Rate at which glutamine enters TCA cycle influences carbon
atom fate in intestinal epithelial cells
J.
Quan,
M. D.
Fitch, and
S. E.
Fleming
Department of Nutritional Sciences, University of California,
Berkeley, California 94720-1304
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ABSTRACT |
Glutamine carbon
entry into the tricarboxylic acid (TCA) cycle was assessed in small
intestinal epithelial cells by measuring CO2 production from
[1-14C]glutamine, and
these data together with
[U-14C]glutamine data
were used to calculate fractional oxidation rates for
glutamine. CO2
production from either
[1-14C]glutamine or
[U-14C]glutamine
showed saturation kinetics, and the concentration needed to achieve the
half-maximal rate of CO2
production was 0.7 and 0.4 mmol/l, respectively. Maximal rate for
[1-14C]glutamine was
twice that for
[U-14C]glutamine.
Increasing glutamine concentration did not cause proportional increases
in glutamine entry into the TCA cycle and glutamine oxidation.
Consequently, fractional oxidation of glutamine decreased with
increasing glutamine concentration. Fractional oxidation could be
predicted from the rate at which glutamine carbon entered the TCA
cycle. (Aminooxy)acetic acid, an aminotransferase inhibitor,
reduced entry of glutamine into the TCA cycle and increased fractional
oxidation of glutamine. Glutamate carbon entered the TCA
cycle at about one-half the rate of glutamine-derived glutamate carbon
and had a higher fractional oxidation rate when provided at equivalent
concentrations to glutamine. These differences in the rate of entry
predictably account for the differences in the metabolic fate of
glutamine vs. glutamate carbon.
jejunum; anaplerosis; energy; alanine; enterocyte
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INTRODUCTION |
THE CLINICAL IMPORTANCE OF glutamine to intestinal
health has been extensively studied over the last decade. This interest was stimulated by earlier observations that glutamine was an important energy-producing substrate for intestinal epithelial cells. Studies (28, 29, 30, 31) performed in vivo demonstrated that glutamine is an
essential respiratory substrate for cells in the small intestinal mucosa, accounting for over one-third of the total
CO2 produced in the small
intestine. In vitro experiments have shown that jejunal epithelial
cells produce more CO2 from
glutamine than from glucose (8, 9) and, when present together,
glutamine and glucose produce similar amounts of ATP (11).
Despite the established importance of glutamine as an energy-providing
substrate, reports of the effects of glutamine on intestinal structure
and function have been inconsistent. Intestinal structure and function
were found to be maintained or improved by glutamine supplementation in
some studies using laboratory animals (5, 22, 33), but not in other
studies (21, 32, 34). Also, glutamine supplementation has been
concluded to be beneficial to the intestinal health of humans in some
review articles (35) but not in others (4). Explanations for the
inconsistent results have not been readily available.
Although glutamine provides energy for the intestinal mucosa, previous
work (10, 15, 18) using the CO2
ratios technique has suggested that glutamine is not completely
oxidized to CO2. As a consequence,
glutamine carbon must efflux from the tricarboxylic acid (TCA) cycle
and be incorporated into synthetic products. In support of this,
glutamine carbon has been shown to be metabolized in vivo to
CO2, amino acids, and organic
acids, including citrate and lactate (28, 31). Through the use of
isolated cells, glutamine carbon has been found in metabolites,
including glutamate, CO2, lactate,
alanine, aspartate, citrulline, proline, succinate, and ornithine (11,
26). The most likely pathway by which glutamine carbon could be
incorporated into several of these metabolites would be via efflux of
intermediates from the TCA cycle. Other minor but physiologically
important synthetic products, such as lipids, would not have been
quantified with the methodologies employed in these studies, although
glutamine would be expected to provide precursors for the synthesis of
any compounds derived from TCA cycle intermediates.
The major objective of these studies was to further evaluate whether or
not the metabolic fate of glutamine carbon is influenced by changing
the rate at which glutamine carbon enters the TCA cycle, as previous
studies have suggested (15). These earlier studies (10, 15)
were based on values for "A + T" (where A is the probability that
carbon entering the TCA cycle will be incorporated into citrate via
acetyl-CoA and T is the probability that carbon will be incorporated
into citrate via oxaloacetate) derived using the
CO2 ratios approach.
In the current study, our first approach was to measure
CO2 production from
[1-14C]glutamine and
[U-14C]glutamine to
quantify glutamine carbon entry into the TCA cycle and glutamine
oxidation, respectively. These data were used to calculate the fraction
of carbon atoms entering the TCA cycle that are metabolized to
CO2, termed "fractional
oxidation" of glutamine. A second approach was to quantify
incorporation of [14C]glutamine into
compounds produced via the TCA cycle. The results suggest that the
metabolic fate of glutamine carbon is a function of the rate at which
glutamine carbon enters the TCA cycle in isolated intestinal epithelial cells.
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MATERIALS AND METHODS |
Animals.
Male Fischer 344 rats (Simonsen Laboratories, Gilroy, CA, or National
Institute on Aging breeding colony, Harlan Industries, Indianapolis,
IN) weighing 265-315 g were allowed access to commercial diets
(Rat Chow no. 5012, Ralston Purina, St. Louis, MO, or NIH 31 stock
diet, Western Research Products, Hayward, CA). All animals were allowed
free access to diet and water. Animal handling procedures were approved
by the Animal Care and Use Committee at the University of California
(Berkeley, CA).
Chemicals.
Radiochemicals, purchased from DuPont NEN (Boston, MA) included
L-[U-14C]glutamate,
L-[1-14C]glutamate,
L-[U-14C]glutamine,
[1,4-14C]succinate,
and
[2,3-14C]succinate.
