1 Department of Radiology, Anaplerotic,
pyruvate recycling, and gluconeogenic fluxes were measured by
13C isotopomer analysis of plasma
glucose, urinary phenylacetylglutamine, and urinary glucuronide in
normal, 24-h-fasted individuals after ingestion of
[U-13C]propionate,
phenylacetate, and acetaminophen. Plasma glucose isotopomer analysis
reported a total anaplerotic flux of 5.92 ± 1.03 (SD) relative to
citrate synthase. This was not significantly different from glucuronide
and phenylacetylglutamine analyses (6.08 ± 1.16 and 7.14 ± 1.94, respectively). Estimates of pyruvate recycling from glucose and
glucuronide isotopomer distributions were almost identical (3.55 ± 0.99 and 3.66 ± 1.11, respectively), whereas phenylacetylglutamine
reported a significantly higher estimate (5.74 ± 2.13). As a
consequence, net gluconeogenic flux reported by phenylacetylglutamine
(1.41 ± 0.28) was significantly less than that reported by glucose
(2.37 ± 0.64) and glucuronide (2.42 ± 0.76). This difference in
fluxes detected by analysis of phenylacetylglutamine vs. hexose is
likely due to compartmentation of hepatic metabolism of propionate. Net
gluconeogenic flux estimates made by use of this stable isotope method
are in good agreement with recent measurements in humans with
[14C]propionate.
glucose; glucuronide; glutamine; isotopomers; nuclear magnetic
resonance
TRACERS OF HEPATIC METABOLISM offer promise for aiding
diagnosis of abnormal glucose metabolism in the clinical setting.
Although radioactive precursors can be used to trace human hepatic
metabolism (28, 30), they are unlikely to be widely adopted in the
clinical environment because of possible risks to the patient and the
impracticality of systematic radiation containment. More recently,
similar metabolic models have been successfully adapted for use with
stable isotope tracers (3, 15), and well-established chemical biopsy
methods have been used to target carbon skeletons of hepatic
UDP-glucose and glutamine (12, 15, 27-30, 40).
Most common tracers of hepatic metabolism, such as the gluconeogenic
substrates lactate and alanine, may be metabolized to some extent by
peripheral tissues and thus possibly confound analysis of liver
metabolism. Ideally, any tracer should be infused directly into the
hepatic portal vein and extracted quantitatively by the liver. Portal
vein infusions are too invasive for human studies, however, and most
hepatic tracers are not quantitatively extracted from portal
circulation by the mammalian liver (34). Propionate may be an important
exception in that it is quantitatively extracted into liver from portal
circulation in a wide range of mammalian species, likely including
humans (2, 8). It is generated in significant quantities by colonic
fermentation of Although deployment of more hepatophilic tracers has minimized labeling
contributions from peripheral metabolism (24, 30, 41), values of
gluconeogenic flux reported by these newer methods remain
controversial. Katz et al. (23) suggested that net gluconeogenic flux
values of about three times the tricarboxylic acid cycle rate are
unsustainable, because the ATP demand for gluconeogenesis exceeds the
total ATP output of the citric acid cycle via substrate level and
oxidative phosphorylation. Others have proposed that sufficient ATP is
available if a significant portion of hepatic acetyl-CoA originates
from Experimental protocol.
Six healthy subjects [5 male and 1 female, 23-39 yr old, and
weighing 54-88 kg (76 ± 12 kg)] were studied under a
protocol approved by the institutional human studies committee.
Subjects were fasted for 24 h before the start of the experiment and
allowed access to water. Figure 1 shows a
flowchart of drug ingestion and blood and urine sampling. From
7:00-8:00 AM on the day of the study, each subject ingested 66 mg/kg phenylacetic acid distributed in 12 gelatin capsules (Gallipot,
St. Paul, MN). At 8:00 AM, each subject ingested 2 tablets (500 mg
each) of acetaminophen, and the first blood sample was also drawn into
heparinized tubes (25 or 30 ml). Over the next hour, five of the
subjects ingested a total of 20 or 25 mg/kg sodium
[U-13C]propionate
distributed in 6 gelatin capsules, and the sixth subject received 17 mg/kg of the sodium
[U-13C]propionate.
After the 8:00 AM sample, blood was drawn at 20-min intervals over the
next 2 h, with additional draws at 2.5 and 3 h (total blood drawn was
225 or 270 ml). Urine was also collected every hour over a 6-h period
starting at 9:00 AM, with the exception of one individual whose urine
was collected every 2 h.
ABSTRACT
Top
Abstract
Introduction
Methods
Results
Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Methods
Results
Discussion
References
-amylase-resistant starch and nonstarch
polysaccharides (2, 8). Propionate has been administered by gastric
infusion to human subjects at rates of 4 mmol/h over a 3-h period with
no ill effects and no alteration of hepatic glucose production (26).
