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Originally published In Press as doi:10.1074/jbc.M309520200 on January 13, 2004

J. Biol. Chem., Vol. 279, Issue 14, 13305-13310, April 2, 2004
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Granule Localization of Glutaminase in Human Neutrophils and the Consequence of Glutamine Utilization for Neutrophil Activity*

Linda Castell{ddagger}§, Caroline Vance{ddagger}, Rachel Abbott{ddagger}, Javier Marquez¶, and Paul Eggleton||**

From the {ddagger}Department of Biochemistry and the ||Medical Research Council Immunochemistry Unit, South Parks Road, University of Oxford OX1 3QU, United Kingdom, the **Institute of Biomedical and Clinical Science, Peninsula Medical School, Heavitree Road, Exeter EX1 2LU, United Kingdom, and the Molecular Biology and Biochemistry Department, University of Malaga, Malaga E-29071, Spain

Received for publication, August 27, 2003 , and in revised form, January 8, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The provision of glutamine in vivo has been observed to reduce to normal levels the neutrophilia observed after exhaustive exercise and to decrease the neutrophil chemoattractant, interleukin-8. Thus, the role for glutamine in the regulation of inflammatory mediators of human neutrophil activation was investigated. The study sought to establish whether glutamine supplementation in vitro affects neutrophil function at rest and whether glutaminase, the major enzyme that metabolizes glutamine, is present in human polymorphonuclear neutrophils (PMN). During in vitro studies, the addition of 2 mM glutamine increased the respiratory burst of human PMN stimulated with both phorbol myristate acetate (PMA) and formyl-methionyl-leucyl-phenylalanine. These observations were made using a highly sensitive, real time chemiluminescent probe, Pholasin®. Glutamine alone did not stimulate the release of reactive oxygen species. In a novel finding using glutaminase-specific antibodies in combination with flow cytometry and confocal microscopy, glutaminase was shown to be present on the surface of human PMN. Subcellular fractionation revealed that the enzyme was enriched in the secondary granules and could be released into cell culture medium upon stimulation with PMA. In conclusion, human PMN appeared to utilize glutamine and possess the appropriate glutaminase enzyme for metabolizing glutamine. This may depress some pro-inflammatory factors that occur during prolonged, exhaustive exercise.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Human polymorphonuclear neutrophils (PMNs)1 are a major class of the nonspecific immune response against infections. This is due, in part, to their ability to produce toxic forms of reactive oxygen species (ROS), including superoxide anions (O2–), hydrogen peroxide (H2O2), and hydroxyl radicals (OH–). The generation of ROS requires the cytosolic NADPH oxidase proteins to form a complex with several of the membrane-bound parts of the oxidases. Once assembled, direct stimulation of cells results in the release of into the extracellular compartment or into phagosomes. In turn, activated PMN synthesize and release a number of pro-inflammatory cytokines, including tumor necrosis factor (TNF), IL-1, IL-6, and IL-8 (1, 2). Recent work has shown that glutamine supplementation in vitro enhances both phagocytosis and ROS in isolated PMN (3) and, in vivo, suppresses IL-8 production by PMN (4). This has important consequences, as glutamine appears to exert a regulatory influence on inflammatory processes by PMN responding to inflammatory and infectious stimuli. For example, PMNs taken from patients with burns (5) or after surgery (6) have been shown to have improved bactericidal activity in vitro when glutamine is added to the culture medium for incubation.

Glutamine is an important substrate for some key cells of the immune system, such as macrophages and lymphocytes (7, 8). It acts as a nitrogen donor for purine and pyrimidine nucleotide synthesis for new DNA synthesis and for mRNA repair. Although classified as non-essential, recent evidence suggests that glutamine is conditionally essential when it becomes rapidly depleted in the blood in stressful situations. Plasma glutamine concentrations are substantially decreased by clinical trauma such as in major surgery by 37% (9) or after prolonged, exhaustive exercise by 20–25% (10). The provision of glutamine or glutamine precursors to endurance athletes has resulted in a decreased self-reported incidence of illness, particularly for upper respiratory tract infections (URTI), in four studies (see Castell, 2003; Ref. 4). Increasing evidence, in vitro and in vivo, suggests that PMN may benefit from exogenous glutamine, which repletes the decrease in the blood concentration observed after stress.

