Laboratoire de Biologie, Conservatoire National de Arts et Métiers, 75003 Paris, France
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ABSTRACT |
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The aim of the present study was to evaluate the effect of cholera toxin on energy balance from intestinal glutamine metabolism and oxidation, glutamine-dependent sodium absorption, and cholera toxin-dependent ion flux. Cholera toxin-stimulated sodium and L-glutamine ileal transport and metabolism were studied in Ussing chambers. Glutamine (10 mM) transport and metabolism were simultaneously studied using 14C flux and HPLC. In the same tissues, the flux of each amino acid was studied by HPLC, and glutamine metabolism and oxidation were studied by the determination of amino acid specific activity and 14CO2 production. In control tissues, glutamine stimulated sodium absorption and was mainly oxidized. The transepithelial flux of intact glutamine represented 45% of glutamine flux across the luminal membrane. The other metabolites were glutamate and, to a lesser degree, citrulline, ornithine, and proline. Cholera toxin did not alter glutamine-stimulated sodium absorption, glutamine oxidation, transport, and metabolism. In conclusion, the present results indicate that cholera toxin does not alter glutamine intestinal function and metabolism. In addition, ~95% of the energy provided by glutamine oxidation remains available to the enterocyte.
electrolyte transport; ATP balance; intestinal mucosa; P5C pathway; rehydration solution
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INTRODUCTION |
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CHOLERA IS OFTEN REGARDED by physiologists as an interesting model for studying the response of intestinal mucosa to a well-defined toxin, cholera toxin (CT). The water and electrolyte intestinal secretion that result from CT challenge have been extensively studied (4). In experimental and human cholera, the relationship between electrolyte and water fluxes across the intestine remains identical to control conditions (5, 32). In addition, the findings that the function of the glucose-Na+ cotransporter at the luminal membrane of the epithelial layer is enhanced by CT (24, 31) provided an interesting explanation for the clinical efficacy of the oral rehydration solution (ORS) recommended by the World Health Organization (35). The World Health Organization ORS contains a concentration of electrolytes approximately equivalent to that lost in stools of patients with severe cholera (90 mmol/l Na+ and 20 mmol/l K+) and 110 mmol/l glucose to stimulate intestinal Na+ absorption (9, 36).
However, cholera remains a widespread and life-threatening disease for several reasons, including the lack of an effective immunization program and a frequent association between cholera and poor nutritional status (36). The treatment of patients with dehydration and malnutrition is particularly complex because the acute metabolic alterations of acute dehydration are superimposed on the long-lasting metabolic disturbances of chronic malnutrition. The present recommendation is to first rehydrate with an ORS, called ReSoMal, that takes into account the amount of water and electrolyte lost in stools and the chronic deficit in K+, Mg2+, and micronutrients and then to feed the patient as soon as possible (36). However, when feeding is initiated, nutrient intestinal absorption may not be optimal as a consequence of malnutrition and partial rehydration (2, 3). At the rehydration phase, except for glucose used to stimulate rehydration, no other nutrient has proved to be functionally effective and useful in the intestinal functions.
Glutamine may be considered a possible candidate to add to ORS for several reasons: 1) it stimulates intestinal Na+ absorption, as demonstrated in an in vitro isolated intestine model (7, 23, 26, 28) and in vivo intestinal perfusion in adult cholera patients in Bangladesh (32); 2) it is also an important source of energy for the intestinal mucosa in animals and humans (6, 30, 33); and 3) it is involved in many important metabolic processes that may favor intestinal repair in malnutrition (15). However, it is not known how much of the energy that glutamine provides to the intestine is used for the stimulation of electrolyte transport and its own metabolism. Also, it is not known whether CT that stimulates electrolyte secretion also alters glutamine metabolism. In addition, most metabolic studies of glutamine are not conducted under conditions that are relevant to a situation where glutamine is used for its functional properties in intestinal Na+ absorption.
