Cyclosporin A inhibits chromium(VI)-induced apoptosis and mitochondrial cytochrome c release and restores clonogenic survival in CHO cells

Daryl E. Pritchard1,2,*, Jatinder Singh1, Diane L. Carlisle1,3 and Steven R. Patierno1,2,3,4

1 Department of Pharmacology,
2 Program in Genetics and
3 Program in Molecular and Cellular Oncology, The George Washington University Medical Center, 2300 Eye Street NW, Washington, DC 20037, USA


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
A variety of key events in the molecular apoptotic pathway involve the mitochondria. Cyclosporin A (csA) affects the mitochondria by inhibiting the mitochondrial permeability transition (MPT), thereby preventing disruption of the transmembrane potential. The role of the MPT in apoptosis is not fully understood, but inhibition of the MPT may prevent the release of mitochondrial caspase activators, such as cytochrome c (cyt c), into the cytosol. Certain hexavalent chromium [Cr(VI)] compounds are known occupational respiratory tract toxins and carcinogens. We have previously shown that these compounds induce apoptosis as a predominant mode of cell death and that this effect can be observed in cell culture using soluble Cr(VI). We show here that Cr(VI)-induced apoptosis in Chinese hamster ovary (CHO) cells involves disruption of mitochondrial stability. Using a cyt c-specific monoclonal antibody, we observed a dose-dependent release of mitochondrial cyt c in cytosolic extracts of CHO cells exposed to apoptogenic doses of sodium chromate. Co-treatment of these cells with csA inhibited the release of cyt c and abrogated Cr(VI)-induced apoptosis as determined by a reduction in internucleosomal DNA fragmentation. Co-treatment with csA also markedly increased clonogenic survival of Cr(VI)-treated CHO cells. In contrast, the general caspase inhibitor Z-VAD-FMK markedly inhibited most of the morphological and biochemical parameters of apoptosis but did not prevent cyt c release and did not increase clonogenic survival. These results suggest that the MPT plays an important role in the regulation of mitochondrial cyt c release and that this may be a critical point in the apoptotic pathway in which cells are irreversibly committed to death.

Abbreviations: csA, cyclosporin A; cyt c, cytochrome c; DMSO, dimethyl sulfoxide; IDF, internucleosomal DNA fragmentation; MPT, mitochondrial permeability transition; PBS, phosphate-buffered saline; Z-VAD-FMK, Z-Val-Ala-Asp(OMe)-FMK.


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The apoptotic signaling pathway consists of a cascade of molecular events which lead from the initiation of cell death to eventual dismantling of the cell. The point at which the cell is irrevocably committed to death is unclear. Inhibitors of the proteolytic caspase cascade are able to block some or all of the morphological consequences associated with apoptosis, but they appear to be unable to maintain clonogenic or long-term survival (13). Cells exposed to pro-apoptotic stimuli in the presence of caspase inhibitors will either undergo terminal growth arrest or will die by some slower non-apoptotic mechanism of cell death (4). Upstream of the caspases, a variety of key events in apoptosis occur which involve the mitochondria, including loss of mitochondrial transmembrane potential, activation of the mitochondrial permeability transition (MPT), the participation of bcl-2 family proteins and the release of caspase activators. If a mechanism for commitment to cell death exists, it may involve these mitochondrial events.

Cytochrome c (cyt c) is released from the mitochondrial intermembrane space to the cytosol where it forms a complex with Apaf-1 and caspase-9, thereby triggering caspase-9 activation and initiating the caspase cascade (57). It has been suggested that cyt c release may occur in response to a collapse of the mitochondrial inner transmembrane potential (3). The loss of membrane potential indicates a MPT caused by opening of a large conductance MPT pore (8). Studies with isolated mitochondria show that the MPT pore favors a closed state, but certain physiological and pathological signals trigger pore opening (911). The fully opened state creates a channel for <=1.5 kDa molecules, resulting in dissipation of the H+ gradient across the membrane and uncoupling of the respiratory chain (12). Two models have been established which describe how MPT pore opening may effect cyt c release. In the first model, opening of the MPT pore may result in mitochondrial volume expansion, eventually causing outer membrane rupture and thereby releasing intermembrane caspase-activating proteins, such as cyt c. The alternative model suggests that the MPT may trigger a signal for the opening of an outer membrane channel which allows cyt c release without concomitant organellar swelling. Cyclosporin A (CsA), an immunosuppressant, can inhibit the MPT. CsA binds to Cyp-M, a cyclophilin-family protein associated with the MPT pore, causing it to dissociate from the pore complex (13,14). This reaction is believed to increase the probability of pore closure. In this study we have examined whether csA can block apoptosis and restore clonogenic survival to cells exposed to apoptogenic doses of hexavalent chromium [Cr(VI)].

