Hypervalent Chromium Mimics Reactive Oxygen ... - ACS Publications

Brooke D. Martin, John A. Schoenhard, and Kent D. Sugden*. Department of Chemistry, 6128 Burke Laboratory, Dartmouth College,. Hanover, New Hampshire ...
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Chem. Res. Toxicol. 1998, 11, 1402-1410

Articles Hypervalent Chromium Mimics Reactive Oxygen Species As Measured by the Oxidant-Sensitive Dyes 2′,7′-Dichlorofluorescin and Dihydrorhodamine Brooke D. Martin, John A. Schoenhard, and Kent D. Sugden* Department of Chemistry, 6128 Burke Laboratory, Dartmouth College, Hanover, New Hampshire 03755-3564 Received June 30, 1998

Intracellular metabolism of the carcinogen chromate [Cr(VI)] produces the oxidative stress and oxidative DNA damage associated with its genotoxicity. Such oxidative stress has previously been measured by fluorescence using oxidant-sensitive dyes and attributed to the formation of reactive oxygen species (ROS). However, metabolism of Cr(VI) also produces Cr(IV) and Cr(V) which can directly damage biological macromolecules without forming ROS. We used the high-valence chromium species, bis(2-ethyl-2-hydroxybutyrato)oxochromate(V) [Cr(V)-EHBA], to test whether high-valence chromium would also react with the oxidantsensitive dyes 2′,7′-dichlorofluorescin (DCFH) and dihydrorhodamine (DHR). Cr(V)-EHBA caused both dyes to fluoresce over a wide dynamic range and under conditions which indicated that Cr(V) had reacted directly with both dyes without first forming a diffusible radical species. Dimethylthiourea (DMTU) and ethanol did not affect Cr(V)-induced fluorescence in vitro or Cr(VI)-induced fluorescence in A549 cells. Under the same conditions, ethanol and DMTU increased the extent of hydrogen peroxide-induced fluorescence. As chromium-induced fluorescence was unaffected by radical scavengers and was qualitatively different from hydrogen peroxide-induced fluorescence, we conclude that DCF and R123 fluorescence in chromatetreated A549 cells is a qualitative and cumulative measure of intracellular Cr(V) formation and not ROS.

Introduction The toxicity, mutagenicity, and carcinogenicity of chromium compounds are well-established phenomena with more than one hundred years of supporting epidemiological evidence (1). Of the two environmentally available forms of chromium, hexavalent and trivalent, only the hexavalent form has been clearly and unequivocally demonstrated to be associated with all toxic parameters and classified as a human carcinogen (2). However, in vitro hexavalent chromium neither reacts with nor binds to DNA but requires reductive metabolism to become mutagenic (3, 4). It is, however, the hexavalent form of chromium which is readily available to living cells as it is isostructural with respect to phosphate and sulfate and enters the cell via the same nonspecific anion transport channel (5, 6). Reductive metabolism of chromium occurs within the cell, generating toxic chromium species intracellularly. Such intracellular reduction necessarily causes intracellular oxidation, and it has been speculated that this leads to concomitant formation of intracellular oxidative stress which should be considered as a part of the cohort of toxic effects exerted by chromium. The observation that the mutagenicity of Cr(VI) in the Salmonella reversion assay was greatly * To whom correspondence should be addressed.

enhanced in the presence of molecular oxygen (7) and that DNA damage was dependent upon the presence of molecular oxygen (8) fueled speculation that activated oxygen may be responsible for the mutagenic and toxic properties of chromate (9, 10). The idea of the involvement of activated oxygen was further supported by the observations that hydroxyl radical scavengers and catalase reduced DNA damage derived from the reduction of chromate by glutathione in the presence of iron (11), and that the catalase inhibitor aminotriazole sensitized cultured rodent FAO cells to chromate toxicity (12). Accordingly, hydrogen peroxide, specifically as hydroxyl radical [hydrogen peroxide does not react with DNA unless transition metals are present (13)], was thought to be the potential ultimate DNA-damaging agent. Hydrogen atom abstraction is one mechanism used to explain DNA damage produced by hydroxyl radical (14, 15). However, hydrogen atom abstraction has also been shown to be the preferred mode of oxidation of aliphatic organic substrates by a synthetic Cr(V) complex bis(2ethyl-2-hydroxybutanoate)oxochromate [[OCr(EHBA)2] or Cr(V)-EHBA]1 (16). It has been shown that hypervalent chromium species can form oxidative lesions on nucleotides and DNA in the presence or in the absence of oxygen (17, 18). These studies failed to detect a diffusible radical species, such as the hydroxyl radical, which could

10.1021/tx9801559 CCC: $15.00 © 1998 American Chemical Society Published on Web 10/17/1998

