Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine, Nashville, Tennessee 37232
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ABSTRACT |
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We previously reported that infection
decreases hepatic glucose uptake when glucose is given as a constant
peripheral glucose infusion (8 mg · kg1 · min
1). This impairment persisted despite greater
hyperinsulinemia in the infected group. In a normal setting, hepatic
glucose uptake can be further enhanced if glucose is given
gastrointestinally. Thus the aim of this study was to determine whether
hepatic glucose uptake is impaired during an infection when glucose is
given gastrointestinally. Thirty-six hours before study, a sham (SH,
n = 7) or Escherichia coli-containing
(2 × 109 organisms/kg; INF; n = 7)
fibrin clot was placed in the peritoneal cavity of chronically
catheterized dogs. After the 36 h, a glucose bolus (150 mg/kg)
followed by a continuous infusion (8 mg · kg
1
· min
1) of glucose was given intraduodenally to
conscious dogs for 240 min. Tracer ([3-3H]glucose and
[U-14C]glucose) and arterial-venous difference techniques
were used to assess hepatic and intestinal glucose metabolism.
Infection increased hepatic blood flow (35 ± 5 vs. 47 ± 3 ml · kg
1 · min
1; SH vs. INF)
and basal glucose rate of appearance (2.1 ± 0.2 vs. 3.3 ± 0.1 mg · kg
1 · min
1).
Arterial insulin concentrations increased similarly in SH and INF
during the last hour of glucose infusion (38 ± 8 vs. 46 ± 20 µU/ml), and arterial glucagon concentrations fell (62 ± 14 to 30 ± 3 vs. 624 ± 191 to 208 ± 97 pg/ml). Net
intestinal glucose absorption was decreased in INF, attenuating the
increase in blood glucose caused by the glucose load. Despite this, net
hepatic glucose uptake (1.6 ± 0.8 vs. 2.4 ± 0.9 mg · kg
1 · min
1; SH vs. INF) and
consequently tracer-determined glycogen synthesis (1.3 ± 0.3 vs.
1.0 ± 0.3 mg · kg
1 · min
1) were similar between groups. In summary, infection
impairs net glucose absorption, but not net hepatic glucose uptake or
glycogen deposition, when glucose is given intraduodenally.
alanine; lactate; inflammation; glycogen
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INTRODUCTION |
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INFECTION LEADS TO MARKED alterations in whole body glucose production and utilization. The increase in glucose production is predominantly caused by acceleration in gluconeogenesis, which is driven by elevated counterregulatory hormones and increased gluconeogenic precursor supply (15). In addition to the increase in glucose flux, whole body energy expenditure is increased (10). To meet the increased caloric requirements, many of these individuals require nutritional support.
Glucose intolerance and insulin resistance are often present during stressful conditions, especially when exogenous glucose must be administered. The hyperglycemia commonly seen in individuals with infections (11, 25) is a result of alterations in both liver and muscle glucose metabolism. Under feeding conditions, the liver is an important site for glucose disposal. We observed during a continuous intravenous infusion of glucose that the liver removed ~30% of the infused glucose in normal dogs (17). The ability of the liver to dispose of the glucose was decreased by infection (17). Net hepatic glucose uptake was decreased because of an attenuated suppression of hepatic glucose production, as well as augmentation of unidirectional hepatic glucose uptake (17). Thus, for the same glucose infusion rate, infection forced insulin-resistant peripheral tissues to dispose of the additional glucose. Failure of the liver to take up glucose predisposes an individual to develop hyperglycemia. In individuals with diabetes (19) who are prone to infections, the severe hyperglycemia can limit the calories that are given.
