The head arterial glucose level is not the reference site for
generation of the portal signal in conscious dogs
Po-Shiuan
Hsieh,
Mary Courtney
Moore,
Bess
Marshall,
Michael J.
Pagliassotti,
Brian
Shay,
Dennis
Szurkus,
Doss W.
Neal, and
Alan D.
Cherrington
Department of Molecular Physiology and Biophysics, and Diabetes
Research and Training Center, Vanderbilt University, Nashville,
Tennessee 37232-0615
 |
ABSTRACT |
Experiments were performed on twelve
42-h-fasted, conscious dogs to determine whether the head arterial
glucose level is used as a reference standard for comparison with the
portal glucose level in bringing about the stimulatory effect of portal
glucose delivery on net hepatic glucose uptake (NHGU). Each experiment consisted of an 80-min equilibration, a 40-min control, and two 90-min
test periods. After the control period, somatostatin was given along
with insulin (7.2 pmol · kg
1 · min
1;
3.5-fold increase) and glucagon (0.6 ng · kg
1 · min
1;
basal) intraportally. Glucose was infused intraportally (22.2 µmol · kg
1 · min
1)
and peripherally as needed to double the hepatic glucose load. In one
test period, glucose was infused into both vertebral and carotid
arteries (HEADG; 22.2 ± 0.8 µmol · kg
1 · min
1);
in the other test period, saline was infused into the head arteries
(HEADS). One-half of the dogs
received HEADG first. When all
dogs are considered, the blood arterial-portal glucose gradients (
0.52 ± 0.07 vs.
0.49 ± 0.03 mM) and
the hepatic glucose loads (339 ± 14 vs. 334 ± 20 µmol · kg
1 · min
1)
were similar in HEADG and
HEADS. NHGU was 24.1 ± 3.8 and
25.1 ± 4.6 µmol · kg
1 · min
1,
and nonhepatic glucose uptake was 46.1 ± 4.2 and 48.8 ± 7.0 µmol · kg
1 · min
1
in HEADG and
HEADS, respectively. The head
arterial glucose level is not the reference standard used for
comparison with the portal glucose level in the generation of the
portal signal.
liver; brain; liver nerve
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INTRODUCTION |
FEEDING STUDIES (7, 28) have shown that splanchnic
removal of glucose is greater after oral glucose ingestion than after peripheral intravenous glucose administration. Moreover, Bergman et al.
(3) and Adkins et al. (1) reported similar hepatic glucose uptake after
intraportal glucose infusion as after oral glucose administration.
Their data thus suggest that the enhanced net hepatic glucose uptake
(NHGU) seen after oral glucose ingestion might occur as a result of the
entry and presence of elevated glucose levels in the portal venous
system. In studies carried out using perfused rat livers (11) or
conscious dogs (24), respectively, Gardemann et al. and Pagliassotti et
al. directly demonstrated that a negative arterial-portal glucose
gradient (glucose concentration in the artery lower than that in the
portal vein) is associated with augmented NHGU. We have attributed this effect to a "portal signal."
To date, only limited insight has been gained into the manner by which
the portal signal is generated. It is possible, therefore, that the
portal glucose level is compared with the arterial glucose level at
some as-yet-undetermined reference site to ascertain whether initiation
of the response is appropriate. It is known that the portal glucose
level can be sensed by glucose-sensitive cells in the portal vein that
signal the brain by use of vagal afferent fibers (20). A likely site
for sensing of the arterial glucose level is within the hypothalamus
(17). Neurophysiological studies have described the existence of neural
pathways that link the brain and the liver (8, 27). Other studies also
have demonstrated that an intact nerve supply to the liver appears to
be important for the normal response to intraduodenal or intraportal glucose delivery (2, 25). Taken together, these observations suggest
that the brain plays an important role in generation of the portal
signal. On the other hand, in studies of isolated perfused rat liver,
Gardemann et al. (11) and Stumpel and Jungermann (30) have suggested
that the portal signal could be sensed and transformed into a metabolic
signal within the liver itself. These observations point to the
potential importance of the hepatic arterial glucose level as a
reference standard used in generation of the portal signal.
The aim of the present study, therefore, was to determine whether
comparison of the brain arterial glucose level with the portal glucose
level initiates the stimulatory effect of portal glucose delivery on NHGU.