Chromium(V) Oxidation of Fluorescent Dyes

be trapped with ethanol and DMPO in EPR spin-trapping experiments (18). Therefore, it was concluded that hypervalent chromium is capable of directly oxidizing biological macromolecules such as DNA without needing to first form diffusible radical species. The reactivity of hypervalent chromium may therefore be analogous to that of reactive oxygen species under certain conditions. Monitoring intracellular oxidants requires expensive and specialized equipment, and it is not always certain whether the oxidants measured are intra- or extracellular in origin. Therefore, the oxidant-sensitive dye 2′,7′dichlorofluorescin diacetate (DCFH-DA) is often used in cells in culture (19) and in tissues (20, 21) to monitor the formation of reactive oxygen species (ROS). Dihydrorhodamine 123 (DHR) is used to measure ROS formation (22) in cells which maintain a transmembrane potential (23, 24). These dyes have several advantages over other methods in that they are straightforward to use and rapidly give reproducible results. To examine the role of ROS in the toxicity and genotoxicity of chromium, the dye DCFH-DA has been used previously as an indicator of intracellular oxidative stress (25) or intracellular hydrogen peroxide levels (26, 27) in chromium-treated cells in culture. However, hypervalent chromium [Cr(V)] has been shown to form intracellularly under the conditions used in those studies (25, 28, 29). Oxidation of DCFH to its fluorescent form, DCF, is thought to proceed via two sequential proton and electron, or if performed as a simultaneous event, hydrogen atom abstractions (20). Similarly, DHR is converted to its fluorescent form rhodamine 123 (R123) by abstraction of a tertiary hydrogen atom (30). As hypervalent chromium species can mimic in part the reactivity of hydroxyl radical, by performing the same hydrogen atom abstraction chemistry, we sought to examine the reactivity of these species with these two oxidant-sensitive dyes. We further sought to determine if it is possible to distinguish the reactivity of hypervalent chromium from that of ROS. Detailed studies on the reactivity of ROS with DCF have already been conducted by Le Bel et al. (20) using a variety of hydroxyl radical scavengers. We investigated these scavengers for their ability to affect the stability of Cr(V) in solution. We included dimethylthiourea (DMTU) in our studies as it is also a potent hydroxyl radical scavenger (31), which was shown to suppress the generation of hydroxyl radicals by Cr(V) and hydrogen peroxide without affecting the formation of Cr(V) in whole cells systems (32). We determined that Cr(V) directly oxidizes both DCFH and DHR to their fluorescent forms under conditions in which ROS cannot form. Using radical scavengers previously studied with respect to DCF fluorescence, we found that those which did not affect the decay rate of Cr(V) did not affect the fluorescence intensity of either dye. However, stabilization of Cr(V) against disproportionation with EHBA led to increased fluorescence. We found that the radical trap mannitol also stabilized Cr(V) against disproportionation by chelating directly to the 1 Abbreviations: EHBA, 2-ethyl-2-hydroxybutyric acid; Cr(V)EHBA, bis(2-ethyl-2-hydroxybutyrato)oxochromate(V); DCFH, 2′,7′dichlorofluorescin; DCF, 2′,7′-dichlorofluorescein; DCF-DA, 2′,7′dichlorofluorescin diacetate; DHR, dihydrorhodamine 123; R123, rhodamine 123; DMSO, dimethyl sulfoxide; EtOH, ethanol; DMTU, 1,3-dimethyl-2-thiourea; HRP, horseradish peroxidase; O2•-, superoxide anion; OH•, hydroxyl radical; ROS, reactive oxygen species; DMPO, 5,5-dimethylpyrroline N-oxide.

Chem. Res. Toxicol., Vol. 11, No. 12, 1998 1403

metal center to form a new Cr(V) species; however, only DHR could react with this new species, and DCF fluorescence was diminished. In cultured human lung A549 cells, we determined that the majority, if not all, of the fluorescence could be attributed to the effects of Cr(V) and that the levels of fluorescence observed were consistent with chromium uptake and intracellular Cr(V) formation. In marked contrast was a strong increase in DCF fluorescence in hydrogen peroxide-treated lung cells which had been preincubated with the radical scavenger ethanol (EtOH) or with DMTU. This increase in fluorescence was ascribed to the ability of the scavengers to trap the radical and form a more stable species, thereby increasing the probability of reaction with the DCFH dye. Trapping of the radical may also prevent degradation of the fluorescent form of the dye (DCF) by excess free radicals.

