(Received for publication, April 30, 1997, and in revised form, May 14, 1997)
From the W. Alton Jones Cell Science Center,
Lake Placid, New York 12946, the ¶ Department of
Chemistry, Clarkson University, Potsdam, New York 13676, and
§ Division of Toxicology, Leiden Amsterdam Center for
Drug Research, Leiden University, Leiden, The Netherlands
Activation of stress response genes can impart cellular tolerance to environmental stress. Iodoacetamide (IDAM) is an alkylating toxicant that up-regulates expression of hsp70 (Liu, H., Lightfoot, D. L., and Stevens, J. L. (1996) J. Biol. Chem. 271, 4805-4812) and grp78 in LLC-PK1 renal epithelial cells. Therefore, we used IDAM to determine the role of these genes in tolerance to toxic chemicals. Prior heat shock did not protect cells from IDAM but pretreatment with trans-4,5-dihydroxy-1,2-dithiane (DTTox), thapsigargin, or tunicamycin enhanced expression of the endoplasmic reticulum (ER) chaperones GRP78 and GRP94 and rendered cells tolerant to IDAM. Cells expressing a 524-base pair antisense grp78 fragment (pkASgrp78) had a diminished capacity to up-regulate grp78 and grp94 expression after ER stress. Protection against IDAM due to prior ER stress was also attenuated in pkASgrp78 cells suggesting that ER chaperones of the GRP family are critical for tolerance. Covalent binding of IDAM to cellular macromolecules and depletion of cellular thiols was similar in tolerant and naïve cells. However, DTTox pretreatment blocked the increases in cellular Ca2+ and lipid peroxidation observed after IDAM treatment. Overexpressing the ER Ca2+-binding protein calreticulin prevented IDAM-induced cell death, the rise in cytosolic Ca2+, and oxidative stress. Although activation of the ER stress response did not prevent toxicity due to Ca2+ influx, EGTA-AM and ruthenium red both blocked cell death suggesting that redistribution of intracellular Ca2+ to the mitochondria may be important in toxicity. The data support a model in which induction of ER stress proteins prevents disturbances of intracellular Ca2+ homeostasis, thus uncoupling toxicant exposure from oxidative stress and cell death. Multiple ER stress proteins are likely to be involved in this tolerance response.
Exposing cells to environmental stress induces expression of stress proteins in various intracellular compartments including the cytoplasm and the ER1 (1-6). In addition, prior treatment with a mild insult that is sufficient to induce stress protein expression renders cells tolerant to a subsequent lethal insult (5, 7). For example, inducing HSPs with mild heat shock treatment confers thermotolerance as well as resistance to damage by cytokines, ischemic injury, and chemicals (8-10). The glucose-regulated proteins (GRPs), a family of molecular chaperones and Ca2+-binding stress proteins located in the endoplasmic reticulum (ER), are also induced by stress (4, 5). Induction of GRPs by ER stress protects cells against a variety of toxic insults including Ca2+ ionophores, oxidative stress, topoisomerase inhibitors, and cytotoxic T-cells (11-19). Thus, multiple stress proteins may be important in the cellular tolerance response.
Chemical toxicants including heavy metals, halogenated hydrocarbons, chemotherapeutic agents, or antibiotics induce stress proteins (1, 3, 5, 6, 20, 21), yet the mechanism(s) by which such a stress response prevents chemical damage in the target organs for these toxicants is not clear. The kidney proximal tubular epithelium is a particularly important target, and much is known about mechanisms of chemically induced cell death in kidney (22) and other cell types (23-26). In general, toxicant exposure initiates a cascade of biochemical events that ultimately cause cell death. For instance, exposing kidney epithelial cells to toxicants that are metabolized to reactive intermediates results in covalent binding of the metabolites to cellular macromolecules, depletion of cellular protein and nonprotein thiols, e.g. glutathione (GSH), increased intracellular Ca2+ concentrations, collapse of the mitochondrial membrane potential, and generation of reactive oxygen species (27-33). In LLC-PK1 cells, blocking any of these events with pharmacological agents blocks the toxicity of reactive metabolites and other toxicants (27-29, 34, 35). Taken together, these biochemical perturbations constitute a sequential and highly interrelated cytotoxic signaling cascade that results in cell death.
