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TOXICOLOGY

Roles of Necrosis, Apoptosis, and Mitochondrial Dysfunction in S-(1,2-Dichlorovinyl)-L-cysteine Sulfoxide-Induced Cytotoxicity in Primary Cultures of Human Renal Proximal Tubular Cells

Department of Pharmacology, Wayne State University School of Medicine, Detroit, Michigan (L.H.L., D.A.P., S.E.H.) and Department of Comparative Biosciences, University of Wisconsin School of Veterinary Medicine, Madison, Wisconsin (R.J.K., A.A.E.)

Received October 25, 2002; accepted February 21, 2003.

	Abstract

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S-(1,2-Dichlorovinyl)-L-cysteine (DCVC) is the penultimate nephrotoxic metabolite of the environmental contaminant trichloroethylene. Although metabolism of DCVC by the cysteine conjugate -lyase is the most studied bioactivation pathway, DCVC may also be metabolized by the flavin-containing monooxygenase (FMO) to yield DCVC sulfoxide (DCVCS). Renal cellular injury induced by DCVCS was investigated in primary cultures of human proximal tubular (hPT) cells by assessment of time- and concentration-dependent effects on cellular morphology, acute cellular necrosis, apoptosis, mitochondrial function, and cellular glutathione (GSH) status. Confluent hPT cells incubated with as little as 10 µM DCVCS for 24 h exhibited morphological changes, although at least 100 µM DCVCS was required to produce marked changes. Acute cellular necrosis did not occur until 48 h with at least 200 µM DCVCS, indicating that this is a high-dose, late response. The extent of necrosis was similar to that with DCVC. In contrast, apoptosis occurred as early as 1 h with as little as 10 µM DCVCS and the extent of apoptosis was much less than that with DCVC. Mitochondrial function was maintained with DCVCS concentrations up to 100 µM, consistent with hPT cells only being competent to undergo apoptosis at early time points and relatively low concentrations. Marked depletion (>50%) of cellular GSH content was only observed with 500 µM DCVCS. These results, combined with previous studies showing protection from DCVC-induced necrosis and apoptosis by the FMO inhibitor methimazole, suggest that formation of DCVCS plays a significant role in trichloroethylene-induced renal cellular injury in hPT cells.

Most TRI toxicity is associated with its metabolism, which occurs by either cytochrome P450-dependent oxidation or glutathione (GSH) conjugation (Lash et al., 2000a). One of the established target organs for TRI is the kidneys, and renal toxicity is associated with metabolism by the GSH conjugation pathway (Lash et al., 2000b). S-(1,2-Dichlorovinyl)-L-cysteine (DCVC), the cysteine conjugate of TRI, was initially considered to be the primary, if not sole, penultimate nephrotoxic metabolite that produces renal cellular injury after bioactivation by the cysteine conjugate -lyase (-lyase). However, DCVC can also be metabolized by the flavin-containing monooxygenase (FMO) to produce DCVC sulfoxide (DCVCS) (Sausen and Elfarra, 1990; Ripp et al., 1997; Krause et al., 2003), which is highly reactive and is a potent nephrotoxicant in rats and is cytotoxic in isolated rat proximal tubular (rPT) cells (Lash et al., 1994).

Isolated rPT cells are susceptible to DCVC-induced necrosis at relatively high doses (i.e., >0.2 mM) (Cummings et al., 2000a,b; Lash and Anders, 1986; Lash et al., 1995b, 2001a,2001b). Primary cultures of rPT cells (van de Water et al., 1996, 1999) and LLC-PK₁ cells (Chen et al., 2001) also undergo apoptosis when exposed to DCVC. The initial step in the GSH conjugation pathway for TRI occurs at comparable rates in both rodent (Lash et al., 1995b, 1998) and human (Lash et al., 1999) kidney and liver. -Lyase activity, however, is much higher in rat kidney than in human kidney (Lash et al., 1990). Although this might be interpreted as meaning that bioactivation of DCVC is much greater in rat kidneys than in human kidneys and that rats are much more susceptible to DCVC-induced renal injury than humans, the role of FMO-dependent bioactivation and toxicity in human kidney has not been previously considered.

