Substrate oxidation by the portal drained viscera of fed
piglets
Barbara
Stoll,
Douglas G.
Burrin,
Joseph
Henry,
Hung
Yu,
Farook
Jahoor, and
Peter J.
Reeds
United States Department of Agriculture, Agricultural Research
Station, Children's Nutrition Research Center, Department of
Pediatrics, Baylor College of Medicine, Houston, Texas 77030
 |
ABSTRACT |
Fully fed piglets
(28 days old, 7-8 kg) bearing portal, arterial, and gastric
catheters and a portal flow probe were infused with enteral
[U-13C]glutamate
(n = 4), enteral
[U-13C]glucose
(n = 4), intravenous
[U-13C]glucose
(n = 4), or intravenous
[U-13C]glutamine
(n = 3). A total of 94% of the
enteral
[U-13C]glutamate but
only 6% of the enteral [U-
13C]glucose was utilized in
first pass by the portal-drained viscera (PDV). The PDV extracted 6.5%
of the arterial flux of
[U-13C]glucose and
20.4% of the arterial flux of
[U-13C]glutamine. The
production of
13CO2
(percentage of dose) by the PDV from enteral glucose (3%), arterial
glucose (27%), enteral glutamate (52%), and arterial glutamine (70%)
varied widely. The substrates contributed 15% (enteral glucose), 19%
(arterial glutamine), 29% (arterial glucose), and 36% (enteral
glutamate) of the total production of
CO2 by the PDV. Enteral glucose
accounted for 18% of the portal alanine and 31% of the portal lactate
carbon outflow. We conclude that, in vivo, three-fourths of the energy
needs of the PDV are satisfied by the oxidation of glucose, glutamate,
and glutamine, and that dietary glutamate is the most important single
contributor to mucosal oxidative energy generation.
gut metabolism; amino acids; glucose; stable isotopes
 |
INTRODUCTION |
THE VISCERAL TISSUES, especially the intestinal mucosa,
pancreas, and spleen, have high rates of metabolism. As a consequence, the contribution of the portal-drained viscera (PDV) to whole body
energy expenditure is much larger than their contribution to body
weight. Understanding the substrates that are used to support this high
oxidative activity is therefore of considerable physiological and
nutritional importance.
The most extensive information on intestinal carbon metabolism in vivo
is contained in a series of publications based on the use of isolated,
in situ, vascularly perfused loops of rat small intestine (26-31).
The authors (31) concluded that, in animals that had been fed up to the
initiation of the tracer infusions, aspartate, glutamate, and glutamine
were the major oxidative substrates used by the small intestinal mucosa
and that glucose was a minor oxidative substrate. The observations of
Windmueller and Spaeth (26-31) with regard to mucosal
glutamine metabolism have received the greatest attention and have
served as the starting point for many investigations of the effects of
glutamine on intestinal growth, metabolism, and function (e.g., Ref.
3). However, their observation (28, 31) that the metabolism of enteral
glutamic acid was of much more quantitative importance to mucosal
energy generation than that of glutamine has been largely overlooked. With the notable exception of their observations on glutamate, studies
in vitro (7, 25, 32) have generally confirmed the findings of
Windmueller and Spaeth, although some recent results (8) imply that the
relative utilization of different substrates by isolated enterocytes
may be a function of their relative concentrations in the extracellular phase.
This last observation is important, because in the fed state, the
intestinal mucosa are presented with a complex mixture of arterial and
luminal substrates, and it is not known with certainty whether, under
fed conditions, the mucosal cells utilize some substrates in preference
to others. In other words, it is not known whether Windmueller and
Spaeth's observations reflected the nature of the experimental
preparation or whether the results revealed a specific feature of
mucosal intermediary metabolism. The main objective of the present work
was to attempt to provide an answer to this question.
