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Carcinogenesis Advance Access originally published online on May 19, 2006
Carcinogenesis 2006 27(11):2316-2321; doi:10.1093/carcin/bgl076
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© The Author 2006. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

Lower mutagenicity but higher stability of Cr–DNA adducts formed during gradual chromate activation with ascorbate

George Quievryn1, Joseph Messer2 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; E-mail: anatoly_zhitkovich{at}brown.edu


   Abstract
Top
 Abstract
Introduction
 Materials and methods
 Results and discussion
 References
 
Recent epidemiological and risk assessment studies have found a very high risk of lung cancer among chromium(VI)-exposed workers even at permissible levels of exposure. However, mechanistic views on the key genotoxic role of transient Cr(V) intermediates were more consistent with the threshold or highly non-linear (heavy dose) models of genetic damage by intracellular Cr(VI). In this work, we examined the production of mutagenic DNA lesions during metabolism of Cr(VI) by its dominant reducer ascorbate (vitamin C) under conditions promoting increased yield of transient Cr forms. We found that slow reductive activation of Cr(VI) by limited concentrations of ascorbate resulted in a greater yield of DCFH-oxidizing Cr intermediates but these species were unable to cause DNA strand breaks. Cr(VI)–ascorbate reactions generated a high number of Cr–DNA adducts that were responsible for all mutagenic responses detected in Cr(VI)-treated pSP189 shuttle plasmids following their replication in human cells. Mutagenicity of DNA damage resulting from the reactions with increased stability of Cr intermediates was approximately four times lower relative to the conditions lacking detectable Cr(V) formation. Unlike other reactions, slow reduction of Cr(VI) with ascorbate produced Cr–DNA adducts that were more resistant to dissociation by chelators, suggesting multicoordinate binding of Cr(III) to DNA. Overall, our findings do not support the possibility that increased Cr(V) formation at depleted ascorbate levels modeling heavy dose exposures causes higher levels of mutagenic DNA damage.


   Introduction
Top
 Abstract
 Introduction
Materials and methods
 Results and discussion
 References
 
Human exposure to metal compounds containing chromium in the +6 oxidation state has been firmly linked to increased cancer incidence in the respiratory system (1–3). The main form of Cr(VI) at neutral pH is the chromate anion that readily enters cells due its structural similarity to the physiological sulfate and phosphate ions. Cr(VI) by itself is unreactive toward DNA and other important biological macromolecules but it is also a short-lived oxidative form. Inside the cells, Cr(VI) undergoes a series of reduction reactions resulting in the formation of thermodynamically stable Cr(III). Cellular reduction of Cr(VI) is predominantly a non-enzymatic process driven by non-protein thiols and ascorbate (4). In cultured cells, glutathione and cysteine are primarily responsible for the reductive conversion of Cr(VI) to Cr(III) but in the target tissues of chromate toxicity, the most important reducer is ascorbate (5–7). The reduction process usually generates a number of unstable intermediates and by-products, such as Cr(V), Cr(IV) and organic radicals. The relative amounts of the transient species vary dramatically depending on the nature of the reducer, its concentration and other experimental conditions. It has been suggested that the yield of the specific reaction intermediates could be a major factor determining DNA damaging activity of Cr(VI) (8).

Reduction reactions of Cr(VI) in the presence of its main biological reducer, ascorbate, are reasonably well characterized (9–11). Under conditions of at least 2-fold molar excess of ascorbate the first reduction step involves a two-electron transfer, which results in the formation of Cr(IV) as the principal Cr intermediate. The presence of Cr(V) in these reactions is undetectable by the ESR approaches. When chromate is present in equimolar or higher concentrations (limited reducer conditions), there is a significant production of Cr(V) originating from the reactions involving long-lived Cr(IV):