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METHODS |
Animals and surgical procedures.
Studies were carried out on twenty-two 42-h-fasted, conscious mongrel
dogs of either sex, weighing between 17 and 27 kg. All animals were
maintained on a diet of meat (Kal Kan, Vernon, CA) and chow (Purina Lab
Canine Diet no. 5006; Purina Mills, St. Louis, MO) composed of 34%
protein, 14.5% fat, 46% carbohydrate, and 5.5% fiber based on dry
weight. The protocols were approved by the Vanderbilt University
Medical Center Animal Care Committee, and animals were housed according
to American Association for the Accreditation of Laboratory Animal Care
International guidelines. Approximately 16 days before study, each dog
underwent a laparotomy under general anesthesia for the insertion of
blood sampling catheters into a hepatic vein, the hepatic portal vein,
and a femoral artery (18). Catheters were also placed in a splenic vein
and a jejunal vein for the infusion of solutions. Ultrasonic flow
probes (Transonic Systems, Ithaca, NY) were positioned around the
portal vein and hepatic artery, and their proximal ends were placed in
subcutaneous pockets. A second surgery was performed 8-9 days
before each experiment. A ventral midline incision was made under
general anesthesia, and Silastic catheters (Dow Corning, Midland, MI)
were inserted into the vertebral and carotid arteries bilaterally (5).
A catheter was also inserted into the left jugular vein to allow blood
sampling so that the success of the head glucose clamp could be
monitored. After insertion, the catheters were filled with glycerin-heparin (1,000 U/ml in a 1:1 ratio), their free ends were
knotted, and they were placed in subcutaneous pockets.
Approximately 2 days before study, blood was drawn to determine the
leukocyte count and the hematocrit of each animal. A dog was studied
only with a leukocyte count
<18,000/mm3, a hematocrit
>35%, normal stools, and if it had consumed all of its daily food
ration. On the morning of the study, the proximal ends of the flow
probes and surgically implanted catheters were exteriorized, the
catheters were cleared, the dog was placed in a Pavlov harness, and
intravenous access was established in three peripheral veins. At the
end of the study, an autopsy was performed on each animal to verify the
position of the carotid and vertebral catheter tips.
Experimental design.
At
120 min, a primed (36 µCi), continuous (0.3 µCi/min)
peripheral infusion of
D-[3-3H]glucose
and a continuous peripheral infusion of indocyanine green dye
(Becton-Dickinson, Cockeysville, MD; 4 µg · kg
1 · min
1)
were begun. The latter provided confirmation of hepatic vein catheter
placement and a second measurement of hepatic blood flow. After 80 min
(
120 to
40) of dye equilibration, there was a 40-min (
40 to 0) basal period, followed by two 90-min experimental
periods. At time 0, constant infusions
of several solutions were begun, and these infusions were continued
throughout the entire experiment. Somatostatin (0.8 µg · kg
1 · min
1;
Bachem, Torrance, CA) was infused to suppress endogenous insulin and
glucagon secretion. Insulin (1.2 mU · kg
1 · min
1)
and glucagon (0.6 ng · kg
1 · min
1)
were infused intraportally to raise the insulin level about three- to
fourfold and to keep the glucagon level basal. In addition, at
time 0, glucose (20% dextrose, Baxter
Healthcare, Deerfield, IL) was infused intraportally (22 µmol · kg
1 · min
1)
to activate the portal signal. The portal signal was then
present throughout both experimental periods in both protocols. At the same time, a primed, continuous peripheral infusion of 50% dextrose was begun so that the glucose load to the liver could quickly be
doubled. At this time the dogs were begun on one of two protocols. In
the first test period of protocol 1,
glucose was infused into the four head arteries to eliminate the
glucose gradient between arterial blood in the head and the portal
vein. The peripheral glucose infusion rate was reduced as required to
maintain the glucose load to the liver (HGL) at twofold basal. In the
second test period, saline was infused into the head instead of
glucose, and again the peripheral glucose infusion rate was adjusted to maintain a similar HGL to that seen in the previous period. The second
protocol was identical to the first, except that the order of the two
test periods was reversed.