The aim of the present study was therefore to evaluate the effect of CT on energy balance from intestinal glutamine metabolism and oxidation, glutamine-dependent Na+ absorption, and CT-dependent ion flux alteration. To do so, we coupled transport studies in Ussing chambers that have proven effective in understanding the nutrient and electrolyte intestinal transport alteration due to CT and HPLC with isotopic tracer methods to identify the glutamine metabolites in isolated rabbit ileum mucosa. Thus the aim was not to duplicate previous studies of the effect of glutamine on Na+ absorption or glutamine metabolism (7, 23, 26, 28, 32, 34). Rather, we wanted to link these approaches to estimate the glutamine-derived energy that is available to the enterocyte after energy expenditure for metabolism and transport.
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MATERIALS AND METHODS |
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Animals. Before death, healthy male weanling New Zealand White rabbits (établissement CEGAV, Saint Mars d'Egrenne, France) were fed with 13% protein standard formula (Lapin entretien 112; UAR, Epinay sur Orge, France). Rabbit weight was 2.15 ± 0.12 kg. They were fasted 18 h before anesthesia with 1 ml/kg of 3% (wt/vol) pentobarbital sodium. Small bowel was washed with 37°C Ringer solution. Two 15-cm-long loops were made in ileum. Intestine was cut between the two loops to block the CT effect via the submucosal nerve plexus. Ringer solution (9 ml) was introduced in each loop, and 20 µg of CT (Sigma, Saint Louis, MO) was added in one of them. After 3 h, rabbits were killed using pentobarbital (2 ml/kg).
Electrical measurements.
For each animal, measurements were assessed in 4-8 controls and
4-8 cholera tissues. Ileal mucosa was mounted between two half-chambers, exposing an area of 1.13 cm2, and bathed at
37°C with 12 ml of oxygenated Ringer buffer containing (in mM) 140 Na+, 5.2 K+, 1.2 Ca2+, 1.2 Mg2+, 120 Cl, 25 HCO
3, 2.4 HPO2
4 and 0.4 H2PO
4. After 30 min, 10 mM
L-glutamine was added to the mucosal side, with equimolar
mannitol added to the serosal side. Potential difference (PD) across
the tissue was measured by calomel electrodes in saturated KCl using
agar bridges positioned near the surface of the tissue. By Ag-AgCl
electrodes connected to the solution via agar bridges, the tissue was
continuously short-circuited with automatic voltage clamps (World
Precision Instruments, Sarasota, FL) that compensated for the fluid
resistance. Ileal sheets were mounted in the chambers within 30 min of
rabbit death and continuously short-circuited, except for a 10-s
interval every 15 min, when the open-circuit PD was measured.
Conductance was calculated from change in current following an imposed
voltage using Ohm's law. An increase in short-circuit
current (Isc) corresponds to cation transepithelial
absorption or anion secretion.
Ion flux measurements.
The transepithelial unidirectional Na+ and
Cl fluxes from mucosa to serosa and from serosa to
mucosa were determined in six rabbits and were both studied in the same
period. For each animal, measurements were assessed in eight control
and eight cholera tissues. The stability of the electrical parameters
was checked for at least 15 min; 1 µCi of
22Na and 2 µCi of 36Cl were then added to the
appropriate reservoir, and a 10-min equilibration period was allowed to
elapse before unidirectional fluxes were determined for five 10-min
periods. After 60 min, 10 mM L-glutamine was
introduced into the mucosal compartment and 10 mM mannitol was
introduced into the serosal side. Electrical parameters and
unidirectional fluxes were determined for three 20-min periods. The
22Na fluxes were measured using a gamma counter (Kontron),
and the 36Cl fluxes were measured using a liquid
scintillation spectrometer (SL 4000 IN Intertechnique PG 4000).
