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Carcinogenesis Advance Access originally published online on February 14, 2007
Carcinogenesis 2007 28(7):1613-1620; doi:10.1093/carcin/bgm031
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Cellular vitamin C increases chromate toxicity via a death program requiring mismatch repair but not p53

Mindy Reynolds and Anatoly Zhitkovich*

Department of Pathology and Laboratory Medicine, Brown University, 70 Ship Street, Room 507, Providence, RI 02912, USA

* To whom correspondence should be addressed. Tel: +401 863 2912; Fax: +401 863 9008; Email: anatoly_zhitkovich{at}brown.edu


   Abstract
Top
 Abstract
Introduction
 Materials and methods
 Results
 Discussion
 References
 
Ascorbate (Asc) plays a key role in reductive activation of carcinogenic chromium(VI) in vivo. In addition to much higher rates (t1/2 = 1 min for 1 mM Asc), its reactions at physiological conditions differ from other reducers by low yields of Cr(V) intermediates. Human cells in culture are severely Asc deficient, which results in distorted metabolism and potentially abnormal responses to Cr(VI). We found that restoration of physiological Asc levels in human lung cells (primary IMR90 fibroblasts and epithelial H460 cells) increased clonogenic lethality and apoptosis by Cr(VI). Enhanced cytotoxicity in mass cultures was more evident after normalization for lower Cr uptake caused by leakage of Asc into media. Asc did not change uptake-adjusted yields of Cr–DNA adducts and had no effect on cytotoxicity when delivered shortly after Cr(VI) exposure. Protein and Ser-15 phosphorylation levels of p53 did not show any association with the presence of Asc and there were no increases in p53-driven reporter activity in Cr-treated cells. Stable silencing of p53 expression by short hairpin RNA (shRNA) had no effect on toxicity of Cr(VI) in both –Asc and +Asc IMR90 and H460 cells. In contrast, shRNA-mediated depletion of essential components of MutS or MutL mismatch repair complexes greatly improved survival of all Cr-treated cells and eliminated Asc-potentiated effects on cell death. Thus, mismatch repair-mediated enhancement of Cr(VI) cytotoxicity by Asc should promote the selection of MSI+/wt-p53 phenotype found among chromate-induced human lung cancers. Our findings also indicate that Asc plays a dual role in Cr(VI) toxicity: protective outside and potentiating inside the cell.

Abbreviations: Asc, ascorbate; DHA, dehydroascorbic acid; PBS, phosphate-buffered saline; SDS, sodium dodecyl sulfate; shRNA, short hairpin RNA


   Introduction
Top
 Abstract
 Introduction
Materials and methods
 Results
 Discussion
 References
 
The carcinogenicity of hexavalent Cr has been characterized extensively through epidemiological, animal and in vitro studies (1–3). Human exposure to Cr(VI) compounds occurs in a large number of industrial workers (4) and has been linked to a significantly elevated risk of developing lung cancer (1–3), as well as other adverse health effects (4,5). There are also recurrent concerns about the potential adverse effects of exposure to Cr(VI) in residential areas from contaminated water and soil sources; however, the extent of exposure and the presence of toxicologically relevant biological damage in individuals residing in the vicinity of Cr(VI)-contaminated sites has been a subject of intense debate (4,6,7). Molecular mechanisms underlying cytotoxic, mutagenic and carcinogenic effects of Cr(VI) are complex, in particular due to its complicated intracellular chemistry and the involvement of multiple intracellular targets and pathways (8,9).

Under physiological conditions, Cr(VI) is unreactive with DNA and requires reductive activation to Cr(III) to induce biological damage. Cr(III) ions form a series of stable coordination complexes with DNA and other intracellular molecules (9). Reduction of Cr(VI) to Cr(III) can be accomplished through non-enzymatic reactions with cysteine and glutathione; however, in the target tissues of chromate toxicity, such as lung, ascorbate (Asc) is the primary reducer of Cr(VI) (9–11). Asc has the highest rate of Cr(VI) reduction among all biological reducing agents (12) and Asc-driven reactions generate high levels of Cr–DNA binding (13,14) and produce mutation-inducing damage (12,15). Mutagenic Asc–Cr–DNA cross-links that are uniquely formed during Cr(VI) reduction with Asc (13) are genotoxic as detected by the highly reduced yield of replicated progeny following propagation of adducted plasmids in human fibroblasts (12). In addition, cellular Asc has been found to strongly enhance the ability of Cr(VI) to cause DNA breakage and mutations (16). Therefore, it is possible that cellular Asc may also act as a potentiator of Cr(VI)-induced death in human cells. Toxicity of Cr(VI) has been linked to its carcinogenicity through the process of selection of resistant but genetically unstable populations (17,18). Given the dominant role of Asc in the reductive metabolism of Cr(VI) to Cr(III), it is important to understand the significance of this reducer in the induction of cell death programs associated with Cr(VI) carcinogenesis. Previous studies have investigated Cr(VI)-induced cytotoxicity under standard tissue culture conditions (19,20), which are severely deficient in Asc and therefore would not address the critical issue of Asc-dependent metabolism in mechanisms of cell death by Cr(VI).