Tracers were purified using conventional TLC (MN 300 Cellulose, Brinkman Instruments, Westbury, NY) and developed in a
mixture of butanol, acetic acid, and water (24:4:10). Labeled substances were detected using a radioactive plate scanner (Bioscan, Washington, DC). The appropriate region of the plate was scraped to
separate the tracer from the impurities. The cellulose containing the
purified tracer was stored at
20°C, then eluted with
Krebs-Henseleit (KH) buffer on the day of experimentation. The
Ca2+-free KH buffer contained the
following (in mmol/l): 121.3 NaCl, 4.85 KCl, 1.21 KH2PO4,
1.21 MgSO4, and
25.5 NaHCO3.
Reagent grade chemicals were obtained from Sigma Chemical (St. Louis,
MO). These included antibiotics, antimycotics (streptomycin sulfate,
penicillin G, kanamycin monosulfate, and amphotericin B), glutamine
synthetase, lactate dehydrogenase assay kits, (aminooxy)acetic acid
(AOA), and 2-amino-2-norbornane-carboxylic acid (BCH; also known as
2-aminobicyclo[2,2,1]heptane-2-carboxylic acid). BSA (fraction V, pH 7.0, lyophilized) was purchased from ICN Biochemicals, and dinitrophenol was purchased from Aldrich (Milwaukee, WI).
Synthesis of
L-[1-14C]glutamine.
L-[1-14C]glutamine
was synthesized according to the procedure of Brosnan and Hall (3).
L-[1-14C]glutamate
(45 µCi/µmol; 10 µCi) was added to 7 ml of an
incubation medium (pH 7.1) containing 50 mmol/l imidazole
hydrochloride, 20 mmol/l MgCl2,
100 mmol/l NH4Cl, 25 mmol/l
mercaptoethanol, 10 mmol/l ATP, and 50 U glutamine synthetase and
incubated at 37°C for 2 h. The entire incubation mixture was passed
through a Dowex 1 Formate column, and the glutamine was eluted with 3 ml of water and stored at
20°C until used. Purity of the
labeled glutamine was confirmed by TLC. Recovery of
L-[1-14C]glutamine
ranged from 92 to 96%.
Preparation of isolated cells.
On the day of experimentation, animals were anesthetized by
intraperitoneal injection of Nembutal (Abbott Laboratories, North Chicago, IL) at 5 mg/100 g rat, and the intestines were
accessed via a midline incision. Cells from the proximal small
intestine were prepared from a 30-cm segment beginning 10 cm distal to
the pylorus; cells from the distal small intestine were prepared from the 25-cm segment proximal to the ileocecal junction. These segments were removed, and the animals were killed by exsanguination and thoracotomy.
The lumen of the isolated segment was washed free of contents. Then the
cells were isolated from the mucosa, using a chemical and mechanical
technique (9, 23), and modified to include antibiotics and antimycotics
(11). The everted segments were filled with a
Ca2+-free KH buffer containing
0.025% BSA, tied off, and incubated in 5 mmol/l EDTA at 37°C for
10 min for the proximal segment and 20 min for the distal segment. The
cells were removed with a pressurized stream of ice-cold cell
suspension buffer (KH buffer supplemented with
CaCl2 to 3.55 mmol/l, 0.25% BSA,
5 mmol/l dithiothreitol, and antibiotics as described above). The cell
suspensions were centrifuged at low speed to sediment intact villi and
crypts and to remove free cells in the supernatant. The final cell
suspensions were kept on ice until the
CO2 assays began. Dry weights of
cells were determined by heating at 100°C for 2 h and were
calculated as the difference between the weight of 1 ml of cell
suspension and 1 ml of
Ca2+-containing KH buffer. Dry
weights of cells ranged from 0.3-4.0 mg/flask.
Lactate dehydrogenase release (Sigma Diagnostics kit DG 1340-UV) was
used to evaluate membrane integrity, as described in detail
previously (10). During the 25- to 30-min incubations, leakage of LDH
into the medium averaged 5-11% of the total cell content.
CO2 production.
CO2 production was measured as
described previously (8). Aliquots of cell suspension were added to
25-ml Erlenmeyer flasks containing substrate (unlabeled substrate and
tracer, in cell suspension buffer) and gassed with 95%
O2-5%
CO2. For most treatments, flasks
contained 2-4 mg dry wt cells in 2 ml substrate medium. When
treatments required using substrate at concentrations of 1 mmol/l or
less, changes in substrate concentration due to its metabolism could be
avoided by reducing cell weights to 0.3 mg/flask, increasing medium
volume to 4 ml, and restricting incubation time to 25 min. Flasks were
sealed with stoppers containing plastic center wells and incubated at
37°C for 25-30 min. After the incubation, the reactions were
stopped by injecting 0.8 ml methanol and l ml
NaH2PO4
(1 mol/l) into the flasks. CO2
evolved during the incubation was trapped by adding 0.45 ml 10 mol/l
NaOH to the center wells. After 2 h, the center wells were transferred
to plastic scintillation vials containing l ml water and 15 ml
scintillation fluid (Hionic Fluor, Packard Instruments, Downers Grove,
IL) and placed in a scintillation counter for radioactivity measurements.
Total CO2 production (in µmol
CO2
produced · g
1 · min
1)
from [U-14C]glutamine
was calculated as follows
where
t is the incubation time in minutes,
C is the dry weight of the cells (in g
dry wt/ml cell suspension), S is the
specific activity of the substrate (in dpm/mol substrate), and
n is the number of carbon atoms per
molecule of substrate.
