Department of Surgery, Medical College of Virginia Campus of Virginia Commonwealth University, Richmond, Virginia 23298
Submitted 7 August 2002 ; accepted in final form 16 May 2003
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ABSTRACT |
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necrosis; caspases; cell culture; prostaglandins
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THE POTENTIAL TOXIC EFFECT of bile salts on gastric mucosal cells is well established both clinically and experimentally (6, 8, 13, 30). In fact, duodenogastric reflux is thought to play a major role in the pathogenesis of human gastric ulceration, and bile seems to be a key participant in this process (8, 30). In previous studies (20, 21) using a human gastric mucosal cell line, known as AGS cells, we have shown that deoxycholate (DC), in concentrations ranging from 200 to 500 µM, consistently injures gastric cells, ultimately leading to irreversible damage and death if exposure is prolonged. The clinical relevance of these findings is obvious when it is realized that DC concentrations approaching 370 µM occur in stomach remnants after distal gastric resection (7).
The precise mechanism of cellular death induced by DC and other bile salts remains controversial. A necrotic pathway has been implicated in many studies evaluating effects of DC on gastric cells (9, 20, 21), but the possibility of apoptosis has also been proposed (26). In hepatocytes, apoptosis appears to be the major mechanism of cell death in response to DC exposure (1618). This study was undertaken to clarify the relative roles of apoptosis vs. necrosis in mediating cellular death induced by DC in gastric mucosal cells and to define the conditions under which a particular pathway is more likely to supervene. AGS cells were used to accomplish these goals as we have employed in our studies previously.
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MATERIALS AND METHODS |
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A human gastric epithelial cell line (AGS), purchased from American Type Culture Collection (Manassas, VA), was used in these experiments. The cells were seeded in 75-cm2 culture flasks (Corning) and maintained in Ham's F-12 (Cellgrow) culture medium supplemented with 10% fetal bovine serum, 100 µg/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml amphotericin B. The cells were grown at 37°C in a humidified atmosphere of 5% CO2. Experiments were performed in 3.5-cm2 dishes or in 96-well tissue culture plates (Costar) at 8090% confluence. Before DC treatment, medium was aspirated and cells were washed with PBS. Experiments were performed under two different conditions: 1) in Ham's F-12 (without serum, growth arrested) or 2) in HBSS (Cellgrow) plus 10 mM HEPES consisting of (in mM) 137 NaCl, 5.7 NaHCO3, 5.3 KCl, 1.26 CaCl2, and 0.8 MgSO4. The cells were treated with DC at a dose range of 50500 µM.
Chemicals and Reagents
The prostaglandin analog 16,16 dimethyl PGE2 (Sigma; dmPGE2) was maintained at -20°C as a stock solution of 1 mg/ml in ethanol. Deoxycholic acid (the sodium salt; Sigma), 10 mM stock solution in PBS, was incubated in water bath at 37°C for 2030 min before each use. Other reagents and chemicals used in our studies included 4',6-diamidino-2 phenylindole (DAPI; Roche), propidium iodide (PI; Molecular Probes), caspase-3 inhibitor IV (Ac-DMQD-CHO, Calbiochem), caspase-6 inhibitor I (Z-VEID-FMK, Calbiochem), and caspase-9 inhibitor I (Z-LEHD-FMK, Calbiochem).
Measurement of Cellular Injury (Cellular Cytotoxicity)
Cellular injury was quantitated by measuring lactate dehydrogenase (LDH) release (5). LDH content was determined by using an LDH-Cytotoxicity Assay Kit (BioVision), which is based on a coupled enzymatic reaction that results in the conversion of a tetrazolium salt into a red formazan product. The increase in the amount of formazan produced in culture supernatant directly correlates with the increase in the number of lysed cells. The formazan was quantified spectrophotometrically by measuring its absorbance at 500 nm. Cytotoxicity in experimental samples measured as %LDH release was compared with cells treated with 1% Triton X-100.
Isolation and Measurement of Fragmented DNA
After treatment, cells were harvested and genomic DNA was isolated using a Suicide-Track DNA ladder isolation kit (Oncogene). Fragmented DNA was resolved on a 1.5% agarose-1x Tris-Acetate-EDTA, visualized by ethidium bromide staining and examined under a ultraviolet light. For detection of histone-associated DNA fragments, a Cell Death Detection ELISA kit (Roche Molecular Biochemicals) was used according to the manufacturer's instructions adapted for suspension cultures (12).
