
by the neonate ceases after weaning, the induction of intestinal polyamine
synthesis from ornithine, arginine and proline may become physiologically
significant for the maintenance of normal intestinal growth and function
(Wu et al., 2000a; 2000b). Furthermore, the induction of intestinal polyamine synthesis is dependent on the weaning-induced cortisol surge.
3.4. Essential amino acid oxidation
The catabolism of essential amino acids by the gut has received little recent
attention, in part because early studies failed to demonstrate the presence of
the catabolic enzymes in the small intestinal mucosa (Wu, 1998). However,
studies based on isotopic tracer kinetics in PDV tissues suggest that dietary
essential amino acids may indeed be oxidized by the intestinal mucosa. Studies in sheep (Pell et al., 1986; Cappelli et al., 1997; Yu et al., 2000), pigs
(Burrin et al., 1999b) and dogs (Yu et al., 1992) have found that approximately 5–10% of whole-body leucine flux is oxidized by the portal-drained
viscera. Recent studies in piglets enterally fed [U-13C] threonine, leucine,
lysine and phenylalanine suggested significant first-pass metabolism by
the intestinal mucosa (Stoll et al., 1998; Stoll et al., 1999; Van Goudoever
et al., 2000; Van der Schoor et al., 2001). In growing pigs, lysine is considered to be the first limiting dietary amino acid. Studies in young piglets
showed that intestinal oxidation of dietary lysine accounted for about
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one-third of whole-body lysine oxidation. Interestingly, although about 10%
of the arterial flux of lysine was taken up by the portal-drained viscera, none
of this was oxidized, suggesting a preferential oxidation of dietary lysine.
Studies in young pigs suggest that approximately 15 to 30% of the wholebody leucine and phenylalanine flux is oxidized by the PDV tissues (Van
der Schoor et al., 2001; Bush et al., 2001). The oxidation of phenylalanine
implies that phenylalanine hydroxylation occurs in the gut and is consistent
with previous observations suggesting de novo gut tyrosine production.
However, threonine seems to be unique among the essential amino acids
in that there is negligible 13C-threonine oxidation to CO2 when given either
enterally or systemically to young piglets. This may be due to the fact that it
is preferentially used for synthesis of mucins.
4. ROLE OF ONTOGENY
The fractional rates of whole-body protein metabolism are highest
during foetal life, and decline progressively with advancing post-conceptual
age, commensurate with fractional growth rates (Waterlow et al., 1978;
Goldspink and Kelly, 1984). In rats, the intestinal protein mass increases
approximately 750-fold (2.5 to 1893 mg between 18 days gestational age
and 105 weeks postpartum age (Goldsplink et al., 1984). During this life
span, however, the fractional protein synthesis rate (FSR) in the small
and large intestine declines by 43%. After weaning, the decline in intestinal
FSR with age is largely due to a decreased synthesis in the muscularis and
serosal layers, whereas the mucosa remains constant (Merry et al., 1992).
Despite the overall age-related decline, studies in neonatal rats and mice
indicate that the FSRs of the stomach, small intestine and pancreas increase
significantly after weaning (Burrin et al., 1991; Burrin et al., 1999a). In
domestic animals, there are few, if any, estimates of gastrointestinal FSRs
before birth and beyond pubertal ages, yet the changes between birth and
weaning in pigs and sheep tend to parallel those in rodents (Seve et al.,
1986; Attaix et al., 1992; Davis et al., 1996). In milk-fed animals, the intestinal FSR declines during the neonatal period, but increases markedly (40–
50%) after weaning. Even more striking is the relative contribution of the
small and large intestine to whole-body FSR, which was found to be
approximately 10% in milk-fed lambs and 20% in weaned lambs. The
explanation for this sharp increase in gut protein synthesis after weaning
is likely the substantial change in the composition diet and resultant stimulation of mucosal cellularity and proliferation (Attaix and Meslin, 1991;
Pluske, 1997; Jiang et al., 2000). The extent to which these changes are
related to alterations in the gut microflora is also a probable factor.
Gastrointestinal protein and amino acid metabolism in growing animals
711
With respect to the ontogenic changes in the metabolism of specific
amino acids, most studies have focused on glutamine, arginine, citrulline,
ornithine and proline. The rates of glutamine and glucose oxidation in isolated enterocytes decrease by roughly 90% in the first 3 weeks of life
(Darcy-Vrillon et al., 1994; Wu et al., 1995). The percentage of glutamine
and glucose oxidized to CO2 at birth is 36 and 21%, respectively, and
declines at 3 weeks of age to 4 and 2%, respectively. After weaning, in
intraepithelial lymphocytes, glutamine is mainly metabolized to glutamate
and ammonia (92%), with minimal oxidation (4%). The decline in glutamine oxidation with age is paralleled by increased activities of glutamine
synthetase, glutaminase, and glutamate dehydrogenase (Hahn et al., 1988;
Shenoy et al., 1996; Madej et al., 1999). Other studies in isolated enterocytes show that the synthesis of citrulline and ornithine increases with
age, particularly after weaning, and proline is a major precursor for their
synthesis (Wu, 1997). Recent studies indicate that this shift in ornithine production is associated with increased ODC activity and is dependent on
increased circulating cortisol (Wu et al., 2000a; Wu et al., 2000b).
5. ROLE OF NUTRITION
5.1. Enteral vs parenteral
Oral feeding is a potent stimulus of intestinal protein and amino acid metabolism in growing animals (McNurlan et al., 1979; Patureau Mirand et al.,
1990; Burrin et al., 1991; Burrin et al., 1995). Prolonged fasting leads to
markedly reduced protein mass in gut tissues via suppressed protein synthesis and increased protein degradation, especially in the small intestine
(Burrin et al., 1991; Samuels et al., 1996). Recent studies have shown that
the stimulatory effect of nutrient intake on gut protein metabolism is highly
dependent on the administration of nutrients via the enteral route (Dudley
et al., 1998; Burrin et al., 2000; Stoll et al., 2000a). These piglet studies
showed that net loss of intestinal protein occurs during 7 days of TPN,
and that intestinal protein accretion is highly correlated with the level of
enteral intake; intestinal protein balance occurred at 20% enteral intake,
and maximal intestinal protein accretion at 60% enteral intake (fig. 9).
Numerous studies have shown positive effects of glutamine-supplemented TPN in preventing atrophy, stimulating protein anabolism and maintaining intestinal permeability (Tamada et al., 1992; Platell et al., 1993; Inoue
et al., 1993; Haque et al., 1996; Naka et al., 1997; Khan et al., 1999). In
contrast, other studies have shown no effect of glutamine on gut growth
or protein metabolism in animals receiving glutamine supplementation
712
Fig. 9.
D. G. Burrin
Small intestinal protein balance in neonatal pigs fed varying proportions of enteral
and parenteral nutrition (Adapted from Stoll et al., 2000a).
parenterally (Spaeth et al., 1993; Burrin et al., 1994; Marchini et al., 1999;
Humbert et al., 2001). Furthermore, parenteral infusion of lipid has been
shown to stimulate intestinal protein synthesis more than either glucose
or glutamine (Stein et al., 1994). Thus, the intestinal trophic effects of glutamine-supplemented TPN are evident under conditions of compromised
gut function, such as sepsis, inflammation, and small-bowel resection, while
in healthy animals, glutamine provides limited benefit.