Exposure to particulate Cr(VI) is strongly associated with lung toxicity and increased incidence of lung cancer (15,16). Soluble sodium chromate (Na2CrO4) can be used to study the effects of Cr(VI) in culture. Cr(VI) is transported into cells where it undergoes metabolic reduction to reactive genotoxic species (1719). This process leads to diverse genotoxic and cytotoxic effects, including oxidative DNA damage (20), DNA–DNA interstrand crosslinks (2123), Cr–DNA adducts (24,25), single-strand breaks (23), DNA–protein crosslinks (25), chromosomal aberrations (26,27), DNA polymerase arrest, RNA polymerase arrest (28) and inhibition of transcription and translation (29). Apoptosis is the predominant cellular response to Cr(VI) in Chinese hamster ovary (CHO) cells in culture (30,31). Therefore, we used CHO cells exposed to Na2CrO4 as a model for apoptotic induction caused by diverse genotoxic exposure.

In this study we show that Cr(VI) exposure causes a dose-dependent release of cyt c and we examine the effect of csA on cyt c release, apoptosis and clonogenic survival in Cr(VI)-exposed CHO cells. csA affects the apoptotic signaling pathway upstream of the caspase cascade, therefore, we investigated the difference in its effects compared with the broad spectrum caspase inhibitor Z-VAD-FMK. The ability of csA to inhibit cyt c release and increase clonogenic survival, whereas Z-VAD-FMK does not, suggests that apoptotic events that occur at the mitochondria may denote a `point of no return' in the cell death pathway. Furthermore, cells that are rescued from Cr-induced apoptosis by treatment with csA would survive despite the genotoxic effects of Cr. This would potentially promote the survival of genetically damaged cells which may contribute to Cr(VI)-induced carcinogenesis.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Cell culture
CHO AA8 cells were maintained in Eagle's basal medium containing 10% fetal bovine serum (Hyclone Laboratories, Logan, UT), 100 U/ml penicillin, 100 µg/ml streptomycin (Life Technologies, Gaithersburg, MD) and 2 mM glutamine (Life Technologies) in a 95% air/5% CO2 humidified atmosphere at 37°C.

Treatment of cells with chromium, Z-VAD-FMK and csA
Sodium chromate (Na2CrO4·4H2O) (J.T. Baker Chemical Co., Phillipsburg, NJ) was dissolved in double distilled H2O and sterilized through a 0.2 µm filter before use. Cells were treated with a final concentration of 0, 3, 6, 9 or 12 µM Na2CrO4 for 24 h in complete medium. After 24 h the medium was replaced and the cells were incubated for an additional 24 h period. csA (Sigma, St Louis. MO) was dissolved in dimethyl sulfoxide (DMSO). Cells were pre-treated for 30 min prior to the Na2CrO4 exposure and co-treated for the 24 h Na2CrO4 exposure period with a final concentration of 5 µM csA. Z-Val-Ala-Asp(OMe)-FMK (Z-VAD-FMK) (Enzyme Systems Products, Livermore, CA) was dissolved in DMSO and added to cells immediately prior to the 24 h Na2CrO4 exposure period at a final concentration of 50 µM. The final concentration of DMSO in the medium never exceeded 0.1%. Control samples were exposed to an equivalent concentration of DMSO.

Internucleosomal DNA fragmentation (IDF) analysis
IDF was analyzed after the 24 h post-Na2CrO4 exposure period. Cells were collected by trypsinization and combined with non-adherent cells from the culture medium. The cells were centrifuged at 600 x g for 5 min. Cell pellets were washed once in phosphate-buffered saline (PBS) and lysed in 500 µl of digestion buffer (100 mM Tris–HCl, pH 8.0, 200 mM NaCl, 10 mM EDTA, 4% SDS and 0.2 mg/ml proteinase K), followed by incubation at 55°C for 18 h. DNA was extracted with phenol/chloroform and then chloroform alone, incubated with 0.1 mg/ml RNase A for 30 min at 37°C and finally precipitated with 100% ethanol. Sample volumes equivalent to 7.5 µg were electrophoresed on a 1% agarose gel with 0.5 µg/ml ethidium bromide and visualized with an Eagle Eye II still video imaging system (Stratagene, La Jolla, CA).

