Department of Molecular Physiology and Biophysics and Diabetes Research and Training Center, Vanderbilt University School of Medicine, Nashville, Tennessee
Submitted 28 July 2003 ; accepted in final form 29 September 2003
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ABSTRACT |
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hypoglycemia; hyperglycemia; portal vein; glucosensor
Recently, it has been reported that portal glucose infusion at a rate equivalent to the basal endogenous glucose production (EGP) caused hypoglycemia in 6-h-fasted conscious mice (7). In contrast, infusion of glucose into the femoral vein at the same rate resulted in significant hyperglycemia (7). Hypoglycemia was averted when somatostatin was infused simultaneously into the portal vein, and it did not occur in GLUT2 knockout mice (6), suggesting that a hepatoportal glucose sensor, requiring GLUT2 for its action, was inhibited by somatostatin (6, 7). These findings led us to conduct analogous studies, i.e., infusion of glucose via the portal or peripheral circulation at a rate designed to mimic EGP in the dog to determine whether the findings in the mouse are applicable to other species.
The dog exhibits very precise regulation of glycemia, in part by coordinating glucose uptake in muscle and liver. The reduction of the proportion of the glucose extracted by nonhepatic tissues concomitant with the enhancement of NHGU by the portal signal is an example of this coordination. Similarly, during peripheral glucose infusion, there is marked enhancement of nonhepatic glucose uptake, with very little NHGU unless sufficient glucose is infused to create unphysiological (>12 mM) glycemic levels (see review in Ref. 29). Because of this precise reciprocity of hepatic and nonhepatic glucose uptake, arterial blood glucose concentrations were virtually identical with the two delivery routes when glucose was infused into conscious dogs at 55.5 µmol·kg1·min1 via the hepatic portal or a peripheral vein. Therefore, we hypothesized that glycemia would be indistinguishable in dogs receiving peripheral or intraportal glucose infusions at a rate equivalent to resting glucose production.
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RESEARCH DESIGN AND METHODS |
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Approximately 16 days before study, each dog underwent a laparotomy, with insertion of sampling catheters in a hepatic vein, the hepatic portal vein, and a femoral artery and infusion catheters into splenic and jejunal veins (38, 39). Ultrasonic flow probes (Transonic Systems, Ithaca, NY) were placed around the portal vein and the hepatic artery. Dogs were used for study only if they met established criteria (38).
On the morning of the study, catheters and flow probe leads were exteriorized from their subcutaneous pockets, and catheter contents were aspirated. The splenic and jejunal catheters were used for intraportal infusion of glucose (when applicable). Angiocaths (Deseret Medical, Sandy, UT) were inserted into two peripheral veins.
Experimental design. Each experiment consisted of a 100-min equilibration period (140 to 40 min), a 40-min basal period (40 to 0 min), and a 180-min experimental period (0 to 180 min). At 140 min, a primed (36 µCi)-constant infusion of [3-3H]glucose (0.35 µCi/min) was initiated via peripheral vein. A constant infusion of cold glucose at 13.7 µmol·kg1·min1 was started at 0 min and continued until 180 min. This was equivalent to the rate of tracer-determined EGP in the basal state in 42-h-fasted dogs previously studied in our laboratory (21, 38). Five of the dogs (PO group) received the glucose intraportally, and five (PE group) received it via a peripheral vein. Femoral artery, portal vein, and hepatic vein blood samples were taken every 20 min during the basal period and every 30 min during the experimental period, as previously described (39).
Processing and analysis of samples. Hematocrit; blood lactate, alanine, and glycerol; plasma glucose, nonesterified fatty acid (NEFA), insulin, and glucagon concentrations, and [3H]glucose were determined as described previously (21, 38, 48).
Calculations and data analysis. The rate of substrate delivery to the liver, or hepatic substrate load, was calculated by a direct (D) method, as loadin (D) = ([S]A x AF) + ([S]P x PF), where [S] is the substrate concentration, A and P refer to artery and portal vein, respectively, and AF and PF refer to blood or plasma flow (as appropriate) through the hepatic artery and portal vein, respectively. Hepatic glucose load was also calculated by an indirect (I) method: loadin (I) = (GA x HF) + GIRPo GUG, where GA is the arterial blood glucose concentration, HF refers to total hepatic blood flow, GIRPo is the intraportal glucose infusion rate, and GUG is the uptake of glucose by the gastrointestinal tract (34).