We have developed a portal catheterized piglet model (6) to study
visceral metabolism in growing, conscious, and fed animals. In past
studies, we have used this model, in combination with enteral infusions
of uniformly 13C-labeled tracers,
to quantify the first-pass utilization of dietary amino acids (16, 17,
21-23). The results of one of these studies (17) confirmed
previous observations of virtually complete first-pass utilization of
dietary glutamate (14, 28, 31) and aspartate (29) by the intestinal
tissues. However, our studies were confined to the measurement of
portal tracer balance and mucosal protein synthesis, and although we
inferred that a high proportion of glutamate and glutamine utilization
was directed to oxidation, no direct measurements were made.
The present paper reports the results of a series of experiments in
which we have measured the oxidative metabolism of
U-13C-labeled glucose, glutamate,
and glutamine by the PDV. The main objective was to quantify their
relative utilization in the synthesis of alanine and lactate and their
contribution to CO2 production. On
the basis of previous results (28, 31) and of our own (17, 18), we
hypothesized that enteral glutamic acid (metabolized in first pass by
the mucosa) would be the single largest contributor to PDV oxidative metabolism.
 |
METHODS |
The protocol received prior approval by the Animal Care and Use
Committee of Baylor College of Medicine. Animal care conformed to
current US Department of Agriculture guidelines.
Feeding and Surgery
All the studies were carried out in female crossbred (Large White × Landrace × Duroc) piglets obtained from the Texas
Department of Criminal Justice (Huntsville, TX). They were received at
the Children's Nutrition Research Center when they were 14 days old, and for the next 10 days they were fed a liquid milk replacer diet
(Litter Life, Merrick, Middleton, WI). This was given at a
rate of 50 g dry
diet · kg
1 · day
1
and provided ~920 kJ and 12.5 g
protein · kg
1 · day
1.
After this preliminary period, the animals were fasted overnight and
were surgically implanted with gastric, portal, jugular, and carotid
catheters (6, 17, 21, 22). An ultrasonic flow probe (model 6S or 6R,
Transonics, Ithaca, NY) was implanted around the common portal vein.
The animals that received enteral tracer glucose were additionally
implanted with a duodenal catheter placed ~4 cm distal to the pyloric
sphincter. The animals were allowed to recover from surgery for a
minimum of 5 days or until they had achieved presurgery weight gain for
2 days. During this time they continued to receive liquid Litter Life
at the presurgery rate.
Tracer Protocol
[U-13C]glutamate (91%
[13C5]glutamate)
was synthesized and donated by Ajinomoto (Tokyo, Japan).
[U-13C]glucose (92%
[13C6]glucose)
and [U-13C]glutamine
(94%
[13C5]glutamine)
were purchased from Cambridge Isotopes (Woburn, MA). The rates
(µmol · kg
1 · h
1)
of tracer (i.e., the U-13C
isotopomer) administration were for intragastric
[U-13C]glutamate, 33 ± 2; for intraduodenal and intravenous
[U-13C]glucose, 173 ± 17; and for intravenous
[U-13C]glutamine, 29 ± 1.
At the time of the tracer infusion, the animals weighed 7.6 ± 0.6 kg. After an overnight fast, baseline samples of portal and
arterial blood were obtained, and the animals were offered a single
meal of one-twelfth of their preceding daily intake. This meal served
to initiate an adequate pattern of gastric emptying. One hour later, a
constant intragastric infusion of liquid Litter Life was
commenced and continued for the next 6 h. The diet infusion provided
nutrients at the same daily rate as the animals had consumed over the
preceding 3 days. In the animals that received an enteral [U-13C]glutamate
infusion, the tracer was mixed with the diet, whereas enteral
[U-13C]glucose was
given intraduodenally. The intravenous and intraduodenal infusions were
started at the same time as the diet infusion. Animals were killed with
an intra-arterial injection of pentobarbital sodium (50 mg/kg body wt)
and sodium phenytoin (5 mg/kg; Beutanasia-D; Schering-Plough Animal
Health, Kenilworth, NJ) after 6 h of tracer infusion.