  1. Cr(IV) + Cr(VI) -> 2Cr(V)
  2. Cr(IV) + Cr(IV) -> Cr(V) + Cr(III)

Some studies have found that a higher production of Cr(V) correlated with elevated levels of certain forms of DNA damage (12–14). Increased production of Cr(V) and potentially greater genotoxicity of the resulting DNA damage could therefore represent a second, intracellular mechanism supporting the threshold model of chromate carcinogenesis (15). However, dose–response curves for mutagenic Cr–DNA adducts were inconsistent with the disproportionately greater yields of DNA damage at the high doses (4,16,17). Furthermore, recent studies on a large cohort of chromate-exposed US workers have found very high risk of lung cancer, with the estimated excess lifetime risk of lung cancer death at the permissible exposure limit being 255 per 1000 workers (3,18). These and other data have led OSHA to the decision earlier this year to lower the ambient standard for Cr(VI) from 52 to 5 µg/m3. Even under the new standard, there would be 10–45 excess deaths per 1000 exposed workers.

In this work, we sought to test the possibility that conditions associated with increased formation of Cr intermediates modeling heavy chromate exposures would generate more DNA damage or a qualitatively different spectrum of mutagenic lesions. Specifically, we assessed the formation of mutation-inducing lesions in DNA templates damaged in Cr(VI) reactions with limited ascorbate concentrations promoting increased stability of intermediate Cr oxidations. These conditions are likely to occur during human exposure to heavy doses of Cr(VI), causing transient depletion of ascorbate levels.


   Materials and methods
Top
 Abstract
 Introduction
 Materials and methods
Results and discussion
 References
 
Reagents
Chelex-100 resin and Bio-Gel P-30 columns were purchased from Bio-Rad (Hercules, CA). Stock solutions of 2',7'-dichlorofluoroscin diacetate (DCFH-DA) were from Molecular Probes (Eugene, OR). Potassium chromate (A.C.S. reagent) and [Cr(H2O)4Cl2]Cl·2H2O were from Aldrich (Milwaukee, WI). Supercoiled {Phi}X174 DNA was obtained from New England Biolabs (Beverly, MA). All other reagents were from Sigma (St Louis, MO). Buffers, ascorbate and chromate solutions (0.5 M, 10 mM and 10 mM stocks, respectively) were purified from the trace amounts of iron using Chelex-100 columns as described previously (19). The pSP189 vector was a gift from Dr Michael Seidman.

Formation of Cr–DNA adducts
For Cr(VI)-based reactions, typical reaction mixtures contained 25 mM MOPS buffer (pH 7.0), 0.2 mM ascorbate, 2 µg pSP189 DNA and various concentrations of K2CrO4. Samples were incubated at 37°C for 30 min, and then unbound Cr was removed by chromatography on Bio-Gel P-30 columns. DNA-containing solutions were supplemented with 200 mM NaCl and then 2 vols of ice-cold 100% ethanol were added. DNA was precipitated overnight at 4°C, collected by centrifugation (10 000x g, 10 min, 4°C) and washed three times with cold 70% ethanol. This purification procedure removes ionic Cr–DNA complexes allowing the determination of stable Cr(III)–DNA adducts (17,19). Air-dried DNA pellets were dissolved in deionized H2O and then used either for cell transfections or quantification of DNA adducts. Some Cr(VI) treatments of DNA were also performed in 25 mM Na-phosphate buffer (pH 7.0) or in 25 mM MOPS buffer containing 5 mM EDTA. For Cr(III)-based reactions, pSP189 DNA was treated with 40 µM [Cr(H2O)4Cl2]Cl·2H2O for 30 min at 37°C in 25 mM MES (pH 6.0). DNA was passed through P-30 columns and precipitated with ethanol as above. The amount of DNA-bound Cr was determined by graphite furnace atomic absorption spectroscopy using Zeeman background correction (model 41002L; Perkin-Elmer GF-AAS). Instrumentation settings were as described previously (17).