Freshly isolated human proximal tubular (hPT) cells undergo necrosis when incubated with relatively high concentrations (>200 µM) of DCVC (Cummings and Lash, 2000) but primary cultures of hPT cells can also undergo apoptosis at relatively low concentrations of DCVC (i.e., 100 µM) and at relatively early incubation times (i.e., 8 h). Comparison of the potency of DCVC and DCVCS in rats shows that the sulfoxide is the more potent nephrotoxicant both in vivo and in vitro. The potency of DCVCS raises the question of the role of this reactive species in the nephrotoxicity of TRI and DCVC. Although studies with selective inhibitors of both the -lyase and FMO suggested that the -lyase plays a more prominent role in DCVC-induced cytotoxicity in rPT cells (Lash et al., 1994), FMO seems to play the more prominent role in bioactivation in hPT cells (Cummings and Lash, 2000; Lash et al., 2001a). This apparent discrepancy highlights the difficulty of extrapolating data from rodents to humans and is a major issue in human health risk assessment for chemicals such as TRI (Maull and Lash, 1998). Furthermore, this suggests that FMO-dependent bioactivation may account for some of the species-dependent differences in the nephrotoxicity of TRI and DCVC. We recently showed that human kidney expresses multiple isoforms of FMO (Krause et al., 2003), supporting the potential for this pathway to function in hPT cells.

In the present study, the importance of the FMO-dependent bioactivation pathway in human kidney was investigated further by studying mechanisms of renal cellular injury in primary cultures of hPT cells induced by DCVCS. The results show that DCVCS is a potent cytotoxicant in hPT cells, inducing acute cellular necrosis and apoptosis. Although the pattern of cellular injury in hPT cells induced by DCVCS and DCVC differs somewhat, the results support the conclusion that FMO-dependent bioactivation of DCVC, along with the -lyase, plays an important role in TRI- and DCVC-induced renal cellular injury in the human kidney.

	Materials and Methods

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Chemicals. TRI (reported to be 98.6% pure, as judged by electron impact ionization mass spectrometry), collagenase type I (EC 3.4.24.3 [EC] ), ascorbate, TMPD, and bovine serum albumin (fraction V) were purchased from Sigma-Aldrich (St. Louis, MO). Annexin V-fluorescein isothiocyanate (FITC) was purchased from BD PharMingen (San Diego, CA). DCVC was first synthesized as described previously (Elfarra et al., 1986) by reacting TRI with L-cysteine in sodium and liquid ammonia. DCVCS was then made by reacting DCVC with H₂O₂, as described previously (Ripp et al., 1997). Purity (>95%) was determined by high-pressure liquid chromatography (HPLC) analysis, and identity was confirmed by proton nuclear magnetic resonance spectroscopy and mass spectrometry. Impurities were likely degradation products of TRI and cysteine and are unlikely to contribute significantly to the observed cytotoxic responses. Because it is very laborious and impractical to try and improve purity of the DCVCS, no additional purification was attempted.

