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MEMBRANE TRANSPORTERS, ION CHANNELS, AND PUMPS

Oxidant-impaired intracellular Ca²⁺ signaling in pancreatic acinar cells: role of the plasma membrane Ca²⁺-ATPase

Faculty of Life Sciences, The University of Manchester, Manchester, United Kingdom

Submitted 20 November 2006 ; accepted in final form 7 May 2007

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Pancreatitis is an inflammatory disease of pancreatic acinar cells whereby intracellular calcium concentration ([Ca²⁺]_i) signaling and enzyme secretion are impaired. Increased oxidative stress has been suggested to mediate the associated cell injury. The present study tested the effects of the oxidant, hydrogen peroxide, on [Ca²⁺]_i signaling in rat pancreatic acinar cells by simultaneously imaging fura-2, to measure [Ca²⁺]_i, and dichlorofluorescein, to measure oxidative stress. Millimolar concentrations of hydrogen peroxide increased cellular oxidative stress and irreversibly increased [Ca²⁺]_i, which was sensitive to antioxidants and removal of external Ca²⁺, and ultimately led to cell lysis. Responses were also abolished by pretreatment with (sarco)endoplasmic reticulum Ca²⁺-ATPase inhibitors, unless cells were prestimulated with cholecystokinin to promote mitochondrial Ca²⁺ uptake. This suggests that hydrogen peroxide promotes Ca²⁺ release from the endoplasmic reticulum and the mitochondria and that it promotes Ca²⁺ influx. Lower concentrations of hydrogen peroxide (10–100 µM) increased [Ca²⁺]_i and altered cholecystokinin-evoked [Ca²⁺]_i oscillations with marked heterogeneity, the severity of which was directly related to oxidative stress, suggesting differences in cellular antioxidant capacity. These changes in [Ca²⁺]_i also upregulated the activity of the plasma membrane Ca²⁺-ATPase in a Ca²⁺-dependent manner, whereas higher concentrations (0.1–1 mM) inactivated the plasma membrane Ca²⁺-ATPase. This may be important in facilitating "Ca²⁺ overload," resulting in cell injury associated with pancreatitis.

oxidant stress; pancreatitis; calcium pump

Although the pathophysiology of pancreatitis remains incompletely understood, increasing evidence points to a role of oxidative stress as an underlying mechanism. In several animal models of pancreatitis, there is clear evidence that antioxidants reduce the associated cell injury (12, 31, 39), which has also proved beneficial in the treatment of human chronic pancreatitis (46). What remains unclear is whether oxidative stress is the trigger or a downstream consequence of an inflammatory response, which promotes the release of further oxidants from activated neutrophils at the site of injury (22, 50). However, there is substantial evidence that acinar cells can be a source of oxidant generation (7, 17, 42). In addition, in some pancreatitis models, lipid peroxidation products were detected prior to any inflammatory changes, suggesting that oxidative stress is an early event, if not the trigger of the disease (32).

The aim of the present study was to characterize in detail the effects of increased oxidative stress, using hydrogen peroxide (H₂O₂), on [Ca²⁺]_i homeostasis and CCK-evoked [Ca²⁺]_i oscillations in acinar cells. The results show that low concentrations of H₂O₂ increased [Ca²⁺]_i and profoundly altered the normal pattern of CCK-evoked [Ca²⁺]_i oscillations. These changes were transformed into an irreversible sustained increase in [Ca²⁺]_i, or Ca²⁺ overload, in an increasing number of cells as the H₂O₂ concentration was increased. This H₂O₂-evoked Ca²⁺ overload also corresponded to inactivation of the plasma membrane Ca²⁺ pump (PMCA), the last "gate keeper" in the control of normal [Ca²⁺]_i homeostasis. This suggests that oxidant-induced inactivation of the PMCA may be an important early event underlying the pathology of pancreatitis.

	MATERIALS AND METHODS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Cell isolation. Small clusters of rat pancreatic acinar cells were isolated by a collagenase digestion procedure similar to that previously described (5). Briefly, rats were killed by cervical dislocation, and the pancreas was dissected, minced with scissors, and digested for 30 min in 10 ml of standard HEPES-buffered physiological saline solution (HEPES-PSS) (composition in mM: 145 Na⁺, 5 K⁺, 1 Mg²⁺, 1.2 Ca²⁺, 148 Cl^–, 1 SO₄^2–, 1 H₂PO₄^–, and 10 HEPES; with a pH of 7.4) containing 800 U collagenase/g of tissue (Sigma Type II), 0.12 mg/ml soybean trypsin inhibitor (Sigma), and 1% (wt/vol) bovine serum albumin (BSA; Fraction V, Sigma). Following digestion, the tissue fragments were washed twice with HEPES-PSS, followed by 3-min incubation with Ca²⁺-free HEPES-PSS (containing 5 mM EDTA). Cells were rinsed with HEPES-PSS and then centrifuged twice through HEPES-PSS containing 4% BSA. Finally, the cells were resuspended in HEPES-PSS containing 0.1% BSA and kept on ice until use.

Digital imaging of fura-2 fluorescence. Cells were loaded with 4 µM fura-2 acetoxymethyl ester (fura-2 AM; TEF Labs/Stratech, Soham, UK) in HEPES-PSS for 30 min at room temperature. Dye-loaded cells were allowed to adhere to a glass coverslip that formed the base of a gravity-fed perfusion chamber, continually perfused with HEPES-PSS with automatic valves for rapid switching of solutions (Harvard Apparatus, Kent, UK). All fluorescence imaging experiments were performed using an inverted epifluorescence Nikon microscope with x40 oil immersion objective (numerical aperture 1.3), charge-coupled device (CCD) camera, and illumination device. Emitted fluorescence was separated from excitation light using a 400-nm diachronic mirror. Earlier experiments utilized a Nikon Diaphot microscope with a slow-scan CCD camera (Digital Pixel, Brighton, UK), xenon arc lamp (Nikon), and Lambda-10 filter wheel (Sutter Instruments, Novato, CA) controlled by Kinetic Imaging image acquisition and analysis software (KI, Nottingham, UK). Later experiments utilized a Nikon TE2000 microscope with a CoolSNAP HQ interline progressive-scan CCD camera (Roper Scientific Photometrics, Tucson, AZ), and Cairn monochromator illumination system (Cairn Research, Kent, UK) controlled by MetaFluor image acquisition and analysis software (Molecular Devices, Downington, CA). Background-subtracted 340-nm and 380-nm fluorescence images were captured with no binning at a rate of 1 Hz, and 340/380 ratiometric images were calculated offline. A x1 relay lens on the microscope side port leading to the camera was used to increase the field of view so that a typical field of view contained 20–35 cells. The fura-2 fluorescence was calibrated into "estimated" [Ca²⁺]_i using the following equation: [Ca²⁺]_i = K_d(R – R_min)/(R_max – R)(S_F380/S_B380) (18). Where K_d is the fura-2 dissociation constant (150 nM) (48), R is any given 340/380 ratio value, S_F380/S_B380 is the ratio of fluorescence measured for Ca²⁺-free and Ca²⁺-bound fura-2, and R_min and R_max are the minimum and maximum ratio values following in situ calibration experiments. This involved treating cells with 10 µM ionomycin, 0.5 µM carbonyl cyanide m-chlorophenylhydrazone (CCCP; to prevent mitochondrial Ca²⁺ uptake), and 30 µM cyclopiazonic acid [CPA; to prevent Ca²⁺ uptake into the endoplasmic reticulum (ER)] in Ca²⁺ free HEPES-PSS (with 1 mM EGTA) to obtain R_min values and Ca²⁺-saturated media (2 mM Ca²⁺) to obtain R_max values. Average values were obtained from periodic in situ calibration experiments throughout the study and for different microscopes and objective lenses. All experiments were carried out at room temperature (20–22°C).