Being a weak acid, propionate is likely to pass rapidly and efficiently
into both splanchnic and portal vein circulation from the stomach after ingestion, in a manner similar to aspirin. These features suggest that
oral administration of propionate may be an effective method for
noninvasive analysis of hepatic gluconeogenesis in humans.
-oxidation (24). Because measurements of gluconeogenesis in
humans to date have been derived by using tracers that enter the citric
acid cycle via pyruvate or acetyl-CoA, it is important to verify these
observations by use of a tracer that enters the citric acid cycle via a
different metabolic pathway. Herein, we demonstrate that such
measurements are easily derived from a single
13C NMR spectrum of human plasma
glucose, urinary glucuronide, and urinary phenylacetylglutamine after
ingestion of
[U-13C]propionate,
phenylacetate, and acetaminophen.
METHODS
Top
Abstract
Introduction
Methods
Results
Discussion
References
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Fig. 1.
Time course for administration of
[U-13C]propionate,
phenylacetate, and acetaminophen and sampling of blood and urine.
Analytic procedures. Blood samples were chilled and centrifuged at 4°C in heparinized tubes immediately after being drawn. The plasma was deproteinized by addition of 0.5 ml of ice-cold 70% perchloric acid, and the precipitated protein was removed by centrifugation. The supernatant was adjusted to pH 5.0 with potassium hydroxide, and the insoluble potassium perchlorate was removed by a second centrifugation. The glucose content of each sample was assayed enzymatically before oxidation to gluconate with glucose oxidase (18). The samples were deproteinized a second time, and the pH was adjusted to 10.0 with KOH before lyophilization. Samples were resuspended in 600 µl 2H2O containing a potassium glycolate standard in which the glycolate carbons had natural 13C abundance (assumed to be 1.11%). For each NMR sample, the amount of glycolate added was equal to the amount of glucose assayed in the parent plasma extract (typically 20-40 µmol).
Our initial studies revealed that lyophilized urine samples were unsuitable for high-resolution 13C NMR analysis of the phenylacetylglutamine and glucuronide moieties, because 1) they were highly viscous, 2) they had a high salt content, and 3) the glucuronide was partially converted into a mixture ofNMR spectroscopy. Proton-decoupled 13C NMR spectra of blood extracts were obtained using a 5-mm probe on a 9.4 T Bruker Omega spectrometer operating at 100.61 MHz. Free-induction decays were digitized into 32 K points and were routinely multiplied by a 0.5- to 1.0-Hz exponential before Fourier transformation. Proton-decoupled 13C NMR spectra of urine extracts were obtained with a Unity INOVA 14.5 T spectrometer operating at 150.9 MHz (Varian Instruments, Palo Alto, CA). Free-induction decays were digitized into 44 K points and were multiplied by a 0.5-Hz exponential before Fourier transformation. Spectra from both systems were analyzed using the curve-fitting routine supplied with the NUTS PC-based NMR spectral analysis program (Acorn NMR, Fremont, CA). Typical numbers of acquisitions were 12,000-18,000 for each blood extract and 6,000-12,000 for each urine extract. Collection times per extract ranged from 4.5 to 13.5 h.
Quantitation of 13C enrichments. 13C enrichments of carbons in the glutamine moiety of phenylacetylglutamine were quantitated directly from the 13C NMR spectrum of the isolated phenylacetylglutamine by use of the method of Dugelay et al. (11). Gluconate 13C enrichments were quantitated directly from the 13C NMR spectrum by comparing the total area of each gluconate resonance with the C-2 resonance of the glycolate standard. Correction factors to account for T1 and nuclear Overhauser enhancement differences between the glycolate C-2 and gluconate carbons were derived from the time 0 blood extract, where all the gluconate carbons are at natural abundance enrichment.
Metabolic flux calculations.
Estimates of total anaplerotic inflow into oxaloacetate
(y), recycling of oxaloacetate carbons via
pyruvate (pk), and net gluconeogenic
flux (g) (all relative to citrate synthase
flux) were calculated from the
13C-13C
spin-coupled multiplets of gluconate C-2, phenylacetylglutamine C-2,
and glucuronate C-5 by use of methods described
previously (18).