Determination of Apoptotic Cells
Cellular DNA was stained with PI or DAPI and examined under fluorescence microscopy. Apoptotic cells were examined by changes in their nuclear morphology, including shrinkage, condensation and margination, and fragmentation of chromatin. Terminal deoxynucleotidyl transferase (TdT)-mediated dUTP deoxyuridine triphosphate nick-end-labeling (TUNEL) assay was used to determine the yield of apoptotic cells (ApoAlert DNA Fragmentation Assay kit, Clontech), according to the manufacturer's recommendations (10). In brief, TUNEL staining was performed in formaldehyde-fixed AGS cells permeabilized with 0.2% Triton X-100. Cells were incubated with TdT buffer containing fluorescein-dUTP and TdT enzyme at 37°C for 1 h in a humid chamber. Incorporation of fluorescein-dUTP at the free 3'-hydroxyl ends of fragmented DNA was terminated by washing with 2x SSC (0.3 M NaCl, 30 mM sodium citrate). The number of TUNEL-positive cells, expressed as percentage of total cells, was counted using a fluorescence microscope (Nikon) equipped with FITC filters.
Caspase Activity Assay
Caspase activation was determined from cytosol of AGS cells treated with DC for the indicated times. The catalytic activities of caspase-2, caspase-3, caspase-6, and caspase-8 were measured using the Apo Target Caspase colorimetric kit (Biosource), which is based on spectrophotometric detection of the chromophore p-nitroanilide, after cleavage from the labeled synthetic caspase substrate (39). Briefly, after the respective treatment cells were collected and spun down, then they were washed with ice-cold PBS and lysed (50 mM HEPES, 1 mM DTT, 0.1 mM EDTA, 10% glycerol, and 0.1% CHAPS, pH 7.4), nuclei were removed (12,000 g at 4°C for 15 min), and the supernatants (cytosol extracts) were frozen and stored at -70°C until usage. Proteolytic reaction was carried out in reaction buffer (50 mM HEPES pH 7.4, 100 mM NaCl, 0.1% CHAPS, 10 mM DTT, 0.1 mM EDTA, 10% glycerol) containing cytosolic protein extract and freshly prepared colorimetric substrate (VDVAD-pNA for caspase-2; DEVD-pNA for caspase-3; VEID-pNA for caspase-6 and IETD-pNA for caspase-8) at 37°C for 1 h. The absorbance at 405 nm was quantified and normalized for protein content. Enzyme activity was expressed as absorbance at 405 nm/mg of protein.
Protein Concentration Assay
Protein concentration was determined by the Bradford method using the Bio-Rad protein assay reagent and BSA as a standard (3).
Statistical Analysis
All data represent at least three independent experiments and are expressed as the means ± SE. Statistical analysis was performed by a paired Student's t-test, with significant differences determined at P < 0.05. For figure preparation, agarose gels were photographed with Gel DOC 1000 (Bio Rad) and PowerPoint software was used for image creation.
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RESULTS |
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Necrosis as a dominant mechanism of cell death. In this study, we investigated the effect of DC on cytotoxicity of AGS cells in two different media conditions. Incubation of human gastric cells (AGS) with DC in HBSS produced a dose- and time-dependent increase in cell cytotoxicity measured as LDH release. The results presented in Figs. 1, A and B, and 2 show that DC concentrations from 50 to 200 µM had no effect on LDH release for up to 2 h, but longer exposure for DC induced a significant increase in LDH release. The most damaging effect was observed with 300 µM DC, with 20% cytotoxicity demonstrated at 30 min and 90% noted at 4 h of treatment. Such response to DC treatment was observed when AGS cells were incubated with DC in buffered HBSS only. Because the toxicity with 300 µM DC was so profound, higher concentrations of DC were not tested.
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Surprisingly, when cells were treated with DC in Ham's F-12 medium, concentrations of DC ranging from 50 to 300 µM demonstrated no evidence of cytotoxicity by the LDH assay for cellular exposure times as long as 8 h (see Fig. 1, C and D). Only at 24 h of exposure did even high concentrations of DC (i.e., >50 µM) demonstrate some degree of cytotoxic damage.
