Zonation of acetate labeling across the liver: implications
for studies of lipogenesis by MIDA
Michelle A.
Puchowicz1,
Ilya R.
Bederman1,
Blandine
Comte1,
Dawei
Yang1,
France
David1,
Eric
Stone2,
Kareem
Jabbour2,
David H.
Wasserman3, and
Henri
Brunengraber1
1 Department of Nutrition, Case
Western Reserve University, Cleveland, Ohio 44106; and Departments
of 2 Surgery and
3 Molecular
Physiology/Biophysics, Vanderbilt University, Nashville, Tennessee
37232
 |
ABSTRACT |
Measurement of fractional lipogenesis by mass
isotopomer distribution analysis (MIDA) of fatty acids or cholesterol
labeled from
[13C]acetate assumes
constant enrichment of lipogenic acetyl-CoA in all hepatocytes. This
would not be the case if uptake and release of acetate by the liver
resulted in transhepatic gradients of acetyl-CoA enrichment. Conscious
dogs, prefitted with transhepatic catheters, were infused with glucose
and
[1,2-13C2]acetate.
Stable concentrations and enrichments of acetate were measured in
artery (17 µM, 36%), portal vein (61 µM, 5.4%), and hepatic vein
(17 µM, 1.0%) and were computed for mixed blood entering the liver
(53 µM, 7.4%). We also measured balances of propionate and butyrate
across gut and liver. All gut release of propionate and butyrate is
taken up by the liver. The threefold decrease in acetate concentration
and the sevenfold decrease in acetate enrichment across the liver
strongly suggest that the enrichment of lipogenic acetyl-CoA decreases
across the liver. Thus fractional hepatic lipogenesis measured in vivo
by MIDA may be underestimated.
acetyl-coenzyme A; mass spectrometry; fatty acid synthesis; cholesterol synthesis
 |
INTRODUCTION |
MASS ISOTOPOMER DISTRIBUTION ANALYSIS (MIDA) was
proposed (11, 17) as a method for estimating the fractional synthetic rate of fatty acids and cholesterol synthesized in vivo and in vitro in
the presence of
[13C]acetate that
labels acetyl-CoA. Computation of the mass isotopomer distribution
(MID) of either fatty acids isolated from very low-density cholesterol
(VLDL)-triacylglycerols or cholesterol yields the fractional synthetic
rates and the enrichment of lipogenic acetyl-CoA. A variant of MIDA,
isotopomer spectral analysis (ISA), was presented by Kelleher and
Masterson (13).
The validity of MIDA and ISA requires that the enrichment of the
acetyl-CoA monomeric precursor be constant in all cells that synthesize
the polymer. When the enrichment of the precursor is not constant,
computation of the MID of the polymer yields erroneous values for the
enrichment of the monomeric precursor and for the fractional synthesis.
In fact, the calculated fractional synthesis is not the integrated
average enrichment of the precursor. This is because the precursor
enrichment appears in the equations for the amount of each isotopomer
as a nonlinear parameter. For a detailed discussion, see Ref. 3.
The goal of the project reported here was to test, in dogs infused with
[13C]acetate, whether
the concentration and 13C
enrichment of acetate vary across the liver. The study was prompted by
the recognition that acetate is generated in the liver by hydrolysis of
cytosolic, peroxisomal, and mitochondrial acetyl-CoA (4-5, 17a,
25, 29). One can therefore wonder whether, during infusion of
[13C]acetate, the
enrichment of blood acetate decreases across the liver lobule because
of the production of unlabeled acetate in the liver. Because acetate is
also generated by intestinal fermentation (4, 22), it was necessary to
monitor acetate concentrations and enrichments in arterial and portal
venous blood. This allowed us to calculate the concentration and
enrichment of acetate in the mixed blood that enters the liver via the
portal vein and hepatic artery. To simulate conditions where
lipogenesis is usually measured, the experiments were conducted under
hyperglycemic clamp (8-10 mM). Our data show a marked decrease in
the enrichment of acetate across the liver.
 |
METHODS |
Materials.