Digital imaging of dichlorofluorescein fluorescence. Cells were loaded with 10 µM dichlorodihydrofluorescein diacetate (H₂DCF-DA) (Molecular Probes/Invitrogen, Paisley, UK) for 30 min at room temperature. Loaded cells exhibited minimal fluorescence until treated with H₂O₂, which oxidizes H₂DCF to the fluorescent 2',7'-dichlorofluorescein (DCF). Images of DCF fluorescence were obtained on the same imaging system as fura-2 images, but they were obtained using a fluorescein filter set. Time courses of normalized DCF fluorescence (%F/F_o, % change in fluorescence divided by the starting fluorescence) were then derived for each individual cell in the field with 4-s time resolution.

Simultaneous measurement of oxidative stress and cytosolic [Ca²⁺]_i. For simultaneous DCF and fura-2 imaging, cells were loaded with 10 µM fura-2 AM for 20 min at room temperature, followed by dual loading with 10 µM fura-2 AM and 10 µM H₂DCF-DA for 20 min at 37°C. These loading conditions were found empirically to be optimal for achieving a measurable fura-2 signal without compromising the DCF signal. Under these conditions, cells were more heavily loaded with fura-2 than cells in experiments measuring [Ca²⁺]_i alone. However, there was no apparent difference in the pattern or frequency of CCK-evoked [Ca²⁺]_i oscillations that would indicate excessive buffering.

For dual-dye imaging, 340 nm, 380 nm (for fura-2), and 490 nm (for DCF) excitation wavelengths were selected using either the filter wheel or monochromator. Emitted fluorescence was captured through the same FITC filter set used for recording DCF fluorescence alone, which captured 35–40% of the emitted fura-2 fluorescence. Ratiometric fura-2 [Ca²⁺]_i images, and normalized DCF images, were calculated offline with 4-s time resolution and analyzed as described above.

Solutions. In all imaging experiments, cells were superfused with a HEPES-PSS. Solutions containing H₂O₂ were made up fresh each day and were periodically assayed using a fluorometric assay, which monitors the conversion of p-hydroxyphenylacetate to a stable colored product during the reduction of H₂O₂ by horseradish peroxidase (20). H₂O₂ was applied to the cells by perfusion of H₂O₂-containing HEPES-PSS. This ensured that the cells were always exposed to a fresh supply of H₂O₂ at the desired concentration, because H₂O₂ degradation by cellular enzymes can markedly alter H₂O₂ concentrations under static incubation conditions (49). For experiments with La³⁺ (1 mM) to block the PMCA, all SO₄^2– and H₂PO₄^– ions were replaced with Cl^– to prevent precipitation of La³⁺ salts. Stock solutions (1 mM) of Ru360 (Calbiochem) were dissolved in deoxygenated water and used immediately before use.

Assessment of H₂O₂-evoked oxidation of CCK. Because some experiments involved simultaneous application of H₂O₂ and CCK to cells, it was necessary to consider the possibility that H₂O₂ might oxidize CCK rather than act directly on the acinar cell. To address this possibility, H₂O₂ (100 µM) was added to HEPES-PSS containing 20 pM CCK. After 5 min, catalase (50 U/ml) was added to remove the H₂O₂. Degradation of H₂O₂ was monitored by periodically assaying aliquots of the medium for H₂O₂ (as above). After all the H₂O₂ had been destroyed, the experimental medium was applied to acinar cells, where it proved equally as potent in eliciting [Ca²⁺]_i oscillations as medium containing 20 pM CCK that had not been exposed to H₂O₂ (data not shown). This confirmed that H₂O₂ does not act by oxidizing CCK.

Data analysis and experimental design. Because of the nature of most experiments, an unpaired experimental design was utilized. Comparisons were made and statistical significance determined between groups of experiments (e.g., control vs. treatment) using, where appropriate, an unpaired Student's t-test or Mann-Whitney test. For any given parameter analyzed, an experimental average was determined from several cells in a particular experiment. These values were in turn averaged to give the true overall average expressed in the text as means ± SE.

	RESULTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Effects of millimolar concentrations of H₂O₂ on [Ca²⁺]_i. To investigate the acute effects of an oxidant on [Ca²⁺]_i in pancreatic acinar cells, fura-2-loaded cells were continuously perfused with high concentrations of H₂O₂ (1–3 mM) as a convenient experimental means of imposing increased intracellular oxidative stress. Fig. 1A demonstrates that H₂O₂ evoked a large, slow and irreversible increase in [Ca²⁺]_i in pancreatic acinar cells (Fig. 1C; maximum increase in [Ca²⁺]_i with 3 mM H₂O₂, 1,167 ± 218 nM, n = 4; with 1 mM H₂O₂, 1,014 ± 176 nM, n = 4). This H₂O₂-induced increase in [Ca²⁺]_i was almost completely abolished by the combined treatment with the thiol-protecting agent and antioxidant, dithiothreitol (DTT) (Fig. 1, A and C; increase in [Ca²⁺]_i with 1 mM H₂O₂ and 2 mM DTT, 81 ± 26 nM, n = 4), suggesting that this was due to increased oxidative stress. In addition, the H₂O₂-induced increase in [Ca²⁺]_i was markedly attenuated when external Ca²⁺ concentration ([Ca²⁺]_o) was removed (Fig. 1, A and C; maximum increase of 317 ± 88 nM, n = 4), suggesting that Ca²⁺ entry contributes to this response. The reduced increase in [Ca²⁺]_i remaining in the absence of [Ca²⁺]_o was presumably derived from an intracellular source, such as agonist-sensitive ER Ca²⁺ stores or mitochondria. To test this further, cells were stimulated with a cocktail of 30 µM 2,5-di(tert-butyl)-hydroquinone (TBQ) and 30 µM CPA, to completely inhibit the (sarco)endoplasmic reticulum Ca²⁺-ATPase (SERCA) and deplete ER Ca²⁺ stores, in the absence of [Ca²⁺]_o. The combined use of TBQ and CPA was shown to be more effective at depleting the ER than applying either drug alone (data not shown). Under these conditions (i.e., in the absence of Ca²⁺ influx and when the ER is depleted), H₂O₂ failed to evoke any increase in [Ca²⁺]_i (see Fig. 1Bii). However, if cells were prestimulated with 20 pM CCK, which evokes oscillations in [Ca²⁺]_i (Fig. 1Bi) and thus causes substantial mitochondrial Ca²⁺ uptake (16), then H₂O₂ did evoke a significant increase in [Ca²⁺]_i (Fig. 1Bi, and 1C, maximum [Ca²⁺]_i change 143 ± 27 nM, n = 5, P < 0.05). This suggests that the H₂O₂-evoked increase in [Ca²⁺]_i under these conditions was due to release of mitochondrial Ca²⁺. To test this directly, these experiments were repeated whereby the cells were treated with a combination of CPA/TBQ (to deplete the ER) and with the mitochondrial uncoupler, CCCP (0.5 µM), which is known to release any Ca²⁺ that had been taken up by the mitochondria during stimulation with CCK (Fig. 1Biii). Similarly to Fig. 1Bii, following this maneuver H₂O₂ failed to evoke any increase in [Ca²⁺]_i. In summary, the experiments in Fig. 1 suggest that the major source of the H₂O₂-evoked increase in [Ca²⁺]_i was from agonist-sensitive ER Ca²⁺ stores and activation of Ca²⁺ entry. Mitochondrial Ca²⁺ release may also contribute to the [Ca²⁺]_i increase following or during agonist stimulation, when the mitochondria have taken up Ca²⁺.

View larger version (26K): [in this window] [in a new window]   Fig. 1. High concentrations of hydrogen peroxide (H2O2; 1–3 mM) evoked a slow and irreversibly sustained increase in intracellular Ca2+ concentration ([Ca2+]i) that was dependent on Ca2+ influx and depletion of intracellular Ca2+ stores. A: effects of 3 mM and 1 mM H2O2, 1 mM H2O2 in the absence of external Ca2+ concentration ([Ca2+]o) and 1 mM H2O2 in the presence of dithiothreitol (DTT; 2 mM) on baseline [Ca2+]i. B: cells were either prestimulated with 20 pM cholecystokinin (CCK; Bi and Biii), to promote mitochondrial Ca2+ uptake, or left unstimulated (Bii) and the effects of H2O2 on baseline [Ca2+]i in the absence of [Ca2+]o and following depletion of the endoplasmic reticulum Ca2+ stores with the (sarco)endoplasmic reticulum Ca2+-ATPase inhibitors 2,5-di(tert-butyl)-hydroquinone (TBQ) and cyclopiazonic acid (CPA) in the presence (Biii) or absence (Bi and Bii) of the mitochondrial uncoupler, carbonyl cyanide m-chlorophenylhydrazone (CCCP). C: summary of the maximum increase in [Ca2+]i evoked by H2O2 (1 or 3 mM) under the various conditions shown in A and B. Values are means ± SE. Max, maximal; pre-stim, prestimulation. *Statistical significance, P < 0.05 (using a 1-sample t-test). Statistical significance compared with the effects of 1 mM H2O2 alone, P < 0.05 (unpaired t-test).

View larger version (21K): [in this window] [in a new window]   Fig. 2. Lower concentrations of H2O2 (10–100 µM) produced heterogeneous effects on baseline [Ca2+]i. Cells were stimulated 20 pM CCK, to test for cell viability, followed by H2O2 (10–100 µM) and then 20 pM and 1 nM CCK, to test for the reversibility and recovery of the H2O2 response. Responses were categorized based on the severity (A, B, and C). Group A response was the most severe and was characterized by an irreversible, sustained increase in [Ca2+]i; group B was characterized by a transient, reversible increase in [Ca2+]i; and finally cells categorized in the group C response exhibited no response. The heterogeneity of responses was quantified at each concentration of H2O2 by counting the number of cells exhibiting each response for every experiment, which is summarized as a percentage (± SE) of the total number of cells treated in each experiment in the bottom right panel. Inset, effects of another oxidant, thimerosal, on baseline [Ca2+]i that evoked [Ca2+]i oscillations in pancreatic acinar cells (n = 5 experiments).

Effects of submillimolar H₂O₂ during CCK-evoked [Ca²⁺]_i oscillations. We also examined the effects of low concentrations of H₂O₂ (50, 75, and 100 µM) during a train of CCK-evoked [Ca²⁺]_i oscillations. Similarly, these effects were heterogeneous, and thus responses were categorized for analysis. In this case, the three main types of response observed were the following: a severely impaired response, whereby oscillations slowly diminished and fused into a sustained increase in baseline [Ca²⁺]_i (group A, Fig. 3); a mildly impaired response, whereby oscillations slowed and diminished and baseline [Ca²⁺]_i transiently increased (group B, Fig. 3); and finally, no observable change in either baseline [Ca²⁺]_i or the oscillation pattern (group C, Fig. 3). Again, the effects of H₂O₂ appeared to be concentration dependent in terms of the number of cells exhibiting each response. Thus a large number of cells exhibited impaired [Ca²⁺]_i oscillations (group A 57 ± 2% and group B 37 ± 2%; n = 8, 143 cells, Fig. 3) in response to the highest concentration of H₂O₂ (100 µM), whereas few cells were unaffected (group C, 6 ± 1%; n = 8, 143, Fig. 3). In contrast, when the concentration of H₂O₂ was reduced to 50 µM, the majority of cells were unaffected (78 ± 2%, n = 5, 152 cells). The intermediate concentration (75 µM) evoked more heterogeneous responses [29 ± 5% (A), 54 ± 3% (B), 18 ± 5% (C); n = 6, 133 cells, Fig. 3].