Glutamine transport and metabolism. Associated with 10 mM L-glutamine (Sigma), 2 µCi glutamine uniformly labeled with 14C (NEN, Boston, MA) was added to the mucosal compartment. The 14C flux across the epithelium representing the overall flux of [14C]glutamine and 14C metabolites was therefore expressed as glutamine equivalent flux. Samples were taken every 30 min in the serosal compartment. The transfer of radioactivity was measured using a liquid scintillation spectrometer. For each sample, two 10-min countings were done. The mean of these two countings was used for flux calculation using the counts per minute accumulation method, taking into account a constant quenching (23).
The glutamine oxidative metabolism produced CO2 that was not included in the glutamine equivalent flux. The oxidative metabolism was therefore estimated by trapping CO2 produced in the two compartments in a series of two tubes containing 1 M NaOH (5 ml). Radioactivity was estimated using the liquid scintillation spectrometer (1-ml sample associated with 4-ml scintillator). The 14CO2 production was calculated from the 14C counted in NaOH and the specific activity of [14C]glutamine, uniformly labeled on its five carbons, as checked by HPLC coupled with liquid scintillation. The fluxes of glutamine metabolites were assessed using a liquid chromatograph with ultraviolet detector (Shimadzu, Kyoto, Japan) coupled with a fraction collector (Foxy 200 ISCO; Isco, Lincoln, NE). Because glutamine and metabolites (glutamate, alanine, ornithine, citrulline, and proline) do not absorb in ultraviolet or in visible light, they were covalently bound to a chromophore. The derivatization reagent was dimethylaminoazobenzene sulfonyl chloride (DABSYL-Cl) with maximal absorbency at 436 nm. Samples (100 µl) were reacted for 10 min at 70°C with 50 µl NaHCO3 (150 mM, pH 8.6) and 150 µl DABSYL-Cl (8 mM; Fluka, Buchs, Switzerland) in acetone. Dilution was made with 300 µl KH2PO4 (12.5 mM, pH 3) and ethanol (50/50 vol/vol). Fifty microliters were injected (Fig. 1). The column was a C8 Kromasil type (250 mm × 45 mm, 5 µm particle size; Eka Chemicals, Bohus, Sweden). Oven temperature was 40°C. Mobile phase was composed of 12.5 mM KH2PO4, pH 3 (solvent A) and acetonitrile 75/25 vol/vol (Prolabo, Paris, France)/isopropanol (Merck, Darmstadt, Germany) (solvent B). Flow rate was 1.3 ml/min. The gradient was 10% of solvent B at 0 min, 30% of solvent B at 2 min, 30% of solvent B at 10 min, 45% of solvent B at 20 min, 55% of solvent B at 43 min, 70% of solvent B at 44 min, 70% of solvent B at 47 min, and 10% of solvent B at 48 min. Run time was 53 min. No other component was eluted at the retention time of glutamine, citrulline, glutamate, alanine, proline, or ornithine. To verify repeatability, a mixture of the six amino acids was derivated 10 times, with a relative standard deviation range from 2.1 to 4.0%. The method was linear in a range of 20-200% of the concentration observed for a flux of 1 µmol · h
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Statistical analysis. Data were analyzed using SAS software (SAS, Cary, NC). Data were summarized as means ± SE with n = number of rabbits. The P values correspond to a general linear model analysis.
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RESULTS |
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Control tissue.
In control tissues, addition of 10 mM glutamine to the mucosal
compartment was followed by an immediate and steady rise in Isc (2.2 ± 0.22 µmol · h1 · cm
2;
Table 1). After 2 h, further addition of 10 mM glucose was again followed by an additional rise in
Isc (2.1 ± 0.59 µmol · h
1 · cm
2;
Table 2). After both substrate additions,
transepithelial conductance increased by 38 and 26% for glutamine and
glucose, respectively.
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Effect of CT.
In CT-treated loops, water secretion was observed after 3 h (7.0 ± 0.72 ml) and water absorption was observed in control Ringer loops
(4.1 ± 0.94 ml). When loops were mounted in Ussing chambers,
Isc was significantly higher (P < 0.004)
in CT than control tissue (Table 1). The conductance was essentially
identical in both conditions. Glutamine equally stimulated
Isc in CT and in control tissues, but conductance
increase was significantly more pronounced in CT than control tissues
(P < 0.001). After 2 h, further addition of 10 mM glucose was
again followed by an additional rise in Isc (2 ± 0.17 µmol · h
1 · cm
2).
