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INTRODUCTION |
In liver, the tricarboxylic acid cycle is not only coupled to
mitochondrial energy production but also provides substrates for
gluconeogenesis and other metabolic pathways. Therefore, in order to
quantify the tricarboxylic acid cycle flux from the labeling kinetics
of glutamate and glutamine in vivo, the influx/efflux of
labeled substrates coupled to the tricarboxylic acid cycle must be
included in the mathematical analysis.
Tricarboxylic acid cycle flux measurements in hepatocyte preparations
have previously been made using a variety of substrates (1), and more
recently measurements of tricarboxylic acid cycle activity in perfused
liver have been made using an isotopomer steady state analysis (2, 3).
Magnusson et al. (4) have measured relative metabolic fluxes
including the tricarboxylic acid cycle flux in human liver via a steady
state isotopomer analysis of glutamine which conjugates to
phenylacetate in liver and is excreted in the urine. Although these
in vivo methods provide relative flux information on a
number of metabolic pathways in liver, they lack the ability to
calculate absolute fluxes unless the absolute flux of one pathway is
known. This is possible in perfused systems but may be difficult in the
whole body.
In vivo kinetic measurements of tricarboxylic acid cycle
activity have been made non-invasively in brain (5-7) and in perfused heart (8-11) and liver (12) by measuring turnover of 13C
label in glutamate by NMR, and absolute tricarboxylic acid cycle fluxes
have been calculated from these measurements in the brain (5, 13) and
heart (8-11) using a variety of mathematical models. In kinetic
experiments of glutamate isotope labeling in brain, label dilution of
the brain glutamate/glutamine pool via exchange with the blood
glutamate/glutamine pool is limited by the blood-brain barrier (14).
Therefore, the tricarboxylic acid cycle flux has been calculated from
the brain [4-13C]glutamate/glutamine turnover data using
a linear metabolic pathway analysis (5, 13) in which to a first
approximation the exchange with blood metabolite pools is neglected.
However in liver, glutamine is in active exchange with blood causing
label dilution of the liver glutamate and glutamine pool (15). Thus in
order to calculate the tricarboxylic acid cycle flux in the liver using
in vivo NMR measurements of 13C label turnover
in glutamate and glutamine, the mathematical analysis must incorporate
labeled liver glutamine exchange with blood.
[2-13C]Ethanol was used as the labeled precursor in this
study, because its metabolism is localized in the liver, and the
decreased redox state (NAD+/NADH) associated with ethanol
metabolism significantly reduces FFA1 oxidation in the liver
thereby limiting label dilution of the [2-13C]acetyl-CoA
pool (Fig. 1).
The present study demonstrates that in vivo 13C
NMR detection of glutamate and glutamine turnover in the liver can be
achieved non-invasively and that a quantitative metabolic steady state analysis used to calculate the tricarboxylic acid cycle flux in liver
must incorporate glutamine exchange with blood. Additional flux
measurements associated with liver such as citrate lyase,
-ketoglutarate
glutamate exchange, glutamine
synthetase/glutaminase, and glutamine influx/efflux may be calculated
as well. Applications of this method to study liver metabolism using
different labeled precursors may be possible.
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EXPERIMENTAL PROCEDURES |
Animal Preparation for in Vivo NMR Experiments--
Eight male
Sprague-Dawley rats (Charles River, Kingston) weighing 280-380 g were
fasted 16-24 h before the experiment. They were anesthetized
intraperitoneally with Inactin (Byk-Guiden Pharmazeutika, Germany) (80 mg/kg). Once anesthetized, an intramuscular injections of xylazine (15 mg/kg) and atropine sulfate (81 µg/kg) were given.
Tracheostomies were performed, and rats were ventilated with a mixture
of room air and oxygen. Both carotid artery and jugular vein were
catheterized. An arterial blood sample was initially taken to measure
blood gases and pH on an ABL-30 physiological monitor (Radiometer,
Copenhagen, Denmark). Oxygen flow and ventilator volume were adjusted
to provide a physiological pO2 (75-100 mm Hg),
pCO2 (35-45 mm Hg), and pH (7.1-7.4). A
non-magnetic blood pressure transducer (Viggo-Spectramed, Oxnard, CA)
was attached to the arterial catheter using a 3-way stopcock to monitor
blood pressure.
Rats were placed prone on a Lucite platform with the liver positioned
over the NMR surface RF coils. After the base-line spectrum, rats were
infused with [2-13C]ethanol (99% enriched, 1 g/kg)
(Cambridge Isotope Laboratories, Woburn, MA) diluted to a 33% v/v
solution in 0.9% NaCl over a 4-min period. A 4-min infusion period was
necessary to minimize blood pressure loss during this period. Arterial
blood samples were drawn at 0, 5, 10, 15 min, and every 15 min
thereafter for blood metabolite measurements. Blood samples were
immediately centrifuged for 5 min in heparinized tubes at 10,000 rpm.
Plasma was removed and refrigerated at
20 °C until further
use.
Livers were in situ freeze-clamped and removed at the end of
the experiment. Rats were euthanized by administering sodium pentobarbital intravenously.