Preparation of cytosolic fractions
Cr-treated cells were harvested by gentle cell scraping and combined with non-adherent cells from the culture medium. The cells were centrifuged at 300 x g for 5 min at 4°C and washed once with ice-cold PBS. Cell pellets were resuspended at 1.0–2.0x107 cells/ml in ice-cold isotonic cytosol buffer [210 mM mannitol, 70 mM sucrose, 5 mM HEPES, pH 7.5, 0.2 mM EGTA, 1 mM MgCl2, 5 mM glutamic acid, 5 mM malate, 0.1 mM phenylmethylsulfonyl fluoride, supplemented with protease inhibitors (5 µg/ml pepstatin A, 10 µg/ml leupeptin, 2 µg/ml aprotonin) (Sigma), and containing.004% digitonin to permeabilize the cells]. Cell suspensions were stirred for 10 min and then centrifuged at 12 000 x g for 15 min at 4°C to remove cell membrane debris, mitochondria and other organelles. The resulting supernatants were concentrated through Millipore Ultrafree-CL filters and stored at –20°C until used for PAGE and western blot analysis.

For measurement of the full cyt c content of a cellular sample, cytoplasmic extracts were isolated in a hypotonic solution. This cell fraction contains lysed mitochondria and pieces of plasma membrane and other organelles, but not whole cells or nuclei. Cell pellets were resuspended at 1.0–2.0x107 cells/ml in ice-cold hypotonic cytosol buffer [20 mM KCl, 20 mM HEPES, pH 7.5, 1.5 mM MgCl2, 1 mM EDTA, 1 mM DTT, 0.1 mM phenylmethylsulfonyl fluoride, 0.2 mM CaCl2, supplemented with protease inhibitors (5 µg/ml pepstatin A, 10 µg/ml leupeptin, 2 µg/ml aprotonin) (Sigma)]. Cell suspensions were vortexed for 10 min and then centrifuged at 5000 x g for 15 min at 4°C. The resulting supernatants were concentrated through Millipore Ultrafree-CL filters and stored at –20°C until used for analysis.

Protein electrophoresis and western blotting
Protein concentrations of the cytosolic fractions were determined with a DC Protein Assay Kit II (Bio-Rad, Hercules, CA). Samples containing an equal amount of concentrated proteins were separated on a 4–20% gradient SDS–polyacrylamide gel and transferred to a polyvinylidene difluoride membrane by electroblotting for 1 h at 30 V at 4°C. Non-specific membrane binding sites were blocked overnight at 4°C with blocking solution [TBS-T (2.7 mM KCl, 138 mM NaCl, 20 mM Tris base, pH 7.4, 0.1% Tween-20) containing 5% non-fat dry milk]. The membrane was incubated with primary mouse anti-cyt c monoclonal antibody (Pharmingen, San Diego, CA) diluted 1:2000 in blocking solution (0.5 µg/ml) for 1 h at room temperature. The membrane was washed thoroughly with TBS-T and then incubated for 1 h with a horseradish peroxidase-conjugated anti-mouse IgG secondary antibody (Amersham, Arlington Heights, IL) diluted 1:2000 in blocking solution. The secondary probe was detected by using the Renaissance enhanced chemiluminescence detection system (NEN, Boston, MA).

Integrated density analysis
Relative densities of protein or DNA bands were determined using an Eagle Eye II still video imaging system with Eaglesight software (Stratagene).

Clonogenicity analysis
CHO cells were seeded at 105 cells/60 mm2 dish and incubated for 24 h prior to csA or Z-VAD-FMK treatment and Na2CrO4 exposure. After the 24 h exposure, cells were collected by trypsinization, counted, and reseeded at 2x102 cells/60 mm2 dish in triplicate. The plates were incubated for 9–10 days, then rinsed with PBS and incubated with crystal violet stain (80% methanol, 2% formaldehyde, 2.5 g/l crystal violet) for 15–30 min at room temperature. The plates were thoroughly rinsed with dH2O and allowed to dry. Colonies were counted, the triplicates were averaged and long-term survival was determined as a percent of control.