After both substrate additions, transepithelial conductance increased
by 83 and 39% for glutamine and glucose, respectively (Tables
1 and 2).
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DISCUSSION |
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The present study confirms that glutamine stimulates electrolyte absorption in control and CT-treated tissues. In both conditions, glucose displays an additional effect on electrolyte absorption. In addition, the present results indicate that CT does not alter effects of glutamine on intestinal function and metabolism. On the basis of an estimate of ATP used for glutamine- and CT-stimulated electrolyte transport and glutamine metabolism, compared with the ATP produced from the glutamine oxidation pathway, we conclude that ~95% of the energy provided by glutamine oxidation remains available to the enterocyte for basic cellular function and repair, leading to increased intestinal nutrient absorption (see below). It is now established that glutamine stimulates Na+ absorption by two mechanisms: an electrogenic and a neutral absorption. However, the relative distribution is unsettled.
Our data show that in control and CT experiments glutamine stimulated
both a neutral NaCl and an electrogenic Na+ absorption in a
2-to-1 ratio. Rhoads et al. (28) found that glutamine promoted 3.5 times more electroneutral than electrogenic Na+ absorption
in the jejunum of 1- to 3-wk-old piglets. In piglet rotavirus
enteritis, glutamine stimulated equal amounts of electrogenic and
electroneutral NaCl absorption (26). In experimental
Cryptosporidium infection in piglet (1),
L-glutamine stimulated neutral NaCl absorption 1.3 times
more than electrogenic Na+ absorption. The magnitude of
stimulation of neutral NaCl absorption in the infected ileum exceeded
that in control tissue, whereas electrogenic Na+ absorption
was smaller. Nath et al. (23) demonstrated that glutamine enhanced
electrogenic Na+ absorption in healthy and diarrheagenic
Escherichia coli-infected rabbits and induced a small (10% of
the electrogenic absorption) electroneutral NaCl absorption in infected
rabbits. In summary, the relative magnitude of the two
glutamine-stimulated Na+ absorptive processes varied from
0.10 to 3.5 (neutral/electrogenic Na+ absorption) according
to animal species, age, or pathological conditions. In addition, in our
experiments glutamine stimulated a large residual electrogenic flux. It
may be related to the large CO2 production (2.2 µmol · h1 · cm
2)
from glutamine metabolism. Whatever the final interpretation of the
isotopic fluxes and electrical parameters, our results indicate that CT
did not alter the net fluxes; it increased the conductance and the
unidirectional fluxes.
For the calculation of energy expenditure related to
glutamine-stimulated Na+ absorption, we used the following:
in control conditions (Fig. 2), 1.31 µmol · h1 · cm
2
glutamine entered the enterocyte with 0.8 µmol · h
1 · cm
2
Na+ as neutral flux and 0.59 µmol · h
1 · cm
2
as electrogenic Na+ flux; in CT (Fig.
3), 1.4 µmol · h
1 · cm
2
glutamine entered with 0.8 µmol · h
1 · cm
2
Na+ as neutral flux and 0.4 µmol · h
1 · cm
2
as electrogenic Na+ flux.
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The ATP cost calculation was based on the observation that