In this work, we examined apoptotic responses and clonogenic survival of human lung cells after exposure to Cr(VI) in the presence and absence of physiological levels of Asc. We found that cellular Asc increased lethality of Cr(VI) due to a stronger activation of apoptosis. The p53 transcriptional factor was dispensable for the induction of cell death, but depletion of MLH1 and MSH2 mismatch repair proteins abrogated the potentiating effects of Asc on Cr(VI) cytotoxicity. These findings indicate that cellular vitamin C enhances the ability of Cr(VI) to select for mismatch repair-deficient cells in a p53-independent manner and are in agreement with a low incidence of p53 mutations in chromate-associated human lung cancers (21). The majority of Cr-related cancers exhibit microsatellite instability caused by the loss of expression of MLH1 mismatch repair protein (22,23).


   Materials and methods
Top
 Abstract
 Introduction
 Materials and methods
Results
 Discussion
 References
 
Cells and exposures
Cells were routinely maintained in a humidified atmosphere containing 95% air and 5% CO2. Human lung epithelial H460 cells and primary human lung IMR90 fibroblasts were obtained from American Type Tissue Collection. IMR90 cells were used at passages 6–9. H460 cells were grown in RPMI medium with 10% fetal bovine serum and penicillin/streptomycin. IMR90 fibroblasts were cultured in Dulbecco’s modified Eagle’s medium supplemented with 10% serum and penicillin/streptomycin. Next day after seeding, cells were loaded with Asc by pre-incubation for 90 min with dehydroascorbic acid (DHA) in Krebs-N-2-hydroxyethyl piperazine-N'-2-ethanesulfonic acid buffer [30 mM HEPES (pH 7.5), 130 mM NaCl, 4 mM KH2PO4, 1 mM MgSO4, 1 mM CaCl2, and 0.5 mM glucose]. To create 1 mM intracellular concentration of Asc, H460 cells were pre-incubated with 2 mM DHA and IMR90 with 5 mM DHA. Exposures to K2CrO4 [Cr(VI)] were for 3 h in serum-free medium.

Determination of cellular Asc
Asc concentrations were measured by high performance liquid chromatography, based on the detection of a specific conjugate with 1,2-diamino-4,5-dimethoxybenzene dihydrochloride (13). Briefly, cells were collected by trypsinization, washed three times with phosphate-buffered saline (PBS) (1100g for 5 min at 4°C) and re-suspended in a solution containing 50 mM methanesulfonic acid and 5 mM diethylenetriaminepentaacetic acid. Samples were subjected to two cycles of freezing (–80°C) and thawing (37°C), and Asc-containing supernatants were collected after centrifugation at 12 000g for 20 min at 4°C. Pellets were re-suspended in 1% sodium dodecyl sulfate (SDS) and 50 mM NaOH for determination of protein concentrations. Derivatization reactions containing 10 µl of Asc solution and 90 µl of a dye solution [0.2 U/µl ascorbate oxidase, 50 mM Na acetate (pH 6.2) and 0.5 mM 1,2-diamino-4,5-dimethoxybenzene dihydrochloride (Molecular Probes)] were incubated for 4 h at room temperature in the dark. HPLC separation of fluorescent derivatives of Asc was performed using isocratic elution with 75% 50 mM phosphoric acid (pH 2.0) and 25% acetonitrile at a flow rate of 1 ml/min. The detection limit for Asc in cells was 0.1 µM.

Clonogenic survival
Cells were seeded onto 60 mm dishes in triplicates and allowed to attach overnight. Cells were pre-loaded with Asc and treated with Cr(VI), as described above. Approximately 7–10 days after exposure, cells were stained with Giemsa solution (Sigma-Aldrich, Saint Louis, MI, USA) and the number of colonies was scored manually.

Cytotoxicity
Short-term viability was measured by a LIVE/DEAD assay (Molecular Probes). Cells were seeded in triplicate in 12-well plates, followed by exposure to Asc and Cr(VI). At the indicated times post-exposure, cells were washed twice with room temperature PBS and incubated with 4 µM calcein for 15 min at 37°C in a humidified 95% air–5% CO2 atmosphere. Fluorescence was measured on a Spectramax M2 (Molecular Devices, Sunnyvale, CA, USA) at excitation 485 nm and emission 525 nm.