View larger version (23K): [in this window] [in a new window]   Fig. 3. Lower concentrations of H2O2 (50–100 µM) produced heterogeneous effects on CCK-evoked [Ca2+]i oscillations. Responses were categorized based on the severity (A, B, and C). Group A response was the most severe and was characterized by a fusing of oscillations into a sustained increase in baseline [Ca2+]i. Group B was characterized by either a decrease in frequency or amplitude of oscillations superimposed over a transient increase in baseline [Ca2+]i. In cells categorized in the group C response, there was no change in the oscillatory [Ca2+]i response. Heterogeneity of responses was quantified by counting the number of cells exhibiting each response for every experiment, which is summarized as a percentage (±SE) of the total number of cells treated in each experiment in the right panel.

Direct measurement of cellular oxidative stress with DCF fluorescence. We next tested whether the observed heterogeneity to H₂O₂ (Figs. 2 and 3) was due to differences in cellular oxidative stress using the oxidizable fluorescein derivative, H₂DCF (23, 42). Following optimization of loading conditions, it was found that cells loaded with H₂DCF initially showed very weak fluorescence (Fig. 4Ai, fluorescent image at 0 min). Because oxidation of the dye is irreversible, normalization was achieved by expressing the data as F/F₀ (Fig. Aii, Bi, Bii, and C). On exposure to 3 mM H₂O₂, cellular fluorescence increased markedly to almost saturated levels (Fig. 4Ai, fluorescent image at 20 min), presumably reflecting intracellular oxidation of H₂DCF to the fluorescent product, DCF [Fig. 4, A and B; (23, 42)]. Monitoring DCF fluorescence in individual cells revealed that the increase in DCF fluorescence was extremely heterogeneous (Fig. 4Aii). Although virtually all cells showed some increase in DCF fluorescence at this high H₂O₂ concentration, the rate of the increase varied considerably between different cells. It was also noticeable that, after 20 min of H₂O₂ exposure, cells often showed a large decrease in DCF fluorescence; sometimes they appeared as a single step (see open arrows in Fig. 4Aii) or as multiple steps. These decreases were not due to a decrease in oxidative stress because oxidation of the dye is irreversible, but rather they were due to cell lysis and thus loss of DCF from the cell. Inspection of phase-contrast images of the cells confirmed that at 40 min cells that had lost DCF fluorescence showed clear evidence of cell lysis (Fig. 4Ai, fluorescent image at 40 min and brightfield image after).

View larger version (39K): [in this window] [in a new window]   Fig. 4. Direct measurement of oxidative stress using DCF fluorescence. A: representative experiment showing the effect of 3 mM H2O2 on DCF fluorescence. Ai: brightfield phase images before and after treatment with H2O2 and 2',7'-dichlorofluorescein (DCF) fluorescent images at 0, 20 and 40 min of H2O2 treatment. At 0 min, DCF fluorescence is very low in all cells; at 20 min, DCF fluorescence has reached almost saturated levels; and at 40 min, many cells have undergone cell lysis. Aii: time course of the same experiment in Ai, showing the normalized relative DCF fluorescence (%F/F0; see MATERIALS AND METHODS) of individual cells in response to 3 mM H2O2. Open arrows indicate a sudden loss of DCF fluorescence due to cell lysis. B: time course showing the average normalized DCF fluorescence (%F/F0) of individual experiments in response to 0 mM (control) to 3 mM H2O2 (Bi) and 3 mM H2O2 in the presence of 2 mM DTT, preincubation with 2 mM DTT (for 60 min), or a combination of both (Bii). C: overall mean data (±SE), showing the change in DCF fluorescence (%F/F0) after 20 min with or without the various treatments shown in B. pre-inc, Preincubation. *Statistical significance, P < 0.05 (using a 1-sample t-test). Statistical significance compared with the effects of 3 mM H2O2 alone, P < 0.01 (using an unpaired t-test).

Simultaneous measurement of oxidative stress and [Ca²⁺]_i; H₂O₂-evoked oxidative stress correlates to the degree of impairment of CCK-evoked [Ca²⁺]_i signaling. As indicated above, the changes in DCF fluorescence and [Ca²⁺]_i evoked by H₂O₂ showed considerable cell-to-cell heterogeneity. It is tempting to speculate that this could have a common cause based on intercellular differences in antioxidant capacity, such that cells with low antioxidant capacity would exhibit a faster rise in DCF fluorescence and a more marked disturbance of [Ca²⁺]_i signaling. This was tested by loading cells with both fura-2 and H₂DCF to allow the simultaneous measurement of cellular oxidative stress and CCK-evoked [Ca²⁺]_i oscillations in individual cells (Fig. 5). Similarly to the experiments shown in Fig. 3, these effects were categorized into three general responses (A to C, Fig. 5A): a severely impaired response, whereby oscillations fused into a sustained increase in baseline [Ca²⁺]_i (group A, Fig. 5Ai); mildly impaired response, whereby the frequency or amplitude of oscillations decreased without any noticeable change in baseline [Ca²⁺]_i (group B, Fig. 5Aii); and finally, no observable change in either baseline [Ca²⁺]_i or the oscillation pattern (group C, Fig. 5iii).

View larger version (34K): [in this window] [in a new window]   Fig. 5. H2O2-evoked oxidative stress correlates with the degree of impairment of CCK-evoked [Ca2+]i signaling. Pancreatic acinar cells were dual loaded with DCF and fura-2 to simultaneously measure oxidative stress (gray trace, A and B) and CCK-evoked [Ca2+]i oscillations (black trace, A and B). A: representative responses to 100 µM in the same experiment that were categorized based on the "severity" of impairment of the CCK-evoked [Ca2+]i oscillations. In group A, oscillations fused into a sustained increase in baseline [Ca2+]i (Ai); in group B the frequency or amplitude of oscillations decreased without any noticeable change in baseline [Ca2+]i (Aii); and finally in group C there was no observable change in the oscillation pattern (Aiii). B: a typical control CCK-evoked [Ca2+]i oscillation response (Bi), and 2 mM DTT prevents the effect of H2O2 on CCK-evoked [Ca2+]i oscillations (Bii). C: mean maximum change in DCF fluorescence (%F/F0) from cells categorized into group A (29.2 ± 2.2%, n = 4, 18 cells; labeled A in C), group B (8.9 ± 1.0%, n = 4, 24 cells; labeled B in C) and group C (4.3 ± 1.1%, n = 4, 12 cells; labeled A in C) in response to 100 µM H2O2 (A, gray bars in C), which is also compared with control cells (Bi; 3.4 ± 1.2%, n = 4, 69 cells; black bar in C) and DTT-treated cells (Bii; 2.7 ± 0.9%, n = 4, 74 cells; hatched bar in C). Statistical analysis (Mann-Whitney test) revealed that the maximum increase in DCF fluorescence in the most severely effected cells (group A, Ai) was significantly greater (**P < 0.05) than the mildly affected cells (group B, Aii), which were both significantly greater than control cells (*P < 0.05). In contrast, the maximum increase in DCF fluorescence in unaffected cells (group C, Aiii), control cells (Bi) and DTT-treated cells (Bii) were statistically indistinguishable.