Para-aminohippuric acid (PAH; Sigma
Chemical, St. Louis, MO; delivered at 1.7 µmol · kg
1 · min
1)
was added to the portal vein infusate to assess mixing of the infused
glucose with blood in the portal and hepatic veins, as described
previously (2, 23).
Processing and analysis of samples.
Plasma glucose was assayed using the glucose oxidase method with a
Beckman Glucose Analyzer II (Fullerton, CA). Plasma insulin and
glucagon concentrations were determined using radioimmunoassays (31).
Blood glucose and blood lactate levels were determined from perchloric
acid-treated samples according to the method of Lloyd et al. (16). PAH
was also measured in perchloric acid-deproteinized blood as previously
described (2, 19, 23).
Calculation.
When substrates are infused intraportally, the possibility of poor
mixing with the blood in the laminar flow of the portal circulation is
of concern. Mixing of the infused glucose in the portal vein was
assessed by comparing the recovery of PAH (which was mixed with the
portal glucose infusate) in the portal and hepatic veins with the PAH
infusion rate (2, 19, 23). Because of the magnitude of the coefficient
of variation for the method used in assessing PAH balance, samples were
considered statistically unmixed (i.e., 95% confidence that mixing did
not occur) if hepatic or portal vein recovery of PAH was 40% greater
or less than the actual amount of PAH infused (2, 19, 23). An
experiment was defined as having poor mixing (and was excluded from the
database) if a PAH recovery-to-infusion ratio of >1.4 or <0.6 was
observed at less than one of the three time points in each test period. Twenty-two dogs were studied; 10 were not included because of poor
mixing or unsuccessful glucose clamping. In the 12 dogs that were used
(n = 6/protocol), the ratio of PAH
recovery in the portal vein to the intraportal PAH infusion rate did
not differ (0.8 ± 0.1 and 0.8 ± 0.1 vs. 0.9 ± 0.1 and
0.9 ± 0.1, respectively) in the two test periods of the
two protocols. The ratios of PAH recovery in the hepatic vein to the
PAH infusion rate were also similar (0.9 ± 0.1 and 0.8 ± 0.1 vs. 1.0 ± 0.1 and 1.0 ± 0.1) during the two test periods of
protocols 1 and
2, respectively; a ratio of 1.0 would
represent perfect mixing. When a dog was retained in the database, all
of the points were used whether they were mixed or not, because mixing
errors occur randomly.
Determination of the rate of glucose infusion into the carotid and
vertebral arteries required to maintain the head arterial glucose level
at a level similar to the portal glucose level was based on the
following principle
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In the calculation of the cerebral glucose infusion rate,
cardiac output (CO) was estimated to be 140 ml · kg
1 · min
1
(26). Although estimates of CO in the dog range from 100 to 140 ml · kg
1 · min
1,
we choose the latter to ensure complete elimination of the glucose gradient between the head arteries and the portal vein. The carotid arterial blood flow was assumed to be 12% of CO; flow through the
vertebral arteries was assumed to be 6% of CO (26). Because glucose
was infused into the plasma compartment in both cases, carotid and
vertebral arterial plasma flows were used in the glucose infusion
calculation. In the study of Matsuhisa et al. (17), the carotid and
vertebral blood flows measured by Doppler flow probes were ~12 and
~5%, respectively, of CO if we assume that the average CO was 100 ml · kg
1 · min
1
and hemotocrit was 40%. If the CO in our dogs had averaged only 100 ml · kg
1 · min
1,
then we would have created a slightly higher glucose level in the head
arteries (
10%) than in the portal vein, as our present data showed.
Hepatic blood flow (HBF) was calculated by two methods, ultrasonic flow
probes and dye extraction (18). The results obtained with ultrasonic
flow probes and dye were not significantly different, but the data
shown in Figs. 1-4 are those obtained with the flow probes,
because their measurement did not require an assumption regarding the
distribution of the arterial and portal contributions to HBF.