Cr uptake analysis
Conical tubes (15 ml) containing 5x105 CHO cells/sample were pre-treated with either 5 µM csA or an equivalent volume of DMSO (final concentration 0.1%) for 30 min at 37°C. The cells were then exposed to appropriate concentrations of Na2CrO4 spiked with Na251CrO4 for 3 h at 37°C. The cells were centrifuged at 300 x g for 5 min at 4°C. Cell pellets were washed twice in PBS and lysed in 500 µl lysis buffer (100 mM Tris–HCl, pH 8.0, 200 mM NaCl, 10 mM EDTA, 4% SDS). An aliquot of 100 µl of each sample was transferred to a 5 ml scintillation vial and combined with 1 ml of Ecolite scintillation cocktail (ICN, Irvine, CA). Values for d.p.m./105 cells were determined in a Beckman LS6500 scintillation counter (Beckman Instruments, Fullerton, CA).


    Results
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 Materials and methods
 Results
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To determine if Cr(VI)-induced apoptosis involves mitochondrial disruption, we isolated the cytosolic fractions of apoptotic CHO cells and analyzed them for the presence of cyt c. We based our Cr(VI) concentrations on previously reported dose-dependant increases in CHO cell apoptosis and decreases in clonogenicity following Cr(VI) exposure (32). We isolated the cytosolic fractions using a method based on selective plasma membrane permeabilization in isotonic buffer containing low concentrations of digitonin (33,34). This method allows isolation of cytosol without contamination with mitochondria-associated cyt c. We carefully titrated the level of digitonin used in the isotonic cytosol buffer to determine the optimal conditions for plasma membrane permeability without mitochondrial membrane permeability in CHO cells. We observed that a 24 h exposure to different concentrations of Na2CrO4 caused a dose-dependent increase in cytosolic levels of cyt c (Figure 1AGo). Densitometric analysis of independent western blots showed that at the 3 µM dose only a minor increase in cytosolic cyt c was observed (Figure 1BGo). However, the cytosolic cyt c levels increased as the Na2CrO4 dose increased from 3 to 12 µM. The increases in cytosolic cyt c levels were statistically significant by Student's t-test at the 6, 9 and 12 µM Na2CrO4 doses (P = 0.024, 2.3x10–7 and 7.4x10–4, respectively). cyt c release was maximal at the highest dose tested (12 µM), which corresponds to nearly 100% apoptotic cell death in CHO cells (data not shown). Cytosolic samples with the full mitochondrial cyt c content isolated using a hypotonic buffer were also analyzed. The level of cyt c release at the 12 µM dose was ~75% of the total cyt c content of the hypotonic control sample (data not shown).



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Fig. 1. Cr(VI) causes dose-dependent release of cyt c. (A) Cytosolic fractions of CHO cells exposed to different concentrations of Na2CrO4 were collected and analyzed for cyt c by western blotting. The Na2CrO4 doses ranged from 0 to 12 µM as indicated. Purified bovine heart cyt c was used as a standard (St). (B) Densitometric analysis of cyt c western blots shows that the relative density of cytosolic cyt c increases significantly at 6, 9 and 12 µM Na2CrO4. Results are the means ± SE of three independent experiments. The * denotes the sample mean of two independent experiments.

 
Cytosolic fractions isolated from cells exposed to 9 µM Na2CrO4 and pre-treated for 30 min and co-treated for 24 h with 5 µM csA were analyzed for the presence of cyt c by western blotting. Treatment with csA markedly inhibited the cyt c release induced by Na2CrO4 (Figure 2AGo). In contrast, the general caspase inhibitor Z-VAD-FMK had no effect on the release of cyt c (Figure 2BGo). Densitometric analysis of independent western blots showed that only 24% of the cyt c release induced by 9 µM Na2CrO4 exposure was observed when cells were treated with csA as compared with cells treated with Z-VAD-FMK or solvent alone (Figure 2CGo).