The load of a substrate exiting the liver was calculated as loadout = ([S]H x HBF), where H represents the hepatic vein. Net hepatic balance was calculated as NHB = loadout - loadin. NHB for glucose (NHGB) was calculated with both the direct and indirect calculations. Only the direct calculation was employed for substrates other than glucose. A negative value indicates net uptake by the liver. Nonhepatic glucose uptake (non-HGU) was calculated as the glucose infusion rate minus NHGB. During the 1st h of glucose infusion, the non-HGU was corrected for the glucose required to fill the pool, with a pool fraction of 0.65 (11) and an assumption that the volume of distribution for glucose equaled the volume of the extracellular fluid, or 22% of the dog's weight (49). Net fractional glucose extraction by the liver (FE) was calculated by direct and indirect methods as NHGB/loadin. Net hepatic carbon retention, an index of the carbon available for glycogen synthesis, equaled NHGU + net hepatic uptake of glycerol + (2 x net hepatic uptake alanine) net hepatic lactate output; we have previously determined that doubling the net hepatic uptake of alanine provides a close approximation of net hepatic amino acid uptake (48). Glucose concentrations were converted from plasma to blood values by using correction factors (ratio of the blood to the plasma concentration) previously described (38). The rates of glucose appearance (Ra) and disappearance (Rd) were calculated with a two-compartment model by use of dog parameters (16, 27). In PE, the glucose infusion rate was subtracted from total Ra to yield endogenous Ra (EndoRa). EndoRa in PO = total Ra [glucose infusion rate x (1 FE)]. Unidirectional hepatic glucose uptake (HGU) was equal to the NHB of [3H]glucose, when the direct calculation was used and when dividing by the inflowing [3-3H]glucose specific activity (in dpm/µmol glucose). Hepatic glucose release (HGR) was NHGB HGU.
Hepatic sinusoidal insulin and glucagon concentrations were calculated as loadin (D)/HF by use of plasma concentrations and plasma flow data. This provides an estimate of the hormone concentrations at the beginning of the sinusoid, when the hepatic artery blood and portal vein blood become confluent.
Area under the curve (AUC) was calculated by the trapezoidal rule, using the change from basal in each animal. Results for the glucose infusion period are the mean for the entire period unless specified otherwise.
Statistical analysis. Data are presented as means ± SE. Time course data were analyzed with repeated-measures analysis of variance, with post hoc comparisons by the Student-Newman-Keuls procedure. A t-test was used for comparisons of AUC. Statistical significance was accepted at P < 0.05.
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RESULTS |
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The hepatic glucose load increased significantly in both groups during glucose infusion, although the change was modest (6 and 22% in PE and PO, respectively; P < 0.05 between groups). Even though the increment in the hepatic glucose load was greater in PO than in PE, the hepatic glucose load itself did not differ significantly between groups (Fig. 1).
NHGB, net hepatic glucose FE, non-HGU, and glucose turnover. The livers in PE and PO exhibited net hepatic glucose output (NHGO) at 8.2 ± 1.9 and 10.6 ± 2.1 µmol·kg1·min1, respectively, during the basal period (Fig. 2). In PE, there was a slight decrease (2.2 ± 1.4 µmol·kg1·min1) in NHGO after 30 min of glucose infusion. NHGO was suppressed to
0 µmol·kg1·min1 by 60 min, and eventually a low rate of NHGU was evident (averaging 2.5 ± 1.5 µmol·kg1·min1 during the last hour). In PO, there was a shift to NHGU at a rate of 2.5 µmol·kg1·min1 by 30 min, creating a change in NHGB of 13.1 ± 2.8 µmol·kg1·min1. By the last hour of the experiment, NHGU had increased to 4.1 ± 2.2 µmol·kg1·min1. The AUC of the change in NHGB from basal was 1.16 ± 0.21 mmol·kg1·3 h in PE vs. 1.94 ± 0.52 (direct calculation) and 2.30 ± 0.49 (indirect calculation) mmol·kg1·3 h in PO (NS between calculations for PO; P < 0.05 for PE vs. PO with both direct and indirect calculations). Because there were no significant differences between the results of the direct and the indirect calculations for any parameter, and the indirect calculation minimizes potential errors introduced by incomplete mixing of the glucose infusate with the blood in the portal vein, the results of the indirect calculation are used in Figs. 1 and 2 and the remainder of the text.
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Non-HGU increased significantly during peripheral glucose infusion, from a basal rate of 8.2 ± 1.9 to a maximum of 16.0 ± 1.9 µmol·kg1·min1, and then declined so that the mean during the last hour was 10.9 ± 1.3 µmol·kg1·min1 (Fig. 2). Non-HGU tended to decrease ( 3.7 ± 3.1 µmol·kg1·min1) initially after the portal glucose infusion began but then returned to basal values for the duration of the infusion (P < 0.05 between groups).