Western blotting
Attached and floating cells were collected by scraping, washed twice with cold PBS and then re-suspended in lysis buffer [50 mM Tris (pH 8.0), 250 mM NaCl, 1% NP-40, 0.1% SDS, 5 mM ethylenediaminetetraacetic acid, 2 mM Na3VO4, 10 mM Na2P2O7, 10 mM NaF, 10 µg/ml aprotinin, 10 µg/ml leupeptin, 0.5 µg/ml pepstatin and 1 mM phenylmethanesulfonyl fluoride. After incubation on ice for 10 min, cellular debris was removed by centrifugation at 10 000g for 10 min at 4°C. Proteins were separated by SDS–polyacrylamide gel electrophoresis and electrotransferred onto ImmunoBlot polyvinylidene difluoride membrane (Bio-Rad, Hercules, CA, USA). Primary antibodies used were anti-MLH1, anti-MSH2 (PharMingen, San Jose, CA, USA), anti-poly(ADP-ribose) polymerase (PARP), anti-phospho-p53 (Ser-15), anti-caspase 3, anti-caspase 7, (Cell Signaling, Beverly, MA, USA), anti-p53 (DO-1) (Santa Cruz Biotechnology, Santa Cruz, CA, USA) and anti-{gamma}-tubulin clone GTU-88 (Sigma). Protein bands were visualized using horseradish peroxidase-conjugated secondary antibodies (Upstate/Millipore, Billerica, MA, USA) and enhanced chemiluminescence kit (Amersham, Piscataway, NJ, USA).

Stable siRNA knock-down of MLH1, MSH2 and p53
Stable depletion of protein levels was achieved by expression of short hairpin RNA (shRNA) from the pSUPER-RETRO retroviral vector (Oligoengine, Seattle, WA, USA). The pSUPER-RETRO plasmid was linearized with HindIII and BglII to permit the insertion of the annealed oligonucleotides. The targeting vectors were constructed from the sense and anti-sense oligos directed at previously validated mRNA regions (16,24,25). Packaging conditions, infections and selection of puromycin-resistant cells were performed as described previously (26). IMR90 cells were infected once and selected in the presence of 1.5 µg/ml of puromycin at 7 days post-infection. The efficiency of knock-downs was determined by western blotting.

p53 reporter assay
After overnight attachment, cells were transfected with p53-Luc (Stratagene, La Jolla, CA, USA) and pL-TK (Promega, Madison, WI, USA) plasmids. The following day, cells were exposed to Asc and Cr(VI) and returned to complete medium. Cellular lysates were prepared 24 h post-Cr exposure. As a positive control, cells were treated with 1 µg/ml doxorubicin for 24 h in complete medium. Luciferase activity was measured on a Tecan Spectrafluor Plus using a Dual-Luciferase Reporter Assay kit (Promega). The p53-driven firefly luciferase activity was normalized using the activity of constitutively expressed Renilla luciferase.

Chromium analyses
Total Cr was measured by graphite furnace atomic absorption spectroscopy, using a model 41002L GF-AAS instrument from PerkinElmer (Waltham, MA, USA) (13). The detection limit was 0.4 pmol of Cr or one Cr adduct per 10 000 nucleotides. For the determination of intracellular Cr concentrations, cells were collected by trypsinization, washed twice with cold PBS and re-suspended in 50 µl of cold water followed by the addition of 50 µl of 10% nitric acid. Samples were frozen at –80°C and then heated to 50°C for 60 min, followed by incubation on ice for 30 min and centrifugation at 10 000g for 10 min at 4°C. Supernatants were diluted with water 2.5 times and analyzed for Cr content. The average recovery of Cr by this procedure was previously found to be 85% (27). For protein determinations, pellets were washed twice with cold 5% HNO3 and then dissolved in 200 µl of 0.5 M NaOH by heating at 37°C for 30 min.

For determination of Cr–DNA adducts, cells were collected with trypsin and washed twice with cold PBS. Pellets were re-suspended in 200 µl of 25 mM HEPES (pH 8.0), 0.5% Triton X-100, 100 µg/ml RNAse A and incubated at 37°C for 30 min. This was followed by the addition of 1% SDS, 200 mM NaCl, sheering of DNA using a 25 gauge needle, addition of 0.2 mg/ml of proteinase K and incubation at 37°C for 1 h. DNA was isolated by phenol–chloroform, followed by overnight precipitation of DNA at 4°C with ethanol. DNA pellets were collected by centrifugation and washed twice with 70% ethanol. Air-dried DNA samples were dissolved overnight in 100 µl water and used for Cr determinations.


   Results
Top
 Abstract
 Introduction
 Materials and methods
 Results
Discussion
 References
 
Asc loading and uptake of Cr(VI)
Human lung tissue on average has 1.3 mM Asc (28), whereas cultured human cells either lack detectable Asc or contain it only at low micromolar concentrations (13,29). Loading of cells directly with Asc is very inefficient and can lead to cellular toxicity (13,30). Pre-treatment of cells with DHA, which readily enters cells and is then reduced to Asc, can rapidly restore physiological concentrations of Asc (13,31). We found that after one day in culture, H460 human lung epithelial cells contained barely detectable levels of Asc (Figure 1A). However, incubation of H460 cells with 2 mM DHA led to the accumulation of 1 mM intracellular Asc (Figure 1A and B). The restoration of intracellular Asc was transient, as levels of Asc decreased by ~90% at 6 h post-DHA loading (Figure 1C). Therefore, to determine the effects of Asc on the cytotoxic potential of Cr(VI), incubations with DHA and Cr(VI) were done sequentially. The presence of Asc in H460 cells proportionally decreased accumulation of Cr and formation of Cr–DNA adducts by 2.3- and 2.1-fold, respectively (Figure 1D and E). Thus, despite the differences in uptake, the normalized yield of Cr–DNA adducts in –Asc and +Asc cells was essentially the same. We attribute diminished accumulation of Cr to leakage of cellular Asc (Figure 1C), resulting in the reduction of Cr(VI) outside the cell.