NMR Spectroscopy--
In vivo NMR spectroscopy was
performed on a Bruker Biospec 7.0T system (horizontal/22 cm diameter
magnet). The magnet was equipped with actively shielded gradients
(Bruker, Oxford, UK). Dual concentric surface RF coils were used. The
outer 1H (30 mm) was tuned to 300.81 MHz, and the inner
13C coil (18 mm) was tuned to 75.65 MHz. The RF isolation
between the two coils was 43 dB.
Global 1H shimming of water was performed to obtain
reasonable line widths for imaging. Axial multislice spin-echo scout
images (8 interleaved slices, 2 mm thickness, repetition time = 1000 ms, echo time = 24 ms) were taken to ensure placement of the liver in
the magnet isocenter. Respiratory gating of the NMR pulse sequence was
triggered using a 2-way switch on the respirator responding to each
piston stroke. This reduced motion artifacts by limiting signal
acquisition to the same period in the respiratory cycle.
Additional localized shimming was performed with a respiratory gated
STEAM sequence (16) over a 2 × 2 × 1-cm volume of the liver.
Water line widths of 35-60 Hz were obtained.
1H-decoupled 13C NMR spectroscopy was used
during the experiment. Localized carbon spectroscopy was necessary to
eliminate strong superficial lipid and abdominal muscle signals.
One-dimensional localized spectroscopy was performed by using a
modified ISIS sequence (17). The ISIS excitation slice was generated
using a 10-ms hyperbolic secant pulse (18) during a
6 G/cm gradient pulse orthogonal to the coil plane. The slice placement, parallel to
the surface coil, was calculated for each rat from its scout image. Due
to the large chemical shift artifact associated with 13C
ISIS, the lipid suppression from ISIS was optimized to the aliphatic region only (23-34 ppm). Additional surface signal suppression was
provided by a magnetic field surface spoiler coil which was placed in
plane between the surface coils and the rat abdomen. The magnetic field
spoiler coil was driven by a DC voltage output (5 V, 200 mA) turned on
during the initial 2 ms of the signal acquisition period.
A 10-min base-line spectrum was acquired (repetition time = 0.5 s,
scan number = 1200, 4K data) with broadband WALTZ-16 decoupling turned
on during the acquisition. Subsequent spectra were accumulated in
10-min acquisition periods over 60-75 min. All spectra were Lorentzian-filtered, Fourier-transformed, and base-line subtracted to
eliminate residual lipid signals. Chemical shifts were referenced to
C-2 ethanol at 17.9 ppm. The average signal to noise (root mean square)
for the C-4 glutamate and C-4 glutamine peaks at the last time point
were 16.5 ± 1.3 and 15.0 ± 1.4, respectively.
Proton-observed carbon-enhanced 1H spectroscopy was
performed on liver and plasma extracts for fractional enrichment (FE)
as described previously (19). The fractional enrichments (given as atom
% excess) of [4-13C]glutamate (2.34 ppm),
[4-13C]glutamine (2.44 ppm), [2-13C]acetate
(1.93 ppm), [3-13C]lactate (1.32 ppm), and
[3-13C]alanine (1.48 ppm) were calculated.
13C NMR analysis of liver tissue extracts was performed on
a Bruker AM 500 spectrometer system to obtain C-2,3,4 glutamate
relative enrichments and isotopomer data from their respective carbon
multiplets. Spectra were acquired with repetition time = 0.5 s,
scan number = 10,000, 16K data, and Waltz-16 broadband proton
decoupling. Peak intensities were corrected for saturation and nuclear
Overhauser effect contributions (with respect to C-4 Glu, 1.24 × C-2 Glu and 0.98 × C-3 Glu; with respect to C-4 Gln, 1.08 × C-2 Gln and 0.81 × C-3 Gln).
In Vivo, Non-NMR Experiments--
The ethanol infusion protocol
described above for the in vivo NMR experiments was
performed on a group of rats to obtain intermediate time point data
with regard to liver glutamate and glutamine pool size as well as their
respective enrichments. Liver and gastrocnemius muscle were
freeze-clamped at 0 (n = 2), 10 (n = 2), 25 (n = 2), and 40 min (n = 2).
Sample Preparation--
Blood plasma extracts were prepared for
NMR analysis by adding 100 µl of plasma to 100 µl of 1.2 N perchloric acid in D2O. Additionally, 10 µl
of 1.6 M sodium formate in D2O was added as an
internal concentration standard. After mixing and centrifugation at
8800 × g for 10 min, 200 µl of the supernatant was
pipetted into a 5-mm micro-NMR tube.
Liver extracts were prepared by homogenizing a 0.50 g sample of
liver with a variable high speed electric homogenizer in a vortex tube
filled with 0.9% perchloric acid (3 v/w) and 100 µl of 1 N sodium formate. The homogenization was performed over ice to keep the sample cold. After homogenization, the sample was centrifuged at 4 °C for 15 min (3400 rpm). KOH (4 N,
0.675 v/w) was added to the supernatant to precipitate excess
perchlorate ions. The sample was centrifuged once more at 4 °C for 5 min (3400 rpm). The supernatant was neutralized and eluted over a
Chelex 100 column (100-200 mesh, Bio-Rad) with 2 ml of distilled
H2O to remove divalent ions. The sample was dried in a
Speed-Vac (Savant, Farmingdale, NY) overnight, and 1 ml of
D2O was added to the dried residue before placing in a 5-mm
NMR tube.