Experimental Section Materials. 2′,7′-Dichlorofluorescin diacetate (DCFH-DA) and dihydrorhodamine 123 (DHR) were purchased from Molecular Probes Inc. (Eugene, OR). 1,3-Dimethyl-2-thiourea (DMTU), dimethyl sulfoxide (DMSO), and 2-ethyl-2-hydroxybutyric acid (EHBA) were from Sigma Chemical Co. Ethanol (EtOH) and mannitol were purchased from Fisher Scientific. Tissue Culture. A549 cells were purchased from the American Type Culture Collection (ATCC) and were maintained in Ham’s 10% media with 10% FBS and antibiotic/antimycotic (Sigma; 10 000 units/mL penicillin, 10 mg/mL streptomycin, and 25 µg/mL amphotericin B) buffered with carbonate. Cells were maintained at 37 °C in a 5% CO2 atmosphere. Synthesis of Cr(V)-EHBA. The sodium salt of bis(2-ethyl2-hydroxybutyrato)oxochromate(V) [Cr(V)-EHBA] was prepared using the method of Krumpolc and Rocek (33) and the purity confirmed by IR spectroscopy in a Nujol mull: 3510, 1680, 1312, 1257, 1177, 1045, 998, 961, 845, 815, and 712 cm-1. Caution: Cr(VI) is a known human carcinogen, and Cr(V) complexes are potentially carcinogenic. Appropriate precautions should be taken in handling these materials. Fluorescence Measurements. For in vitro studies, DCFHDA must be de-esterified to generate the oxidation substrate DCFH. The de-esterification reaction was performed by the method of Cathcart et al. (34) by mixing 125 µL of a 1.5 mM DCFH-DA solution in EtOH with 0.5 mL of 0.01 N NaOH for 30 min at room temperature in the dark. The mixture was then neutralized with 2.5 mL of demetalated 20 mM NaH2PO4 buffer (pH 7.0) to give 60 µM activated DCFH dye stock. This activated stock was then diluted so as to give a final dye concentration of 10 µM for each set of reaction conditions. DHR does not require activation prior to in vitro use. Radical quenching reactions were carried out in 96-well Nunclon plates using 50 µL aliquots of the DCFH or DHR dye, 50 µL of chromium, and 50 µL of the radical scavenger where appropriate, with a final volume of 150 µL. For the in vitro Cr(V) reactions, 50 µL aliquots of the dye were used, and reactions were initiated by addition of 25 µL of Cr(V) to give a final volume of 75 µL. All Cr(V) dilutions were performed in water as rapidly as possible to prevent loss of Cr(V) due to disproportionation. Argon studies were conducted using rigorously argon-degassed solutions, and all dilutions were conducted in a glovebag under an argon atmosphere. The 96-well plate was additionally maintained under an argon blanket within the glovebag and the plate sealed with Dura-Seal Laboratory Stretch Film prior to the first fluorescence reading. Fluorescence studies in cell culture were carried out on cells passaged at a density of 40 000 cells/well into a Nunclon 96well plate and incubated for 72 h at 37 °C under a 5% CO2 atmosphere. After 72 h, the cells were gently washed three times with sterile PBS to remove residual media. The washed cells were pre-equilibrated with 15 DCFH-DA or DHR in PBS

1404 Chem. Res. Toxicol., Vol. 11, No. 12, 1998 containing 5 mM glucose, 0.3 mM CaCl2, and 0.62 mM MgCl2 (PBS+). For the whole cell radical quenching reactions, the solution also contained 5 mM EtOH or 5 mM DMTU as indicated. After 30 min, 25 µL of Cr(VI) or hydrogen peroxide was added to give the final concentration indicated. The final concentrations of the dyes were 10 µM, and the total volume for these experiments was 75 µL to minimize solution volume above the adherent monolayer of cells. Fluorescence studies for both the in vitro and cell culture assays were carried out using a Perseptive Biosystems Cytofluor II instrument with an excitation wavelength of 485 nm and an emission wavelength of 530 nm for both DCF and R123. EPR Conditions. Cell-free EPR spectra were recorded using a Bruker ESP-300 spectrometer. The spectral parameters were 100 kHz field modulation, 1.0 G modulation amplitude, 5.12 ms time constant, 9.768-9.772 microwave frequency, 1 × 105 receiver gain, 2 mW microwave power, and 3380-3580 G sweep width with a 21 s scan time. Single scans were collected every minute using an automation subroutine. Typical reactions were carried out on 2.0 mL volumes at room temperature (RT) in chelexed 20 mM phosphate buffer at the appropriate pH. Measurements were carried out on ca. 100 µL volumes drawn into a capillary tube sealed on one end with Dow-Corning highvacuum grease and placed in a quartz EPR tube. The g values were determined with respect to 2,2-diphenyl-1-picrylhydrazyl radical (DPPH, g ) 2.0036). Frozen whole cell EPR spectra were collected using a liquid nitrogen finger dewar. Spectral parameters were 100 kHz field modulation, 1.0 G modulation amplitude, 5.12 ms time constant, 9.439-9.464 microwave frequency, 1 × 105 receiver gain, 12 mW microwave power, and 3350-3450 G sweep width and a 168 s scan time. The signals were averaged over nine scans, and all scans were adjusted for a chromium-free background signal. The g values were determined with respect to 2,2-diphenyl-1picrylhydrazyl radical (DPPH, g ) 2.0036). Cells were harvested by scraping from two T-75 flasks and resuspended in 10.0 mL of complete media for counting using a Coulter Counter II. Cells were immediately recentrifuged and resuspended in 200300 µL of complete media, and Cr(VI) was added to give the final concentration indicated. Cell suspensions containing Cr(VI) were incubated at 37 °C for 25 min before decanting into quartz EPR tubes and flash-freezing in liquid nitrogen. A frozen Cr(V) standard curve was obtained for quantitation by using solutions of Cr(V)-EHBA in 100 mM EHBA (pH ∼3-4). Toxicity Assays. Twenty-four hour postconfluent cultures of A549 cells between passage numbers 10 and 20 were treated with sterile filtered solutions of Cr(VI) (from a sodium dichromate stock solution) or hydrogen peroxide. After 4 h, cells were harvested by trypsinization and seeded at a density of 200 cells/ plate [400 cells/plate for the 80 µM Cr(VI) dose]. Cells were incubated in Ham’s media as described above for 10-14 days. The medium was aspirated off, and the individual colonies were visualized by crystal violet staining. Colonies were counted, and survival was measured in comparison to control plates. Whole Cell Chromium Measurements. Twenty-four hour postconfluent A549 cells were incubated with a filter-sterilized solution of 10 µM Cr(VI) (as sodium dichromate). At the indicated time, cells were washed three times with PBS and harvested by trypsinization and either the cells were counted by a Coulter Counter II or an aliquot was removed for protein determination by Coomassie Blue. Cells were resuspended in ultrapure nitric acid (ULREX II, Ultrapure reagent), and the sample was digested until clear of particulate material. Chromium was measured by atomic absorption spectroscopy using a Perkin-Elmer 503 atomic absorption spectrophotometer with an HGA-2100 graphite furnace. Furnace settings were taken from ref 35. Absorption, measured as peak height, was monitored at 357.8 nm and compared to a standard curve prepared by dilution of a 1000 ppm Cr standard (Aldrich). Chromium content was normalized for cell number and/or protein content. Chromium content was converted to intracellular concentration using the numbers 0.323 µg of protein/106 cells and 3.56 pL/