During the infusion, arterial and portal blood samples (3 ml) were
taken every hour for 5 h and then at 15-min intervals over the last
hour. Blood gases (Chiron Diagnostics, Halstead, Essex, UK), glucose,
and lactate (YSI analyzer, Yellow Springs, OH) were determined
immediately in all blood samples. An aliquot of whole blood (0.5 ml)
was immediately placed in an evacuated tube (5 ml capacity) for
subsequent analysis of the isotopic enrichment of blood bicarbonate,
and two aliquots (1 ml) were immediately frozen for subsequent
measurements of amino acid, glucose and lactate concentrations, and
labeling. The remaining blood was centrifuged (3,000 g, 10 min at 4°C), and the plasma
was frozen in liquid nitrogen until taken for the analysis of its
ammonia concentration.
Sample Analysis
Plasma ammonia was measured with a clinical analyzer (COBOS FARA, Roche
Diagnostics, Nutley, NJ) with the manufacturer's standard protocol.
Blood samples were prepared for amino acid analysis and mass
spectrometry as described previously (10, 21-23). Gas chromatography-mass spectrometry was performed with the pentaacetate derivative of glucose, the pentafluorobenzyl derivative of lactate, and
the heptafluorobutyramide derivative of the amino acids. The analyses
were performed with a 5890 series II gas chromatograph linked to a
model 5989B (Hewlett-Packard, Palo Alto, CA) quadrupole mass
spectrometer. We used methane negative chemical ionization for amino
acids and lactate and methane positive chemical ionization for glucose.
To estimate the isotopic enrichment of blood bicarbonate, 0.5 ml of
perchloric acid (1 mol/l) was injected into the evacuated tube
containing the blood sample. The contents of the tube were mixed on a
vortex mixer, and the head space was carefully removed with a 5-ml
syringe. The gas sample was then injected into a second evacuated tube
(15 ml volume). The isotopic enrichment of the CO2 was then determined on a
continuous flow gas isotope ratio machine (ANCA, Europa Instruments,
Crewe, UK). The between-sample standard deviation was 0.001 atoms
percent excess (1
).
Calculations
The crude ion spectra were converted to tracer-to-tracee ratios (mol
per 100 mol of the U-12C-labeled
compound) by use of a matrix approach (17). The baseline spectrum was
that of analytes isolated from each pig before tracer infusion.
Portal balance.
|
(1)
|
in which concentration is expressed as micromoles per liter, and portal
blood flow is expressed as liter per kilogram per hour.
|
(2)
|
in
which t/T is the tracer-to-tracee ratio of the
U-13C-labeled substrate (glucose,
glutamate, or glutamine) or the product (alanine or lactate).
Throughout this article, a positive portal balance signifies the
addition of a compound to the portal blood, and a negative balance
signifies the removal of the compound from the arterial input.
|
(3)
|
|
(4)
|
For the enteral tracers, input in Eq. 3 is the rate of tracer infusion, and for the
intravenous tracers, input is the arterial tracer flux, i.e.,
(concnART × t/TART) × portal blood flow.
End product synthesis.
The contribution of each substrate to total lactate, alanine, or
CO2 production was calculated as
follows. The portal balance of the
13C3
isotopomers of lactate of alanine was calculated with
Eq. 2 and expressed as a fraction of
the input of the respective U-13C
tracers. For enteral glutamate and glucose, which we assume were
completely removed from the intestinal lumen, the denominator was the
rate of tracer infusion. For intravenous glucose and glutamine, the
input was the portal tracer balance. On the assumption that tracer and
tracee are metabolized identically, total alanine or lactate production
was calculated as
|
(5)
|
Once again, for enteral glutamate and glucose, the input was the
dietary intake, whereas for intravenous glucose or glutamine, the input
was the estimated uptake from the arterial supply. It is important to
note that in calculating the total production of lactate or alanine
from glucose, each molecule of glucose yields two molecules of lactate
or alanine.
The contribution of each substrate to
CO2 production was calculated in a
similar fashion, with the inputs (either tracer or tracee) expressed in
terms of carbon, i.e., glutamate or glutamine × 5 or glucose × 6.
First-pass metabolism of enteral glucose.
The calculation of the metabolism of enterally administered
[U-13C]glucose to
lactate, alanine, and CO2 is
complicated by the fact that the large majority of the tracer is
absorbed and thereby labels arterial glucose. A portion of this labeled
arterial glucose is extracted and metabolized by the PDV and
contributes to the production of glycolytic end products and
13CO2.