Figure 1
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  		Fig. 1. Asc loading and Cr(VI) uptake in H460 cells. (A) Representative HPLC profiles for Asc determination in protein-normalized extracts from H460 cells. (B) Dose-dependent accumulation of Asc in H460 cells. Data are means ± SDs for three independent samples. (C) Time-dependent loss of intracellular Asc from H460 cells following removal of DHA. Data are means ± SDs for two independent samples. Cells loaded with 1 mM Asc had (D) lower uptake of Cr(VI) and (E) decreased formation of Cr–DNA adducts. Results are means ± SDs for two to five independent samples. When not visible, error bars were smaller than the data symbol.

 
Increased toxicity of Cr(VI) in human H460 cells containing Asc
To determine the effects of Asc on cytotoxicity of Cr(VI), we first examined clonogenic survival of cells as a function of Cr(VI) dose. We found that H460 cells pre-loaded with 1 mM Asc showed decreased clonogenicity compared with –Asc control cells (Figure 2A). Asc increased the clonogenic lethality of Cr(VI) in a concentration-dependent manner (Figure 2B). We found that even the presence of low 0.3 mM Asc led to decreased cell survival, and Cr was even more toxic in cells containing 0.6 and 1 mM Asc (Figure 2B). As an additional indicator of toxicity, we measured cellular viability using a LIVE/DEAD assay that monitors intracellular esterase activity. Because +Asc and –Asc H460 cells accumulate different amounts of Cr (Figure 1D), we analyzed cellular viability as a function of cellular Cr content (Figure 2C). We found that at 48 h post-exposure, Cr-treated +Asc cells exhibited approximately two times lower viability compared with their –Asc controls. Decreased viability of +Asc cells was also observed at 24, 72 and 96 h post-exposure (data not shown).


Figure 2
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  		Fig. 2. Cellular Asc increases clonogenic lethality and apoptosis in Cr-treated H460 cells. (A) Clonogenic survival of Cr(VI) in control and 1 mM Asc-pre-loaded H460 cells. Data are means ± SDs for three independent experiments (each with three dishes per dose). (B) Clonogenic survival of H460 cells loaded with different Asc concentrations prior to Cr exposure. Cells contained 0, 0.3, 0.6, or 1 mM Asc. Shown are means ± SDs for three dishes. (C) Viability of control and 1 mM Asc-pre-loaded H460 cells after Cr exposure. Cells were assayed for viability at 48 h post-Cr exposure using a LIVE/DEAD assay. Results are means ± SDs for six independent measurements. When not visible, error bars were smaller than the data symbol. (D) Western blots for intact and cleaved PARP in cellular extracts collected at different times following exposure to uptake-normalized doses of Cr(VI) (10 µM for –Asc and 23 µM for +Asc cells, as determined by uptake measurements in Figure 1D). (E) Western blots for caspases 3 and 7 in cell lysates collected 24 h after Cr exposure. *, cells were treated with uptake-normalized doses of Cr(VI) (2.3 times higher than for –Asc samples).

 
We next sought to determine whether the observed differences in cell survival were related to the alterations in apoptotic responses. Because of lower accumulation of Cr(VI) in +Asc H460 cells (Figure 1D), we treated these cells with uptake-normalized doses of Cr(VI). Time course studies showed that Asc promoted enhanced PARP cleavage as early as 9 h post-exposure in Cr-exposed cells, whereas cleavage of PARP in –Asc cells was not detected until 24 h post-exposure (Figure 2D). Even at the latest time point, apoptotic PARP cleavage in –Asc cells was still significantly lower than in +Asc cells. To further investigate the apoptotic responses, we analyzed the activation of executioner caspases 3 and 7. At the highest dose of Cr(VI), both pro-caspases underwent activating cleavage in +Asc cells, whereas the presence of activated caspases 3 and 7 was almost undetectable in –Asc cells (Figure 2E). Thus, the presence of Asc during the reduction of Cr(VI) enhances Cr-induced apoptosis, as measured by three biochemical markers.