Martin et al.

Figure 1. Concentration-dependent and oxygen-independent Cr(V)-EHBA-induced formation of fluorescent R123 and DCF: (A) formation of fluorescent R123 with Cr(V)-EHBA in the absence (9) or presence (b) of oxygen and (B) formation of fluorescent DCF with Cr(V)-EHBA in the absence (9) or presence (b) of oxygen. cell volume obtained by Shellard et al. for A549 cells (36).

Results and Discussion Concentration and Oxygen Dependence of Cr(V) Fluorescence Induction. Studies were carried out using the Cr(V)-EHBA complex as a model for the highvalence chromium observed upon intracellular reduction of Cr(VI). This Cr(V) complex has well-characterized aqueous chemistry (37), is able to directly oxidize DNA (18), and is considered to resemble the Cr(V)-ascorbate complex formed during the reduction of Cr(V) by ascorbate (vitamin C) (38, 39). Reaction of 0.1-1000 µM Cr(V)-EHBA with DHR and DCFH at pH 7.0 generated the fluorescent species R123 and DCF, respectively, in a concentration-dependent manner (panels A and B of Figure 1). The fluorescence produced in this reaction had a broad dynamic range of measurement with a lower detection limit of less than 100 nM and an upper detection range of greater than 1 mM (range of 600 nM to 100 µM shown). Neither Cr(VI), Cr(III), nor an aged solution of Cr(V)-EHBA [in which all of the Cr(V) is converted to Cr(VI) and Cr(III)EHBA through disproportionation] induced fluorescence. The impact of molecular oxygen was determined by carrying out the fluorescence assay in an identical manner but in a rigorously argon-degassed glovebag. The reaction of Cr(V)-EHBA with both dyes (panels A and B of Figure 1) in the absence of molecular oxygen was identical to that in air. This lack of an oxygen dependence for dye oxidation by Cr(V)-EHBA was consistent with that seen previously with EPR spin traps (40) and indicates that activation of molecular oxygen is not involved in the mechanism of oxidation.

Chromium(V) Oxidation of Fluorescent Dyes

Chem. Res. Toxicol., Vol. 11, No. 12, 1998 1405

Scheme 1

Figure 2. In vitro (A) R123 or (B) DCF fluorescence induced by 10 µM Cr(V)-EHBA in 20 mM chelexed phosphate buffer: as is (9) or in 40 mM Tris buffer ([), 5 mM DMTU (0), 5 mM mannitol (2), 5 mM EHBA (b), 5 mM DMSO (4), or 5 mM ethanol (O).