The energy substrate for PMN has traditionally been thought to be glucose. However, it is possible that PMN could use glutamine, particularly in cases such as severe infection in which glucose is restricted. Evidence for a direct effect of glutamine on neutrophil function has been demonstrated in rats, where it was shown that isolated PMN utilized glutamine at a rate of 12.8 nmol min–1 mg–1 protein (11) in the absence of glucose. The same workers further confirmed the presence of phosphate-dependent glutaminase (GA) in rat PMN using Western blots and immunocytochemistry methods. To date, the presence of GA in human PMN has not been demonstrated, despite several attempts to do so.

Strenuous exercise stimulates leukocytosis and neutrophilia (12–14) and the release of immature PMN from bone marrow (15, 16). However, there are conflicting reports upon the effects of exercise on ROS release by PMN (17). Significant increases have been observed in the plasma concentration of IL-8 (~2-fold) immediately after and 1.5 h after a marathon (18) and in cell IL-8 production after a rowing ergo test (19). This suggests enhanced post-exercise activation of PMN. Significant decreases in post-exercise neutrophilia (~1.75-fold) and cell IL-8 production (~3.5-fold) have been observed after a marathon in runners taking glutamine compared with a placebo (20). Since then, in three further studies in our laboratory, a reduction in IL-8 associated with glutamine feeding after exercise has been observed (see Castell, 2003; Ref. 4). Clinical studies (21, 22) established that IL-8 production was reduced in both surgical patients and patients with acute pancreatitis who received glutamine-enriched parenteral nutrition. IL-8 is a potent chemoattractant that perpetuates the inflammatory response and attracts PMN to the site of tissue damage.

The present study was set up to establish whether glutamine supplementation affects human neutrophil respiratory burst via in vitro incubation of blood with/without glutamine and to determine whether human PMN utilizes glutamine via the presence of GA, using both immunochemical techniques and confocal microscopy.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Subjects and Blood Samples—Ethical permission for the studies was obtained from the Central Oxford Research Ethics Committee (COREC). Resting blood samples were taken from healthy controls recruited from colleagues. Neutrophil function was measured with a novel chemiluminescence assay as described below.

Isolation of PMN—Isolated whole cells and subcellular neutrophil fractions were assayed for the presence of glutaminase. Neutrophils were isolated as described previously (23). Briefly, 5 ml of anticoagulated (potassium EDTA, 1.5 mg/ml) blood was layered on top of 4 ml of PolymorphPrepTM (Nycomed Pharma AS) in a 15-ml conical tube that was then spun at 1000 x g for 45 min. The layer of PMN was removed using a Pasteur pipette and placed with an equal volume of hypertonic sodium chloride solution (0.45% w/v) in another 15-ml tube to reduce viscosity. Following another spin, the supernatant was aspirated and the pellet resuspended in lysis buffer (isotonic NH4Cl) and spun again. The pellet was resuspended in 0.5 ml of phosphate-buffered saline without calcium or magnesium (PBS2–), and the cells were counted and checked for viability by the exclusion of 0.02% (v/v) trypan blue and stored at 4 °C until required.

Subcellular Fractionation of PMN—Subcellular fractionation of human PMN by nitrogen cavitation is an established technique for elucidating PMN structural composition (24). Briefly, freshly isolated, unstimulated PMN (2 x 108 per condition) were resuspended in 5 ml of nitrogen cavitation buffer (0.34 M sucrose, 10 mM Hepes, 1 mM EDTA, 0.1 mM MgCl2, and 1 mM Na2ATP, pH 7.4), followed by lysis by nitrogen cavitation (15 min, 400 p.s.i.) (4 °C). The lysates were centrifuged (1000 x g), and supernatants were then subjected to isopycnic sucrose density gradient fractionation on linear 20–55% sucrose gradients in a Beckman SW-28 swinging bucket rotor (100,000 x g; 3 h, 4 °C) exactly as described previously. 1.5-ml fractions were collected from each gradient and analyzed for marker proteins, plasma membrane (alkaline phosphatase), primary granules (myeloperoxidase), and secondary granules (lactoferrin) as described previously (25).

