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Nutrient Metabolism

Net Interorgan Transport of L-Glutamate in Rats Occurs via the Plasma, Not via Erythrocytes¹

Department of Nutritional Sciences, Thompson Hall, Cook College, Rutgers, The State University, New Brunswick, NJ 08901

²To whom correspondence should be addressed. E-mail: Watford{at}AESOP.Rutgers.edu.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

 
Glutamate is concentrated within RBC, but this intracellular glutamate is often ignored in studies of glutamate metabolism in vivo. The objective of this work was to determine the size of the plasma and cellular glutamate pools in rat blood and to clarify the role of RBC in the interorgan transport of glutamate. Approximately 20% of whole-blood glutamate was associated with isolated RBC membranes, but this was easily removed by washing with high salt solutions. Arterial plasma glutamate levels were relatively stable and did not show marked differences with starvation, streptozotocin diabetes or feeding 60% casein diets. In rats fed 5% casein, the plasma glutamate level was slightly higher (P < 0.05) than in most other groups. In contrast, RBC glutamate levels showed considerable variation. In rats consuming 5% casein, cellular glutamate levels were 100% higher (P < 0.05) than in control, starved, diabetic or 20 or 60% casein-fed rats. Cellular glutamate levels were also higher (P < 0.05) in rats fed 60% casein than in those consuming 20% casein or the control diet. Rat erythrocytes in vitro did not take up or release free glutamate, confirming that they do not possess a glutamate transporter. Arteriovenous difference measurements across the portal drained viscera indicated a net glutamate release into the portal vein in control, 60% casein-fed and diabetic rats. In all cases, the net change in blood glutamate across the tissue occurred via the plasma, with no change in cellular glutamate levels. Therefore analyses of glutamate metabolism in rats in vivo may be made confidently using measurements of either whole-blood or plasma glutamate concentrations.

KEY WORDS: • glutamate • amino acids • blood • plasma • erythrocytes • rats

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

 
Glutamate can account for up to 20% of dietary protein but it is generally considered that these large amounts of dietary glutamate are metabolized effectively during absorption in the intestinal mucosa and little or no glutamate enters the portal circulation under normal conditions (1 ,2 ). Similarly, although involved in many reactions of intermediary metabolism, glutamate metabolism is predominantly intracellular and glutamate is considered to play a minor role in interorgan nitrogen transport (3 ). However, most studies of interorgan glutamate flux and whole-body turnover have been based on measurements of plasma glutamate, ignoring the potential role of RBC.

Glutamate is concentrated within RBC (4 –7 ) and there have been reports that the plasma and intracellular pools can redistribute as the cells pass through tissue capillary beds in vivo (4 ,5 ,8 ,9 ). Although erythrocytes from some species, e.g., dogs and frogs (10 ,11 ), do express a glutamate transporter, there is no evidence for any glutamate transport in cells from rats and humans (12 –16 ). Indeed, it is reported to be very difficult to change intracellular glutamate pools in rat erythrocytes in vitro, and the absence of any demonstrable glutamate transport has provided support for those studies in vivo based on plasma glutamate, in which intracellular glutamate was ignored. However, studies in vitro cannot address the question of glutamate exchange as the cell is deformed by passage through capillaries. Similarly, even if the cells do not show glutamate uptake, they still may be able to release glutamate formed intracellularly from the metabolism of other amino acids.

A number of recent reports have suggested that the size of the intracellular glutamate pool is subject to regulation and that insulin, and possibly insulin-like growth factor (IGF-1), may act to move glutamate out of the blood cells and into tissues in vivo (17 ,18 ). Other studies have reported that as much as 15% of RBC glutamate may be adsorbed on the cell membrane, which raises the possibility that this, not intracellular, glutamate may exchange as the cell passes through a capillary bed (19 –21 ). The aims of this work were to carefully determine the size of the glutamate pools in rat blood and to clarify the role of RBC in the interorgan transport of glutamate. The results confirm that rat RBC are not capable of transporting glutamate and that net exchange of glutamate across tissues occurs via the plasma compartment. However, the size of the intracellular glutamate pool is subject to considerable variation even though the plasma pool remains relatively stable.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

 
Animals.