Although the provision of nutrition via the enteral route is critical for
intestinal growth, the effects of first-pass metabolism can significantly
increase whole-body requirements for arginine, threonine, and methionine
(Brunton et al., 2000). Moreover, studies in piglets have shown that when
first-pass intestinal metabolism is bypassed during TPN, arginine and proline become essential amino acids because their biosynthesis requires the
enteral input of precursors (Bertolo et al., 1999a). In contrast, the consequences of reduced gut mass following TPN may enhance the absorption
of nutrients when enteral feeding is resumed. Our preliminary studies indicated that the 52% reduction in intestinal mass observed after 7 days of
TPN was associated with a 30% increase in the net portal absorption of enterally administered leucine (Burrin et al., 1999b). Moreover, the increased
absorption was largely due to reduced portal utilization of arterial leucine,
Gastrointestinal protein and amino acid metabolism in growing animals
713
secondary to the reduced intestinal mass. This finding indicates that a 50%
reduction in gut mass actually increased the net systemic amino acid availability for growth. Interestingly, these results provide a metabolic explanation for the phenomenon of compensatory growth, which occurs when
animals are fed ad libitum following periods of restricted food intake. Previous studies in ruminants have shown that the reduction in the mass and
energy expenditure of the visceral organs during nutrient restriction results
in increased dietary energy availability for compensatory growth (Ferrell,
1988). It would appear equally plausible that the increased systemic amino
acid availability resulting from a reduction in intestinal mass and amino acid
metabolism also could lead to enhanced efficiency of dietary protein use and
growth rate, typical of compensatory growth.
5.2. Nutrient Composition
In addition to the quantity of dietary intake, the composition of dietary
nutrients can also substantially affect gut protein and amino acid metabolism. There is an extensive literature describing the impact of dietary
composition on gut growth and function (Spector et al., 1977; Weser
et al., 1986; Buts et al., 1990; Bragg et al., 1991; Jenkins and Thompson,
1994). However, considerably less is known about how the composition
of the diet alters gut protein metabolism. Some studies have shown that
restriction of dietary protein has limited effects on intestinal protein synthesis and growth (Seve et al., 1986; Ebner et al., 1994), whereas others have
reported a decrease in protein synthesis (McNurlan and Garlick, 1981;
Wykes et al., 1996). The study with neonatal pigs demonstrated that,
although protein malnutrition significantly reduced whole-body growth
and amino acid absorption, this was associated with reduced carcass growth,
since gut tissue growth was maintained (Ebner et al., 1994). A recent study
based on metabolism of enterally fed 13C-lysine showed that in pigs fed
low-protein diets, the PDV tissues utilized more than 75% of the lysine
intake, compared to 45% in high- protein-fed pigs (Van Goudoever et al.,
2000). Another important finding of this study was that on the high-protein
diet, all of the PDV lysine utilization was derived from the arterial circulation, whereas on the low-protein diet, the gut used lysine equally from the
enteral and arterial input. This study highlights the preferential use of dietary amino acids by the gut and how this limits the systemic availability of
lysine for lean tissue growth during protein malnutrition.
Studies focusing on the impact of dietary macronutrients found that enteral amino acids, but not carbohydrate or lipid, stimulated intestinal protein
synthesis, whereas each of these stimulated gut protein accretion, suggesting
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D. G. Burrin
that carbohydrate and lipid suppressed gut proteolysis (Stoll et al., 2000).
In contrast, in other studies, enteral amino acids rapidly decreased intestinal
protein synthesis and proteolysis in young pigs (Adegoke et al., 1999).
Feeding a high- fat versus high- carbohydrate diet to young pigs stimulated
intestinal protein synthesis (Ponter et al., 1994). Numerous studies have
examined the effect of dietary supplementation with specific amino acids
on intestinal growth and metabolism. Supplementing glutamine, glutamate,
and arginine has been shown to be stimulatory to the gut in some cases,
but not in others (Michail et al., 1995; Wu et al., 1996; Horvath et al.,
1996; Vanderhoof et al., 1997; Hasebe et al., 1999; Bertolo et al., 1999b;
Ewtushik et al., 2000). Dietary factors that stimulate mucosal proliferation
and cell turnover also tend to increase protein synthesis. Feeding fibre and
lectins have been shown to stimulate intestinal protein synthesis in some
cases (Southern et al., 1985; Palmer et al., 1987), but not in others (Nyachoti
et al., 2000). Studies show that neonatal pigs fed elemental diets have higher
rates of protein synthesis and cell proliferation than those fed cow’s milk
formula, even though intestinal mucosal growth and villus morphology
are similar (Stoll et al., 2000c).
6. INFECTION AND INFLAMMATION
Infestation with pathogenic microbial and viral organisms, or exposure to
the toxins they produce, is known to adversely affect both intestinal structure and function. Studies with growing animals indicate that exposure to
pathogenic and non-pathogenic organisms impart a protein metabolic cost
to the animal associated with stimulation and maintenance of the proinflammatory, acute-phase response (Von Allmen et al., 1992; MacRae,
1993; Higashiguchi et al., 1994; Johnson, 1997; Breuille et al., 1998; Wang
et al., 1998; Breuille et al., 1999; Mack et al., 1999). The studies demonstrate that treatment with pro-inflammatory stimuli such as bacteria, enteric
parasites, endotoxin and cytokines significantly increases protein synthesis
in visceral tissues, especially the liver, gut and spleen, while increasing catabolism and net loss of muscle protein body weight. Recent work in sheep
demonstrated that parasitic infection increases the rate of leucine utilization
and oxidation by PDV tissues, thereby reducing the systemic availability of
dietary amino acids by 20–30% (Yu et al., 2000). In addition, most of the
increased PDV leucine utilization is either oxidized or lost as endogenous
protein secretion; together, these losses account for most of the reduced
nitrogen retention associated with infection (Yu et al., 2000).
These results illustrate a mechanism whereby the gastrointestinal microflora activate the immune system and suppress the growth rate in domestic
Gastrointestinal protein and amino acid metabolism in growing animals
715
animals. Antimicrobial compounds are fed to domestic animals in order to
suppress the activity of the gut microflora and enhance growth; however, the
exact mechanism for this effect is unknown. It has been shown that by suppressing microbial activity, antimicrobials reduce the luminal concentration
and associated toxic insult of ammonia, and thereby diminish the thickness
and mass of the intestinal mucosa and associated lymphoid tissue (Visek,
1978). Studies in pigs and chickens show that feeding antimicrobial compounds significantly reduces small intestinal mass, cell proliferation and
intestinal ammonia absorption (Yen et al., 1987; Yen and Pond, 1990;
Krinke and Jamroz, 1996). Additional evidence indicates that much of the
luminal ammonia originates from bacterial hydrolysis of urea and deamination of dietary amino acids. Thus, given the recent evidence that the gut may
be an important site of dietary amino acid catabolism, the question arises as
to whether this activity is associated with the luminal microbes or the cell
populations of the mucosa. Although it is likely that both possibilities exist,
the end result is the catabolism and loss of dietary amino acids, especially
those that are essential, that otherwise would be used for growth.
