Increased Hepatic Fructose 2,6-Bisphosphate after an Oral Glucose Load Does Not Affect Gluconeogenesis*
Eunsook S. Jin
,
Kosaku Uyeda
¶,
Takumi Kawaguchi
,
Shawn C. Burgess
,
Craig R. Malloy
and
A. Dean Sherry
|| **
From the
The Mary Nell and Ralph B. Rogers
Magnetic Resonance Center, Department of Radiology and the
¶Department of Biochemistry, University of Texas
Southwestern Medical Center, Dallas, Texas 75235, the
Veterans Affairs North Texas Health Care System,
Dallas, Texas 75216, and the ||Department of
Chemistry, University of Texas at Dallas, Richardson, Texas 75083
Received for publication, February 28, 2003
, and in revised form, May 9, 2003.
 |
ABSTRACT
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The generally accepted metabolic concept that fructose 2,6-bisphosphate
(Fru-2,6-P2) inhibits gluconeogenesis by directly inhibiting
fructose 1,6-bisphosphatase is based entirely on in vitro
observations. To establish whether gluconeogenesis is indeed inhibited by
Fru-2,6-P2 in intact animals, a novel NMR method was developed
using [U-13C]glucose and 2H2O as tracers. The
method was used to estimate the sources of plasma glucose from gastric
absorption of oral [U-13C]glucose, from gluconeogenesis, and from
glycogen in 24-h fasted rats. Liver Fru-2,6-P2 increased
10-fold shortly after the glucose load, reached a maximum at 60 min, and
then dropped to base-line levels by 150 min. The gastric contribution to
plasma glucose reached
50% at 30 min after the glucose load and gradually
decreased thereafter. Although the contribution of glycogen to plasma glucose
was small, glucose formed from gluconeogenesis was substantial throughout the
study period even when liver Fru-2,6-P2 was high. Liver glycogen
repletion was also brisk throughout the study period, reaching
30
µmol/g at 3 h. These data demonstrate that Fru-2,6-P2 does not
inhibit gluconeogenesis significantly in vivo.
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INTRODUCTION
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Plasma glucose is preserved by gluconeogenesis after exhaustion of glycogen
stores during a moderate fast. Following an oral glucose load, gluconeogenesis
is thought to be modulated by allosteric regulation of
fructose-1,6-bisphosphatase
(1). This is an eminently
satisfying model because a key regulatory site in glycolysis and
gluconeogenesis occurs at level of fructose-6-P and fructose-1,6-P2
(Fig. 1).
Phosphofructokinase-1, the glycolytic enzyme, is potently activated by
fructose-2,6-P2, whereas fructose-1,6-bisphosphatase, the
gluconeogenic enzyme, is thought to be inhibited by this same effector
molecule (2,
3). Thus, by regulating the
activities of phosphofructo-1-kinase and fructose-1,6-bisphosphatase in a
reciprocal manner, Fru-2,6-P2 is thought to serve as an elegant
regulator of glucose usage/production by the liver after an oral glucose
load.

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FIG. 1. Metabolic pathways and sources of deuterium enrichment. In the
presence of 2H2O, glucose derived from glycogen will be
enriched at H2 (assumes rapid equilibration between Glc-6-P and Fru-6-P),
whereas glucose coming from gluconeogenesis will also be enriched at H5.
Enzymes: PF-1-k, phosphofructo-1-kinase; PF-2-k,
phosphofructo-2-kinase; F-2,6-P2ase,
fructose-2,6-bisphosphatase;
F-1,6-P2ase,
fructose-1,6-bisphosphatase. TCA, tricarboxylic acid; DHAP,
dihydroxy-acetone phosphate.
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This generally accepted model is based on kinetic analysis of
fructose-1,6-bisphosphatase in vitro, which shows that
Fru-2,6-P2 competes with Fru-1,6-P2 for the active site
of fructose-1,6-bisphosphatase and that both molecules have similar affinity
constants, 15 µM
(46).