For all the data in Figs. 2–5, the observed heterogeneity was also apparent within cells from the same experiment (see mean data) and sometimes from the same acinus in any given experiment. In addition the type of response observed did not seem to depend on the extent of cell isolation as it was also noticed that some single cells were relatively "protected" compared with cells that were part of a large acinus. However, these are qualitative observations, and it would be difficult to completely rule out the effect of cell isolation on the heterogeneity without comparing the responses with those in vivo, which is technically very demanding. Furthermore, any approach to prevent oxidative damage during the isolation procedure may artificially improve the cells' antioxidant capacity and therefore artificially eliminate any heterogeneity that may have existed in vivo.

H₂O₂ regulates PMCA activity. We next attempted to identify the underlying mechanism responsible for the H₂O₂-evoked irreversible increase in [Ca²⁺]_i (Ca²⁺ overload). Rather than identifying the source of the increase in [Ca²⁺]_i, we reasoned that the mechanism of impaired [Ca²⁺]_i removal may be more important in converting a "mildly" effected cell to a severely effected cell, thus facilitating irreversible Ca²⁺ overload. We therefore tested the effects of H₂O₂ on PMCA activity utilizing an in situ assay, similar to that used in previous studies in parotid acinar cells (6). This involved depleting ER Ca²⁺ stores by inhibition of SERCA with TBQ and CPA. This increases [Ca²⁺]_i due to Ca²⁺ leak from the ER and activation of store-operated Ca²⁺ entry (SOCE) (36). Subsequent removal of [Ca²⁺]_o, and thus the Ca²⁺ influx component, essentially isolates Ca²⁺ efflux due to PMCA activity (Fig. 6A). This was quantified by fitting the falling phase of the [Ca²⁺]_i clearance to an exponential decay, which yielded an average time constant () of 90 ± 10 s (n = 4, 43 cells, Fig. 6A) Under these conditions, all other [Ca²⁺]_i clearance pathways are either inhibited (SERCA) or are expected to contribute little to the [Ca²⁺]_i clearance [Na⁺-Ca²⁺-exchanger (30) and mitochondria (10)]. To validate experimentally that the major [Ca²⁺]_i clearance pathway under these conditions was PMCA activity, we used two basic experimental approaches. First, to test whether mitochondrial Ca²⁺ uptake contributed to the [Ca²⁺]_i clearance, cells were preincubated with the specific mitochondrial Ca²⁺ uptake inhibitor, Ru360 (10 µM), for 30 min before starting the "[Ca²⁺]_i clearance assay". This has been shown to markedly inhibit mitochondrial Ca²⁺ uptake in pancreatic (21) and parotid acinar cells (6). Although there appeared to be a slowing of the clearance rate in some cells, on average this did not reach statistical significance ( = 114 ± 27 s, n = 5, 57 cells; Fig. 6B), suggesting that mitochondrial Ca²⁺ uptake is not the major [Ca²⁺]_i clearance pathway under the conditions of these experiments. Second, to further validate that the [Ca²⁺]_i clearance is due to the PMCA, La³⁺ (1 mM) was used to inhibit the PMCA (3, 8). This was applied either before the addition of CPA/TBQ (Fig. 6C) or during the increasing phase of the CPA/TBQ-evoked Ca²⁺ response (Fig. 6D). At this concentration La³⁺ inhibits both Ca²⁺ influx and Ca²⁺ efflux, effectively "sealing" the cell so that Ca²⁺ remains trapped so that it can neither enter nor leave the cell. Therefore, in the continued presence of La³⁺, CPA/TBQ-evoked a substantial increase in [Ca²⁺]_i that remained elevated despite the subsequent removal of external Ca²⁺, presumably due to complete inhibition of the PMCA (see Fig. 6C). However, removal of La²⁺ using EGTA, which binds La³⁺ with high affinity, slowly reversed the inhibition of the PMCA, which then started to rapidly clear Ca²⁺ from the cytosol (see Fig. 6A). Furthermore, on average CPA/TBQ evoked a much larger increase in [Ca²⁺]_i in the presence of La³⁺ (407 ± 51 nM, n = 4, 54 cells; compared with control cells, 219 ± 34 nM, n = 4, 43 cells), suggesting that PMCA actively removes Ca²⁺ from the cytosol as Ca²⁺ continues to leak from the ER, thereby reducing the net magnitude of the CPA/TBQ-evoked [Ca²⁺]_i response. Addition of La³⁺ during the rising phase of the CPA/TBQ-evoked [Ca²⁺]_i response caused an initial decrease, probably due to the rapid inhibition of Ca²⁺ influx, followed by a further increase that then rapidly reached a steady state, presumably due to the gradual inhibition of the PMCA (see Fig. 6D). One possible explanation for these observations is that La²⁺ enters the cell and interferes with fura-2 fluorescence, because it is known to bind to fura-2 with pM affinity (25). This could occur either by leak of La³⁺ or by La³⁺ entry through SOCE channels. However, this is highly unlikely because addition of La³⁺ neither quenched nor saturated the fura-2 signal, which would be expected even if small amounts of La³⁺ entered the cell (25). Furthermore, La³⁺ is thought to inhibit SOCE channels by a similar mechanism to voltage-operated Ca²⁺ channels (VOCCs) by blocking the pore of the channel (33), and it has recently been shown that La³⁺ cannot enter chromaffin cells through VOCCs (25). Therefore, these data provide convincing evidence that the PMCA is the major [Ca²⁺]_i clearance pathway under the conditions of the experiments shown in Fig. 6.