Despite the integration of the cell death cascade, activation of stress response genes in kidney epithelial cells is linked to specific perturbations suggesting that discrete signals within the cell death pathway are linked to specific genomic responses. For example, activation of hsp70 expression by iodoacetamide (IDAM) or the nephrotoxicant S-(1,2-dichlorovinyl)-L-cysteine is caused by oxidation or depletion of protein and nonprotein thiols and not directly by the covalent binding, Ca2+ disturbances, or oxidant production that also occur as part of the cell death pathway (21, 36). On the other hand, c-myc mRNA induction by S-(1,2-dichlorovinyl)-L-cysteine appears to be linked, at least in part, to an the increase in cellular free Ca2+ levels (37). Alkylation of cellular macromolecules may be sufficient to induce expression of c-fos and gadd153 (37, 38). Thus, biochemical perturbations caused by toxicant exposure serve both as discrete signals that activate specific stress response genes and as integrated components of a cell death pathway.
Intracellular Ca2+ homeostasis has received considerable attention as a cell death signal and as an activator of gene expression, yet consensus has not emerged regarding its role in either process (25, 26, 39, 40). Nevertheless, maintaining intracellular free Ca2+ levels at about 100 nM in the face of 1-2 mM extracellular Ca2+ is important for cell survival, and toxicant treatment generally causes an increase in free Ca2+ levels (26, 39). Membrane pumps in the ER, mitochondria, and plasma membranes work in concert to maintain intracellular Ca2+ levels (41, 42). Failure of Ca2+ pumping at any of these sites could contribute to an increase in free Ca2+ (26, 43). At physiological intracellular Ca2+ concentrations, the ER is a major intracellular Ca2+ storage site in nonmuscle cells (41, 42), and high lumenal Ca2+ is essential for normal ER function (44-46). Abundant ER Ca2+-binding proteins, including GRP78, GRP94, calreticulin, and calnexin, may help sequester ER Ca2+ (47-50). For example, calreticulin provides up to 45% of the Ca2+ buffering capacity in the inositol 1,4,5-trisphosphate-sensitive Ca2+ pool (51) and facilitates protein processing in the ER (52). Increasing or decreasing calreticulin expression also modulates physiological Ca2+ release from the hormone-sensitive pool (51, 53-55). Thapsigargin or calcium ionophores deplete ER Ca2+ thereby inhibiting ER protein processing and cellular protein synthesis in general (45, 46, 56, 57). Induction of ER chaperones renders cells tolerant to Ca2+ depletion (4, 5, 19, 56). Thus, a general increase in cellular Ca2+ and/or depletion of intracellular Ca2+ stores can cause cell death. Because ER chaperones are important both in cellular tolerance and in regulating cellular Ca2+, it seems possible that ER stress might protect cells by helping maintain cellular Ca2+ homeostasis.
The goal of these studies was to address the role of stress proteins in tolerance to chemical damage using the alkylating toxicant IDAM and the renal epithelial cell line LLC-PK1 as a model. These cells have been used extensively to investigate cytotoxicity and stress gene activation (21, 27, 28, 36-38, 58, 59). Herein, we show that conditioning LLC-PK1 cells with mild ER stress, but not heat shock, increases expression of ER stress proteins and prevents IDAM-induced cell death. Increasing expression of ER stress proteins apparently helps control intracellular Ca2+ levels following IDAM exposure preventing oxidative stress. The results provide new insights into the role of ER stress proteins in cellular Ca2+ homeostasis and cell death as well as in tolerance to chemical damage.
Fetal bovine serum and Dulbecco's modified
Eagle's medium (DMEM) were obtained from Life Technologies, Inc.
LLC-PK1 cells, a porcine renal epithelial cell line with proximal
tubule epithelial characteristics (60, 61), were obtained from American
Type Culture Collection (Rockville, MD) at passage 195 and were used from passage 205-215.
N,N-Diphenyl-p-phenylenediamine (DPPD) was
obtained from Eastman Kodak. The acetoxymethyl ester of EGTA (EGTA-AM)
and Fura-2 (Fura-2AM) and Pluoronic F-127 were purchased from Molecular
Probes (Eugene, OR). Radiochemicals were obtained from NEN Life Science
Products. All other chemicals were obtained from commercial
sources.