However, the in vivo concentrations of Fru-1,6-P2 in
fasted and fed livers are 20 and 35 µM, respectively, whereas
those of Fru-2,6-P2 are 1 and 8 µM, respectively
(79).
This suggests that it would be difficult for Fru-2,6-P2 to have a
significant direct effect on fructose-1,6-bisphosphatase activity in
vivo based simply upon concentration differences. Some evidence has been
presented that suggests Fru-2,6-P2 is not a potent inhibitor of
gluconeogenesis in intact animals. For example, Kuwajima et al.
(10) reported continual
production of liver glycogen in sucrose-fed rats despite high levels of
Fru-2,6-P2, and Hue and Bartrons
(11) observed stimulated
glucose production by glucagon in isolated hepatocytes regardless of the
levels of Fru-2,6-P2. Levels of Fru-2,6-P2 have also
been manipulated by recombinant adenovirus overexpression of the bifunctional
enzyme phosphofructo-2-kinase:fructose-2,6-bisphosphatase (the enzyme that
catalyzes both synthesis and degradation of Fru-2,6-P2) in mice and
rats in vivo
(1214).
Here, increased hepatic Fru-2,6-P2 in vivo actually
resulted in increased glycogen synthesis from [1-13C]glucose via
the indirect pathway (14),
thereby suggesting that activation of glycolysis by Fru-2,6-P2 is
more important than inhibition of glyconeogenesis in vivo. The
relationship between glucose production by liver and hepatic
[Fru-2,6-P2] after an oral glucose load typical of that used in a
tolerance test (OGTT)1
is even less well defined. It has also been shown that hepatic glucose output
in 2430-h fasted rats is not suppressed after an oral glucose load
(15,
16), but
[Fru-2,6-P2] was not measured.
Continual production of glucose by the liver may play a role in diabetes,
and so a simple method to detect persistent gluconeogenesis after an oral
glucose load or after administration of a hypoglycemic agent may assist in
therapy of this epidemic disease
(17). Numerous methods to
monitor sources of plasma glucose have been described. Classical metabolite
balance studies across the liver or across the entire splanchnic circulation
are not optimal because measuring hepatic glucose production requires access
to the portal vein, and analysis of splanchnic glucose balance is limited by
uncertainties about glucose uptake in the gut. Detection of hepatic glycogen
by 13C NMR offers direct, noninvasive, and serial measurements of
hepatic glycogen mobilization, but the method is not widely available and is
difficult to apply to small animals without prelabeling of hepatic glycogen.
Sophisticated isotope tracer studies rely on incorporation of 13C
gluconeogenic precursors into plasma glucose, incorporation of deuterium or
tritium from body water into specific sites in plasma glucose, or the
redistribution of 13C label within plasma glucose molecules
(18). These methods assume
metabolic steady state and require metabolic models of variable
sophistication.
The present study had two purposes: 1) to distinguish the sources of plasma
glucose (gastric absorption, gluconeogenesis, glycogenolysis) in fasted rats
during an oral glucose load using a simple combination of 13C and
2H tracers; and 2) to determine whether hepatic
Fru-2,6-P2, elevated after an oral glucose load, alters the
contribution of gluconeogenesis to plasma glucose. Here, a combination of
1H and J-resolved heteronuclear single quantum coherence
(J-HSQC) spectroscopy was used to evaluate the contributions of hepatic
versus gastric glucose over time following an oral glucose load,
whereas 2H enrichments at the H5 versus H2 positions of
plasma glucose as determined by 2H NMR gave a direct measure of the
glycogenolysis versus gluconeogenesis contributions to plasma
glucose. The data show that gluconeogenesis is not inhibited significantly
in vivo following a glucose load in 24-h starved rats and that
gluconeogenesis remains active even when Fru-2,6-P2 is elevated
10-fold.
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MATERIALS AND METHODS
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ProtocolThe study was approved by the Institutional Animal
Care and Use Committee at the University of Texas Southwestern Medical Center.