Net hepatic carbon retention during the first 90 min averaged 0.5 ± 1.0 and 5.6 ± 2.8 µmol·kg1·min1 in PE and PO, respectively (P < 0.05). During the last 90 min, it was very similar between groups, at 6.0 ± 1.2 and 8.2 ± 2.8 µmol·kg1·min1 in PE and PO, respectively.
Glucose EndoRa decreased similarly in both groups during glucose infusion (16.5 ± 0.7 to 6.9 ± 0.5 µmol·kg1·min1 in PE and 15.8 ± 0.8 to 7.5 ± 0.7 µmol·kg1·min1 in PO; Fig. 3). HGU did not differ between groups during the basal or glucose infusion periods (Fig. 4). HGR tended to be suppressed less in the PE than in the PO group ( from basal 5.7 ± 3.2 vs. 11.1 ± 2.0 µmol·kg1·min1; P = 0.07). Glucose Rd increased similarly in the two groups (
4.4 ± 1.4 and
4.7 ± 1.0 µmol·kg1·min1 in PE and PO, respectively; Fig. 3).
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Plasma hormone concentrations. Glucose infusion induced a significant rise in the insulin concentrations in both groups. The arterial plasma insulin concentration in PE increased from 41 ± 6 to a peak of 94 ± 34 pmol/l, with the AUC of the change from basal being 4,706 ± 2,391 pmol·l1·3 h (Fig. 5). In PO, the arterial concentration increased from 55 ± 14 to a peak of 120 ± 41 pmol/l, with the AUC of the change from basal being 7,502 ± 2,471 pmol·l1·3 h (P = 0.2 between groups). The hepatic sinusoidal insulin concentrations in PE increased from 113 ± 31 to a peak of 198 ± 70 pmol/l, with the AUC of the change totaling 13,849 ± 5,656 pmol·l1·3 h. The hepatic sinusoidal insulin concentrations in PO increased from 214 ± 82 to a peak of 427 ± 151 pmol/l (AUC of the change: 20,407 ± 11,425 pmol·l1·3 h; P = 0.4 between groups).
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Arterial and hepatic sinusoidal glucagon concentrations declined 1020% in both groups and were indistinguishable between the two groups throughout the studies (Table 1).
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Arterial concentrations and net hepatic balances of nonglucose substrates. Arterial blood concentrations of lactate increased 3040% in both groups during glucose infusion (Table 1). The groups exhibited net hepatic lactate uptake during the basal period and then shifted to a low rate of net hepatic lactate output in response to glucose infusion. Net hepatic lactate balance was near 0 µmol·kg1·min1 during the last hour of study in both groups. Neither the arterial concentrations nor the net hepatic balances of lactate differed between groups at any time.
Arterial alanine concentrations and net hepatic alanine uptake remained stable in both groups throughout the studies (Table 1). Arterial blood glycerol concentrations and net hepatic glycerol uptakes fell by 30% in both groups, and arterial plasma NEFA concentrations and net hepatic NEFA uptakes declined by
5565%.
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DISCUSSION |
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Portal glucose infusion resulted in a change in NHGB equivalent to 96% of the glucose infusion rate within 30 min. This change resulted from stimulation of HGU (5.2 ± 5.1 µmol·kg1·min1) and suppression of HGR (
7.8 ± 4.2 µmol·kg1·min1). Non-HGU had been suppressed by 3.7 ± 3.1 µmol·kg1·min1 by 30 min, however. Consequently, the arterial glucose concentration increased despite the near-complete compensation by the liver for the exogenous glucose. It is well known that, under hyperglycemic conditions, dogs demonstrate a rapid enhancement of NHGU and concomitant suppression of non-HGU in response to portally delivered glucose (18, 38). In this study, we demonstrate for the first time that both the liver and the nonhepatic tissues respond to the portal signal in the same way in the presence of only very mild hyperglycemia. It is worth noting that, whereas HGU was near-maximal at 30 min, HGR continued to decrease (nadir 0.3 ± 3.2 µmol·kg1·min1 at 150 min).