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RESULTS |
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13C NMR spectra from blood samples. Figure 2 shows 13C NMR spectra of gluconate C-1 and C-2 from blood drawn from an individual at various times after oral ingestion of [U-13C]propionate. Initially, only singlets from natural abundance 13C were detected, but by 20 and 40 min [U-13C]propionate conversion into glucose was already evident from the appearance of 13C-13C spin-coupled multiplets in all six gluconate resonances. The intensity of the multiplet resonances increased gradually for 120 min and then remained relatively constant between 120 and 180 min. In all subjects, the nine-line C-2 gluconate multiplet was clearly formed at 120 min, and the ratios of multiplet components remained constant between 150 and 180 min. The time course of gluconate fractional enrichment from 0 to 180 min is shown in Fig. 3. Carbons 1, 2, 5, and 6 of gluconate showed identical enrichment kinetics, reaching a peak enrichment of ~4.5%1, whereas carbons 3 and 4 reached peak enrichments of ~2.5%. Between 120 and 180 min, enrichment of all six gluconate carbons remained relatively constant.
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Urine analysis. The time course of urinary phenylacetylglutamine enrichment was similar to that of blood glucose, with peak enrichments being achieved 2-3 h after ingestion of propionate (Fig. 5). However, the 13C fractional enrichment in C-1, C-2, and C-3 of phenylacetylglutamine was significantly lower than that of C-1, C-2, C-5, or C-6 of glucose or glucuronate (compare Figs. 3 and 5). This difference is thought to be due to dilution of the glutamine pool by inflow of unenriched glutamine from peripheral tissues such as skeletal muscle (24, 30). Figure 6 shows the 20- to 100-ppm region of a 13C NMR spectrum of urine collected 2 h after the last propionate ingestion. The C-2, C-3, and C-4 resonances of the glutamine moiety of phenylacetylglutamine are prominent, as is the natural abundance signal of the phenylacetyl methylene carbon at 43 ppm. The phenylacetylglutamine C-2 and C-3 resonances appear as multiplets arising from 13C-13C spin-coupling, whereas the C-4 resonance is largely a singlet (flanked by a weak doublet) arising primarily from natural abundance levels of 13C. This indicates that few of the 13C-enriched gluconeogenic precursors that leave the cycle actually reenter the cycle pools via acetyl-CoA, thus validating one important simplification of the metabolic model (18).
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Metabolic analysis.
For calculations of y,
pk, and
g, we selected blood spectra from 120 to 180 min, when the 13C
enrichment was relatively constant. These were compared with the 2- to
3-h and 3- to 4-h urine samples, which sampled the hepatic metabolite
pools at ~1.25-2.25 h and 2.25-3.25 h, respectively (7).
The gluconate C-2, glucuronate C-5, and phenylacetylglutamine C-2
multiplet areas (Table 1) were substituted into equations previously
presented (18) to give estimates of y,
pk, and
g (Table
2). For all three metabolites, total
anaplerotic flux relative to citrate synthase was estimated at
6.0-7.0. However, the phenylacetylglutamine analysis reported a
significantly higher OAA-PYR recycling flux than either glucose and
glucuronate, and this resulted in a significantly lower estimate of net
gluconeogenesis compared with the hexose analyses (1.4 vs. 2.4). These
data parallel the results of Landau et al. (24), in which net
gluconeogenic fluxes reported by glucose and phenylacetylglutamine
after infusion of
[14C]propionate were
~3.0 and ~1.8, respectively (24). With the exception of one
individual, the multiplet ratios of glucuronate C-5
and gluconate
C-2 in all subjects were similar, providing consistent values for
y,
pk, and
g (see Table 2). This indicates that
both triose components of the parent hepatic glucose-6-phosphate molecule had similar 13C
isotopomer distributions.
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DISCUSSION |
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Analytic considerations.
We have shown that complex 13C
isotopomer distributions of phenylacetylglutamine, glucuronate, and
glucose produced by hepatic metabolism of
[U-13C]propionate can
be analyzed in a straightforward manner by use of
13C NMR spectroscopy. The analyte
pools, particularly the urinary products, provide sufficient material
to quantitate isotopomer distributions by
13C NMR at
13C enrichment levels only
1-2% above natural abundance.
13C NMR proved to be a
particularly valuable tool for direct isotopomer analysis of the
glucuronide hexose skeleton after its enzymatic hydrolysis from the
parent acetaminophen -glucuronide. There are three advantages to the
methods reported here compared with previously published methods.
First, the determination of positional 13C glucose isotopomers appears to
be much simpler by 13C NMR than by
gas chromatography-mass spectrometry. Second, others have isolated
acetaminophen-
-glucuronide from urine and converted the glycone
moiety to glucose by chemical reduction of the carboxy glycone C-6
followed by enzymatic cleavage of the
-glucoside product (27, 28).
Again, the chemical approach reported here appears simpler and provides
complete recovery of labeled urinary glycones, an important
consideration given the relatively low sensitivity of
13C NMR compared with MS methods.