Cell culture and treatment of LLC-PK1 cells with IDAM were carried out as described (27, 36). LLC-PK1 cells were maintained in Dublecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (complete medium). Confluent LLC-PK1 cells were treated with IDAM for 15 min in Earle's balanced salt solution (EBSS), then washed with phosphate-buffered saline (PBS), and allowed to recover in complete medium. Where appropriate, the antioxidant DPPD, prepared as a 20 mM stock in ethanol, was added to the medium at a concentration of 20 µM during the treatment period and/or during the recovery period. Cell were treated with DTTox (10 mM) for 2 or 3 h in EBSS and returned to complete medium for 12 h. Cells treated for 12 h in complete medium containing thapsigargin (0.3 µg/ml) or tunicamycin (1.5 µg/ml) were washed with PBS and returned to complete medium. For heat shock treatment, confluent LLC-PK1 cells in 10-cm dishes were incubated for 1 h in a water bath maintained at 43 or 45 °C in a humidified incubator at the same temperature and then returned to 37 °C for either 12 or 24 h.
Cytotoxicity, determined by measuring release of lactate dehydrogenase (LDH), covalent binding of [14C]IDAM to cellular macromolecules, as well as depletion of protein and nonprotein thiols were measured as described (36). Lipid peroxidation was determined by the formation of thiobarbituric acid-reactive substances (TBARS) as before (27).
Preparation of Antisense grp78 CellsAn antisense
grp78 expression vector was constructed in pcDNA3
(Invitrogen). A 524-base pair fragment from a hamster grp78 cDNA (62), a gift from Dr. Amy Lee, was digested with
NaeI (+145 to +669) and inserted into the EcoRV
site of pcDNA3 in a 3 to 5
orientation to create the antisense
grp78 expression plasmid pASgrp78. pASgrp78 or the
pcDNA3 empty vector was transfected into LLC-PK1 cells using
Lipofectin (Life Technologies, Inc.), and a mass culture of cells that
expressed the 0.5-kb antisense RNA (pkASgrp78 cells) was selected in
800 µg/ml G418 (Sigma) and maintained in 500 µg/ml G418. Multiple
clones of pkASgrp78 were selected from the mass culture by ring
cloning. Empty vector clones, termed pkNEO cells, were selected at the
same time. Five pkASgrp78 clones were screened further for expression
of GRPs following DTTox treatment by [35S]methionine and
[35S]cysteine metabolic labeling (see below). Bands on
autoradiograms representing 35S-labeled GRP78 were
quantitated by densitometric scanning using a BioImage Densitometer
(BioImage, Ann Arbor, MI) as described previously (36). The integrated
optical densities were normalized by taking the ratio of the GRP78 and
actin signals in each lane, and the data were expressed as the fold
increase in GRP78 relative to untreated cells. Three clones,
pkASgrp78-5, -8, and -10, showed markedly reduced GRP78 synthesis and
were further tested for the presence of 0.5-kb grp78
cDNA fragment by Southern blot analysis. Genomic DNA (20 µg) was
digested with ApaI and BamHI; fragments were
separated by electrophoresis, transferred to nitrocellulose membranes,
and blotted with a hamster grp78 cDNA probe according to
standard procedures. In experiments in which the response of pkNEO and
pkASgrp78 clones was compared, three pkNEO clones, 2, 9, and 10, were
compared with three pkASgrp78 clones, 5, 8, and 10. The response for
the individual clones was determined in at least two separate
experiments, and the mean of each clone was used as a single data point
to calculate the mean of the three clonal lines.
An expression vector, pRC/CMV, containing a full-length (1.9 kb) human calreticulin cDNA (63) was provided by Dr. S. Dedhar. After transfection, calreticulin overexpressing cells (pkCRT) were selected for G418 resistance and were ring cloned as described above. Again, pkNEO cells were selected under identical conditions. Individual clones were tested for the expression of calreticulin by immunofluorescence and Western blot analysis using an antibody against calreticulin (StressGen, Vancouver, British Columbia). Clones overexpressing calreticulin were analyzed further for sensitivity to IDAM. Biological responses in three pkNEO clones, 1, 2, and 3, were compared with the pkCRT clones, 2, 3, and 5, as described above for pkASgrp78 cells.