A parallel equation was used to calculate the entry of glutamine carbon
atoms (given in µmol carbon
atoms · g
1 · min
1)
into the TCA cycle. In this case, however, glutamine was labeled only
in the carbon-1 position.
Values for fractional oxidation were calculated using the following
equation
where
the numerator represents the oxidation of glutamine (using
[U-14C]glutamine or
[U-14C]glutamate) and
the denominator equals entry of glutamine into the TCA cycle (using
[1-14C]glutamine or
[1-14C]glutamate).
With [U-14C]glutamine
as the tracer, specific activities ranged from 3 × 105 dpm/µmol for glutamine at 5 mmol/l to 3 × 106 dpm/µmol
for glutamine at 0.1 mmol/l. With
[1-14C]glutamine as
the tracer, specific activities ranged from 1 × 105 dpm/µmol for glutamine at 5 mmol/l to 5 × 104 dpm/µmol
for glutamine at 0.1 mmol/l.
Values for A + T were calculated using the formula derived by Kelleher
(13) and Mallet et al. (18) as
follows
Care
was taken to ensure that equivalent quantities of these two tracers
were added to flasks. Tracer was added at ~1 × 107 dpm/flask.
Incorporation of glutamine carbon into metabolites.
Epithelial cells were incubated with substrates and trace quantities of
[U-14C]glutamine for
30 min as described above, then metabolism was stopped with 3×
vol 100% methanol. Each incubation included 2-4 mg cells (dry
wt), tracer (6 × 106 dpm),
and 5 mmol/l glutamine or 5 mmol/l glutamine plus 2 mmol/l AOA. The
production of CO2 was determined
in one set of flasks using the procedures described above.
Incorporation of glutamine carbon into
non-CO2 metabolites was determined
in a second set of flasks.
To determine the incorporation of substrate into
non-CO2 metabolites, we
centrifuged the alcoholic incubates at 4,300 g for 2 min. Supernatants were
lyophilized and extracted three times with 93% ethanol. The ethanol
extracts were applied to 20 × 20 cm cellulose TLC plates and
chromatographed using two-dimensional chromatography
(solvent
1 composed of
n-butanol, acetic acid, and
H2O, 24:8:6, vol/vol/vol;
solvent
2 composed of phenol,
H2O, and NaCN, 3:1:0.003,
wt/vol/wt; Ref. 28). The Rf values for key amino and key
organic acids were previously determined. Samples were
spiked with a mixture of unlabeled amino acids to facilitate visualization by spraying the developed plates with 0.05%
fluorescamine (28). Radioactivity was visualized using a radioactive
plate scanner. Regions of the plate corresponding to known metabolites were scraped and placed in scintillation vials, and compounds were
eluted with 500 µl of 0.1 mmol/l NaOH, 1 ml
H2O, and aqueous compatible
scintillation cocktail (Hionic Fluor). The specific activity of
substrate (dpm/µmol carbon of substrate = dpm · µmol substrate
1 · no.
of carbons per molecule of
substrate
1) and the
amount of radioactivity in each metabolite were used to calculate the
incorporation of substrate carbon into metabolites.
Experimental design and statistical analysis.
Figure 1 depicts the pathways involved and
the metabolites examined in the eight experiments performed in this
study.

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Fig. 1.
Glutamine metabolic pathway. Intermediates and end products analyzed in
these experiments are shown in boldface type. ALAT, alanine
aminotransferase; ME, malic enzyme; PAG, phosphate-activated
glutaminase.
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The objective of experiment
1 was to determine whether fractional
oxidation of glutamine could be reliably measured using [U-14C]glutamine to
measure oxidation to CO2 and using
[1- 14C]glutamate to
measure glutamine carbon entry into the TCA cycle. In this same
experiment, cells were incubated with
[1,4-14C]succinate and
[2,3-14C]succinate to
simultaneously determine values for A + T, facilitating comparisons
between the two approaches. In this experiment, 14 rats were used and
observations per treatment ranged from three to eight.
The objective of experiments
2 and
3 was to evaluate the kinetics of
glutamine entry into the TCA cycle and its oxidation to
CO2. To do this, we incubated
cells with either
[1-14C]glutamine or
[U-14C]glutamine as
tracers. Glutamine concentrations ranged from 0.1 to 20 mmol/l in
experiment
2 and from 0.01 to 5 mmol/l in
experiment 3. In both experiments, cells from
three animals were exposed to all treatments. The Lineweaver-Burk
equation was used to calculate values for maximal rate
(Vmax) and the
glutamine concentration needed to achieve the half-maximal rate of
CO2 production from either
[U-14C]glutamine or
[1-14C]glutamine
(Kox).
The objective of experiment
4 was to determine whether reducing
entry of glutamine carbon into the TCA cycle with a transaminase inhibitor, AOA, would increase the fractional oxidation of glutamine as
predicted from the relationship between these variables that was
determined in the previous experiments. AOA is known to be a general
inhibitor of aminotransferases and other pyridoxal phosphate-dependent enzymes (12), but alanine aminotransferase and aspartate
aminotransferase are the only pyridoxal phosphate-dependent enzymes
known to be important for the metabolism of glutamine. AOA has been
used previously in studies of enterocyte metabolism (2, 16, 19) and has been shown to reduce the relative rate of glutamine carbon entry into
the TCA cycle using the CO2 ratios
approach (15). In this experiment, cells were incubated with either
[1-14C]glutamine or
[U-14C]glutamine as
tracers, with either 5 mmol/l glutamine or 5 mmol/l each of glutamine
and glucose, and in either the absence or presence of 2 mmol/l AOA.