This consideration does not apply to glutamate and glutamine metabolism
because 1) very little of the
enteral glutamate tracer is absorbed (16) and
2) metabolism of glutamine occurs
essentially only from the arterial input. However, the fractional
extraction of the intravenously infused
[U-13C]glucose by the
PDV is known (from Eq. 3), so that
the first-pass metabolism of enteral
[U-13C]glucose can be
calculated from the product of the arterial flux of
[U-13C]glucose during
the enteral glucose infusion and the fractional extraction of arterial
[U-13C]glucose during
the intravenous infusion.
|
(6)
|
Thus
in which the concentration and tracer-to-tracee ratios are those
measured during the enteral
[U-13C]glucose
infusion. Similarly, with the proportion of the extraction of arterial
glucose that was metabolized to lactate, alanine, and
CO2 known, the contribution of the
metabolism of absorbed [U-13C]glucose to
CO2, alanine, and lactate
production during the enteral [U-13C]glucose
infusion can also be calculated as
|
(7)
|
Absorption of dietary glutamine.
Although in our past studies, and in this study, the portal mass
balance of glutamine is negative, (i.e., there is net removal of
arterial glutamine by the PDV), there is the possibility that there is
simultaneous absorption of dietary glutamine into the portal vein and
extraction of arterial glutamine. The absorption of dietary glutamine
can be estimated from the difference between the portal glutamine
fractional mass and fractional tracer balances. Thus
|
(8)
|
in which arterial glutamine flux is
concnART × portal blood flow.
All results are expressed as means ± SD. Where fractional extraction
or production is shown, the average has been derived from the mean of
individual values and not from the mean values of balance and input.
 |
RESULTS |
The body weights and rates of substrate intake, tracer infusion, portal
blood flow, and total CO2
production by the PDV are shown in Table 1.
There were no significant differences among the mean weights of the
animals subjected to different tracer infusions. Over the postoperative
period (6 ± 1 day), the daily weight gain (293 ± 49 g/day; 38 ± 6 g · kg
1 · day
1)
was also not different among the groups. The absolute values for the
portal blood flow (22.3 ± 1.4 l/h), total
CO2 production (38 ± 2 mmol/h),
and O2 consumption (28 ± 2 mmol/h) were also similar, and consequently there was a negative
relationship between body weight and weight-specific portal blood flow,
CO2 production, and
O2 consumption. In unpublished
studies (T. A. Davis, D. Wray-Cahen, P. R. Beckett, and P. J. Reeds)
with 4-wk-old piglets receiving nutrients at a rate similar to that in
the present study, we found that whole body
CO2 production is ~30 mmol
CO2 · kg
1 · h
1.
Thus the observed CO2 production
by the PDV (4.92 ± 0.97 mmol · kg
1 · h
1)
accounted for 16% of total body
CO2 production. The respiratory quotient across the PDV was 1.38 ± 0.11, a value that was
significantly (P < 0.001) higher
than unity.
Mass and Tracer Balance of Primary Substrates
The mass balances of enteral glutamate, enteral glucose, and
circulating glutamine are shown in Table 2.
A high proportion of the calculated intake of lactose-bound glucose (on
average, 85% of intake in groups 3 and 4) appeared in the portal blood. As found in previous studies (16, 17), the PDV were in glutamate equilibrium. Portal glutamine balance was negative and accounted for
12.7% of the arterial glutamine flux.
The portal balance of the U-13C
tracers is shown in Table 3. There was a
small but significant (3.7% of dose;
P < 0.05) net absorption of
[U-13C]glutamate;
93.6% of the enteral dose of
[U-13C]glucose
appeared in the portal vein. The PDV extracted 20.4% of the arterial
[U-13C]glutamine flux
and 6.5% of the arterial flux of intravenously infused
[U-13C]glucose. The
glutamine fractional tracer balance was significantly (P < 0.001) greater than its
fractional mass balance, and in molar terms, the difference was 72 ± 10 µmol · kg
1 · h
1.
