Role of glutamine and arginase in protection against ammonia-induced cell death in gastric epithelial cells

Eiji Nakamura and Susan J. Hagen

Department of Surgery, Beth Israel Deaconess Medical Center, Boston, Massachusetts 02215


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Ammonia is a cytotoxic factor produced during Helicobacter pylori infection that may reduce the survival of surface epithelial cells. Here we examine whether ammonia kills cells and whether L-glutamine (L-Gln) protects against cell death by stimulating ammonia detoxification pathways. Cell viability and vacuolation were quantified in rat gastric epithelial (RGM1) cells incubated with ammonium chloride at pH 7.4 in the presence or absence of L-Gln. Incubation of RGM1 cells with ammonium chloride caused a dose-dependent increase in cell death and vacuolation, which were both inhibited by L-Gln. We show that RGM1 cells metabolize ammonia to urea via arginase, a process that is stimulated by L-Gln and results in reduced ammonia cytotoxicity. L-Gln also inhibits the uptake and facilitates the extrusion of ammonia from cells. Blockade of glutamine synthetase did not reduce the survival of RGM1 cells, demonstrating that the conversion of L-glutamate and ammonia to L-Gln is not involved in ammonia detoxification. Thus our data support a role for L-Gln and arginase in protection against ammonia-induced cell death in gastric epithelial cells.

Helicobacter pylori; rat; rat gastric epithelial 1 cells; NH4Cl


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

INFECTION OF THE STOMACH by Helicobacter pylori causes chronic-active gastritis and peptic/duodenal ulcer disease in humans and in many animal models. Terminal deoxynucleotidyltransferase-mediated dUTP nick-end labeling staining of the mucosa has shown that apoptosis increases significantly during H. pylori infection and is most prevalent in surface epithelial cells (42, 51). Although apoptosis occurs, it is not clear what factors associated with H. pylori infection cause injury and death of surface epithelial cells. It has been reported that H. pylori produce many deleterious factors for gastric epithelial cells, such as vacuolating cytotoxin (VacA), gene products of the Cag pathogenicity island (PAI), and urease. Although VacA was initially thought to be cytotoxic to gastric epithelial cells, compelling evidence against this contention was recently shown in the gerbil model of infection by targeted deletion of the VacA gene in H. pylori, in which severe gastritis and mucosal injury were present (36, 53). In addition, H. felis, a related gastric Helicobacter that does not possess VacA or the Cag PAI but has potent urease activity (33), induces severe gastritis and injury to gastric epithelial cells in several animal models (12, 30). These combined results suggest that injury to epithelial cells during H. pylori infection may occur from urease-derived ammonia (NH3) rather than from VacA or gene products of the Cag PAI. In fact, the severity of gastric injury during H. pylori infection is correlated with the concentration of NH3 in the gastric juice (47) or the urease activity of H. pylori (26). Patients with H. pylori infection show a significant increase in gastric juice NH3 compared with uninfected control patients (13, 14, 23, 24, 27, 32, 34, 35, 46, 50, 58).

A number of studies recently showed that NH3 affects the gastric mucosa in vivo and gastric epithelial cells in vitro. NH3, at a concentration below that detected in H. pylori-infected patients, inhibits oxygen consumption (48), cell proliferation (31), and acid secretion (17, 18, 57). In addition, NH3 kills parietal and chief cells in isolated gastric glands by necrosis and apoptosis, respectively (17). NH3, generated by using ammonium chloride (NH4Cl) or urea/urease, kills gastric MKN 45 cells alone and in combination with cytokines, such as tumor necrosis factor-alpha or interferon-gamma (21). Furthermore, NH3 retards restitution of the injured gastric mucosa (43), leading to impaired barrier function. Thus NH3 may significantly impair mucosal homeostasis, resulting in injury and death of gastric epithelial cells during H. pylori infection.

Although gastric surface epithelial cells are exposed to high levels of NH3 during H. pylori infection, it is not established whether these cells are injured by NH3 or whether they possess any mechanism(s) to protect against NH3-induced injury. In the liver, systemic NH3 detoxification occurs in metabolic zones, where periportal and periveneous hepatocytes have unique enzymatic pathways for the production of nontoxic NH3 metabolites such as urea and glutamine, respectively (20, 59). Brain glial cells also produce glutamine from glutamate and NH3 to protect neurons from NH3-induced cytotoxicity (9, 59). Thus it is possible that gastric epithelial cells have the ability to process NH3, either by facilitating the production of urea from NH3 or by converting glutamate and NH3 to glutamine. Either detoxification pathway would be beneficial to protect surface epithelial cells against the cytotoxic effects of NH3 in the gastric lumen, in general, and during H. pylori infection, in particular.

Thus the purpose of this study was to determine whether NH3 affects the survival of gastric surface epithelial cells and, if so, to determine whether glutamine protects surface epithelial cells from injury by facilitating NH3 detoxification. To accomplish this, we measured cell viability and the degree of vacuolation in rat gastric epithelial (RGM1) cells that were exposed to NH4Cl, producing NH3 and ammonium, with or without L-glutamine (L-Gln). Our results indicate that NH4Cl significantly reduces the viability of RGM1 cells and that L-Gln and L-glutamate (L-Glu) both protect RGM1 cells against NH4Cl-induced cell death. Our results establish that RGM1 cells metabolize NH3 to urea, that L-Gln protects by decreasing the intracellular accumulation of NH3 and increasing NH3 metabolism, and that the conversion of NH3 and L-Glu to L-Gln via glutamine synthetase does not protect RGM1 cells. Because L-Gln completely reverses the cytotoxic effects of NH3 in our study, it is proposed that L-Gln supplementation may be beneficial to reduce mucosal injury during H. pylori infection.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
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REFERENCES

Preparation of RGM1 cell cultures. Rat gastric epithelial cell line, RGM1 cells, established by Dr. H. Matsui, Institute of Physical and Chemical Science (RIKEN) Cell Bank and Institute of Clinical Medicine, University of Tsukuba, Tsukuba, Japan (29), are nontransformed gastric surface epithelial cells. RGM1 cells were cultured in DMEM-F12 (1:1) supplemented with heat-inactivated 10% fetal bovine serum (FBS; GIBCO/BRL, Gaithersburg, MD), 100 U/ml penicillin, 100 U/ml streptomycin, and 0.25 µg/ml amphotericin B. Confluent monolayers of RGM1 cells were starved for 24 h in culture medium without FBS (DMEM-F12 containing 15 mM HEPES at pH 7.4) at 37°C under 5% CO2 in air and then used for experiments. All experiments were performed in standard (STD) buffer at pH 7.4 in the presence and/or absence of reagent(s) without FBS.