The mechanism for the formation of the fluorescent dye species from a direct oxidation reaction with Cr(V) is shown in Scheme 1 with DCFH. A sequential hydrogen atom abstraction by Cr(V) to form the radical DCF• followed by a further electron abstraction and rearrangement gives the resulting fluorescent dye species DCF. This mechanism is similar to that proposed previously for ROS (20, 30) and is consistent with the known hydrogen atom abstraction chemistry for Cr(V) (16-18). Induction of DCF and R123 Fluorescence by Cr(V)-EHBA in the Presence of Radical Scavengers. We examined the effect of a number of radical scavengers that have been used previously to trap radicals in this fluorescent dye system. We have also examined the effects of excess 2-ethyl-2-hydroxybutyric acid (EHBA), which chelates Cr(V) to stabilize it against disproportionation (37), and Tris buffer as it is a radical scavenger (20) and a weak Cr(V) chelator (20, 38, 41). Of the radical scavengers used, none substantially affected R123 fluorescence induced by 10 µM Cr(V)-EHBA, although a minor effect was observed for 5 mM DMTU and 40 mM Tris buffer (Figure 2A). DCF fluorescence induced by 10 µM Cr(V)-EHBA was affected by some of the radical

scavengers, with 5 mM DMTU displaying the least effect, followed by 5 mM DMSO and 5 mM EtOH (Figure 2B). EtOH at 1 mM did not affect DCF fluorescence (data not shown). Mannitol significantly affected Cr(V)-EHBAinduced DCF fluorescence, and the addition of Tris buffer led to a high initial value for fluorescence (time point t ) 0 min is actually at 1-2 min due to experimental time constraints) which plateaued more rapidly and at a lower value. For both dyes, the addition of excess EHBA led to significantly higher final fluorescence values (off the scale for both dyes; not shown for DCF), and lower fluorescence for DCF was exhibited at earlier time points. This result suggests a differential reactivity of the two dyes toward the Cr(V)-EHBA complex in the presence of radical scavengers. Final fluorescence results are higher that those for equimolar reactions in Figure 1 due to the increased well volume required for rapid and accurate addition of the different solutions. EPR Stability of Cr(V)-EHBA in the Presence of Selected Radical Quenchers. The stability of 10 µM Cr(V) in the presence of the radical scavengers or chelators was monitored by EPR to aid interpretation of the fluorescence results. The decay rate of an EPR signal of 10 µM Cr(V)-EHBA was almost identical in the presence or absence of 5 mM EtOH or DMSO (Figure 3A). However, we observed a reduction in DCF fluorescence, but not in R123 fluorescence, in the presence of radical quenchers at these concentrations, with a lesser effect at 1 mM DMSO and no effect at 1 mM EtOH (data not shown). While this result reflects those obtained by LeBel et al. (20), in this case we could determine by EPR

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Figure 4. DCF (O) or R123 (0) fluorescence in chromatetreated monolayers of A549 cells compared with toxicity (b) assessed by clonogenic assay.

Figure 3. (A) Stability of 10 µM Cr(V)-EHBA in 20 mM phosphate buffer (pH 7.4) measured by EPR at room temperature: as is (9) or in 40 mM Tris buffer ([), 5 mM DMTU (0), 5 mM mannitol (2), 5 mM EHBA (b), 5 mM DMSO (4), or 5 mM ethanol (O). (B) EPR signal of a quasistable Cr(V)-EHBA or Cr(V)-bis-mannitol species.

that the concentration of the DCFH-reactive species, namely Cr(V), was not affected by the presence of these solvents. Of the radical quenchers and chelators used in this study, EtOH and DMSO were water-miscible organic solvents, and the remainder aqueous salt solutions. EtOH and DMSO did not affect the stability of Cr(V) in 20 mM phosphate buffer, and the decrease in fluorescence was dependent upon their concentration, which suggested a solvent effect for these two radical scavengers. A likely explanation is that the xanthene dyes, such as fluorescein and rhodamine, can be converted from the colored zwitterionic (non-lactone) form of the dye to a colorless inner lactone at the carboxylate moiety in the presence of non-hydrogen-bonding donor solvents, of which DMSO is a specific example (42). Ethanol also stabilizes the colored, zwitterionic form to a lesser extent than water alone. To support this hypothesis, we also observed a pH-dependent decrease in DCF fluorescence below pH 7.0 (not shown). There was no analogous decrease below pH 7 in a previous study in which the pH dependence on DHR fluorescence was examined (30). Unlike DCF, the carboxylate moiety of the rhodamine dye used in this study, dihydrorhodamine, is esterified and is therefore stable against formation of this colorless inner lactone. DMTU (5 mM) slightly destabilized Cr(V)-EHBA, although this effect was not substantial in either dye. This was consistent with the finding of Ueno et al. (32),