In contrast to the effect with portal glucose delivery, peripheral glucose infusion resulted in a much smaller decline in NHGO (equivalent to 16% of the glucose infusion rate) by 30 min. At that time, there was no stimulation of HGU (in fact, the rate tended to decline, with a change of 1.3 ± 4.1 µmol·kg1·min1 from basal) and a decline of only 3.4 ± 4.8 µmol·kg1·min1 in HGR. However, non-HGU had increased by 8.9 ± 1.2 µmol·kg1·min1, an amount equal to 65% of the glucose infusion rate. As a consequence, the arterial glucose concentration in PE increased to almost the same extent as in PO. Whether the enhancement of NHGU in PO vs. PE at 30 min resulted from the slightly larger increment in hepatic glucose load (despite the similarity between the absolute rate of glucose delivery to the liver during the PO and PE glucose infusion), modestly higher insulin levels, and/or the presence of the portal signal is not clear.
A striking difference in non-HGU was evident between the two groups for 90 min. Despite similar arterial glucose levels and a tendency toward higher arterial insulin concentrations in the presence of portal glucose delivery, the PO group displayed blunted non-HGU compared with that observed in PE. Relative peripheral insulin resistance persisted throughout the glucose infusion period in the PO group, with no stimulation of non-HGU evident despite continuously elevated insulin and glucose levels. The rapid enhancement of non-HGU in PE paralleled the increase in plasma insulin, and the gradual fall in non-HGU in that group followed the decline in the arterial insulin concentration. If the insulin concentrations had remained high in PE, as in PO, the difference in non-HGU between the groups evident over the first 90 min would likely have been sustained. The mechanism for the peripheral insulin resistance caused by portal glucose delivery is not clear but may be reciprocally linked to the increase in NHGU or a change in the central nervous system sympathetic outflow, as we will discuss.
In contrast to the findings in the dog, pig, and human, infusion of glucose at a rate equivalent to basal EGP in the mouse resulted in significant hyperglycemia (as high as 7.7 mmol/l), during peripheral glucose infusion, and hypoglycemia (as low as 2.3 mmol/l) during portal glucose infusion (7). Similar findings were previously reported in anesthetized cats (44). In six of the eight cats examined, slow intraportal injection (1 min) of small amounts (5.5, 27.8, or 55.5 µmol/kg) of an isotonic glucose solution induced a reduction (mean
1.1 mmol/l) in arterial blood glucose within 5 min that lasted >30 min. Conversely, injection of 5.5 and 27.8 µmol/kg of glucose into the radial vein had a mild hyperglycemic effect. Only at the 55.5 µmol/kg dose did peripheral glucose delivery produce a hypoglycemic effect, and even then, the decrease in blood glucose was much smaller (
0.09 vs. 0.28 mmol/l) and more transient (<15 vs. >30 min) than the corresponding response to portal glucose injection. It is difficult to put the data from cats into context, because no pancreatic hormone data were obtained and the presence of anesthesia complicates interpretation of the data. However, the discrepant findings in the response to low-rate peripheral and portal glucose infusions in dogs, pigs, and humans vs. mice and cats suggest species differences.
Species-specific responses to portal and peripheral glucose infusion might be explained by differences in the predominant tissues involved in glucose disposal among species. In the mouse, both peripheral and portal glucose infusion stimulated glucose utilization in the heart, brown adipose tissue (BAT), diaphragm, and soleus muscle, but not in the liver (7). Moreover, in the mouse, unlike the dog (Fig. 3), whole body glucose turnover was significantly greater with portal vs. peripheral glucose infusion. The heart and the BAT exhibited the greatest enhancement in glucose clearance during portal vs. peripheral glucose infusion in the mouse (7), suggesting that glucose uptake by these tissues was a major contributor to the development of hypoglycemia. Burcelin et al. (6, 7) proposed that the stimulation of peripheral glucose uptake was activated by a hepatoportal glucose sensor and that peripheral glucose delivery did not trigger the response. In contrast to the findings in the mouse, the liver in the dog has significant involvement in disposal of a glucose load whichever route of delivery is used, although it plays a much larger role in response to portal glucose. Myocardial glucose extraction is relatively low in both humans and dogs except under ischemic conditions (14, 33, 53), and adult humans and dogs have little or no BAT (9, 22, 23). Instead, skeletal muscle is the predominant nonhepatic tissue responsible for insulin-stimulated glucose disposal in the dog (18).
Portal glucose delivery is associated with a decrease in the firing rate in the hepatic branch of the splanchnic nerve and in the adrenal nerve (both primarily sympathetic) and an increase in efferent firing in the pancreatic branch of the vagus (primarily parasympathetic) nerve (36). Presumably, these effects remove an inhibition to hepatic glucose uptake (31) and stimulate insulin secretion (17).