Cell suspensions were prepared from four animals, and each cell
suspension was exposed to the eight treatments.
Experiment
5 was designed to expand on the
objective of experiment
4. In
experiment
5, however, incorporation of glutamine carbon into CO2 as well as into
non-CO2 metabolites was quantified since this provided an alternative approach to assessing changes in
fractional oxidation. To do this, we incubated cells, with [U-14C]glutamine and 5 mmol/l glutamine in either the absence or presence of 2 mmol/l AOA.
Cell suspensions were prepared from four animals, and each cell
suspension was exposed to the eight treatments.
The objective of experiment
6 was to determine whether stimulating
the entry of glutamate carbon into the TCA cycle would decrease
fractional oxidation of glutamine, as was predicted by the hypothesis
developed from the previous data. Because no stimulators of
aminotransferase activity were known, we approached this with the
glutamate dehydrogenase (GDH) stimulator BCH. Although
glutamine carbon has been thought to enter the TCA cycle primarily via
aminotransferase pathways, there is evidence of flux through GDH (2,
6). Previous observations from our laboratory and others (6, 15) have
shown also that aminotransferase inhibitors reduce but do not prevent
glutamine oxidation, further suggesting that some flux through GDH may
occur. In preliminary experiments (data not shown), the potential
toxicity of BCH to isolated intestinal cells was evaluated by
quantifying CO2 production from
glucose
([U-14C]glucose) and
glutamine
([U-14C]glutamine). In
experiment
6, cells were incubated with either [1-14C]glutamine or
[U-14C]glutamine as
tracers and 5 mmol/l glutamine, in either the absence or presence of 5 mmol/l BCH. Also, cells were incubated with
[U-14C]glutamate and 5 mmol/l glutamate, in either the absence or presence of 5 mmol/l BCH.
Cell suspensions were prepared from three animals, and each cell
suspension was exposed to the six treatments.
The objective of experiment
7 was to determine whether the
relationship between glutamine entry into the TCA cycle and its fractional oxidation was constant along the length of the small intestine. Therefore, we isolated cells from the proximal and distal
small intestine from four animals. Each cell preparation was then
incubated with either
[1-14C]glutamine or
[U-14C]glutamine as
tracers and with either 20 mmol/l glutamine or 20 mmol/l each of
glutamine and glucose. Each of the four cell suspensions was exposed to
all four treatments.
The objective of experiment
8 was to determine whether the low
rate of glutamate oxidation by intestinal epithelial cells is
associated with low rates of entry into the TCA cycle. Cells from the
proximal small intestine were incubated with 5 mmol/l glutamate, and
CO2 production from both
[U-14C]glutamate and
[1-14C]glutamate was
measured to estimate glutamate oxidation to
CO2 and glutamate carbon entry
into the TCA cycle, respectively. Cells were isolated from six animals,
and each cell suspension was exposed to the two treatments.
In all experiments, values for each treatment were calculated as the
mean of duplicate analyses for each cell preparation. Data are
presented as means ± SE. In most experiments, differences were
determined using ANOVA techniques (repeated measures, one-way, and
two-way ANOVA) and, when present, differences were identified using the
Tukey's Studentized range test. Before undertaking ANOVA, variables
were checked for equal variance and normal distribution of data to
ensure suitability for that analysis. When there were significant
interaction effects using ANOVA, follow-up tests were completed on the
appropriate cell means rather than on the marginal means. In other
experiments, pair-wise comparisons were made using the
t-test, or the relationship between
two variables was described using regression. Computer programs were
used to perform computations (25). Results were considered
statistically significant at P < 0.05.
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RESULTS |
Use of [14C]glutamate to
estimate entry into TCA cycle of glutamine-derived
glutamate.
When both glutamine and glucose were present at 5 mmol/l,
CO2 was produced from
[U-14C]glutamine at
~50% of the rate at which CO2
was produced from [1-14C]glutamate
(Table 1). Assuming that
[1-14C]glutamate is a
reliable predictor of the rate at which glutamine-derived glutamate
enters the TCA cycle, then it can be estimated that 50% of the
glutamine carbons entering the TCA cycle were oxidized to
CO2. This proportion is in
reasonable agreement with the A + T value of 0.46, obtained by using
the succinate ratio method.
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Table 1.
Influence of substrate concentration on fractional oxidation rates
estimated using [14C]glutamate and
[14C]glutamine vs. CO2 ratio technique
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When glutamine was present at 0.3 mmol/l and glucose was present at 0.1 mmol/l, CO2 production from
[U-14C]glutamine
exceeded by more than twofold the rate of glutamine carbon entry into
the TCA cycle when determined using
[1-14C]glutamate as
tracer (Table 1). This result was unreasonable and also was not in
agreement with the A + T value of 0.50 determined simultaneously. These
results ruled out the possibility of using [1-14C]glutamate to
measure TCA cycle entry of glutamine in concentrations <5 mmol/l and
thus also the
[1-14C]glutamate and
[U-14C]glutamate pair
as tracers to calculate fractional oxidation rates for
glutamine. In all subsequent experiments,
[U-14C]glutamine and
[1-14C]glutamine were
used to measure glutamine oxidation to
CO2 and glutamine carbon entry
into the TCA cycle, respectively.
Influence of glutamine concentration on glutamine oxidation and
entry into TCA cycle.
The production of
14CO2
from [U-14C]glutamine
increased with increasing glutamine concentration and reached a plateau
at ~5 mmol/l glutamine (Fig. 2),
suggesting saturation kinetics. A plot of 1/S vs. 1/v (where S refers
to the substrate concentration and v refers to the rate of
CO2 production) gave a line with a
regression coefficient >0.999.