Apoptosis as a dominant mechanism of cell death. To determine whether apoptosis is activated as a result of DC treatment, cells were treated with 100 and 300 µM DC in Ham's F-12. Cells were harvested for the indicated times, spun, and DNA was isolated from cell pellets and from supernatants. Figure 3, A and B, shows an agarose gel electrophoresis of DNA with the characteristic laddering pattern of fragmented DNA. Each band represents polynucleosomal DNA in its typical multiple 200-bp fragments. As shown in Fig. 3B, 300 µM DC caused a rapid increase ( 50%) in the rate of DNA fragmentation over the basal value, which was sustained over a period of 112 h. For 100 µM DC (Fig. 3A), longer time periods (412 h) were required to initiate
1020% apoptosis. When cells were exposed to concentrations of 100300 µM DC for 24 h, clear evidence of cytotoxicity (by LDH assay and DNA gel electrophoresis, Figs. 1, C and D, and 3, A and B, bottom, respectively) was present, indicating necrosis as well.
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Because the morphological changes in the cells undergoing apoptosis show characteristic nuclear condensation and formation of apoptotic bodies, we used two DNA-binding dyes to examine nuclei. PI- and DAPI-stained cells examined under the fluorescent light indicated fragmented nuclei compared with unfragmented ones in untreated cells (data not shown). Another indicator of apoptosis, internucleosomal cleavage of DNA, was determined and quantified by counting cells stained by the TUNEL method as described in MATERIALS AND METHODS. We found a concentration-dependent increase in TUNEL-positive cells exposed to DC (Fig. 4). Concentrations ranging from 100 to 300 µM caused significant increases in the percentage of apoptotic cells, from 4.5 ± 2.3 to 45.2 ± 3.6, respectively. Findings with 400500 µM DC were not different from those with 300 µM DC.
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We also assessed histone-associated DNA using an ELISA assay. In the early stage of apoptosis, the endogenous endonucleases cleave double-stranded DNA at the most accessible internucleosomal linker regions, generating mono- and oligonucleosomes (4, 29). In contrast, the DNA of the nucleosomes is tightly complexed with the core histones H2A, H2B, H3, and H4 and therefore is protected from cleavage by endonucleases (4, 29). ELISA analysis showed time- and concentration-dependent responses in the enrichment of histone-associated DNA fragments (see Fig. 5, A and B). A concentration of 100 µM DC mediated a small but detectable and proportional increase in histone-DNA formation over a period of 8 h. However, a higher DC concentration (300 µM) induced a rapid and 50-fold increase in histone-DNA formation, compared with control levels, after 13 h of DC-induced apoptosis. Longer times of exposure to 300 µM DC showed decreases in histone-DNA formation, which indicates the stage when nucleosomal histones and DNA are no longer tightly complexed.
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Acidification and apoptosis. An additional series of experiments was conducted to determine the effect of acidification on apoptosis. Studies were performed in identical fashion to those described except that the pH of the cellular bathing solution was altered by adding HCl. A range of pHs were evaluated from pH 3 to 7.4. Control studies assessed the effect of pH change alone, whereas experimental studies evaluated the combination of acid and DC. Apoptosis in these studies was determined by DNA laddering.
Acidification by itself had no effects on AGS cells. No evidence of DNA laddering was noted at either 1 or 4 h regardless of the pH range tested (i.e., pH 36, and 7.4; data not shown). When DC (300 µM) was added to the bathing solution, the resultant response was pH dependent. At neutral pH (i.e., 7.4), DC initiated DNA laddering at both 1 and 4 h, with the response being more pronounced at the latter time period. With acidification, evidence of DNA laddering could no longer be demonstrated. In fact, with prolonged acidification (4 h), DC in acid medium at pH 4 actually induced clear evidence of necrosis in contrast to apoptosis when the pH was 7.4. At 1 h of acidification at pH 4, this necrotic response was present but not near as pronounced as at 4 h. These results indicate that acidification shifts the pattern of cell death from apoptosis to necrosis when AGS cells are exposed to DC. These findings are shown in Fig. 6.