In situ hydrolysis of the
-glucuronide may also have advantages in
the clinical setting. A wide range of medications and drugs are cleared
by glucuronidation; hence, a patient has the potential to produce a
complex cocktail of urinary
-glucuronides even in the absence of
acetaminophen. For such an individual, collective hydrolysis in situ
should be far easier than quantitative isolation, reduction, and
hydrolysis of the complex glucuronide mixture. Finally, our simple
urine purification procedure retains both phenylacetylglutamine and
glucuronide, thereby allowing a
13C isotopomer analysis of both
metabolites from a single 13C NMR spectrum.
Ingestion of gram quantities of [U-13C]propionate. Because 13C is a nonradioactive isotope and bulk substitution of 12C by 13C is not known to have a significant kinetic isotopic effect, substantial quantities of 13C-enriched substrates can be safely given to humans (7, 13, 14, 35). However, such experiments can generate supraphysiological substrate levels that could alter basal metabolic activities. In our study, fasted individuals ingested an average of 1.5 g of propionate over 1 h. With the assumptions of quantitative capture of propionate by portal circulation and a portal vein blood flow of ~700 ml/min (10), one can estimate that the average portal propionate concentration is no higher than 0.36 mM over the 1-h period. Portal vein propionate concentrations averaging 0.03 mM have been reported in patients undergoing gall-bladder surgery (9), but these probably represent fasting levels, which are likely to be much less than postprandial concentrations. The only other measurements of human portal vein propionate concentrations have been obtained by autopsy of sudden death victims, for whom levels ranging from 0.02 to 0.19 mM were reported (8). Measurements of portal vein propionate levels in the pig, a good model for human digestion, indicated levels of 0.08 mM under fasting conditions and 0.25-0.40 mM after feeding (38). These postprandial levels span the estimated portal vein propionate levels generated during the experimental protocol. Nonetheless, the fact that urinary 3-[U-13C]hydroxypropionate was detected in several subjects suggests that the hepatic propionyl-CoA carboxylase can become saturated (1), and physiological consequences of saturating propionyl-CoA carboxylase in normal fasted humans are not known. In rat hepatocytes, high levels of propionate can inhibit the activities of several enzymes by lowering the free CoA levels (4, 5, 33), including pyruvate carboxylase. Because pyruvate carboxylase is the predominant anaplerotic pathway in the liver, inhibition of this enzyme can result in decreased gluconeogenic flux (4, 6). However, in fasted humans, the provision of 1.2 g of propionate over 3 h did not alter hepatic glucose production in fasted volunteers (26), suggesting that hepatic gluconeogenesis was maintained under these conditions. Our study featured comparable amounts of ingested propionate (~1.5 g), but the shorter ingestion period of our protocol probably generated higher levels of propionate in portal circulation. Nonetheless, our estimated portal vein propionate levels of 0.36 mM are still far below the 5 to 10 mM concentrations reported to inhibit gluconeogenesis and citric acid cycle fluxes (4, 6, 32).
The maximal contribution of propionate carbons to the total anaplerotic flux can also be estimated. From the study of Chandramouli et al. (6a), gluconeogenic flux in a 24-h-fasted individual is ~5.6 µmol · kgMetabolic flux estimates. Anaplerotic flux (y), determined here by 13C NMR analysis of glucose/glucuronate, is slightly lower (6.0 vs. 7.0) than that reported by Magnusson et al. (30), whereas pyruvate-oxaloacetate recycling (pk) flux was similar (~3.6 vs. 3.7). Although estimates of pk may be inflated by flux from the Cori cycle (23, 30), we believe that our study protocol minimized any contribution from extrahepatic pathways. The Cori cycle has been estimated to contribute maximally ~15% to the total hepatic glucose output in starved rats (23). Any lactate generated in extrahepatic tissues from 13C-enriched glucose and returned to the liver via the Cori cycle would supplement estimates of intrahepatic pyruvate-oxaloacetate recycling. With the assumption of steady-state conditions and no glycogenolysis, a 15% Cori cycle contribution would result in a 15% underestimation of g, and pk would be correspondingly increased (y being unchanged). The effects of this level of Cori cycling on pk are rather modest; 15% of g amounts to 0.36 units or ~10% of our pk estimate. However, it has been estimated that about one-third of the total hepatic glucose output is derived from glycogenolysis (6a) after a 24-h fast, so dilution of blood glucose with unlabeled glucose via glycogenolysis would further attenuate the 10% overestimate of pk to ~3%. Thus we conclude that the bulk of the observed pk flux activity detected in this study originates from hepatic metabolism.