Whole Blood Neutrophil Oxidative Burst Measurement with and without Glutamine, Using a Microplate Luminometer—The novel chemiluminescent method is designed to analyze the oxidative burst in real time by emitted light (ABEL®, Knight Scientific Ltd, Plymouth, United Kingdom). This technique uses the photoprotein Pholasin®, which is isolated from the marine, rock-boring, bioluminescent mollusc Pholas dactylus, the common piddock. Pholasin® only emits light when activated by free radicals and other reactive oxygen species such as those released by activated leukocytes.

Whole blood was diluted 1:10 in RPMI 1640 (Sigma) with or without glutamine (final concentration 2 mM) and with antibiotics (1:1000 dilution of streptomycin and penicillin). After incubating for 3 h at 37 °C with continuous shaking, the blood was assayed for PMN activity using the chemiluminescence assay and measuring with the Anthos Lucy 1 luminometer (Labtech, Uckfield, United Kingdom) as outlined below. Aliquots (20 µl) were further diluted in 2 ml of blood dilution buffer (Hanks' buffered saline plus 20 mM HEPES). Using an opaque, white, 96-well plate, 90 µl of reconstitution and assay buffer was added to each well, together with 20 µl of Adjuvant-K to enhance the luminescence, 50 µl of Pholasin®, and 20 µl of diluted whole blood. The cells were stimulated by the addition of 20 µl of either formyl-methionyl-leucylphenylalanine (fMLP) (final concentration 1 µM) or PMA (final concentration 0.16 µM), and the response was measured until the reaction had passed (5 min with fMLP and 30 min with PMA) using a microplate luminometer (Anthos Lucy 1).

Production of Anti-GST-LGA347–602 Antibodies—An EcoRI-XhoI fragment containing nucleotides 1074–1909 of the human liver-type GA (LGA) cloned from ZR-75 breast cancer cells (26) was PCR-amplified and ligated in-frame in the EcoRI/XhoI site of expression vector pGEX-6P-1 (Amersham Biosciences). This fragment, encoding for the amino acids 347–602 of the C-terminal half of human LGA, was expressed in Escherichia coli as a GST fusion protein (GST-GA347–602). Protein expression and affinity purification were performed as described previously (26) using a glutathione-Sepharose affinity column. The purified recombinant protein was used for hyperimmunization of New Zealand White rabbits, and polyclonal antibodies were generated as described elsewhere (27).

SDS-PAGE and Western Blot of Glutaminase—Approximately 40 µg of neutrophil membrane sub-fractions, prepared by nitrogen cavitation as described previously (25), were suspended in sample buffer (62 mmol/liter Tris-Cl, pH 6.8, 2% SDS, 10% glycerol, and 0.01% bromphenol blue with 5% 2-mercaptoethanol), incubated for 5 min at 100 °C, and then applied to lanes of a 12% polyacrylamide mini-gel; electrophoresis was performed in Tris-buffered saline (TBS) at a constant current of 60 mA for 2 h. Molecular weight standards (Bio-Rad) were run simultaneously. The gel was stained with Coomassie Blue and dried. A parallel SDS-PAGE gel was run as above, and the separated proteins were transferred directly by semi-dry blotting onto polyvinyl difluoride (PVDF) transfer membrane for 90 min at a constant current of 0.8 mA. After saturation of the nonspecific sites with 5% nonfat milk-TBS overnight, the proteins were probed with a 1:200 dilution of rabbit anti-human glutaminase antibody prepared as described below. The blot was then washed in 20 mM Tris-HCl, pH 7.5, and 0.14 mM NaCl containing 0.4% Tween 20 (TBS-Tween) and then incubated for 1 h in an anti-rabbit peroxidase-conjugated IgG antibody diluted 1:1000 in TBS-Tween; the immunoblot was exposed to an ECL immunoassay substrate reagent (Amersham Biosciences) for 1 min to detect any signal, and then the membrane was exposed to x-ray film for 5 min.