This presumably represented absorption of dietary glutamine, so it
appeared that ~50% of the dietary glutamine was absorbed intact.
Lactate and Alanine Production
The portal mass and tracer (i.e.,
U-13C isotopomer) balances of
alanine and lactate are shown in Tables 4
and 5, respectively. The portal balance of
ammonia is also shown. The portal balance of alanine (623 ± 132 µmol · kg
1 · h
1)
was ~120% higher than the calculated intake of alanine (285 µmol · kg
1 · h
1).
There was also substantial lactate production (483 ± 52 µmol · kg
1 · h
1)
by the PDV and a positive portal balance of
[U-13C]alanine and
[U-13C]lactate in all
four groups. The production of ammonia by the PDV (968 µmol · kg
1 · h
1)
was the equivalent of 21% of the protein-nitrogen intake of the
animals. Given that the net synthesis of alanine requires a net input
of nitrogen, it would appear that
1,300
µmol · kg
1 · h
1
of amino acids were catabolized by the PDV. Approximately one-half of
this (680 µmol
N · kg
1 · h
1)
could be ascribed to catabolism of glutamate and glutamine.
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Table 5.
Portal balance of [U-13C]alanine and
[U-13C]lactate derived from metabolism of
[U-13C]glutamate, [U-13C]glutamine, or
[U-13C]glucose
|
|
The proportion of the input of any given substrate metabolized to
alanine and lactate was quite low (2.9-6.6% of dose). However, because the quantities of each substrate presented to the PDV were
markedly different (ranging from 191 µmol · kg
1 · h
1
for arterial glutamine to 3,406 µmol · kg
1 · h
1
for dietary glucose), the contribution of each substrate to total alanine and lactate production (Table 6)
varied widely. Even after adjustment for the secondary metabolism of
absorbed glucose, the first-pass metabolism of enteral glucose
contributed 18% of the portal alanine balance and 31% of the balance
of lactate. A further 22% of total alanine production and 14% of the
lactate production derived from arterial glucose metabolism. Thus
glucose metabolism accounted for 40% of the portal alanine carbon and 53% of the portal lactate carbon.
CO2 Production
The data on the portal balance of
13CO2
are summarized in Table 7. There was net
13CO2
production from all four 13C
substrates. A very low proportion (4%) of the enteral dose of [U-13C]glucose
appeared in portal
13CO2,
and when adjusted for the secondary metabolism of the recycled [U-13C]glucose, only
1.6% of the enteral glucose dose was metabolized to
CO2 in first pass. In contrast,
27% of the arterial glucose extracted by the PDV was metabolized to
13CO2.
Fifty-two percent of the enteral dose of
[U-13C]glutamate and
70% of the uptake of arterial
[U-13C]glutamine
appeared in portal
13CO2.
Estimated contributions of the substrates to total
CO2 production by the PDV are
shown in Table 8. The most important
sources were enteral glutamate (36% of total) and arterial glucose
(29% of total). Arterial glutamine (15%) made a smaller contribution, and the contribution of enteral glucose (6% of
CO2 production) was minimal. The
oxidation of the four substrates accounted for 76% of total
CO2 production by the PDV.
Other Nitrogenous Products
The data in Table 9 show that during the
enteral
[U-13C]glutamate
infusion, there was significant portal outflow of
[U-13C]arginine (3.9%
of dose) and proline (5.2% of dose). These observations confirm in
vitro studies with isolated porcine enterocytes (2, 32).
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Table 9.