Isolation of hPT Cells from Human Kidneys. hPT cells were derived from human kidney cortical slices obtained from the Human Tissue Resources Core of Wayne State University (Department of Pathology, Harper Hospital, Detroit, MI) or from whole human kidneys procured by International Bioresearch Solutions (Pasadena, CA). Kidneys or kidney slices were obtained from a total of 17 donors, with the following characteristics: seven female (mean age ± S.E., 56.9 ± 3.5; age range, 40–68; six Caucasian, one Hispanic); 10 male (mean age ± S.E., 54.9 ± 6.6; age range, 16–75; six Caucasian, two African American, one Hispanic, one Pacific Islander). All tissue was scored by a pathologist as normal (i.e., derived from noncancerous, nondiseased tissue). Cell isolation procedures are based on those originally described by Todd et al. (1995) and modified (Cummings and Lash, 2000; Cummings et al., 2000c). Whole kidneys were perfused with Wisconsin medium and were kept on wet ice until they arrived at the laboratory, which was usually within 24 h of removal from the donor. All buffers were continuously bubbled with 95% O₂, 5% CO₂ and were maintained at 37°C. Tissue slices were washed with sterile phosphate-buffered saline (PBS), minced, and the pieces were placed in a trypsinization flask filled with 40 or 300 ml (for slices from the Tissue Resources Core or slices from the whole kidneys, respectively) of sterile, filtered Hanks' buffer, containing 25 mM NaHCO₃, 25 mM HEPES, pH 7.4, 0.5 mM EGTA, 0.2% (w/v) bovine serum albumin, 50 µg/ml gentamicin, 1.3 mg/ml collagenase, and 0.59 mg/ml CaCl₂, which was filtered before use. Minced cortical pieces from either slices or whole kidneys were subjected to collagenase digestion for 60 min, after which the supernatant was filtered through a 105-µm mesh filter to remove tissue fragments, centrifuged at 150g for 7 min, and the pellet resuspended in Dulbecco's modified Eagle's medium/Ham's F-12 medium (DMEM/F-12; 1:1). Approximately 5 to 7 x 10⁶ cells were obtained per 1 g of human kidney cortical tissue.

Culturing of hPT Cells. Isolation of hPT cells was achieved as explained above with use of sterile conditions (i.e., all instruments and glassware were autoclaved and all buffers were filtered through a 0.2-µm pore-size filter). After isolation, cells were resuspended in 2 ml of DMEM/F-12 and diluted to 500 ml with cell culture media. Basal medium was a 1:1 mixture of DMEM/F-12. Standard supplementation included 15 mM HEPES, pH 7.4, 20 mM NaHCO₃, anti-biotics for day 0 through day 3 only (192 IU of penicillin G/ml + 200 µg of streptomycin sulfate/ml or 50 µg of gentamicin/ml) to inhibit bacterial growth, 2.5 µg of amphotericin B/ml to inhibit fungal growth, 5 µg of bovine insulin/ml (0.87 µM), 5 µg of human transferrin/ml (66 nM), 30 nM sodium selenite, 100 ng of hydrocortisone/ml (0.28 µM), 100 ng of epidermal growth factor/ml (17 nM), and 7.5 pg of 3,3',5-triiodo-DL-thyronine/ml (111 nM) (Lash et al., 1995a). Cells were seeded at densities of 50 to 100 µg of protein/cm² (0.5–1.0 x 10⁶ cells/ml) on either collagen-coated polystyrene culture dishes or collagen-coated polystyrene tissue culture flasks (T-25, T-75, or T-175). Cultures were grown at 37°C in a humidified incubator under an atmosphere of 95% air, 5% CO₂ at pH 7.4. Cultures were grown to confluence (generally 5 to 7 days) before treatment with any agent. Cells were harvested from the dishes by either scraping the plates with a Teflon scraper or by brief incubation with Cellstripper (Cellgro, Herndon, VA) (in Ca²⁺- and Mg²⁺-free Hanks' buffer).

Protein determination was done using the bicinchoninic acid protein determination kit (Sigma-Aldrich), using bovine serum albumin as a standard.

Confocal Microscopy. hPT cells were grown on collagen-coated, 35-mm culture dishes and were viewed with a triple-laser scanning confocal microscope (LSM 310; Carl Zeiss, Thornwood, NY) with integrated workstation at the Confocal Imaging Core Facility in the School of Medicine at Wayne State University (Detroit, MI). This is a core facility of the National Institute of Environmental Health Sciences Center for Molecular Toxicology with Human Application at Wayne State. Initial magnification was 196x.