In contrast to the decrease in sympathetic signaling to the liver, sympathetic nervous system (SNS) input to peripheral tissues, as evidenced by norepinephrine (NE) levels and turnover, is increased after mixed meals (40, 50) and oral carbohydrate intake (41, 47, 51, 52). Similarly, circulating serotonin (5-hydroxytryptamine, or 5-HT) concentrations rise following high-carbohydrate meals or glucose ingestion (3). NE is generally regarded as having an inhibitory effect on glucose disposal in vivo (10, 24), but this effect has not always been observed (46). The disparate findings may relate to dosages utilized in the different studies, because the dose response of NE on glucose and oxygen uptake in vitro is bell shaped (43), and/or to the differing study conditions, e.g., insulinemia and glycemia. Nevertheless, studies on isolated tissues indicate that sympathetic neurotransmitters can have different effects on different tissues. NE (25) and the 3-adrenoreceptor-specific agonist CL-316243 (15, 26) stimulate adipose tissue glucose uptake in the rat, with BAT having 100-fold greater capacity for NE stimulation of glucose uptake than white adipose tissue (25). On the other hand, NE reduces insulin-mediated glucose uptake in the perfused rat hindlimb (43). In regard to a potential effect of 5-HT on extrahepatic glucose uptake, it has been demonstrated that dexfenfluramine, a stimulator of 5-HT release and inhibitor of 5-HT reuptake, can increase glucose uptake in perfused rat white adipose tissue (4). However, 5-HT produced acute insulin resistance in the perfused rat hindlimb (35, 42). When individual muscles were examined, it was determined that those rich in slow-twitch oxidative fibers actually increased their glucose uptake during 5-HT perfusion, whereas the muscles with predominantly fast-twitch glycolytic fibers showed suppressed glucose uptake (42). Because the latter fiber type predominated in the hindlimb, the net effect was a reduction in glucose uptake (42). Thus a likely explanation of the discrepant findings of Burcelin et al. (7) and the current report is that SNS activity is responsible for the extrahepatic effects in both cases, and the results of the increase in SNS activity are dependent on the predominant tissues involved in glucose disposal. SNS activity markedly stimulates BAT glucose uptake, where that tissue is present, but can suppress insulin-mediated glucose uptake in skeletal muscle. Interestingly, the only skeletal muscle that Burcelin et al. observed to exhibit a substantial enhancement of glucose uptake after portal glucose infusion was the soleus (composed primarily of oxidative fibers), suggesting that 5-HT is involved in the extrahepatic effects of the portal signal.
In conclusion, during both portal and peripheral glucose delivery at a rate approximating the basal rate of endogenous glucose release, the dog was able to maintain blood glucose at a near-euglycemic level. There was, however, a pronounced difference in the adaptive mechanisms employed in the PE and PO groups, particularly during the early portion of the infusion period. During peripheral glucose infusion, there was a marked increase in glucose uptake by the nonhepatic tissues, with only a small suppression of NHGO occurring as a consequence of a fall in HGR. Portal glucose infusion, on the other hand, prompted a rapid conversion to NHGU stemming from both enhancement of HGU and suppression of HGR, but there was no stimulation (and in fact slight suppression) of non-HGU. This difference resulted in a marked alteration in the site of glucose storage, with an augmentation of the liver's role during portal glucose delivery, such that net hepatic carbon retention was equivalent to 41% of the infused glucose during the first 90 min in PO vs. 4% in PE. Because overall glucose Rd was similar in the two protocols, it is clear that muscle and liver played reciprocal roles. There is general agreement (6, 20, 28) that a hepatoportal glucose sensor is involved in the regulation of glucose metabolism, with both canine and murine data indicating that this sensor is involved in regulation of the disposition of portally delivered glucose in nonhepatic tissues. Nevertheless, it would appear that the sensor may operate differently in the mouse (where it directs glucose into the extrahepatic tissues even at the expense of hypoglycemia) and in the dog (where it partitions glucose between the liver and extrahepatic tissues in a manner that sustains near-euglycemia). Although further study of the hepatoportal glucose sensor mechanism in the mouse will undoubtedly yield useful insights, glucoregulation in the mouse does not appear to be as rigorous as it is in the dog (current data) or the human (54). Therefore, data regarding glucose sensing in the mouse must be viewed in context with data from other models before it can be extrapolated to human physiology.
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ACKNOWLEDGMENTS |
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Present address of S. Cardin: Wellstat Therapeutics Corporation, Gaithersburg, MD.
GRANTS
This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant R01-DK-18243 and Diabetes Research and Training Center Grant SP-60-AM-20593.
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FOOTNOTES |
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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. Section 1734 solely to indicate this fact.
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REFERENCES |
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