Treatment of RGM1 cells with NH4Cl or methylamine with or without L-Gln or L-Glu. Starved RGM1 cells were transferred to STD buffer containing (in mM) 147 Na+, 5.0 K+, 131 Cl-, 1.3 Mg2+, 1.3 SO<UP><SUB>4</SUB><SUP>2−</SUP></UP>, 2 Ca2+, 25 HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>, 15 HEPES, and 20 D-glucose at pH 7.4 and incubated with 0-100 mM NH4Cl or 0-30 mM methylamine hydrochloride (MeNH2) in the presence or absence of 0-20 mM L-Gln. NH4Cl was used as the source of NH3, a primary amine weak base. Because NH3 is at equilibrium with its protonated form (NH<UP><SUB>4</SUB><SUP>+</SUP></UP>) in a pH-dependent manner, 30 mM NH4Cl (pKa is 9.24) results in 0.44 mM NH3 (1.4%) and 29.56 mM NH<UP><SUB>4</SUB><SUP>+</SUP></UP> (98.6%) at pH 7.4. MeNH2 is also a primary amine weak base that is at equilibrium with its protonated form (MeNH<UP><SUB>2</SUB><SUP>+</SUP></UP>) in a pH-dependent manner. MeNH2 at 3 mM (pKa is 10.66) results in 0.0017 mM MeNH2 (0.03%) and 2.9993 mM MeNH<UP><SUB>2</SUB><SUP>+</SUP></UP> (99.97%) at pH 7.4. Both NH3 and MeNH2 enter cells and result in intracellular and lysosomal alkalinization (10). However, NH3 is a substrate in intracellular metabolic pathways, and MeNH2 is not. Thus MeNH2 was used as a control weak base for all experiments. To examine (indirectly) the role of urea production in protection of RGM1 cells against NH3, RGM1 cells were incubated with 0.1-1 mM Nomega -hydroxy-nor-L-arginine (nor-NOHA; Calbiochem, San Diego, CA) in the presence of 30 mM NH4Cl or 3 mM MeNH2, with or without L-Gln. nor-NOHA is a potent and specific competitive inhibitor of arginase (15, 45), a key enzyme in the production of urea (from arginine) in the urea cycle. To examine whether the conversion of L-Glu and NH3 into L-Gln is involved in NH3 detoxification, RGM1 cells were incubated with 0.1- 10 mM L-S-[3-amino-3-carboxypropyl]-S-methylsulfoximine [methionine sulfoximine (MS)], a potent inhibitor of glutamine synthetase (52). Glutamine synthetase is responsible for the conversion of L-Glu plus NH3 to L-Gln. L-Gln is an amino acid with an uncharged R group. L-Glu is an amino acid with a charged polar group. NH4Cl, MeNH2, L-Gln, L-Glu, MS, and all other buffer components were purchased from Sigma (St. Louis, MO).

Measurement of cell viability. The viability of RGM1 cells was evaluated by a colorimetric assay by using crystal violet (25), a cytochemical stain that binds to chromatin. For this assay, RGM1 cells were washed once with PBS to remove dead cells, fixed with methanol for 15 min, and then air-dried. The dried cells were stained with 0.1% crystal violet for 5 min at room temperature, washed twice with PBS, and then air-dried. Stained cells were solubilized with 0.5% SDS for 30 min with slight agitation. Lysates were diluted with 0.5% SDS, and the absorbance was measured at 590 nm by using a microplate reader. Crystal violet stain was purchased from Sigma.

Measurement of cell vacuolation. Intracellular acidic vacuoles, containing H+ generated by the vacuolar ATPase, expand in the presence of a weak base (in a concentration-dependent manner) because the unprotonated weak base freely partitions into the acidic space, is protonated by H+, and cannot freely exit (10). The resulting loss of H+ alkalinizes the vacuole and initiates further H+ generation by the vacuolar ATPase, which is followed by water movement into the vacuole and vacuole expansion (10). Because vacuolation is an indicator of intracellular weak base concentration, we evaluated the intracellular concentration of NH3 or MeNH2 by quantifying vacuolation.

To quantify vacuolation in RGM1 cells, uptake of neutral red into vacuoles was determined as described by Cover et al. (7, 8), with slight modification. In brief, RGM1 cells were incubated for 10 min at 37°C with 0.005% neutral red in STD buffer and then washed twice with PBS containing 0.3% BSA. The dye was extracted with isopropyl alcohol containing 0.04 M HCl. The extract was diluted, and the absorbance was measured at a test wavelength of 540 nm and a reference wavelength of 650 nm by using a microplate reader.

Assay for the extrusion of NH4Cl or MeNH2 from vacuoles. After the induction of vacuoles for 6 h with 30 mM NH4Cl or 3 mM MeNH2, RGM1 cells were incubated for 1 h in STD buffer with or without 0-20 mM L-Gln, in the absence of NH4Cl or MeNH2. Vacuolation was quantified as described above.

Measurement of MeNH2 accumulation in RGM1 cells. Intracellular accumulation of [14C]MeNH2 was measured in RGM1 cells that were incubated for 3 h, at 37°C, with 3 mM MeNH2 containing 0.5 µCi of [14C]MeNH2 · HCl (NEN Life Science Products, Boston, MA) and 0-20 mM L-Gln. Washing the cells with ice-cold PBS terminated the reaction. The cells were solubilized with 0.3 N NaOH, and the radioactivity was measured by liquid scintillation (Packard Instruments, Downers Grove, IL).

Measurement of urea production in RGM1 cells. Urea concentration in the culture supernatant was measured in two ways. First, by using a commercially available assay kit (Sigma), which follows the procedure of Ormsby (37), and second, by measuring the conversion of L-[guanido-14C]arginine into [14C]urea as described below for the measurement of arginase activity. For measurement of urea by the Sigma assay kit, RGM1 cells were cultured in 100-mm dishes to obtain 4 × 106 cells/dish. After starvation for 24 h, the cells were incubated with or without 20 mM L-Gln in the presence or absence of 30 mM NH4Cl for 6 h. The culture supernatant was collected from four dishes, combined into one sample, and lyophilized. The lyophilized sample was solubilized in PBS and used for the urea assay, where the absorbance at 540 nm of hydroxylamine generated by the reaction of urea with diacetylmonoxime was measured. This assay is not affected by other nitrogen compounds such as NH3 or nitrogen oxides (37). The urea concentration was determined from a standard curve by using urea purchased from Sigma.

Measurement of arginase activity in RGM1 cells. Starved RGM1 cells were incubated at 37°C for 6 h with STD buffer. The cells were solubilized and sonicated in lysis buffer containing 0.01% Triton X-100, 2 mg/ml BSA, 10 mM MnCl2, and 12 mM Na maleate (pH 7.5). After centrifugation at 1,000 g at 4°C, arginase activity was determined in the supernatant by measuring the conversion of L-[guanido-14C]arginine to [14C]urea (6, 39, 40). In brief, the supernatant was added to reaction buffer (100 mM glycine, pH 7.4) in the presence or absence of 30 mM NH4Cl, 20 mM L-Gln, and 1 mM nor-NOHA, and the reaction was started by the addition of 250 mM L-arginine containing 0.05 µCi of L-[guanido-14C]arginine. After 90 min at 37°C, the reaction was terminated by the addition of 0.8 ml of stop buffer containing 250 mM acetic acid, 100 mM urea, 10 mM L-arginine (pH 4.5), and a 50% suspension of Dowex 50-WX8 resin (H+ form). After centrifugation, the supernatant containing [14C]urea (500 µl) was measured by liquid scintillation. Under these conditions, the resin removed 99.8% of the arginine substrate and 99.0% of the converted ornithine. Arginase activity in the supernatant was extrapolated from a standard curve by using purified arginase (Sigma).

Morphological analysis of cell cultures. Cell morphology was evaluated in cultured RGM1 cells at 6 and 24 h in STD buffer or in STD buffer containing 30 mM NH4Cl or 3 mM MeNH2 with or without 20 mM L-Gln. Cells were photographed with a Nikon TE300 microscope (MicroVideo Instruments, Avon, MA) outfitted with an Orca charge-coupled device camera (Hamamatsu Photonics) and IP laboratory software (Scanalytics, Fairfax, VA).

Statistical analysis. The data represent means ± SE for four wells of RGM1 cells from three different experiments. Statistical differences were evaluated by using Dunnett's multiple comparison test and Student's t-test, with a value of P < 0.05 regarded as significant.