Analytical Analysis--
Glutamate and glutamine concentrations
in liver extracts were calculated using an NMR assay as follows: a
fully relaxed 1H NMR spectrum (repetition time = 30 s)
of the liver extract was acquired, and the C-4 Glu, C-4 Gln, and C-3
Lac peak areas were integrated. Lactate concentrations in the extracts
were measured using a 2300 STAT PLUS lactate analyzer (Yellow Springs
Instruments, Yellow Springs, OH), and the glutamate and glutamine
concentrations were calculated using the ratios of peak areas and the
known lactate concentration.
Data Analysis--
Steady state isotopomer analysis was
performed on the glutamate 13C multiplet data obtained from
liver extracts as described previously by Malloy et al.
(20). This analysis was used to indirectly obtain the
[2-13C]acetyl-CoA FE. The individual glutamate C-2, C-3,
and C-4 multiplet peak areas were integrated with respect to their
respective homonuclear coupling patterns (i.e. singlet,
doublet, triplet, and quartet). The isotopomer analysis allowed for the
steady state analysis of all 13C labeling permutation of
glutamate described by Malloy et al. (20). With
simplifications (using [2-13C]ethanol which provides only
[2-13C]acetyl-CoA, and using only C-2, C-3, C-4 Glu to
avoid T1 differences), the following Equations
1-5 were used to calculate the C-2 acetyl-CoA FE.
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(Eq. 1)
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(Eq. 2)
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(Eq. 3)
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(Eq. 4)
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(Eq. 5)
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where C2S indicates C-2 Glu singlet; C2D23 indicates C-2 Glu
doublet (C-2, C-3 coupled); C3S indicates C-3 Glu singlet; C3T indicates C-3 triplet (C-3, C-2, C-4 coupled); and C4D34 indicates C-4
Glu doublet (C-4, C-3 coupled). Fc0 indicates
fraction of acetyl-CoA that is unlabeled; Fc2
indicates fraction of acetyl-CoA that is C-2-labeled;
Fa1 indicates fraction of labeled anaplerotic substrate that will yield either C-2 or C-3 oxaloacetate in the first
span of the tricarboxylic acid cycle; and y indicates ratio of anaplerotic versus citrate synthase flux.
Tricarboxylic Acid Cycle Flux (Vtca)
Calculation--
Metabolic steady state equations may be derived for
mass flow into the tricarboxylic acid cycle as described previously for brain (13) and heart (10). From the labeled substrate flow schematic
shown in Fig. 1, we may obtain isotopic
mass balance Equations 6-15.
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(Eq. 6)
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(Eq. 7)
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(Eq. 8)
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(Eq. 9)
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(Eq. 10)
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(Eq. 11)
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(Eq. 12)
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(Eq. 13)
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(Eq. 14)
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(Eq. 15)
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where the C2, C3, or C4 prefix refers to 13C label
at the respective carbon isotope positions. Vpdh
indicates pyruvate dehydrogenase flux; Vacet
indicates acetate thiokinase flux; Vcs indicates
citrate synthase flux; Vffa indicates fatty acid
synthase flux; Vket indicates acetoacetate
synthesis flux; Vtca indicates tricarboxylic
acid cycle flux; Vlyase indicates citrate lyase
flux; Vglu and
Vglu
1 indicate aminotransferase
and glutamate dehydrogenase flux; Vgln and
Vgln
1 indicate glutamine
synthetase and glutaminase flux, respectively; Vout and Vin indicate
glutamine efflux and influx, respectively; Vasp
indicates aspartate aminotransferase flux;
Vpepck indicates phosphoenolpyruvate
carboxykinase flux; Vpc indicates pyruvate carboxylase flux. Both glutamate and glutamine pool concentrations were
measured, and acetyl-CoA, citrate,
-ketoglutarate, and oxaloacetate pool concentrations in liver during ethanol metabolism were obtained from literature and converted to µmol/g wet weight (21). The pyruvate
dehydrogenase flux (Vpdh) was set equal to 0 as
this flux is very low in liver (2, 3, 15) especially during ethanol
metabolism (12). The acetate thiokinase flux
(Vacet) as a minimum estimate was set equal to
citrate synthase flux (Vcs). Varying
Vffa and Vket
indiscriminately had no effect on C-2 AcCoA turnover, because of the
small AcCoA pool size. The
-KG
Glu exchange via glutamate
dehydrogenase and/or aminotransferase reaction (Vglu and
Vglu
1) is rapid with respect to
Vtca in the brain (13) but has been shown to be
significantly slower in heart (10, 11, 22). Therefore, it was necessary
to include both C-4 and C-2 Glu turnover data in the mathematical
analysis to discriminate between Vtca and
Vglu. C-2 Glu time points were extrapolated from
the in vivo turnover of the C-2 Glu/Gln peak (not resolved
at 55.5 ppm) and ratio of C-4 Glu to C-4 Gln at their respective time
points. C-2 GlnB was extrapolated from C-4 GlnB
and the ratio of C-2 Glu to C-4 Glu at their respective time points. No
labeling of aspartate was detected in vivo, and minimal C-3
Asp was detected in liver extracts, so the aspartate aminotransferase
flux (Vasp) was set to 0. Lactate and pyruvate
are believed to be in fast exchange in the liver (15), so the C-3 Pyr
turnover was set equal to the C-3 Lac turnover which was determined
from liver extracts at intermediate time points. Pyruvate carboxylase
flux (Vpc) was set equal to the measured
anaplerotic flux. Malic enzyme flux (Vmalic) was
set to 0, because it was shown to be low following a fast (23).