Measurement of Necrosis by Lactate Dehydrogenase (LDH) Release. hPT cells were incubated with either medium or medium containing the indicated concentrations of DCVCS for up to 48 h. Cell viability at the conclusion of these incubations was estimated by determining the release of LDH from cells after various incubations and at various times (Lash et al., 1995a). LDH release from cells was measured by determining LDH activity (measured spectrophotometrically as NADH oxidation at 340 nm) in media and, after removal of media, washing cells with PBS, and solubilization of cells with 0.1% (v/v) Triton X-100, in total cells. The fraction of LDH release was an index of irreversible injury or necrosis: %LDH release = LDH activity in media/(LDH activity in media + LDH activity in total cells) x 100%.

Flow Cytometry Analysis of Cell Cycle. Cell cultures were washed twice with sample buffer [PBS plus glucose (1 g/l) filtered through a 0.22-µm filter], dislodged by trypsin/EDTA (0.1% w/v) incubation, centrifuged at 400g for 10 min, and resuspended in sample buffer. Cell concentrations were adjusted to 1 to 3 x 10⁶ cells/ml with sample buffer and 1 ml of the cell suspension was centrifuged at 400g for 10 min. All of the supernatant except 0.1 ml/10⁶ cells was removed and the remaining cells were mixed on a vortex mixer in the remaining fluid for 10 s. Next, 1 ml of ice-cold ethanol (70%, v/v) was added to the sample drop by drop, with samples being mixed for 10 s between drops. The tubes were capped and fixed in ethanol at 4°C. After fixing, the cells were stained in propidium iodide (50 µg/ml) containing RNase A (100 U/ml). Samples were then mixed, centrifuged at 1000g for 5 min and all the ethanol except 0.1 ml was removed. Cells were mixed in the residual ethanol and 1 ml of the propidium iodide staining solution was added to each tube. After mixing again, cells were incubated at room temperature for at least 30 min. Samples were analyzed within 24 h by flow cytometry using an FACSCalibur flow cytometer (BD Biosciences, San Jose, CA), which is a core facility of the National Institute of Environmental Health Sciences Center for Molecular Toxicology with Human Application at Wayne State. Analysis was performed with 20,000 events/sample using the ModFit LT v. 2 for Macintosh data acquisition software package (Verity Software House, Inc., Topsham, ME; distributed by BD Immunocytometry System BDIS). Propidium iodide was detected by the FL-2 photomultiplier tube. Fractions of apoptotic cells were quantified by analysis of the subG₁ (subdiploid) peak with ModFit cell cycle analysis. The percent distribution of cells in the various stages of the cell cycle (G₀/G₁, S, and G₂/M) were also calculated. Cell aggregates were discarded in the flow cytometry analysis by post-fixation aggregate discrimination.

Annexin V-FITC Binding Assay. Induction of apoptosis was also analyzed by annexin V-FITC binding and flow cytometry. hPT cells grown in T-25 flasks were washed with 3 ml of ice-cold PBS. Ice-cold binding buffer (1.5 ml; 10 mM HEPES, 140 mM NaCl, and 2.5 mM CaCl₂, pH 7.4) was added to each flask, to which was then added 5 µl of annexin V-FITC. After a 15-min incubation with shaking in the dark at room temperature, the annexin was removed, and 1 ml of ice-cold binding buffer was added. Cells were gently scraped from the flasks, 1 ml of cells was placed in the flow cytometry tubes, 18 µl of a 20x stock solution of propidium iodide (50 µg/ml in water, final concentration) was added, and tubes were placed on ice in the dark for at most 1 h until analysis on an FACSCalibur flow cytometer (BD Biosciences). Control samples included unstained cells, cells stained with annexin V-FITC but no propidium iodide, and cells stained with propidium iodide but no annexin V-FITC.

Measurement of Mitochondrial Membrane Potential () with JC-1. Confluent, primary cultures of hPT cells were grown on collagen-coated, glass coverslips in 35-mm tissue culture dishes and were incubated for 2 or 4 h with 0 (control), 10, or 50 µM DCVCS. JC-1 fluorescence was measured by confocal microscopy assessing the emission shift from green (530 nm) to red (590 nm) in polarized mitochondria using 488-nm excitation. Polarized mitochondria are indicated by yellow-red punctate staining.