Vmax was
calculated to be 15.0 µmol
CO2 · g
1 · min
1,
and Kox was calculated to be 0.4 mmol/l.

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Fig. 2.
Influence of glutamine concentration on glutamine oxidation
(14CO2
from [U-14C]glutamine)
and on the entry of glutamine carbon atoms into the tricarboxylic acid
(TCA) cycle
(14CO2
from
[1-14C]glutamine).
Cells, isolated from the jejunum of fed rats, were incubated with
glutamine at concentrations ranging from 0.1 to 20.0 mmol/l and trace
quantities of
[U-14C]glutamine or
[1-14C]glutamine.
Production of
14CO2
was used to calculate entry of glutamine carbon into the TCA cycle
using equations presented in MATERIALS AND
METHODS. Each data point represents mean ± SE;
n = 3. Data were taken from
experiment
2.
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The plot of
14CO2
production from
[1-14C]glutamine vs.
glutamine concentration also suggested saturation kinetics (Fig. 2) and the plot of 1/S vs. 1/v gave a line with a regression coefficient >0.999. Using the Lineweaver-Burk equation,
Vmax was
calculated to be 28.7 µmol carbon
atoms · g
1 · min
1
and Kox was calculated to be 0.7 mmol/l. In a separate experiment (data not shown),
CO2 production from
[1-14C]glutamine was
measured at low glutamine concentrations of 0.01 to 5 mmol/l. The
regression coefficient of the Lineweaver-Burk plot was >0.999.
Vmax was
calculated to be 30.2 µmol carbon
atoms · g
1 · min
,
and Kox was calculated to be 0.6 mmol/l.
Increasing the glutamine concentration of the incubation medium was
associated with decreasing fractional oxidation rates of glutamine
(Fig. 3). A linear and inverse relationship
was observed between the rate of glutamine entry into the TCA cycle and
the fractional oxidation rate of glutamine (Fig.
4). These two variables (glutamine entry
rate and fractional oxidation of glutamine) had a significant
correlation coefficient of
0.96. Using regression analysis, the
best-fit line had a slope of
0.012 and a
y-intercept of 0.85. These constants
were subsequently used to predict fractional oxidation rates using TCA
cycle entry data from other experiments to determine whether this
relationship could be more generally applied.

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Fig. 3.
Influence of glutamine concentration on fractional oxidation of
glutamine. Cells, isolated from the jejunum of fed rats, were incubated
with glutamine at concentrations of 0.1, 1.0, 5.0, 10.0, and 20.0 mmol/l and trace quantities of
[U-14C]glutamine or
[1-14C]glutamine.
Production of
14CO2
was measured, and fractional oxidation was calculated as
14CO2
from
[U-14C]glutamine/14CO2
from [1-14C]glutamine.
Each data point was calculated from the mean of duplicate analyses.
Data were taken from experiment
2.
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Fig. 4.
Influence of the rate at which glutamine carbon enters the TCA cycle
(14CO2
from [1-14C]glutamine)
on fractional oxidation of glutamine. Cells, isolated from the jejunum
of fed rats, were incubated with glutamine at concentrations of 0.1, 1.0, 5.0, 10.0, and 20.0 mmol/l and trace quantities of
[U-14C]glutamine or
[1-14C]glutamine.
Production of
14CO2
was measured, and fractional oxidation was calculated as
14CO2
from
[U-14C]glutamine/14CO2
from [1-14C]glutamine.
Each data point was calculated from the mean of duplicate analyses.
Data were taken from experiment
2. Linear regression was used to
characterize the relationship between these variables
(r, 0.96; slope, 0.012;
y-intercept, 0.85).
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In cells of the proximal small intestine, AOA reduced glutamine
oxidation to CO2 (using
[U-14C]glutamine as
tracer) by 51% when glutamine was the sole substrate, and reduced
glutamine carbon entry into the TCA cycle (using
[1-14C]glutamine as
tracer) by 64% (Table 2). Consequently,
the fractional oxidation of glutamine increased from 0.52 in the
absence of AOA to 0.71 in the presence of AOA. The regression equation
of glutamine entry into the TCA cycle vs. fractional oxidation rate,
calculated for the data from
experiment
2 (Fig. 4), was used to predict
fractional oxidation rates using the glutamine entry data for
experiment 4 (Table 2). The predicted fractional
oxidation rate for the glutamine-only treatment (no AOA) was calculated
to be 0.47 (vs. 0.52 as measured) and for the glutamine plus AOA
treatment was calculated to be 0.71 (vs. 0.71 as measured). In the
presence of glucose, AOA increased the fractional oxidation of
glutamine from 0.53 to 0.75 (Table 2). Corresponding fractional
oxidation rates, predicted from the data shown in Fig. 4, were 0.43 and 0.78, respectively.
The incorporation of 14C from
[U-14C]glutamine into
CO2 was compared with
14C incorporation into alanine,
aspartate, lactate, and succinate since the TCA cycle is needed to
convert glutamine into these metabolites. Including AOA in the medium
reduced incorporation of glutamine carbon into each of these compounds,
but the effects were most profound for alanine, succinate, and
CO2. AOA decreased glutamine
carbon incorporation into the
non-CO2 TCA cycle requiring metabolites by 66%, whereas the decrease in
CO2 production averaged 49%
(Table 3). Label incorporation into
glutamate was increased by 21%. Trace amounts of label were also found
in proline, ornithine, and citrulline (data not shown), and the values
obtained agree with those previously published (11).