Asc increases toxicity of Cr(VI) in primary human cells
To determine whether Asc can also influence toxic activity of Cr(VI) in primary cells, we examined viability of IMR90 lung fibroblasts. IMR90 cells pre-loaded with 1 mM Asc exhibited a much lower viability compared with control cells containing barely detectable Asc concentrations (Figure 3A). Based on the slopes of dose–response curves, the presence of Asc increased toxicity of Cr(VI) by 3.6-fold. Asc-mediated enhancement of Cr toxicity was detected as early as 24 h post-exposure (data not shown). Increased toxicity in +Asc cells did not result from the elevated uptake of Cr(VI), which in fact was lower in +Asc IMR90 cells by 2.2-fold (Figure 3B). Analysis of toxicity as a function of cellular doses of Cr (Figure 3C) showed that Asc-dependent potentiation of cell death was very strong, producing the enhancement factor of 6.8.


Figure 3
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  		Fig. 3. Asc increases toxicity of Cr(VI) in primary lung fibroblasts. DHA-pretreated cells had 1 mM Asc, whereas mock-treated cells typically contained 2–5 µM Asc. When not visible, error bars were smaller than the data symbol. (A) IMR90 cells pre-loaded with Asc had lower viability after Cr(VI) treatments. Viability was measured by a LIVE/DEAD assay at 48 h post-Cr exposure. (B) IMR90 cells pre-loaded with Asc had lower uptake of Cr(VI). Data are means ± SDs for six independent samples. (C) Viability of control and Asc-loaded IMR90 cells as a function of cellular Cr concentrations. The graph was generated using data from panels (A and B).

 
Post-Cr loading with Asc had no effect on cell survival
Increased sensitivity of Asc-containing cells to Cr(VI)-induced lethality could potentially stem from either stronger toxic responses to the same type of Cr–DNA damage (general sensitization effect) or the differences in Cr(VI) metabolism and resulting DNA damage in –Asc and +Asc cells. To differentiate between these possibilities, we loaded H460 and IMR90 cells with Asc following exposure to Cr(VI) and examined cell viability and clonogenic survival. We found that delivery of Asc at 1 h post-Cr exposure had no effect on clonogenic survival of H460 cells or viability of H460 and primary IMR90 cells at 48 h post-Cr exposure (Figure 4A, B and C). Thus, cellular Asc does not alter cytotoxic responses to Cr–DNA damage that was already formed in cells. These results indicate that in order to cause increased cell death, Asc must be present in cells during Cr(VI) reduction.


Figure 4
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  		Fig. 4. Loading of Asc after Cr(VI) exposure has no effect on toxicity. Loading with Asc was performed 1 h after removal of Cr-containing media. When not visible, error bars were smaller than the data symbol. (A) Clonogenic survival of H460 cells with and without post-Cr loading with 1 mM Asc. Data are means ± SDs for two independent experiments, each with three dishes per dose. Loading of cells with Asc after Cr exposure had no effect on viability of (B) H460 cells or (C) IMR90 cells. Cytotoxicity was evaluated by a LIVE/DEAD assay at 48 h post-Cr exposure. Data are means ± SDs for three to nine measurements.

 
Enhanced cytotoxicity by Asc is independent of p53
Induction of cell death in response to many types of DNA damage is mediated by p53-dependent expression of pro-apoptotic genes (32). Therefore, we decided to examine whether Asc-enhanced lethality also required activation of the p53 transcription factor. We assessed two important indicators of p53 activation: levels of p53 protein and the extent of its phosphorylation at Ser-15. These parameters were measured 48 h post-exposure to Cr(VI), which was the time corresponding to the maximal induction of apoptosis. We did observe modest increases in p53 protein levels and phosphorylation of Ser-15 in H460 cells, which appeared to be unrelated to the presence of Asc (Figure 5A). Cr treatments of either +Asc or –Asc IMR90 cells did not lead to significant alterations in protein levels of p53 (Figure 5B). Ser-15 phosphorylation also remained essentially unchanged in these primary cells. Next, we tested whether Cr(VI) stimulated p53 transactivation ability. The levels of p53-dependent transcription were assessed using a p53-Luc reporter plasmid containing multiple copies of the p53 response element linked to the firefly luciferase gene. As a positive control, we used doxorubicin-treated cells, which showed an ~25-fold increase in p53-dependent transcription of the luciferase gene over untreated cells. We found that regardless of their Asc status, increasing doses of Cr did not induce p53 transcriptional activity over the untreated control (Figure 5C).