Our estimate of net gluconeogenic flux was somewhat lower than the value (~2.4 vs. 3.3) reported by Magnusson et al. (30). One explanation could be the difference between the fasting periods in the two studies (24 h here vs. 60 h in Magnusson et al.). It has been shown that 3-carbon gluconeogenic sources contribute 67% of the total hepatic glucose output after a 22-h fasting period but >90% after a 42-h fasting period (6a). We would expect that this increased demand for 3-carbon precursors would be fulfilled in part by an increased ratio of gluconeogenic to citrate synthase flux. However, the additional gluconeogenic precursors could also be provided by a systematic increase in flux through all the pathways without any change in flux ratios. Hence, the modest differences between our flux estimates and those of Magnusson et al. may simply be methodological in origin. Conversion of relative to absolute fluxes by indexing g to the gluconeogenic portion of hepatic glucose output by use of additional tracers (15, 25, 30) will better resolve this uncertainty and is the focus of our ongoing studies. Finally, because net anaplerotic efflux can also be directed into nongluconeogenic pathways, such as glutathione biosynthesis, g represents the upper limit of glucose output from the tricarboxylic acid cycle.Effects of 13C reincorporation via bicarbonate on the measurements. Measurements of gluconeogenesis by use of singly labeled substrates such as [3-14C]lactate rely on determination of the ratio of label in glucose C-1/C-3 or glutamate C-2/C-1. These measurements are exquisitely sensitive to the reincorporation of label into glucose C-3 and glutamate C-1 via carboxylation of labeled bicarbonate (24, 30). To obtain the necessary correction factors, an additional labeling experiment using labeled bicarbonate must be performed (24, 30). By comparison, the glucose and phenylacetylglutamine multiplets observed in the 13C NMR spectra of these metabolites have differing degrees of sensitivity to enrichment from [13C]bicarbonate. The glucose C-1 (and C-6) and phenylacetylglutamine C-3 multiplets are completely insensitive to contributions from [13C]bicarbonate, because they originate from carbons 2 and 3 of oxaloacetate (18). After subtraction of natural abundance contributions, these multiplets give an estimate of g that requires no correction for contributions from [13C]bicarbonate. Estimates of g from the gluconate C-6 multiplets3 (2.39 ± 0.84) and the phenylacetylglutamine C-3 multiplets (1.67 ± 0.71) are in good agreement with estimates of g from the C-2 multiplets (see Table 2). This suggests that the C-2 isotopomer ratios of these metabolites were not significantly weighted by 13C enrichment from bicarbonate.
Conflicting isotopomer distributions and flux estimates from hexose and phenylacetylglutamine. The different flux estimates reported by hexose and phenylacetylglutamine indicate that hepatic glucose-6-phosphate and glutamine are sampling oxaloacetate pools that are not isotopically equivalent. Phenylacetylglutamine appears to be sampling a citric acid cycle pool that has a higher level of oxaloacetate-pyruvate recycling than the one sampled by either glucose or glucuronide. Labeling contributions from peripheral tissues such as the heart and skeletal muscle are unlikely to skew the flux estimates in this direction, because these tissues possess only modest anaplerosis and no significant OAA-PYR recycling activities (16, 31, 36, 37). Therefore, the oxaloacetate pool reported by phenylacetylglutamine is probably hepatic in origin, demonstrating some degree of metabolic zonation or compartmentation within the intact human liver, at least with respect to propionate. It is well established that hepatic glucose synthesis is highest in the periportal region, whereas glutamine synthetase is highly localized in the distal perivenous region. The high PYR-OAA recycling flux reported by phenylacetylglutamine does match an important characteristic of perivenous metabolism: a higher ratio of pyruvate kinase to phosphoenolpyruvate-carboxykinase activity compared with that found in the periportal zone (17, 19, 20, 22). This suggestion has to be tempered by the fact that the perivenous zone is also reported to have relatively high pyruvate dehydrogenase activity (22), which would be expected to result in significant acetyl-CoA and phenylacetylglutamine C-4 enrichment. Also, a significant fraction of glutamine carbon skeletons are thought to originate from outside the perivenous region, including periportal contributions (in the form of glutamate).
