Department of Surgery, Beth Israel Deaconess Medical Center, Boston, Massachusetts 02215
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ABSTRACT |
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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
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INTRODUCTION |
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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- or interferon-
(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.
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MATERIALS AND METHODS |
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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
-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.
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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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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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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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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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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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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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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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DISCUSSION |
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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
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 · min1 · 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
-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.
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ACKNOWLEDGEMENTS |
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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.
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FOOTNOTES |
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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.
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