Portal balance of amino acid products of mucosal metabolism of enteral
glutamate in pigs receiving an ig infusion of
[U-13C]glutamate
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|
 |
DISCUSSION |
Largely as a result of the pioneering work of Windmueller and Spaeth
(summarized in Ref. 26), considerable attention,
particularly in the clinical literature (3), has been paid to the role
of glutamine in the small intestinal mucosa, and it is often stated that glutamine is the "major" energy source for these cells. In vitro studies (24, 25) appear to confirm this comment, because isolated
enterocytes transport glutamine at a rate that is about fourfold higher
than that of glutamate. In vivo, however, the same considerations may
not apply. First, despite the fact that isolated enterocytes will
readily metabolize glucose (8), a number of studies in adult humans
(e.g., Ref. 13) have shown that the splanchnic extraction of oral or
intraduodenal glucose is minimal. Second, other studies in humans (1,
14) have shown that the splanchnic extraction of enteral glutamate
exceeds that of glutamine. Third, it has been repeatedly demonstrated (16-18) that in fed animals, even though the removal of lumenal glutamate (both free and protein-bound) is efficient, little dietary glutamate appears in the portal circulation. Finally, Windmueller and
Spaeth (28) explicitly commented on the fact that lumenal glutamate
appeared to be a more important oxidative substrate than glutamine. The
present data, obtained with conscious animals receiving a mixed
high-protein diet, confirm the large majority of Windmueller and
Spaeth's results and, in particular, their assertion about the
importance of glutamate to the energy economy of the small intestinal
mucosa. The fact that such similar results have been obtained in
different species, at different stages of development, and under quite
different experimental circumstances, strongly suggests that the
results reflect a specific feature of mucosal metabolism, i.e., a
critical role of amino acids as energy sources.
In drawing conclusions about the broader significance of the results,
there are two points that require discussion. The first concerns the
outflow of nitrogenous substances in the portal vein. Although we have
not directly quantified its metabolism, there was no release of
aspartate to the portal circulation, so that it is probable that
dietary aspartate is catabolized to the same extent as glutamate and
glutamine (see also Refs. 29, 31). About 10% of aspartate nitrogen
metabolism would have been directed to arginine biosynthesis, thereby
generating fumarate, but the majority of the aspartate would been
metabolized via transamination with
-ketoglutarate to generate
cytoplasmic oxaloacetate. That being so, we estimate that the exit of
nitrogen in newly synthesized alanine (338 µmol · kg
1 · h
1) accounted for
only 47% of the nitrogen generated from the estimated catabolism of
glutamate, aspartate, and glutamine (715 µmol · kg
1 · h
1).
The remainder apparently exited as ammonia, which in the present study
was produced by the PDV at a rate of 968 µmol · kg
1 · h
1.
Critically, the rate of ammonia production not only exceeded glutamine
deamidation (190 µmol · kg
1 · h
1)
by a factor of five, but the sum of alanine synthesis and ammonia production exceeded the quantity of nitrogen that would have been generated from aspartate, glutamate, and glutamine catabolism. These
observations are particularly important because they imply that, in
vivo and in the fed state, 1) there
is a metabolically significant glutamate deamination (i.e., an active
glutamate dehydrogenase) in the mucosa and
2) other dietary amino acids are
catabolized in first pass by the mucosa.
On the basis of the results of Windmueller and Spaeth (28, 31) and from
results with isolated enterocytes (25), it is generally assumed (see
Ref. 11) that glutamate dehydrogenase is either absent from the mucosa
or present in negligible quantities. However, it is possible to
question this assumption. First, in at least one detailed study of the
metabolism of isolated enterocytes (25), ~35% of cellular glutamate
uptake appeared as ammonia. Second, direct measurements in both porcine
(4) and rodent small intestinal mucosa (5, 9, 19) have demonstrated the presence of low, but significant, amounts of glutamic dehydrogenase. We
find it noteworthy that if the portal ammonia production that we
observed is expressed per unit mucosa, then the value [1.3 µmol · g (mucosal
weight)
1 · min
1]
is remarkably similar to that measured directly (1.5 µmol · g
1 · min
1)
in mucosa isolated from porcine jejunum (4).
The second observation that requires comment is the difference in the
distribution of the metabolism of glucose, glutamate, and glutamine in
the production of lactate and alanine ("incomplete" oxidation;
Ref. 24) compared with that of CO2
(complete oxidation). With glucose, the majority of the pyruvate was
converted to lactate and alanine and exported to the portal
circulation, whereas with glutamate or glutamine, only a minor portion
followed the same pathway. Windmueller and Spaeth (31) made similar observations.