Measurement of Intracellular CalciumIntracellular free
Ca2+ was determined with the Ca2+-sensitive
fluorescent dye Fura-2 according to Chen et al. (28) with
modifications. Cells grown on coverslips coated with bovine collagen
type I were rinsed with PBS and loaded with Fura-2AM in EBSS to achieve
a final concentration of 3 µM. A 1:1,000 (v/v) dilution
of 20% Pluoronic F-127 was added to EBSS to dissolve Fura-2AM and
facilitate cell loading. In addition, probenecid, an inhibitor of
organic ion transport, was included at a concentration of 2 mM to prevent intracellular transport or extrusion of
Fura-2 free acid (33). Loading with Fura-2 was carried out at room
temperature. After loading cells with Fura-2AM for 1 h, cells were
washed four times with EBSS in the presence of 2 mM
probenecid to prevent leakage. The coverslips were positioned in a
quartz cuvette containing 3.5 ml of EBSS with probenecid for
fluorescence analysis using a Shimadzu RF-5000 spectrofluorophotometer
(Shimadzu, Columbia, MD). The calcium concentration was calculated as
Kd (224 nM)× (R Rmin)/(Rmax
R) according to Grynkiewicz et al. (64) as
described previously (28). R is the ratio
(F1/F2) of the fluorescence at excitation (ex) 340 nm, emission 505 nm over that of
the fluorescence at excitation 380 nm. In some experiments, Ca2+ concentrations were also determined using digital
fluorescence imaging as described (30).
When spectrofluorometric measurements were used to quantitate
intracellular free Ca2+, the distribution of Fura-2 between
the cytosol and intracellular compartments was determined in cells
loaded as described above. Cytoplasmic Fura-2 was released by adding
buffer A (250 mM sucrose, 20 mM KCl, 3 mM EGTA, 10 mM K2HPO4,
5 mM MgCl2, 5 mM succinate) containing 50 µM digitonin for 5 min to permeabilize the
plasma membrane. The supernatant was collected, and the cells were
lysed with 0.1% Triton X-100 in buffer A. Fura-2 fluorescence in the digitonin (cytosolic Fura-2) and Triton X-100 fractions (total remaining) were monitored at the calcium-independent wavelength ex = 362 nm. Using this procedure, we found that over
75% of the Fura-2 was in the cytosol, i.e. released by
digitonin.
Preparation of mRNA was carried out as described
previously (21). cDNA probes were labeled with
[32P]dCTP (NEN Life Science Products) by random priming
using a kit (Boehringer Mannheim). Blots were probed with a hamster
grp78 cDNA probe and then with -actin cDNA as an
internal control. Western blot analysis for stress-inducible HSP70,
also called HSP72, was carried out essentially as described (36) using
a monoclonal antibody (Amersham Corp.). For detection of calreticulin, anti-calreticulin polyclonal antibody (StressGen) was used.
Nitrocellulose membranes were blocked with 5% nonfat milk and probed
with antibody in the presence of 5% nonfat milk. Detection of
endogenous calreticulin by immunoblotting required an anti-calreticulin
antibody dilution of 1:250, but with overexpressing cells a 1:5000
dilution was used. Appropriate secondary antibodies and the enhanced
chemiluminescence system (Amersham Corp.) were used to develop the
blots.
Immunofluorescence analysis of calreticulin was done using the same
polyclonal anti-calreticulin antibody. Confluent cells on
collagen-coated glass coverslips were rinsed in PBS and fixed with
methanol at 20 °C for 10 min. After blocking with 2% horse serum
in PBS for 45 min, the coverslips were incubated for 1 h with
anti-calreticulin antibody (1:50) followed by dichlorotriazinyl aminofluorescein-conjugated goat anti-rabbit IgG (Jackson
ImmunoResearch, West Grove, PA), diluted 1:250 in PBS containing 1%
bovine serum albumin. Coverslips were mounted on slides and observed
with a Nikon episcopic fluorescence microscope using a 60 × objective.