Male Sprague-Dawley rats weighing 100140 g (120 ± 9 g) were
fasted for 24 h with free access to water. To initiate the study (t =
0), a bolus of glucose (2 g/kg; enriched with [U-13C]glucose to 5
or 10%) was administered by oral gavage, and 2 ml of
2H2O (99.9%; Cambridge Isotopes, Andover, MA) was
injected into the intraperitoneal cavity. This was done without anesthesia. At
30, 60, 90, 120, 150, or 180 min, the animals were anesthetized by ether
inhalation, a small portion of liver (0.20.3 g) was immediately
freeze-clamped, and as much blood as possible (
23 ml) was
collected from the descending aorta into a heparinized syringe. Following
exsanguination, the remaining liver tissue was also quickly freeze-clamped.
Assays for hepatic metabolites were performed on the first small section of
liver, whereas the larger freeze-clamped portion was used to isolate liver
glycogen. The blood was centrifuged and the plasma divided into 3 aliquots; 10
µl was used to assay for glucose, 10 µl for a 2H2O
enrichment measurement, and the remaining plasma for isolation of glucose for
NMR spectroscopy. A separate group of animals was studied to determine the
2H enrichment pattern in blood glucose in the absence of a glucose
load; these rats received only 2 ml of 2H2O and were
sacrificed at 50 min.
Analytical ProceduresWhole blood was immediately
centrifuged, and the plasma was separated and deproteinized with perchloric
acid (70%), neutralized with KOH solution, and lyophilized. Plasma glucose was
converted to monoacetone glucose (MAG) using the method of Landau et
al. (19). The lyophilized
extract was suspended in 1.0 ml of acetone containing 40 µl of concentrated
sulfuric acid and stirred at room temperature for 5 h. The pH of the mixture
was adjusted with 50% NaOH until it became mildly basic. The acetone
supernatant was transferred into another tube, and the remaining precipitate
was washed three times using 1-ml aliquots of acetone. The supernatant and
washings were combined and evaporated under a stream of dry nitrogen gas. The
resulting dried residue was suspended in 5 ml of water, adjusted to pH 2.0
with dilute HCl, and incubated at 40 °C for 5 h. Thereafter, the solution
was adjusted to pH 8.0 with NaOH and lyophilized.
Glycogen was extracted from the liver and purified as described previously
(20). Isolated glycogen was
dissolved in 5 ml of 10 mM sodium acetate solution (pH 5) and
hydrolyzed by incubating for 4 h at 50 °C with 20 units of
amyloglucosidase (Sigma). After freeze-drying, the glucose was converted to
MAG as described above.
NMR SpectroscopyMAG derived from plasma glucose was
dissolved into 180 µl of 90% acetonitrile, 10% deuterium-depleted water
(Cambridge Isotopes) along with a few grains of sodium bicarbonate
(NaHCO3). 2H NMR spectra were collected at 50 °C
using a Varian INOVA 14.1 T spectrometer (Varian Instruments, Palo Alto, CA)
equipped with 3-mm broadband probe tuned to 2H (92.1 MHz). Shimming
was performed by visual inspection of select 1H resonances of MAG
using the decoupler coil for detection. Proton-decoupled 2H NMR
spectra were acquired using a 90° pulse and a sweep width of 920 Hz and
1984 digitized points. Typical 2H spectra required the sum of
5,00020,000 scans. Proton decoupling was performed using a standard
WALTZ-16 pulse sequence.
1H NMR and J-HSQC spectra were obtained on MAG dissolved in 120
µl of acetonitrile (natural abundance), 20 µl of deionized water, and 40
µl of deuterated acetonitrile to provide a 2H lock. All
1H and J-HSQC spectra were collected using a 3-mm inverse probe
(Nalorac, Inc. Martinez, CA) on the same spectrometer. 1H NMR
spectra were acquired using a 90° pulse and a 1-s interpulse delay,
averaged over 64 scans (the acetonitrile signal was presaturated using a
frequency selective pulse). The J-HSQC sequence using REBURP-shaped refocusing
pulses was described previously
(21). The 180° null method
was used to determine the 1H90° pulse width, and the null of
the acetonitrile 13C satellites was used to determine the
13C 90° pulse width. A spectral width of 4000 Hz digitized into
2394 points was used for the 1H dimension (F2), whereas 64
increments covering a spectral width of 120 Hz were used for the
13C dimension (F1).