Dissociation of chromium–DNA complexes
The formation of Cr–DNA adducts was performed with 50 µM Cr(VI) in 25 mM MOPS buffer as described above. Unbound Cr was removed by the Bio-Gel P-30 chromatography followed by the ethanol/NaCl precipitation. Cr-modified plasmids were dissolved in 50 mM sodium phosphate (pH 7.0) and then incubated for 24 h at 37°C. Released Cr(III) was removed by a passage through P-30 columns and purified DNA was assayed for Cr or used for mutagenesis experiments. A control set of Cr-adducted DNA was incubated in 50 mM MOPS (pH 7.0). The amount of DNA-bound Cr was determined by GF-AAS.

Shuttle-vector mutagenesis experiments
The SV40-based pSP189 vector containing the supF gene as a mutagenic target was used to assess the formation of mutagenic DNA lesions (20). This vector contains the SV40 origin of replication and encodes large T-antigen, which permits replication of this plasmid in a broad variety of human cell lines. Control and Cr-modified plasmids were replicated in human diploid fibroblasts immortalized with SV40 virus (HF/SV cell line). Detection of supF mutants was performed using Escherichia coli MBL50 strain (a gift from Dr C. Pueyo). This indicator E.coli strain contains dual araC and lacZ amber mutations suppressed by the supF gene (21). Inactivating mutations in the araC gene allow selection of the colonies with a mutated supF gene on plates containing L-arabinose. Amber mutation in the lacZ gene permits white/blue color screening of colonies. Control and Cr-treated plasmids were transfected into the cells using TransFast reagent (Promega) according to manufacturer's recommendations. Plasmids were propagated for 48 h and their replicated progeny was recovered by a plasmid isolation kit from Qiagen. DNA was precipitated with ethanol and dissolved in deionized H2O. Plasmids were electroporated into the E.coli MBL50 according to Hanahan et al. (22). The total number of bacterial transformants was determined on the minimal agar plates containing 30 µg/ml ampicillin and 0.5 µg/ml chloramphenicol. Mutant selection was performed on plates additionally containing 2 mg/ml L-arabinose. Mutation frequency was calculated as the ratio of ampicillin/arabinose resistant to ampicillin resistant colonies in each sample.

Kinetics of chromate reduction
Time-course of chromate reduction was monitored by recording absorbance at 372 nm using a Shimadzu UV1601 spectrophotometer. Reaction mixtures contained 25 mM MOPS or phosphate buffer (pH 7.0), 50 µM chromate and indicated concentrations of ascorbate. To ensure a rapid mixing of the reagents, the reduction was initiated by the addition of 0.5 ml of 100 µM Cr(VI) solution to the cuvette containing 0.5 ml of pre-warmed buffer-ascorbate mixtures. All reactions were performed at 37°C using electronically controlled temperature unit. Absorbance values were recorded every 10 s.

Detection of oxidants by DCF fluorescence
The production of oxidizing species during chromate reduction was monitored by the formation of fluorescent 2',7'-dichlorofluoroscein (DCF), which is a product of DCFH oxidation. DCFH-diacetate ester was hydrolyzed in the presence of 10 mM NaOH for 30 min followed by neutralization with Chelex-treated phosphate buffer (pH 7.0). Samples contained 10 µM DCFH, 25 mM buffer (pH 7.0), 0.2 or 1 mM ascorbate and various concentrations of chromate. All reactions were performed at 37°C in 96-well plates that were incubated inside the microplate reader. Fluorescence measurements were recorded using 485 nm excitation and 535 nm emission filters (SpectraFluor Plus microplate fluorometer, Tecan).

Detection of DNA strand breaks
The presence of DNA breaks was detected based on the conversion of supercoiled {Phi}X174 DNA into a nicked, open circular form. Reaction samples contained 0.3 µg supercoiled {Phi}X174, 25 mM MOPS or phosphate buffer (pH 7.0), 0.2 mM ascorbate and 0–200 µM chromate in the total volume of 25 µl. After incubation at 37°C for 30 min, the samples were placed on ice, mixed with 6x Ficoll buffer (15% Ficoll 400, 0.25% bromphenol blue and 0.25% xylene cyanol) and loaded on to 1% agarose-TAE gels. DNA bands were visualized with ethidium bromide and analyzed using GelDoc 2000 (Bio-Rad).