The rate of substrate delivery to the liver, or hepatic substrate load,
was calculated by a direct (D) method as
where
[S] is the substrate concentration, A and P refer to artery
and portal vein, respectively, and ABF and PBF refer to blood flow
through the hepatic artery and portal vein, respectively. A similar
method was used to calculate the hepatic sinusoidal insulin and
glucagon concentrations
where
[H] is the hormone concentration and HS refers to the
hepatic sinusoids. To avoid any potential errors arising from either
incomplete mixing of glucose during intraportal glucose infusion or a
lack of precise measurement of the distribution of HBF, HGL was also
calculated by an indirect (I) method during the portal glucose infusion
period
where
G is the blood glucose concentration,
GIRPO is the intraportal glucose
infusion rate, and GUG is the uptake of glucose by the gastrointestinal
tract, calculated on the basis of the previously described relationship
between the arterial blood glucose concentration and GUG (2, 19, 23).
HGL did not differ whether measured using direct or indirect methods.
The load of a substrate exiting the liver was calculated as
where
H represents the hepatic vein.
Direct and indirect methods were used in calculation of net hepatic
glucose balance (NHGB). The direct calculation was
NHGBD = loadout
loadin (D). The indirect
calculation was NHGBI = loadout
loadin (I). The glucose data in
Figs. 1-4 represent those calculated with the indirect method, but
the values were not significantly different from these calculations
with the direct method. Lactate balance was calculated by the direct
method. Net fractional glucose extraction by the liver was calculated
as the ratio of NHGB (I) to loadin
(I). Nonhepatic glucose uptake (non-HGU) was calculated by subtracting
the rate of NHGU (I) from the total GIR. The net hepatic balance of
glucose equivalents was calculated as the sum of the balances of NHGB
(I) and lactate, once the latter had been converted to glucose
equivalents. This calculation ignores carbon derived from gluconeogenic
precursor uptake (
2.5
µmol · kg
1 · min
1)
and glucose used for oxidation (
1.5
µmol · kg
1 · min
1),
which tend to offset one another. Nevertheless, it provides an estimate
of the carbon available for glycogen deposition in the liver.
To calculate glucose balance, plasma glucose values were converted to
whole blood glucose values by using correction factors, as previously
described (23). Use of whole blood glucose ensures accurate NHGB
measurements regardless of the characteristics of glucose entry into
the erythrocyte.
Data are presented as means ± SE. SYSTAT (Evanston, IL) was used
for statistical analysis. The time course data were analyzed with
repeated-measures ANOVA, with post hoc analysis by univariate F tests. Results were considered
statistically significant at P < 0.05.
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RESULTS |
Plasma insulin and glucagon concentrations.
Arterial and liver sinusoidal insulin concentrations rose
similarly (
3.5-fold) in both groups during the experimental periods (Table 1 and Fig.
1), thus mimicking the insulin
concentrations seen in the postprandial state. Arterial and liver
sinusoidal glucagon levels remained basal and did not differ between
groups.
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Table 1.
The average of arterial plasma insulin and glucagon concentrations,
total glucose infusion rate, and net hepatic lactate balance during 2 test periods in 2 groups of 42-h-fasted, conscious dogs
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Fig. 1.
Calculated plasma insulin (A) and
glucagon (B) levels in the hepatic
sinusoid in 42-h-fasted conscious dogs during the basal and 2 test
periods in protocols 1 [head
glucose infusion (INF) followed by head saline] and
2 (head saline INF followed by head
glucose). Values are means ± SE; n = 6 for each period/protocol, except n = 5 in the 2nd test period of protocol
1. There were no significant differences between
groups.
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Blood glucose levels and HBF.
Blood glucose levels in the femoral artery and portal vein were
increased about twofold over basal and were not significantly different
between the two groups at any time (Fig.
2). In the first test period, intraportal
glucose infusion produced an arterial-portal blood glucose gradient of
0.55 ± 0.07 and
0.53 ± 0.08 mM in the
presence of head glucose and head saline infusion, respectively (Fig.
2). Likewise, in the second test period, the arterial-portal glucose
gradient was
0.62 ± 0.09 and
0.43 ± 0.05 mM in
the presence of head glucose and head saline infusion, respectively
(Fig. 2). Head glucose infusion, on the other hand, completely
eliminated the negative blood glucose gradient between the head
arteries (estimated) and the portal vein (0.23 ± 0.14 and 0.19 ± 0.20 mM in the first and second test periods of
protocols 1 and
2, respectively).

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Fig. 2.