Chemicals were purchased from Sigma-Aldrich. Sodium
[1,2-13C2]acetate
and sodium
[1-13C,2H3]acetate
(99%) were from Isotec.
Animal experiments.
Mongrel dogs (18-26 kg) of either gender that had been fed a
standard diet (Kal Kan beef dinner, Vernon, CA, and Wayne Lab Blox:
51% carbohydrate, 31% protein, 11% fat, and 7% fiber based on dry
weight, Allied Mills, Chicago, IL) were studied. The dogs were housed
in a facility that met the American Association for Accreditation of
Laboratory Animal Care guidelines, and the protocols were approved by
Vanderbilt University's Institutional Animal Care and Use
Subcommittee. At least 16 days before each experiment, a laparotomy was
performed under general anesthesia (0.04 mg/kg of atropine and 15 mg/kg
pentothal sodium presurgery, and 1.0% isoflurane inhalation anesthetic
during surgery). Silastic catheters (0.03 in. ID) were inserted into
the vena cava for infusions. Silastic catheters (0.04 in. ID) were
inserted into the portal vein and left common hepatic vein for blood
sampling. Incisions were also made in the neck region for the placement
of a sampling catheter in the carotid artery. The carotid artery was
isolated, and a Silastic catheter (0.04 in. ID) was inserted so that
its tip rested in the aortic arch. After insertion, the catheters were
filled with saline containing heparin (200 U/ml, Abbott Laboratories, North Chicago, IL), and their free ends were knotted.
Ultrasonic flow probes (Transonic Systems, Ithaca, NY) were used to
measure portal vein and hepatic artery blood flows. Briefly, a small
section of the portal vein, upstream from its junction with the
gastroduodenal vein, was cleared of tissue, and a flow cuff (7.0 mm ID)
was placed around the vessel and secured. The gastroduodenal vein was
isolated and then ligated proximal to its coalescence with the portal
vein. A section of the main hepatic artery lying proximal to the portal
vein was isolated, and a flow cuff (3.0 mm ID) was placed around the
vessel and secured. The Doppler probe leads and the knotted free
catheter ends, with the exception of the carotid artery, were stored in
a subcutaneous pocket in the abdominal region so that complete closure
of the skin incision was possible. The free end of the carotid artery catheter was stored under the skin of the neck.
On the day of the experiment, the subcutaneous ends of the catheters
were freed through small skin incisions made while the dogs were under
local anesthesia (2% lidocaine, Astra Pharmaceutical Products,
Worcester, MA) over the subcutaneous pockets in which catheters were
stored. The contents of each catheter were aspirated, and catheters
were flushed with saline. Silastic tubing was connected to the exposed
catheters and brought to the back of the dog, where they were secured
with quick-drying glue. Saline was infused in the arterial catheter
throughout experiments (0.1 ml/min).
Indocyanine green (ICG) was infused at a rate of 0.1 mg · min
1 · m
2.
ICG was purchased from Hynson, Westcott, and Dunning (Baltimore, MD).
ICG was used as an independent back-up measurement of hepatic blood
flow measured using flow probes and as a means of confirming hepatic
vein catheter placement. Flow probe measurements of portal vein and
hepatic artery blood flows were continuously monitored on-line.
From 0 to 6 h, the dogs were infused intravenously with glucose at
variable rates (0.034 to 0.24 mmol · min
1 · kg
1)
to set up and maintain a stable plasma glucose concentration of 10 mM.
Also, from 0.5 to 6 h, 99% enriched
[1,2-13C2]acetate
was infused at 1.5 µmol · min
1 · kg
1
as a 150 mM solution. Blood was sampled from the carotid artery, portal
vein, and hepatic vein at regular intervals, treated with heparin, and
centrifuged, and plasma was frozen until analysis. Care was taken to
withdraw hepatic vein blood very slowly to prevent contaminating the
sample with blood drawn from the inferior vena cava.
