INTRODUCTION

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THE VISCERAL TISSUES, especially the intestinal mucosa, pancreas, and spleen, have high rates of metabolism. As a consequence, the contribution of the portal-drained viscera (PDV) to whole body energy expenditure is much larger than their contribution to body weight. Understanding the substrates that are used to support this high oxidative activity is therefore of considerable physiological and nutritional importance.

The most extensive information on intestinal carbon metabolism in vivo is contained in a series of publications based on the use of isolated, in situ, vascularly perfused loops of rat small intestine (26-31). The authors (31) concluded that, in animals that had been fed up to the initiation of the tracer infusions, aspartate, glutamate, and glutamine were the major oxidative substrates used by the small intestinal mucosa and that glucose was a minor oxidative substrate. The observations of Windmueller and Spaeth (26-31) with regard to mucosal glutamine metabolism have received the greatest attention and have served as the starting point for many investigations of the effects of glutamine on intestinal growth, metabolism, and function (e.g., Ref. 3). However, their observation (28, 31) that the metabolism of enteral glutamic acid was of much more quantitative importance to mucosal energy generation than that of glutamine has been largely overlooked. With the notable exception of their observations on glutamate, studies in vitro (7, 25, 32) have generally confirmed the findings of Windmueller and Spaeth, although some recent results (8) imply that the relative utilization of different substrates by isolated enterocytes may be a function of their relative concentrations in the extracellular phase.

This last observation is important, because in the fed state, the intestinal mucosa are presented with a complex mixture of arterial and luminal substrates, and it is not known with certainty whether, under fed conditions, the mucosal cells utilize some substrates in preference to others. In other words, it is not known whether Windmueller and Spaeth's observations reflected the nature of the experimental preparation or whether the results revealed a specific feature of mucosal intermediary metabolism. The main objective of the present work was to attempt to provide an answer to this question.

We have developed a portal catheterized piglet model (6) to study visceral metabolism in growing, conscious, and fed animals. In past studies, we have used this model, in combination with enteral infusions of uniformly ¹³C-labeled tracers, to quantify the first-pass utilization of dietary amino acids (16, 17, 21-23). The results of one of these studies (17) confirmed previous observations of virtually complete first-pass utilization of dietary glutamate (14, 28, 31) and aspartate (29) by the intestinal tissues. However, our studies were confined to the measurement of portal tracer balance and mucosal protein synthesis, and although we inferred that a high proportion of glutamate and glutamine utilization was directed to oxidation, no direct measurements were made.

The present paper reports the results of a series of experiments in which we have measured the oxidative metabolism of U-¹³C-labeled glucose, glutamate, and glutamine by the PDV. The main objective was to quantify their relative utilization in the synthesis of alanine and lactate and their contribution to CO₂ production. On the basis of previous results (28, 31) and of our own (17, 18), we hypothesized that enteral glutamic acid (metabolized in first pass by the mucosa) would be the single largest contributor to PDV oxidative metabolism.

	METHODS

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The protocol received prior approval by the Animal Care and Use Committee of Baylor College of Medicine. Animal care conformed to current US Department of Agriculture guidelines.

Feeding and Surgery

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Tracer Protocol

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At the time of the tracer infusion, the animals weighed 7.6 ± 0.6 kg. After an overnight fast, baseline samples of portal and arterial blood were obtained, and the animals were offered a single meal of one-twelfth of their preceding daily intake. This meal served to initiate an adequate pattern of gastric emptying. One hour later, a constant intragastric infusion of liquid Litter Life was commenced and continued for the next 6 h. The diet infusion provided nutrients at the same daily rate as the animals had consumed over the preceding 3 days. In the animals that received an enteral [U-¹³C]glutamate infusion, the tracer was mixed with the diet, whereas enteral [U-¹³C]glucose was given intraduodenally. The intravenous and intraduodenal infusions were started at the same time as the diet infusion. Animals were killed with an intra-arterial injection of pentobarbital sodium (50 mg/kg body wt) and sodium phenytoin (5 mg/kg; Beutanasia-D; Schering-Plough Animal Health, Kenilworth, NJ) after 6 h of tracer infusion.

During the infusion, arterial and portal blood samples (3 ml) were taken every hour for 5 h and then at 15-min intervals over the last hour. Blood gases (Chiron Diagnostics, Halstead, Essex, UK), glucose, and lactate (YSI analyzer, Yellow Springs, OH) were determined immediately in all blood samples. An aliquot of whole blood (0.5 ml) was immediately placed in an evacuated tube (5 ml capacity) for subsequent analysis of the isotopic enrichment of blood bicarbonate, and two aliquots (1 ml) were immediately frozen for subsequent measurements of amino acid, glucose and lactate concentrations, and labeling. The remaining blood was centrifuged (3,000 g, 10 min at 4°C), and the plasma was frozen in liquid nitrogen until taken for the analysis of its ammonia concentration.