Na+-K+-ATPase is the main energy-driven
transport process on which all electrolyte transport is dependent. The
well-known stoichiometry of 3 Na+ expelled from the cell
for 2 K+ entering at the expense of one ATP provides a
reasonable estimate of energy used for transport (17). In control
conditions, the presence of 10 mM glutamine at the luminal membrane was
associated with an increase of 1.39 µmol · h1 · cm
2
Na+ absorption that hydrolyzed 0.46 µmol · h
1 · cm
2
ATP (Fig. 2). In CT-treated intestine, 0.3 µmol · h
1 · cm
2
Na+ entered the cell associated with 0.6 µmol · h
1 · cm
2
Cl
through basolateral
Na+-K+-2Cl
transport; it was
then expelled through the Na+-K+-ATPase, which
hydrolyzed an additional 0.1 µmol · h
1 · cm
2 ATP.
From a methodological point of view, we did not intend to describe the
different metabolic steps; rather, we clearly identified the oxidative
pathway. Although measured exclusively in the serosal compartment, the
-pyrroline-5-carboxylate (P5C) pathway metabolites were identified
in very small quantity. Most studies found such a metabolism, except
one in rabbit ileum (23). The unexpected findings of that study may be
explained by the methodology used, in which metabolism was measured by
comparing [15N]glutamine and
[14C]glutamine. However, the measurements were
not performed on the same piece of intestine. Thus the variance due to
the variable "rabbit" or "tissue" may have masked the
difference related to metabolism.
It has previously been reported that the rate of glutamine metabolism
and oxidation depends on the intraluminal glutamine concentration (20,
33). About 70% of the glutamine transported across the brush border
was not metabolized in the intestine when the intraluminal glutamine
concentration was 45 mM, but only 34% was transported intact to the
serosal side when the concentration was 6 mM (33). Our results are in
agreement with these results because 48% of glutamine was not
metabolized when the intraluminal concentration was 10 mM. The reason
for increased intact glutamine transport with increasing glutamine
luminal concentration is to be found in the kinetic parameters of the
metabolic and transport processes. In agreement with this hypothesis,
we found that 0.33 ± 0.08 µmol · h1 · cm
2
of [14C]glutamine metabolites were present
before intact glutamine was transported (Fig.
4). In addition, a transepithelial
diffusional pathway is frequently observed with increasing luminal
concentration (8). Thus it seems that for a luminal concentration as
low as ~0.5-1 mM, the energy provided by glutamine has reached a
maximal value. Above this threshold value, glutamine that is not
metabolized is transported intact to the blood side.
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We do not have quantitative determination of the concentration of glutamine in the human intestinal lumen after a meal or a glutamine-containing oral rehydration solution. However, human perfusion studies indicate that jejunal absorptive capacity of glutamine and glucose is essentially the same in control and cholera conditions (8, 22, 32). Direct measurement of glucose concentration in rat small intestinal lumen after a meal indicated a mean value of 0.2-24 mM; it ranged with time and small intestinal region from 0.2 mM to a maximum of 48 mM (11). Thus if glutamine luminal concentration varies in a similar range, the energy derived from glutamine in the intestinal cell may not be very different from what we measured using 10 mM glutamine.
For the energy produced by glutamine metabolism, we counted that one
mole of ATP was used for each mole of glutamate produced. The
production of glutamate was estimated to be 0.68 µmol · h1 · cm
2,
assuming that all the difference between glutamine entering the cell
(1.31 µmol · h
1 · cm
2)
minus the glutamine found in the serosal compartment (0.63 µmol · h
1 · cm
2)
has been deaminated to glutamate (Fig. 2)
The other metabolic pathway is the P5C synthetase pathway leading to ornithine, citrulline, and proline synthesis (34). In terms of energy consumption it may be regarded as negligible, because the metabolic flux is very low. However, the P5C pathway is functionally important, as exemplified by the report concerning two brothers with severe clinical symptoms related to P5C synthase deficiency (19).
Windmueller and Spaeth (34) demonstrated that [14C]glutamine synthesis leads to labeled lactate acid, suggesting that part of the glutamine oxidation is incomplete. Stoll et al. (30) indicated that this pathway is more important with glucose than with glutamate or glutamine, as observed by the presence of lactate or alanine. Our results are in agreement with these studies; despite the 14CO2 production (complete oxidation), a small amount of [14C]alanine was present, indicating that metabolically significant quantities of glutamine carbons were transformed to pyruvate in enterocytes and that its oxidation was thereby incomplete.