Porcine GRP78 did not cross-react with any available GRP78 antibodies tested; therefore, increased synthesis of stress proteins was determined by [35S]methionine and [35S]cysteine labeling. For short term labeling, confluent LLC-PK1 cells were incubated with methionine- and cysteine-free DMEM for 20 min followed by a 1-h incubation with [35S]methionine and [35S]cysteine (100 µCi/ml) in methionine- and cysteine-free DMEM. For the long term labeling, cells were incubated with [35S]methionine and [35S]cysteine (50 µCi/ml) in normal DMEM for 4 h. After radiolabeling, cells were lysed in hypotonic buffer (0.25 M sucrose, 25 mM Tris, pH 7.4, 2.5 mM magnesium acetate, 2.0 mM dithiothreitol), and proteins were solubilized in SDS sample preparation buffer. Radiolabeled proteins were resolved by SDS-polyacrylamide gel electrophoresis and protein bands visualized by autoradiography.
Statistical AnalysesStudent's t test was used to determine if there was a significant difference between the two groups (p < 0.05). When multiple means were compared, significance (p < 0.05) was determined by ANOVA followed by the Student-Newman-Keul's test. For ANOVA analysis, letter designations are used to indicate significant differences. Means with a common letter designation are not different, and those with a different letter designation are significantly different from all other means with different letter designations. Means with more than one letter designation are not different from groups with either letter designation. In cases where statistical analysis is shown for two different parameters in a single table or figure, i.e. Ca2+, thiobarbituric acid-reactive substances or LDH release, letters indicating significant differences apply only within that measurement group.
IDAM treatment
increases expression of hsp70 in LLC-PK1 cells (36). Since
induction of HSP expression is linked to tolerance, we evaluated the
effect of heat shock on IDAM cytotoxicity. Although heat shock induced
HSP70 in LLC-PK1 cells (data not shown), it did not protect against
IDAM-induced cell death (Fig. 1). IDAM treatment also increased expression of the mRNA for prototypical ER
stress protein grp78 in a time- and
concentration-dependent manner (Fig.
2). Treating cells with DTTox,
tunicamycin, or thapsigargin, agents that cause ER stress (5, 65),
increased mRNA for grp78 and synthesis of both GRP78 and
GRP94 proteins (Fig. 3, A and B) in LLC-PK1 cells. Pretreatment with all three agents
prevented IDAM-induced cell death without altering
[14C]IDAM covalent binding to macromolecules (Fig.
3C). There was also a good correlation between the peak of
GRP78 and GRP94 biosynthesis and the onset of the tolerant phenotype
after DTTox treatment (Fig. 4,
A and B). The cells maintained the tolerant
phenotype up to 24 h, probably due to the long half-life (>36 h)
of ER stress proteins such as GRP78 (18). Thus, conditioning cells with
ER stress protected them against IDAM toxicity without affecting toxicant entry and covalent binding.
Blocking Expression of grp78 Disrupts the ER Stress Response and Tolerance
Antisense and ribozyme strategies directed against
grp78 and grp94, respectively, have been
effective in probing the role of ER stress proteins in tolerance and
protein secretion (12, 13). Selective targeting of grp78
with antisense interferes with induction of both grp78 and
grp94 and disrupts the ER stress response (12). We targeted
grp78 using a 0.5-kb antisense grp78 fragment
that spanned the translation start site. After transfection, G418-resistant pkASgrp78 clones were tested for induction of GRP78 and
GRP94. In pkASgrp78 clones, GRP78 synthesis after DTTox treatment was
attenuated compared with empty vector pkNEO clones (Fig.
5, A and B). All
the pkASgrp78 clones had integrated the antisense fragment (Fig.
5C). Although it appeared that induction of
35S-labeled GRP94 was also reduced (Fig. 5B),
the band could not be quantitated accurately by densitometry due to its
proximity to other bands.
The pkASgrp and pkNEO clones were tested for IDAM sensitivity. Covalent
binding of [14C]IDAM was equivalent in pkASgrp78 and
pkNeo cells, 407 ± 113 versus 448 ± 14 pmol/mg
protein, respectively, indicating that both took up IDAM equally well.
LDH release 1-2 h after IDAM treatment was higher in pkASgrp78 clones
compared with pkNeo clones, but there was no difference in maximum LDH
release observed at 6 h (Fig.
6A). Unlike pkNEO cells,
pkASgrp78 cells had a reduced capacity to develop tolerance after DTTox
(Fig. 6B), nor did they develop tolerance after treatment
with thapsigargin and tunicamycin (Fig. 7). Thus, expression of grps
is important for tolerance to IDAM. The data clearly suggest that GRP78
is important in the ER stress response and cytoprotection, but we
cannot exclude a role for GRP94 as well.