Figure 5
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  		Fig. 5. Effect of Asc on Cr(VI) toxicity is independent of p53. Western blots of p53 protein and phosphorylated Ser-15 of p53 in (A) H460 cells and (B) IMR90 cells. H460 and IMR90 cells were pre-loaded with 1 mM Asc and protein extracts were prepared 48 h post-Cr exposure. *, H460 cells were treated with uptake-normalized doses of Cr(VI) (2.3 times higher than for –Asc samples). IMR90 cells were treated with the same (not normalized) doses of Cr(VI). (C) Luciferase reporter assay for p53 transcriptional activity. H460 cells were pre-loaded with 1 mM Asc, treated with 0–10 µM Cr(VI), and luciferase activity was assayed 24 h later. As a positive control, cells were treated with 1 µg/ml doxorubicin for 24 h. Results are means ± SDs for three independent experiments, each with four wells per dose. (D) Knock-down of p53 protein by stable expression of shRNA in H460 cells. A non-specific (Luc) shRNA was used as a control. Clonogenic lethality of Cr(VI) in shRNA-expressing H460 cells (E) without and (F) with 1 mM Asc pre-loading. (G) Effect of p53 depletion on clonogenic survival of H460 cells treated with hydrogen peroxide. Cells were treated H2O2 for 1 h in serum-free medium. Results are means ± SDs from two independent clonogenic assays that had three dishes per dose. (H) Western blotting for p53 protein in parental and shRNA-expressing IMR90 cells. Toxicity of Cr(VI) in IMR90 cells expressing Luc and p53 siRNA cultured (I) without Asc or (J) pre-loaded with 1 mM Asc. Cellular viability was measured 48 h post-Cr exposure using a LIVE/DEAD assay. Shown are means ± SDs for three independent samples. When not visible, error bars were smaller than the symbol.

 
To test a potential role of p53 in Asc-mediated Cr cytotoxicity more directly, we depleted p53 protein levels in H460 and IMR90 cells by expressing shRNA from a chromosomally integrated retroviral vector. Western blots revealed knock-down of p53 to almost undetectable levels in cells expressing targeting shRNA, whereas the amount of p53 protein in cells infected with a non-specific, luciferase-targeting (Luc) shRNA was similar to that in parental cells (Figure 5D and H). To validate the functionality of p53 knock-downs, we measured clonogenic survival of H460 cells exposed to H2O2. As expected, H460 cells expressing p53-targeting small interfering RNA (siRNA) exhibited decreased clonogenic lethality compared with control cells infected with the vector encoding a non-specific shRNA (Figure 5G). At the highest dose of H2O2, p53 deficiency increased the number of surviving cells by >6-fold. In contrast to oxidative damage, depletion of p53 showed no significant effect on clonogenic survival or short-term viability of Cr-treated Asc-deficient (Figure 5E and I) and Asc-supplemented cells (Figure 5F and J). Therefore, enhanced cytotoxicity of Cr(VI) in the presence of cellular Asc appears to be completely independent of p53 protein.

Loss of mismatch repair blocks potentiating effects of Asc
Mismatch repair is an important trigger of toxic responses to Cr–DNA damage in cells cultured under standard conditions (17). This prompted us to investigate whether the effects of Asc on cytotoxicity of Cr(VI) also involve mismatch repair. To generate mismatch repair-deficient cells, we created stable knock-downs of MSH2 and MLH1 using retroviral vectors expressing shRNA. MSH2 and MLH1 proteins are essential components of MutS and MutL mismatch repair complexes, respectively (33). Western blot analyses showed that our shRNA constructs were highly effective and decreased MLH1 and MSH2 proteins to very low levels in IMR90 and H460 cells (Figure 6A and D). We found that depletion of MLH1 or MSH2 proteins in IMR90 cells led to almost complete loss of potentiating effects of Asc on Cr toxicity (compare Figure 6B versus C). Knock-down of MLH1 in H460 cells also resulted in essentially indistinguishable clonogenic lethality of Cr(VI) in the presence or absence of cellular Asc (Figure 6E versus F). To further confirm our findings, we compared clonogenic survival of parental H460 cells and their derivatives expressing MSH2 targeting and a different non-specific shRNA (Figure 6G and H). Silencing of MSH2 again resulted in the increased viability of both –Asc and +Asc cells and the disappearance of Asc dependence. Control experiments showed that our manipulations of MLH1 and MSH2 levels did not change Cr(VI) accumulation by H460 in the presence or absence of Asc (Figure 6I).


Figure 6
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  		Fig. 6. Down-regulation of mismatch repair blocks potentiating effects of Asc on Cr(VI) toxicity. (A) Western blots demonstrating efficiency of siRNA-mediated depletion of MLH1 and MSH2 proteins in IMR90 cells. Viability of IMR90 cells expressing (B) non-specific (Luc) siRNA or (C) MLH1 and MSH2-targeting siRNA. Cells were either cultured under standard conditions (–Asc) or pre-loaded with 1 mM Asc before Cr exposure (+Asc). Cytotoxicity was measured 72 h after Cr exposure. Data are means ± SDs for six independent samples. (D) Western blots for MLH1 and MSH2 in H460 cells expressing targeting and non-specific (Luc) siRNA. MSH2 knock-down is shown for two independently infected populations. Clonogenic survival of H460 cells expressing (E) Luc or (F) MLH1 targeting siRNA with and without pre-loading with 1 mM Asc. Results are means ± SDs from two independent clonogenic experiments, each with three dishes per dose. When not visible, error bars were smaller than the symbol. (G) Clonogenic survival of parental (no siRNA) and non-specific green fluorescence protein (GFP) siRNA expressing H460 cells without (–Asc) or with (+Asc) 1 mM Asc pre-loading. (H) Silencing of MSH2 by shRNA abrogated survival differences between Asc-deficient (–Asc) and 1 mM Asc-pre-loaded (+Asc) H460 cells. (I) Depletion of MLH1 and MSH2 proteins does not change uptake of Cr(VI) by H460 cells. Open symbols, control (–Asc) cells, closed symbols, Asc-loaded (+Asc) cells; closed triangles and open triangles, non-specific (GFP) siRNA; open squares and closed squares, MLH1 siRNA; closed circles and open circles, MSH2 siRNA. Results are means from three independent samples. Error bars were 5–15% of the means and not shown for clarity.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
References
 