Why are these labeling differences revealed with propionate but not with lactate? A key difference in the hepatic metabolism of propionate vs. lactate is that the former is quantitatively extracted from the circulation, whereas the latter is not (34). Because mitochondrial density is highest in the periportal region of the liver (39), it is possible that a propionate tracer is preferentially consumed by the periportal cells compared with a lactate tracer. Therefore, downstream regions could receive little direct labeling from propionate while receiving significant contributions from secondary labeled products of periportal propionate metabolism, including glucose. Reutilization of hepatic glucose by the perivenous region is consistent with the zonal model of regulated hepatic glucose output proposed by Jungermann and colleagues (19-22), in which periportal glucose synthesis is appropriately attenuated by perivenal glucose reutilization. Utilization of secondary labeled products by the perivenous region could generate quite a different oxaloacetate isotopomer distribution than that produced from periportal metabolism of [U-13C]propionate if the two regions have different metabolic flux characteristics. In contrast, a lactate tracer is metabolized more uniformly across the lobule and therefore generates an averaged labeling distribution in glucose and glutamine. The fact that glucose and glutamate were labeled equivalently in perfused rat livers supplied with supraphysiological and saturating levels of [U-13C]propionate, lactate, and pyruvate (18) suggests that intralobular substrate gradients may be an important determinant of labeling heterogeneity among different metabolites in the liver. In conclusion, we have shown that a detailed analysis of the relative rates of anaplerosis, oxaloacetate-pyruvate cycling, and gluconeogenesis can be obtained in a rather simple procedure by 13C NMR analysis of blood and urine after ingestion of [U-13C]propionate, phenylacetate, and acetaminophen. Our results are in good agreement with other recently published studies that used 14C-labeled radioisotopes. Finally, the simple processing protocols for both blood and urine are well within the capability of a standard medical analytic laboratory and its personnel. ![]() |
ACKNOWLEDGEMENTS |
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We acknowledge the excellent technical assistance and support provided by the staff of the General Research Clinical Center at University of Texas Southwestern.
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FOOTNOTES |
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This research was supported by National Institutes of Health Grants RR-02584, HL-34557, and M01-RR-00633 and a Clinical Investigator Award from the Department of Veterans Affairs.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
1 Enrichment values include the natural abundance 13C contribution (assumed to be 1.11%).
2 Key assumptions are 1) isotopic steady-state conditions, 2) negligible labeling of acetyl-CoA, and 3) complete randomization of oxaloacetate carbons via fumarate.
3 Gluconate C-6 analysis was chosen over the C-1 carboxy terminus because of its higher signal-to-noise ratio and the possibility of faster relaxation of the C-1 doublet relative to the C-1 singlet.
Address for reprint requests: J. G. Jones, Mary Nell and Ralph B. Rogers Magnetic Resonance Center, 5801 Forest Park Rd., Dallas, TX 75235-9085.
Received 29 April 1998; accepted in final form 23 July 1998.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Ando, T.,
K. Rasmussen,
W. L. Nyhan,
and
D. Hull.
3-[U-13C]hydroxypropionate: significance of -oxidation of propionate in patients with propionic acidemia and methylmalonic acidemia.
Proc. Natl. Acad. Sci. USA
69:
2807-2811,
1972[Abstract].
2.
Bergman, E. N.
Energy contributions of volatile fatty acids from the gastrointestinal tract in various species.
Physiol. Rev.
70:
567-590,
1990
3.
Beylot, M.,
M. V. Soloviev,
F. David,
B. R. Landau,
and
H. Brunengraber.
Tracing hepatic gluconeogenesis relative to citric acid cycle activity in vitro and in vivo. Comparisons in the use of [3-13C]lactate, [2-13C]acetate, and alpha-keto[3-13C]isocaproate.
J. Biol. Chem.
270:
1509-1514,
1995
4.
Blair, J. B.,
D. E. Cook,
and
H. A. Lardy.
Interaction of propionate and lactate in the perfused rat liver. Effects of glucagon and oleate.
J. Biol. Chem.
248:
3608-3614,
1973
5.
Brass, E. P.
Interaction of carnitine and propionate with pyruvate oxidation by hepatocytes from clofibrate-treated rats: importance of coenzyme A availability.
J. Nutr.
122:
234-240,
1992[Medline].
6.
Chan, T. M.,
and
R. A. Freedland.
The effect of propionate on the metabolism of pyruvate and lactate in the perfused rat liver.
Biochem. J.
127:
539-543,
1972[Medline].
6a.
Chandramouli, V.,
K. Ekberg,
W. C. Schumann,
S. C. Kalhan,
J. Wahren,
and
B. R. Landau.
Quantifying gluconeogenesis during fasting.
Am. J. Physiol.
273 (Endocrinol. Metab. 36):
E1209-E1215,
1997[Medline].
7.
Cline, G. W.,
D. L. Rothman,
I. Magnusson,
L. D. Katz,
and
G. I. Shulman.
13C-nuclear magnetic resonance spectroscopy studies of hepatic glucose metabolism in normal subjects and subjects with insulin-dependent diabetes mellitus.
J. Clin. Invest.
94:
2369-2376,
1994[Medline].
8.
Cummings, J. H.,
E. W. Pomare,
W. J. Branch,
C. P. Naylor,
and
G. T. Macfarlane.
Short chain fatty acids in human large intestine, portal, hepatic and venous blood.
Gut
28:
1221-1227,
1987[Abstract].
9.
Dankert, J.,
J. B. Zijlstra,
and
B. G. Wolthers.
Volatile fatty acids in human peripheral and portal blood: quantitative determination vacuum distillation and gas chromatography.
Clin. Chim. Acta
110:
301-307,
1981[Medline].