Measurement of Cellular Respiration. Confluent, primary cultures of hPT cells were grown on collagen-coated T-25 flasks. Cells were incubated for various times up to 48 h with either PBS or the indicated concentration of DCVCS. At the end of the incubation period, cells were released from the culture surface by brief trypsinization and resuspended in PBS at a concentration of 2 x 10⁶ cells/ml. Succinate-, (glutamate + malate)-, or (ascorbate + TMPD)-stimulated O₂ consumption was measured in a 5/6H oxygraph (Gilson Medical Electronics, Middleton, WI), as described previously (Lash and Anders, 1986; Lash and Tokarz, 1989).

Measurement of Cellular ATP Concentrations. Intracellular contents of adenine nucleotides were measured in neutralized perchloric acid extracts by the HPLC method of Jones (1981), as described previously (Lash and Tokarz, 1989). Separation of ATP, ADP, and AMP was achieved by reversed-phase HPLC with a µBondapak C₁₈ cartridge (8 mm x 10 cm) (Waters, Milford, MA) and detection was by absorbance at 260 nm.

Measurement of Cellular GSH Concentrations. Cellular content of GSH was determined by ion-exchange HPLC on a Waters µBondapak amine cartridge (8 mm x 10 cm) (Waters, Milford, MA) after derivatization of thiols with iodoacetate and amine groups with 1-fluoro-2,4-dinitrobenzene (Fariss and Reed, 1987; Visarius et al., 1996). Derivatives were detected by absorbance at 365 nm and were compared with authentic standards (limit of detection, 50 pmol).

Data Analysis. Measurements of apoptosis, necrosis, and cell proliferation were performed on at least three separate cell cultures, except where indicated. Results are expressed as means ± S.E., except where inclusion of error bars made the figures unclear. In those cases, the degree of variation is indicated in the legends. Significant differences for means were first assessed by a one- or two-way analysis of variance. When significant F values were obtained with the analysis of variance, the Fisher's protected least-significance t test was performed to determine which means were significantly different from one another, with two-tail probabilities <0.05 considered significant.

	Results

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Cellular Morphology. Incubation of confluent primary cultures of hPT cells for 24 h with a range of concentrations of DCVCS produced obvious cell injury (Fig. 1). Significant changes in cell shape (i.e., elongation and irregularity) and an increase in intracellular vesicularization were observed at a concentration of DCVCS as low as 10 µM. These changes progressed in severity with increasing concentrations of DCVCS, such that at a concentration of 500 µM, little recognizable cellular structure was evident.

View larger version (100K): [in this window] [in a new window]   Fig. 1. Morphology of primary cultures of hPT cells incubated with DCVCS. Confluent, primary cultures of hPT cells were incubated for 24 h with either media (control) or the indicated concentrations of DCVCS. Photomicrographs were obtained on a triple-laser scanning confocal microscope (Carl Zeiss) at an initial magnification of 196x. Scale bar, 5 µm.

Time and Concentration Dependence of LDH Release from hPT Cells. Confluent primary cultures of hPT cells were incubated with a wide range of concentrations of DCVCS, ranging from 10 to 500 µM, and release of LDH was determined at various times as a measurement of acute cellular necrosis (Fig. 2). Little effect of DCVCS on LDH release was observed at concentrations of 100 µM or less throughout the 48-h incubation. The lowest concentration of DCVCS and time point that showed a significant increase in LDH release was 200 µM at 48 h (approximately 35% LDH release compared with 10% for control cells), whereas the earliest significant increase in LDH release occurred after a 2-h incubation with 500 µM DCVCS (approximately 19% LDH release compared with 12% for control cells). A DCVCS concentration of at least 200 µM was thus required to elicit any significant increase in LDH release.

View larger version (16K): [in this window] [in a new window]   Fig. 2. DCVCS-induced LDH release in primary cultures of hPT cells. Cellular necrosis was measured as LDH release after incubation of confluent, primary cultures of hPT cells with 10 to 500 µM DCVCS for various times up to 48 h. Results are means of measurements from three cell cultures. S.E.M. values, which were omitted for clarity, were generally 5% to at most 20% of means. Significantly different (P < 0.05) from corresponding control: 1 h, none; 2 h, 500 µM; 4, 8, 24, and 48 h, 200 and 500 µM.