Asc and Cr(VI) cytotoxicity
All human cells and non-hepatic cells from other mammalian species lack the ability to make their own vitamin C and therefore must obtain it from exogenous sources. Standard tissue culture conditions provide only a limited supply of Asc (13,34) because the majority of synthetic media formulations do not include vitamin C and the typical amounts of added serum would provide only ~10% of the normal Asc concentrations. The actual levels of total vitamin C in tissue culture are even lower because of the losses of Asc/DHA during serum preparations and storage, as well as due to irreversible oxidation of Asc in the oxygenated media. Thus, in order to understand the role of Asc in Cr(VI) genotoxicity, cultured cells must be supplemented with this vitamin. Incubations with the reduced form of vitamin C do not lead to the restoration of physiological levels of intracellular Asc due to a very inefficient transport of Asc into the majority of human cells (13,30). Additionally, the presence of extracellular Asc in high concentrations has been associated with toxic effects (30), resulting from the production of H2O2. The most effective approach to cellular delivery of Asc was found to be incubations with DHA, the oxidized form of vitamin C (13,16), which is rapidly taken up by all cells via major glucose transporters (35). This delivery method also eliminates undesirable redox reactions outside the cells. The inability of previous studies to observe significant differences in toxicity of Cr(VI) in human cells pre-incubated with 1 mM reduced vitamin C was most probably the result of too low cellular concentrations of Asc (36,37). Interestingly, one of these studies (37) has determined that the presence of ~80 µM Asc in 1 mM vitamin C-treated A549 cells has been sufficient to alter the formation of intermediate Cr oxidations. Using a more efficient DHA-based loading procedure, we found that cellular Asc acts as an enhancer of cell death in Cr(VI)-treated human lung cells, such as primary IMR90 fibroblasts and H460 epithelial cells. We have also observed increased lethality of Cr(VI) in other human cell lines pre-loaded with physiological levels of Asc (Reynolds M. and Zhitkovich A., unpublished data). One difficulty in the evaluation of the effects of cellular Asc on Cr(VI) toxicity is the leakage of Asc into the medium, which results in the extracellular reduction of Cr(VI) leading to its diminished uptake. This phenomenon is probably less pronounced in the clonogenic experiments because leakage of Asc from 200 to 500 cells would not create significant extracellular concentrations of this reducer in the entire dish (although reduction in the close proximity of individual cells would still be possible due to locally higher Asc levels). Studies on the biochemical mechanisms of cell death require large numbers of cells per dish and the comparison of toxic responses between –Asc and +Asc cells necessitated corrections for differences in Cr(VI) uptake. Primary IMR90 cells with restored Asc levels exhibited lower survival in response to Cr(VI) even without uptake corrections, but analysis of apoptosis in H460 cells without adjustments for lower cellular doses of Cr would have missed the potentiating ability of intracellular Asc.

Cr(VI) metabolism by Asc is very different from that by other biological reducers. At physiological concentrations and reactant ratios, Asc reduces Cr(VI) >10 times faster than do thiols (12) and Asc-driven two-electron reduction reactions lacked Cr(V)-specific electron spin resonance signals (38,39). Our highest doses led to the accumulation of <50 or 150 µM cellular Cr concentrations in H460 or IMR90 cells, respectively (Figures 1D and 3B). Since at the start of exposures cells contained 1 mM Asc and Cr(VI) was entering gradually, it is likely that the Asc:Cr(VI) ratio in cells was >10:1 at all time points. Considering that in vitro reactions with the same or even significantly lower ratio of reactants had no detectable Cr(V) (38,39) and taking into account our findings that toxicity of Cr(VI) in +Asc cells was higher, it appears that Cr(V)-generating reduction pathways are less toxic.

Increased death of cells that metabolized Cr(VI) in the presence of Asc was probably caused by the altered spectrum of Cr–DNA damage. Experimental findings supporting this interpretation include the same levels of uptake-normalized Cr–DNA binding in –Asc and +Asc cells and lack of any effect of Asc introduced into cells following the completion of Cr(VI) reduction. The latter result demonstrates that cellular stress responses to Cr–DNA damage are not sensitized by the mere presence of Asc. Mismatch repair dependence of cell death points to the altered spectrum of Cr–DNA adducts as the most likely cause of increased toxicity of Cr(VI) by Asc. The status of mismatch repair plays a major role in the inhibition of replication of Cr/Asc-modified plasmids in human cells (17) but it has no significant effect on toxicity of oxidative DNA damage (17,40). The absence of p53 dependence in Asc-potentiated cell death further supports the argument that oxidative damage is not a significant factor in Cr(VI) lethality within our dose ranges. Processing of Cr–DNA damage by mismatch repair proteins generates DNA double-strand breaks as secondary genetic lesions (16,17). It is likely that increased formation of these breaks in response to the production of Asc–Cr–DNA cross-links (13) and a potentially increased yield of other ternary Cr–DNA adducts (41,42) is a major cause of enhanced lethality of Cr(VI) in cells containing normal levels of Asc. However, we cannot exclude the possibility that Cr(VI) reduction by Asc can selectively promote the formation of some forms of oxidized DNA bases that are abnormally processed by mismatch repair into more toxic damage.