    RESULTS
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INTRODUCTION
MATERIALS AND METHODS
RESULTS
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NH4Cl and MeNH2 reduce the viability of cultured RGM1 cells. Treatment of RGM1 cells with NH4Cl or MeNH2 at pH 7.4 reduced viability in a concentration-dependent manner (Fig. 1). The viability of RGM1 cells was significantly reduced with 3, 10, 30, and 100 mM NH4Cl (Fig. 1A). Similarly, the viability of RGM1 cells was significantly reduced with 1, 3, 10, and 30 mM MeNH2 (Fig. 1B). For all further experiments, we used 30 mM NH4Cl, which reduced viability to 37.6 ± 1.1% of control, and 3 mM MeNH2, which reduced viability to 54.6 ± 0.5% of control. It should be noted that a significantly greater concentration of NH4Cl was required to produce the same reduction in viability compared with MeNH2.


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Fig. 1.   Effect of NH4Cl (A) or MeNH2 (B) on cell viability. Rat gastric epithelial (RGM1) cells were incubated with 1-100 mM NH4Cl (A) or 0.3-30 mM MeNH2 (B) for 24 h, and then the number of viable cells was quantified by the crystal violet assay. The data show that both NH4Cl and MeNH2 kill RGM1 cells in a dose-dependent manner. Values are means ± SE of 4 wells from 3 different experiments and are expressed as the percentage of control cells that were incubated with standard (STD) buffer alone. * Significant decrease in viability compared with control cells, P < 0.05.

Time course studies showed that the viability of RGM1 cells decreased over time in the presence of 30 mM NH4Cl or 3 mM MeNH2 (Fig. 2). The viability of RGM1 cells was significantly reduced at 12, 24, and 36 h, when incubated with 30 mM NH4Cl or 3 mM MeNH2.


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Fig. 2.   Time course analysis of cell viability in the presence of NH4Cl (A) or MeNH2 (B). RGM1 cells were incubated for 36 h in the presence or absence of 30 mM NH4Cl (A) or 3 mM MeNH2 (B), and the number of viable cells was quantified by the crystal violet assay. These data demonstrate that NH4Cl and MeNH2 significantly reduce the viability of RGM1 cells over time. Values are means ± SE of 4 wells from 3 different experiments and are expressed as the percentage of initial viability (0 h) in each group. * Significant decrease in viability compared with time-matched control cells, P < 0.05.

L-Gln improves the viability of RGM1 cells in the presence of NH4Cl and MeNH2. Treatment of RGM1 cells with L-Gln prevented the reduction in cell viability induced by 30 mM NH4Cl or 3 mM MeNH2 in a concentration-dependent manner (Fig. 3). In the presence of NH4Cl, significant protection occurred with 0.2, 2, and 20 mM L-Gln (Fig. 3A). In fact, 20 mM L-Gln completely (100.3 ± 1.1% of the initial value) protected RGM1 cells that were incubated with 30 mM NH4Cl (Fig. 3A). Similarly, significant protection occurred with 0.2, 2, and 20 mM L-Gln in RGM1 cells that were incubated with 3 mM MeNH2. Like with NH4Cl, 20 mM L-Gln completely (99.1 ± 1.1%) protected RGM1 cells that were incubated with 3 mM MeNH2 (Fig. 3B). L-Gln had no effect on the viability of RGM1 cells in the absence of NH4Cl or MeNH2 at 24 h (104.2 ± 3.6% viability with L-Gln vs. 100.0 ± 4.1% with STD buffer alone). In addition, treatment with 20 mM mannitol, used to control for osmotic changes produced by 20 mM L-Gln, had no effect on the viability of RGM1 cells that were treated with 30 mM NH4Cl (42.2 ± 1.9% viability with mannitol and NH4Cl vs. 45.0 ± 2.0% with NH4Cl alone).


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Fig. 3.   Effect of L-glutamine (L-Gln) on the viability of RGM1 cells incubated with NH4Cl (A) or MeNH2 (B). RGM1 cells were incubated with 0.02-20 mM L-Gln in the absence or presence of 30 mM NH4Cl (A) or 3 mM MeNH2 (B) for 24 h, and then cell viability was quantified by the crystal violet assay. The data show that L-Gln significantly improves the viability of RGM1 cells in a dose-dependent manner. Values are means ± SE of 4 wells from 3 different experiments and are expressed as percentage of control cells that were incubated with STD buffer alone. * Significant decrease in viability compared with control cells, P < 0.05. dagger  Significant increase in viability compared with cells treated with NH4Cl (A) or MeNH2 (B), P < 0.05.

NH4Cl or MeNH2 causes vacuolation of RGM1 cells that is reduced by L-Gln. Incubation of RGM1 cells for 6 h with NH4Cl or MeNH2 resulted in the vacuolation of RGM1 cells in a concentration-dependent manner (Table 1). Vacuolation increased significantly in the presence of 0.3-30 mM NH4Cl, resulting in a maximum increase of 86.4 ± 4.5% compared with control cells treated with buffer alone (Table 1). Similarly, vacuolation increased significantly in the presence of 0.3-10 mM MeNH2, resulting in a maximum increase of 181.6 ± 9.1% compared with control cells treated with buffer alone (Table 1). In all cases, the percentage of vacuolation induced by NH4Cl was significantly less than with an equal concentration of MeNH2 (Table 1).

                              
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Table 1.   Vacuolation of RGMI cells by NH4Cl or MeNH2

When RGM1 cells were treated with L-Gln in the presence of 30 mM NH4Cl for 6 h, vacuolation was reduced in a concentration-dependent manner (Fig. 4A). The vacuolation induced by NH4Cl decreased significantly with 0.02, 0.2, 2, and 20 mM L-Gln (Fig. 4A). Treatment with L-Gln also resulted in a decrease in vacuolation induced by 3 mM MeNH2 (Fig. 4B). L-Gln significantly reduced MeNH2-induced vacuolation at 0.2, 2, and 20 mM (Fig. 4B).


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Fig. 4.   Effect of L-Gln on NH4Cl- (A) and MeNH2-induced vacuolation (B). RGM1 cells were incubated for 6 h with 0.02-20 mM L-Gln in the absence or presence of 30 mM NH4Cl (A) or 3 mM MeNH2 (B), and then vacuolation was quantified by the neutral red uptake assay. These data show that both NH4Cl and MeNH2 significantly increase the number and size of vacuoles in RGM1 cells and that L-Gln significantly decreases the degree to which RGM1 cell vacuolation occurs in the presence of NH4Cl (A) or MeNH2 (B). Values are means ± SE of 4 wells from 3 different experiments and are expressed as percentage of control cells incubated with STD buffer alone. * Significant increase in vacuolation compared with control cells, P < 0.05. dagger  Significant reduction in vacuolation compared with cells incubated with NH4Cl (A) or MeNH2 (B) alone, P < 0.05.

Time course experiments with 3 mM MeNH2 showed that vacuolation increased rapidly for the first hour and then increased slowly 2-6 h thereafter (Fig. 5A). Treatment of RGM1 cells with L-Gln in the presence of MeNH2 significantly reduced vacuolation (by > 60%) in RGM1 cells (Fig. 5A). However, some vacuolation was always present with MeNH2 and L-Gln, compared with control cells incubated with buffer alone (Fig. 5A).


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Fig. 5.   Time course analysis of MeNH2- (A) and NH4Cl-induced vacuolation (B) of RGM1 cells in the presence or absence of L-Gln. RGM1 cells were incubated with 20 mM L-Gln in the presence of 3 mM MeNH2 (A), 30 mM NH4Cl (B), or STD buffer alone (control) for 6 h, and the rate of vacuolation was quantified by the neutral red uptake assay. Values are means ± SE of 4 wells from 3 different experiments, and the data are expressed as the change in optical density (OD) compared with the initial value at 0 h. The results demonstrate that L-Gln significantly decreases the rate of vacuolation in RGM1 cells. In addition, the data in B show that, when RGM1 cells are incubated with NH4Cl and L-Gln, the L-Gln-dependent response, identified in A, is greater than the "predicted response" to L-Gln. Thus there is a significant difference in the response of MeNH2 and NH4Cl to L-Gln. * Significant decrease in vacuolation compared with cells treated with MeNH2 (A) or NH4Cl (B) alone, P < 0.05.