7. FUTURE PERSPECTIVES
It is evident from past studies that the tissues of the gastrointestinal tract
play an integral role in the whole-body protein and amino acid metabolism
of growing animals. It has also become increasingly evident that the gut not
only consumes a substantial fraction of the dietary intake, but is critical for
the metabolism of key amino acids, such as arginine and threonine, that are
essential for the growth of developing animals. There has been considerable
progress in our understanding of intestinal amino acid metabolism, especially nonessential amino acids such as glutamate, glutamine, citrulline,
ornithine, and proline. However, with the exception of arginine and, to some
extent, threonine, there is only a very limited understanding of the cellular
basis for how other essential amino acids are metabolized within the gut tissues. With the expanding range of analytical tools, from the molecular to the
whole-organ level, there is a clear rationale and opportunity to explore these
issues in growing animals. For example, many of the biochemical pathways
responsible for oxidation of essential amino acids have either not been well
characterized or even shown to exist in intestinal tissues. Although it is well
established that glutamine and glutamate are major oxidative fuels for the
gut, the metabolic fate of nitrogen from these substrates is only vaguely
understood. There is also a compelling need to identify the cellular signaling mechanisms whereby nutrients and other extracellular molecules affect
the rates of protein and amino acid metabolism of intestinal cells. Many of
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D. G. Burrin
the pathways and cellular targets that have been shown to be important in
other cell types are obvious candidates (e.g., MAP kinases, PI3 kinase,
mTOR, proteasome, NF-kB). As we begin to establish the underlying cellular basis of protein metabolism, it should become increasingly evident as to
which specific nutrients are important for intestinal growth and function;
this will then lead to more rational diet formulation. In addition, continued
efforts should be aimed at establishing how regulatory elements, such as
nutrients, hormones and environmental factors, modulate the growth and
metabolism of the gut tissues, and then quantifying their impact on nutrient
availability and growth of the animal. Further studies that couple the use of
trans-organ substrate balance and isotopic kinetic measurements will be particularly useful in attempting to answer these questions. An especially crucial area will be the impact of the gut microflora, and the extent to which it
modifies gut amino acid utilization and the nitrogen economy of the animal.
REFERENCES
Adegoke, O.A., McBurney, M.I., Samuels, S.E., Baracos, V.E., 1999. Luminal amino acids
acutely decrease intestinal mucosal protein synthesis and protease mRNA in piglets.
J. Nutr. 129, 1871–1878.
Alpers, D.H., 1972. Protein synthesis in intestinal mucosa: the effect of route of administration of precursor amino acids. J. Clin. Invest. 51, 167–173.
Anthony, J.C., Anthony, T.G., Kimball, S.R., Jefferson, L.S., 2001. Signaling pathways
involved in translational control of protein synthesis in skeletal muscle by leucine.
J. Nutr. 131, 856S–860S.
Attaix, D., Manghebati, A., Grizard, J., Arnal, M., 1986. Assessment of in vivo protein synthesis
in lamb tissues with [3H] valine flooding doses. Biochim. Biophys. Acta 882, 389–397.
Attaix, D., Arnal, M., 1987. Protein synthesis and growth in the gastrointestinal tract of the
young preruminant lamb. Brit. J. Nutr. 58, 159–169.
Attaix, D., Meslin, J.C., 1991. Changes in small intestinal mucosa morphology and cell
renewal in suckling, prolonged-suckling, and weaned lambs. Amer. J. Physiol. 261,
R811–R818.
Attaix, D., Aurousseau, E., Rosolowska-Huszcz, D., Bayle, G., Arnal, M., 1992. In vivo
longitudinal variations in protein synthesis in developing ovine intestines. Amer. J. Physiol. 263, R1318–R1323.
Ball, R.O., Law, G., Bertolo, R.F.P., Pencharz, P.B., 1999. Adequate oral threonine is critical for
mucin production and mucosal growth by the neonatal piglet gut. VIIIth International Symposium on Protein Metabolism and Nutrition, Aberdeen, United Kingdom. (Abstr.), p. 31.
Baracos, V.E., Samuels, S.E., Adegoke, O.A., 2000. Anabolic and catabolic mediators of
intestinal protein turnover: a new experimental approach. Curr. Opin. Clin. Nutr. Metab.
Care 3, 183–189.
Battezzati, A., Brillon, D.J., Matthews, D.E., 1995. Oxidation of glutamic acid by the
splanchnic bed in humans. Amer. J. Physiol. 269, E269–E276.
Bertolo, R.F., Chen, C.Z., Law, G., Pencharz, P.B., Ball, R.O., 1998. Threonine requirement
of neonatal piglets receiving total parenteral nutrition is considerably lower than that of
piglets receiving an identical diet intragastrically. J. Nutr. 128, 1752–1759.
Gastrointestinal protein and amino acid metabolism in growing animals
717
Bertolo, R.F., Brunton, J.A., Pencharz, P.B., Ball, R.O., 1999a. Small intestinal metabolism
is necessary for the conversion between arginine and proline in orally fed piglets. VIIIth
International Symposium on Protein Metabolism and Nutrition, Aberdeen, United Kingdom. (Abstr.), p. 33.
Bertolo, R.F., Pencharz, P.B., Ball, R.O., 1999b. A comparison of parenteral and enteral
feeding in neonatal piglets, including an assessment of the utilization of a glutamine-rich,
pediatric elemental diet. J. Parent. Enter. Nutr. 23, 47–55.
Blachier, F., Darcy-Vrillon, B., Sener, A., Duee, P.H., Malaisse, W.J., 1991. Arginine metabolism in rat enterocytes. Biochim. Biophys. Acta 1092, 304–310.
Blachier, F., M’Rabet-Touil, H., Posho, L., Morel, M.T., Bernard, F., Darcy-Vrillon, B.,
Duee, P.H., 1992. Polyamine metabolism in enterocytes isolated from newborn pigs. Biochim. Biophys. Acta 1175, 21–26.
Blachier, F., M’Rabet-Touil, H., Posho, L., Darcy-Vrillon, B., Duee, P.H., 1993. Intestinal
arginine metabolism during development. Evidence for de novo synthesis of L-arginine
in newborn pig enterocytes. Eur. J. Biochem. 216, 109–117.
Blachier, F., Selamnia, M., Robert, V., M’Rabet-Touil, H., Duee, P.H., 1995. Metabolism of
L-arginine through polyamine and nitric oxide synthase pathways in proliferative or differentiated human colon carcinoma cells. Biochim. Biophys. Acta 1268, 255–262.
Bragg, L.E., Thompson, J.S., Rikkers, L.F., 1991. Influence of nutrient delivery on gut structure and function. Nutrition 7, 237–243.
Breuille, D., Arnal, M., Rambourdin, F., Bayle, G., Levieux, D., Obled, C., 1998. Sustained
modifications of protein metabolism in various tissues in a rat model of long-lasting sepsis. Clin. Sci. 94, 413–423.
Bruielle, D., Voisin, L., Contrepois, M., Arnal, M., Rose, F., Obled, C., 1999. A sustained rat
model for studying the long-lasting catabolic state of sepsis. Infec. Immunity. 67, 1079–1085.