ER Stress Prevents Ca2+ Accumulation and Oxidative Stress
Having established a role for ER stress proteins in cellular tolerance, we went on to investigate the mechanism of protection. As shown in Fig. 3C, and in previous reports (29, 36), IDAM covalently modifies cysteinyl thiol groups in proteins. However, IDAM also elicits secondary effects in LLC-PK1 cells including depletion of GSH and oxidation of protein thiols (29, 36). Since ER stress did not affect covalent binding of [14C]IDAM (Fig. 3C), we determined if it diminished thiol-disulfide redox perturbations. However, depletion of cellular nonprotein and protein thiols after IDAM treatment was not altered by DTTox (Table I). Similar results were obtained in cells rendered tolerant by thapsigargin or tunicamycin treatment (data not shown).
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Elevation of cytosolic Ca2+ is important in
toxicant-induced cell death in renal epithelial cells (28, 30, 37), and
other cell types (26, 39). Therefore, we determined if the cellular free Ca2+ surge observed after IDAM treatment was
attenuated in tolerant cells. There was a sustained increase in
intracellular free Ca2+ after IDAM treatment (Fig.
8, and data not shown). DTTox
pretreatment blocked the increase in intracellular Ca2+.
Lipid peroxidation also increased within 30 min after IDAM treatment followed by LDH release; both were prevented by DTTox pretreatment (Fig. 9). Thus, conditioning cells with
ER stress blocked the IDAM-induced Ca2+ surge, lipid
peroxidation, and cell death.
Loss of membrane integrity due to lipid peroxidation can cause
extracellular Ca2+ influx (40). If this were the case, then
the antioxidant,
N,N-diphenyl-p-phenylenediamine (DPPD), which blocks lipid peroxidation after IDAM treatment (29), should block Ca2+ entry. DPPD treatment blocked much of the
increase in intracellular Ca2+; however, Ca2+
still increased 3-fold from 64 to 189 nM (Fig.
10). When DPPD and DTTox treatments
were combined, Ca2+ remained at control levels (Fig. 10).
Removing extracellular Ca2+ also prevented the increase in
free Ca2+ after IDAM treatment (data not shown), consistent
with a role for oxidative stress in influx of extracellular
Ca2+.
Increased Expression of Calreticulin Prevents IDAM Cytotoxicity
The data suggested that there might be a connection
between ER stress, induction of Ca2+ binding chaperone
proteins, and blockade of an IDAM-induced Ca2+ surge linked
to oxidative stress. If the mechanism underlying this effect was
dependent on an increase in Ca2+-binding proteins in the
ER, then artificially increasing the level of ER
Ca2+-binding proteins might produce the same effect.
Overexpression of calreticulin, the major ER Ca2+-binding
protein in nonmuscle cells (49), has been shown to increase ER
Ca2+ stores and to modulate ER Ca2+ release
(53, 55); therefore, we determined the effect of calreticulin
overexpression on IDAM toxicity. We prepared three clones of LLC-PK1
cells, designated pkCRT-2, -3, and -5, all of which expressed high
levels of calreticulin (Fig.
11A) in the ER (Fig.
11B). Compared with pkNEO cells, pkCRT cells were less
sensitive to IDAM-induced cell death (Fig. 11C), although
covalent binding of [14C]IDAM was unchanged;
i.e. pkCRT, 398 ± 11 pmol/mg/protein; pkNEO clones,
399 ± 14 pmol/mg protein. Thus, enforced expression of calreticulin produced a tolerant phenotype indicating that ER proteins
other than GRP78 could participate in cellular tolerance. Although we
could not determine GRP78 levels by Western blotting due to a lack of
antibodies (see "Experimental Procedures"), CRT expression did not
alter the basal level of GRP94 (data not shown), indicating that CRT
expression may not have a global effect on other ER stress
proteins.