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Fig. 1.
13C label flow schematic in liver
originating from [2-13C]ethanol. This schematic
illustrates the flow of 13C label (in bold) from
[2-13C]ethanol to [4-13C]glutamate and
[4-13C]glutamine which are coupled to the tricarboxylic
acid cycle. Label from [4-13C] -ketoglutarate becomes
incorporated into [2-13C] or [3-13C]malate
and oxaloacetate, and in subsequent turns of the tricarboxylic acid
cycle, [2-13C] and [3-13C]citrate,
-ketoglutarate, glutamate, and glutamine become labeled (not all
shown). [2-13C]Alanine and aspartate are not shown, but
will appear when their respective aminotransferase reactions occur with
[2-13C]glutamate. The relevant fluxes to be used in the
mathematical analysis are shown next to arrows representing
label flow. Bi-directional arrows indicate flux in both
directions. AcAc, acetoacetate; Mal, malate;
PEP, phosphoenolpyruvate; and the 2-, 3-, or
4-13C refers to label at the respective carbon isotope
positions. Vacet, acetate thiokinase flux;
Vffa, fatty acid synthase flux;
Vket, acetoacetate synthesis flux;
Vcs, citrate synthase flux;
Vtca, tricarboxylic acid cycle flux;
Vlyase, citrate lyase flux;
Vgln and
Vgln 1, aminotransferase and
glutamate dehydrogenase flux; Vgln and
Vgln 1, glutamine synthetase and
glutaminase flux respectively; Vout and
Vin, glutamine efflux and influx respectively;
Vmalic, malic enzyme flux;
Vasp, aspartate aminotransferase flux;
Vpepck, phosphoenolpyruvate carboxykinase flux;
Vpc, pyruvate carboxylase flux;
Vpdh, pyruvate dehydrogenase flux.
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Mathcad (MathSoft Inc., Cambridge, MA) software was used to
simultaneously solve the differential equations described above. The
computer-generated Glu and Gln turnover curves determined from the
series of equations were fit to the NMR data via a non-linear least
squares best fit analysis (modified Levenberg-Marquardt process).
Random errors determined from S/N measurements were included with each
data point in the fit. All fits had a correlation value
R2
0.90. Intra-rat data were used to calculate
each Vtca. The error in the calculated fluxes
represent the mean ± S.D. reflecting the accuracy of the
measurement resulting from each curve fit.
Student's t test analysis was performed on data to
determine significance at a p
0.05 threshold.
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RESULTS |
A localized 13C NMR pulse sequence was used to
suppress the strong superficial lipid signals that overlapped the peaks
of interest. Fig. 2A
represents a non-localized carbon spectrum from the liver, abdominal
muscle, and superficial lipid (large lipid signals present at 23-34
ppm) in rat postmortem. These peaks obscure the C-4 resonances of
glutamate and glutamine at 34.4 and 31.3 ppm, respectively. The
modified one-dimensional ISIS localized pulse sequence was used to
produce the spectrum in Fig. 2B. Additional lipid
suppression was provided using the combined magnetic field spoiler
coil/one-dimensional ISIS localization sequence as shown in Fig.
2C. No decrease in the C-2 EtOH peak at 17.9 ppm in Fig. 2,
B and C, indicates that there was excellent
signal localization to the liver.

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Fig. 2.
In vivo 13C NMR superficial
lipid suppression techniques used in the label turnover experiments.
A illustrates a 10-min acquisition base-line spectrum of the
intact rat liver and superficial tissues after infusion with 1 g/kg
[2-13C]ethanol. The C-2 ethanol (EtOH) signal was
assigned to 17.9 ppm. The predominantly superficial aliphatic lipid
signals appeared at approximately 23-34 ppm. B illustrates
a 10-min acquisition spectrum using a modified one-dimensional ISIS
localized pulse sequence on the same liver. This was accomplished by
using a 10-ms hyperbolic secant inversion pulse during a 6 G/cm
magnetic field gradient orthogonal to the RF coil plane. This sequence
accounted for an approximately 50% (CH2)n lipid
signal reduction with no reduction in the C-2 ethanol signal. Spectrum
C was the result of the combined modified one-dimensional
ISIS and magnetic field spoiler coil localized pulse sequence. The
result was an approximately 68% reduction of the
(CH2)n resonance from that of spectrum A. All
measurements were made postmortem to ensure ethanol signal
stability.