Brunton, J.A., Bertolo, R.F.P., Pencharz, P.B., Ball, R.O., 1999. Proline ameliorates arginine
deficiency during enteral but not parenteral feeding in neonatal pigs. Amer. J. Physiol.
277, E223–E231.
Brunton, J.A., Ball, R.O., Pencharz, P.B., 2000. Current total parenteral nutrition solutions
for the neonate are inadequate. Curr. Opin. Clin. Nutr. Metab. Care 3, 299–304.
Burrin, D.G., Ferrell, C.L., Eisemann, J.H., Britton, R.A., Nienaber, J.A., 1989. Effect of
level of nutrition on splanchnic blood flow and oxygen consumption in sheep. Brit. J.
Nutr. 62, 23–34.
Burrin, D.G., Davis, T.A., Fiorotto, M.L., Reeds, P.J., 1991. Stage of development and fasting affect protein synthetic activity in the gastrointestinal tissues of suckling pigs. J. Nutr.
121, 1099–1108.
Burrin, D.G., Shulman, R.J., Reeds, P.J., Davis, T.A., Gravitt, K.R., 1992. Porcine colostrum
and milk stimulate visceral organ and skeletal muscle synthesis in neonatal piglets.
J. Nutr. 122, 1205–1213.
Burrin, D.G., Shulman, R.J., Langston, C., Storm, M.C., 1994. Supplemental alanylglutamine, organ growth and nitrogen metabolism in neonatal pigs fed by total parenteral
nutrition. J. Parent. Enter. Nutr. 18, 313–319.
Burrin, D.G., Davis, T.A., Ebner, S., Schoknecht, P.A., Fiorotto, M.L., Reeds, P.J., McAvoy,
S., 1995. Nutrient-independent and nutrient-dependent factors stimulate protein synthesis
in colostrum-fed newborn pigs. Pediat. Res. 37, 593–599.
Burrin, D.G., Fiorotto, M.L., Hadsell, D.L., 1999a. Transgenic hypersection of des(1-3)
human insulin-like growth factor I in mouse milk has limited effects on the gastrointestinal tract in suckling pups. J. Nutr. 129, 51–56.
Burrin, D.G., Stoll, B., Chang, X., Yu, H., Reeds, P.J., 1999b. Total parenteral nutrition does
not compromise digestion or absorption of dietary protein in neonatal pigs. FASEB J. 13,
A1024.
718
D. G. Burrin
Burrin, D.G., Stoll, B., Jiang, R., Chang, X., Hartmann, B., Holst, J.J., Greeley, G.H. Jr.,
Reeds, P.J., 2000. Minimal entral nutrient requirements for intestinal growth in neonatal
piglets: How much is enough? Amer. J. Clin. Nutr. 71, 1603–1610.
Burrin, D.G., 2001. Nutrient requirements for intestinal metabolism and growth in the neonatal pig. In: Lindberg, J.E. (Ed.), Digestive Physiology of the Pig Symposium, Proceedings of the 8th International Symposium on Digestive Physiology in Pigs., EAAP Publ.
No. 99, Uppsala, Sweden, pp. 75–88.
Bush, J.A., Nguyen, H.V., Suryawan, A., O’Connor, P.M.J., Burrin, D.G., Reeds, P.J., Liu,
C.W., Davis, T.A., 2001. Tissue-specific response of protein metabolism to somatotropin
treatment in growing pigs. FASEB J. 15, (Abstr.) A730.
Buts, J.P., Vijverman, V., Barudi, C., De Keyser, N., Maldague, P., Dive, C., 1990. Refeeding after starvation in the rat: comparative effects of lipids, proteins and carbohydrates on
jejunal and ileal mucosal adaptation. Eur. J. Clin. Invest. 20, 441–452.
Buts, J.P., De Keyser, N., De Raedemaeker, L., Collette, E., Sokal, E.M., 1995. Polyamine
profiles in human milk, infant artificial formulas, and semi-elemental diets. J. Pediat. Gastroenterol. Nutr. 21, 44–49.
Capano, G., Bloch, K.J., Carter, E.A., Dascoli, J.A., Schoenfeld, D., Harmatz, P.R., 1998.
Polyamines in human and rat milk influence intestinal cell growth in vitro. J. Pediat. Gastroenterol. Nutr. 27, 281–286.
Cappelli, F.P., Seal, C.J., Parker, D.S., 1997. Glucose and [13C] leucine metabolism by the
portal-drained viscera of sheep fed on dried grass with acute intravenous and intraduodenal infusions of glucose. Brit. J. Nutr. 78, 931–946.
Darcy-Vrillon, B., Posho, L., Morel, M-T., Bernard, F., Blachier, F., Meslin, J-C., Duee,
P-H., 1994. Glucose, galactose, and glutamine metabolism in pig isolated enterocytes
during development. Pediat. Res. 36, 175–181.
Darmaun, D., Roig, J.C., Auestad, N., Sager, B.K., Neu, J., 1997. Glutamine metabolism in
very low birth weight infants. Pediat. Res. 41, 391–396.
Davis, T.A., Nguyen, H.V., Garcia-Bravo, R., Fiorotto, M.L., Jackson, E.M., Lewis, D.S.,
Lee, D.R., Reeds, P.J., 1994. Amino acid composition of human milk is not unique.
J. Nutr. 124, 1126–1132.
Davis, T.A., Burrin, D.G., Fiorotto, M.L., Nguyen, H.V., 1996. Protein synthesis in skeletal
muscle and jejunum is more responsive to feeding in 7- than in 26-day-old pigs. Amer. J.
Physiol. 270, E802–E809.
Davis, T.A., Nguyen, H.V., Suryawan, A., Bush, J.A., Jefferson, L.S., Kimball, S.R., 2000.
Developmental changes in the feeding-induced stimulation of translation initiation in
muscle of neonatal pigs. Amer. J. Physiol. 279, E1226–E1234.
DeJonge, W.J., Dingemanse, M.A., de Boer, P.A., Lamers, W.H., Moorman, A.F., 1998.
Arginine-metabolizing enzymes in the developing rat small intestine. Pediat. Res. 43,
442–451.
Dudley, M.A., Burrin, D.G., Wykes, L.J., Toffolo, G., Cobelli, C., Nichols, B.L., Rosenberger, J., Jahoor, F., Reeds, P.J., 1998. Protein kinetics determined in vivo with a multipletracer, single-sample protocol: application to lactase synthesis. Amer. J. Physiol. 274,
G591–G598.
Dufour, C., Dandrifosse, G., Forget, P., Vermesse, F., Romain, N., Lepoint, P., 1988. Spermine
and spermidine induce intestinal maturation in the rat. Gastroenterology 95, 112–116.
Dugan, M.E.R.., Knabe, D.A., Wu, G., 1994. Glutamine and glucose metabolism in intraepithelial lymphocytes from pre- and post-weaning pigs. Comp. Biochem. Physiol. 109B,
675–681.
Dugan, M.E.R., Knabe, D.A., Wu, G., 1995. The induction of citrulline synthesis from glutamine in enterocytes of weaned pigs is not due primarily to age or change in diet. J. Nutr.
125, 2388–2393.
Gastrointestinal protein and amino acid metabolism in growing animals
719
Ebner, S., Schoknecht, P., Reeds, P.J., Burrin, D.G., 1994. Growth and metabolism of gastrointestinal and skeletal muscle tissues in protein-malnourished neonatal pigs. Amer. J.