We also determined the effect of calreticulin overexpression on intracellular Ca2+ and oxidative stress after IDAM treatment (Table II). Without IDAM treatment, there was no difference in resting Ca2+ levels in pkCRT and pkNEO clones. However, after IDAM treatment, there was a significant increase in intracellular Ca2+ in pkCRT cells, but not nearly to the level seen in pkNEO cells. In addition, lipid peroxidation was prevented in pkCRT but not pkNEO clones after IDAM exposure. Thus, overexpression of calreticulin blocked the IDAM-induced increase in intracellular Ca2+ and oxidative stress indicating that the presence of Ca2+-binding proteins in the ER was important in preventing both responses.
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Although prior ER stress blocked the increase in cellular Ca2+ and prevented oxidative stress, much of the Ca2+ surge was due to entry from the extracellular pool, i.e. outside-in Ca2+ flux. To address the role of Ca2+ influx in IDAM toxicity, we compared the effect of removing extracellular Ca2+ on cell death caused by treatment with IDAM or the Ca2+ ionophore, ionomycin. Removing extracellular Ca2+ blocked cell death caused by the Ca2+ ionophore ionomycin but had no effect on IDAM-induced cell death (Table III). We next determined if DTTox pretreatment or calreticulin overexpression had any effect on toxicity due to influx of extracellular Ca2+ caused by ionomycin. pkCRT cells were less sensitive to ionomycin, indicating that pkCRT cells had an enhanced capacity to buffer extracellular Ca2+ (Fig. 12). However, the protection was not as dramatic as observed for IDAM (Fig. 11). DTTox treatment had no effect on ionomycin toxicity (data not shown).
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Several lines of evidence suggest that inside-out Ca2+ flux also might be important in cell death (39). Thapsigargin releases ER Ca2+ and causes apoptosis in LLC-PK1 cells, but prior ER stress blocks this response.2 Since increasing cytosolic Ca2+ results in mitochondrial Ca2+ uptake and increased oxidant production (30), efflux of ER Ca2+ could stimulate mitochondrial oxidant production providing an inside-out mechanism of Ca2+ flux in cell death. Agents that buffer intracellular Ca2+ (EGTA-AM) or prevent mitochondrial Ca2+ uptake (ruthenium red) prevent oxidative stress and cell death in renal epithelial cells (28, 30). Loading cells with EGTA, using EGTA-AM, or adding ruthenium red prevented IDAM-induced cell death (Table IV). Thus, a disturbance of the ER Ca2+ pool (inside-out signaling) may be more important in the cell death pathway than the influx of extracellular Ca2+ (outside-in signaling).
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A number of useful conclusions can be drawn from these studies. First, induction of ER stress proteins protects cells against alkylating chemicals. Preliminary studies show that ER stress also protected LLC-PK1 cells from nephrotoxic cysteine conjugates (65), t-butylhydroperoxide, and thapsigargin toxicity as well.3 Second, multiple ER proteins may be important since blocking induction of GRPs prevented tolerance while overexpressing calreticulin protected cells. To our knowledge, calreticulin has not been shown to play a role in tolerance to chemical damage. A role for GRP78 also seems clear, but GRP94 may play a role as well, nor can we exclude the possibility that altering expression of one ER stress protein has an indirect but significant effect on another. Third, ER tolerance depends in part on maintaining cellular Ca2+ homeostasis and preventing oxidative stress. Although the importance of ER Ca2+ in protein processing (45, 46, 57), translational control (56), and regulation of grp78 transcription (66) is well known, the role of the ER in regulating cellular Ca2+ homeostasis and oxidative stress after chemical damage has not been shown previously. Thus, our data shed new light on the role of the ER in control of cellular Ca2+ and cytotoxicity.
Our data also support a model of IDAM-induced cell death in which an
increase in cytoplasmic Ca2+ leads to mitochondrial
Ca2+ uptake, induction of oxidative stress, membrane
peroxidation, and cell death (Fig. 13),
a model supported by data from other studies in renal epithelial cells
(27, 28, 30). Mitochondria sense cytosolic Ca2+
fluctuations by accumulating Ca2+ and thus tune energy
production to meet the biological responses initiated by
Ca2+ signaling (41). However, Ca2+ buffering by
mitochondria must be coupled to extrusion across the plasma membrane
and/or re-uptake of Ca2+ into the ER, otherwise
mitochondria will accumulate a lethal load of Ca2+ (30, 67,
68). If the latter happens, membrane potential collapses, reduced
pyridine nucleotide pools are depleted, phospholipases are activated,
and large pores open in the mitochondrial inner membrane (67). When
cellular GSH has been depleted, mitochondrial Ca2+ overload
can cause excess oxidant production, oxidative stress, and plasma
membrane rupture. Thus, buffering intracellular Ca2+ and
preventing mitochondrial Ca2+ accumulation and/or cycling
blocks cell death following chemical exposure by uncoupling
Ca2+ perturbations from oxidative stress (30, 67).