Male Sprague-Dawley rats (200–300 g) were obtained from Taconic Farms (Germantown, NY). Rats were maintained in a room at 22°C with lighting from 0700 to 1900 h and all procedures were approved by the Rutgers University Institutional Review Board for the Use and Care of Animals. Routinely, rats had free access to a standard diet (LabDiet 5012, 23% crude protein, 6% crude fiber, 4.5% fat) and water. Rats were made diabetic by injection of streptozotocin (60 mg/kg body) directly into a tail vein. Diabetic rats had free access to the standard diet and drinking water and were used 6–8 d after the streptozotocin injection. For some experiments, rats were starved by removal of the food for 48 h with drinking water available at all times. The effect of dietary protein level was assessed using rats that had free access for 8–10 d to pelleted semipurified diets containing 5, 20 or 60 g/100 g casein by weight (Research Diets, New Brunswick, NJ) as previously described (22 ). On the day of experimentation, rats were anesthetized with an intrapertioneal injection of an aqueous solution of sodium pentobarbitol (60 mg/kg body) between 0900 and 1000 h.

Materials.

Radiolabeled L-[2,3-³H]-glutamic acid (888 GBq/mmol) and sucrose [glucose-¹⁴C(U)] (9065 MBq/mmol) were obtained from NEN Life Science Products (Boston, MA). Streptozotocin, glutamate dehydrogenase and NAD were from Sigma (St. Louis, MO).

Distribution of glutamate in blood.

Arterial blood (6–8 mL) was drawn from the aorta into heparinized syringes and immediately mixed with a trace amount of dry heparin to prevent clotting. Samples were taken in capillary tubes for hematocrit determination and a 0.2-mL sample (whole blood) was immediately deproteinized with an equal volume of ice-cold perchloric acid (100 g/L). Four milliliters of the remaining blood was added to 4.0 mL Saponin (10 g/L) solution and placed on ice to lyse the cells (20 ). Fifty microliters of ¹⁴C sucrose (1 kBq/mL), a marker of extracellular space, was added to the remaining blood which was then centrifuged at 1000 x g for 10 min to separate plasma and cells. A 0.5-mL sample of plasma (plasma) and a 0.2-mL sample of packed cells (packed cells) were taken and deproteinized with equal volumes of perchloric acid (100 g/L). The buffy coat was removed and the remaining cells were washed three times with ice-cold saline (9 g/L NaCl) and the resultant pellet suspended in 1.0 mL saline and deproteinized with 1.0 mL perchloric acid (100 g/L) (washed cells). Samples were taken for the determination of ¹⁴C at all steps of the procedure. The saponin-treated sample was centrifuged at 15,000 x g for 30 min to isolate membranes (20 ,21 ). A sample (0.5 mL) of the supernatant (lysed supernatant) was taken and deproteinized with 0.5 mL perchloric acid (100 g/L). The remaining supernatant was discarded and the pellet resuspended in 2.0 mL 5mmol/L TrisHCl, pH 7.4. A sample (0.5 mL) of the resuspended membranes (membranes) was deproteinized with an equal volume of perchloric acid (100 g/L) and the remainder centrifuged to reisolate the membranes. These membranes were then washed three times with 5 mmol/L TrisHCl, pH 7.4, 154 mmol/L NaCl or 1 mol/L NaHCO₃. The final pellet was resuspended in 0.5 mL of 5 mmol/L TrisHCl, pH 7.4, and deproteinized with an equal volume of perchloric acid (100 g/L) (washed membranes).

Uptake and release of glutamate by red blood cells in vitro.