View larger version (18K): [in this window] [in a new window]   Fig. 6. Functional [Ca2+]i clearance is unaffected by mitochondrial Ca2+ uptake inhibitors (Ru360) but markedly inhibited by the PMCA inhibitor, La3+. Pancreatic acinar cells were treated with 30 µM TBQ and 30 µM CPA [to inhibit (sarco)endoplasmic reticulum Ca2+-ATPase], followed by removal of external Ca2+ ([Ca2+]o) to initiate the measurement of [Ca2+]i clearance (see RESULTS). Cells were pretreated with (B) or without (A) 10 µM Ru360 for 30 min, to prevent mitochondrial Ca2+ uptake, before beginning the "[Ca2+]i clearance assay." [Ca2+]i clearance was quantified by fitting the falling phase to a single exponential decay to give the time constant () (control A, = 90 ± 10 s, n = 4, 43 cells; Ru360 B, = 114 ± 27 s, n = 5, 57 cells). [Ca2+]i clearance was inhibited when 1 mM La3+ was either applied before the addition of CPA/TBQ (C, n = 4, 56 cells) or during the rising phase of the CPA/TBQ-evoked [Ca2+]i response (D, n = 4, 47 cells). The inhibition was reversed when cells were perfused with 1 mM EGTA to remove the La3+ (C and D).

View larger version (22K): [in this window] [in a new window]   Fig. 7. H2O2 regulates [Ca2+]i clearance. Pancreatic acinar cells were treated with or without (control) H2O2 for 15 min before beginning the [Ca2+]i clearance assay (see RESULTS). A.: representative traces showing the effects of various concentrations of H2O2 pretreatment on [Ca2+]i clearance (PMCA activity), plotted on the same [Ca2+]i scale. [Ca2+]i clearance was quantified by either fitting the falling phase to a single exponential decay to give (mean data shown in B) or alternatively by computing the maximum rate of change in [Ca2+]i (over 30 s) following the removal of [Ca2+]o (max rate; mean data shown in C). In cells treated with 1 mM H2O2, data could not be fitted to a single exponential decay. C: mean data of the starting [Ca2+]i (shown as the crosshairs in A) at which [Ca2+]o was removed and [Ca2+]i clearance measurements began, plotted against each concentration of H2O2. Statistical analysis utilized an unpaired t-test to compare each parameter (, max rate, or start [Ca2+]i) following treatment with H2O2 to the corresponding parameter in untreated control cells (*P < 0.05).

The most important observation from these experiments was that there was a transition at or above 100 µM H₂O₂, whereby [Ca²⁺]_i clearance, and thus PMCA activity, was dramatically reduced despite a further increase in preclearance start [Ca²⁺]_i (see Fig. 7C). Higher concentrations of H₂O₂ reduced [Ca²⁺]_i clearance from 140 ± 16 nM/min (control, n = 4, 43 cells) to 87 ± 16 nM/min (300 µM H₂O₂, n = 4, 46 cells, P = 0.057) and 42 ± 11 nM/min (1,000 µM, n = 4, 52 cells, P < 0.001; see Fig. 7C). Similar exponential decay data revealed that 300 µM H₂O₂ significantly increased from 90 ± 10 s to 327 ± 67 s (P = 0.0012; see Fig. 6B), and the data with 1 mM H₂O₂ could not be fitted to a single exponential decay, suggesting almost complete inactivation of the PMCA. The effect of 100 µM H₂O₂ on PMCA activity ( = 112 ± 46, Fig. 7B; maximum rate = 239 ± 56 nM/min; n = 4, 42 cells) was not significantly different from the control using either analytic method. However, this effect was likely a mixture of responses from two subpopulations of cells, whereby H₂O₂ either increased or decreased PMCA activity. It is also worth noting that the progressive increase in preclearance start [Ca²⁺]_i was due to a greater extent to an increase in resting [Ca²⁺]_i (>100 µM), rather than an enhanced CPA/TBQ-evoked Ca²⁺ leak (10–50 µM). This suggests that the irreversible nature of the H₂O₂-evoked Ca²⁺ overload response was due to inhibition of the PMCA.

Although under control conditions the major [Ca²⁺]_i clearance pathway appears to be due to the PMCA (Fig. 6), evidence from Fig. 7 suggests that H₂O₂ may promote a greater ER Ca²⁺ leak, which could influence the clearance rate. Therefore, to test the direct effect of H₂O₂ on PMCA activity, the following experimental approach was applied (see Fig. 8). First, cells were preincubated with 10 µM Ru360 (similarly to Fig. 6) to inhibit mitochondrial Ca²⁺ uptake and subsequent Ca²⁺ release. Cells were then treated with CPA/TBQ in the absence of external Ca²⁺ to deplete ER Ca²⁺ (similarly to Fig. 1B), and following the recovery of [Ca²⁺]_i, H₂O₂ was applied before external Ca²⁺ was added back which increased [Ca²⁺]_i due to SOCE (Fig. 8). [Ca²⁺]_i clearance (and thus PMCA activity) was then measured following a further removal of external Ca²⁺. The direct effects of 50 µM and 1 mM H₂O₂ was assessed because these concentrations caused the maximum effect on [Ca²⁺]_i clearance in Fig. 7. Under these conditions 50 µM H₂O₂ had no direct effect on [Ca²⁺]_i clearance (rate = 90 ± 16 nM/min, = 49 ± 5 s; n = 5, 62 cells) compared with control cells (rate = 86 ± 5 nM/min; = 41 ± 4, n = 6, 99 cells). This suggests that the increased [Ca²⁺]_i clearance following treatment with 50 µM H₂O₂ (observed in Fig. 7) was Ca²⁺-dependent due to enhanced Ca²⁺ leak from the ER. However, 1 mM H₂O₂ significantly inhibited [Ca²⁺]_i clearance (rate = 8 ± 3 nM/min; n = 4, 47 cells), consistent with experiments in Fig. 7 and the conclusion that inactivation of the PMCA is responsible for the H₂O₂-evoked irreversible Ca²⁺ overload.

View larger version (15K): [in this window] [in a new window]   Fig. 8. High concentrations of H2O2 (1 mM) inactivate plasma membrane Ca2+ pump activity. Pancreatic acinar cells were pretreated with Ru360 (similarly to Fig. 6) to inhibit mitochondrial Ca2+ uptake before addition of CPA/TBQ in the absence of [Ca2+]o to deplete endoplasmic reticulum [Ca2+]. [Ca2+]o was added back, which increased [Ca2+]i, because of activation of store-operated Ca2+ entry. Subsequent removal of [Ca2+]o evoked [Ca2+]i clearance that was either fitted to a single exponential decay (A, control, = 41 ± 4, n = 6, 99 cells) or the maximum rate in [Ca2+]i (over 30 s) following the removal of [Ca2+]o (A, control 86 ± 5 nM/min). H2O2 (B, 50 µM and D, 1 mM) was added just before the addition of [Ca2+]o to test the direct effect on the [Ca2+]i clearance following the subsequent removal of [Ca2+]o. 1 mM H2O2 markedly inhibited the rate of [Ca2+]i clearance (8 ± 3 nM/min, P < 0.001).