Analytical procedures.
One set of plasma samples (0.25 ml) was spiked with internal standards
of
[1-13C,2H3]acetate
(50 nmol),
[2H5]propionate
(10 nmol), and
[2H7]butyrate
(10 nmol). The concentrations of plasma acetate, propionate, and
butyrate were determined by gas chromatography-mass spectrometry of the
2,4-difluoroaniline derivative (24). A second set of plasma samples
without internal standards was used to assay the molar percent
enrichments (MPE) of acetate (24). To avoid contaminations by
ubiquitous acetate, we dedicated an isolated laboratory to the assays.
For more details, see Ref. 24. Glucose was assayed enzymatically.
Calculations.
Isotopic enrichments are expressed as MPE, that is, the percentage of
molecules that are labeled. Some authors (18) express enrichments as
tracer-to-tracee ratios (TTR). The two expressions of enrichment are
related by Eq. 1
|
(1)
|
Because
the concentration and the enrichment of acetate are very different in
arterial and portal vein blood, the concentration and enrichment of
acetate in the mixed blood entering the liver were calculated using
Eqs. 2 and 3, in which PV and HA represent the
portal vein and the hepatic artery, respectively. Blood flows are
expressed in milliliters per minute per kilogram.
|
(2)
|
|
(3)
|
Note that the numerator of Eq. 2
represents the nanomoles of total acetate (unlabeled + labeled) that
enter the liver per minute. The numerator of Eq. 3 represents the nanomoles of labeled acetate that
enter the liver per minute. The denominator of Eq. 3 represents the nanomoles of total acetate (unlabeled + labeled) that enter the liver per minute. The extent of dilution of
acetate labeling across the liver and the hepatic acetate uptake were calculated from Eqs. 4 and 5, where HV represents the hepatic vein.
|
(4)
|
|
(5)
|
|
(6)
|
|
(7)
|
|
(8)
|
Equations 2, 5, 6, and 8 are also used to calculate
parameters of propionate and butyrate metabolism.
 |
RESULTS |
The variable glucose infusion resulted in a stable arterial glucose
concentration of 10-10.4 mM during the last 4.5 h of the experiment (not shown). The mean glucose infusion rate necessary to
achieve this glucose concentration was 0.11 ± 0.03 mmol · min
1 · kg
1
(SE, n = 6). Total hepatic
blood flow averaged 37 ± 2 ml · min
1 · kg
1,
and the portal vein flow contributed 80-85% of the total hepatic flow.
Figures
1-3
show the measured concentrations of acetate, propionate, and butyrate,
respectively, in the artery, portal vein, and hepatic vein, as well as
the calculated concentrations in the mixed blood entering the liver
(Eq. 2). The concentrations of the
three short-chain fatty acids are markedly higher in the portal vein
compared with the artery. For this reason, and because most of the
blood supply to the liver is through the portal vein, the
concentrations of these compounds in the mixed liver influent are close
to those in the portal vein. For each compound, the concentrations in
the artery and hepatic vein are very similar. For butyrate, the
arterial concentration was slightly, but significantly, higher than in
the hepatic vein. Figure 4 shows the fluxes
of acetate, propionate, and butyrate in and out of the liver, as well
as the percentages of the rates of inflowing substrates that are
released in the hepatic vein. Figure 5
compares the releases of acetate, propionate, and butyrate by the gut
to the uptake of these substrates by the liver. Within experimental
errors, all the gut production of the three compounds is taken up by
the liver in a single pass of the blood through the organ.

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Fig. 1.
Profile of acetate concentration in artery (ART), portal vein (PV),
hepatic vein (HV), and mixed liver influent (MIXED)
(Eq. 2). Data are presented as means ± SE (n = 6).