Physiol. 266, R1736–R1743.
Ewtushik, A.L., Bertolo, R.F.P., Ball, R.O., 2000. Intestinal development of early-weaned
piglets receiving diets supplemented with selected amino acids or polyamines. Can. J.
Anim. Sci. 80, 653–662.
Fan, M.Z., Stoll, B., Jiang, R., Burrin, D.G., 2001. Enterocyte digestive enzyme activity
along the crypt-villus and longitudinal axes in the neonatal pig small intestine. J. Anim.
Sci. 79, 371–381.
Ferrell, C.L., 1988. Contribution of visceral organs to animal energy expenditures. J. Anim.
Sci. 66, 23–34.
Fleming, S.E., Fitch, M.D., DeVries, S., Liu, M.L., Kight, C., 1991. Nutrient utilization by
cells isolated from rat jejunum, cecum, and colon. J. Nutr. 121, 869–878.
Ganessunker, D., Gaskins, H.R., Zuckermann, F.A., Donovan, S.M., 1999. Total parenteral
nutrition alters molecular and cellular indices of intestinal inflammation in neonatal piglets. J. Parent. Enter. Nutr. 23, 337–344.
Gate, J.J., Parker, D.S., Lobley, G.E., 1999. The metabolic fate of the amido-N group of glutamine in the tissues of the gastrointestinal tract in 24 h-fasted sheep. Brit. J. Nutr. 81,
297–306.
Goldspink, D.F., Lewis, S.E.M., Kelly, F.J., 1984. Protein synthesis during the developmental growth of the small and large intestine of the rat. Biochem. J. 217, 527–534.
Goldspink, D.F., Kelly, F.J., 1984. Protein turnover and growth in the whole body, liver and
kidney of the rat from the foetus to senility. Biochem. J. 217, 507–516.
Goldstein, R.M., Hebiguchi, T., Luk, G.D., Taqi, F., Guilarte, T.R., Franklin, F.A. Jr., Niemiec, P.W., Dudgeon, D.L., 1985. The effects of total parenteral nutrition on gastrointestinal growth and development. J. Pediat. Surg. 20, 785–791.
Grant, A.L., Thomas, J.W., King, K.J., Liesman, J.S., 1990. Effects of dietary amines on small
intestinal variables in neonatal pigs fed soy protein isolate. J. Anim. Sci. 68, 363–371.
Hahn, P., Taller, M., Chan, H., 1988. Pyruvate carboxylase, phosphate-dependent glutaminase and glutamate dehydrogenase in the developing rat small intestinal mucosa. Biol.
Neonate 53, 362–366.
Haisch, M., Fukagawa, N.K., Matthews, D.E., 2000. Oxidation of glutamine by the splanchnic bed in humans. Amer. J. Physiol. 278, E593–E602.
Haque, S.M., Chen, K., Usui, N., Iiboshi, Y., Okuyama, H., Masunari, A., Cui, L., Nezu, R.,
Takagi, Y., Okada, A., 1996. Alanyl-glutamine dipeptide-supplemented parenteral nutrition improves intestinal metabolism and prevents increased permeability in rats. Ann.
Surg. 223, 334–341.
Hasebe, M., Suzuki, H., Mori, E., Furukawa, J., Kobayashi, K., Ueda, Y., 1999. Glutamate in
enteral nutrition: can glutamate replace glutamine in supplementation to enteral nutrition
in burned rats? J. Parent. Enter. Nutr. 23, S78S82.
Higashiguchi, T., Hasselgren, P.O., Wagner, K., Fischer, J.E., 1993. Effect of glutamine on
protein synthesis in isolated intestinal epithelial cells. J. Parent. Enter. Nutr. 17, 307–314.
Higashiguchi, T., Noguchi, Y., O’Brien, W., Wagner, K., Fischer, J.E., Hasselgren, P.O.,
1994. Effect of sepsis on mucosal protein synthesis in different parts of the gastrointestinal tract in rats. Clin. Sci. 87, 207–211.
Horvath, K., Jami, M., Hill, I.D., Papadimitriou, J.C., Magder, L.S., Chanasongcram, S.,
1996. Isocaloric glutamine-free diet and the morphology and function of rat small intestine. J. Parent. Enter. Nutr. 20, 128–134.
Humbert, B., LeBacquer, O., Nguyen, P., Dumon, H., Darmaun, D., 2001. Protein restriction
and dexamethasone as a model of protein hypercatabolism in dogs: effect of glutamine on
leucine turnover. Metabolism 50, 293–298.
720
D. G. Burrin
Hurwitz, R., Kretchmer, N., 1986. Development of arginine-synthesizing enzymes in mouse
intestine. Amer. J. Physiol. 251, G103–G110.
Inoue, Y., Grant, J.P., Snyder, P.J., 1993. Effect of glutamine-supplemented total parenteral
nutrition on recovery of the small intestine after starvation atrophy. J. Parent. Enter. Nutr.
17, 165–170.
Jenkins, A.P., Thompson, R.P.H., 1994. Mechanisms of small intestinal adaptation. Digest.
Dis. 12, 15–27.
Jiang, R., Chang, X., Stoll, B., Ellis, K.J., Shypailo, R.J., Weaver, E., Campbell J., Burrin,
D.G., 2000. Dietary plasma protein is used more efficiently than extruded soy protein
for lean tissue growth in early-weaned pigs. J. Nutr. 130, 2016–2019.
Johnson, R.W., 1997. Inhibition of growth by pro-inflammatory cytokines: An integrated
view. J. Anim. Sci. 75, 1244–1255.
Kahn, J., Iiboshi, Y., Cui, L., Wasa, M., Sando, K., Takagi, Y., Okada, A., 1999. Alanylglutamine-supplemented parenteral nutrition increases luminal mucus gel and decreases
permeability in the rat small intestine. J. Parent. Enter. Nutr. 23, 24–31.
Kandil, H.M., Argenzio, R.A., Chen, W., Berschneider, H.M., Stiles, A.D., Westwick, J.K.,
Rippe, R.A., Brenner, D.A., Rhoads, J.M., 1995. L-glutamine and L-asparagine stimulate
ODC activity and proliferation in a porcine jejunal enterocyte line. Amer. J. Physiol. 269,
G591–G599.
Kight, C.E., Fleming, S.E., 1995. Oxidation of glucose carbon entering the TCA cycle is
reduced by glutamine in small intestine epithelial cells. Amer. J. Physiol. 268, G879–G888.
Kimball, S.R., Horetsky, R.L., Jefferson, L.S., 1998. Signal transduction pathways involved
in the regulation of protein synthesis by insulin in L6 myoblasts. Amer. J. Physiol. 274,
C221–C228.
Kimball, S.R., Jefferson, L.S., Nguyen, H.V., Suryawan, A., Bush, J.A., Davis, T.A., 2000.
Feeding stimulates protein synthesis in muscle and liver of neonatal pigs through an
mTOR-dependent process. Amer. J. Physiol. 279, E1080–E1087.
Klimberg, V.S., 1996. Glutamine, cancer, and its therapy. Amer. J. Surg. 172, 418–424.