   Results and discussion
Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
References
 
To create conditions with the increased yield and stability of intermediate Cr oxidations, we chose to use reactions with the low ascorbate concentration of 0.2 mM. This experimental set-up would mimic the reducer-depleting conditions of exposure to heavy Cr(VI) doses and minimize the reactions of oligomerization of different Cr species (4). The alternative approach based on the use of millimolar concentrations of Cr(VI) and ascorbate near 1:1 ratio would not be biologically relevant and would cause massive amounts of DNA damage, precluding plasmids from being able to replicate in human cells. We have found previously that modification of pSP189 DNA in the presence of 0.2 mM Cr(VI)–1 mM ascorbate created sufficient levels of DNA lesions to block replication of >99.5% of these plasmids (7). Such low recoveries of replicated plasmids make it difficult to perform reliable measurements of mutation frequencies and raise concerns about potentially non-specific effects caused by excessively heavy damage to the DNA templates.

Kinetics of chromate reduction and generation of DCFH-oxidizing species
First, we examined the time-course of Cr(VI) reduction and the formation of DCFH-oxidizing Cr intermediates in the reactions containing 0.2 and 1 mM ascorbate. Reduction kinetics of Cr(VI) in the presence of 0.2 mM ascorbate was monophasic and essentially identical in both MOPS and phosphate buffers (Figure 1A). In this regard, ascorbate reactions were similar to those containing cysteine (23) but differed from glutathione-driven reductions that are biphasic, particularly at low concentrations and in the presence of phosphate ions (24). Reduction rates for Cr(VI) in the samples containing 0.2 mM ascorbate were approximately five times slower relative to 1 mM (Figure 1B), which is consistent with the first-order reaction with respect to reducer. Using the rates of reduction for thiols and their typical cellular concentrations (7,25), we calculated that 0.2 mM ascorbate would still be responsible for metabolism of 88.5% of cellular Cr(VI) (97.5% for 1 mM). Slower reduction with the lower 0.2 mM concentration resulted in the increased formation of intermediate Cr species, detected by the oxidation of the redox-sensitive dye DCFH to its fluorescent form, DCF (Figure 2A and B). Oxidation of DCFH in chromate reactions containing Chelex-purified reactants has been found previously to reflect the levels of Cr(V) production (25,26). In comparison with 1 mM, samples containing 0.2 mM ascorbate generated three to five times higher amounts of oxidized DCFH (Figure 2B), which is consistent with the ability of slow reduction reactions to yield increased amounts of intermediate Cr forms.


Figure 1
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  		Fig. 1 Reduction of chromate by ascorbate. All reactions contained 50 µM chromate and were performed in either 25 mM MOPS (pH 7.0) or 25 mM sodium phosphate buffer (pH 7.0). (A) Time-course of chromate reduction by 0.2 mM ascorbate. (B) Half-life values for chromate in the presence of different concentrations of ascorbate. Values are means ±SD of three to six determinations.

 

Figure 2
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  		Fig. 2 Generation of DCFH-oxidizing species during Cr(VI) reduction. Samples contained 25 mM MOPS or phosphate buffer (pH 7.0), 10 µM DCFH, 0.2 or 1 mM ascorbate and 0–200 µM chromate. Data are means ± SD of three independent determinations. (A) Time-course of DCFH oxidation by 50 µM chromate. (B) Dose-dependent oxidation of DCFH by chromate. DCF fluorescence was recorded after 30 min incubation.