13C NMR spectra of hepatic glutamate (purified from liver
extracts) were obtained in 2H2O using a Varian INOVA
11.75 T spectrometer (Varian Instruments) equipped with a 5-mm broadband
probe. Proton-decoupled spectra were acquired over 28,000 Hz (sweep width)
digitized into 16,000 points using a 45° pulse and a 4-s interpulse delay.
Spectra were typically averaged over 10,000 to 40,000 scans.
2H Enrichment in Plasma WaterThe 2H
enrichment of plasma water was determined as described previously
(22). Proton-decoupled
2H NMR spectra (128 scans) were acquired using a 30° pulse and
a sweep width of 920 Hz digitized into 3776 points. An interpulse delay of 8 s
was used to avoid partial saturation effects. 2H spectra were
collected on acetone samples (990 µl) containing 10 µl of plasma. The
percent 2H in plasma water was determined by comparing the areas of
the H2O/acetone resonances with resonance areas measured in a
series of standards. Standards were prepared with 2H enrichments
ranging from 0 to 2.5% using 2H2O (99.9%) and natural
abundance water.
Analysis of SpectraThe 2H resonance intensities
in spectra of MAG were determined by Bayesian analysis (Varian Instruments) of
the raw time domain data. Peak areas in all other NMR spectra were measured
after Fourier transformation by using the line-fitting subroutine in the
PC-based NMR spectral analysis program, NUTSTM (Acorn NMR Inc., Freemont,
CA).
Plasma Glucose Originating from the Oral Load: 13C
AnalysisAt each time point after an oral glucose load, it was
assumed that plasma glucose resulted from a combination of gastric absorption,
glycogenolysis, and gluconeogenesis from trioses. Given that the oral glucose
contained 5% [U-13C]glucose, the probability that two
[U-13C3]triose units originally derived from oral
glucose recombined to form [U-13C]glucose is at most 0.25% (0.05
x 0.05 less the amount of label lost as a result of entry into the
tricarboxylic acid cycle). Thus, it was assumed that all
[U-13C]glucose present in plasma glucose originated from gastric
absorption of oral glucose and not from partial degradation and
recombination.
The fraction of plasma glucose with an enriched 13C at carbon 1
(g,as defined in Equation
1) was determined by 1H NMR as the fraction of the
doublet (because of JCH coupling) in the H1 resonance.
Algebraically, this is expressed as
 | (Eq. 1) |
The contribution of [U-13C]glucose to this fraction was determined
from multiplet areas in the J-HSQC spectrum where the coupling constant
J1,3 is small but both J1,2 and
J1,5 are easily detected. We assumed that any molecule
with enrichment in carbon 1, carbon 2, and carbon 5 must reflect
[U-13C]glucose. Therefore, the fraction of C-1-enriched glucose
contributed by [U-13C]glucose (h) is defined as
 | (Eq. 2) |
To illustrate one example of these calculations, the fraction of plasma
glucose enriched in carbon 1 (g) was 0.041 as determined by the
1H NMR spectrum of MAG (not shown). A J-HSQC spectrum of the same
sample (Fig. 2, spectrum at 30
min) indicates that the fraction of glucose labeled in C-1 contributed by
[U-13C]glucose (h) was 0.59, and the fraction of glucose
that was [1,2-13C2]- or
[1,2,3-13C3]glucose (i) was 0.12. From these
data, the fraction of plasma glucose that is [U-13C]glucose is
g x h = 0.041 x 0.59 = 0.02419. However, in this
sample the oral glucose contained only 5% [U-13C]glucose, and so
the fraction of plasma glucose originating from the stomach,
Fgastric, was (g x h)/0.05 or 48%.
Because total plasma glucose measured 15.8 mM, then 7.6
mM originated from oral glucose.