who reported that the intensity of a Cr(V) EPR signal, derived from in situ reduction of Cr(VI) by glutathione, was unaffected by the presence of 5 mM DMTU both in vitro and in cellular systems. Tris buffer (40 mM) significantly destabilized the EPR signal of the Cr(V)EHBA complex which was consistent with the observed DCF fluorescence (Figure 2B). As expected, 5 mM EHBA substantially stabilized the Cr(V) EPR signal against decay by disproportionation. This increased lifetime of Cr(V) in solution with excess EHBA explained the offscale fluorescence results for both dyes and suggests that fluorescence intensity is a qualitative and not quantitative measure of Cr(V) levels. EPR studies also demonstrated that 5 mM mannitol significantly stabilized Cr(V) in solution. As shown in Figure 3B, mannitol exchanged for the EHBA ligand coordinated to the metal center to give a new EPR signal. The signal showed a characteristic 1:4:6:4:1 hyperfine splitting pattern, with an AH of 1.1 G and a g of 1.982. The chromium hyperfine splitting with an ACr of 17.0 G indicated that the axial site is likely to be unoccupied (37). The hyperfine splitting pattern was accordingly attributed to the protons of two mannitol ligands, and the new Cr(V) species was assigned as a bismannitol species chelated to the Cr(V) metal center through the internal diols. As the intensity of R123 fluorescence was relatively unaffected by mannitol, it was clear that the nonfluorescent form of this dye can react with the Cr(V)-mannitol complex whereas DCFH was less reactive toward the new Cr(V) species. These results imply that there is a differential reactivity between the two dyes which is sensitive to the structure of the Cr(V) complexes formed in the reaction. Chromate-Induced Fluorescence and Toxicity in Human Lung A549 Cells. Adherent monolayers of A549 cells in culture were pretreated with DCFH-DA or DHR and exposed to different doses of Cr(VI) for 4 h. A dose dependence was observed for both toxicity and dye fluorescence (Figure 4). Significant DCF and R123 fluorescence was observed at relatively nontoxic doses of Cr(VI), namely 5 and 10 µM. These results demonstrated a strong correlation between Cr(VI) toxicity and fluorescence of DCF and R123. In cells, R123 fluorescence was approximately the same, or even slightly higher than that of DCF, in contrast to the in vitro results. Cr(V)chelating species analogous to mannitol and EHBA undoubtedly exist within the cell, and the effects these endogenous chelators may have with regard to DCFH and DHR oxidation are not known.

Chromium(V) Oxidation of Fluorescent Dyes

Chem. Res. Toxicol., Vol. 11, No. 12, 1998 1407 Table 2. Effect of 5 mM EtOH or 5 mM DMTU on Hydrogen Peroxide-Induced DCF Fluorescence in Postconfluent Monolayers of A549 Cells at 90 min [H2O2] (µM)

with 5 mM EtOHa

with 5 mM DMTUa

0 50 100 200 350 500

141 ( 17% 119 ( 20% 122 ( 12% 137 ( 11% 163 ( 8% 142.5 ( 6.7%

81 ( 6.4% 107 ( 20% 171 ( 43% 218 ( 24% 472 ( 14% 552 ( 41%

a

Figure 5. DCF (O) fluorescence in hydrogen peroxide-treated monolayers of A549 cells compared with toxicity (b) assessed by clonogenic assay. Table 1. Effect of 5 mM EtOH or 5 mM DMTU on Cr(VI)-Induced DCF and R123 Fluorescence in Postconfluent Monolayers of A549 Cells at 240 min (4 h) [Cr(VI)] (µM) 0 5 10 20 40 80

with 5 mM EtOHa R123

DCF

105 ( 11.4% 114 ( 10.4% 97 ( 12% 98.25 ( 2% 95.7 ( 5.4% 102 ( 5% 113 ( 9.8% 97.25 ( 4% 101.3 ( 5.4% 98.5 ( 2.5% 100.6 ( 5% 96.5 ( 6%

Table 3. Rate of Increase of Fluorescence Induced by Cr(VI) in Postconfluent Monolayers of A549 Cells Compared with a Calculation of the Extent of Cr(V) Formation Corresponding to Fluorescence Based upon Fluorescence Numbers Obtained for Cr(V)-EHBA R123a

R123

fluorescence per hour (%)

Cr(V) per hour (µM)

fluorescence per hour (%)

Cr(V) per hour (µM)

5 10 20 40 80

93 ( 9 98.5 ( 7.6 207.5 ( 16 341 ( 35 494 ( 40

0.167 0.178 0.373 0.612 0.886

114 ( 4 125 ( 5 217.7 ( 8.1 296 ( 11 506.5 ( 28.5

0.035 0.046 0.131 0.202 0.395

DCF

a Measurements are the mean ( SD of two experiments of 10 points each (20 replications total).