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Fig. 2.
Profile of propionate concentration in artery, portal vein, hepatic
vein, and mixed liver influent (Eq. 2). Data are presented as means ± SE
(n = 6).
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Fig. 3.
Profile of butyrate concentration in artery, portal vein, hepatic vein,
and mixed liver influent (Eq. 2).
Data are presented as means ± SE (n = 6).
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Fig. 4.
Fluxes of acetate (unlabeled and labeled), propionate, and butyrate in
and out of liver. Fluxes of substrate entering liver (IN) were
calculated from numerator of Eq. 2. In
the case of acetate, this represents the total flux of unlabeled + labeled substrate. The flux of
[1,2-13C2]acetate
entering the liver (IN) is calculated from the numerator of
Eq. 3. Fluxes of substrate exiting the
liver (OUT) are calculated from hepatic vein concentrations and blood
flows. Percentages of inflows that are released in the hepatic vein,
indicated above OUT bars, were calculated using Eqs.
6-8.
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Fig. 5.
Comparison between releases of acetate, propionate, and butyrate by gut
(left, Eq. 8) and uptakes of the same substrates by liver
(right, Eq. 6). Percentages of rates of substrate release by gut
that are taken up by liver are indicated on
right above each bar.
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Figure 6 shows the measured MPE of acetate
in the artery, portal vein, and hepatic vein, as well as the calculated
MPE of acetate in the mixed blood entering the liver
(Eq. 3). The highest acetate
enrichment was in arterial blood (36.1 ± 2.5% from 270 to 360 min,
n = 6). The acetate enrichment in the
portal vein (5.4 ± 0.8%) was 6.7 times lower than in arterial blood.
The enrichment of acetate in the mixed liver influent (7.4 ± 1.0%)
was only 1.4 times that in the portal vein. The enrichment of acetate
in the hepatic vein (1.0 ± 0.3%) was 7.4 times lower than in the
mixed liver influent and 36 times lower than in the artery.

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Fig. 6.
Profile of acetate molar percent enrichment (MPE) in artery, portal
vein, hepatic vein, and mixed liver influent (Eq. 3) of conscious dogs infused with glucose and
[1,2-13C2]acetate
in a peripheral vein.
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Figure 4 (left) shows another aspect
of the dilution of acetate enrichment across the liver. Although 32%
of the total acetate entering the liver left the organ through the
hepatic vein, the corresponding value for
[1,2-13C2]acetate
was 4.7%.
 |
DISCUSSION |
Balances of acetate, propionate, and butyrate across the dog gut and
liver (Fig. 5) confirm the notion (4, 22) that the gut is a major site
of production of these short-chain fatty acids. The very similar
concentrations of these compounds in the arterial and hepatic venous
blood (Figs. 1-3), as well as the equivalence between net gut
production and net hepatic uptake (Fig. 5) could lead one to believe
that these compounds are made only by the gut and taken up only by the
liver. However, the sevenfold decrease in acetate enrichment across the
liver (Fig. 4) reveals that the liver also generates unlabeled acetate.
Furthermore, because the acetate concentration in the hepatic vein is
one-third of that of the liver influent, most of the unlabeled acetate
generated in the liver is taken up by the same organ.
In mammalian cells, the only known mechanisms for acetate production
are the hydrolysis of acetyl-CoA (4, 5) and the metabolism of acetone
in the liver by the C2 pathway (8). In our dogs infused with glucose,
concentrations of ketone bodies were very low; thus acetyl-CoA
hydrolysis was probably the only source of acetate in liver. Our data
confirm that the liver is the site of an intense acetate
acetyl-CoA
acetate substrate cycle (5). Indeed, acetyl-CoA
synthetases and hydrolases have been described in liver mitochondria
and cytosol (4, 5, 25, 29). In perfused rat liver and hepatocyte
experiments conducted in the presence of nonacetate
14C or
13C tracers that label acetyl-CoA
in liver mitochondria, cytosol, or peroxisomes, labeled acetate was
released (5, 8, 14, 19, 30). Thus acetate
acetyl-CoA
acetate substrate cycles occur in liver cytosol and mitochondria.