Time course experiments revealed that the formation of vacuoles induced by 30 mM NH4Cl was slower than with MeNH2, whereby vacuoles increased rapidly for the first 2 h and then increased slowly for 2-6 h thereafter (Fig. 5B). When RGM1 cells were treated with 30 mM NH4Cl in the presence of 20 mM L-Gln, the response to L-Gln was significantly different than described in Fig. 5A for MeNH2 and L-Gln (L-Gln-dependent response). For the first 2 h, vacuole formation with 30 mM NH4Cl was significantly reduced by 20 mM L-Gln (Fig. 5B) and would be a "predicted response" if the L-Gln-dependent response were to continue through the 6-h experiment. However, there was an actual decline in vacuole formation 2-6 h after the addition of 20 mM L-Gln to cells that were treated with 30 mM NH4Cl. In fact, if the decline in vacuolation were to be extrapolated to the x-axis in Fig. 5B, these data would suggest that, after 10 h of incubation with NH4Cl and L-Gln, vacuolation would be the same as in control cells treated with buffer alone.

Morphological studies show that L-Gln protects RGM1 cells against vacuolation, cell rounding, and detachment in the presence of NH4Cl and MeNH2. RGM1 cells in culture formed a confluent monolayer that was unchanged by incubation with STD buffer for 6 h (Fig. 6A). By 24 h after the addition of STD buffer, some cell death occurred, as demonstrated by cell rounding and loss of attachment to the culture dish (Fig. 6B). Cultures incubated for 6 and 24 h in STD buffer containing 20 mM L-Gln were nearly identical to cultures incubated in STD buffer alone (Fig. 6, C and D). In contrast, RGM1 cells incubated with 30 mM NH4Cl or 3 mM MeNH2 showed significant vacuolation by 6 h (Fig. 6, E and G). By 24 h in NH4Cl or MeNH2, >70 and 60%, respectively, of cells were rounded and/or detached from the culture plate (Fig. 6, F and H). In cultures incubated with NH4Cl or MeNH2 containing L-Gln for 6 h, vacuolation was significantly reduced (Fig. 6, I and K). In cultures incubated with NH4Cl or MeNH2 containing L-Gln for 24 h, cultures were confluent with cell rounding and detachment from the culture dish not significantly different from that of control cultures (Fig. 6, J and L).


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Fig. 6.   Phase-contrast images of RGM1 cells that were incubated with or without L-Gln in the absence or presence of NH4Cl or MeNH2. A and B: RGM1 cells maintained a confluent monolayer in STD buffer for 6 and 24 h, respectively. At 24 h, some dying cells (arrows) were present in the monolayer from control cells. C and D: L-Gln alone resulted in no change in the monolayer of cells for 6 and 24 h, respectively. Like in control cells, some dying cells (arrows) were present in the monolayer at 24 h. E and G: addition of 30 mM NH4Cl or 3 mM MeNH2, respectively, for 6 h resulted in the vacuolation of RGM1 cells (arrowheads). F and H: by 24 h, the number of adherent cells decreased significantly in the presence of 30 mM NH4Cl or 3 mM MeNH2, respectively. Many cells were rounded (arrows), and the few remaining attached cells (arrowheads) had large vacuoles. I and K: L-Gln at 20 mM protected RGM1 cells against NH4Cl- or MeNH2-induced vacuolation, respectively, at 6 h. The number of rounded and detached cells (arrows) with L-Gln was not different from that of control cells in B. J and L: L-Gln at 20 mM protected RGM1 cells against NH4Cl- or MeNH2-induced vacuolation, respectively, and cell death at 24 h. The number of rounded and detached cells (arrows) was not different from that of control cells in B. Bar = 20 µm.

Identification of the mechanism(s) by which L-Gln protects RGM1 cells in the presence of NH4Cl and MeNH2. To determine the mechanism by which L-Gln improves viability and decreases the vacuolation of RGM1 cells, we studied two potential pathways. First, we investigated whether L-Gln inhibits the intracellular accumulation of MeNH2 in RGM1 cells. The intracellular accumulation of MeNH2, and not NH3, was done because radiolabeled [15N]NH3 is not commercially available. In addition, MeNH2 is a weak base with no potential for entry into an intracellular metabolic pathway, so that intracellular reduction in weak base concentration by metabolism is not a factor in the experiment. Second, we determined whether RGM1 cells, like liver or brain cells, utilize the urea cycle and/or glutamine synthetase, with or without L-Gln, as potential NH3 detoxification pathways. NH3 detoxification would reduce the intracellular concentration of NH3 in cells, resulting in less cell death and vacuolation.

L-Gln reduces the intracellular accumulation of MeNH2 in RGM1 cells. When RGM1 cells were incubated with 3 mM MeNH2, containing 0.5 µCi of [14C]MeNH2, accumulation of MeNH2 was 6.28 ± 0.13 µmol · 5 × 105 cells/well (Fig. 7). When RGM1 cells were treated with 3 mM MeNH2 containing L-Gln, [14C]MeNH2 accumulation was reduced in a concentration-dependent manner (Fig. 7). Treatment with L-Gln resulted in a significant reduction in the accumulation of [14C]MeNH2 by 9.8 ± 2.7, 30.0 ± 3.6, and 45.8 ± 3.2% for 0.2, 2.0, and 20 mM L-Gln, respectively, compared with 3 mM MeNH2 alone. Incubation with STD buffer containing 0.5 µCi [14C]MeNH2 and no additional L-Gln resulted in little accumulation of [14C]MeNH2 (0.021 ± 0.001 µmol · 5 × 105 cells/well) or vacuolation (data not shown) in RGM1 cells.


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Fig. 7.   Effect of L-Gln on the intracellular accumulation of radiolabeled MeNH2 ([14C]MeNH2) in RGM1 cells. RGM1 cells were incubated for 3 h with 0.02-20 mM L-Gln in the presence of 3 mM MeNH2 containing 0.5 µCi of [14C]MeNH2, and then the intracellular accumulation of [14C]MeNH2 was measured. The data show that L-Gln significantly decreased the intracellular concentration of [14C]MeNH2 in a concentration-dependent manner. Values are means ± SE. * Significant increase of intracellular [14C]MeNH2 compared with control cells, P < 0.05. dagger  Significant decrease of intracellular [14C]MeNH2 compared with cells treated with 3 mM MeNH2 and no L-Gln, P < 0.05.

The intracellular concentration of a weak base is regulated by the equilibrium between entry (uptake) and extrusion. We showed in Fig. 7 that L-Gln reduced the accumulation of MeNH2 in RGM1 cells. However, these results cannot distinguish between inhibited uptake or facilitated extrusion. To examine the role of L-Gln in facilitated extrusion of weak bases, we used RGM1 cells preloaded with NH4Cl or MeNH2.

Preloading RGM1 cells for 6 h with 30 mM NH4Cl or 3 mM MeNH2 caused a significant increase in vacuolation (195.6 ± 2.8 and 213.6 ± 2.2%, respectively) compared with control cells incubated with STD buffer alone (Fig. 8, A and B, respectively). When NH4Cl or MeNH2 preloaded cells were incubated for 1 h with STD buffer alone, vacuolation decreased by 64.5% in NH4Cl-treated cells and 78.6% in MeNH2-treated cells (Fig. 8, A and B, respectively). This reduction in vacuolation reflects a decrease in the intracellular concentration of NH4Cl or MeNH2 due to diffusion into the culture medium (10), resulting in smaller vacuoles. When the preloaded RGM1 cells were incubated with STD buffer containing 0.2-20 mM L-Gln, vacuolation was significantly reduced in a concentration-dependent manner (Fig. 8). In both NH4Cl and MeNH2 preloaded cells, vacuolation was significantly reduced by 0.2, 2, and 20 mM L-Gln (Fig. 8, A and B, respectively). Substitution of 20 mM mannitol for 20 mM L-Gln, to control for the osmotic effects of L-Gln, resulted in no reduction in the size of vacuoles compared with STD buffer alone (20 mM mannitol is 34.8 ± 5.0% vs. STD buffer of 35.0 ± 1.0%). Thus these results demonstrate that L-Gln stimulates the extrusion of both NH4Cl and MeNH2 from RGM1 cells.