Krinke, A.L., Jamroz, D., 1996. Effects of feed antibiotic avoparcine on organ morphology in
broiler chickens. Poultry Sci. 75, 705–710.
Lien, K.A., Sauer, W.C., Fenton, M., 1997. Mucin output in ileal digesta of pigs fed a
protein-free diet. Z. Ernahrungswiss 36, 182–190.
Lobley, G.E., Milne, V., Lovie, J.M., Reeds, P.J., Pennie, K., 1980. Whole body and tissue
protein synthesis in cattle. Brit. J. Nutr. 43, 491–502.
Lobley, G.E., Harris, P.M., Skene, P.A., Brown, D., Milne, E., Calder, A.G., Anderson, S.E.,
Garlick, P.J., Nevison, I., Connell, A., 1992. Responses in tissue protein synthesis to suband supra-maintenance intake in young growing sheep: comparison of large-dose and
continuous-infusion techniques. Brit. J. Nutr. 68, 373–388.
Mack, D.R., Michail, S., Wei, S., McDougall, L., Hollingsworth, M.A., 1999. Probiotics
inhibit enteropathogenic E. coli adherence in vitro by inducing intestinal mucin gene
expression. Amer. J. Physiol. 39, G941–G950.
MacRae, J.C., 1993. Metabolic consequences of intestinal parasitism. Proceedings of the
Nutrition Society 52, 121–130.
Madej, M., Lundh, T., Lindberg, J.E., 1999. Activities of enzymes involved in glutamine
metabolism in connection with energy production in the gastrointestinal tract epithelium
of newborn, suckling and weaned piglets. Biol. Neonate 75, 250–258.
Marchini, J.S., Nguyen, P., Deschamps, J-Y., Maugere, P., Krempf, M., Darmaun, D., 1999.
Effect of intratvenous glutamine on duodenal mucosa protein synthesis in healthy growing dogs. Amer. J. Physiol. 276, E747–E753.
Martensson, J., Jain, A., Meister, A., 1991. Glutathione is required for intestinal function.
Proc. Nat. Acad. Sci. USA, 87, 1715–1719.
Gastrointestinal protein and amino acid metabolism in growing animals
721
McNurlan, M.A., Tomkins, A.M., Garlick, P.J., 1979. The effect of starvation on the rate of
protein-synthesis in rat liver and small intestine. Biochem. J. 178, 371–379.
McNurlan, M.A., Garlick, P.J., 1981. Protein synthesis in liver and small intestine in protein
deprevation and diabetes. Amer. J. Physiol. 241, E238–E245.
Merry, B.J., Lewis, S.E., Goldspink, D.F., 1992. The influence of age and chronic
restricted feeding on protein synthesis in the small intestine of the rat. Exp. Gerontol.
27, 191–200.
Michail, S., Mohammadpour, H., Park, J.H., Vanderhoof, J.A., 1995. Effect of glutaminesupplemented elemental diet on mucosal adaptation following bowel resection in rats.
J. Pediat. Gastroenterol. Nutr. 21, 394–398.
Montagne, L., Toullee, R., Lalles, J.P., 2000. Calf intestinal mucin: isolation, partial characterization, and measurement in ileal digesta with an enzyme-linked immunosorbent assay.
J. Dairy Sci. 83, 507–517.
Mordrelle, A., Jullian, E., Costa, C., Cormet-Boyaka, E., Benamouzig, R., Rome, D.,
Huneau, J.F., 2000. EAAT1 is involved in transport of L-glutamate during differntiation
of the Caco-2 cell line. Amer. J. Physiol. 279, G366–G373.
Morgan, W. 3d, Yardley, J., Luk, G., Niemiec, P., Dudgeon, D., 1987. Total parenteral nutrition and intestinal development: A neonatal model. J. Pediatr. Surg. 22, 541–545.
Mouille, B., Morel, E., Robert, V., Guihot-Joubrel, G., Blachier, F., 1999. Metabolic capacity
for L-citrulline synthesis from ammonia in rat isolated colonocytes. Biochim. Biophys.
Acta 24, 401–407.
Murphy, J.M., Murch, S.J., Ball, R.O., 1996. Proline is synthesized from glutamate during
intragastric infusion but not during intravenous infusion in neonatal piglets. J. Nutr.
26, 878–286.
Naka, S., Saito, H., Hashiguchi, Y., Lin, M.T., Furukawa, S., Inaba, T., Fukushima, R.,
Wada, N., Muto, T., 1997. Alanylglutamine-enriched total parenteral nutrition improves
protein metabolism more than branched chain amino acid-enriched total parenteral
nutrition in protracted peritonitis. J. Trauma 42, 183–190.
Nieto, R., Lobley, G.E., 1999. Integration of protein metabolism within the whole body and
between organs. In: Lobely, G.E., White, A., MacRae, J.C. (Eds.), Proceedings of the 8th
International Symposium on Protein Metabolism and Nutrition. EAAP Publication No.
96, 69–99.
Nyachoti, C.M., de Lange, C.F., McBride, B.W., Leeson, S., Gabert, V.M., 2000. Endogenous gut nitrogen losses in growing pigs are not caused by increased protein synthesis rates
in the small intestine. J. Nutr. 130, 566–572.
Palmer, R.M., Pusztal, A., Bain, P., Grant, G., 1987. Changes in rates of tissue protein synthesis in rats induced in vivo by consumption of kidney bean lectins. Comp. Biochem.
Physiol. 88C 179–183.
Papaconstantinou, H.T., Hwang, K.O., Rajaraman, S., Hellmich, M.R., Townsend, C.M. Jr.,
Ko, T.C., 1998. Glutamine deprivation induces apoptosis in intestinal epithelial cells. Surgery 124, 152–159.
Patureau-Mirand, P., Mosoni, L., Levieux, D., Attaix, D., Bayle, G., Bonnet, Y., 1990. Effect
of colostrum feeding on protein metabolism in the small intestine of newborn lambs. Biol.
Neonate 57, 30–36.
Pell, J.M., Caldarone, E.M., Bergman, E.N., 1986. Leucine and-ketoisocaproate metabolism
and interconversion in fed and fasted sheep. Metabolism 35, 1005–1016.
Platell, C., McCauley, R., McCulloch, R., Hall, J., 1993. The influence of parenteral glutamine and branched-chain amino acids on total parenteral nutrition-induced atrophy of the
gut. J. Parent. Enter. Nutr. 17, 348–354.
Pluske, J.R., Hampson, D.J., Williams, I.H., 1997. Factors influencing the structure and function of the small intestine in the weaned pig: a review. Livest. Prod. Sci. 51, 215–236.
722
D. G. Burrin
Pollack, P.F., Koldovsky, O., Nishioka, K., 1992. Polyamines in human and rat milk and in
infant formulas. Amer. J. Clin. Nutr. 56, 371–375.
Ponter, A.A., Cortamira, N.O., Seve, B., Salter, D.N., Morgan, L.M., 1994. The effects of
energy source and tryptophan on the rate of protein synthesis and on hormones of the
entero-insular axis in the piglet. Brit. J. Nutr. 71, 661–674.
Quan, J., Fitch, M.D., Flemming, S.E., 1998. Rate at which glutamine enters TCA cycle
influences carbon atom fate in intestinal epithelial cells. Amer. J. Physiol. 275,
G1299–G1308.