Role of p53 in Cr(VI)-induced cell death
Induction of apoptotic responses to genotoxic insults is frequently caused by increased expression of p53-dependent pro-apoptotic genes (32). Generation of DNA breaks by ionizing radiation is a classic inducer of p53-mediated programmed cell death (43). We found that depletion of p53 by stable expression of shRNA had no significant effect on the levels of Cr(VI)-induced cytotoxicity in H460 and IMR90 cells irrespective of the presence or absence of intracellular Asc. These findings are not likely to be the result of some unusual phenotype of our cell lines because IMR90 are well-characterized primary cells and a recent screening of numerous established human cell lines has determined that unlike other transformed human cells, H460 cells accurately recapitulate normal p53-dependent stress responses to DNA damage (44). We have determined previously that clonogenic survival and apoptotic death were not affected by the p53 status in a pair of MLH1+ and MLH1–/– human colon HCT116 cells cultured without Asc supplementation (17). Lack of changes in protein levels and the extent of Ser-15 phosphorylation in IMR90 cells indicated the absence of activation of p53 signaling by Cr–DNA damage. In H460 cells, Cr(VI) did cause p53 stabilization and increased Ser-15 phosphorylation; however, these responses were independent of the cellular Asc status. Elevated levels of p53 did not lead to its transactivation in H460 cells because a reporter assay did not detect any increases in p53-regulated promoter activity. Increased protein levels and Ser-15 phosphorylation of p53 have also been observed previously in Cr(VI)-treated A549 lung cells grown under Asc-deficient conditions (19). Our results on unperturbed cell death in p53-depleted H460 cells indicate that neither protein nor Ser-15 phosphorylation levels are indicative of the activation of the p53-dependent apoptotic program. This interpretation is well supported by recent studies demonstrating that expression of pro-apoptotic gene targets requires involvement of other tightly regulated proteins and the presence of additional post-translational modifications at Lys residues of p53 (45–47). The observed inhibition of early apoptosis in Asc-deficient human fibroblasts transiently transfected with an E6-expessing vector (48) may not have necessarily been a consequence of low p53 levels, as E6 also inactivates a related pro-apoptotic transcriptional factor, p73 (49). In another case of mismatch repair-dependent toxicity, p73, not p53, played a key role in apoptosis of cisplatin-treated human cells (50). Since DNA breaks are potent inducers of p53 and its pro-apoptotic functions, it is possible that p53 can participate in the execution of death programs induced by high doses of Cr(VI). The threshold for the activation of p53-dependent apoptosis is expected to be dependent on the rate of Cr(VI) influx and the levels of glutathione, since the latter controls the production of oxidant-generated, toxic DNA single-strand breaks (27). Consequently, for cells containing low glutathione levels, the induction of p53-mediated toxic responses should occur at lower doses and a rapid loading of large amounts of Cr(VI) (for example, incubations with 0.2 mM Cr(VI) as in ref. 20) can also cause a shift toward oxidative DNA damage.

Overall, our findings on mismatch repair dependence but p53 independence of Cr(VI) toxicity in lung cells are in very good concordance with the molecular characteristics of chromate cancers that were found to exhibit high frequency of mismatch repair deficiency but contain a low number of p53 mutations (21–23). Thus, the loss of apoptotic signaling in response to Cr–DNA damage through inactivation of the mismatch repair system appears to be a very important event for the selective expansion of pre-malignant cells. Inactivation of p53 does not confer resistance to Cr(VI) toxicity, which explains why there is no significant selection for the loss of p53 during Cr(VI)-induced transformation process. Metabolism of Cr(VI) by cellular Asc produces toxic DNA lesions that are particularly strong activators of mismatch repair-mediated cell death. This would put additional selective pressure for outgrowth of mismatch repair-deficient clones exhibiting high rates of spontaneous mutagenesis.


   Acknowledgments
 
This work was supported by research grants ES008786 and ES012915 from National Institute of Environmental Health Sciences.

Conflict of Interest Statement: None declared.


   References
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received December 12, 2006; revised January 25, 2007; accepted February 3, 2007.


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M. F. Reynolds, E. C. Peterson-Roth, I. A. Bespalov, T. Johnston, V. M. Gurel, H. L. Menard, and A. Zhitkovich
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