Flow Cytometry and Confocal Microscopy—Detection of GA on the surface of human PMN was achieved using both flow cytometry and a Bio-Rad Radiance 2000 Confocal Laser Scanning Microscope (Zeiss Axiostar) and an Image Analysis work station. PMNs were incubated with 1:50 diluted rabbit anti-human glutaminase for 30 min, and, after three washes in PBS2–, the cells were incubated with 1:50 goat-anti rabbit-FITC-conjugated IgG for 30 min at 4 °C and then washed three times in PBS2–. In the flow cytometry studies, the PMNs were stimulated with either 0.16 µM (100 ng/ml) PMA or 0.1 µM fMLP or left in phosphate-buffered saline with calcium and magnesium (PBS2+) for 30 min at 37 °C before probing with test antibodies. The cells were then either fixed in 2% (v/v) paraformaldehyde and analyzed on a BD Biosciences FACScan or spun onto microscope slides at 500 rpm in a Cytospin centrifuge (Shandon Southern Instruments), and cover slips were mounted over the slides using an anti-quenching reagent Citifluor (Citifluor UK Chemical Lab, Canterbury, United Kingdom) and sealed with nail polish. The cells were viewed on the Zeiss microscope fitted with a confocal argon-krypton mixed gas laser. Images were taken serially from the top to the bottom of each cell using a z plane motorized sub-stage. The appropriate excitation and emission filters for FITC were employed. Several data sets were collected for each experiment.

Statistical Analysis—Results were compared using non-parametric analysis with Wilcoxon paired t tests.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Real Time Oxidative Response of Stimulated Leukocytes in the Presence of Glutamine—Whole blood leukocytes from healthy control individuals stimulated with PMA (final concentration 0.16 µM) and fMLP (final concentration 1 µM) were incubated with the highly sensitive real time chemiluminescence probe Pholasin® to examine ROS release in the presence and absence of glutamine. Four of six samples gave an increased response (39.2%) to fMLP when incubated with glutamine compared with no glutamine. In the fMLP experiments there were two individuals who did not respond to the addition of glutamine. However, the response was not diminished whether or not glutamine was present. In a separate group, four of four samples gave an increased response to PMA (31.1%) (Fig. 1).



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  		FIG. 1.
Effect of glutamine on real time release of ROS from whole blood human leukocytes stimulated with PMA and fMLP and detected by Pholasin®-dependent chemiluminescence. Aliquots of diluted whole blood (1:200) were incubated for 30 min at 37 °C before incubation with Pholasin® (0.5 µg/well) before the addition of PMA (0.16 µM) (A) or fMLP (1 µM) (B). The effect of glutamine (solid line) on ROS release was monitored for up to 30 min for PMA or 5 min for fMLP at 12 s intervals. Results are shown for single experiments performed in duplicate on four different healthy individuals. ALU, actual light units.

 
Glutamine stimulated the oxidative burst from PMA-stimulated cells ~2-fold without affecting the speed of response (Fig 1A). However, fMLP-stimulated cells (Fig. 1B) increased both the speed of response and the oxidative burst in the presence of glutamine.

Detection of Glutaminase on the Surface of Human PMN— The observation that glutamine had a pro-stimulatory effect on oxidative burst from leukocytes prompted us to examine whether human PMN, as the predominant generator of superoxide, possessed the glutamine-metabolizing enzyme GA on its cell surface. Intact PMNs were isolated as described above and assessed for the presence of GA by flow cytometry. As shown in Fig. 2a, resting cells appeared as two populations of cells (denoted M1 and M2) with respect to surface GA. Approximately 44% of PMN had very little surface GA mean fluorescence intensity (MFI = 8.4), whereas 56% of the cells were highly stained for GA (MFI = 100). Nonspecific staining accounted for an MFI of 4.8. Treatment of the cells with PMA led to loss of surface GA and the loss of highly stained GA populations of cells (Fig 2b). In contrast, fMLP had very little effect on surface GA levels in the two populations, M1 and M2 (Fig 2c). Imaging the surface of PMN in 0.5 micron sections labeled with FITC-anti-GA (Fig. 3) showed that the GA was on or near the cell surface and could be detected 3–3.5 microns into the cell surface but, most intensely, at the surface of the cell.