 
Cr(VI) reduction causes extensive Cr–DNA binding but not DNA breaks
A potential formation of DNA nicking species during Cr(VI) reduction was assessed by a sensitive plasmid relaxation assay that detects DNA breaks by the conversion of supercoiled DNA molecules into their open circular form. The sensitivity of this assay is 0.5–1 break per 50 000 DNA base pairs (25). Neither MOPS- nor phosphate-based reactions generated detectable amounts of DNA breakage during chromate reduction (Figure 3A). In MOPS-containing samples, however, supercoiled (intact) DNA molecules exhibited progressively slower electrophoretic mobility with increasing doses of Cr(VI). The corresponding bands also displayed broadening and much weaker staining with ethidium bromide. All these changes reflected structural distortions caused by Cr–DNA binding because Cr(III) sequestration by phosphate ions prevented any alterations in the supercoiled DNA bands. The ability of phosphate buffer to block the formation of Cr–DNA was directly confirmed by measurements of DNA-bound Cr (Figure 3B). Both diminished electrophoretic mobility and decreased ethidium bromide staining of Cr(VI)-treated supercoiled plasmids are caused by DNA-bound Cr(III), inducing unwinding of the helix and base destacking, respectively (23). EDTA, another Cr(III) chelator, was also highly effective in eliminating the formation of Cr–DNA adducts (Figure 3B). Reactions in MOPS buffer, which has a low ability to bind Cr(III) (25), generated a very high number of Cr–DNA adducts in a dose-dependent manner. The highest Cr(VI) concentration of 200 µM produced 9600 Cr adducts/50 000 DNA base pairs. Using a conservative estimate of the detection sensitivity of the plasmid relaxation assay (1 break/50 000 bp), these results indicate that Cr–DNA adducts exceeded the number of breaks by at least 10 000:1. Similar estimates for the relative yields of Cr–DNA complexes and strand nicking were obtained for iron-free Cr(VI) reactions containing cysteine (19,25), glutathione (24) or millimolar concentrations of ascorbate (7). Induction of modest levels of DNA breaks observed in earlier studies (27,28) has been traced recently to the presence of residual amounts of iron in the commercial sources of the reducers (19,24). Ascorbate stocks used in this work were rigorously purified from any residual metals by Chelex-100 chromatography and used immediately to avoid autooxidation, which results in the production of H2O2 (29,30) and is the main cause of DNA nicking species in Cr(VI) reactions (24).


Figure 3
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  		Fig. 3 Formation of DNA damage in chromate–ascorbate reactions. Reactions contained 0.2 mM ascorbate, 0.3 µg {Phi}X174 DNA (breakage assay) or 2 µg pSP189 DNA (Cr–DNA binding), 25 mM buffer (MOPS or phosphate or MOPS + EDTA) and 0–200 µM chromate. Results are means ± SD from three to six independent determinations. (A) Agarose gels of Cr(VI)-damaged plasmid DNA. (B) Chromium-DNA binding.

 
Mutagenic damage resulting from Cr(VI)–ascorbate reactions
Measurements of individual forms of DNA damage are either incomplete or suspected to be incomplete, because it is usually difficult to ensure that all types of DNA lesions have been detected. For example, our analyses of plasmid integrity by agarose gels would have missed the presence of oxidized DNA bases, although their formation is typically accompanied by the generation of DNA strand breaks (31,32). Chemical approaches are also limited by their inability to predict the biological importance of one class of DNA lesions over another. Therefore, to further characterize the nature of genotoxic damage resulting from chromate–ascorbate reactions, we employed the pSP189 shuttle vector to examine the presence of mutagenic damage. The principal advantage of this approach is that it allows the assessment of mutagenicity of DNA damage in intact human cells while permitting manipulations of the reactions to selectively alter the formation of different types of reactive species or DNA lesions.

We found that modification of pSP189 plasmids with Cr(VI)–ascorbate in MOPS buffer resulted in the extensive formation of mutagenic lesions (Figure 4). At the highest Cr(VI) concentration, there was ~30-fold increase in the number of supF mutants over control. HF/SV cells used for replication of pSP189 vectors are DNA repair-competent (33) and, therefore, mutagenic responses detected in our experiments were not amplified by the unusually long persistence of any particular form of DNA damage. Based on the slopes of dose–response curves, we calculated that the yield of mutations in the samples treated with Cr(VI)–0.2 mM ascorbate was 4.4 times lower in comparison with the reactions containing 1 mM ascorbate (7). Thus, the slow reduction process allowing greater stability of intermediate Cr forms is actually less mutagenic than a more rapid process lacking significant production of Cr(V). Increased mutagenicity of DNA templates damaged in the presence of millimolar ascorbate is related to the elevated formation of highly mutagenic ascorbate–Cr(III)–DNA crosslinks in these reactions (7,17).