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A series of base-line subtracted 10 min acquired spectra is shown in
Fig. 3. The times indicated are the
median time of acquisition. All experiments showed an immediate
increase in the 13C NMR signal at 17.9 ppm as a result of
the ethanol bolus. In subsequent spectra, the ethanol signal decreased
indicating ethanol clearance. C-4 Glu and C-4 Gln signals appeared at
34.4 and 31.3 ppm, respectively, in the first spectrum and increased
until steady state signal intensities were achieved. Labeling of C-4
Glu and C-4 Gln was the result of [2-13C]acetyl-CoA
condensing with oxaloacetate to produce [4-13C]citrate
which in turn labeled
-ketoglutarate, glutamate, and glutamine at
the C-4 position (Fig. 1). C-2 Glu and C-2 Gln at 55.5 and 55.0 ppm,
respectively (unresolved), began to appear at 5-15 min and increased
at a slower rate than C-4 Glu and C-4 Gln. These peaks appeared during
subsequent turns of the tricarboxylic acid cycle which results in
scrambling the C-4 label of Glu and Gln to C-2 and C-3 Glu and Gln. The
peaks of C-3 Glu and Gln at 27.9 and 27.0 ppm, respectively, and C-2
acetate at 24.2 ppm were difficult to observe, because these
frequencies are in the area where the ISIS suppression was optimized.
The ISIS suppression was optimized by placing the downfield edge of the
frequency selective inversion pulse at the C-2 fatty acyl chain peak
(~34.2 ppm) of the triglycerides. Therefore, using a magnetic field
gradient of
6 G/cm during the frequency selective pulse, signal
suppression at 27.9 and 27.0 ppm was 0.46 and 0.52 cm deep into the
liver, respectively, if the ISIS suppression was optimized at the
liver/abdominal muscle interface. The C-4 Glu, C-4 Gln, and C-2 Glu/Gln
peaks of all the base-line subtracted spectra were integrated, and the time point enrichments of C-4 Glu, C-4 Gln, and C-2 Glu extrapolated from tissue extract enrichment and in vivo label turnover
data are shown in Fig. 4A.

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Fig. 3.
A series of base-line subtracted
13C NMR spectra from a label turnover experiment. Each
spectrum represented a 10-min acquisition period after a bolus
[2-13C]ethanol infusion. The median acquisition times are
indicated to the left of the spectra. The C-2 ethanol (EtOH)
peak was chemical shift referenced to 17.9 ppm. C-4 glutamate (Glu) and
C-4 glutamine (Gln) peaks appeared at 34.4 and 31.3 ppm, respectively,
in the initial spectrum and increased until isotopic steady state was
achieved. The appearance of C-2 Glu and C-2 Gln at 55.5 and 55.0 ppm,
respectively (unresolved), in later spectra illustrates that label
scrambling through tricarboxylic acid cycle intermediates
occurred.
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Fig. 4.
Extrapolated substrate enrichment time
courses in liver and blood. A, the C-4 glutamate ( ) and
C-4 glutamine ( ) time points were extrapolated from their respective
integrated peak areas, and final substrate fractional enrichments
(APE) were measured in the liver extracts. The C-2 glutamate
( ) time points were extrapolated from the integrated C-2 Glu/Gln
peak (not resolved, Fig. 3) and ratio of C-4 Glu to C-4 Gln at their
respective time points. These data were subjected to a least squares
fit analysis of the mathematical model generated turnover curves. All
fits had a best fit correlation value R2 0.90.
The errors are given as S.E. for each data point. B, the C-2
acetate ( ) and C-4 glutamine ( ) enrichment time courses measured
in blood.
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The individual C-2, C-3, and C-4 Glu multiplet peak areas
(Fig. 5) represented as a fraction of
entire isotopomer peak area are given in Table
I. Steady state isotopomer analysis of
these glutamate peaks was used to calculate the C-2 acetyl-CoA
enrichment (Fc2), labeled anaplerotic substrate
enrichment (Fa1), and ratio of anaplerotic
versus citrate synthase flux (y) (Table I).

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Fig. 5.
13C NMR spectrum of a liver
extract. Steady state isotopomer analysis of glutamate was used to
calculate the C-2 acetyl-CoA enrichment (Fc2),
labeled anaplerotic substrate enrichment (Fa1),
and ratio of anaplerotic versus citrate synthase flux. The
individual C-2, C-3, and C-4 glutamate multiplet peak areas
(inset) were integrated, and the data were used in the
isotopomer analysis (results shown in Table I). Homonuclear
J coupling patterns are denoted above the individual
multiplets, and coupled nuclei are indicated.
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Table I
13C NMR steady state isotopomer analysis of glutamate in liver
extracts (% of multiplet peak areas)
C-2, C-3, and C-4 Glu refer to the entire multiplet peak resulting from
homonuclear scalar coupling of neighboring nuclei. Doublet 1-2 refers
to the doublet arising from the C-1 Glu to C-2 Glu scalar interaction,
and other doublets are similarly represented. The C-2 Glu doublet 1-2
and quartet were not detected in our samples. There was no C-4 doublet
4-5 as a consequence of essentially no C-1 acetyl CoA enrichment.