Our data do not address the production of acetate by peripheral tissues
other than the gut. Bleiberg et al. (1) infused [1-14C]acetate in
normal dogs and demonstrated the simultaneous uptake and release of
acetate in intestine, liver, kidney, and hindlimb. Mittendorfer et al.
(18) infused
[1,2-13C2]acetate
in humans and found that the acetate enrichment and concentration in
the femoral vein were one-half of those in the artery. Thus muscle is
also a site of acetate cycling. The physiological role of this
substrate cycle is not known.
Based on the enrichment of acetate in arterial blood, the apparent
turnover rate of acetate was 2.8 ± 0.3 µmol · min
1 · kg
1,
which is similar to values obtained by Mittendorfer et al. in humans
(3.3 µmol · min
1 · kg
1,
Ref. 18) and by others using
[14C]acetate (1, 28).
Pouteau et al. (23) reported much higher rates measured with
[13C]acetate. The
apparent turnover rate we calculated is most likely underestimated
because of the combination of 1) the
very short apparent half-life of plasma acetate, which is of the order
of the circulatory time, ~30 s (6), and
2) the intravenous route of tracer
infusion with arterial blood sampling (V-A mode, Ref. 12). The
contribution of the gut to this apparent acetate turnover (Fig. 5) is
~50%.
Mittendorfer et al. (18) infused
[1,2-13C2]acetate
in normal humans at the same rate as we did in dogs (1.5 µmol · min
1 · kg
1).
The enrichment of arterial acetate they observed (0.447 TTR, equivalent
to 30.9 MPE) was similar to ours (36.1 MPE, Fig. 6). However, they
reported much higher acetate enrichment in the hepatic vein (0.134 TTR,
equivalent to 11.2% MPE) compared with our data (1% MPE). Therefore,
their artery-to-hepatic vein acetate enrichment ratio was ~3, whereas
ours was ~36. The reason for the discrepancy between data by
Mittendorfer et al. and ours is not clear. In addition to the species
difference of human vs. dog, the Mittendorfer et al. subjects were
studied after an overnight fast without glucose infusion. Our dogs were
infused with glucose so that the extent of acetate dilution across the
liver could be studied under lipogenic conditions. Conceivably, the
infusion of glucose increased acetyl-CoA production in the liver and
may have resulted in increased acetate
acetyl-CoA
acetate cycling. This would be a possible explanation of the difference
in acetate dilution across the liver between the Mittendorfer et al.
report and ours. However, against this interpretation are data from one
experiment we conducted on an overnight-fasted, anesthetized dog
without glucose infusion. There, the artery-to-hepatic vein enrichment
ratio was 27, and acetate enrichment across the liver decreased
7.2-fold, as in our experiments on conscious dogs infused with glucose.
The 7.4-fold decrease in acetate enrichment across the liver (Fig. 6)
must result in some decrease in the enrichment of lipogenic (cytosolic)
acetyl-CoA between periportal and perivenous hepatocytes. The extent of
decrease in acetyl-CoA enrichment and the shape of this decrease cannot
be deduced from the overall drop in acetate enrichment across the
liver. Although there is evidence that the activity of cytosolic
acetyl-CoA synthetase is two times greater in periportal than in
perivenous hepatocytes (15), we could find no information on the degree
of zonation of the activities of the mitochondrial acetyl-CoA
synthetase and mitochondrial/cytosolic/peroxisomal acetyl-CoA
hydrolases. There is evidence that both cytosolic and mitochondrial
acetyl-CoA synthetases participate in the activation of labeled acetate
that is incorporated into fatty acids and cholesterol (9). Cytosolic
acetyl-CoA synthetase labels directly the lipogenic acetyl-CoA pool. In
contrast, mitochondrial acetyl-CoA synthetase forms acetyl-CoA that
mixes with unlabeled acetyl-CoA derived from pyruvate and fatty acid
oxidations. The resulting mitochondrial acetyl-CoA is transferred to
the cytosol via citrate and ATP-citrate lyase (9).