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FIG. 2. H1C1 projections of J-HSQC spectra of monoacetone glucose
converted from plasma glucose of rats sacrificed at 30-min interval after a 5%
[U-13C]glucose load and intraperitoneal injection of
2H2O. Q, quartet due to coupling between
C-1 and C-2 plus C-1 and C-5; D12, doublet, due to coupling between
C-1 and C-2; S, singlet, due to natural abundance of 13C
plus any singly labeled glucose generated in the tricarboxylic acid
(TCA) cycle. A shaded circle represents 13C, and
an open circle represents 12C.
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Plasma Glucose Originating from Glycogenolysis or Gluconeogenesis:
2H NMR AnalysisFrom the combined information in the
1H and J-HSQC spectra, one can conclude that about half of plasma
glucose arose from gastric absorption of oral glucose. The remaining glucose
resulted from either degradation of glycogen or gluconeogenesis, and this is
reported in the 2H NMR spectrum of MAG
(23) as
 | (Eq. 3) |
 | (Eq. 4) |
Integration of 2H and 13C Tracer
ObservationsTogether, these spectra allow a measure of the sources
of plasma glucose.
 | (Eq. 5) |
 | (Eq. 6) |
 | (Eq. 7) |
where Fgastric + Fglycogen +
Fgluconeogenesis = 1. For example, the 2H
spectrum in Fig. 3 indicates
that the contribution of glycogen to hepatic glucose production was
1(H5/H2) = 0.18. Thus, the fraction of plasma glucose that originated
from glycogen 30 min after the oral glucose load was (0.18)(10.48) or
9.4%. The assumptions that underlie this calculation are reviewed under
"Discussion."

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FIG. 3. 2H NMR spectrum of monoacetone glucose derived from plasma
glucose isolated from a rat sacrificed at 60 min after a 5%
[U-13C]glucose load and intraperitoneal injection of
2H2O. The inset shows H5/H2 ratio as
measured by 2H NMR as a function of time after the oral glucose
load. Each point represents means ± S.D. for 35
measurements.
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Metabolite AssaysFreeze-clamped liver was homogenized in
0.1 N NaOH solution, and the supernatant was incubated at 80 °C
for 5 min after centrifugation. Fru-2,6-P2 was assayed by taking
advantage of the sensitivity of pyrophosphate:fructose-6-phosphate
phosphotransferase to Fru-2,6-P2 as described by Van Schaftingen
et al. (24). Liver
glycogen was assayed enzymatically
(25), and plasma glucose was
measured using a HemoCue glucose analyzer (HemoCue AB, Angelholm, Sweden).
Statistical AnalysisThe data are expressed as means
± S.D. using Microsoft Excel. Linear regression analysis for standard
curves was also performed with the same program.
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RESULTS
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Influence of an Oral Glucose Load on Liver
Fru-2,6-P2 and Other MetabolitesThe
influence of the oral glucose load on hepatic glycogen, Fru-2,6-P2,
and plasma glucose is shown in Fig.
4. Plasma glucose peaked at 30 min after the glucose load and
decreased gradually thereafter. Liver glycogen was low in 24-h fasted animals
but was gradually replenished throughout the study period, reaching a maximum
of 31.5 µmol/g at 3 h. This was a significant increase compared with time
zero but considerably below the normal level in fed rats (
200 µmol/g)
(26). Fru-2,6-P2
also increased significantly after the glucose load
(Fig. 4). The initial
concentration prior to glucose administration was 0.4 ± 0.3 nmol/g of
liver, but this was followed by a
10-fold increase by 30 min after the
glucose load. The time courses of hepatic Fru-2,6-P2 and plasma
glucose were roughly parallel. Finally, the amount of
2H2O in plasma water (1.61.8%) did not change
significantly throughout the study period
(Fig. 4).

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FIG. 4. Liver glycogen (µmol/g wet weight), liver Fru-2,6-P2
(nmol/g wet weight), plasma glucose (mM), and 2H
enrichment in plasma water as a function of time after an oral glucose load in
24-h fasted rats. At t = 0, animals were given an oral glucose
load and injected intraperitoneally with a bolus of
2H2O. Animals were then sacrificed at each time point to
collect the data shown. Each point represents the mean ± S.D. for three
measurements.