	DISCUSSION

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The present study clearly demonstrates that H₂O₂ profoundly alters hormone-evoked [Ca²⁺]_i signaling and resting [Ca²⁺]_i homeostasis in pancreatic acinar cells. These changes were due directly to increases in intracellular oxidative stress as assessed by DCF fluorescence and sensitivity to antioxidants. High levels of oxidative stress (1–3 mM H₂O₂) evoked a large, slow, and irreversible increase in [Ca²⁺]_i that after prolonged periods (>20 min) ultimately led to cell lysis, as indicated by the rapid loss of DCF dye. In addition, and almost certainly more important, lower levels of oxidative stress (10–100 µM H₂O₂) were also capable of evoking similar irreversible increases in [Ca²⁺]_i in subpopulations of cells. These observations are important because cell lysis is a common feature of necrotic cell death and may underlie the pathology of pancreatitis. In addition, an irreversible increase in [Ca²⁺]_i, or Ca²⁺ overload, is also thought to be a common feature of pancreatitis whatever the causal agent (35).

Ca²⁺ overload is a common pathological precursor that mediates a variety of disease states, such as excitotoxicity during neurodegeneration (1). A common strategy for understanding this Ca²⁺ overload, either during excitotoxicity or during pancreatitis, has been to identify the sources of the [Ca²⁺]_i increase, including most notably, mitochondria (15, 34) and Ca²⁺ entry pathways (1, 37). However, the irreversible nature of this Ca²⁺ overload suggests that perhaps a critical mechanism underlying this response is impairment of [Ca²⁺]_i clearance pathways. In the present study, we have demonstrated that a critical concentration of H₂O₂ (>100 µM) dramatically and irreversibly inhibits or inactivates Ca²⁺ efflux. Interestingly, this corresponds to the H₂O₂ concentration that produced an irreversible Ca²⁺ overload in the vast majority of cells.

Oxidant-mediated regulation of PMCA has been directly demonstrated in neuronal and liver membranes (54) and has been suggested to occur in pancreatic acinar cells (34). The mechanism could be due to direct redox modulation of critical thiol groups within the PMCA or within calmodulin (CaM) (54) both of which could inhibit Ca²⁺/CaM-dependent activation of the PMCA (13). Another possibility is that the H₂O₂-evoked increase in [Ca²⁺]_i activates Ca²⁺-dependent proteases, such as caspases or calpain, which are known to cleave and thus inactivate the PMCA (40). A similar mechanism has been described in neurons, whereby caspase cleavage of the Na⁺/Ca²⁺ exchanger, was shown to mediate the Ca²⁺ overload during the excitotoxicity associated with brain ischemia (2). Moreover, and of direct relevance to the present study, two groups have demonstrated that Ca²⁺-dependent activation of calpain leads to activation of the cytokine transcription factor, NF-KB, and pancreatic acinar cell injury (47, 51). In addition, cell injury associated with cerulean-induced pancreatitis was markedly ameliorated by calpain inhibitors (47, 52). Finally, there is now evidence that H₂O₂ can directly activate calpain, and that this leads to similar cell injury to that observed during pancreatitis (51). However, the time frame over which calpain became significantly activated (60-min exposure to 0.5 mM H₂O₂) in these experiments (51) occurred much later than when we observed PMCA inactivation (10-min exposure to 1 mM H₂O₂).

In addition, low concentrations of H₂O₂ (10–50 µM) transiently increased baseline [Ca²⁺]_i in a large proportion of cells, and it is also worth noting that pretreatment of similar concentrations significantly increased PMCA activity in a Ca²⁺-dependent manner due to enhanced CPA/TBQ-evoked Ca²⁺ leak (see Fig. 7). Such Ca²⁺- and time-dependent modulation of the PMCA has been observed previously in other cells (3) and is thought to be due to increased CaM binding (13). This Ca²⁺-dependent modulation of the PMCA makes physiological sense; low-level oxidants increase PMCA activity in an attempt to restore low resting [Ca²⁺]_i in the face of other impaired Ca²⁺ transport pathways, thereby preventing cell injury. However, as oxidative stress increases further, the PMCA becomes irreversibly inactivated, leaving the cell unable to restore resting [Ca²⁺]_i and thereby leading to "Ca²⁺ overload" and thus substantial cell injury.

It is not surprising that millimolar concentrations of H₂O₂ produced such pathologically clear-cut responses. However, the striking observation was that lower concentrations of H₂O₂ (10–100 µM) produced markedly heterogeneous responses such that there was a population shift in favor of the most severe [Ca²⁺]_i response with increasing concentrations of H₂O₂. Furthermore, the degree of impairment of [Ca²⁺]_i signaling in any given cell was directly proportional to the increase in cellular oxidative stress, suggesting that the heterogeneity was due to differences in cellular antioxidant capacity. This suggests that the concentration of H₂O₂ that produces any given response is less important than the proportion of cells that exhibit that response. This has important pathological implications, because only a fraction of cells with low antioxidant capacity may be sufficient to cause significant tissue damage within the whole organ in the face of an oxidant insult. This is because these highly sensitive cells would likely undergo an irreversible Ca²⁺ overload in response to low-level oxidant stress, causing cell lysis and necrotic cell death. This may cause inflammatory cytokine release and infiltration of activated neutrophils that further release oxidants (22, 50), culminating in a spiral of self-perpetuating cytotoxic tissue damage and ultimately organ failure, the hallmarks of severe acute pancreatitis (38). It is tempting to hypothesize that low level oxidants released from within acinar cells (7, 17, 42) could conceivably initiate pancreatitis if the antioxidant capacity of those cells or neighboring cells is low. It is also of interest that a lowered pancreatic antioxidant capacity has been suggested to be a predisposing factor for chronic pancreatitis (4).

The major source of the oxidant-evoked [Ca²⁺]_i increase in the present study is likely to be due to Ca²⁺ leak from agonist-sensitive Ca²⁺ stores, such as the ER. The is because pretreatment with H₂O₂ enhanced the CPA/TBQ-evoked [Ca²⁺]_i increase and the direct H₂O₂-evoked increase in [Ca²⁺]_i was completely abolished under conditions where ER Ca²⁺ was depleted and Ca²⁺ influx was prevented. However, under similar conditions if cells were first stimulated with CCK, which evokes oscillatory [Ca²⁺]_i signals and thus substantial mitochondrial Ca²⁺ uptake (17), H₂O₂ caused mitochondrial Ca²⁺ release that was abolished by the mitochondrial uncoupler, CCCP. This is consistent with other studies that showed oxidant-evoked mitochondrial depolarization and release of mitochondrial Ca²⁺ via the opening of the permeability transition pore (15, 16, 34). However, in naive unstimulated resting cells, where mitochondrial Ca²⁺ uptake does not occur, mitochondria are unlikely to be a major source of the oxidant-evoked [Ca²⁺]_i increase.