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Fig. 8.   Effect of L-Gln on vacuolation in the NH4Cl- (A) or MeNH2-preloaded cells (B). RGM1 cells were incubated for 6 h with 30 mM NH4Cl (A) or 3 mM MeNH2 (B) to create large vacuoles (preloaded condition). Next, the preloaded cells were incubated for 1 h with STD buffer containing 0-20 mM L-Gln and no NH4Cl or MeNH2. Vacuolation was quantified by the neutral red uptake assay and expressed as means ± SE of 4 wells from 3 different experiments. The results show that vacuolation decreased faster in the presence of L-Gln, suggesting that L-Gln accelerates the extrusion of both NH4Cl and MeNH2. * Significant increase in vacuolation compared with control cells that were not exposed to NH4Cl (A) or MeNH2 (B), P < 0.05. dagger  Significant reduction in vacuolation compared with cells in STD buffer alone, P < 0.05.

Production of urea contributes to L-Gln-induced protection against NH4Cl but not MeNH2 in RGM1 cells. To determine whether gastric epithelial cells are protected from NH3 (but not methylamine) cytotoxicity by utilizing NH3 to form urea, we blocked arginase activity, a key enzyme in the urea synthetic pathway, with nor-NOHA. In RGM1 cells treated with 30 mM NH4Cl, cell viability was reduced significantly in the presence of 0.01-1 mM nor-NOHA (Fig. 9A). In fact, cell viability was reduced to 5.5 ± 0.8% in the presence of 1 mM nor-NOHA, a concentration that did not affect cell viability in the absence of NH4Cl (Fig. 9A). When RGM1 cells were treated with NH4Cl in the presence of 20 mM L-Gln and nor-NOHA, protection induced by L-Gln was abolished (Fig. 9B). In contrast, nor-NOHA (at 1 mM) had no effect on viability in the presence of MeNH2 (Fig. 9C) or on L-Gln-induced protection against MeNH2 (Fig. 9D). RGM1 cells in STD buffer had arginase activity (87.2 ± 3.1 mU/ml) that increased significantly in the presence of NH4Cl (112.9 ± 4.9 mU/ml). nor-NOHA blocked arginase activity in a dose-dependent manner by 39.9, 63.5, and 98.0% at 0.01, 0.1, and 1 mM, respectively, in the presence of NH4Cl, and by 38.3, 71.6, and 100.3% at 0.01, 0.1, and 1 mM, respectively, in the presence of NH4Cl and L-Gln.


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Fig. 9.   Viability of RGM1 cells in the presence of Nomega -hydroxy-nor-L-arginine (nor-NOHA), a potent arginase inhibitor that blocks urea formation from ammonia. RGM1 cells were incubated with 30 mM NH4Cl (A) or 30 mM NH4Cl and 20 mM L-Gln (B) in the presence of 0.01-1 mM nor-NOHA, and cell viability was determined by the crystal violet assay. A: NH4Cl decreased viability that was further reduced when urea production was blocked by nor-NOHA. B: blockade of urea production with nor-NOHA completely reversed the protection in viability provided by 20 mM L-Gln. In contrast, either alone (C) or in combination with L-Gln (D), nor-NOHA had no effect on viability in the presence of 3 mM MeNH2. Values are means ± SE. * Significant decrease in viability compared with control cells, P < 0.05. dagger  Significant increase in viability compared with cells treated with NH4Cl (B) or MeNH2 (D) alone, P < 0.05. # Significant reduction in viability compared with cells treated with NH4Cl alone, P < 0.05. § Significant reduction in viability compared with cells treated with NH4Cl and L-Gln alone (P < 0.05). Note that 1 mM nor-NOHA has no effect on the viability of untreated RGM1 cells (A and C).

L-Glu improves the viability of RGM1 cells in the presence of NH4Cl but does not improve viability by the conversion of L-Glu and NH3 to L-Gln. Treatment of RGM1 cells with L-Glu prevented the reduction in cell viability induced by 30 mM NH4Cl in a concentration-dependent manner (Fig. 10A). In the presence of NH4Cl, significant protection occurred with 0.02, 0.2, 2, and 20 mM L-Glu (Fig. 10A). In fact, 20 mM L-Glu completely (100.3 ± 1.1% of the initial value) protected RGM1 cells that were incubated with 30 mM NH4Cl (Fig. 10A).


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Fig. 10.   Effect of L-glutamate (L-Glu) with or without methionine sulfoximine (MS) on cell viability in the presence of NH4Cl. A: RGM1 cells were incubated with 30 mM NH4Cl with or without 0.02-20 mM L-Glu for 24 h, and the number of viable cells was quantified by the crystal violet assay. The data show that L-Glu protects RGM1 cell viability in the presence of NH4Cl in a dose-dependent manner. B: RGM1 cells were incubated with 30 mM NH4Cl and 20 mM L-Glu in the presence of 0.1-10 mM MS for 24 h, and the number of viable cells was quantified by the crystal violet assay. The data show that there was no difference in viability with NH4Cl, L-Glu, and MS compared with RGM1 cells incubated in NH4Cl and L-Glu alone. Values represent means ± SE of 4 wells from 3 different experiments and are expressed as the percentage of control cells that were incubated with STD buffer alone. * Significant decrease in viability compared with control cells, P < 0.05. dagger  Significant increase in viability compared with cells incubated with 30 mM NH4Cl without L-Glu, P < 0.05.

To determine whether the conversion of L-Glu and NH3 to L-Gln contributes to protection by L-Glu, we blocked this conversion with MS (Fig. 10B), a potent inhibitor of glutamine synthetase activity (52). If the conversion of L-Glu and NH3 to L-Gln is involved in NH3 detoxification, blockade of the pathway with MS would reduce viability in the presence of L-Glu. However, we found that there was no significant difference in survival with 30 mM NH4Cl and 20 mM L-Glu containing 0.1-10 mM MS compared with RGM1 cells incubated with 30 mM NH4Cl and 20 mM L-Glu alone (Fig. 10B). These results demonstrate that RGM1 cells do not convert L-Glu and NH3 to L-Gln to protect against NH3-induced cell death.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The present study shows that apical exposure of gastric surface epithelial (RGM1) cells to NH3 significantly reduces cell viability within 24 h. The mean concentration of NH3 (measured as NH<UP><SUB>4</SUB><SUP>+</SUP></UP>) in the gastric juice of H. pylori-infected patients is from 3.4 to 22.8 mM (13, 23, 24, 27, 32, 34, 35, 46, 50, 58) but may be much higher next to surface epithelial cells, because most H. pylori colonize the adherent mucous layer in vivo. Our study demonstrates that surface epithelial cells possess an active NH3 detoxification pathway that provides some level of protection against NH3-induced cytotoxicity. When the luminal NH3 concentration increases, our results demonstrate that L-Gln facilitates NH3 detoxification and improves cell survival.