BCH was not found (data not shown) to significantly influence glutamate
oxidation to CO2
([U-14C]glutamate, 5 mmol/l glutamate), glutamate entry into the TCA cycle ([1-
14C]glutamate), or
fractional oxidation of glutamate (glutamate oxidation/glutamate carbon
entry into the TCA cycle). Also, BCH was found to not significantly
influence glutamine oxidation to CO2, glutamine entry into the TCA
cycle, or fractional oxidation of glutamine. Consequently, we were
unable to use this approach to study further the relationship between
glutamate entry into the TCA cycle and fractional oxidation rates.
Relationship between glutamine entry into TCA cycle and fractional
oxidation: proximal vs. distal small intestine.
In the presence of either glutamine alone or glutamine plus glucose,
CO2 production from
[U-14C]glutamine was
significantly higher for cells from the proximal than distal small
intestine (Table 4). Similarly, glutamine carbon entry,
determined using
[1-14C]glutamine, was
significantly higher in the proximal than in the distal segment for
both substrate treatments. The measured fractional oxidation values
were not significantly different for the two segments.
The regression equation of glutamine entry into the TCA cycle vs.
fractional oxidation rate (Fig. 4) was used to predict fractional oxidation rates for the data from
experiment
7. The predicted fractional
oxidation rates for data taken from cells of the proximal segment
agreed within 15% with the measured fractional oxidation values (Table
4). Fractional oxidation rates measured for cells taken from the
distal small intestine were not accurately predicted by this equation,
however, as measured values differed from predicted values by >40%.
Fractional oxidation of glutamate determined using
[14C]glutamate tracers.
The fractional oxidation rate for glutamate averaged 0.77 in cells of
the proximal small intestine when glutamate was provided at 5 mmol/l
(Table 5). The regression equation of glutamine entry into the TCA cycle vs. fractional oxidation rate that was calculated for the data from experiment
2 (Fig. 4) was used to predict
fractional oxidation rates for these glutamate data (Table 5). The
predicted fractional oxidation rate (0.74) for glutamate differed by
<4% from the measured fractional oxidation rate (0.77).
View this table:
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|
Table 5.
Fractional oxidation of glutamate using [14C]glutamate
tracers, and comparison to fractional oxidation predicted from
regression analysis using glutamine substrate and tracers
|
|
 |
DISCUSSION |
Use of [U-14C]glutamine and
[1-14C]glutamine to measure
fractional oxidation of glutamine.
For several reasons, it appears that entry of glutamine-derived
glutamate into the TCA cycle and fractional oxidation of glutamine can
be accurately measured using
[U-14C]glutamine and
[1-14C]glutamine as
tracers. The observations using
[14C]glutamine tracers
provided data that were intuitively correct, as
14CO2
production from both
[1-14C]glutamine and
[U-14C]glutamine
showed saturation kinetics with increasing glutamine concentrations
(Fig. 2), and fractional oxidation never exceeded the theoretical
maximum of 100%. Furthermore, the value of 27.7 µmol C
atoms · g
1 · min
1,
calculated as the total of the five TCA cycle-requiring metabolites (alanine, aspartate, CO2, lactate,
and succinate; Table 3) approximates the value of 32.5 µmol C
atoms · g
1 · min
1,
calculated for glutamine carbon entry determined using
[1-14C]glutamine
(Table 2). Fractional oxidation values decreased with increasing
concentrations of glutamine (Fig. 3), supporting the trend observed
previously using the CO2 ratios
approach (10).
It was surprising to observe that fractional oxidation of glutamine
could not be accurately measured using
[1-14C]glutamate and
[U-14C]glutamate as
tracers. The CO2 ratios technique,
pioneered many years ago (27) and universally utilized, is based on the
principle that pathway intermediates can be used to quantify the extent to which a substrate is oxidized in the TCA cycle to
CO2. We reasoned that the same
approach could be used to assess the relative oxidation of glutamine
but with isotopes of glutamate as tracers. This proved not to be the
case, suggesting that there are multiple intracellular glutamate pools
and that glutamine concentration influences the relative size and
specific activity of these pools. Glutamine and glutamate
are known to be transported across enterocyte plasma and
subcellular membranes via different mechanisms and at different rates
(6, 20, 29). Although care was taken to ensure that the ratio of
exogenously provided glutamine to glutamate was constant for the
[U-14C]glutamate and
[1-14C]glutamate
treatments, it appears that transport or other factors caused
differences in the specific activities of intermediates in key
intracellular compartments or metabolic pools. Isotopes of glutamate
could not be used to measure the fractional oxidation of glutamine.
Fractional oxidation rates for glutamine, using
[U-14C]glutamine and
[1-14C]glutamine as
tracers, were in good agreement with measurements made using the
principles of the CO2 ratios
technique.1
When both glutamine and glucose were present at 5 mmol/l, the value for
A + T was 0.46 (Table 1) and, using
[U-14C]glutamine and
[1-14C]glutamine as
tracers, fractional oxidation of glutamine was found to be 0.53 (Table
2). These values are within 15% and indicate close agreement between
these two quite different approaches to determining the extent to which
the glutamine molecule is oxidized to
CO2 by jejunal epithelial cells.
The fractional oxidation technique that utilizes
[14C]glutamine tracers
is a more direct approach, however, and avoids having to make
assumptions regarding the homogeneity of the pools of intermediates
originating from different substrates.
As further evidence that
[U-14C]glutamine and
[1-14C]glutamine data
can be used to measure fractional oxidation of glutamine, an experiment
was conducted to directly measure the incorporation of glutamine carbon
into metabolites of the TCA cycle. This experiment was conducted in the
absence and presence of AOA, since AOA was found to reduce by ~50%
the entry of glutamine carbon into the TCA cycle and to increase
fractional oxidation of glutamine (Table 2). Overall, AOA reduced
CO2 production by 49% and reduced
non-CO2 metabolite production by
66% (Table 3). The greater influence of AOA on
non-CO2 metabolites vs.