Sample Analysis

To estimate the isotopic enrichment of blood bicarbonate, 0.5 ml of perchloric acid (1 mol/l) was injected into the evacuated tube containing the blood sample. The contents of the tube were mixed on a vortex mixer, and the head space was carefully removed with a 5-ml syringe. The gas sample was then injected into a second evacuated tube (15 ml volume). The isotopic enrichment of the CO₂ was then determined on a continuous flow gas isotope ratio machine (ANCA, Europa Instruments, Crewe, UK). The between-sample standard deviation was 0.001 atoms percent excess (1).

Calculations

Portal balance.

(1)

in which concentration is expressed as micromoles per liter, and portal blood flow is expressed as liter per kilogram per hour.

(2)

in which t/T is the tracer-to-tracee ratio of the U-¹³C-labeled substrate (glucose, glutamate, or glutamine) or the product (alanine or lactate). Throughout this article, a positive portal balance signifies the addition of a compound to the portal blood, and a negative balance signifies the removal of the compound from the arterial input.

(3)

(4)

For the enteral tracers, input in Eq. 3 is the rate of tracer infusion, and for the intravenous tracers, input is the arterial tracer flux, i.e., (concn_ART × t/T_ART) × portal blood flow.

End product synthesis. The contribution of each substrate to total lactate, alanine, or CO₂ production was calculated as follows. The portal balance of the ¹³C₃ isotopomers of lactate of alanine was calculated with Eq. 2 and expressed as a fraction of the input of the respective U-¹³C tracers. For enteral glutamate and glucose, which we assume were completely removed from the intestinal lumen, the denominator was the rate of tracer infusion. For intravenous glucose and glutamine, the input was the portal tracer balance. On the assumption that tracer and tracee are metabolized identically, total alanine or lactate production was calculated as

(5)

Once again, for enteral glutamate and glucose, the input was the dietary intake, whereas for intravenous glucose or glutamine, the input was the estimated uptake from the arterial supply. It is important to note that in calculating the total production of lactate or alanine from glucose, each molecule of glucose yields two molecules of lactate or alanine.

First-pass metabolism of enteral glucose. The calculation of the metabolism of enterally administered [U-¹³C]glucose to lactate, alanine, and CO₂ is complicated by the fact that the large majority of the tracer is absorbed and thereby labels arterial glucose. A portion of this labeled arterial glucose is extracted and metabolized by the PDV and contributes to the production of glycolytic end products and ¹³CO₂. This consideration does not apply to glutamate and glutamine metabolism because 1) very little of the enteral glutamate tracer is absorbed (16) and 2) metabolism of glutamine occurs essentially only from the arterial input. However, the fractional extraction of the intravenously infused [U-¹³C]glucose by the PDV is known (from Eq. 3), so that the first-pass metabolism of enteral [U-¹³C]glucose can be calculated from the product of the arterial flux of [U-¹³C]glucose during the enteral glucose infusion and the fractional extraction of arterial [U-¹³C]glucose during the intravenous infusion.

(6)

(7)

Absorption of dietary glutamine. Although in our past studies, and in this study, the portal mass balance of glutamine is negative, (i.e., there is net removal of arterial glutamine by the PDV), there is the possibility that there is simultaneous absorption of dietary glutamine into the portal vein and extraction of arterial glutamine. The absorption of dietary glutamine can be estimated from the difference between the portal glutamine fractional mass and fractional tracer balances. Thus

(8)

in which arterial glutamine flux is concn_ART × portal blood flow.

	RESULTS

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The body weights and rates of substrate intake, tracer infusion, portal blood flow, and total CO₂ production by the PDV are shown in Table 1. There were no significant differences among the mean weights of the animals subjected to different tracer infusions. Over the postoperative period (6 ± 1 day), the daily weight gain (293 ± 49 g/day; 38 ± 6 g · kg¹ · day¹) was also not different among the groups. The absolute values for the portal blood flow (22.3 ± 1.4 l/h), total CO₂ production (38 ± 2 mmol/h), and O₂ consumption (28 ± 2 mmol/h) were also similar, and consequently there was a negative relationship between body weight and weight-specific portal blood flow, CO₂ production, and O₂ consumption. In unpublished studies (T. A. Davis, D. Wray-Cahen, P. R. Beckett, and P. J. Reeds) with 4-wk-old piglets receiving nutrients at a rate similar to that in the present study, we found that whole body CO₂ production is ~30 mmol CO₂ · kg¹ · h¹. Thus the observed CO₂ production by the PDV (4.92 ± 0.97 mmol · kg¹ · h¹) accounted for 16% of total body CO₂ production. The respiratory quotient across the PDV was 1.38 ± 0.11, a value that was significantly (P < 0.001) higher than unity.