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  		FIG. 2.
Effect of Protein Kinase C activators PMA and fMLP on the cell surface expression of glutaminase and its release into cell-free media. PMNs were treated with no stimulation (a), stimulated with 0.16 µM PMA (b), or 0.1 µM fMLP (c) for 30 min at 37 °C. Non-stimulated PMNs, as depicted in panel a, were probed with irrelevant rabbit anti-human IgG (unfilled fluorescence peak). In parallel, non-stimulated and stimulated PMNs in panels a–c were analyzed by a fluorescence-activated cell sorter using a rabbit-specific, affinity-purified polyclonal antibody against human glutaminase. This was detected using an anti-rabbit FITC-conjugated second antibody. Representative results of three independent experiments are shown. d, PMNs from the same individual (2.5 x 106) in 50-µl aliquots were treated as described above and under "Materials and Methods." Equal amounts of cell-free supernatants were separated by SDS-PAGE gels and immunoblotted onto PVDF membranes, which were then probed with an anti-glutaminase antibody. Glutaminase was only detected in cell-free supernatants removed from PMA-treated cells (panel d, middle lane).

 



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  		FIG. 3.
Localization of glutaminase at the ruffled cell surface of human neutrophils. a, confocal microscopic images of isolated neutrophils were taken consecutively from the top of the cell every 0.5 µM below the surface level (z plane) and show that the glutaminase is distributed over the pseudopodia of the cells, which appears to extend ~3.5 µM deep. b, transmission electron microscope image of neutrophils displaying pseudopodia.

 
Secretion of Glutaminase—A number of human cells, including liver and kidney, have been reported to produce GA, but to date GA has not been identified in human PMN. To investigate if the source of the GA came from the PMNs themselves, we stimulated the cells with PMA (100 ng/ml) and fMLP for 30 min. The PBS in which 5 x 106 cells had been incubated were collected and concentrated to 100 µl volumes. Then, 50 µl aliquots of these were loaded on a 12% v/v SDS-PAGE gel, together with purified GA, and immunoblotted onto PVDF membranes and probed with rabbit anti-human glutaminase polyclonal antisera. As shown in Fig. 2d, Western blot analysis revealed that the cell-free supernatants from cells that had been treated with PMA contained GA. In contrast, supernatants from resting cells and fMLP-treated cells contained no GA as probed by immunoblotting. The presence of GA in the cell media suggested that it might be secreted from the cells. Thus, it was next decided to examine neutrophil subfractions, which had been prepared previously by nitrogen cavitation into four main fractions, namely primary granules ({alpha}), secondary granules ({beta}), endoplasmic and plasma membrane-enriched fraction ({gamma}), and cell supernatant (S). The protein profiles of these four fractions are shown in Fig 4a, and the immunoblot of these protein fractions (Fig 4b) reveal that a protein 65–70 kDa was detected in the secondary granules but not in any of the other purified subcellular fractions.



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  		FIG. 4.
Localization of glutaminase in unstimulated PMN by subcellular fractionation. a, PMN (2 x 108) were lysed by nitrogen cavitation, centrifuged on an isopycnic sucrose gradient, and fractionated as described under "Materials and Methods." a, subcellular organelles (40 µg/fraction) were pooled according to their marker activities, run on SDS-PAGE gels, and stained with Coomassie Blue. The different bands are indicated by arrows and consist of primary granules ({alpha}), secondary granules ({beta}), plasma membrane ({gamma}) cytosolic fraction (S), and whole neutrophil lysates (NL), together with markers (M). b, an immunoblot of a parallel set of subcellular organelles was probed with anti-human glutaminase to determine which subfraction contained glutaminase. Glutaminase was detected in the secondary granules (shown by the arrow at the top of the panel) as a single band between 50 and 80 kDa.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The fact that lymphocytes and macrophages use high rates of glutamine has been known for a number of years (7, 8). More recent observations have provided evidence that glutamine is also utilized by rat PMN (28). Blannin et al. (42) observed no uptake of glutamine by human neutrophils stimulated in vitro with lipopolysaccharide. More recently (43), Healy et al. reported glutamine utilization by human neutrophils (apparent Km 1.29 mM). The ability of glutamine metabolism to enhance the bacterial function of PMN has been investigated in burn patients (5) and postoperative individuals (3). The molecular mechanism of the production of enhanced reactive oxygen intermediates by glutamine is only just being characterized. Recent data suggest that glutamine can increase the expression of some of the proteins that together form the NADPH oxidase components, which, in turn, regulate the production of the superoxide anion (29). In particular, glutamine appears to increase the expression of the cytosolic protein gp47phox as well as the membrane-associated components and NADPH oxidases p22phox and p47phox. Moreover, blocking glutamine metabolism by inhibiting GA in PMN causes a significant decrease in superoxide production. Therefore, the presence and subcellular location of glutamine appears to be important for glutamine-dependent superoxide production by PMN.