The impaired liver glucose uptake seen in stressed patients may be improved if glucose is given via the oral route. Previous studies indicate that the route of glucose delivery is important in determining the magnitude of net hepatic glucose uptake. Indeed, oral administration of glucose enhances net hepatic glucose uptake to a greater extent than when the glucose is administered via a peripheral vein (23). If the oral delivery of glucose can overcome the infection-induced impairment in liver glucose uptake, glucose carbon can be diverted to the liver and away from the insulin-resistant tissues in the stressed individual. This, in turn, could limit the hyperglycemia and the insulin required to maintain normoglycemia. Thus the aim of this study was to determine whether the infection-induced impairment in net hepatic glucose uptake seen after peripheral infusion of glucose could be overcome by intraduodenal administration of glucose.
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METHODS |
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Animal Preparation
Experiments were carried out on 14 conscious female mongrel dogs (21 ± 1 kg). Before being studied, they received a diet consisting of Kal-Kan meat (Vernon, CA) and Purina dog chow (St. Louis, MO) once daily. The composition of the diet was 52% carbohydrate, 31% protein, 11% fat, and 6% fiber, based on dry weight. The dogs were housed in a facility that met the guidelines of the Association for Assessment and Accreditation of Laboratory Animal Care International. The Vanderbilt University Animal Care and Use Committee approved the experimental protocols.Surgical Preparation
Fourteen to seventeen days before study, a laparotomy was performed under general anesthesia (isoflurane). Sampling catheters (0.04 inch ID) were inserted into the portal vein and the left common hepatic vein for blood sampling. An infusion catheter was placed into the duodenum. Additional catheters (0.04 inch ID) for blood sampling were inserted into the femoral artery after an incision in the left inguinal area. The catheters were then filled with saline containing heparin (200 U/ml). Doppler flow probes were placed around the portal vein and the hepatic artery after the gastroduodenal vein had been ligated. The portal and hepatic vein sampling catheters, the intraduodenal infusion catheter, and the Doppler flow probe leads were exteriorized and placed in a subcutaneous pocket in the abdominal area. The femoral artery sampling catheter was placed under the skin in the inguinal region (18).Two weeks after catheter implantation, all animals had
1) a good appetite (consuming the entire daily ration),
2) a normal stool, 3) a hematocrit >35%, and
4) a leukocyte count <18,000 mm3.
Induction of Infection
The model used is nonlethal (15) and is similar to the model of Fink et al. (7). Approximately two weeks after catheter implantation, a fibrin clot was prepared from a 1% bovine fibrinogen solution (10 ml/kg; Sigma Chemical, St. Louis, MO), which was then filtered through a sterile 0.2-µm filter. On the day before clot implantation, 1 liter of trypticase soy broth (Becton Dickinson, Cockeysville, MD) was inoculated with bacteria and incubated overnight at 37°C. The next day, the bacteria were pelleted by centrifugation, washed with sterile saline, and reconstituted in 20 ml of sterile saline. The dose of bacteria (Escherichia coli; American Type Tissue Culture #25922) was 2 × 109 organisms/kg. The concentration of bacteria was determined by serial dilution of the bacteria followed by plating. The bacteria were mixed with the fibrinogen, and thrombin (1,000 U) was added to initiate clot formation. The sham group received a fibrin clot that did not contain bacteria. After an overnight fast (18 h), dogs were placed under general anesthesia. An abdominal midline laparotomy incision was made at a point below that made 2 wk earlier, and the fibrin clot was placed into the peritoneal cavity. Dogs received 500 ml of saline immediately after clot implantation. An additional 1,000 ml were given the next morning. The dogs were fasted after implantation of the clot.Experimental Protocol
Thirty-six hours after implantation of the clot, the sampling and infusion catheters and the free ends of the transonic flow probes were removed from the subcutaneous pockets under local anesthesia (2% lidocaine). The dog was then placed in a Pavlov harness, and angiocaths were inserted percutaneously into the right and left cephalic veins. AtProcessing of Blood Samples