Mass and Tracer Balance of Primary Substrates

The portal balance of the U-¹³C tracers is shown in Table 3. There was a small but significant (3.7% of dose; P < 0.05) net absorption of [U-¹³C]glutamate; 93.6% of the enteral dose of [U-¹³C]glucose appeared in the portal vein. The PDV extracted 20.4% of the arterial [U-¹³C]glutamine flux and 6.5% of the arterial flux of intravenously infused [U-¹³C]glucose. The glutamine fractional tracer balance was significantly (P < 0.001) greater than its fractional mass balance, and in molar terms, the difference was 72 ± 10 µmol · kg¹ · h¹. This presumably represented absorption of dietary glutamine, so it appeared that ~50% of the dietary glutamine was absorbed intact.

Lactate and Alanine Production

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The proportion of the input of any given substrate metabolized to alanine and lactate was quite low (2.9-6.6% of dose). However, because the quantities of each substrate presented to the PDV were markedly different (ranging from 191 µmol · kg¹ · h¹ for arterial glutamine to 3,406 µmol · kg¹ · h¹ for dietary glucose), the contribution of each substrate to total alanine and lactate production (Table 6) varied widely. Even after adjustment for the secondary metabolism of absorbed glucose, the first-pass metabolism of enteral glucose contributed 18% of the portal alanine balance and 31% of the balance of lactate. A further 22% of total alanine production and 14% of the lactate production derived from arterial glucose metabolism. Thus glucose metabolism accounted for 40% of the portal alanine carbon and 53% of the portal lactate carbon.

CO₂ Production

Estimated contributions of the substrates to total CO₂ production by the PDV are shown in Table 8. The most important sources were enteral glutamate (36% of total) and arterial glucose (29% of total). Arterial glutamine (15%) made a smaller contribution, and the contribution of enteral glucose (6% of CO₂ production) was minimal. The oxidation of the four substrates accounted for 76% of total CO₂ production by the PDV.

Other Nitrogenous Products

	DISCUSSION

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Largely as a result of the pioneering work of Windmueller and Spaeth (summarized in Ref. 26), considerable attention, particularly in the clinical literature (3), has been paid to the role of glutamine in the small intestinal mucosa, and it is often stated that glutamine is the "major" energy source for these cells. In vitro studies (24, 25) appear to confirm this comment, because isolated enterocytes transport glutamine at a rate that is about fourfold higher than that of glutamate. In vivo, however, the same considerations may not apply. First, despite the fact that isolated enterocytes will readily metabolize glucose (8), a number of studies in adult humans (e.g., Ref. 13) have shown that the splanchnic extraction of oral or intraduodenal glucose is minimal. Second, other studies in humans (1, 14) have shown that the splanchnic extraction of enteral glutamate exceeds that of glutamine. Third, it has been repeatedly demonstrated (16-18) that in fed animals, even though the removal of lumenal glutamate (both free and protein-bound) is efficient, little dietary glutamate appears in the portal circulation. Finally, Windmueller and Spaeth (28) explicitly commented on the fact that lumenal glutamate appeared to be a more important oxidative substrate than glutamine. The present data, obtained with conscious animals receiving a mixed high-protein diet, confirm the large majority of Windmueller and Spaeth's results and, in particular, their assertion about the importance of glutamate to the energy economy of the small intestinal mucosa. The fact that such similar results have been obtained in different species, at different stages of development, and under quite different experimental circumstances, strongly suggests that the results reflect a specific feature of mucosal metabolism, i.e., a critical role of amino acids as energy sources.

In drawing conclusions about the broader significance of the results, there are two points that require discussion. The first concerns the outflow of nitrogenous substances in the portal vein. Although we have not directly quantified its metabolism, there was no release of aspartate to the portal circulation, so that it is probable that dietary aspartate is catabolized to the same extent as glutamate and glutamine (see also Refs. 29, 31). About 10% of aspartate nitrogen metabolism would have been directed to arginine biosynthesis, thereby generating fumarate, but the majority of the aspartate would been metabolized via transamination with -ketoglutarate to generate cytoplasmic oxaloacetate. That being so, we estimate that the exit of nitrogen in newly synthesized alanine (338 µmol · kg¹ · h¹) accounted for only 47% of the nitrogen generated from the estimated catabolism of glutamate, aspartate, and glutamine (715 µmol · kg¹ · h¹). The remainder apparently exited as ammonia, which in the present study was produced by the PDV at a rate of 968 µmol · kg¹ · h¹. Critically, the rate of ammonia production not only exceeded glutamine deamidation (190 µmol · kg¹ · h¹) by a factor of five, but the sum of alanine synthesis and ammonia production exceeded the quantity of nitrogen that would have been generated from aspartate, glutamate, and glutamine catabolism. These observations are particularly important because they imply that, in vivo and in the fed state, 1) there is a metabolically significant glutamate deamination (i.e., an active glutamate dehydrogenase) in the mucosa and 2) other dietary amino acids are catabolized in first pass by the mucosa.