RBC were isolated from heparinized arterial blood obtained from fed rats. The cells were washed three times with saline (9 g/L) and resuspended in saline to give a hematocrit between 0.15 and 0.30. Aliquots (0.1 mL) of resuspended cells were added to 0.2 mL of a mixture containing ³H glutamate (0.5 kBq), ¹⁴C sucrose (0.2 kBq), and unlabeled glutamate to give final glutamate concentrations from 3 to 24 mmol/L. All incubation mixtures were made isotonic with NaCl. At timed intervals (30 s to 180 min) 1.0 mL stop mix (405 mmol/L NaCl, 15 mmol/L KCl, 45 mmol/L MOPS, pH 7.5, 6 mmol/L MgCl₂, 20 mmol/L glucose) (12 ) was added and the mixture rapidly centrifuged. The supernatant was removed and the pellet resuspended in 0.5 mL Triton X 100 (50 g/L). Samples of the supernatant were deproteinized with an equal volume of perchloric acid (100 g/L) and counted for radioactivity. Tritiated glutamate uptake into the cells was calculated from the ³H content corrected for extracellular water as indicated by the ¹⁴C content.

Glutamate efflux was studied in resealed RBC ghosts (10 ). Washed blood cells were lysed in 40 volumes of hypotonic buffer (5 mmol/L potassium phosphate/5 mmol/L Tris pH 7.4, 1 mmol/L MgCl₂, 0.8 mmol/L phenylmethylsulfonyl fluoride on ice for 10 min. After lysis, the membranes were isolated by centrifugation at 15,000 x g for 20 min. The membranes were resuspended in a volume of hypotonic buffer equal to the original cell volume followed by the addition of 0.5 volumes of reconstitution buffer (750 mmol/L KCl, 100 mmol/L MOPS/Tris, pH 7.4, plus ³H glutamate and ¹⁴C sucrose). After incubation at 37°C for 45 min, the sealed ghosts were isolated by centrifugation at 15,000 x g for 5 min. Ghosts were washed once with normal saline, resuspended to give a hematocrit of 0.30 and incubated with shaking for up to 3 h at 37°C. Samples (0.2 mL) were removed and the ghosts pelleted by centrifugation and the radioactivity determined in the supernatant.

Arteriovenous differences for glutamate.

Blood (2 mL) was simultaneously drawn into heparinized syringes from the aorta and either the portal vein or the left femoral vein. Samples were taken for hematocrit determination. One 0.5-mL sample of whole blood was added to 0.5 mL ice-cold perchloric acid (100 g/L) and the remaining blood was centrifuged at 1000 x g for 10 min to isolate plasma. A sample of plasma was deproteinized with an equal volume of ice-cold perchloric acid (100 g/L).

Metabolite assays.

The supernatants from the percholoric acid extracts were neutralized with KOH. Glutamate and ß-hydroxybutyrate were assayed on the day of experiment by enzymatic spectrophotometric methods as previously described (23 ). Whole-blood glucose was determined using a One Touch glucose meter (LifeScan, Milpitas, CA) immediately after blood was obtained.

Calculations and statistical analysis.

For most experiments, cellular glutamate was calculated from the value for whole blood and plasma with a correction made for the hematocrit (Hct) according to the formula:

Data were evaluated by ANOVA followed by the Student-Newman-Keuls Multiple Comparison Procedure. Plasma and red cell, and arterial and venous glutamate values were compared to zero by t test. All results are reported as means ± SEM with the significance level of differences set at P < 0.05.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

 
No problems were encountered with the different treatment groups. Rats consumed equal amounts of the three casein diets as reported previously (22 ). Diabetic rats (n = 18) had plasma ß hydroxybutyrate levels of 1.21 ± 0.37 mmol/L and blood glucose of 22.2 ± 0.06mmol/L vs. 0.09 ± 0.02 mmol/L and 6.71 ± 0.26 mmol/L, respectively, for control rats (n = 4).

Distribution of glutamate in blood.

The concentration of glutamate was higher in RBC than in plasma (Table 1 ) for all conditions studied. The determination of cellular glutamate based on direct measurement of either glutamate in the packed cell fraction, with correction for extracellular glutamate using ¹⁴C sucrose, or the washed cells gave varying results that were always lower than those calculated from whole-blood and plasma values (results not shown). Because the direct measurement procedures were complex and involved radioactivity, they were not used in subsequent experiments. Isolated cellular membrane fractions contained between 13 and 30% of the total whole-blood glutamate and most of this could be washed off with 5 mmol/L TrisHCl, pH 7.4 (Table 1) . Washing membranes with 154 mmol/L NaCl or 1 mol/L NaCO₃ decreased the glutamate associated with the membranes to below the limit of detection (results not shown). Although the glutamate levels associated with membranes from diabetic rats were greater (P < 0.05) than the other groups, all of this glutamate was removed by the more stringent washing procedures.