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The plasma glucose concentration was somewhat higher than expected for a
typical OGTT. Earlier studies
(27) used indwelling venous
catheters for blood drawing in conscious animals, in contrast to our study, in
which the liver was removed from anesthetized animals. Therefore, in a smaller
group of 24-h fasted rats, glucose (2 g/kg body weight) containing natural
abundance levels of 13C was dissolved in H2O and
administered by oral gavage, 2 ml of 2H2O was injected
into the intraperitoneal cavity, and blood was withdrawn at various time
points via the tail vein in the complete absence of anesthesia. In this group,
plasma glucose peaked at 7.5 mM at 30 min and decreased to
6mM by 180 min (data not shown). This indicates that the high
plasma glucose levels reported in Fig.
4 may be attributed in part to the known effects of anesthesia
(28,
29).
Effects of an Oral Glucose Load on the 1H, J-HSQC, and
2H SpectraHigh resolution 1H NMR spectra of
monoacetone glucose showed a fully resolved H1 resonance with well resolved
13C satellite peaks (not shown). The total areas of the
13C satellite wings in these spectra were typically
34%
or
78% for rats given an oral glucose load containing either 5%
[U-13C]glucose or 10% enriched glucose, respectively. The H1
projection of typical J-HSQC spectrum is shown in
Fig. 2. Here, the peak labeled
quartet (Q) is the dominant multiplet. At a minimum, this glucose
isotopomer must be enriched in C-1, C-2, and C-5, and thus it represents
[U-13C]glucose from oral glucose that has not undergone metabolism.
The doublet (D12) is the signal from glucose isotopomers with
13C at C-1 and C-2 representing [1,2,3-13C3]-
or [1,2-13C2]glucose. D12 reflects oral glucose that had
been taken either to the level of a triose or the citric acid cycle before
being resynthesized to glucose. The singlet (S) represents glucose
isotopomers with 13C only at C-1. This peak has contributions from
naturally abundant glucose and from singly enriched isotopomers derived from
the citric acid cycle. As illustrated in
Fig. 2, the Q fraction in these
spectra was maximal at 30 min and decreased gradually thereafter, whereas the
D12 fraction was minimal at 30 min and increased thereafter.
Intraperitoneal injection of 2H2O resulted in rapid
equilibration (within 30 min) of 2H into plasma water to a level of
1.61.8% excess enrichment (Fig.
4). 2H incorporation into glucose has been reported to
occur at various steps along the gluconeogenic and glycogenolytic pathways
(Fig. 1), and the 2H
spectrum of MAG provides a convenient readout of those exchanges. In the 24-h
fasted animals prior to oral glucose, an H5/H2 ratio of 0.79 ± 0.17
indicated that 79% of all plasma glucose containing 2H was produced
via gluconeogenesis, whereas the remaining
21% came from glycogen.
Somewhat surprisingly, the deuterium spectrum did not change significantly
after the oral glucose load. The H5/H2 ratio remained relatively constant (0.8
± 0.2) throughout the study period
(Fig. 3, inset), again
suggesting that
80% of all plasma glucose containing 2H was
produced via gluconeogenesis. That portion of plasma glucose arising from the
oral load would not be detected by 2H NMR in this experiment unless
glucose cycling between plasma and liver was active. The 2H NMR
spectra of MAG derived from liver glycogen isolated at 120180 min also
confirmed that hepatic glycogen was derived via gluconeogenesis (H5/H2 = 0.90
± 0.05).
Sources of Plasma GlucoseThe contributions of oral glucose,
gluconeogenesis, and glycogenolysis to plasma glucose pools are summarized in
Fig. 5. At 30 min, about half
of the plasma glucose originated from oral glucose, and its contribution
decreased gradually thereafter. The gluconeogenic contribution to plasma
glucose was substantial throughout the study period (
5.56.7
mM), whereas the contribution of glycogenolysis to plasma glucose
was
2mM and constant throughout the study period.