Figure 4
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  		Fig. 4 Production of mutagenic DNA lesions requires Cr–DNA binding. The presence of mutagenic DNA damage was analyzed using the pSP189 shuttle vector. Plasmids were modified in the reactions containing 0.2 mM ascorbate, 0–200 µM chromate and 25 mM buffer, pH 7.0. Purified vectors were transfected into human HF/SV fibroblasts, replicated progeny was recovered 48 h later and the number of the supF mutants was determined following electroporation of isolated plasmids into MBL50 E.coli strain. Results are means ± SD from four independent DNA transfections. MOPS—reactions contained 25 mM MOPS, Pi—reactions contained 25 mM phosphate, EDTA—reactions contained 25 mM MOPS and 5 mM EDTA.

 
Addition of Cr(III) chelators phosphate or EDTA to reduction reactions essentially abolished mutagenic responses (Figure 4), demonstrating the crucial role of Cr–DNA adducts in the induction of mutations by Cr(VI). Plasmids incubated with 100–200 µM Cr(VI) in phosphate buffer did show ~2-fold increase in the mutation frequency, which probably reflected the presence of residual amounts of Cr–DNA complexes under these conditions (4.5–6 adducts/1000 bp, 2.3–4.4% of MOPS samples). EDTA samples had no detectable Cr–DNA binding (Figure 3) and showed no residual mutagenic activity. The dependence of mutagenic responses on Cr–DNA binding is also consistent with the absence of DNA strand breaks (Figure 3A), which is the most common marker of oxidative DNA damage. Although we did not directly test the formation of oxidized DNA bases, their potential presence would have been revealed by the mutagenicity of plasmids lacking Cr–DNA adducts. Other investigators have detected recently a significant formation of oxidized guanine products in Cr(VI)–ascorbate reactions employing concentrations of the reagents similar to those in our work (14). It remain unclear whether dG oxidation did not occur in our iron-free reactions or the extent of this process was too low to cause mutagenic responses in vectors replicated in repair-proficient human cells. It is also possible that advanced products of dG oxidation are primarily toxic (replication-blocking) but not very mutagenic.

Stability of Cr–DNA adducts
In these experiments, we sought to explore the possibility that Cr(VI) reduction by limited concentrations of ascorbate could result in a more stable coordination of Cr(III) with DNA duplex. The stability of Cr–DNA binding was assessed by the susceptibility of DNA-bound Cr to release by phosphate ions (23). Time-course experiments showed that the majority of Cr–DNA adducts generated in the reactions containing Cr(VI)–1 mM ascorbate were disrupted during incubation in phosphate buffer for 24 h (Figure 5A). Since ~75% of all adducts produced in 1 mM ascorbate-containing reactions are binary Cr(III)–DNA complexes (7), these results indicated that in addition to ternary ascorbate–Cr(III)–DNA complexes, binary adducts were also susceptible to phosphate-induced dissociation. However, for reactions with 0.2 mM ascorbate, we found that on average 66.2 ± 8.5% of Cr–DNA complexes were still present after phosphate post-treatments (Figure 5B). To further test the stability of DNA adducts, we examined the ability of pSP189 plasmids to induce mutations before and after incubation with phosphate. In accordance with the weak dissociation of adducts, post-incubation with phosphate led to only a modest 1.5-fold reduction in mutational responses using DNA templates treated with Cr(VI)–0.2 mM ascorbate (Figure 5C). Control experiments showed that selectively formed binary Cr(III)–DNA adducts were sensitive to dissociation by chelators as evidenced by a >5-fold decrease in the mutagenicity of phosphate-treated samples (Figure 5C). Thus, binary adducts formed in chromate–0.2 mM ascorbate samples were more stable than the same type of Cr–DNA complexes produced with Cr(III) chloride or in reactions with Cr(VI)–1 mM ascorbate. It appears that under conditions of low concentrations of a competing ligand, such as ascorbate, newly formed Cr(III) has more than one reactive coordination site permitting bifunctional mode of binding to DNA. Bidentate coordination to DNA is believed to involve the formation of microchelates between phosphate group and N7-dG (23,34). A fraction of DNA-bound Cr(III) in low ascorbate samples could also be present in the form of multicoordinated oligomers (4). The presence of poorly leaving water ligands in Formula and [Cr(H2O)4]Cl+ complexes that exist in Cr(III) chloride solutions apparently favored the monodentate type of DNA binding, which is more easily reversable by phosphate ions. Therefore, reactions with inorganic Cr(III) complexes do not appear to model completely the range of potential Cr–DNA interactions occurring during chromate reduction with limited amounts of reducers. Differential stability of Cr–DNA complexes to dissociation during sorbtion-elution cycles at NENsorb columns was probably responsible for the reported dependence of Cr–DNA binding on the presence of Cr(V) (35).