Blood samples were drawn into heparinized syringes and transferred to chilled tubes containing potassium EDTA (15 mg). The collection and immediate processing of blood samples has been previously described (15). 14CO2 in blood was assessed in triplicate by acidifying blood with HCl and trapping the 14CO2 on filter paper saturated with hyamine hydroxide (8). Blood glucose, lactate, glycerol, and alanine were analyzed according to the method of Lloyd et al. (13) on a Monarch 2000 centrifugal analyzer (Lexington, MA). Plasma glucose was assayed immediately with a Beckman Glucose Analyzer II (Beckman Instruments, Fullerton, CA). Immunoreactive plasma insulin (27) was assayed by means of a double antibody technique [Pharmacia Diagnostics, Piscataway, NJ; intra-assay coefficient of variation (CV) of 11%]. Plasma (1 ml) treated with 500 kallikrein inhibitor units of Trasylol (Miles, Kankakee, IL) was assayed for immunoreactive glucagon (1) with a similar procedure as for insulin (intra-assay CV of 8%). Plasma cortisol (5) was assayed with a Clinical Assays Gamma Coat RIA kit (intra-assay CV of 6%). Plasma collected from blood samples that had been immediately treated with EGTA and glutathione were assayed for epinephrine and norepinephrine with HPLC [CV of 14%;(14)]. Hepatic glycogen was assayed according to the method of Chan and Exton (3).Calculations
Net hepatic glucose uptake was calculated by use of the formula [(Fa × A) + (Fp × P)The rates of total glucose appearance (Ra) and clearance
rates and [14C]glucose appearance rate were calculated
according to the method of Wall et al. (26a), as simplified by Debodo
et al. (4a). Endogenous glucose production was calculated as the
difference between Ra and meal-derived glucose appearance
rate. The meal-derived glucose appearance rate was calculated as the
total [14C]glucose appearance rate (dpm · kg1 · min
1) divided by the meal
[14C]glucose specific activity (dpm/mg), where dpm is
disintegrations per minute. Hepatic glucose oxidation was calculated
during the last hour of the experimental period as the ratio of the net
hepatic 14CO2 output and the average inflowing
[14C]glucose specific activity. Tracer-determined
glycogen synthesis (mg · kg
1 · min
1) was calculated as the ratio of hepatic
[14C]glycogen content [(14C dpm/g
liver × liver wt)/(body wt × 240 min)] and the
average inflowing [14C]glucose specific activity
(dpm/mg).
Intestinal substrate output was calculated as (P A) × Fp × HBF. Because the intestine can be both a
consumer and a producer of glucose during intraduodenal glucose
delivery, we calculated the unidirectional absorption of glucose (Gut
Ra) as the sum of meal-derived gut glucose output and
unidirectional gut glucose uptake. Because the absorbed glucose
contained [14C]glucose, the meal-derived gut glucose
output was calculated as the ratio of net gut
[14C]glucose output and meal glucose specific activity.
Unidirectional gut glucose uptake was calculated as the ratio of net
gut [3H]glucose uptake and arterial glucose specific
activity. If the site of glucose uptake were common with or distal to
the site of glucose absorption, the use of inflowing glucose specific
activity would underestimate gut glucose output and, consequently,
absorption. In the case of glucose, the fall in
[3H]glucose specific activity in the portal vein was
small, and therefore, the potential error is small.
Statistics
Hepatic blood flow and substrate flux are expressed as kilograms of body weight. Data are expressed as means ± SE. Data presented for the experimental period are the average of the last 60 min of the experimental period unless otherwise indicated. Statistical comparisons were made by means of analysis of variance (Systat for Windows; Systat, Evanston, IL). A univariate post hoc F-test was used when a significant F ratio was found. Statistical significance was accepted at P < 0.05. ![]() |
RESULTS |
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Basal Period
Hemodynamic parameters.
Infection increased body temperature, heart rate, and hepatic arterial
flow; mean arterial blood pressure and portal vein blood flow remained
unaltered (Table 1; Fig.
1). Consequently, total hepatic blood
flow increased by ~30% (34.9 ± 4.6 vs. 47.2 ± 3.2 ml · kg1 · min
1, sham vs.
infected, respectively; P < 0.05).
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Metabolic parameters.
Arterial plasma glucose concentrations (Table 1) were similar in the
two groups, whereas whole body glucose appearance (2.1 ± 0.2 vs.