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Fig. 2. Inhibition of Cr(VI)-induced mitochondrial cyt c release by csA but not by Z-VAD-FMK. (A) A 30 min pre-treatment and 24 h co-treatment with 5 µM csA was given to CHO cells exposed to 0 or 9 µM Na2CrO4. Cytosolic fractions of these cells were compared with cytosols of cells without csA treatment for the presence of cyt c by western blot analysis. An aliquot of 10 ng of purified bovine heart cyt c was used as a standard (St) in the first lane. (B) CHO cells exposed to 0 or 9 µM Na2CrO4 were either treated with 50 µM Z-VAD-FMK (Z-VAD) or a solvent control and the cytosols were compared for the presence of cyt c by western blot analysis. An aliquot of 10 ng of purified bovine heart cyt c was used as a standard (St) in the first lane. (C) Densitometric analysis of western blots shows the percentage of cytosolic cyt c in samples treated with 9 µM Na2CrO4 and co-treated with either csA or Z-VAD-FMK compared with the samples which received solvent only. Results are the means ± SE of three independent experiments.

 
Degradation of DNA into 180–200 bp oligonucleosomal fragments is a hallmark indicator of apoptosis (35). IDF is considered a late stage apoptotic event and is induced by active caspases (36). CHO cells exposed to 9 µM Na2CrO4 undergo extensive IDF (Figure 3AGo). Inhibition of the caspases with 50 µM Z-VAD-FMK was able to markedly reduce Cr-induced IDF, verifying the dependence for IDF on caspase activity. Treatment with 5 µM csA was also able to reduce IDF induced by 9 µM Na2CrO4, indicating that cyt c release may also be a necessary component of the IDF pathway. Densitometric analysis of three independent IDF gels showed that Z-VAD-FMK is able to reduce the amount of IDF to 36% of that caused by 9 µM Na2CrO4 alone, while csA can reduce Cr-induced IDF to a comparable 41% (Figure 3BGo).



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Fig. 3. Reduction of Cr-induced apoptosis by csA and Z-VAD-FMK. (A) Genomic DNA was isolated from CHO cells exposed to 0 or 9 µM Na2CrO4 for 24 h, and either treated with 5 µM csA, 50 µM Z-VAD-FMK or a solvent control. The DNA was isolated 24 h after exposure and analyzed for IDF. A 1 kb ladder was used as a DNA marker in lane 1. (B) Densitometric analysis of IDF shows the percentage of cytosolic cyt c from samples treated with 9 µM Na2CrO4 and co-treated with either csA or Z-VAD-FMK compared with the samples which received solvent only. Results are the means ± SE of three independent experiments.

 
Since csA was shown to block mitochondrial cyt c release and reduce apoptosis, we examined whether it could restore clonogenic survival to Cr-treated CHO cells. Clonogenicity is an indicator of a cell's long-term survival and ability to grow and form colonies in culture after exposure to a toxic agent. Clonogenicity reflects both apoptosis and long-term growth arrest caused by the toxic exposure. The clonogenicity of CHO cells exposed to different concentrations of Na2CrO4 alone and with pre- and co-treatments with csA or Z-VAD-FMK was determined (Figure 4Go). At the 3 and 6 µM doses csA was able to increase the number of colonies that formed. The increase in clonogenicity was statistically significant by Student's t-test at the 3 and 6 µM Na2CrO4 doses (P = 0.021 and 0.003, respectively). Exposure to 3 µM Na2CrO4 decreased clonogenic survival to 57% compared with the 0 µM control. Pre - and co-treatment with 5 µM csA at 3 µM Na2CrO4 increased the clonogenicity to nearly 100% of control. The 6 µM dose reduced clonogenicity to 9% of control, but csA increased this clonogenic potential to 30%. We observed the same trend at the highly toxic 9 µM dose, although it was not statistically significant. In contrast, Z-VAD-FMK had no effect on clonogenicity of Cr-exposed CHO cells.



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Fig. 4. Increased clonogenicity of Cr-exposed CHO cells by csA but not by Z-VAD-FMK. CHO cells exposed to increasing concentrations of Na2CrO4 for 24 h, and either treated with 5 µM csA, 50 µM Z-VAD-FMK or a solvent control were analyzed for cloning efficiency. The number of colonies for the indicated concentrations of Na2CrO4 are expressed as a percentage of the 0 µM control for each group. Results are the means ± SE of three independent experiments.