On the basis of the results of Windmueller and Spaeth (28, 31) and from results with isolated enterocytes (25), it is generally assumed (see Ref. 11) that glutamate dehydrogenase is either absent from the mucosa or present in negligible quantities. However, it is possible to question this assumption. First, in at least one detailed study of the metabolism of isolated enterocytes (25), ~35% of cellular glutamate uptake appeared as ammonia. Second, direct measurements in both porcine (4) and rodent small intestinal mucosa (5, 9, 19) have demonstrated the presence of low, but significant, amounts of glutamic dehydrogenase. We find it noteworthy that if the portal ammonia production that we observed is expressed per unit mucosa, then the value [1.3 µmol · g (mucosal weight)¹ · min¹] is remarkably similar to that measured directly (1.5 µmol · g¹ · min¹) in mucosa isolated from porcine jejunum (4).

The second observation that requires comment is the difference in the distribution of the metabolism of glucose, glutamate, and glutamine in the production of lactate and alanine ("incomplete" oxidation; Ref. 24) compared with that of CO₂ (complete oxidation). With glucose, the majority of the pyruvate was converted to lactate and alanine and exported to the portal circulation, whereas with glutamate or glutamine, only a minor portion followed the same pathway. Windmueller and Spaeth (31) made similar observations.

In considering this observation, it is important to emphasize that, for an amino acid that enters intermediary metabolism directly via the tricarboxylic acid cycle to be completely oxidized (20), it must be metabolized via acetyl-CoA and hence via pyruvate. Thus in this respect pyruvate is a common product of glucose, glutamate, and glutamine metabolism. However, because the pyruvate pool derived from glutamate and glutamine metabolism appears to be channeled to oxidation, whereas that derived from glucose is channeled to lactate and alanine formation, it seems reasonable to argue that the pyruvate derived from glutamate and glutamine metabolism is generated in the mitochondrion.

This proposition, however, begs the question of the enzymes responsible. The first possibility is via phosphoenolpyruvate carboxykinase. This leads to the cytoplasmic generation of phosphoenolpyruvate. However, although phosphoenolpyruvate carboxykinase is active in the enterocyte, alanine production from glutamine is not blocked by inhibition of phosphoenolpyruvate carboxykinase (26). The second possibility is via the decarboxylation of malate by NADP-linked malic enzyme, an enzyme also present in the cytoplasm of enterocytes (25). This would also lead to the generation of cytoplasmic pyruvate, which, in the absence of further cytoplasmic compartmentation, should follow the pathway of pyruvate generated from glycolysis. The third, and intriguing, possibility is that the synthesis of pyruvate from the tricarboxylic acid cycle intermediates generated from glutamate and glutamine occurs within the mitochondrion and that the reaction is catalyzed by a malic enzyme (20), termed NAD(P)⁺-malic enzyme by Sauer and colleagues (15, 20). This enzyme differs from the cytoplasmic malic enzyme by having a mixed specificity for NAD and NADP and, critically, is expressed only in rapidly proliferating cells, including those of the intestinal mucosa (15). Although speculative, this proposition is, in our opinion, the most compatible with our data.

As we have pointed out elsewhere (21-23), our observations regarding amino acid metabolism by the intestinal tissues have substantial nutritional implications. First, if as we believe, dietary amino acids are the major source of mucosal energy generation, then other factors that affect mucosal mass, such as parasitic infestation and dietary toxins, could have a substantial effect on the systemic availability of dietary amino acids. Second, the results imply that a considerable portion of the splanchnic metabolism of dietary amino acids occurs in the intestine rather than in the liver. Leucine catabolism by the canine intestine has been observed (34). However, whether other essential amino acids are catabolized in the mucosa is a controversial point, for no other reason than the fact that there is little direct evidence for the presence of the appropriate enzymes in the intestinal mucosa. Finally, we should emphasize that the animals that we studied had habitually received very high protein intakes, and our observations could reflect metabolic adaptation to this dietary regimen. In view of this, at least two further questions now need to be investigated. First, do the mucosa catabolize and oxidize dietary essential amino acids in vivo? Second, is mucosal amino acid metabolism sensitive to the level of protein in the diet?