Fc2 refers to the C-2 acetyl-CoA FE arising from C-2
acetate. Anaplerosis (y) refers to the percent ratio of
total anaplerotic versus citrate synthase flux.
Fa1 refers to the fraction of labeled anaplerotic
substrate that will yield either C-2 or C-3 oxaloacetate in the first
span of the tricarboxylic acid cycle. Results are mean ± S.E.
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Fig. 6 illustrates the three spectra
obtained when performing Proton-observed carbon-enhanced spectroscopy.
Spectrum A represents the heteronuclear non
J-inverted spectrum, and subspectrum B represents the heteronuclear J-inverted spectrum. Spectrum C
represents the difference (A and B). These
spectra were used to calculate the fractional enrichments of C-4
Glu = 23.0 ± 1.1%, C-4 Gln = 17.2 ± 1.5%
(p < 0.005 versus C-4 Glu FE), C-3 Ala = 1.9 ± 0.4%, C-3 Lac = 2.0 ± 0.6%, and C-2
Acet = 49.9 ± 3.9% (Table
II). The C-2 Glu (7.7 ± 0.5%) FE
was calculated from the known C-4 Glu FE and relative C-2 to C-4 Glu
was measured in tissue extracts. The Glu and Gln concentrations were
4.02 ± 0.35 and 4.64 ± 0.35 µmol/g wet weight,
respectively.

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Fig. 6.
1H NMR spectra of a liver
extract. Proton-observed carbon-enhanced NMR analysis was
performed on liver extracts to determine the C-4 glutamate, C-4
glutamine, C-2 acetate, C-3 alanine, and C-3 lactate fractional
enrichments. Spectrum A represents the heteronuclear non
J-inverted spectrum, and subspectrum B represents
the heteronuclear J-inverted spectrum. The broadband
13C inversion pulse frequency was placed between C-4
glutamate and C-3 alanine (~26.2 ppm). Spectrum C
represents the difference (A and B). These
spectra were used to calculate the fractional enrichment of the
relevant enriched metabolites.
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Table II
Liver extract metabolite fractional enrichments and calculated
metabolic fluxes
Glu, glutamate; Gln, glutamine; Acet, acetate; AcCoA, acetyl-CoA; Lac,
lactate; Ala, alanine; FE, fractional enrichment as % APE (atom % excess). Data are given as mean ± S.E.
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The best fit C-4 Glu, C-4 Gln, and C-2 Glu turnover curves generated by
the computer analysis of the data above are shown in Fig.
4A. Table II summarizes the flux results from the
mathematical fitting procedure. Vtca = 0.33 ± 0.09 µmol/g wet weight/min, and the
-KG
Glu exchange rate
was an order of magnitude greater (Vglu,
Vglu
1 =3.80 ± 1.2 µmol/g
wet weight/min). These fluxes were relatively insensitive to changes in
the Cit,
-KG, and Oaa pools as a doubling of these concentrations
had no effect on Vtca.
A separate group of rats were sacrificed for base-line and intermediate
time point (10, 25, and 40 min) measurements after [2-13C]ethanol infusion to obtain liver glutamate and
glutamine enrichments and concentrations at various times. The C-4 Glu
and C-4 Gln enrichments were 14, 20.3, 24.1, and 23% and 7.2, 14.4, 16.9, and 17.2% at 10, 25, 40, and 55 min, respectively. These
enrichments correspond to C-4 Glu and C-4 Gln label turnover rates that
are not significantly different than those obtained by in
vivo NMR. The glutamate and glutamine concentrations were
2.49 ± 0.10 and 5.02 ± 3.10 µmol/g, respectively, at base
line and 3.63 ± 0.20 and 3.53 ± 0.78 µmol/g, respectively, at 10 min and did not vary throughout the remaining time
points. Therefore, the maximum overestimation in
Vtca assuming that the Glu pool increased from
2.49 ± 10 µmol/g at base line to 4.02 ± 0.35 µmol/g at
the end of the study would be 27% (0.24 versus 0.33 µmol/g wet weight/min).
 |
DISCUSSION |
In this study, the non-invasive 13C NMR detection of
glutamate and glutamine labeling in liver was achieved using
physiologically allowable concentrations of
[2-13C]ethanol (~0.1% in blood). A metabolic steady
state mathematical analysis similar to that previously used by Mason
et al. (13) and Yu et al. (10) was applied to the
kinetic and steady state labeling data to calculate a number of
metabolic fluxes coupled to the tricarboxylic acid cycle in liver. We
have combined isotopomer steady state (1H and
13C NMR of liver extracts) and non-steady state (in
vivo 13C NMR) measurements to calculate the absolute
substrate flux through the tricarboxylic acid cycle directly in liver
for the first time.
Oxygen consumption rates have previously been measured in perfused
liver in situ following ethanol addition to the perfusate (24-26). Thurman and Scholz (24) measured O2 consumption
rates of 1.83 µmol/g/min during a 1 mM EtOH perfusion.