Blood glucose levels in femoral artery, portal vein, and head arteries
(estimated) in 42-h-fasted conscious dogs during the basal and 2 test
periods in protocols 1 (A) and
2 (B). Values are means ± SE of 6 dogs for each period/protocol, except there were 5 dogs in the 2nd test
period of protocol 1. There were no
significant differences in blood glucose levels in femoral artery and
portal vein between groups.
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HBF was not different (32 ± 1 and 33 ± 2 vs. 34 ± 3 and 38 ± 3 ml · kg
1 · min
1
in periods 1 and
2 of protocols
1 and 2, respectively)
between test periods or between protocols.
HGL and NHGB.
The HGLs increased twofold in both groups (150 ± 10 to 310 ± 14 and 338 ± 22 vs. 155 ± 9 to 328 ± 26 and 367 ± 27 µmol · kg
1 · min
1;
basal to period 1 and
period 2) in
protocols 1 and
2, respectively (Fig.
3). NHGB changed from net outputs of 7 ± 2 and 11 ± 1 to net uptakes of 22 ± 3 and 21 ± 2 µmol · kg
1 · min
1
during the first test period in the presence and absence, respectively, of head glucose infusion. Likewise, in the second test period, NHGU was
26 ± 3 and 29 ± 6 µmol · kg
1 · min
1
in the presence and absence, respectively, of head glucose infusion. As
expected, on the basis of these data, the net fractional extraction of
glucose by the liver was not significantly different between groups
(data not shown). If one compares data within each group (Fig. 3), it
is also clear that head glucose infusion did not alter NHGU. Likewise,
if the data from both periods of both groups are pooled, there was no
effect of head glucose infusion on NHGU (24 ± 4 vs. 25 ± 5 µmol · kg
1 · min
1)
in the presence and absence, respectively, of head glucose infusion. When the direct method of calculation was used, NHGU was slightly but
not significantly less than with the indirect method; however, again
there was no difference between the presence (18 ± 2 µmol · kg
1 · min
1)
and absence (18 ± 3 µmol · kg
1 · min
1)
of head glucose infusion.

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Fig. 3.
Hepatic glucose load (A) and net
hepatic glucose balance (NHGB, B) in
42-h-fasted conscious dogs during the basal and 2 test periods in
protocols 1 and
2. Values are means ± SE of 6 dogs
for each protocol/period, except there were 5 dogs in 2nd test period
of protocol 1. There were no
significant differences between groups.
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Non-HGU.
Mean non-HGU increased to 43.6 ± 8.8 and 44.3 ± 7.0 µmol · kg
1 · min
1
during the first test period in the presence and absence, respectively, of head glucose infusion (Fig. 4) and did
not differ between the two groups. Similarly, in the
second test period, non-HGU was 51.7 ± 7.9 and 47.4 ± 12.3 µmol · kg
1 · min
1
in the presence and absence, respectively, of head glucose infusion. Likewise, if one examines the data within each protocol, one finds no
difference. If the data from both groups are pooled, non-HGU was 46.1 ± 4.2 and 48.8 ± 7.0 µmol · kg
1 · min
1
in the presence and absence, respectively, of head glucose infusion.

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Fig. 4.
Nonhepatic glucose uptake in 42-h-fasted conscious dogs during the
basal and 2 test periods in protocols
1 and 2. Values are
means ± SE of 6 dogs, except there were 5 dogs in 2nd test period of
protocol 1. There were no significant
differences between groups.
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In the first test period, the rates of total glucose infusion were not
different in the presence (65 ± 8 µmol · kg
1 · min
1)
and absence (65 ± 8 µmol · kg
1 · min
1)
of head glucose infusion. The glucose infusion rates rose modestly in
the second period of each protocol [85 ± 7 and 78 ± 9 µmol · kg
1 · min
1,
nonsignificant (NS)], presumably as a result of the progressively increasing effect of insulin. When the data were pooled, there were no
differences in the GIRs in the presence or absence of head glucose
infusion (72 ± 5 and 73 ± 7 µmol · kg
1 · min
1).
Net hepatic lactate balance.