Although our work and other reports conclude that a high concentration of NH3 is cytotoxic to cells in vitro, several findings that address the role of NH3 or NH4Cl in vivo are not consistent with these results. For instance, intragastric administration of urea (6%)/urease (100 units) or concentrations of NH4Cl up to 3% (560 mM, pH 4.8 or pH 8.0) for 1 h in the rat in vivo caused no damage to epithelial or other cells in the stomach (44). Tsujii et al. (49) showed that 187.5-250 mM of NH3 decreased oxygen consumption, energy charge, and the survival of isolated mucosal cells, but the same concentrations of NH4Cl at pH 7.4 in vivo did not (48). From these results, it is tempting to conclude that NH3 plays no role in gastric epithelial injury under physiological conditions. However, we propose that the intact mucosa must be incubated with NH4Cl for many hours before cell death is evident. To support this contention, the present study shows that RGM1 cells must be incubated with a high concentration of NH4Cl (from the luminal surface) for at least 12 h before NH3 initiates cell death. Although it is not known why it takes NH4Cl so long to kill cells from the luminal surface, parietal and chief cells in gastric glands have a permeability barrier to NH3 (3) that may also occur in surface epithelial cells. Thus, with the slow paracellular flux of a weak base that occurs in gastric tissues (18), NH3 may move from the lumen to the basolateral compartment and kill cells after entry from the basolateral surface. Alternatively, our results suggest that gastric surface cells are protected from NH3 by the metabolic elimination of NH3 to urea, via arginase activity. The rate at which arginase metabolizes NH3 may determine ultimate cell fate.

That gastric surface epithelial cells can metabolize NH3 to urea is a concept demonstrated, for the first time, in the present study. The use of an intracellular detoxification pathway in RGM1 cells was suggested in our study because cell death occurs with ~10-fold higher concentrations of NH4Cl than MeNH2, a primary amine weak base with similar properties to NH4Cl. In addition, NH4Cl and MeNH2 should cause the same degree of vacuolation due to similar properties as weak bases, but it requires 10-fold higher concentrations of NH4Cl to cause the same degree of vacuolation as with MeNH2. These results are even more significant if the weak base concentration is taken into consideration, where the NH3 concentration in 30 mM NH4Cl (pH 7.4) is 0.44 and the MeNH2 (weak base) concentration in 3 mM MeNH2 (pH 7.4) is 0.0017 mM. Because NH3 can enter intracellular metabolic pathways and MeNH2 cannot, we suggest that the metabolism of NH3 to urea must lower the effective concentration of NH3 in cells, causing less vacuolation and cytotoxicity.

The results presented here demonstrate that gastric RGM1 cells have arginase activity that is inhibitable by nor-NOHA, a selective arginase inhibitor (15, 45). Arginase, an enzyme that catalyzes the hydrolysis of L-arginine to urea and L-ornithine, is a key enzyme in NH3 detoxification via the urea cycle (22). Arginase exists in two isoforms. Arginase I, a cytosolic enzyme, is expressed exclusively in liver as a component of the urea cycle (16, 22, 38). In contrast, arginase II is a mitochondrial enzyme that is expressed in many tissues, including the stomach (16, 38). Compared with that in the intestine and liver, arginase activity is extremely low in the stomach, and the glandular stomach (as a whole) produces very little urea (19). We also found this to be true in our study, because urea production by RGM1 cells, even in the presence of NH4Cl, was below detectable levels using the commercial urea assay kit (Sigma). This finding was not surprising, because the urea kit measures between 1,650 and 3,300 µM of urea (37), a concentration that can easily be measured in blood, urine, and liver, a tissue that produces urea at a rate of 158 µmol · min-1 · g-1 of tissue (19). Because the RGM1 cells in our study produced urea at a rate of 1 nmol · min-1 · g-1 of cells, it would take 27.5 h to generate enough urea to measure using the commercial urea assay kit, which would not be possible in the presence of NH4Cl. Thus it was necessary to use a radioactive procedure, developed by Ruegg and Russell (39), to measure urea that is produced (by arginase activity) by the conversion of L-[guanido-14C]arginine to [14C]urea. Byrne et al. (4) showed that arginase activity in the stomach is found predominantly in a low-density fraction that contains 84 ± 2% parietal cells. Our study shows that arginase II activity must be present in surface epithelial cells and that arginase II activity may increase in the presence of L-Gln or other amino acids that regulate urea cycle activity. In the liver, there are five urea cycle enzymes that contribute to the synthesis of urea for NH3 detoxification (38). Because no other urea cycle intermediates have been described in gastric tissues, further studies will be necessary to complete our understanding of the active components of the urea cycle in gastric mucosal cells.

In this study, we show that L-Gln protects RGM1 cells against NH4Cl-induced cell death. It is noteworthy that protection was observed even at very low concentrations of L-Gln (0.2 mM), as shown in Fig. 3A. Because the plasma level of L-Gln is 0.5-0.8 mM (52), physiological concentrations of L-Gln may protect against the cytotoxic effects of NH3 in daily life. In general, L-Gln is involved in a wide variety of metabolic processes, such as the synthesis of proteins and nucleotides, and in energy metabolism (41, 59). L-Gln plays an essential role in intestinal mucosal protection in many animal models of critical illness, including burns, trauma, obstruction, radiation damage, cytotoxic chemotherapy, and sepsis (5, 11, 28). Cellular ATP levels are maintained in the presence of L-Gln, which protects mitochondria from damage and partially protects alpha -ketoglutarate dehydrogenase activity in the TCA cycle (1). L-Gln also induces heat shock protein expression to protect cells against injury (54, 56). Furthermore, L-Gln reduces the expression of proinflammatory cytokines (55), which may reduce inflammatory cell-induced mucosal damage in vivo. Although it is not known how L-Gln protects against the cytotoxic effects of NH4Cl, our study suggests that it inhibits the uptake and/or facilitates the extrusion of NH3 from cells and increases cellular metabolism of NH3 via arginase. Although L-Glu does not protect RGM1 by NH3 detoxification via glutamine synthetase activity per se, it may act to increase cellular metabolism and ATP production, inhibit uptake and/or increase extrusion, or facilitate some other process that facilitates protection against NH3.

The results presented here clearly show that L-Gln protects cells differently in the presence of NH4Cl than in the presence of MeNH2. Our data suggest that L-Gln, in some way, accelerates cellular NH3 metabolism to reduce both the intracellular concentration of NH3 and NH3 cytotoxicity. Our data in Fig. 5B suggest that accelerated NH3 metabolism occurs 2 h after the addition of NH4Cl and L-Gln to RGM1 cells. In addition, blockade of urea production with nor-NOHA completely reversed the protective effect of L-Gln (against NH4Cl-induced death), suggesting that NH3 detoxification via arginase activity is paramount in protecting RGM1 cells against NH4Cl-induced cell death. In contrast, even though L-Gln completely protects against the cytotoxic effects of MeNH2, our data clearly show that urea cycle activity is not involved in L-Gln protection against MeNH2. This is because nor-NOHA did not reverse the protective effect of L-Gln against MeNH2-induced cell death. If exposure to L-Gln protected RGM1 cells solely by extrusion of weak base, increased expression of heat shock proteins, and/or by increased cellular ATP production, it is likely that protection would be similar with both NH4Cl and MeNH2. Thus the differential effect of weak bases on viability in RGM1 cells may lend important insights into the mechanism by which L-Gln protects against injury in gastric and other tissues.

In summary, we demonstrate that L-Gln and L-Glu protect gastric epithelial RGM1 cells against NH4Cl-induced cell death. Because L-Gln alimentation is used routinely in human patients (2), it is possible that L-Gln alone or in combination with L-Glu would be effective as a therapeutic treatment for gastric epithelial damage induced by NH3 during H. pylori infection.


    ACKNOWLEDGEMENTS

The authors thank Sarah W. Morrison for technical help with cell culture and Marianne Smith and Dr. Kimihito Tashima for critical evaluation of the manuscript. We are especially grateful to Dr. Koji Takeuchi for critical reading of the manuscript and for helpful discussions concerning the results of this study.