CO2 would be expected, based on
the increased fractional oxidation of glutamine observed using the
[U-14C]glutamine/[1-14C]glutamine
approach (Table 2). This further supports the use of
[14C]glutamine tracers
for calculating fractional oxidation.
Factors influencing fractional oxidation rate of glutamine.
Fractional oxidation of glutamine was influenced by the rate at which
glutamine carbon entered the TCA cycle. At low rates of entry, the
glutamine molecule may be 80-90% oxidized to
CO2 (Fig. 4). At high rates of
entry there may be little, if any, reentry of carbon into the TCA cycle
as the molecule appears under these conditions to be only 40% oxidized.
Entry of glutamine carbon into the TCA cycle was markedly influenced by
changes in the glutamine concentration of the medium (Fig. 2). The
Kox for glutamine carbon entry
into the TCA cycle was calculated to be 0.6-0.7 mmol/l, and this
function was saturated at 5 mmol/l. Only at near saturating
concentrations of glutamine can glutamine carbon be substantially
incorporated into three-carbon intermediates, including lactate and
alanine (26). The Kox of 0.6-0.7 mmol/l for CO2
production from
[1-14C]glutamine was
somewhat higher than the Kox of
0.4 (see RESULTS) or 0.45 mmol/l
(14) for production of CO2 from
[U-14C]glutamine.
These values suggest that in fed rats, in which the normal arterial
glutamine concentration is ~0.50 mmol/l (31), glutamine will be
oxidized to CO2 at ~50% of the
maximal oxidation rate, whereas entry of glutamine carbon will be
<50% of maximal entry rates. Glutamine may enter the TCA cycle at
maximum rates and may be maximally oxidized to
CO2, however, if luminal glutamine concentrations increase to 5 mmol/l or more after meal feeding. The
glutamine plus asparagine concentration in jejunal contents has been
reported to be 1.2 mmol/l after consumption of a high protein meal (1),
but less destructive hydrolysis techniques would need to be employed to
determine with certainty luminal glutamine concentrations.
The maximum rate of entry of glutamine into the TCA cycle was twofold
higher than the rate at which CO2
was produced from the glutamine molecule (Fig. 2). This is
comparable to the ratio of 2.5:1 observed previously (26). These ratios
indicate that, under conditions when the substrate is present at
concentrations that saturate the pathways, each glutamine molecule that
enters the TCA cycle as
-ketoglutarate undergoes two
decarboxylations to yield CO2. The
remaining carbons were found previously in three carbon intermediates,
such as alanine and lactate (26). Our results (Table 3) support and
expand on this previous observation, because we show that glutamine
availability influences glutamine carbon entry into the TCA cycle,
which, in turn, influences the fractional oxidation of the glutamine
molecule. Thus glutamine availability influences the metabolic fate of
glutamine carbon entering the TCA cycle.
The presence of glucose stimulated glutamine entry into the TCA cycle
(Tables 2 and 4). Pyruvate-producing compounds other than
glucose could also be expected to increase glutamine entry into the TCA
cycle since it is thought that glucose stimulates TCA cycle entry by
providing a pyruvate source for transamination with glutamine-derived
glutamate (15). Accordingly, compounds that would inhibit transaminase
activity would be expected to reduce the rate of entry into the TCA
cycle and increase fractional oxidation. This is supported by our
observations that AOA decreased entry, increased fractional oxidation
(Table 2), and decreased incorporation of glutamine into
CO2 to a lesser extent than into non-CO2 metabolites of the TCA
cycle (Table 3). The degree of change agreed very well with the
predicted fractional oxidation rate, based on the relationship between
these two variables that was established in a previous experiment (Fig.
4). Our attempt to stimulate glutamine carbon entry into the TCA cycle
with the purported stimulator of GDH, BCH, was not successful, however (see RESULTS). Our results suggest
that BCH did not stimulate GDH activity in intestinal epithelial cells
as it has been shown to do in other systems (7, 17, 24).
Metabolism of glutamate.
The current studies expand our understanding of glutamate metabolism in
the following way. When glutamine or glutamate is present in cell
suspensions at equimolar concentrations (5 mmol/l), glutamate carbon
enters the TCA cycle at approximately one-third the rate of glutamine
carbon (Table 2 vs. Table 5). Because glutamate requires only to be
transported to the site of its conversion to
-ketoglutarate (Fig.
1), whereas glutamine must be both transported and deamidated, one
might expect that glutamate carbon would enter the TCA cycle at a
greater rate than glutamine carbon. This was not observed here (Tables
2 and 5) or elsewhere (15, 29). The lower rate of entry of glutamate
carbon into the TCA cycle may be due to the slower absorption rate into
intestinal cells or mitochondria of glutamate compared with glutamine
(6, 29). At 5 mmol/l, glutamate carbon also was more completely
oxidized than glutamine carbon, as indicated by a higher fractional
oxidation value of 0.77 for glutamate (Table 5) vs. 0.52 for glutamine (Table 2). When the regression equation of glutamine
carbon entry vs. fractional oxidation (Fig. 4) was used to predict the
fractional oxidation rate of glutamate from the glutamate carbon entry
data reported here (Table 5), the measured and predicted values for fractional oxidation were in good agreement. This suggests that the
high fractional oxidation rate for glutamate vs. glutamine is probably
a consequence of the lower rate at which glutamate carbon enters the
TCA cycle in intestinal cells. These observations with glutamate,
together with our observations with glutamine, support a general
conclusion that the fractional oxidation within the TCA cycle of these
molecules is determined by their rate of entry into the TCA cycle.