In the present study, the results obtained support the notion that glutamine can enhance oxidative burst by human PMN in the presence of phorbol esters and formyl tripeptide stimulants. These data were achieved using a real time in vitro assay of oxidative burst. The supplementation of whole blood leukocytes with 2 mM glutamine not only led to a greater production of superoxide but also increased the initial rate of activation. GA activity has been reported in rat PMN (28), and numerous studies have detected GA in human tumor cells (30, 31), where GA is an essential requirement for cell growth.

In vitro data from oxidative burst measurements in culture medium, with or without glutamine used to incubate whole blood samples from marathon runners, suggested a role for glutamine and glutaminase in neutrophil metabolism during inflammation. However, during inflammation, phagosomes only fuse with the extracellular matrix at the cytosolic and membrane components, and the NADPH oxidase pathway translocates to form the complex upon alteration. Therefore, if glutamine plays a role in production, one might expect to find GA in a similar location. The present results indicate that GA is present on the surface of cells and is released into the medium upon PMA, but not fMLP, stimulation. In the present investigation, more precise information was obtained on the localization of GA by employing confocal microscopy. By generating 0.5 µM optical sections through individual PMN, it was observed that GA was distributed 0–3.5 µM into the cell surface. PMNs are ruffled cells, and the GA appeared to be predominantly localized in pseudo lamellipodia at or near the plasma membrane.

In an initial study, little or no evidence of the presence of GA in human PMN was found using three different methods, namely freeze-thaw, homogenization, and nebulization. None of these techniques was apparently able to break open the granules. Consequently, to determine in more detail the subcellular source of glutamine in human PMN, other techniques were employed. Nitrogen cavitation and discontinuous Percoll density gradient centrifugation provided a simple and rapid means of separating azurophil granules ({alpha}), specific granules ({beta}), and plasma membrane ({gamma}) vesicles, as well as a cytoplasm fraction. GA was detected in the secondary granules but, surprisingly, not in the plasma membrane fractions. Our data suggest that, in intact PMN, GA is secreted from the secondary granules and binds to the cell surface. Following PMA but not fMLP stimulation, the GA is removed from the membrane fractions and can be found in the extracellular culture medium. It seems that, as granules fuse with the plasma membrane at the sites of developing phagosomes, the NADPH oxidase and GA enzymes would be in close proximity to the plasma membrane in response to stimulation. This would bring them into close contact with their target substrates. Interestingly, a number of groups have recently identified isoforms of glutaminase in human tumor cell lines at the mRNA and protein level (31) and in mammalian brain cells (32). In the present study, treatment of human PMN with PMA, a degranulating agent, resulted in the release of glutaminase from the cell surface into cell culture media. However, three bands were recognized by the polyclonal anti-glutaminase antibody. This is consistent with the results of previous investigators that revealed limited proteolytic digestion of isoforms of GA (33). When intact secondary granules were subsequently examined by Western blotting, a single non-proteolytic glutaminase band of ~65 kDa was observed and compared favorably with that found for rat neutrophil GA, which is ~65 kDa (11).