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Effect of DC on Caspase Activity
Caspases that are involved in the execution of apoptosis exist in living cells as inactive zymogens that become activated through intracellular caspase cascades. There are two caspase cascades (1, 14). One is initiated by the activation of cell-surface death receptors, such as Fas and tissue necrosis factor, leading to caspase-8 activation, which, in turn, cleaves and activates downstream caspases such as caspase-3, -6, and -7. In this pathway, mitochondrial signaling seems to be unnecessary but may contribute to apoptosis at a later time as an amplification mechanism (24, 34, 43). The second general apoptotic pathway involves cytochrome c release from mitochondria, which promotes the activation of caspase-9 through APAF-1 (1, 14, 23, 25, 33, 37, 45). Recent observations suggest that bile salt-induced apoptosis in hepatocytes is mediated by the Fas-receptor signaling pathway and caspase-8 activation (11). There is also evidence that bile salts can directly mediate mitochondrial permeability transition in isolated mitochondria from hepatocytes (2, 32) and from the HT-29 cell line (36). Enhanced activation of caspase-8 during apoptosis after mitochondrial dysfunction has been reported in specific cell types (36). In addition, release of the zymogen form of caspase-2 from mitochondria and caspase-2 expression is reported in HT-29 colon carcinoma cells treated with DC (36, 40). Recent findings also suggest that caspase-2 may be a key initiator of the mitochondrial pathway of apoptosis (23). We therefore analyzed the activation of caspase-2, -3, -6, and -8 in the cytoplasm of DC-treated AGS cells by colorimetric assays. Caspase-3 activity was tested by its ability to cleave synthetic peptide substrate DEVD-pNA and was noted to be detectable within 15 min after the start of DC treatment, with maximum activity at 1 h (see Fig. 7). Interestingly, we also found caspase-2, -6, and -8 activation. Caspase-2 cleaved synthetic substrate VDVAD-pNA, and a sustained increase of activity was detectable over a period of 2 h. The same pattern of enzyme activation was present for caspase-6 and -8, which cleaved VEID-pNA and IETD-pNA synthetic peptide substrate, respectively (see Fig. 7).
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Experiments with selected caspase inhibitors have shown that activation of the caspase cascade is functionally involved because inhibition of caspases reversed the proapoptotic effect of DC. The most inhibitory effect was observed with the caspase-6 inhibitor, which at 100 µM concentration abolished completely DC-induced DNA fragmentation. Caspase-9 and -3 inhibitors were also effective and reversed DC-mediated apoptosis by 70 and 60%, respectively (see Fig. 8). Enhanced activation of caspase-3, measurable just after 15 min of DC treatment, indicates that this effector caspase is involved early in the signaling cascade. On the other hand, inhibition of another effector caspase, namely caspase-6, which completely prevented DC-induced apoptosis, indicates the particular and important role that this caspase also plays in transferring the signal downstream to DNA degradation. Our observations demonstrate that DC-induced apoptosis in the AGS cell line is mediated by both mitochondrial (caspase-2 and -9) and receptor (caspase-8) pathways; however, further study is necessary to elucidate the precise mechanisms of this process.
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Effect of PGE2 on Cytotoxicity and Apoptosis Induced by DC
We also investigated the effect of dmPGE2 pretreatment on cytotoxicity and apoptosis in cells treated with DC. Several studies have implicated PG as a protective agent against cell injury (19, 20, 22, 44). Our findings demonstrated that the cytotoxic effect of DC, measured as LDH release, is attenuated when cells are pretreated with dmPGE2. Figure 9 showed that a 30-min pretreatment with 2.6 µM dmPGE2 decreased the cytotoxic effect of 300 µM DC at 0.5 to 2 h in cells incubated in HBSS/HEPES. However, we did not observe any protective effect of dmPGE2 against apoptosis induced by DC in Ham's F-12 media (Fig. 10) when using a 2.6 µM concentration or even higher doses of this agent.
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DISCUSSION |
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In previous studies, we grew our cells in Ham's F-12 media and then conducted our experimental studies in HBSS. With this approach, we found that low concentrations of DC (i.e., <200 µM) were generally nontoxic until cells had been exposed to such concentrations for extended periods (20, 21). In contrast, higher concentrations could evoke significant damage within 2030 min and often irreversibly killed the cells in a short period of time thereafter. Employing LDH release as a marker of necrotic damage, we were able to validate these earlier findings in this study, and we noted that concentrations of DC <200 µM were generally non-damaging until such cells were exposed to this chemical insult for 4 h. If a higher concentration of DC (namely 300 µM) was used to challenge our cells, significant damage was noted as early as 30 min, which became consistently more pronounced over time so that by 4 h,
90% of the cells had died a necrotic death.