Time and Concentration Dependence of Apoptosis in hPT Cells. In contrast to cell swelling and the collapse of membrane permeability that is characteristic of acute cellular necrosis, cells may also undergo apoptosis. The process of apoptosis differs from that of cellular necrosis in several ways, such as a requirement for metabolic energy (i.e., ATP), formation of apoptotic bodies, nuclear condensation, and regulation by expression of specific proteins and/or signaling molecules. Confluent hPT cells were stained with propidium iodide and subjected to FACS analysis and flow cytometry to quantitate the proportion of cells that are subdiploid. DCVCS was shown to cause time- and concentration-dependent increases in the proportion of hPT cells undergoing apoptosis (Fig. 3). The general pattern was that the proportion of cells undergoing apoptosis increased up to a DCVCS concentration of 50 to 100 µM and increased with time up to 4 to 8 h; at higher concentrations and later incubation times, the proportion of cells undergoing apoptosis decreased. A concentration of DCVCS of as low as 10 µM induced a significant increase in the proportion of apoptotic cells as early as 1 h. Maximal levels of apoptotic cells occurred in the range of 4 to 8 h of incubation, with approximately 4 to 10% apoptotic cells compared with untreated hPT cells, which exhibited less than 1% apoptotic cells. The proportion of apoptotic cells tended to decrease after 24 or 48 h of incubation with concentrations of DCVCS of 100 µM. This response is likely due to cells undergoing necrosis rather than apoptosis when exposed to higher concentrations of toxicant and/or for longer incubation times, when cells are likely to be incompetent to undergo apoptosis.

View larger version (44K): [in this window] [in a new window]   Fig. 3. DCVCS-induced apoptosis in primary cultures of hPT cells as indicated by FACS analysis and flow cytometry. Confluent hPT cells were grown on 35-mm polystyrene culture dishes and treated for the indicated times with PBS (control) or DCVCS. Cells were then processed as described under Materials and Methods and analyzed within 24 h by flow cytometry with an FACSCalibur flow cytometer (BD Biosciences). Results are means ± S.E.M. of measurements from four separate cell cultures. The percentages of subdiploid (i.e., apoptotic) cells in samples incubated with DCVCS (10–500 µM) were significantly greater (P < 0.05) than the control value at each incubation time. Percentage of cells in subdiploid, G0/G1, G2/M, and S phases are as follows: control, 1.39, 71.08, 12.29, and 16.63; 10 µM, 7.98, 77.10, 12.64, and 10.26; 50 µM, 12.17, 75.37, 12.37, and 12.26; 100 µM, 12.41, 74.97, 11.24, and 13.79; 200 µM, 7.22, 75.50, 11.95, and 12.54; and 500 µM, 6.09, 72.31, 21.58, and 6.10.

From the plot in Fig. 3, the 4-h incubation time seems to elicit the optimal response for cells to undergo apoptosis. An example of the DNA histograms for hPT cells incubated with various concentrations of DCVCS for 4 h is shown in Fig. 4. Note that G₀/G₁ peaks comprise the majority of cells in most incubations and are somewhat broad because the cells are confluent. A progressive increase in the proportion of subdiploid cells up to 100 µM DCVCS was observed. This was followed by a modest decline in the proportion of subdiploid or apoptotic cells. The only significant changes in the proportion of hPT cells in the other phases of the cell cycle were observed in cells incubated with 500 µM DCVCS, which exhibited a nearly doubling in the proportion of cells in the G₂/M phase and a decrease by >50% in the proportion of cells in the S phase. Also note that the total number of viable cells, as indicated by the scale for the y-axes in each panel of Fig. 4, was significantly decreased with DCVCS concentrations 100 µM. This is consistent with cell loss due to necrosis or aggregation at higher DCVCS concentrations.