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FIG. 5. Changes in contributions of oral glucose (open bar),
gluconeogenesis (cross-hatched), and glycogenolysis (shaded)
to plasma glucose based upon analysis of the C-1 projections of J-HSQC and
2H NMR spectra. The height of each bar represents the mean
± S.D. for 35 measurements.
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DISCUSSION
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The purpose of this study was to measure sources of plasma glucose after a
gastric load and to determine whether the contribution of gluconeogenesis to
hepatic glucose production is suppressed by increased hepatic
Fru-2,6-P2. Based on early in vitro reports that
Fru-2,6-P2 inhibits fructose-1,6-bisphosphatase, it is now standard
teaching (30) that this
inhibition holds true in vivo as well. The 2H NMR results
of Fig. 3 show that
gluconeogenesis continues even in the presence of substantially increased
levels of hepatic Fru-2,6-P2
(Fig. 4), and thus inhibition
of gluconeogenesis by Fru-2,6-P2 does not appear to be important
in vivo. This result is consistent with a recent report showing that
increasing hepatic levels of Fru-2,6-P2 by overexpression of the
kinase isoform of phosphofructo-2-kinase resulted in increased glycogen
synthesis from [1-13C]glucose via the indirect pathway
(14). This result indicates
that Fru-2,6-P2 stimulates glycolysis in vivo but does not
inhibit production of glycogen via the indirect pathway. The current
investigation also demonstrated that gluconeogenesis is not inhibited
substantially after an oral glucose load typical of an OGTT even though liver
Fru-2,6-P2 was increased
10-fold above fasting levels. This
indicates that any inhibition of fructose-1,6-bisphosphatase by
Fru-2,6-P2 in the in vivo rat liver is not enough to alter
gluconeogenesis.
Plasma glucose reached higher levels than expected at 30 min post-oral
glucose and did not return to pre-oral glucose levels even at 180 min. Thus,
the curve shown in Fig. 4 is
somewhat elevated compared with that observed for an OGTT in conscious animals
(27). Dohm et al.
(31) reported that both
methoxyflurane (inhalation) and Innovar (intramuscular injection) induce
glycogenolysis in rats, with the effect being higher in fed animals than in
fasted animals. Thus, the somewhat enhanced levels of plasma glucose found
here during the OGTT likely resulted from increased glycogenolysis during
exposure of the animals to ether prior to collection of plasma glucose.
Increased glycogenolysis, however, does not detract from the primary
conclusion of this study because any glycogen degraded in response to
anesthesia was synthesized de novo during the OGTT (see
Fig. 4, bottom panel).
The observation that the H5/H2 ratio in MAG derived from liver glycogen at 180
min was similar to that of plasma glucose demonstrates that gluconeogenesis
contributed equally to both. Thus, any glycogenolysis that may have occurred
during the short period of anesthesia would have reported the same H5/H2
ratio.
The NMR method reported here for detecting persistent gluconeogenesis
requires three reasonable assumptions. First, all glucose isotopomers enriched
in 13C at carbons 1, 2, and 5 reflect only
[U-13C]glucose from oral glucose. The chance that this group of
isotopomers could arise from [U-13C]glucose resynthesized after
metabolism to a triose is small. If the entire oral glucose load was
metabolized to a triose and resynthesized, the chances of
[U-13C]glucose reforming would be at best 0.25%, if oral glucose
consisted of 5% [U-13C]glucose, and at most 1%, if oral glucose
consisted of 10% [U-13C]glucose. This lower limit would be reduced
even further in vivo because of dilution of the triose pools by
endogenous gluconeogenic precursors. It was also assumed that plasma glucose
only arises from three possible sources: gastric absorption, liver
glycogenolysis, or liver gluconeogenesis. This assumption excludes other
organs as origins of endogenous glucose production. The kidney is also a
gluconeogenic organ, but its contribution to blood glucose is not considered
significant except during unusual circumstances such as prolonged fasting or
acidosis (32,
33). A third assumption was
that the 2H labeling in glucose reflected recent glucose synthesis.