10.
Doi, R.,
K. Inoue,
M. Kogire,
S. Sumi,
K. Takaori,
T. Suzuki,
and
T. Tobe.
Study on splanchnic circulation: measurement of the liver blood flow.
Nippon Geka. Gakkai. Zasshi.
89:
560-567,
1988[Medline].
11.
Dugelay, S.,
D. Yang,
M. V. Soloviev,
S. F. Previs,
K. C. Agarwal,
C. A. Fernandez,
and
H. Brunengraber.
Assay of the concentration and 13C-labeling pattern of phenylacetylglutamine by nuclear magnetic resonance.
Anal. Biochem.
221:
368-373,
1994[Medline].
12.
Ekberg, K.,
V. Chandramouli,
K. Kumaran,
W. C. Schumann,
J. Wahren,
and
B. R. Landau.
Gluconeogenesis and glucuronidation in liver in vivo and the heterogeneity of hepatocyte function.
J. Biol. Chem.
270:
21715-21717,
1995
13.
Gruetter, R.,
E. J. Novotny,
S. D. Boulware,
G. F. Mason,
D. L. Rothman,
G. I. Shulman,
J. W. Prichard,
and
R. G. Shulman.
Localized 13C NMR spectroscopy in the human brain of amino acid labeling from D-[1-13C]glucose.
J. Neurochem.
63:
1377-1385,
1994[Medline].
14.
Gruetter, R.,
E. J. Novotny,
S. D. Boulware,
D. L. Rothman,
G. F. Mason,
G. I. Shulman,
R. G. Shulman,
and
W. V. Tamborlane.
Direct measurement of brain glucose concentrations in humans by 13C NMR spectroscopy.
Proc. Natl. Acad. Sci. USA
89:
1109-1112,
1992[Abstract].
15.
Hellerstein, M. K.,
R. A. Neese,
P. Linfoot,
M. Christiansen,
S. Turner,
and
A. Letscher.
Hepatic gluconeogenic fluxes and glycogen turnover during fasting in humans. A stable isotope study.
J. Clin. Invest.
100:
1305-1319,
1997
16.
Jeffrey, F. M. H.,
C. J. Storey,
A. D. Sherry,
and
C. R. Malloy.
13C isotopomer model for estimation of anaplerotic substrate oxidation via acetyl-CoA.
Am. J. Physiol.
271 (Endocrinol. Metab. 34):
E788-E799,
1996
17.
Jones, C. G.,
and
M. A. Titheradge.
Measurement of metabolic fluxes through pyruvate kinase, phosphoenolpyruvate carboxykinase, pyruvate dehydrogenase, and pyruvate carboxylate in hepatocytes of different acinar origin.
Arch. Biochem. Biophys.
326:
202-206,
1996[Medline].
18.
Jones, J. G.,
R. Naidoo,
A. D. Sherry,
F. M. H. Jeffrey,
G. L. Cottam,
and
C. R. Malloy.
Measurement of gluconeogenesis and pyruvate recycling in the rat liver: a simple analysis of glucose and glutamate isotopomers during metabolism of [1,2,3-13C3]propionate.
FEBS Lett.
412:
131-137,
1997[Medline].
19.
Jungermann, K.
Metabolic zonation of liver parenchyma. Regulation of the glucostat of the liver.
Naturwissenschaften
72:
76-84,
1985[Medline].
20.
Jungermann, K.
Metabolic zonation of liver parenchyma: significance for the regulation of glycogen metabolism, gluconeogenesis, and glycolysis.
Diabetes. Metab. Rev.
3:
269-293,
1987[Medline].
21.
Jungermann, K.,
R. Heilbronn,
N. Katz,
and
D. Sasse.
The glucose/glucose-6-phosphate cycle in the periportal and perivenous zone of rat liver.
Eur. J. Biochem.
123:
429-436,
1982[Abstract].
22.
Jungermann, K.,
and
N. Katz.
Functional specialization of different hepatocyte populations.
Physiol. Rev.
69:
708-764,
1989
23.
Katz, J.,
P. Wals,
and
W. N. P. Lee.
Isotopomer studies of gluconeogenesis and the Krebs cycle with 13C-labeled lactate.
J. Biol. Chem.
268:
25509-25521,
1993
24.
Landau, B. R.,
W. C. Schumann,
V. Chandramouli,
I. Magnusson,
K. Kumaran,
and
J. Wahren.
14C-labeled propionate metabolism in vivo and estimates of hepatic gluconeogenesis relative to Krebs cycle flux.
Am. J. Physiol.
265 (Endocrinol. Metab. 28):
E636-E647,
1993
26.
Laurent, C.,
C. Simoneau,
L. Marks,
S. Braschi,
M. Champ,
B. Charbonnel,
and
M. Krempf.
Effect of acetate and propionate on fasting hepatic glucose production in humans.