The ATP production derived from glutamine metabolism was essentially
derived from 14CO2 production, using the
stoichiometric ratio of 2 µmol · h1 · cm
2
CO2 for 9 µmol · h
1 · cm
2
ATP. This low rate of oxidation would take into account the incomplete glutamine oxidation. The same figure has been used to take into account
the probability that a molecule entering the tricarboxylic acid cycle
will be oxidized in rat intestinal cells (12). In addition, the
glutaminase activity hydrolyzed 0.68 µmol · h
1 · cm
2.
We did not use other correcting factors that are probably small and
uncertain in these experimental conditions (12). Thus in both control
(Fig. 2) and CT (Fig. 3) intestine, the flux of ATP production was
~11
µmol · h
1 · cm
2.
The main objective of the study was to evaluate the effect of CT on energy balance from glutamine metabolism and transport functions. Our results strongly suggest that CT does not alter or minimally alters the energy balance derived from glutamine; in control conditions, the estimated ATP production was 9.8 for 0.46 used for transport, i.e., 95% of the glutamine-derived energy remains available to the mucosa (Fig. 2). Very similar figures were obtained in CT-treated tissues (Fig. 3).
It is likely that the present figures obtained in an isolated piece of intestine mounted in an Ussing chamber are lower than in in vivo conditions. Clearly, the tissue is deprived of its blood supply, and, although oxygenated at pH 7.4 and circulated at 37°C, Ringer solution in the mucosal and serosal compartments does not match the luminal fluid and blood circulation. However, in these well-defined conditions it was possible to study both glutamine metabolism and electrolyte transport in the same piece of tissue. In addition to stimulating Na+ absorption, glutamine may also provide energy in CT-treated intestine for other useful functions, including synthesis of key molecules such as glutathione and nucleotides, protein synthesis, epithelial repair, and improvement of barrier function (10, 13, 16, 21, 27).
Finally, we used glutamine as the only energetic substrate in Ringer solution; glutamine was considered as an ingredient for ORS rather than a nutrient provided with food. In addition to stimulating Na+ absorption, our results indicate that glutamine given during the rehydration period may have a beneficial effect on intestinal function by providing energy directly to the gut epithelium. These experiments may provide a rationale to test the effect of glutamine in the rehydration period on the absorptive capacity of the intestine during the subsequent feeding period.
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FOOTNOTES |
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* The first and second authors contributed equally to this work. The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: M. Abely, Laboratoire de Biologie, Conservatoire National des Arts et Métiers, 2 rue Conté, 75003 Paris, France (E-mail: abely{at}cnam.fr).
Received 8 June 1999; accepted in final form 16 December 1999.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Argenzio, RA,
Rhoads JM,
Armstrong M,
and
Gomez G.
Glutamine stimulates prostaglandin-sensitive Na(+)-H(+) exchange in experimental porcine cryptosporidiosis.
Gastroenterology
106:
1418-1428,
1994[ISI][Medline].
2.
Butzner, JD,
Butler DG,
Miniats OP,
and
Hamilton JR.
Impact of chronic protein-calorie malnutrition on small intestinal repair after acute viral enteritis: a study in gnotobiotic piglets.
Pediatr Res
19:
476-481,
1985[Abstract].
3.
Butzner, JD,
and
Gall DG.
Refeeding enhances intestinal repair during an acute enteritis in infant rabbits subjected to protein-energy malnutrition.
Pediatr Res
29:
594-600,
1991[Abstract].
4.
Crowe, SE,
and
Powell DW.
Fluid and electrolyte transport during enteric infections.
In: Infections of the Gastrointestinal Tract, edited by Blaser MJ,
Smith PD,
Ravdin JI,
Greenberg HB,
and Guerrant RL.. New York: Raven, 1995, p. 107-141.
5.
Curran PF. Solute-solvent interactions and water transport.
Role of Membranes in Secretory Processes edited by Bolis
L, Keynes RD, and Wilbrantdt W. Amsterdam: 1972, p. 409-419.
6.