View larger version (33K): [in this window] [in a new window]   Fig. 4. Cell cycle analysis of hPT cells incubated for 4 h with DCVCS. Cells were incubated with PBS (control) or the indicated concentration of DCVCS for 4 h and were processed for analysis of DNA content as described under Materials and Methods and the legend to Fig. 3. Peaks from left to right represent apoptotic cells, cells in G0/G1, cells in S phase, and cells in G2/M. Results are from a single preparation and are typical of those from four separate experiments. The coefficient of variation (CV) for these peaks ranged between 3 and 14%, with approximately 80% of the peaks in samples exhibiting CV values of 5 to 8%. Insets, distribution of cells according to fluorescence intensity. Cells outside the box are those that were excluded from the analysis due to aggregation. Values above the arrows indicate the percentage of cells that are subdiploid.

As an additional determination of apoptosis, we assessed staining of hPT cells with annexin V-FITC (Fig. 5). Significant increases in the proportion of annexin V-positive/propidium iodide-negative cells was observed at all concentrations of DCVCS (i.e., 10–500 µM) after both 2- and 4-h incubations, providing additional evidence of the induction of apoptosis.

View larger version (17K): [in this window] [in a new window]   Fig. 5. DCVCS-induced apoptosis in primary cultures of hPT cells as indicated by annexin V-FITC staining and flow cytometry. Confluent hPT cells were grown in T-25 flasks and were treated for 2 or 4 h with either PBS (control) or the indicated concentrations of DCVCS. Samples were processed for analysis of annexin V-FITC staining, as described under Materials and Methods and were analyzed by flow cytometry. Values shown are the percentage of cells that are apoptotic, and represent cells in the lower right quadrant that are annexin V-positive and propidium iodide-negative. Results are the means ± S.E.M. of measurements from three separate cell cultures. All values for cells incubated with 10 to 500 µM DCVCS for either 2 or 4 h were significantly greater (P < 0.05) than those for the corresponding control.

Alterations in Mitochondrial Function of hPT Cells Induced by DCVCS. Early stages of induction of apoptosis involve, in many cases, alterations in mitochondrial function. Some of the effects serve as signaling processes in the induction of apoptosis. For cells to be competent to undergo apoptosis, adequate supplies of ATP and the ability to generate a must exist. Hence, delineation of the time and concentration dependence at which changes in mitochondrial function occur can provide information about competence of cells to undergo apoptosis.

Mitochondrial in confluent hPT cell cultures was qualitatively assessed with the fluorescent dye JC-1 and confocal microscopy (Fig. 6). JC-1 accumulates in mitochondria of cells and undergoes a fluorescence emission shift from green (530 nm) to red (590 nm) in polarized mitochondria. Accordingly, a cell population with polarized and functional mitochondria will exhibit red punctate staining, whereas mitochondria that are depolarized exhibit yellow-green staining. Photomicrographs from an exemplary set of JC-1 fluorescence measurements of hPT cells treated for 2 h with various concentrations of DCVCS indicate little effect at DCVCS concentrations of 100 µM and maintenance of some even at 200 µM DCVCS (Fig. 6A). In contrast, a marked decrease in the appearance of red staining in cells incubated for 4 h with 50 µM DCVCS was observed and at 200 µM DCVCS, no yellow or red staining was observed. Hence, this shows that hPT cells treated with up to 200 µM DCVCS for 2 h or up to 50 µM DCVCS for 4 h retain the ability to generate a mitochondrial .

View larger version (70K): [in this window] [in a new window]   Fig. 6. Effect of DCVCS on mitochondrial membrane potential in primary cultures of hPT cells. Confluent, primary cultures of hPT cells were grown on collagen-coated glass coverslips in 35-mm tissue culture dishes and were incubated for 2 or 4 h with 0 (control), 10, 50, 100, or 200 µM DCVCS. JC-1 fluorescence was measured by confocal microscopy assessing the emission shift from green (530 nm) to red (590 nm) in polarized mitochondria using 488-nm excitation. Composite red and green fluorescence is shown. Polarized mitochondria are indicated by yellow-red punctate staining. White scale bars, 5 µm. Results are from one cell culture and is representative of those from three separate cultures.