Hydrogen Peroxide-Induced Fluorescence and Toxicity in Human Lung A549 Cells. Hydrogen peroxide exposure to adherent monolayers of A549 cells was monitored for toxicity and dye fluorescence in a manner similar to that of Cr(VI). A strong dose-dependent increase in fluorescence was observed, which plateaued upon reaching 100% toxicity, in a manner similar to that for Cr(VI)-induced fluorescence (Figure 5). Fluorescence measurements were taken after 90 min rather than after 4 h, as for Cr(VI), as dye fluorescence in the hydrogen peroxide-treated system decreased after longer periods of time. This loss in fluorescence after longer periods of time was considered to arise from degradation of the fluorescent form of the dye by excess reactive oxygen species, which we have also observed in the in vitro horseradish peroxidase-hydrogen peroxide system (unpublished data). Effect of Radical Scavengers on Hydrogen Peroxide versus Cr(V) Fluorescence. To distinguish Cr(V)-induced fluorescence from that induced by reactive oxygen species (ROS), we repeated the experiment with prior incubation of the A549 cells with the radical scavengers 5 mM DMTU and 5 mM EtOH, which were shown to have little effect on the stability of Cr(V) at physiological pH. As seen in Table 1, neither radical scavenger had a significant effect on Cr(VI)-induced fluorescence in this cell line, although a slight decrease in R123 fluorescence was observed in cells pretreated with 5 mM DMTU. There was little difference between the results for a 1 versus 5 mM EtOH preincubation (not shown). As DCF fluorescence has been used as a measure of the extent of intracellular hydrogen peroxide formation, we examined the effect of radical scavengers on hydrogen peroxide-induced DCF fluorescence. As shown in Table

DCFa

[Cr(VI)] (µM)

with 5 mM DMTUa 82 ( 12% 101 ( 4.6% 83 ( 12% 95 ( 16% 80 ( 4.8% 103 ( 12% 96 ( 7.3% 97.5 ( 14% 91 ( 7.5% 94.5 ( 1.5% 88 ( 5.7% 93 ( 12%

Measurements are the mean ( SD of 10 replications.

a

Measurements are the mean ( SD of 10 replications.

2, DMTU markedly enhanced DCF fluorescence. We attributed this increase in fluorescence to the ability of the radical scavengers to trap the hydroxyl radical, thereby forming a more stable species with a longer lifetime in the cell and ipso facto a higher probability that it can react with the dye. An alternative mechanism was that, by trapping the radical, the radical scavenger ameliorated the destruction of the fluorescent form of the dye by excess free hydroxyl radical. This alternative mechanism may account for the DMTU studies as fluorescence continued to increase after 90 min rather than decreasing as it had in non-DMTU-treated systems. The effect for EtOH was, however, much less marked. An increase in fluorescence was observed in the presence of 5 mM EtOH, although of a lesser magnitude. This result was somewhat unexpected as EtOH has been used as a hydroxyl radical scavenger in EPR spin-trapping studies where an R-alcohol radical species is formed. With regard to the effect of radical scavengers on the intracellular oxidation of DCFH and DHR, it is clear that hydrogen peroxide and Cr(VI) induced intracellular oxidative bursts of a qualitatively different nature. Intracellular Chromium Uptake versus Fluorescence. Hydrogen peroxide fluorescence of the two dyes exhibited a decrease after reaching a maximum at 90 min, while Cr(VI)-induced fluorescence was approximately linear for all concentrations used in this system. The rate of increase in fluorescence is listed in Table 3 for all concentrations. As shown in Figure 6 (panels A and B), a time-dependent increase in fluorescence following exposure to 10 µM Cr(VI) corresponded to the linear uptake of chromium from the media as measured by atomic absorption spectroscopy. For a 10 µM dose of Cr(VI), uptake of chromium occurred at ∼200 µM/h as chromium was concentrated from the whole media into the much smaller total cell volume of the cell monolayer. The rate of uptake was dose-dependent at 4 h with a plateau observed at the 40 µM Cr(VI) dose and above (Figure 6C). This was attributed to the toxicity of these doses at 4 h (Figure 4). Quantitative measurement of

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Figure 7. Cr(V) signal detected in A549 cells treated with 50 or 200 µM Cr(VI) and incubated at 37 °C for 25 min followed by flash-freezing and EPR measurement at 77 K.

Figure 6. DCF (b) or R123 (0) fluorescence in whole cells vs time in A549 cells treated with 10 µM chromate (A) compared with the extent of chromium uptake into A549 cells treated with 10 µM chromate (B). The extent of uptake at 4 h is linearly dependent on dose until chromium becomes toxic to the cells (C).

the intracellular concentrations of Cr(V) by fluorescence is impossible as Cr(VI) was actively reduced intracellularly in a manner other than by the disproportionation decay kinetics of the model Cr(V)-EHBA complex shown in Figures 1-3. In addition, chelating species analogous to mannitol and EHBA undoubtedly exist within the cell. These chelators may stabilize the Cr(V) metal center against disproportionation as well as provide competition for DCFH as seen for mannitol. Nevertheless, the basic assumption that the Cr(V) concentration can be represented by fluorescence was used to tabulate Cr(V) levels which would correspond to the fluorescence observed in Figure 1. These numbers are given in Table 3. As expected, the values were significantly lower than the levels obtained for total chromium uptake by atomic absorption spectroscopy. This was almost certainly due to the strong reducing environment within the cell where reduction of intracellular Cr(V) to Cr(III) would compete with dye oxidation. Intracellular Formation of Cr(V) in A549 Cells. Whole cell frozen EPR spectroscopy was used to detect a Cr(V) signal of approximately 1.4 µM for 11.68 million