Darmaun, D,
Messing B,
Just B,
Rongier M,
and
Desjeux JF.
Glutamine metabolism after small intestinal resection in humans.
Metabolism
40:
42-44,
1991[ISI][Medline].
7.
Dechelotte, P,
Darmaun D,
Rongier M,
and
Desjeux JF.
Glutamine transport in isolated rabbit ileal epithelium.
Gastroenterol Clin Biol
13:
816-821,
1989[ISI][Medline].
8.
Dechelotte, P,
Darmaun D,
Rongier M,
Hecketsweiler B,
Rigal O,
and
Desjeux JF.
Absorption and metabolic effects of enterally administered glutamine in humans.
Am J Physiol Gastrointest Liver Physiol
260:
G677-G682,
1991
9.
Desjeux, JF,
Briend A,
and
Butzner JD.
Oral rehydration solution in the year 2000: pathophysiology, efficacy and effectiveness.
Baillieres Clin Gastroenterol
11:
509-527,
1997[ISI][Medline].
10.
Dugan, ME,
and
McBurney MI.
Luminal glutamine perfusion alters endotoxin-related changes in ileal permeability of the piglet.
JPEN J Parenter Enteral Nutr
19:
83-87,
1995[Abstract].
11.
Ferraris, RP,
Yasharpour S,
Lloyd KC,
Mirzayan R,
and
Diamond JM.
Luminal glucose concentrations in the gut under normal conditions.
Am J Physiol Gastrointest Liver Physiol
259:
G822-G837,
1990
12.
Fleming, SE,
Zambell KL,
and
Fitch MD.
Glucose and glutamine provide similar proportions of energy to mucosal cells of rat small intestine.
Am J Physiol Gastrointest Liver Physiol
273:
G968-G978,
1997
13.
Gianotti, L,
Alexander JW,
Gennari R,
Pyles T,
and
Babcock GF.
Oral glutamine decreases bacterial translocation and improves survival in experimental gut-origin sepsis.
JPEN J Parenter Enteral Nutr
19:
69-74,
1995[Abstract].
14.
Hahn, P,
Taller M,
and
Chan H.
Pyruvate carboxylase, phosphate-dependent glutaminase and glutamate deshydrogenase in the developing rat small intestinal mucosa.
Biol Neonate
53:
362-366,
1988[ISI][Medline].
15.
Hall, JC,
Heel K,
and
McCauley R.
Glutamine.
Br J Surg
83:
305-312,
1996[ISI][Medline].
16.
Higashiguchi, T,
Hasselgren PO,
Wagner K,
and
Fischer JE.
Effect of glutamine on protein synthesis in isolated intestinal epithelial cells.
JPEN J Parenter Enteral Nutr
17:
307-314,
1993[Abstract].
17.
Horisberger, JD,
Lemas V,
Kraehenbühl-P J,
and
Rossier BC.
Structure-function relationship of Na,K-ATPase.
Annu Rev Physiol
53:
565-584,
1999[ISI][Medline].
18.
James, LA,
Lunn PG,
Middleton S,
and
Elia M.
Distribution of glutaminase and glutamine synthetase activities in the human gastrointestinal tract.
Clin Sci
94:
313-319,
1998[ISI][Medline].
19.
Kamoun, P,
Aral B,
and
Saudubray JM.
A new inherited metabolic disease: delta1-pyrroline 5-carboxylate synthetase deficiency.
Bull Acad Natl Med
182:
131-137,
1998[ISI][Medline].
20.
Kight, CE,
and
Fleming SE.
Nutrient oxidation by rat intestinal epithelial cells is concentration dependent.
J Nutr
123:
876-882,
1993[ISI][Medline].
21.
Li, J,
King BK,
Janu PG,
Renegar KB,
and
Kudsk KA.
Glycyl-L-glutamine-enriched total parenteral nutrition maintains small intestine gut-associated lymphoid tissue and upper respiratory tract immunity.
JPEN J Parenter Enteral Nutr
22:
31-36,
1998[Abstract].