3.3 ± 0.1 mg · kg1 · min
1) and clearance (2.2 ± 0.2 vs. 3.7 ± 0.2 ml · kg
1 · min
1) were
characteristically increased (P < 0.05) by infection.
The liver was a net consumer of lactate (9.5 ± 2.1 vs. 15 ± 2.5 µmol · kg
1 · min
1, sham
vs. infected; P < 0.05) and alanine (2.7 ± 0.4 vs. 3.6 ± 0.7 µmol · kg
1 · min
1) in the basal period. Arterial insulin
concentrations were similar (8.0 ± 1.6 vs. 8.2 ± 1.5 µU/ml; sham vs. infected), whereas arterial plasma glucagon
concentrations were markedly elevated (53 ± 8 vs. 680 ± 207 pg/ml; P < 0.05; Fig.
2). Arterial cortisol concentrations were
also elevated (1.3 ± 0.2 vs. 4.2 ± 0.7 µg/dl;
P < 0.05). Arterial epinephrine concentrations were
not altered (108 ± 30 vs. 124 ± 50 pg/ml), whereas arterial
plasma norepinephrine concentrations were elevated (268 ± 69 vs.
430 ± 89 pg/ml; P < 0.05).
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Response to Intraduodenal Glucose Infusion
Hepatic blood flow and pancreatic hormones. Hepatic arterial and portal vein blood flows were not altered by intraduodenal glucose infusion (Fig. 1). Arterial plasma insulin concentration increased to similar concentrations in both groups; however, the initial rise was faster in the sham group (Fig. 2). Arterial plasma glucagon concentrations decreased in both groups (53 ± 8 to 30 ± 2 vs. 680 ± 207 to 269 ± 128 pg/ml; P < 0.05; basal period to average of last 60 min of the experimental period); however, the absolute decrease in glucagon was greater in the infected group (Fig. 2).
Intestinal substrate kinetics.
Although arterial glucose concentrations increased rapidly in both
groups, the initial rise in glucose was faster in the sham group (Fig.
3). This was paralleled by a faster rise
in both net intestinal glucose output and unidirectional intestinal
glucose output (P < 0.05). During the last hour of the
study, we could account for 75 and 55% of the glucose infused in the
sham and infected groups, respectively. The absorption of glucose
created a negative arterial-portal blood glucose gradient of 20 ± 4 vs. 13 ± 3 mg/dl (sham vs. infected) during the last hour of
the study. Thus the portal vein blood glucose concentrations tended to
be lower in the infected group (150 ± 10 vs. 135 ± 7 mg/dl;
Fig. 3) during the last hour of the study.
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Hepatic substrate kinetics.
The liver switched from being a net producer (net hepatic glucose
utpake is negative) to a net consumer of glucose (net hepatic glucose
uptake is positive) in both groups (Fig.
4) during glucose infusion. Net hepatic
glucose fractional extraction and hepatic glucose load were similar in
both groups. Tracer-determined endogenous glucose production decreased
in both groups (1.0 ± 0.2 vs. 1.2 ± 0.2 mg · kg1 · min
1, sham vs. infected).
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DISCUSSION |
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In the present study, infection did not attenuate the increase in net hepatic glucose uptake seen when glucose was administered via the intraduodenal route. On the basis of our previous work, when glucose was infused via the peripheral route, infection decreased net hepatic glucose uptake (17). The normal liver glucose uptake during duodenal glucose infusion is even more surprising, given that infection delayed net intestinal glucose uptake. However, not all aspects of liver metabolism were normalized; the liver of the infected animal remained a net consumer of lactate, whereas the liver of the sham animal was not a consumer of lactate. These studies suggest that, despite infection-induced alterations in intestinal absorption, when glucose is administered via the oral (duodenal) route, the presence of infection does not impair net hepatic glucose uptake.