Reeds, P.J., Burrin, D.G., Davis, T.A., Fiorotto, M.L., 1993. Postnatal growth of gut and
muscle: competitors or collaborators. Proc. Nutr. Soc. 52, 57–67.
Reeds, P.J., Burrin, D.G., Jahoor, F., Wykes, L., Henry, J., Frazer, E.M., 1996. Enteral glutamate is almost completely metabolized in first pass by the gastrointestinal tract of infant
pigs. Amer. J. Physiol. 270, E413–E418.
Reeds, P.J., Burrin, D.G., Stoll, B., Jahoor, F., Wykes, L., Henry, J., Frazer, M.E., 1997. Enteral glutamate is the preferential source for mucosal glutathione synthesis in fed piglets.
Amer. J. Physiol. 273, E408–E415.
Reeds, P.J., Burrin, D.G., Stoll, B., van Goudoever, J.B., 1999. Consequences and regulation
of gut metabolism. In: Lobley, G.E., White, A., MacRae, J.C. (Eds.), Protein Metabolism
and Nutrition. EAAP Publication No. 96. pp.127–153.
Rhoads, M., 1999. Glutamine signaling in intestinal cells. J. Parent. Enter. Nutr. 23, S38–
S40.
Rhoads, J.M., Argenzio, R.A., Chen, W., Graves, L.M., Licato, L.L., Blikslager, A.T.,
Smith, J., Gatzy, J., Brenner, D.A., 2000. Glutamine metabolism stimulates intestinal cell
MAPKs by a cAMP-inhibitable, Raf-independent mechanism. Gastroenterology 118,
90–100.
Rouse, K., Nwokedi, E., Woodliff, J.E., Epstein, J., Klimberg, V.S., 1995. Glutamine
enhances selectivity of chemotherapy through changes in glutathione metabolism. Ann.
Surg. 221, 420–426.
Samuels, S.E., Taillandier, D., Aurousseau, E., Cherel, Y., Maho, L., Arnal, M., Attaix, D.,
1996. Gastrointestinal tract protein synthesis and mRNA levels for proteolytic systems in
adult fasted rats. Amer. J. Physiol. 271, E232–238.
Seidel, E.R., Haddox, M.K., Johnson, L.R., 1985. Ileal mucosal growth during intraluminal
infusion of ethylamine or putrescine. Amer. J. Physiol. 249, G434–G438.
Selamnia, M., Robert, V., Mayeur, C., Delpal, S., Blachier, F., 1998. De novo synthesis of
arginine and ornithine from citrulline in human colon carcinoma cells: metabolic fate of
L-ornithine. Biochim. Biophys. Acta 1425, 93–102.
Seve, I., Reeds, P.J., Fuller, M.F., Cadenhead, A., Hay, S.M., 1986. Protein synthesis and
retention in some tissues of the young pig as influenced by dietary protein intake after
early-weaning. Possible connection to the energy metabolism. Reprod. Nutr. Develop.
26, 849–861.
Shenoy, V., Roig, J.C., Chakrabarti, R., Kubilis, P., Neu, J., 1996. Ontogeny of glutamine
synthetase in rat small intestine. Pediat. Res. 39, 643–648.
Shoveller, A.K., Ball, R.O., Pencharz, P.B., 2000. The methionine requirement is 35% lower
during parenteral versus oral feeding in neonatal piglets. FASEB J. 14, A558.
Shoveller, A.K., Pencharz, P.B., Ball, R.O., 2001. First pass metabolism affects methionine
(MET) requirement but not the proportion spared by cysteine (CYS) in intravenously (IV)
and intragastrically (IG) fed piglets. FASEB J. 15, A265.
Smith, M.W., Jarvis, L.G., 1978. Growth and cell replacement in the new-born pig intestine.
Proc. R. Soc. Lond. B. Biol. Sci. 203, 69–89.
Smith, R.J., 1990. Glutamine metabolism and its physiologic importance. J. Parent. Enteral.
Nutr. 14, 40S–44S.
Gastrointestinal protein and amino acid metabolism in growing animals
723
Souba, W.W., Klimberg, V.S., Plumley, D.A., Salloum, R.M., Flynn, T.C., Bland, K.I.,
Copeland, E.M. 3d., 1990. The role of glutamine in maintaining a healthy gut and supporting the metabolic response to injury and infection. J. Surg. Res. 48, 383–391.
Southon, S., Livesey, G., Gee, J.M., Johnson, I.T., 1985. Differences in intestinal protein
synthesis and cellular proliferation in well-nourished rats consuming conventional laboratory diets. Brit. J. Nutr. 53, 87–95.
Spaeth, G., Gottwald, T., Haas, W., Holmer, M., 1993. Glutamine peptide does not improve
gut barrier function and mucosal immunity in total parenteral nutrition. J. Parent. Enter.
Nutr. 17, 317–323.
Spector, M.H., Levine, G.M., Deren, J.J., 1977. Direct and indirect effects of dextrose and
amino acids on gut mass. Gastroenterology 72, 706–710.
Stein, T.P., Yoshida, S., Schluter, M.D., Drews, D., Assimon, S.A., Leskiw, M.D., 1994.
Comparison of intravenous nutrients on gut mucosal protein synthesis. J. Parent. Enter.
Nutr. 18, 447–452.
Stoll, B., Henry, J., Reeds, P.J., Yu, H., Jahoor, F., Burrin, D.G., 1998. Catabolism dominates
the first-pass intestinal metabolism of dietary essential amino acids in milk protein-fed
piglets. J. Nutr. 128, 606–614.
Stoll, B., Burrin, D.G., Henry, J., Yu, H., Jahoor, F., Reeds, P.J., 1999a. Substrate oxidation
by the portal drained viscera of fed piglets. Amer. J. Physiol. 277, E168–E175.
Stoll, B., Burrin, D.G., Henry, J., Jahoor, F., Reeds, P.J., 1999b. Dietary and systemic phenylalanine utilization for mucosal and hepatic constitutive protein synthesis in pigs.
Amer. J. Physiol. 276, G49–G57.
Stoll, B., Chang, X., Fan, M.Z., Reeds, P.J., Burrin, D.G., 2000a. Enteral nutrient intake
determines the rate of intestinal protein synthesis and accretion in neonatal pigs. Amer.
J. Physiol. 279, G288–G294.
Stoll, B., Chang, S., Jiang, R., Van Goudoever, J.B., Reeds, P.J., Burrin, D.G., 2000b. Enteral
carbohydrate and lipid inhibit small intestinal proteolysis in neonatal pigs. FASEB J. 14,
A558.
Stoll, B., Price, P., Reeds, P.J., Henry, J., Burrin, D.G., 2000c. Feeding an elemental diet versus a milk-based formula does not decrease intestinal mucosal growth in infant pigs.
J. Pediat. Gastroenterol. Nutr. 31, S169.
Szondy, Z., Newsholme, E.A., 1990. The effect of various concentrations of nucleobases,
nucleosides or glutamine on the incorporation of [3H]thymidine into DNA in rat mesenteric-lymph-node lymphocytes stimulated by phytohaemagglutinin. Biochem. J. 270,
437–440.
Tamada, H., Nezu, R., Imamura, I., Matsuo, Y., Takagi, Y., Kamata, S., Okada, A., 1992.