Glutamate transport.

Washed RBC incubated with ³H-labeled glutamate took up between 0.1 and 0.5% of the glutamate during the course of a 3-h incubation (results not shown). Similarly, there were no differences in release of ³H-labeled glutamate and ¹⁴C-labeled sucrose from resealed RBC ghosts over a 3-h period (results not shown). Thus, we were unable to demonstrate any inward or outward specific transport of glutamate in rat RBC.

Arteriovenous differences.

The distribution of glutamate has been reported to change as blood passes through tissues in dogs and humans (4 ,5 ,8 ,9 ). This raises the possibility that erythrocytes play a major role in the interorgan transport of glutamate and thus measurements made on plasma may be inaccurate. To determine whether RBC played a role in interorgan glutamate transport, we measured the glutamate exchange across two tissue beds, the leg (sampled at the left femoral vein) and the portal drained viscera (sampled at the portal vein) in the following groups of rats: controls, diabetic, 60% casein-fed and 5% casein-fed. No net glutamate exchange was seen across the femoral vein in any group (results not shown) but in control, diabetic and 60% protein-fed rats, there was a net output of glutamate into the portal vein (Table 3 ). In rats consuming the 5% casein diet, there was no net glutamate exchange across the portal drained viscera (results not shown). In each case in which a net difference was found, this was always due to changes in plasma glutamate. When the contribution of the plasma in 1 L of whole blood was calculated, this was approximately equal to the output as measured for 1 L of whole blood. There was no difference between arterial and venous RBC glutamate content in any of the rats studied.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

In humans and rats, consumption of single meals containing high levels of protein or free glutamate causes a transient rise in plasma glutamate levels but RBC glutamate levels remain unchanged (24 –27 ). The results shown in Table 2 illustrate that long-term arterial plasma glutamate levels are relatively stable, showing little variation in a number of physiologic and pathologic conditions. In contrast, cellular glutamate levels do vary; they tend to be higher with starvation, diabetes and high protein feeding, and are much higher when low protein diets are consumed. High levels of cellular glutamate have also been reported in humans fed a zero protein diet for 3 d and in rats fed low protein diets for 47 d (28 ). Pico et al. (6 ) found higher ratios of glutamate in cells vs. plasma in starved compared with fed rats, and Blackshear and Alberti (29 ) found higher red cell glutamate in ketotic streptozotocin diabetic rats. Such studies suggest that insulin levels are related to RBC glutamate levels and that conditions with lowered insulin levels such as diabetes or starvation result in higher glutamate concentrations in erythrocytes. Similarly, Divino Filho et al. (17 ) found that human red cell glutamate concentration correlated positively with the plasma levels of IGF-1 binding protein but negatively with the concentration of free IGF-1. They also showed that in rats, long-term feeding of diets with differing protein contents resulted in changes in RBC glutamate concentrations, whereas plasma glutamate levels were unaffected. As in the present work, Divino Filho et al. (17 ) reported the highest cellular glutamate levels in the rats fed the lowest protein diets. Again, erythrocyte glutamate levels correlated positively with plasma IGF-1 binding protein levels and negatively with free IGF-1 levels. Although Divino Filho et al. (17 ) proposed that IGF-1 plays a direct role in the transport of glutamate out of the red cells and into the tissue, the findings reported in this study do not support such a mechanism. It is more likely that IGF-1 and insulin have effects on erythrocyte glutamate synthesis and/or utilization.