    FOOTNOTES

This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants R01-DK-15681 (to S. J. Hagen) and DK-34854 (to Harvard Digestive Diseases Center).

Address for reprint requests and other correspondence: S. J. Hagen, Dept. of Surgery, Dana 805, Beth Israel Deaconess Medical Center, 330 Brookline Ave., Boston MA 02215 (E-mail: shagen{at}caregroup.harvard.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. Section 1734 solely to indicate this fact.

August 28, 2002;10.1152/ajpgi.00235.2002

Received 17 June 2002; accepted in final form 20 August 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Ahmad, S, White CW, Chang LY, Schneider BK, and Allen CB. Glutamine protects mitochondrial structure and function in oxygen toxicitity. Am J Physiol Lung Cell Mol Physiol 280: L779-L791, 2001[Abstract/Free Full Text].

2.   Boelens, PG, Nijveldt RJ, Houdijk APJ, Meijer S, and van Leeuwen PAM Glutamine alimentation in catabolic states. J Nutr 131: 2569S-2577S, 2001[Abstract/Free Full Text].

3.   Boron, WF, Waisbren SJ, Bodlin IM, and Geibel JP. Unique permeability barrier of the apical surface of parietal and chief cells in isolated perfused gastric glands. J Exp Biol 196: 347-360, 1994[Abstract/Free Full Text].

4.   Byrne, CR, Price KJ, Williams JM, Brown JF, Hanson PJ, and Whittle BJR Nitric oxide synthase and arginase in cells isolated from the rat gastric mucosa. Biochim Biophys Acta 1356: 131-139, 1997[ISI][Medline].

5.   Chang, TM, Lu RH, and Tsai LM. Glutamine ameliorates mechanical obstruction-induced intestinal injury. J Surg Res 95: 133-140, 2001[ISI][Medline].

6.   Cook, TH, Jansen A, Lewis S, Largen P, O'Donnell M, Reaveley D, and Cattell V. Arginine metabolism in experimental glomerulonephritis: interaction between nitric oxide synthase and arginase. Am J Physiol Renal Fluid Electrolyte Physiol 267: F646-F653, 1994[Abstract/Free Full Text].

7.   Cover, TL, Halter SA, and Blaser MJ. Characterization of HeLa cell vacuoles induced by Helicobacter pylori broth culture supernatant. Hum Pathol 23: 1004-1010, 1992[ISI][Medline].

8.   Cover, TL, Puryear W, Perez-Perez GI, and Blaser MJ. Effect of urease on HeLa cell vacuolation induced by Helicobacter pylori cytotoxin. Infect Immun 59: 1264-1270, 1991[ISI][Medline].

9.   Daikhin, Y, and Yudoff M. Compartmentation of brain glutamate metabolism in neurons and glia. J Nutr 131: 1026S-1031S, 2001.

10.   DeDuve, C, DeBarsy T, Poole B, Trouet A, Tulkens P, and VanHoof F. Lysosomotropic agents. Biochem Pharmacol 23: 2495-2531, 1974[ISI][Medline].

11.   Fink, MP. Gastrointestinal mucosal injury in experimental models of shock, trauma, and sepsis. Crit Care Med 19: 627-641, 1991[ISI][Medline].

12.   Fox, JG, Lee A, Otto G, Taylor NS, and Murphy J. Helicobacter felis gastritis in gnotobiotic rats: an animal model of Helicobacter pylori gastritis. Infect Immun 59: 785-791, 1991[ISI][Medline].

13.   Furuta, T, Baba S, Takashima M, Futami H, Arai H, Kajimura M, Hanai H, and Kaneko E. Effect of Helicobacter pylori infection on gastric juice pH. Scand J Gastroenterol 33: 357-363, 1998[ISI][Medline].

14.   Gillen, D, Wirz AA, Neithercut WD, Ardill JES, and McColl KEL Helicobacter pylori infection potentiates the inhibition of gastric acid secretion by omeprazole. Gut 44: 468-475, 1999[Abstract/Free Full Text].

15.   Gobert, AP, Daulouede S, Lepoivre M, Boucher JL, Bouteille B, Buguet A, Cespuglio R, Beyret B, and Vincendeau P. L-Arginine availability modulates local nitric oxide production and parasite killing in experimental trypanosomiasis. Infect Immun 68: 4653-4657, 2000[Abstract/Free Full Text].

16.   Gotoh, T, Araki M, and Mori M. Chromosomal localization of the human arginase II gene and tissue distribution of its mRNA. Biochem Biophys Res Commun 233: 487-491, 1997[ISI][Medline].

17.   Hagen, SJ, Takahashi S, and Jansons R. Role of vacuolation in the death of gastric epithelial cells. Am J Physiol Cell Physiol 272: C48-C58, 1997[Abstract/Free Full Text].

18.   Hagen, SJ, Wu H, and Morrison SW. NH4Cl inhibition of acid secretion: possible involvement of an apical K+-channel in bullfrog oxyntic cells. Am J Physiol Gastrointest Liver Physiol 279: G400-G410, 2000[Abstract/Free Full Text].

19.   Harri, MP, and Hartiala K. Arginase activity in rat small intestinal mucosa. Acta Physiol Scand 89: 126-128, 1973[ISI][Medline].

20.   Häussinger, D. Liver glutamine metabolism. JPEN J Parenter Enteral Nutr 14: 56S-62S, 1990[Medline].

21.   Igarashi, M, Kitada Y, Yoshiyama H, Takagi A, Miwa T, and Koga Y. Ammonia as an accelerator of tumor necrosis factor alpha-induced apoptosis of gastric epithelial cells in Helicobacter pylori infection. Infect Immun 69: 816-821, 2001[Abstract/Free Full Text].

22.   Jenkinson, CP, Grody WW, and Cederbaum SD. Comparative properties of arginases. Comp Biochem Physiol 114B: 107-132, 1996[ISI].

23.   Kaneko, H, Nakada K, Mitsuma T, Uchida K, Furusawa A, Maeda Y, and Morise K. Helicobacter pylori infection induces a decrease in immunoreactive-somatostatin concentrations of human stomach. Dig Dis Sci 37: 409-416, 1992[ISI][Medline].

24.   Kearney, DJ, Ritchie K, and Peacock JS. Gastric-juice ammonia assay for diagnosis of Helicobacter pylori infection and the relationship of ammonia concentration to gastritic severity. Am J Gastroenterol 95: 3399-3403, 2000[ISI][Medline].

25.   Keisari, Y. A colorimetric microtiter assay for the quantitation of cytokine activity on adherent cells in tissue culture. J Immunol Methods 146: 151-161, 1992.

26.   Khoda, K, Tanaka K, Aiba Y, Yasuda M, Miwa T, and Koga Y. Role of apoptosis induced by Helicobacter pylori infection in the development of duodenal ulcer. Gut 44: 456-462, 1999[Abstract/Free Full Text].

27.   Kim, H, Park C, Jang WI, Lee KH, Kwon SO, Robey-Cafferty SS, Ro JY, and Lee YB. The gastric juice urea and ammonia levels in patients with Campylobacter pylori. Am J Clin Pathol 94: 187-191, 1990[ISI][Medline].

28.   Klimberg, VS, and Souba WW. The importance of intestinal glutamine metabolism in maintaining a healthy gastrointestinal tract and supporting the body's response to injury and illness. Surg Annu 22: 61-76, 1990[Medline].

29.   Kobayashi, I, Kawano S, Tsuji S, Matsui H, Nakama A, Sawaoka H, Masuda E, Takei Y, Nagano K, Fusamoto H, Ohno T, Fukutomi H, and Kamada T. RGM1, a cell line derived from normal gastric mucosa of the rat. In Vitro Cell Dev Biol Anim 32: 259-261, 1996[ISI][Medline].