The exact nature of the H₂O₂-evoked ER Ca²⁺ release in rat pancreatic acinar cells is unclear from the present study. We observed [Ca²⁺]_i oscillations in response to another oxidant, thimerosal (Fig. 2, inset), in rat pancreatic acinar cells and also in response to low concentrations of H₂O₂ in mouse pancreatic acinar cells (3–30 µM, data not shown), consistent with other studies (16, 44). However, we never observed [Ca²⁺]_i oscillations in response to H₂O₂ at any of the concentrations tested (10 µM-1 mM) in rat pancreatic acinar cells, suggesting that the nature of the response to oxidants depends on type of oxidant and the species in which it is tested. In fact, the time course of the H₂O₂-evoked increase in [Ca²⁺]_i in rat pancreatic acinar cells was more reminiscent of a slow leak of Ca²⁺ from the ER, caused either by inhibition of SERCA (26, 29), or a direct effect on the passive leak pathway, due most likely to the translocon complex (27). However, further work is required to verify this mechanism and is beyond the scope of this study.

The H₂O₂-evoked increase in [Ca²⁺]_i was also dependent on Ca²⁺ influx, since the response was markedly attenuated by removal of external [Ca²⁺]. However, this is likely due to depletion of ER Ca²⁺ and an indirect activation of SOCE. A direct effect on SOCE channels is unlikely since addition of H₂O₂ failed to affect the rate of [Ca²⁺]_i increase when external Ca²⁺ was added back to ER-depleted cells, an assay routinely used as an indirect measure of SOCE (36). Notwithstanding this, a direct effect of H₂O₂ on noncapacitative Ca²⁺ influx pathways cannot be completely ruled out.

An alternative possibility is an effect of cellular ATP depletion, which is classically viewed as one of the earliest changes in oxidative injury (41). Depletion of 90% of cellular ATP has been reported to increase basal [Ca²⁺]_i and inhibit [Ca²⁺]_i oscillations in pancreatic acinar cells (45). However, studies on cultured intestinal epithelial cells suggest that even high millimolar concentrations of H₂O₂ do not deplete cellular ATP significantly over the first few minutes of exposure (49). In addition, ³¹P NMR measurements have shown that intracellular ATP levels in rat pancreas change by no more than 20% even during maximal agonist stimulation, when metabolic energy demand is presumably high (28). Given that H₂O₂ increased [Ca²⁺]_i almost immediately, albeit relatively slowly, it seems unlikely that global ATP depletion is the primary mechanism for the increase in [Ca²⁺]_i or inhibition of the PMCA, although local changes in ATP cannot be ruled out.

Low concentrations of H₂O₂ (10–100 µM) also profoundly altered CCK-evoked [Ca²⁺]_i oscillations. As indicated above, ER store depletion and inactivation of the PMCA are also likely explanations for the most severe cases of Ca²⁺ overload under these conditions. In addition, under the conditions of these experiments, impaired mitochondrial function likely contributes to the impaired CCK-evoked [Ca²⁺]_i oscillations consistent with other studies (16). Furthermore, the oxidant, tert-butylhydroperoxide, was also shown to impair carbachol-evoked [Ca²⁺]_i oscillations, which fused into a sustained response in pancreatic acinar cells (43). These observations also corresponded to a marked impairment of agonist-evoked fluid and amylase secretion from the perfused pancreas (43).

Another interesting observation from the present study was that there were striking differences in the proportion of cells showing moderate to severe damage when H₂O₂ was applied during CCK-evoked [Ca²⁺]_i oscillations compared with when H₂O₂ was added to resting cells. In particular, 50 µM H₂O₂ produced a Ca²⁺ overload response in approximately half of resting cells compared with none of the stimulated cells. These observations may be important pathologically, because they suggest that cells are more protected from the effects of H₂O₂ when stimulated with CCK. To our knowledge, this is a novel finding that suggests either that CCK couples to or enhances an antioxidant pathway within the cells, or that some aspect of the Ca²⁺ signaling machinery confers use-dependent protection from oxidant attack. However, further work is required to fully elucidate the mechanism, which is clearly beyond the scope of this study.

In summary, the present study shows that lower concentrations of H₂O₂ transiently increased [Ca²⁺]_i due in the most part to depletion of ER [Ca²⁺] and activation of Ca²⁺ influx (SOCE). In addition, H₂O₂ also profoundly altered the normal pattern of CCK-evoked [Ca²⁺]_i oscillations, reducing the amplitude and frequency of oscillations that are sometimes superimposed over a rising baseline. Disruption of mitochondrial Ca²⁺ handling is also likely to contribute to these responses. Similarly, the enhanced ER Ca²⁺ leak evoked by pretreatment with these low concentrations of H₂O₂ also indirectly increased the activity of the PMCA, which is the last gatekeeper for the control of low resting [Ca²⁺]_i. During oxidant attack, we suggest that the PMCA works hard to maintain [Ca²⁺]_i homeostasis in the face of disregulated Ca²⁺ transport pathways. However, at a critical H₂O₂ concentration (depending on the antioxidant capacity of the cell), PMCA activity rapidly and irreversibly declined. This likely converts the cell from a mildly effected cell, that can maintain [Ca²⁺]_i homeostasis to some degree, to a severely effected cell, where [Ca²⁺]_i is uncontrolled and thus irreversibly elevated. This mechanism may therefore be important in facilitating the Ca²⁺ overload, thereby resulting in a spiral of self-perpetuating cellular injury.

Finally, the effects of lower concentrations of H₂O₂ gave rise to a large degree of cellular heterogeneity that was likely due to differences in cellular antioxidant capacity. This further illustrates that the transition from a "mildly" effected to a "severely" effected cell potentially need only occur in a fraction of cells with low antioxidant capacity to trigger a cytotoxic inflammatory response. Inactivation of the PMCA may be a critical mechanism underlying these events and may well be important during the pathology of pancreatitis.

	GRANTS

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This work was supported by the Biotechnology and Biological Sciences Research Council (BBSRC) and the Wellcome Trust.

	FOOTNOTES

 

Address for reprint requests and other correspondence: J. I. E. Bruce, Faculty of Life Sciences, 2nd Floor Core Technology Facility, 46 Grafton St., The Univ. of Manchester, Manchester M13 9NT, UK (e-mail: jason.bruce{at}manchester.ac.uk)

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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