Net hepatic lactate balance switched to output (4.0 ± 3.0 vs. 5.5 ± .3.5 µmol · kg
1 · min
1)
in the first test period in the presence and absence, respectively, of
head glucose (Table 1). In the second test period, net hepatic lactate
output was slightly lower (
1.2 ± 2.7 vs. 5.0 ± 3.1 µmol · kg
1 · min
1,
P = 0.17) during head saline than head
glucose infusion. When the data were pooled there was no difference in
the net hepatic lactate balance in the presence or absence of head
glucose infusion (4.8 ± 1.7 and 2.8 ± 1.2 µmol · kg
1 · min
1).
The net balance of glucose equivalents across the liver, which
represents the combination of glucose and lactate balance (after the
latter is converted to glucose equivalents) serves as an estimate of
the carbon used for glycogen deposition. Net balance of glucose equivalents across the liver switched to uptake of 17.8 ± 3.5 and
19.1 ± 3.4 µmol · kg
1 · min
1
(NS) during the first test period in the presence and absence, respectively, of head glucose infusion. Likewise, in the second test
period, the net balance of glucose equivalents across the liver was
23.9 ± 3.4 vs. 29.2 ± 5.4 µmol · kg
1 · min
1
(NS) in the presence and absence, respectively, of head glucose infusion. When all dogs are considered, the net balance of glucose equivalents across the liver was 21.5 ± 2.8 and 22.9 ± 3.5 µmol · kg
1 · min
1
in the presence and absence, respectively, of head glucose infusion. Head glucose infusion thus had no effect on the amount of carbon available for glycogen deposition in response to portal glucose infusion.
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DISCUSSION |
Under postprandial conditions, the liver responds rapidly and uniquely
to portal glucose delivery, suggesting that a signal in addition to
insulin, which acts slowly, is involved in stimulating hepatic glucose
uptake (4, 14, 15, 23) after feeding. Our previous studies (12, 13, 23)
have demonstrated that a signal generated when the portal glucose level
exceeds the arterial level plays such a role. Previous studies (10, 11,
17) have suggested that the two most likely reference sites for
comparison of arterial and portal glucose levels are in the brain or
liver. The present results suggest that the brain arterial glucose
level is not used as a reference standard for comparison with the
portal glucose level in generation of the portal signal and its effect on the liver. They leave open the possibility, as suggested by Gardemann et al. (11) and Stumpel et al. (30), that under postprandial
conditions it is the hepatic arterial glucose level that provides the
required reference information.
Previous studies (1, 10, 12, 13, 23) have demonstrated that the portal
signal not only enhances hepatic glucose uptake but also suppresses
non-HGU. The potential mechanism by which the extrahepatic effect of
portal glucose delivery occurs is still unknown. The present data
clearly indicate that the brain arterial glucose level did not provide
the reference information required to initiate the suppressive effect
of portal glucose delivery on peripheral glucose uptake. Thus neither
the effects of the portal signal on the liver nor those on muscle were
altered by eliminating the glucose gradient between the brain and the portal vein. Xie et al. (32-37) have demonstrated that hepatic denervation per se can produce insulin resistance in the skeletal muscle of the cat and rat. Their data suggest that the signal that
brings about the suppressive effect of portal glucose delivery on
peripheral glucose uptake may originate within the liver itself. Our
data thus are consistent with this hypothesis.
Although certain neurons in specific hypothalamic regions (21, 22, 29)
appear to be sensitive to changes in local and/or plasma glucose
concentrations, only one study by Matsuhisa et al. (17) has suggested
that the brain arterial glucose level is involved in the effect of the
portal signal on glucose uptake by the liver. Matsuhisa et al. utilized
conscious dogs and infused somatostatin along with intraportal
infusions of insulin (to create marked hyperinsulinemia) and glucagon
(at basal rates). In one test period, glucose was infused intraportally
(55.6 µmol · kg
1 · min
1);
in the next test period, the portal glucose infusion was continued, and
head glucose infusion was added through one carotid and one vertebral
artery at a rate calculated to eliminate the portal vein-to-head
arterial glucose gradient. Moderate hyperglycemia (8 mmol/l) was
maintained throughout the experiment. Eliminating the gradient between
the portal vein and the central nervous system diminished NHGU by
50% (42 ± 5 to 22 ± 3 µmol · kg
1 · min
1)
when the data from the last 30 min of each test period are considered, and the authors concluded that the brain was involved as a reference site for the portal signal. Caution should be used, however, in interpreting the findings of this study. First, although the authors infused the correct amount of glucose into the head, it was only given
in one carotid and one vertebral artery, so that the extent of glucose
mixing is unclear, and it seems likely that certain areas were above
and others below the portal glucose level. Second, the quantitative
accuracy of their balance data is not clear. Because the portal glucose
infusion rate and the portal blood flow were the same in the presence
and absence of the head glucose infusion, one would have expected the
glucose gradient between the femoral artery and the portal vein to be
the same in each period (
of
2.3 mmol/l). Instead, they were
different (
of 2.1 vs. 1.8 mmol/l), suggesting incomplete mixing of
the infusate in portal blood. This random A-P glucose difference
accounted for >40% of the difference in NHGU in the two periods.