Figure 5
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  		Fig. 5 Mutagenic responses and levels of Cr–DNA adducts in pSP189 plasmids post-incubated with phosphate. The pSP189 DNA was modified with Cr(VI) or Cr(III) as described in Materials and Methods. One half of the purified DNA was incubated with 50 mM phosphate (pH 7.0) for the indicated periods of time at 37°C followed by passing through Bio-Gel P-30 columns. DNA was ethanol-precipitated, dissolved in deionized water and tested for the amount of bound Cr and mutagenicity. (A) Time-course of Cr release from DNA complexes formed in the reaction of 100 µM chromate with 1 mM ascorbate. Results are means ± SD from four independent analyses. (B) Stability of Cr–DNA complexes formed in chromate–0.2 mM ascorbate reactions. Cr-modified DNA was incubated with phosphate for 24 h. Shown are means ± SD from three independent determinations. (C) Mutagenic responses before and after incubation of Cr-modified pSP189 plasmids with phosphate for 24 h. CrVI - plasmids were modified with 50 µM chromate–0.2 mM ascorbate. CrIII - pSP189 DNA was reacted with 40 µM chromium(III) chloride. Results are means ± SD from four independent transfections.

 
Conclusions
Reductive activation of Cr(VI) by low ascorbate concentrations generates Cr–DNA damage with lower mutagenicity than the corresponding reactions with physiological amounts of this reducer. Our findings do not support the hypothesis that increased production of intermediate Cr oxidations in ascorbate-driven reactions causes higher levels or more potent forms of mutation-inducing DNA lesions by Cr(VI). Thus, from the point of view of intracellular Cr(VI) metabolism by its dominant reducer, heavy dose exposures associated with depletion of ascorbate would not cause a disproportionately higher formation of biologically important DNA damage. Our results point to a potential possibility that linear extrapolations from high doses may actually underestimate the levels of certain forms of genetic damage occurring at low-level exposures. Decreased yields of mutagenic ternary Cr–DNA adducts at high Cr(VI) doses were also detected in experiments with cultured cells (16). Chromate carcinogenesis is certainly a much more complex process than just formation of mutagenic DNA damage, and it also involves a selection for microsatellite unstable, mismatch repair-deficient cells (36,37). Further studies are needed to assess the ability of low versus high doses of Cr(VI) to promote post-DNA damage steps of chromate carcinogenesis, such as the rate of outgrowth of cellular clones lacking mismatch repair.


   Footnotes
 
1Present address: Draeger Medical Systems, Danvers, MA 01923, Back

2Present address: Genzyme, Framingham, MA 01701, USA Back


   Acknowledgments
 
This work was supported by grants R01 ES008786 and P42 ES013660 from the National Institute of Environmental Health Sciences.

Conflict of Interest Statement: None declared.


   References
Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 

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Received March 23, 2006; revised May 10, 2006; accepted May 15, 2006.


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