The results presented here confirm that rat RBC do not take up glutamate and our findings with resealed RBC ghosts indicate that they are also incapable of releasing intracellular glutamate directly. Rats therefore lack a glutamate transporter in their erythrocytes in common with humans, sheep and rabbits (13 ,14 ,16 ) but in contrast to dogs, frogs and some species of kangaroos (10 –12 ), which express a sodium-linked glutamate transporter. Thus, the high level of glutamate found inside rat erythrocytes was not taken up directly. The origin of this cellular glutamate is not known, but it has been proposed to arise as a by-product of NAD synthesis through a glutamine amidotransferase reaction. In sheep, labeled glutamate infused into the circulation does not enter the RBC, but the infusion of labeled glutamine does lead to the intracellular accumulation of labeled glutamate, again suggesting glutamine as the origin of RBC glutamate (16 ). A further possibility, not addressed here, is that glutamyl peptides maybe taken up from the circulation and hydrolyzed within the RBC. The fate of cellular glutamate is also not known, although it is generally thought that some of it may be used for glutathione synthesis. The consistent finding that intracellular glutamate is markedly higher in rats fed protein-deficient diets indicates changes in erythrocyte glutamate metabolism under such conditions. But such simple concentration measurements offer no insight into the source or ultimate fate of that glutamate, or whether the changes are due to changes in glutamate synthesis, degradation or both. Similarly, all of the observations made in this work used rats adapted to their conditions for a number of days, and thus we are not able to determine how quickly the changes in intracellular glutamate levels arose.

In studies of amino acid metabolism, it is often assumed that plasma is the medium of interorgan transfer and that the cellular components of blood may be ignored. However, Elwyn and colleagues (8 ,9 ) showed that there was a redistribution of glutamate to and from RBC as the blood passed through tissues in dogs. Similarly, Aoki et al. (4 ,5 ) studying fasted humans reported that as blood passed through skeletal muscle (forearm), there was a decrease in plasma glutamate due to an uptake by both skeletal muscle and RBC. Because human blood cells, like those of rats, do not express a glutamate transporter, it is difficult to envision a mechanism for such changes. Moreover, in humans, labeled glutamate infused into the circulation does not enter RBC (30 ). In rats (Table 3) , we clearly found that any net change in whole-blood glutamate across a tissue bed occurred exclusively via the plasma compartment with no change observed in the level of cellular glutamate. Although the possibility exists that there was concomitant influx and efflux of glutamate in erythrocytes in vivo, this is unlikely given the lack of a suitable transporter. Thus, there is no evidence in rats that RBC play any role in the net flux of glutamate between organs, and measurements based solely on plasma may be considered valid. This is also likely to be true in humans but it may not be true in dogs because this species does express a RBC glutamate transporter .

The experiments reported here were not designed to determine arteriovenous differences among groups of rats, and the limited number in each group precludes any statistical assessment at this level. However, it is apparent from Table 3 that net glutamate output into the portal vein is higher in diabetic rats (which consume more food) and much higher in rats consuming very high (60%) protein diets than in the controls, and is not seen in rats fed protein-deficient (5%) diets. Thus, there appears to be a correlation with dietary protein intake, which raises the possibility that some of the glutamate released into the portal vein arose directly from the diet. This is unexpected from tracer studies of dietary glutamate utilization in pigs (2 ). This may be due to species differences because glutamate release into the portal circulation has been extensively reported for rats (24 ,31 ,32 ), particularly when large amounts of glutamate are fed (31 ). However, without suitable tracer studies, it is not possible to unequivocally identify the source of portal vein glutamate in rats.

The experiments reported here clearly show that rat RBC concentrate glutamate at levels more than twofold that of plasma and that the cellular glutamate levels are subject to long-term changes with diet and physiologic condition. In contrast, long-term plasma glutamate levels are relatively constant. Furthermore, rat RBC cannot take up or release glutamate because they lack a specific glutamate transporter. Despite earlier reports that RBC glutamate played a role in the interorgan transport of glutamate, this was not found to be true in any of the conditions studied. Therefore, analyses of glutamate metabolism in rats may confidently be made using measurements of either whole-blood or plasma glutamate levels.

	ACKNOWLEDGMENTS

 
The author thanks Yin Xiu Zhou for excellent technical assistance throughout this work.

	FOOTNOTES

 
¹ Supported in part by grants from the International Glutamate Technical Committee and the New Jersey Agricultural Experiment Station project # 14161.

Manuscript received 30 August 2001. Initial review completed 17 November 2001. Revision accepted 7 February 2002.

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TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

 

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