 
Because csA, but not Z-VAD-FMK, increased clonogenic survival after chromium treatment, we sought to determine whether csA affected uptake of Cr(VI) from the medium. Importantly, csA did not alter Cr uptake by CHO cells (Figure 5Go). CHO cells treated with csA and exposed to different levels of Na251CrO4 for 3 h had similar levels of intracellular 51Cr to the solvent controls.



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Fig. 5. CsA has no effect on Cr uptake. CHO cells exposed to increasing concentrations of Na2CrO4 spiked with Na251CrO4 for 3 h were either pre-treated for 30 min and co-treated with 5 µM csA or treated with a solvent control. The cells were then lysed and analyzed for intracellular 51Cr on a scintillation counter. Results are the means ± SE of three independent experiments.

 

    Discussion
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 Introduction
 Materials and methods
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 References
 
Dysregulation of the homeostasis between cell proliferation and cell death is involved in a vast array of human health consequences, including tumorigenesis. If apoptosis fails to occur normally when cells have been damaged or injured, then it may contribute to the progression of cancer. In studying the relationship between apoptosis and cancer, genotoxic carcinogens are particularly interesting because in order to induce neoplastic transformation, doses that induce a certain level of apoptosis are often required (16). In the case of Cr(VI), exposure to levels of particulate Cr compounds that are associated with lung cancer usually also cause respiratory toxicity that manifests as perforation of the nasal septum and/or respiratory tract ulcerations (16). These conditions are associated with high levels of cell death, thus a key question is how and why do certain cells exposed to a carcinogen escape death when others die? In tissue exposed to a Cr(VI) compound, individual cells may experience different local concentrations that are possibly genotoxic but not apoptotic. The populations of cells which either escape or are resistant to apoptosis may become the precursor pools out of which neoplastic variants will emerge. Therefore, it is important to determine the point in the apoptotic pathway when a cell is irrevocably committed to death. Our results here suggest that the `point of no return' for chromium-induced apoptosis occurs at the mitochondria and that cells that survive an apoptotic challenge may have a selective advantage in their ability to maintain mitochondrial stability.

Exposure to apoptotic doses of Cr(VI) causes mitochondrial instability, as indicated by the release of cyt c into the cytosol (Figure 1Go). It is believed that cyt c release is necessary for activation of the caspase cascade or, in some cases, at least for the amplification of caspase activity required for a normal apoptotic response (6,7,12). Here we show that treatment with the caspase inhibitor Z-VAD-FMK is able to inhibit DNA fragmentation associated with apoptosis (Figure 3Go) but does not affect long-term clonogenic survival (Figure 4Go). Despite inhibition of the morphological apoptotic characteristics caused by active caspases, the cells never regain replicative competency and eventually die. The point at which a cell is committed to death should, therefore, occur upstream of the caspase cascade.

csA functions at the mitochondria by promoting mitochondrial stability and preventing the release of caspase activators. It is believed to bind at the MPT pore, thereby increasing the probability of pore closure and preventing disruption of the mitochondrial inner membrane potential (13,14). Our results indicate that treatment with csA is able to inhibit apoptosis (Figure 3Go) and block mitochondrial cyt c release (Figure 2Go). Most importantly, csA treatment is able to restore long-term clonogenic survival to cells that have been exposed to apoptotic doses of Cr(VI) (Figure 4Go). These results suggest that maintaining mitochondrial stability may allow cells to escape apoptotic death and that the point along the apoptotic pathway in which cells are committed to death involves mitochondria. It is unclear whether the `point of no return' in cell death is disruption of the mitochondrial membrane potential or release of mitochondrial caspase activators. These two events may be intricately linked, however, some studies have shown that the release of cyt c can occur before or in the absence of disruption of the transmembrane potential (37,38). It is possible that different regulatory events which control mitochondrial membrane permeability are dependent on cell type. Nevertheless, it seems likely that uncoupling of the respiratory chain and subsequent release of cyt c from the mitochondria would prevent long-term cell survival. Release of cyt c from a mitochondrion should cause an irreversible disruption of oxidative phosphorylation, rendering the mitochondrion non-functional for energy metabolism and cell growth. The extent of cyt c release may vary for each cell depending on the individual molecular and cellular factors that govern apoptosis. Thus it is possible that when a critical amount of cyt c is released from mitochondria the cell is committed to death, either apoptotic or otherwise.