Unexpectedly, a different pattern emerged if cells were not only grown in Ham's F-12 media but continually maintained in this environment for our experiments. Under these conditions, a concentration of DC as high as 300 µM elicited no evidence of cellular toxicity, as measured by LDH release, even after continuous exposure for 8 h. After 24 h, this concentration evoked considerable cellular death by necrosis, but a lower concentration such as 100 µM DC, which induced necrosis in 30% of cells within 4 h when maintained in Hank's solution, only elicited a 20% necrotic response at 24 h in Ham's F-12 media. Of further note, clear evidence of apoptosis could be demonstrated in cells exposed to DC in Ham's F-12 media that was both concentration and time related. Thus, at 4 h of incubation in this media, an apoptotic response was evoked in fewer than 10% of cells if the concentration of DC was 150 µM or less. With a concentration of 200 µM, the apoptotic response involved 25% of cells. At higher concentrations, such as 400 and 500 µM, this response reached a level of
50%.
To dissect out the events leading to apoptosis, we employed indexes of this type of cellular death that measured both early and late-term responses. Early markers that we employed included measurement of caspase activity and histone-associated DNA fragmentation. Depending on the concentration of DC used to damage gastric cells, caspase activation could be identified in <1 h after exposing cells to DC. For low concentrations of DC, such as 50 and 100 µM DC, no caspase activity was noted by 60 min following DC challenge. In contrast, caspase-3 activation was profoundly enhanced in cells exposed to 200 or 300 µMDC in this time frame (data not shown). In fact, in cells exposed to 300 µM DC, activation of caspase-3 was evident as early as 15 min after exposing cells to this damaging agent and reached its highest level by 60 min. Interestingly, caspase-2, caspase-6, and caspase-8 activation also occurred as briskly but did not achieve the levels observed with caspase-3. Because caspase-2 and caspase-8 are upstream initiators of caspase-3 and caspase-6 activation via both a receptor-mediated (i.e., caspase-8) and mitochondrial (i.e., caspase-2) mechanism, involvement of these caspases in bile-salt-induced apoptosis indicates that the processes through which DC mediates its apoptotic effects presumably employ both pathways.
Histone-associated DNA fragmentation was also markedly increased at 1 h following exposure to 300 µM DC. This effect remained sustained for at least 3 h and then slowly decreased. In agreement with our caspase data, no histone-associated DNA fragmentation was discernable with concentrations of DC in the range of 50100 µM DC at either 1 or 4 h. Similar to 300 µM DC, however, 200 µM DC also elicited significant histone-associated DNA fragmentation as early as 1 h. Time course findings suggested that caspase activation precedes the DNA fragmentation. Furthermore, the fact that caspase inhibitors (such as caspase-3, caspase-6, and caspase-9) can significantly prevent the formation of DNA fragmentation lends support to the contention that caspase activation is essential for the subsequent effects on DNA. Moreover, whereas blockades of caspases-3 and -6 are not surprising in their ability to prevent DNA fragmentation because they are both downstream effector caspases, the fact that caspase-9 inhibition is also effective suggests that the mitochondrion may also play a role because this caspase is an upstream initiator that orchestrates effects induced by this intracellular organelle. On the basis of these findings, a schematic representation of the possible pathways through which DC may trigger apoptosis in AGS cells is shown in Fig. 11.
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In addition to these early markers of apoptosis, two late markers were also measured, namely DNA laddering and the TUNEL assay. TUNEL positivity was not consistently evident until 4 h of exposure to DC and was clearly concentration related as observed with the earlier markers. Thus low concentrations of DC, such as 100 and 150 µM, showed mild degrees of apoptosis by the TUNEL assay, whereas concentrations of 200 µM or greater demonstrated more pronounced effects. In concentrations of DC approaching 300 µM or greater, 50% of cells were observed to be apoptotic by this assay. The results for DNA laddering were similar. Cells exposed to 100 µM DC demonstrated no consistent evidence of laddering until 4 h of exposure. In contrast, cells exposed to 300 µM DC had clear evidence of DNA laddering as early as 1 h following exposure to this damaging agent.