The infection-induced decrease in intestinal glucose absorption
(~25%) after glucose infusion explained the attenuated rise in
arterial blood glucose concentration in the infected animals. The
decrease in net glucose absorption was caused by an impairment in
unidirectional intestinal glucose output rather than by a stimulation of intestinal glucose utilization. The mechanism for this attenuation is unknown. Intestinal glucose absorption requires a sodium-dependent glucose transporter (SGLT1) (6). It is unclear whether the impairment in glucose absorption represents a specific impairment in
SGLT1, a nonspecific impairment in sodium-coupled transport, or a more
general defect in gut absorption. The latter could be caused by
impairments in mucosal mass or mucosal blood flow or to increases in
intestinal transit time (6). Although intestinal hyperpermeability to large macromolecules is suspected to occur during
systemic infections (22), sepsis can delay the absorption of small molecules such as glucose (12).
Despite the blunted gut glucose absorption, net hepatic glucose uptake was not diminished by infection. The two determinants of liver glucose uptake in the normal setting are hepatic glucose load and net hepatic glucose fractional extraction. Hepatic glucose load is dependent on liver blood flow and glucose concentration. In our previous report (17), liver blood flow was not increased in this model of infection. In the present study, liver blood flow was increased by 30% in the infected group. We do not have an explanation for this difference; however, one possibility is that we used Transonic rather than Doppler flow probes in the present study. The Transonic flow probes are not subject to problems with turbulence and orientation of the probe with respect to the vessel. Regardless of the reason, an underestimation of liver blood flow cannot explain the differences in the two routes of administration. Recalculation of the previous data by use of the mean flow obtained in this study had minimal effects on the calculated net hepatic glucose uptake. This is because the primary contributor to the infection-induced alterations in liver glucose uptake during peripheral glucose infusion (17) was a marked decrease in net hepatic glucose fractional extraction.
Infection did not impair net hepatic glucose uptake when glucose was given intraduodenally, because net hepatic glucose fractional extraction was not altered. In the previous study, hepatic glucose uptake was decreased when glucose was given into a peripheral vein, because net hepatic glucose fractional extraction was markedly decreased by infection (17). The mechanism whereby duodenal glucose delivery overcame the defect in net hepatic glucose fractional extraction is unclear. It is well known that oral delivery of glucose enhances net hepatic glucose fractional extraction, and consequently net hepatic glucose uptake, to a greater extent than when the glucose is given into a peripheral vein (23). The enhancement of liver glucose uptake seen with the oral route does not require the gut; bypassing the gut by infusing the glucose directly into the portal vein mimics the response. The ability of portal delivery of glucose to augment liver glucose uptake has been termed the "portal signal" (23). Activation of the portal signal may be capable of overriding the infection-induced impairment in liver uptake and fractional extraction of glucose by the liver. Thus, although the mechanism by which the portal signal enhances liver glucose uptake is unknown, it is clear that some aspect of duodenal glucose delivery (possibly the portal signal) can play a dominant role in facilitating liver glucose uptake during infection.
The effective suppression of hepatic glucose production seen during intraduodenal glucose infusion contributed to the improved hepatic glucose disposal during infection. During a peripheral glucose infusion, the lack of suppression of hepatic glucose production was as important as the lack of stimulation of unidirectional hepatic glucose uptake in limiting net hepatic glucose uptake during infection (17). The inability of a peripheral infusion of exogenous glucose to suppress endogenous glucose production has been observed in stressed humans as well (26). It is unclear how intraduodenal infusion of glucose is more effective than peripheral glucose infusion in suppressing hepatic glucose production during infection.
Although hepatic glucose uptake was not altered by infection, the liver of the infected animal remained a net lactate consumer. In a normal animal, when the liver is a net consumer of glucose, the liver is either releasing lactate or at least is not consuming lactate, indicating that glycolysis is increased and gluconeogenesis from lactate is suppressed. However, during infection, the liver exhibited net lactate consumption, which may reflect a persistent stimulation of gluconeogenesis. In noninfected animals, the switch to lactate production is thought to reflect diversion of a portion of the glucose carbons to glycolysis. Chronic exposure to high cortisol concentrations amplifies glycolysis in the liver, but this can be reversed by excess glucagon (16). Thus hyperglucagonemia, a characteristic response to infection, may explain the maintenance of substantial lactate consumption in the face of normal hepatic glucose uptake.