Eur. J. Clin. Nutr.
49:
484-491,
1995[Medline].
27.
Magnusson, I.,
V. Chandramouli,
W. C. Schumann,
K. Kumaran,
J. Wahren,
and
B. R. Landau.
Quantitation of the pathways of hepatic glycogen formation on ingesting a glucose load.
J. Clin. Invest.
80:
1748-1754,
1987[Medline].
28.
Magnusson, I.,
V. Chandramouli,
W. C. Schumann,
K. Kumaran,
J. Wahren,
and
B. R. Landau.
Pentose pathway in human liver.
Proc. Natl. Acad. Sci. USA
85:
4682-4685,
1988[Abstract].
29.
Magnusson, I.,
V. Chandramouli,
W. C. Schumann,
K. Kumaran,
J. Wahren,
and
B. R. Landau.
Pathways of hepatic glycogen formation in humans following ingestion of a glucose load in the fed state.
Metabolism
38:
583-585,
1989[Medline].
30.
Magnusson, I.,
W. C. Schumann,
G. E. Bartsch,
V. Chandramouli,
K. Kumaran,
J. Wahren,
and
B. R. Landau.
Noninvasive tracing of Krebs cycle metabolism in liver.
J. Biol. Chem.
266:
6975-6984,
1991
31.
Malloy, C. R.,
A. D. Sherry,
and
F. M. H. Jeffrey.
Carbon flux through citric acid cycle pathways in perfused heart by 13C NMR spectroscopy.
FEBS Lett.
212:
58-62,
1987[Medline].
32.
Matsuishi, T.,
D. A. Stumpf,
M. Seliem,
L. A. Eguren,
and
K. Chrislip.
Propionate mitochondrial toxicity in liver and skeletal muscle: acyl CoA levels.
Biochem. Med. Metab. Biol.
45:
244-253,
1991[Medline].
33.
Patel, T. B.,
M. S. DeBuysere,
and
M. S. Olson.
The effect of propionate on the regulation of the pyruvate dehydrogenase complex in the rat liver.
Arch. Biochem. Biophys.
220:
405-414,
1983[Medline].
34.
Remesy, C.,
and
C. Demigne.
Changes in availability of glucogenic and ketogenic substrates and liver metabolism in fed or starved rats.
Ann. Nutr. Metab.
27:
57-70,
1983[Medline].
35.
Rothman, D. L.,
E. J. Novotny,
G. I. Shulman,
A. M. Howseman,
O. A. Petroff,
G. Mason,
T. Nixon,
C. C. Hanstock,
J. W. Prichard,
and
R. G. Shulman.
1H-{13C}NMR measurements of [4-13C]glutamate turnover in human brain.
Proc. Natl. Acad. Sci. USA
89:
9603-9606,
1992[Abstract].
36.
Sherry, A. D.,
C. R. Malloy,
R. E. Roby,
A. Rajagopal,
and
F. M. H. Jeffrey.
Propionate metabolism in the rat heart by 13C NMR spectroscopy.
Biochem. J.
254:
593-598,
1988[Medline].
37.
Szczepaniak, L.,
E. E. Babcock,
C. R. Malloy,
and
A. D. Sherry.
Oxidation of acetate in rabbit skeletal muscle: detection by 13C NMR spectroscopy in vivo.
Magn. Reson. Med.
36:
451-457,
1996[Medline].
38.
Topping, D. L.,
R. J. Illman,
M. N. Taylor,
and
G. H. McIntosh.
Effects of wheat bran and porridge oats on hepatic portal venous volatile fatty acids in the pig.
Ann. Nutr. Metab.
29:
325-331,
1985[Medline].
39.
Wimmer, M.,
and
D. Pette.
Microphotometric studies on intraacinar enzyme distribution in rat liver.
Histochemistry
64:
23-33,
1979[Medline].
40.
Yang, D.,
M. Beylot,
K. C. Agarwal,
M. V. Soloviev,
and
H. Brunengraber.
Assay of the human liver citric acid cycle probe phenylacetylglutamine and of phenylacetate in plasma by gas chromatography-mass spectrometry.
Anal. Biochem.
212:
277-282,
1993[Medline].
41.
Yang, D.,
S. F. Previs,
C. A. Fernandez,
S. Dugelay,
M. V. Soloviev,
J. W. Hazey,
K. C. Agarwal,
W. C. Levine,
F. David,
P. Rinaldo,
M. Beylot,
and
H. Brunengraber.
Noninvasive probing of citric acid cycle intermediates in primate liver with phenylacetylglutamine.
Am. J. Physiol.
270 (Endocrinol. Metab. 33):
E882-E889,
1996