22.
Modigliani, R,
and
Bernier JJ.
Absorption of glucose, sodium, and water by the human jejunum studied by intestinal perfusion with a proximal occluding balloon and at variable flow rates.
Gut
12:
184-193,
1971[ISI][Medline].
23.
Nath, SK,
Dechelotte P,
Darmaun D,
Gotteland M,
Rongier M,
and
Desjeux JF.
[15N] and [14C]glutamine fluxes across rabbit ileum in experimental bacterial diarrhea.
Am J Physiol Gastrointest Liver Physiol
262:
G312-G318,
1992
24.
Nath, SK,
Rautureau M,
Heyman M,
Reggio H,
L'Helgoualc'h A,
and
Desjeux JF.
Emergence of Na+-glucose cotransport in an epithelial secretory cell line sensitive to cholera toxin.
Am J Physiol Gastrointest Liver Physiol
256:
G335-G341,
1989
25.
Newsholme, EA,
and
Carrie AL.
Quantitative aspects of glucose and glutamine metabolism by intestinal cells.
Gut
35 Suppl:
S13-S17,
1994[ISI][Medline].
26.
Rhoads, J,
Keku O,
Quinn J,
Woosely J,
and
Lecce J.
L-Glutamine stimulates jejunal sodium and chloride absorption in pig rotavirus enteritis.
Gastroenterology
100:
683-691,
1991[ISI][Medline].
27.
Rhoads, JM,
Argenzio RA,
Chen W,
Rippe RA,
Westwick JK,
Cox AD,
Berschneider HM,
and
Brenner DA.
L-Glutamine stimulates intestinal cell proliferation and activates mitogen-activated protein kinases.
Am J Physiol Gastrointest Liver Physiol
272:
G943-G953,
1997
28.
Rhoads, JM,
Keku EO,
Bennett LE,
Quinn J,
and
Lecce JG.
Development of L-glutamine-stimulated electroneutral sodium absorption in piglet jejunum.
Am J Physiol Gastrointest Liver Physiol
259:
G99-G107,
1990
29.
Riby, JE,
Hurwitz RE,
and
Kretchmer N.
Development of ornithine metabolism in the mouse intestine.
Pediatr Res
28:
261-265,
1990[Abstract].
30.
Stoll, B,
Burrin DG,
Hung Yu JH,
Jahoor F,
and
Reeds P.
Substrate oxidation by the portal drained viscera of fed piglets.
Am J Physiol Endocrinol Metab
277:
E168-E175,
1999
31.
Tai, YH,
Perez E,
and
Desjeux JF.
Cholera toxin and cyclic AMP stimulate D-glucose absorption in rat ileum.
In: Ion Gradient-Coupled Transport, edited by Alvarado F,
and van Os CH.. Amsterdam: Elsevier, 1986, p. 403-406.
32.
Van Loon, FP,
Banik AK,
Nath SK,
Patra FC,
Wahed MA,
Darmaun D,
Desjeux JF,
and
Mahalanabis D.
The effect of L-glutamine on salt and water absorption: a jejunal perfusion study in cholera in humans.
Eur J Gastroenterol Hepatol
8:
443-448,
1996[ISI][Medline].
33.
Windmueller, H.
Glutamine utilization by the small intestine.
Adv Enzymol Relat Areas Mol Biol
53:
201-237,
1982[ISI][Medline].
34.
Windmueller, HG,
and
Spaeth AE.
Intestinal metabolism of glutamine and glutamate from the lumen as compared to glutamine from blood.
Arch Biochem Biophys
171:
662-672,
1975[ISI][Medline].
35.
World Health Organization.
A manual for the Treatment of Acute Diarrhea for Use by Physicians and Other Senior Health Workers. Geneva: World Health Organization, 1984, p. p.72.
36.
World Health Organization.
Management of Severe Malnutrition: a Manual for Physicians and Other Health Workers. Geneva: World Health Organization, 1999, p. 60.