At lower doses (3 µM) of Cr(VI) only a small amount of cyt c release (reproducible but not statistically significant) could be observed at one particular time point (Figure 1Go). Interestingly, these low doses caused a significant decrease in clonogenic survival (57% of control) over a longer course of time. Cyt c release at lower doses of Cr(VI) may occur gradually followed by a protracted increase in apoptosis. Therefore, this response accounts for the significant decrease in clonogenic survival, but a marked release of cyt c at any one moment in time would not be expected. Nevertheless, treatment with csA was able to significantly increase the long-term survival of the cells, raising clonogenicity from 57 to nearly 100% (Figure 4Go). csA was not able to significantly increase the clonogenic survival of cells treated with the highest dose of Cr(VI) (9 µM). This is undoubtedly due to the presence of an insurmountable amount of damage to critical macromolecules making it impossible for the cells to regain replicative competence even if apoptosis is prevented.

Here we have shown that the anti-apoptotic and pro-survival effects of csA are not the result of altered cellular uptake of Cr(VI) (Figure 5Go). We have previously shown that treatment of cells with Cr(VI) causes structural DNA damage in the form of adducts, single-strand breaks, DNA crosslinks and chromosomal abnormalities (39). Some of this structural damage leads to functional damage in the form of base-specific DNA and RNA polymerase arrest and inhibition of DNA replication and transcription. The cellular response to this damage includes altered gene expression, G1/S and S phase cell cycle arrest, terminal growth arrest and apoptosis (21,23,2831). Survivors exhibit DNA deletion mutations and may progress to neoplastic transformation (39). We have observed that p53 levels are increased in Cr(VI)-treated cells (40) and, presumably, this increase would be part of the cellular response to genotoxicity, contributing to G1/S checkpoint arrest and apoptosis in the presence of excessive unrepairable damage. CHO cells express a mutant p53 which alters the G1/S checkpoint arrest arm of the p53 molecular response pathway (41,42), but they still undergo apoptosis after Cr(VI) treatment. Nevertheless, the p53 status of the cell line does not negate the observation that csA inhibited apoptosis and increased clonogenic survival, whereas Z-VAD-FMK inhibited apoptosis but had no effect on clonogenic survival. Experiments on p53 wild-type diploid human lung cells are in progress but are difficult and slow due to the nature of the primary cell cultures.

The ability of cells to undergo apoptosis in response to genotoxic insult may represent a protective mechanism against neoplastic transformation in the organism by eliminating cells that contain unrepaired genetic lesions. The cells that survive genotoxic damage by escaping apoptosis and gradually regaining replicative competence may harbor the deleterious effects of the apoptotic insult that induced death in the rest of the cell population. Thus, these cells may be predisposed to further progression towards a neoplastic phenotype. CsA, widely used as an immunosuppressive drug to treat autoimmune diseases and prevent rejection in organ transplantation, has many common side-effects, including an increased risk of cancer (43). The ability of csA to reduce apoptosis may be a consideration as a mechanism for the increased risk of cancer in patients who take csA long term. In this study, cells acquired resistance to apoptosis by interference with the mitochondrial instability conveyed by the pharmacological treatment. However, this raises the interesting question of whether the cells that survive apoptotic doses of a chemical carcinogen such as Cr(VI), in the absence of any pharmacological intervention, have a stochastic resistance to mitochondrial instability. By way of this resistance, such a sub-population of cells may be selected for by the apoptotic inducer, thereby increasing the population of cells resistant to apoptosis. If early tumor growth requires a net accumulation of cells due to alterations in the growth/death ratio, then selection of cells with resistance to apoptosis may facilitate the early steps of carcinogenesis.


    Notes
 
* This work was conducted in partial fulfilment of the requirements for the PhD degree in Genetics, Columbian Graduate School of Arts and Sciences, The George Washington University. Back

4 To whom correspondence should be addressed at: Department of Pharmacology, The George Washington University Medical Center, 2300 Eye Street NW, Washington, DC 20037, USA Email: phmsrp{at}gwumc.edu Back


    Acknowledgments
 
The authors thank Bronwyn Owens and Kristy Markos for their expert technical assistance and Dr. Anne Murphy for her valuable assistance in method development.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received February 4, 2000; revised July 6, 2000; accepted July 11, 2000.