Our calculation of the enrichment of acetate that enters the liver
assumes that the portal venous and hepatic arterial influents mix
perfectly at the entrance of the liver sinusoid. Mixing of the arterial
and portal influents might be incomplete, resulting in streaming of
portal and arterial blood through part of the length of the sinusoid.
If this were the case, acetyl-CoA would be much more enriched in
hepatocytes in contact with arterial blood than in adjacent hepatocytes
in contact with portal venous blood.
What are the implications of our data for measurements of fractional
lipogenesis (11) by MIDA of fatty acids isolated from VLDL-triglycerides or of plasma cholesterol after infusion of [13C]acetate? An
absolute condition of validity of this technique requires that the
enrichment of lipogenic acetyl-CoA be constant in all hepatocytes that
synthesize lipids (3). Our data suggest that this condition may not be
met when acetyl-CoA is labeled from a tracer of acetate. MIDA probably
underestimates to some extent fractional hepatic lipogenesis traced
with [13C]acetate
unless fatty acid synthesis is confined to a narrow band of hepatocytes
in the liver lobule. This is unlikely to be the case, because the
activity of the lipogenic enzyme acetyl-CoA carboxylase is only about
twofold higher in periportal than in pericentral hepatocytes (7). Thus
it is likely that lipogenesis extends, to some degree, into the
perivenous area of the lobule. Also, there is no variation in VLDL-
triacylglycerol secretion between periportal and perivenous hepatocytes
(10).
Conceptually, the possible underestimation of fractional lipogenesis
traced with
[13C]acetate is
similar to the underestimation of fractional gluconeogenesis traced
with [13C]glycerol.
The latter, like
[13C]acetate,
undergoes considerable dilution across the liver (27). In the case of
fractional gluconeogenesis, the degree of underestimation can be
estimated in humans and animals whose liver glycogen has been depleted
by fasting. Then the apparent contribution of gluconeogenesis to
glucose production, calculated from MIDA of glucose labeled from
[13C]glycerol, ranges
from 27 to 90% (16, 20, 21, 26, 27) instead of 100%. High apparent
values for fractional gluconeogenesis result from very high rates of
[13C]glycerol infusion
that blunt the periportal-to-perivenous gradient of glycerol enrichment
(20, 21).
In the case of lipogenesis traced with
[13C]acetate, the
technique just described is not applicable, because it is impossible to
set up conditions where all the fatty acids in VLDL-triglycerides would
be derived from de novo synthesis. However, a decrease in enrichment of
lipogenic acetyl-CoA across the liver could be detected by ISA (13) of
fatty acids isolated from VLDL-triglycerides. This requires that the
enrichment of acetate in the portal venous blood be sufficiently high
that at least three labeled mass isotopomers are present in the
long-chain fatty acids isolated from VLDL-triglycerides. If these
conditions are met, ISA could reveal nonhomogeneity in labeling of
lipogenic acetyl-CoA. We are presently testing this strategy.
 |
ACKNOWLEDGEMENTS |
The assistance of Anna Williams and Kevin Cherrington is gratefully acknowledged.
 |
FOOTNOTES |
This work was supported by grants from National Institute of Diabetes
and Digestive and Kidney Diseases (DK-35543) and the Cleveland Mt.
Sinai Medical Center.
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for correspondence and reprint requests: H. Brunengraber, Dept.
of Nutrition, Case Western Reserve University, Cleveland, OH
44106-4906.
Received 7 December 1998; accepted in final form 12 July 1999.
 |
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