Furthermore, if the indirect approach to calculating NHGU is used
(which eliminates the need to use the portal glucose level), the
difference in NHGU between the last two test periods is reduced to
15% and was probably not significant. Although these authors
assessed glucose mixing in the portal vein, they used the changes in
glucose to do so, rather than an independent measurement such as PAH.
The problem with this approach is that it does not allow the assessment
of mixing in the hepatic vein. The latter is critical if the accuracy of both the direct and indirect estimates of NHGB is to be validated.
The effect of the portal signal on hepatic glucose uptake has been
shown to turn on and off within 15 min (12, 13, 23). Knowing that is
the case, one would have expected elimination of the brain-portal
glucose gradient, if it were key to the initiation of the portal
signal, to quickly reduce NHGU. On the contrary, in the study of
Matsuhisa et al. (17) it took almost 1 h to see a diminution in NHGU.
Finally, Matsuhisa et al. administered all treatments in the same order
in each animal, and thus the lack of a time-matched control raises the
issue of what would have happened over time in the absence of head
glucose infusion. All of the above caveats weaken the conclusions that
can be drawn from the study of Matsuhisa et al.
To maximize the mixing of glucose in the head in the present study, we
infused glucose bilaterally through both carotid and both vertebral
arteries (5, 9). We then assessed the head glucose clamp, using a
catheter inserted into a jugular vein. With the assumption that 83% of
the infused glucose escapes the head on first pass (5, 6), the
estimated head arterial blood glucose levels were 11.0 ± 0.5 and
11.1 ± 0.2 mM in the presence of head glucose infusion in
protocols 1 and
2, respectively. This confirms that we
completely eliminated the negative glucose gradient between the head
arteries and portal vein. According to the calculation we have shown,
the head arterial glucose levels were 11 and 10% higher than the
portal vein glucose levels in protocols
1 and 2, respectively.
This was probably a result of the fact that we used a conservative
estimate of CO in our studies. We assumed a CO of 140 ml · kg
1 · min
1
to ensure complete elimination of the gradient in each dog. Some estimates of CO in the dog have been as low as 100 ml · kg
1 · min
1
(44). It is unlikely that a slight excess of glucose in the brain would
have had any effect, because cerebral glucose infusion itself is not
thought to affect NHGU (17).
In summary, under hyperglycemic hyperinsulinemic conditions, the
elimination of the negative glucose gradient between portal vein and
head arteries did not alter the effect of the portal signal on hepatic
or peripheral glucose uptake. This suggests that another reference site
must play an important role in sensing the arterial glucose level and
thereby triggering the response to portal glucose delivery.
 |
ACKNOWLEDGEMENTS |
We acknowledge the technical assistance of Wanda Snead and Pam
Venson in the Hormone Core Laboratory of the Vanderbilt University Medical Center Diabetes Research and Training Center.
 |
FOOTNOTES |
This work was supported by National Institute of Diabetes and Digestive
and Kidney Diseases Grant R-01-DK-43706 and Diabetes Research and
Training Center Grant SP-60-AM-20593.
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: M. C. Moore, 702 Light
Hall, Dept. of Molecular Physiology and Biophysics, Vanderbilt Univ.
School of Medicine, Nashville, TN 37232-0615.
Received 22 February 1999; accepted in final form 2 June 1999.
 |
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