Within 30 min after an oral glucose load, glucose turnover increases from
about 15 mg/kg/min at base line to more than 50 mg/kg/min
(15). Although glucose
turnover was not measured in this study, the gluconeogenic contribution to
plasma glucose at 30 min after the oral glucose and 2H2O
loads was already significant. Consequently, the gluconeogenic contribution at
later time points must also reflect the metabolic activity of each individual
time point rather than an accumulated result over the previous periods.
Further evidence for rapid glucose turnover is shown by the rapid decline in
[U-13C]glucose (originating in the oral load) observed in plasma
(Fig. 5), consistent with the
high turnover of glucose reported in earlier studies of oral glucose loading
in rats (15).
Perturbation of glucose metabolism with an oral load continues to attract
interest because it is thought that the post-prandial state accounts for much
of the duration of hyperglycemia in patients with diabetes, and because the
OGTT is a standard method for diagnosis of abnormalities in carbohydrate
metabolism. Despite intensive work, the fate of oral glucose remains
surprisingly controversial. Reports of the cumulative appearance of oral
glucose in plasma have varied from about 70%
(34,
35) to nearly 100%
(36,
37), and the maximal rates of
glucose appearance have varied about 2-fold. Livesey et al.
(36) reported a study of
glucose kinetics in 12-h fasted humans after an oral glucose load, using
stable isotopes and mass spectrometry to detect gastric absorption of
[13C6]glucose oral glucose. They report that the
contribution of hepatic glucose to plasma glucose began to decrease shortly
after the oral load from an initial value of near 5 mM to a nadir
of 1.1 mM (36),
suggesting that gluconeogenesis may be more highly regulated in humans after
an oral glucose load. However, that study differed from ours in two respects.
First, the contribution of hepatic glucose production to plasma glucose was
not measured directly but rather was obtained by difference between total
plasma glucose (measured analytically) and plasma glucose derived from the
oral load (measured as M + 6 by mass spectrometry). It is important to point
out that this method cannot distinguish hepatic glucose production from
glycogenolysis versus gluconeogenesis, and it is well known that
substantial liver glycogen remains after a short 12-h fast in humans. In our
experiments with rats, liver glycogen was low after a 24-h fast, and
gluconeogenesis was measured directly using the combined 13C and
2H tracers. This has allowed us to demonstrate that gluconeogenesis
is not altered after an oral glucose load in this animal model. These study
differences emphasize the need to apply simple tracer methods such as the
method demonstrated here to assess post-prandial glucose metabolism in humans.
The method reported here could easily be applied during an OGTT in humans. A
dual isotope technique commonly used for the measurement of oral or endogenous
glucose appearance
(3840)
requires intravenous infusion at a constant rate of glucose tracer
([3H]glucose) and a glucose load of the other tracer
([14C]glucose). In comparison, the approach presented here requires
only ingestion of [U-13C]glucose (and 2H2O),
yet provides detailed information about the sources of plasma glucose. Such a
study in humans could be especially timely because persistent hepatic glucose
production after a meal may be an attractive therapeutic target for diabetic
patients (17).
 |
FOOTNOTES
|
---|
* This study was supported by grants from the National Institutes of Health
(RR-02584, DK-16194, HL-34557) and by Merit Review support from the Department
of Veterans Affairs. The costs of publication of this article were defrayed in
part by the payment of page charges. This article must therefore be hereby
marked "advertisement" in accordance with 18 U.S.C.
Section 1734 solely to indicate this fact. 
**
To whom correspondence should be addressed: Mary Nell and Ralph B. Rogers
Magnetic Resonance Center, University of Texas Southwestern Medical Center,
5801 Forest Park Rd., Dallas, TX 75235-9085. Tel.: 214-648-5886; Fax:
214-648-5881; E-mail:
dean.sherry{at}utsouthwestern.edu.
1 The abbreviations used are: OGTT, oral glucose tolerance test; J-HSQC,
J-resolved heteronuclear single quantum coherence; MAG, monoacetone
glucose. 
 |
ACKNOWLEDGMENTS
|
---|
We appreciate the review of this manuscript by Dr. Brian Weis.
 |
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