Entry rate, in turn, is influenced by their extracellular
concentration, their rate of transport into the cell, and the rates at
which they are metabolized to
-ketoglutarate.
When glutamine is the sole substrate, in addition to entry into the TCA
cycle, glutamine-derived glutamate can also accumulate in the cells or
be secreted back into the medium. Even though intracellular glutamate
cannot be distinguished from labeled glutamate released into the
medium, the total nonglutamine radioactivity can serve as a measure of
glutamine processed through glutaminase. In
experiment
5, labeled glutamate (see
RESULTS) amounted to an additional
41% above the value for glutamine entry into the TCA cycle (using
Table 2 data). At a substrate concentration of 5 mmol/l (saturating
conditions), the capacity of glutaminase to generate glutamate exceeded
the capacity of the TCA cycle to process it further. Inhibiting the
transaminase removal of glutamate by using AOA reduced the TCA cycle
entry to 36% of the uninhibited level but produced only a 21%
increase in
[14C]glutamate. Total
glutamine deamidation was reduced to 60% of the uninhibited level,
reflecting a possible inhibitory effect of intracellular glutamate
accumulation on the action of glutaminase.
Glutamine metabolism in proximal vs. distal small intestine.
Cells from the proximal small intestine showed a significantly greater
rate of CO2 production from
glutamine than did cells from the distal segment (Table 4). These
results are consistent with previous observations (15). Also, the rate
of glutamine carbon entry into the TCA cycle was approximately twofold
higher for cells of the proximal vs. distal small intestine, resulting in similar values for fractional oxidation (Table 4). The measured values for fractional oxidation were not in good agreement with the
values predicted using the regression equation of glutamine entry vs.
fractional oxidation (Fig. 4) generated using cells of the proximal
small intestine. These few results in the distal small intestine
suggest that the relationship between fractional oxidation and
glutamine entry rates may differ substantially along the length of the
small intestine.
In conclusion, our results suggest that the availability of glutamine
has a marked influence on the metabolic fate of its carbons that enter
the TCA cycle. Increasing entry of glutamine into the TCA cycle was
associated with decreased fractional oxidation rates of glutamine under
several conditions, resulting in greater incorporation into synthetic
products such as lactate, alanine, and succinate. It can be expected
that there would be increased incorporation also into minor, but
physiologically important, synthetic products, including lipids or
other compounds using TCA cycle-derived precursors. These results will
have clinical implications if such precursors are essential for repair
or replacement of diseased or damaged intestinal tissues.
 |
ACKNOWLEDGEMENTS |
We thank R. Gill, K. L. Zambell, and A. Lee for technical
assistance and M. Hudes for statistical consultation.
 |
FOOTNOTES |
This study was funded by National Institutes of Health Competitive
Grant R01-AG-10765 and the Agriculture Experiment Station.
1
The
CO2 ratio is defined as the
steady-state production of
14CO2
from [1-14C]acetate or
[1,4-14C]succinate
divided by that from
[2-14C]acetate or
[2,3-14C]succinate
(13, 27). While
[2-14C]acetate or
[2,3-14C]succinate
must remain in the cycle for a complete turn before any
14CO2
is released,
[1-14C]acetate or
[1,4-14C]succinate is
oxidized to
14CO2
during the first turn of the cycle. Thus more
14CO2
can be produced from
[1,4-14C]succinate
than from
[2,3-14C]succinate
when efflux of TCA cycle intermediates occurs. The probability that a
compound is oxidized rather than incorporated into a product after its
efflux from the TCA cycle can be predicted by calculating the variable
previously referred to as A + T (13), where A is the probability that
carbon entering the TCA cycle will be incorporated into citrate via
acetyl-CoA, and T is the probability that carbon will be incorporated
into citrate via oxaloacetate. The variable A + T thus equals the
probability that carbon will complete one turn of the TCA cycle by
either remaining in the cycle (path T) or leaving as a four-carbon
fragment and reentering via acetyl-CoA (path A). Thus a higher A + T
value represents a greater probability of complete oxidation and lower efflux of TCA cycle intermediates. A + T predicts the probability that
carbon atoms entering the TCA cycle will remain in the TCA cycle for
one complete turn and, as such, is an index of oxidation of the
substrate molecule. The theoretical basis for use of
14CO2
production ratios to estimate flux into the TCA cycle and net oxidation
of a substrate is derived from classic work on steady-state TCA cycle
labeling patterns (27). This approach requires making the following
assumptions (13): 1) the system is
in metabolic and isotopic steady state when data are collected so that
the specific activity ratios among the citrate carbons are constant, 2) sources of radioactivity may be
varied without altering the chemical composition of the medium,
3) the specific activity of any pool
of TCA cycle intermediates cannot be determined by direct experimental
measurement, 4) TCA cycle metabolism
is not compartmentalized in a manner that causes the fate of
intermediates to be dependent on the precursors from which they were
formed, and 5) the system is not
compartmentalized in such a way that the tracer (acetate or succinate)
and substrate (glutamine) are metabolized by separate types of cells or
mitochondria. A study by Mallet et al. (18) in rat enterocytes provided
evidence substantiating these assumptions.
Address for reprint requests: S. E. Fleming, Dept. of Nutritional
Sciences, Univ. of California, 119 Morgan Hall, Berkeley, CA
94702-1304.
Received 9 October 1997; accepted in final form 15 July 1998.
 |
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