Taylor et al. (25) measured O2 consumption rates
of approximately 1.6 µmol/g/min during a 2.5 mM EtOH
perfusion, and Eriksson et al. (26) measured O2
consumption rates of approximately 1.5 µmol/g/min after 20 min during
a 4 mM EtOH perfusion. Typically, it is difficult to extrapolate tricarboxylic acid cycle flux rates from tissue oxygen consumption rates unless the exact profile of substrates oxidized by
the tricarboxylic acid cycle is known due to differences in stoichiometric coupling. Assuming O2 consumption was
aerobically coupled to acetate oxidation in the above-mentioned ethanol
perfusion studies, then the oxygen consumption rate
(VO2) is stoichiometrically coupled to
Vtca by VO2 = 2 Vtca. Therefore, from these measurements we
estimate Vtca equal to 0.92, 0.80, and 0.75 µmol/g/min, respectively. These rates are more than double our
measured value of 0.33 ± 0.09 µmol/g/min. Unfortunately,
VO2 extrapolated measurements are not accurate
during conditions where there is elevated production of reducing
equivalents, and the respiration quotient is very low as is the case
during ethanol oxidation. It has been shown that although oxygen
consumption rates are maintained, CO2 production is
significantly inhibited by ethanol metabolism (27). We estimate that
greater than 90% of the acetate formed in our study was not oxidized
in the liver, yet 2 NADH reducing equivalents were produced for every
acetate formed via ethanol oxidation in the liver. Therefore, the vast
majority of reducing potential necessary for ATP synthesis was provided
by ethanol oxidation and not tricarboxylic acid cycle turnover as
reflected by our low tricarboxylic acid cycle flux measurements.
In perfused liver, the tricarboxylic acid cycle flux has previously
been quantified using a mass spectrometric assay (2, 3) of
13C labeling of glutamate during conditions of steady state
labeling and using equations developed by Magnusson et al.
(4). Beylot et al. (2) reported a tricarboxylic acid cycle
flux of 0.34 µmol/g/min following [3-13C]lactate
administration, and DiDonato et al. (3) reported a value of
0.25 µmol/g/min following combined [3-13C]lactate and
[3-13C]pyruvate administration. A mathematical model by
Vogt et al. (28) was also established to calculate relative
tricarboxylic acid cycle flux using data obtained from 13C
NMR isotopomer analysis of glutamine in liver extracts.
The heterogeneous liver acinus is spatially comprised of periportal and
perivenous zones, and it has been proposed that the periportal zone is
primarily gluconeogenic and the perivenous zone is glycolytic (32). In
our experiment, periportal and perivenous metabolism cannot be
distinguished, but Cline and Shulman (33) have shown in vivo
that there is no difference in tricarboxylic acid cycle substrate
labeling patterns in periportal versus perivenous zones
following a [U-13C]glucose administration. This would
imply that our tricarboxylic acid cycle flux measurement was
representative of the global hepatocyte tricarboxylic acid cycle
flux.
Glutamine is transported across the cell membrane by the following two
glutamine transport systems: the Na+-dependent
N transport system, and the Na+-independent system which
involves facilitated diffusion of glutamine in hepatocytes (34-36).
There was a difference in C-4 Glu and C-4 Gln enrichment (23.0 versus 17.2%, respectively) in our experiment which is
evidence that liver glutamine was in exchange with lower 13C-enriched glutamine in blood. Previous in
vivo measurements of net glutamine flux across the liver in fasted
rats resulted in a small net uptake of 1.7 (37) to 2.4 (38) nmol/g/min.
By using the glutamine extraction data established by Pardridge (39) and a plasma glutamine concentration of 0.4-0.6 mM
(37,38), the unidirectional glutamine influx would be 260-390
nmol/g/min, and efflux would be ~258-388 nmol/g/min by using the
above net uptake data. These values are similar to our calculated
values for Vin and Vout
(250 ± 50 nmol/g/min) assuming 0 net glutamine flux across the
membrane. If ethanol metabolism did change the net flux of glutamine, a
large increase in net glutamine efflux of, for example, 100 nmol/g/min
would have only a small effect on the calculated Vtca (
9%, 0.30 versus 0.33 nmol/g/min) assuming that endogenous glutamate or glutamine provided
the necessary substrate for increased net glutamine efflux.
Häussinger and Gerok (40) measured a glutamate uptake and net
glutamate release of approximately 0.03 µmol/g/min for each at
physiological glutamate concentrations in perfused livers, and Low
et al. (41) measured glutamate influx kinetics
(Km = 0.25 mM,
Vmax = 0.046 µmol/g protein/min) suggesting an
even lower uptake at physiological concentrations in sinusoidal
membrane vesicles. Since these transport rates are 1 to 2 orders of
magnitude slower than the calculated Vtca and
glutamine transport rates, this transport mechanism was omitted from
the model (Fig. 1).
[2-13C]Ethanol was used as the labeled precursor in this
experiment to minimize [2-13C]acetyl-CoA label dilution
resulting from fat oxidation due to the decreased redox potential
[NAD+/NADH] (29-31) resulting from ethanol metabolism.