Tracer-determined hepatic glycogen synthesis was also normal after intraduodenal glucose delivery. After peripheral glucose delivery, tracer-determined glycogen synthesis was decreased, primarily because of a decrease in net hepatic glucose uptake (17). Because liver glucose uptake was not impaired during duodenal glucose delivery, and because glycogen synthesis is a major fate for liver glucose uptake, it is not surprising that glycogen synthesis was normal. Some aspect of duodenal glucose delivery, possibly the portal signal, may have helped facilitate hepatic glycogen synthesis. Diversion of gluconeogenic carbon to glycogen (i.e., indirect glycogen synthesis) can amplify glycogen deposition. Indirect glycogen synthesis is significant even in the normal dog (20), and the tracer method we used does not distinguish between direct and indirect glycogen synthesis. Approximately 60% of the net glucose uptake could be accounted for by tracer-determined glycogen deposition in both groups. In the infected group, the liver remained a net hepatic lactate consumer; thus it is likely that indirect hepatic glycogen synthesis was amplified. Although hepatic glycogen content was greater in the sham group after glucose infusion, this does not mean that net glycogen synthesis was higher in the sham group. Without knowing the glycogen content before glucose infusion, net hepatic glycogen synthesis could not be calculated. However, it is likely that the hepatic glycogen content was lower in the infected group, which may have facilitated hepatic glycogen synthesis (9). Although previous work has demonstrated that glycogen synthesis is impaired by infection during peripheral glucose infusion (4), we found no evidence that it is impaired when glucose is given intraduodenally (2). The mechanism by which the portal signal activates hepatic glucose entry is unknown. It requires an intact hepatic neural supply, and it facilitates hepatic glycogen synthesis and lactate release in the normal animal (23). Thus it likely acts at an early point in the metabolic pathway such as activation of glucokinase (23). Generation of such a signal may be sufficient to normalize hepatic glucose entry during infection.
For the stressed patient, improving liver glucose uptake by delivering glucose via the enteral route may help limit the hyperglycemia associated with nutrient intake. This is most critical in individuals with preexisting glucose intolerance (19). By enhancing liver glucose uptake, the mass of glucose that has to be removed by the insulin-resistant peripheral tissues will be lessened. In turn, whole body insulin requirements may be lessened as well. Although enteral delivery is the preferred route of nutrient delivery, in part because of its trophic effects on the intestinal mucosal barrier (24, 28), the complications of malabsorption in the stressed patient have been an obstacle to its widespread use. The additional benefit of improved hepatic glucose disposal observed in this study further amplifies the need to overcome the complications associated with enteral feeding to achieve the multiple benefits of enteral nutrient delivery on intestine and hepatic function.
In summary, net hepatic glucose uptake is not decreased by infection when glucose is administered via the gastrointestinal route. This occurs despite delayed absorption of glucose by the gut during infection. The infection-induced enhancement in net hepatic lactate uptake persists, however, suggesting that oral glucose delivery cannot override the characteristic augmentation of gluconeogenesis.
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ACKNOWLEDGEMENTS |
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The authors are grateful for the technical assistance of Pamela Venson and Eric Allen from the hormone core laboratory of the Vanderbilt University Diabetes Research and Training Center.
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FOOTNOTES |
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This study was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-43748 (O. P. McGuinness), with support from the Clinical Nutrition Research Center (DK-26657) and Diabetes Research and Training Center Grant P60-DK-20593.
Address for reprint requests and other correspondence: O. P. McGuinness, Dept. of Molecular Physiology and Biophysics, 702 Light Hall, Vanderbilt University, Nashville, TN 37232-0615 (E-mail: owen.mcguinness{at}mcmail.vanderbilt.edu).
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.
Received 29 November 1999; accepted in final form 2 February 2000.
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