cells treated with 50 µM Cr(VI) and 1.65 µM for 5.41 million cells treated with 200 µM Cr(VI) (Figure 7). The g value of 1.987 for this Cr(V) signal was identical in each sample. This signal was the same as that for a chromium-glutathione complex formed in vitro (43-45), indicating the potential importance of glutathione in reducing Cr(VI) within the cell. Cr(V) signals were also detected in cells which were treated with 50 µM to 5 mM Cr(VI) as adherent monolayers and subsequently collected by scraping or trypsinization after 20 min, counted, and frozen for EPR measurements. A frozen Cr(V) signal represents a time slice in the steady state formation and decay of Cr(V), and the values of 1-2 µM were not unrealistic for the levels of uptake. When corrected for the smaller total volume of cells using a cell volume of 3.56 pL/cell as calculated by Shellard et al. (36) for A549 cells, this Cr(V) concentration increases approximately 10-fold. These results, coupled with the lack of an effect of radical scavengers compared to that observed for hydrogen peroxide, suggested that Cr(V) is indeed produced at intracellular levels which would account for much, if not all, of the observed fluorescence in Cr(VI)treated cells.

Conclusions It is clear from the results presented in Figure 1 that both intracellular dyes studied are sensitive measures of the extent of Cr(V) formation and that fluorescence does not require the formation of ROS. In cultured A549 cells, we observed a dose-dependent increase in fluorescence with Cr(VI), in agreement with previous studies on Cr(VI)-treated cells in culture (25-27). However, the use of radical scavengers demonstrated that the intracellular oxidation induced by Cr(VI) was qualitatively different from that induced by hydrogen peroxide. The inability of EtOH and DMTU to significantly affect Cr(VI)-induced fluorescence in A549 cells suggests that the observed fluorescence was due to intracellular formation of Cr(V) and not ROS as previously believed. This nonROS pathway of cellular oxidative stress is consistent with the work of Dubrovskaya and Wetterhahn (46), who determined that none of seven ROS-inducible genes, including catalase and the Cu- and Mn-superoxide dismutases, were upregulated in response to Cr(VI) treatment in A549 cells or in a normal human lung fibroblast line.

Chromium(V) Oxidation of Fluorescent Dyes

The intracellular level of Cr(VI)-induced fluorescence was lower than that expected on the basis of cellular chromium uptake and the intensity of the Cr(V) EPR signal. The g value of the intracellular Cr(V) signal indicates that it may be a chromium-glutathione complex which would decay to Cr(III) at a rate that competes with the reaction between Cr(V) and DCFH or DHR. Such competition for the dye-reactive species by free sulfhydryls has been observed for DHR (30). However, the extents of uptake and fluorescence both display a linear dependence upon time and Cr(VI) dose. Thus, DCF and R123 fluorescence are qualitative and cumulative measures of the extent of intracellular Cr(V) formation. To our knowledge, this is the first study in which a non-ROS-mediated mechanism for dye fluorescence has been unequivocally established. However, the suggestion that hypervalent metals may be responsible for DCF fluorescence is not a novel one. Detailed studies directed toward defining the ROS responsible for DCFH and DHR fluorescence led to the conclusion that a ferryl species (hypervalent iron) was responsible for the observed fluorescence (20, 47, 48). As neither DCFH nor DHR fluoresces in the presence of hydrogen peroxide alone, but requires a Fenton metal or a metalloenzyme (20, 30, 47), a catalytic metal center is an important component of oxidant-sensitive dye fluorescence. The classification of DCFH and DHR as hydrogen peroxide-sensitive probes is misleading in the context of Cr(VI)-treated cell systems. The conversion of DCFH and DHR to their fluorescent forms is, however, an oxidation reaction, and the intracellular fluorescence of these dyes in cultured cells is a clear indicator of intracellular oxidation. However, the nature of the oxidant needs to be examined when dealing with chromium, as hypervalent chromium is capable of performing many of the oxidation reactions usually ascribed to reactive oxygen. We wish with this study to alert the biological chromium community to the dose-dependent reactivity between Cr(V) and the oxidant-sensitive dyes, 2′,7′-dichlorofluorescin and dihydrorhodamine. It is our hope that this study will assist in the interpretation of intracellular fluorescence studies in chromate-treated biological systems.

Acknowledgment. This research was funded by U.S. Public Health Service Grant ES07167 from the National Institute of Environmental Health Sciences, DHHS, and NIH Grant CA45735 which were awarded to Karen E. Wetterhahn. J.A.S. was a Dartmouth Undergraduate Presidential Scholar in the Department of Chemistry and also acknowledges the NSF Research Experience for Undergraduates program, (CHE-9322017 provided to Dartmouth College). The EPR spectrometer was purchased with NSF Grant CHE-8701406. This paper is dedicated to the memory of Karen Wetterhahn and her infectious enthusiasm for science.

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