30.   Lee, A, Fox JG, Otto G, and Murphy J. A small animal model of human Helicobacter pylori active chronic gastritis. Gastroenterology 99: 1315-1323, 1990[ISI][Medline].

31.   Matsui, T, Yoshizumi Y, Sakai T, Nakamura E, Aoike A, and Kwai E. Ammonia inhibits proliferation and cell cycle progression at S-phase in human gastric cells. Dig Dis Sci 42: 1394-1399, 1997[ISI][Medline].

32.   Miyaji, H, Ito S, Azuma T, Ito Y, Yamazaki Y, Ohtaki Y, Sato F, Hirai M, Kuriyama M, and Kohli Y. Effects of Helicobacter pylori eradication therapy on hyperammonaemia in patients with liver cirrhosis. Gut 40: 726-730, 1997[Abstract].

33.   Mohammadi, M, Redline R, Nedrud J, and Czinn S. Role of the host in pathogenesis of Helicobacter-associated gastritis. H. felis infection of inbred and congenic mouse strains. Infect Immun 64: 238-245, 1996[Abstract].

34.   Mokuolu, AO, Sigal SH, and Lieber CS. Gastric juice urease activity as a diagnostic test for Helicobacter pylori infection. Am J Gastroenterol 92: 644-648, 1997[ISI][Medline].

35.   Nakamura, M, Haruma K, Kamada T, Mihara M, Yoshihara M, Sumioka M, Fukuhara T, and Chayama K. Cigarette smoking promotes atrophic gastritis in Helicobacter pylori-positive subjects. Dig Dis Sci 47: 675-681, 2002[ISI][Medline].

36.   Ogura, K, Maeda S, Nakao M, Watanabe T, Tada M, Kyutoku T, Yoshida H, Shiratori Y, and Omata M. Virulence factors of Helicobacter pylori responsible for gastric diseases in Mongolian gerbils. J Exp Med 192: 1601-1609, 2000[Abstract/Free Full Text].

37.   Ormsby, AA. A direct colorimetric method for the determination of urea in blood and urine. J Biol Chem 146: 595-604, 1942.

38.   Osaki, M, Gotoh T, Nagasaki A, Minyanaka K, Takeya M, Fujiyama S, Tomita K, and Mori M. Expression of arginase II and related enzymes in the rat small intestine and kidney. Biochem J 125: 586-593, 1999.

39.   Ruegg, UT, and Russell AS. A rapid and sensitive assay for arginase. Anal Biochem 102: 206-212, 1980[ISI][Medline].

40.   Russell, AS, and Ruegg UT. Arginase production by peritoneal macrophages: a new assay. J Immunol Methods 32: 375-382, 1980[ISI][Medline].

41.   Souba, WW, Herskowitz K, Salloum RM, Chen MK, and Austgen TR. Gut glutamine metabolism. JPEN J Parenter Enteral Nutr 14: 45S-50S, 1990[Medline].

42.   Suzuki, H, Miyazawa M, Nagahashi S, Mori M, Seto K, Kai A, Suzuki M, Miura S, and Ishii H. Attenuated apoptosis in H. pylori-colonized gastric mucosa of Mongolian gerbils in comparison with mice. Dig Dis Sci 47: 90-99, 2002[ISI][Medline].

43.   Suzuki, H, Yanaka A, and Muto H. Luminal ammonia retards restitution of guinea pig injured gastric mucosa in vitro. Am J Physiol Gastrointest Liver Physiol 279: G107-G117, 2000[Abstract/Free Full Text].

44.   Takeuchi, K, Ohuchi T, Harada H, and Okabe S. Irritant and protective action of urea-urease ammonia in rat gastric mucosa: different effects of ammonia and ammonium ion. Dig Dis Sci 40: 274-281, 1995[ISI][Medline].

45.   Tenu, JP, Lepoivre M, Moali C, Brollo M, Manusy D, and Boucher JL. Effects of the new arginase inhibitor Nomega -hydroxy-nor-L-arginine on NO synthase activity in murine macrophages. Nitric Oxide 3: 427-438, 1999[ISI][Medline].

46.   Tokushima, H, Tamura H, Murakawa M, Matsumura O, Itakura Y, Itoyama S, Mitari T, and Isoda K. Eradication of Helicobacter pylori restores elevation of serum gastrin concentrations in patients with end-stage renal disease. Intern Med 37: 435-439, 1998[ISI][Medline].

47.   Triebling, AT, Korsten MA, Dlugosz JW, Paronetto F, and Lieber CS. Severity of Helicobacter-induced gastric injury correlates with gastric juice ammonia. Dig Dis Sci 36: 1089-1096, 1991[ISI][Medline].

48.   Tsujii, M, Kawano S, Tsuji S, Fusamoto H, Kamada T, and Sato N. Mechanism of gastric mucosal damage induced by ammonia. Gastroenterology 102: 1881-1888, 1992[ISI][Medline].

49.   Tsujii, M, Kawano S, Tsuji S, Ito T, Nagano K, Sasaki Y, Hayashi N, Fusamoto H, and Kamada T. Cell kinetics of mucosal atrophy in rat stomach induced by long-term administration of ammonia. Gastroenterology 104: 796-801, 1993[ISI][Medline].

50.   Verdu, EF, Armstrong D, Sabovcikova L, Idström JP, Cederberg C, Blum AL, and Bericik P. High concentrations of ammonia, but not volatile amines, in gastric juice of subjects with Helicobacter pylori infection. Helicobacter 3: 97-102, 1998[ISI][Medline].

51.   Von Herbay, A, and Rudi J. Role of apoptosis in gastric epithelial turnover. Microsc Res Tech 48: 303-311, 2000[ISI][Medline].

52.   Weiss, MD, DeMarco V, Strauss DM, Samuelson DA, Lane ME, and Neu J. Glutamine synthetase: a key enzyme for intestinal epithelial differentiation. JPEN J Parenter Enteral Nutr 23: 140-146, 1999[Abstract].

53.   Wirth, HP, Beins MH, Yang M, Tham KT, and Blaser MJ. Experimental infection of Mongolian gerbils with wild-type and mutant Helicobacter pylori strains. Infect Immun 66: 4856-4866, 1998[Abstract/Free Full Text].

54.   Wischmeyer, PE, Kahana M, Wolfson R, Ren H, Musch MM, and Chang EB. Glutamine induces heat shock protein and protects against endotoxin shock in the rat. J Appl Physiol 90: 2403-2410, 2001[Abstract/Free Full Text].

55.   Wischmeyer, PE, Khana M, Wolfson R, Ren H, Musch MM, and Chang EB. Glutamine reduces cytokine release, organ damage, and mortality in a rat model of endotoxemia. Shock 16: 398-402, 2001[ISI][Medline].

56.   Wischmeyer, PE, Musch MM, Madonna MB, Thisted R, and Chang EB. Glutamine protects intestinal epithelial cells: role of inducible HSP70. Am J Physiol Gastrointest Liver Physiol 272: G879-G884, 1997[Abstract/Free Full Text].

57.   Yanaka, A, Muto H, Ito S, and Silen W. Effects of ammonium ion and ammonia on function and morphology of in vitro frog gastric mucosa. Am J Physiol Gastrointest Liver Physiol 265: G277-G288, 1993[Abstract/Free Full Text].

58.   Yang, DH, Bom HS, Joo YE, Choi SK, Rew JS, and Yoon CM. Gastric juice ammonia as CLO test for diagnosis of Helicobacter pylori infection. Dig Dis Sci 40: 1083-1086, 1995[ISI][Medline].

59.   Young, VR, and Ajami AM. Glutamine: the emperor and his clothes? J Nutr 131: 2449S-2459S, 2001[Abstract/Free Full Text].


Am J Physiol Gastrointest Liver Physiol 283(6):G1264-G1275
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