We previously reported that the prostaglandin dmPGE2 could prevent DC-induced gastric damage in cells in which a necrotic mechanism of death occurred (20, 21). We confirmed this finding in the present study. Thus cells exposed to 300 µM DC in Hank's buffered salt solution demonstrated a time-related necrotic response with >30% of cells undergoing necrosis by 30 min and >85% of cells experiencing necrotic death by 4 h. Pretreatment with PGE2 significantly attenuated the magnitude of this necrotic response during the first 2 h of DC exposure. By 4 h, this efficacy was overcome. Interestingly, in cells exposed to this same concentration of DC and maintained in Ham's culture media in which apoptosis occurred, pretreatment with dmPGE2 was ineffective in attenuating the apoptotic response (see Fig. 10). This being the case, such findings suggest that PGE2 can prevent the necrotic pathway of death but apparently has no effect on the apoptotic pathway. The mechanism(s) responsible for this difference in action remains to be elucidated.
As shown in Fig. 6, an ambient pH of 4.0 altered the method of cell death in response to DC exposure to one of necrosis in contrast to an apoptotic pathway at pH 7.4. There are several possible explanations for this finding. First, it is well known that when gastric cells are bathed with luminal acid, some of this acid back-diffuses into these cells. Normally, this is not a problem because intracellular buffering mechanisms adequately neutralize this influx and thereby maintain the intracellular pH. In the presence of bile, however, this back-diffusion is substantially increased, making it more difficult for the cell to maintain acid-base balance. Second, bile is also known to inhibit active transport mechanisms within gastric cells under acidic conditions. This effect would further make it difficult to neutralize influxing hydrogen ions. Third, the generation of an "alkaline tide," which is present in an acid-secreting gastric epithelium and assists in maintaining an acid-base balance in the gastric epithelium when challenged by a toxic substance such as bile, is absent under resting conditions and would clearly be absent in our model because it contains only gastric surface mucous cells. These mechanisms have been previously reviewed (27, 28, 42) and almost certainly are responsible, at least in part, for the differences in cell death induced by DC that we observed under acidic and nonacidic ambient environments. Whether other mechanisms are also operational will require further elucidation. Thus, whereas DC clearly induces apoptosis in gastric mucosal cells, this process can revert to a necrotic pathway of cell death under acidic conditions. Consistent with our findings, ammonia-induced apoptosis in gastric mucosal cells has also been shown to be pH dependent and to optimally occur in a neutral environment (41).
Finally, why DC initiates necrosis when cells are maintained in Hank's balanced salt solution but seems to be more resistant to injury and undergo primarily an apoptotic mechanism of death in Ham's medium remains uncertain. Because Ham's solution is primarily an amino acid-rich media, whereas Hank's solution is primarily a balanced salt solution, it is possible that the presumed growth-enhancing capabilities of Ham's solution enable cells to be more resistant to a noxious insult than what occurs under the other experimental conditions. Glutamine is a particularly important amino acid in Ham's solution and has been shown to be an essential nutrient in maintaining the health of gastrointestinal cells (15, 38). Thus it seemed likely that the rich amino acid mixture comprising Ham's solution, and particularly glutamine, provided a more healthy environment for cells, making it a more appropriate medium to study potentially injurious processes than the Hank's solution, which has been commonly employed in such research. To test the potential role that glutamine might play in protecting cells exposed to DC from following a necrotic pathway toward cell death, we added glutamine to Hank's buffer before challenging cells with DC when studied in this solution. Such glutamine addition did not prevent the induction of the necrotic pathway of cell death (data not shown). This being the case, further study is obviously needed to dissect out the parameters responsible for the different results that we observed with DC when cells were maintained in these differing culture media.
In conclusion, we have shown that DC can induce both necrosis and apoptosis in gastric epithelial cells and that the predominant mechanism of cell death is dependent upon the environment in which the cells are cultured. Because Ham's culture media more likely simulates conditions that would exist under in vivo circumstances, we believe that apoptosis is the major mechanism by which gastric cells die when exposed to damaging concentrations of DC. When cells are housed in an environment less likely to exist under normal living conditions, such as Hank's balanced salt solution, they become less resistant to the potential toxic effects of DC and die a necrotic death.
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DISCLOSURES |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
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REFERENCES |
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