The dipeptide alanyl-glutamine prevents intestinal mucosal atrophy in parenterally fed
rats. J. Parent. Enter. Nutr. 16, 110–116.
Taupin, D., Podolsky, D.K., 1999. Mitogen-activated protein kinase activation regulates
intestinal epithelial differentiation. Gastroenterology 116, 1072–1080.
Van Goudoever, J.B., Stoll, B., Henry, J.F., Burrin, D.G., Reeds, P.J., 2000. Adaptive regulation of intestinal lysine metabolism. Proc. Natl. Acad. Sci. 97, 11620–11625.
Van Klinken, B.J.-W., Dekker, J., Büller, H.A., de Bolòs, C., Einerhand, A.W.C., 1997.
Biosynthesis of mucins (MUC2-6) along the longitudinal axis of the human gastrointestinal tract. Amer. J. Physiol. 273, G296–G302.
Vanderhoof, J.A., Kollman, K.A., Griffin, S., Adrian, T.E., 1997. Growth hormone and glutamine do not stimulate intestinal adaptation following massive small bowel resection in
the rat. J. Pediat. Gastroenterol. Nutr. 25, 327–331.
Van der Schoor, S.R.D., van Goudoever, J.B., Stoll, B., Henry, J.H., Rosenberger, J., Burrin,
D.B., Reeds, P.J., 2001. The pattern of intestinal substrate oxidation is altered by protein
restriction in pigs. Gastroenterology 121, 67–1175.
724
D. G. Burrin
Visek, W.J., 1978. The mode of growth promotion by antibiotics. J. Anim. Sci. 4,
1447–1469.
Von Allmen, D., Hasselgren, P-O., Higashiguchi, T., Frederick, J., Zamir, O., Fischer, J.E.,
1992. Increased intestinal protein synthesis during sepsis and following the administration
of tumour necrosis factor ‘‘or interleukin-1’’. Biochem. J. 286, 585–589.
Wakabayashi, Y., Yamada, E., Yoshida, T., Takahashi, N., 1995. Effect of intestinal resection and arginine-free diet on rat physiology. Amer. J. Physiol. 269, G313–G318.
Wang, Q., Meyer, T.A., Boyce, S.T., Wang, J.J., Sun, X., Tiao, G., Fischer, J.E., Hasselgren,
P.-O., 1998. Endotoxemia in mice stimultes production of complement C3 and serum
amyloid jA in mucosa of small intestine. Amer. J. Physiol. 275, R1584–R1592.
Wang, Q., Wang, X., Hernandez, A., Kim, S., Evers, B.M., 2001. Inhibition of the phosphatidylinositol 3-kinase pathway contributes to HT29 and Caco-2 intestinal cell differentiation. Gastroenterology 120, 1065–1066.
Wang, J-Y, Viar, M.J., Blanner, P.M., Johnson, L.R., 1996. Expression of the ornithine decarboxylase gene in response to asparagine in intestinal epithelial cells. Amer. J. Physiol.
271, G164–G171.
Waterlow, J.C., Garlick, P.J., Millward, D.J., 1978. Protein Turnover in Mammalian Tissues
and in the Whole Body. Elsevier, Amsterdam.
Weser, E., Babbitt, J., Hoban, M., Vandeventer, A., 1986. Intestinal adaptation. Different
growth resopnses to disaccharides compared with monosaccharides in rat small bowel.
Gastroenterology 91, 1521–1527.
Windmueller, H., Spaeth, A.E., 1980. Respiratory fuels and nitrogen metabolism in vivo in
small intestine of fed rats. Quantitative importance of glutamine, glutamate, and aspartate. J. Biol. Chem. 255, 107–112.
Windmueller, H.G., Spaeth, A.E., 1975. Intestinal metabolism of glutamine and glutamate
from the lumen as compared to glutamine from blood. Arch. Biochem. Biophys. 171,
662–672.
Wu, G., 1995. Urea synthesis in enterocytes of developing pigs. Biochem. J. 312, 717–723.
Wu, G., Knabe, D.A., Yan, W., Flynn, N.E., 1995. Glutamine and glucose metabolism in
enterocytes of the neonatal pig. Amer. J. Physiol. 268, R334–R342.
Wu, G., Meier, S.A., Knabe, D.A., 1996. Dietary glutamine supplementation prevents jejunal
atrophy in weaned pigs. J. Nutr. 126, 2578–2584.
Wu, G., 1997. Synthesis of citrulline and arginine from proline in enterocytes of postnatal
pigs. Amer. J. Physiol. 272, G1382–G1390.
Wu, G., 1998. Intestinal mucosal amino acid catabolism. J. Nutr. 128, 1249–1252.
Wu, G., Morris, S.M. Jr., 1998. Arginine metabolism: nitric oxide and beyond. Biochem. J.
336, 1–17.
Wu, G., Flynn, N.E., Knabe, D.A., 2000a. Enhanced intestinal synthesis of polyamines from
proline in cortisol-treated piglets. Amer. J. Physiol. 279, E395–E402.
Wu, G., Flynn, N.E., Knabe, D.A., Jaeger, L.A., 2000b. A cortisol surge mediates the
enhanced polyamine synthesis in procine enterocytes during weaning. Amer. J. Physiol.
279, R554–R559.
Wu, G., Haynes, T.E., Li, H., Yan, W., Meininger, C.J., 2001. Glutamine metabolism to glucosamine is necessary for glutamine inhibition of endothelial nitric oxide synthesis.
Biochem. J. 353, 245–252.
Wykes, L.J., Fiorotto, M., Burrin, D.G., Del Rosario, M., Frazer, M.E., Pond, W.G., Jahoor,
F., 1996. Chronic low protein intake reduces tissue protein synthesis in a pig model of
protein malnutrition. J. Nutr. 126, 1481–1488.
Yen, J.T., Nienaber, J.A., Pond, W.G., 1987. Effect of neomycin, carbadox and length of
adaptation to calorimeter on performance, fasting metabolism and gastrointestinal tract
of young pigs. J. Anim. Sci. 65, 1243–1248.
Gastrointestinal protein and amino acid metabolism in growing animals
725
Yen, J.T., Nienaber, J.A., Hill, D.A., Pond, W.G., 1989. Oxygen consumption by portal veindrained organs and by whole animal in conscious growing swine. Proc. Soc. Exp. Biol.
Med. 190, 393–398.
Yen, J.T., Pond, W.G., 1990. Effect of carbadox on net absorption of ammonia and glucose
into hepatic protal vein of growing pigs. J. Anim. Sci. 68, 4236–4242.
Yu, F., Bruce, L.A., Calder, A.G., Milne, E., Coop, R.L., Jackson, F., Horgan, G.W.,
MacRae, J.C., 2000. Subclinical infection with the nematode Trichostrongylus colubriformis increases gastrointestinal tract leucine metabolism and reduces availability of leucine
for other tissues. J. Anim. Sci. 78, 380–390.
Yu, Y.-M., Burke, J.F., Vogt, J.A., Chambers, L., Young,V.R., 1992. Splanchnic and whole
body L-[1-13C,15N] leucine kinetics in relation to enteral and parenteral amino acid supply. Amer. J. Physiol. 262, E687–E694.