Although ethanol is not a preferred substrate used to study normal
liver physiology, this choice nevertheless allowed adequate in
vivo 13C NMR detection of liver glutamate and
glutamine labeling. With the current sensitivity of NMR detection of
13C-labeled glutamate and glutamine in liver, it may be
possible to make these 13C NMR measurements using
[2-13C]acetate precursor infusion. It may also be
possible to observe [2-13C]glutamate turnover if
[3-13C]lactate precursor is used. Schumann et
al. (42) suggested that the distribution of 13C label
in liver glutamate derived from [2-13C]acetate and
[2-13C]ethanol precursors was incompatible with
[2-13C]acetate metabolism being primarily in the human
liver. They suggested that acetate was also metabolized to glutamate by
peripheral skeletal muscle and cycled back to the liver via glutamine.
Numerous studies have observed acetate metabolism as occurring in
peripheral tissues (43-45). At 40 min, we measured a
[4-13C]glutamine FE of 11.8% in blood (Fig.
4B) but a [4-13C]glutamine FE of only 3.7% in
skeletal muscle. This suggests that the glutamine label in blood was
predominantly a result of label contribution from liver rather than
from peripheral muscle. Therefore, when comparing the relatively high
enrichment of C-4 Glu in liver (23.0%) versus C-4 Gln in
muscle (3.7%), we can be reasonably confident that liver and not
muscle tricarboxylic acid cycle activity was detected in
vivo.
The liver [2-13C]acetate represented in Fig. 3 as a small
peak at 24.2 ppm remained stable throughout the experiment, and
therefore the [2-13C]acetyl-CoA FE most likely remained
stable. Additionally, the [2-13C]acetate FE (49.9 ± 3.9%) in liver extracts equalled the [2-13C]acetyl-CoA
FE (51.4 ± 3.4%). It is not known whether the label dilution of
these pools was due to incompletely suppressed FFA oxidation and
acetyl-CoA hydrolase activity in liver or from unlabeled acetate
originating in peripheral tissues. Label incorporation into
[2-13C]acetyl-CoA via pyruvate recycling (pyruvate
oxaloacetate
phosphoenolpyruvate
pyruvate) or via malic enzyme
which are both active in liver was most likely negligible as pyruvate
dehydrogenase (2, 3, 15) and malic (23) enzyme activity in fasted liver
is low. Additionally, [3-13C]lactate and
[3-13C]alanine FE in the liver extracts were low
(2.0 ± 0.6 and 1.9 ± 0.4%, respectively) and therefore
could have had only a minimal effect on varying the
[2-13C]acetyl-CoA enrichment throughout the experiment.
Although we could not explicitly calculate Vpdh
from the model, independently varying this flux while maintaining the
[2-13C]acetyl-CoA FE did not alter
Vtca because of the rapid turnover of this small
pool. The dilution of [2-13C]acetyl-CoA (51.4%) to
[4-13C]glutamate (23.0%) could not be entirely accounted
for by liver glutamine exchange with blood. It is possible that
endogenous proteolysis in liver may have occurred. This unexplained
dilution was similarly observed by Des Rosiers et al. (46)
in liver. The anaplerotic flux (0.9 × Vcs)
determined from the isotopic steady state analysis of glutamate (20)
was set equal to the pyruvate carboxylase (Vpc)
flux. This may be an overestimation of Vpc as
malic enzyme (Vmalic) flux may also contribute
to anaplerosis, but the distribution of anaplerotic flux between
Vpc and Vmalic has no
effect on Vtca. Citrate lyase flux
(Vlyase) provides acetyl-CoA substrate for FFA
synthesis, and although this flux contributed negligibly to the rate of
C-4 and C-2 Glu turnover in liver, varying this flux affected C-2 Glu
FE considerably as pyruvate carboxylase (Vpc) is
coupled to Vlyase. The
-KG
Glu exchange
flux (Vglu, Vglu
1) which comprises both
aminotransferase and glutamate dehydrogenase fluxes and
mitochondrial/cytosolic transfer of
-KG and Glu was the source for
the greatest error in Vtca. It was necessary to model both C-2 and C-4 Glu turnover to calculate this flux. This exchange rate was determined to be very rapid from in vitro
kinetic measurements (47) (~640 µmol/g/min) and determined to be
much slower in vivo, but an order of magnitude greater than
Vtca in our experiments (3.8 ± 1.2 µmol/g/min).
In summary, It is possible to detect 13C labeling of
glutamate and glutamine in liver via non-invasive 13C NMR.
The in vivo 13C labeling kinetics of glutamate
and glutamine in liver and glutamine in blood combined with a metabolic
steady state mathematical analysis allows for rapid, direct calculation
of the liver tricarboxylic acid cycle flux. This study illustrates how
non-invasive metabolic flux measurements in tissues other than brain
can be made by including label exchange with blood in the mathematical
analysis.
We are grateful to electrical engineers Terry
Nixon and Peter Brown for NMR technical improvements and maintenance of
the high field NMR systems.