In considering this observation, it is important to emphasize that, for
an amino acid that enters intermediary metabolism directly via the
tricarboxylic acid cycle to be completely oxidized (20), it must be
metabolized via acetyl-CoA and hence via pyruvate. Thus in this respect
pyruvate is a common product of glucose, glutamate, and glutamine
metabolism. However, because the pyruvate pool derived from glutamate
and glutamine metabolism appears to be channeled to oxidation, whereas
that derived from glucose is channeled to lactate and alanine
formation, it seems reasonable to argue that the pyruvate derived from
glutamate and glutamine metabolism is generated in the mitochondrion.
This proposition, however, begs the question of the enzymes
responsible. The first possibility is via
phosphoenolpyruvate carboxykinase.
This leads to the cytoplasmic generation of
phosphoenolpyruvate. However, although
phosphoenolpyruvate carboxykinase is
active in the enterocyte, alanine production from glutamine is not
blocked by inhibition of
phosphoenolpyruvate carboxykinase
(26). The second possibility is via the decarboxylation of malate by
NADP-linked malic enzyme, an enzyme also present in the cytoplasm of
enterocytes (25). This would also lead to the generation of cytoplasmic pyruvate, which, in the absence of further cytoplasmic
compartmentation, should follow the pathway of pyruvate generated from
glycolysis. The third, and intriguing, possibility is that the
synthesis of pyruvate from the tricarboxylic acid cycle intermediates
generated from glutamate and glutamine occurs within the mitochondrion
and that the reaction is catalyzed by a malic enzyme (20), termed NAD(P)+-malic enzyme by Sauer and
colleagues (15, 20). This enzyme differs from the cytoplasmic malic
enzyme by having a mixed specificity for NAD and NADP and, critically,
is expressed only in rapidly proliferating cells, including those of
the intestinal mucosa (15). Although speculative, this proposition is,
in our opinion, the most compatible with our data.
As we have pointed out elsewhere (21-23), our observations
regarding amino acid metabolism by the intestinal tissues have
substantial nutritional implications. First, if as we believe, dietary
amino acids are the major source of mucosal energy generation, then other factors that affect mucosal mass, such as parasitic infestation and dietary toxins, could have a substantial effect on the systemic availability of dietary amino acids. Second, the results imply that a
considerable portion of the splanchnic metabolism of dietary amino
acids occurs in the intestine rather than in the liver. Leucine
catabolism by the canine intestine has been observed (34). However,
whether other essential amino acids are catabolized in the mucosa is a
controversial point, for no other reason than the fact that there is
little direct evidence for the presence of the appropriate enzymes in
the intestinal mucosa. Finally, we should emphasize that the animals
that we studied had habitually received very high protein intakes, and
our observations could reflect metabolic adaptation to this dietary
regimen. In view of this, at least two further questions now need to be
investigated. First, do the mucosa catabolize and oxidize dietary
essential amino acids in vivo? Second, is mucosal amino acid metabolism sensitive to the level of protein in the diet?
 |
ACKNOWLEDGEMENTS |
We are particularly grateful to the Ajinomoto Co. for synthesizing
[U-13C]glutamate and
to Drs. T. Kimura and D. M. Bier for many helpful discussions. Finally,
we thank L. Loddeke for careful editing of this paper.
 |
FOOTNOTES |
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 USDA/ARS Cooperative Agreement no.
58-6258-6001, by the National Institute of Child Health and
Human Development (R01-HD-33920 and RO1-HD-35679) and by the
International Glutamate Technical Committee. The contents of this
publication do not necessarily reflect the views or policies of the US
Department of Agriculture, nor does mention of trade names, commercial
products, or organizations imply endorsement by the US Government.
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 correspondence and reprint requests: P. J. Reeds, USDA/ARS
Children's Nutrition Research Center, Dept. of Pediatrics, Baylor
College of Medicine, 1100 Bates, Houston, TX 77030 (E-mail:
preeds{at}bcm.tmc.edu).
Received 4 November 1998; accepted in final form 9 March 1999.
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