Although we did not directly assess the changes in ER Ca2+ stores and the effect of chaperone expression on ER Ca2+ stores during toxicant treatment, it may be that the ability of the ER to release or buffer intracellular Ca2+ modulates cell death (Fig. 13). The ER is the major intracellular Ca2+ storage site in nonmuscle cells and could be a target for toxic damage. The ER Ca2+-ATPase is inhibited by carbon tetrachloride treatment in vivo (69), and toxicants have been shown to impair operation of ER Ca2+ release channels (26, 70, 71). Moreover, thapsigargin treatment causes apoptosis suggesting that loss of ER Ca2+ is a cell death signal (72-75). Interestingly, bcl-2 expression can block thapsigargin-induced cell death in some cells (73, 74). Mitochondria from bcl-2 overexpressing cells have an increased capacity to accumulate Ca2+ (68) indicating that there could be a link between ER and mitochondrial Ca2+ pools in cell death. In addition, toxicants that modify protein sulfhydryls release ER Ca2+ (70, 71) and prevent extrusion of Ca2+ across the plasma membrane (43) generally perturbing Ca2+ signaling. Depleting ER Ca2+ also disrupts ER protein processing and general protein synthesis and activates expression of grps genes (41, 45, 46, 66), effects that are blocked by expression of ER chaperones (5, 11, 19, 56). Thus, in naive cells, disrupting ER Ca2+ buffering may contribute to cell death, whereas inducing ER chaperones and Ca2+-binding proteins prevents cell death. Although it is not clear from our studies if the protection caused by induction of ER stress proteins is due to their ability to modulate ER Ca2+ stores directly or indirectly, it is apparent that induction of ER stress proteins helps control general intracellular Ca2+ homeostasis preventing toxicant-induced cell death.
It has been suggested that accumulation of intracellular Ca2+ is merely a secondary effect of membrane damage and influx from the extracellular pool (40). Indeed, in our studies removing extracellular Ca2+ or adding antioxidants blocked Ca2+ accumulation after IDAM treatment. However, this outside-in Ca2+ surge was not responsible for IDAM-induced cell death. Yet, buffering intracellular Ca2+ by treating cells with EGTA-AM prevented IDAM toxicity arguing that Ca2+ does play a role. Here again the model in Fig. 13 accounts for these observations since release of intracellular Ca2+ would lead to secondary influx from the extracellular pool due to oxidative stress and membrane damage (43). Cooperation between ER Ca2+ efflux and extracellular Ca2+ influx is well know during hormone-induced capacitative Ca2+ entry (76).
Although, we addressed toxicant-induced necrosis, our data may provide general support for an ER-mitochondrial Ca2+ axis in cell death. Disturbances in Ca2+ pumping in the ER and mitochondria also cause apoptotic as well as necrotic cell death (26, 39). bcl-2 prevents this apoptosis, perhaps by increasing the capacity of mitochondria to buffer Ca2+ (68). This connection between bcl-2 and mitochondrial Ca2+ is particularly important given that mitochondrial damage is linked to activation of proapoptotic protease cascades (77, 78). Our recent finding that prior ER stress or overexpression of calreticulin protects against thapsigargin-induced apoptosis in LLC-PK1 cells2 further supports the notion that efflux of ER Ca2+ is an early event in cell death. Even if this model does not hold for all cells (79), when taken in context, our data point toward a link between ER and mitochondrial Ca2+ handling and cell death signals.
We thank Drs. Amy Lee, Randy Kaufman, and Sri Prakash Srivastava for discussing unpublished studies, for helpful comments, and for providing reagents. We also thank Drs. John Subjeck, Martin Tenniswood, Denry Sato, and Susan Jaken as well for helpful discussions and Ellen Miller for technical assistance. Special thanks to Margaretann Halleck, Senait Asmellash, and to members of the laboratory for continued comments and support.