Somewhat surprisingly, although a marked increase in glutaminase expression in human neutrophils on the cell surface occurred following stimulation with PMA, there was no change in expression after stimulation with fMLP. Both fMLP and PMA are well known proinflammatory agonists that activate protein kinase C (PKC) by different cellular pathways. fMLP activates PKC indirectly via interactions with surface fMLP receptors and G-protein activation and subsequent inositol phosphate hydrolysis. The phorbol ester PMA, on the other hand, passes directly through the cell membrane and mimics diacylglycerol, a natural ligand and activator of protein kinase C. Both of these agonists can activate the release of various enzymes and inflammatory mediators into the extracellular fluid of stimulated cells. In this study, the source of glutaminase appears to be located in the secondary granules of leukocytes and expressed on the cell surface, where it is available for glutamine metabolism. The origin of the PMA-induced change observed might be distal to the stage of PKC activation in the process that results in glutaminase-containing vesicles moving toward and fusing with the plasma membrane. Differences in PKC activity and translocation have been reported previously (34). Alternatively, PMA can activate intracellular enzymes other than PKC, which may explain the release of glutaminase by PMA as compared with fMLP (35). Further studies will be necessary to elucidate the cause of this difference.

There is an increase in the plasma concentration of neutrophil granule contents after strenuous exercise (36). Large numbers of PMN are produced by bone marrow on a daily basis in healthy humans (37). It is possible that this rapid turnover relates to constant immunosurveillance as the PMN pass through the capillary bed. In these circumstances, glutamine utilization might play an important role in maintaining these cells in a constant state of "awareness." PMN also contain large reserves of the endogenous antioxidant glutathione (38), for which glutamine is a precursor.

The fact that in vivo glutamine feeding has affected cell production of IL-8 in several in vitro studies is clearly important. IL-8 acts as a potent chemoattractant for PMNs and induces them to leave the blood stream and migrate into damaged or infected tissue. IL-8 also activates NADPH oxidase and nitric oxide synthase, which, in turn, causes the release of ROS from PMN granules.

Results from recent studies suggest that glutamine has a modulating effect on IL-8 cytokine production in various cell types during activation by inflammatory mediators (4, 21, 22, 39–41). The signaling of this neutrophil chemoattractant might be diminished because of the effect of additional glutamine in the blood upon circulating mature or immature PMN. Increased numbers of the latter are recruited during the leukocytosis that occurs in response to strenuous exercise. However, glutamine supplementation has been shown to help reduce the number of circulating PMNs to near normal levels compared with a placebo 16 h after a race (20). The mechanism of glutamine-mediated regulation of IL-8 production by peripheral blood mononucleocytes is not yet known. It could reflect changes in cell receptor signaling, transcription, and/or translation expression by modulatory anti-inflammatory cytokines such as IL-10 (39).

In conclusion, the present study suggests that glutamine supplementation might enhance the respiratory burst of human PMN to specific inflammatory stimuli in vitro. Moreover, the novel finding has been made that there is a sub-cellular source of GA, the major enzyme responsible for glutamine metabolism, in the secretory granules of human PMN. This is likely to contribute to the modulation of immune function of PMN in clinical and exercise-induced stress.


   FOOTNOTES
 
* This work was supported by University Research Funds (to L. C.), funding from the Medical Research Council Immunochemistry Unit, University of Oxford, and Arthritis Research Campaign Grant E0543 (to P. E.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ To whom correspondence should be addressed: Nuffield Dept. of Anaesthetics, University of Oxford, Radcliffe Infirmary, Oxford OX2 6HE, United Kingdom. E-mail: lindy.castell{at}nda.ox.ac.uk.

1 The abbreviations used are: PMN, polymorphonuclear neutrophils; FITC, fluorescein isothiocyanate; fMLP, formyl-methionyl-leucyl-phenylalanine; GA, glutaminase; GST, glutathione S-transferase; IL, interleukin; LGA, liver-type GA; MFI, mean fluorescence intensity; PBS, phosphate-buffered saline; PKC, protein kinase C; PMA, phorbol-12-myristate-13-acetate; PVDF, polyvinylidene fluoride; ROS, reactive oxygen species; TBS, Tris-buffered saline. Back


   ACKNOWLEDGMENTS
 
We are grateful to Dr Jan Knight of Knight Scientific Limited, Plymouth, United Kingdom for advice and help with the microplate Pholasin® assay and to LabTech for the loan of the Lucy Anthos microplate luminometer.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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