Mitochondrial integrity of hPT cells treated with DCVCS was also assessed by measurement of cellular respiration (Fig. 7). hPT cells incubated for up to 48 h with 10 to 500 µM DCVCS exhibited time- and concentration-dependent decreases in the rate of succinate-stimulated oxygen consumption. Maximal decreases in respiration rate of approximately 30% were observed after up to 2 h of incubation with as much as 500 µM DCVCS. This modest extent of functional decrement suggests that the mitochondria and the cells are still largely functional. Incubations with DCVCS concentrations of 100 µM for 4 h produced >50% decreases in respiration rates, suggesting that mitochondrial and cellular function are compromised. These time and dose dependence patterns are consistent with the diminished ability of hPT cells treated with 100 µM DCVCS at later incubation times to undergo apoptosis (cf. Fig. 3).

View larger version (22K): [in this window] [in a new window]   Fig. 7. Inhibition of cellular respiration in hPT cells by DCVCS. Confluent, primary cultures of hPT cells were incubated with the indicated concentrations of DCVCS for up to 48 h. At specified times, succinate-stimulated O2 consumption was measured in a 5/6H oxygraph (Gilson Medical Electronics). Results are averages of measurements from two separate cell cultures.

Although succinate-stimulated respiration was clearly altered by DCVCS in a time- and concentration-dependent manner and respiration coupled to site II (i.e., succinate/ubiquinone oxidoreductase) substrates such as succinate were previously shown to be the primary form inhibited by DCVC in rat kidney cells (Lash and Anders, 1986), we also examined respiration coupled to site I (i.e., NADH dehydrogenase) and site III (i.e., ubiquinol/cytochrome c oxidoreductase) substrates (Fig. 8). hPT cells were incubated for up to 4 h with either PBS or 50 µM DCVCS. At the indicated times, effects on respiration coupled to either glutamate + malate (site I substrates), succinate (site II substrate), or ascorbate + TMPD (site III substrates) was determined. As previously shown with DCVC, succinate-stimulated respiration was the earliest and most potently inhibited form. Although respiration coupled to glutamate + malate was also inhibited by DCVCS, the extent of inhibition was less and seemed to occur on a later time scale that that with succinate as the respiratory substrate. Respiration coupled to ascorbate + TMPD was not inhibited by 50 µM DCVCS at any incubation time.

View larger version (27K): [in this window] [in a new window]   Fig. 8. Respiration coupling site specificity of DCVCS in hPT cells. Confluent, primary cultures of hPT cells were incubated with 50 µM DCVCS for up to 4 h. At specified times, O2 consumption stimulated by either 1 mM L-glutamate + 1 mM malate, 3 mM succinate, or 1 mM ascorbate + 0.2 mM TMPD was measured in a 5/6H oxygraph (Gilson Medical Electronics). Results are means ± S.E.M. of measurements from three separate cell cultures. Significantly different (*, P < 0.05) from corresponding control sample.

Finally, mitochondrial function was assessed by measurement of cellular concentrations of ATP (Fig. 9). Incubation of hPT cells for 0.5 h with 10 to 200 µM DCVCS exhibited relatively modest decreases in ATP concentration of 50%. In contrast, incubation of hPT cells with as low a concentration of DCVCS as 10 µM for 1 h caused approximately 60% ATP depletion, whereas incubation with 10 µM or higher concentrations of DCVCS for 2 h or longer caused >70% ATP depletion. As with mitochondrial respiration, the large extent of ATP depletion induced by DCVCS concentrations >100 µM at early time points and 10 µM at later time points is consistent with a diminished competence of the cells to undergo apoptosis under these conditions.

View larger version (24K): [in this window] [in a new window]   Fig. 9. ATP depletion in hPT cells incubated with DCVCS. Confluent hPT cells grown on collagen-coated, polystyrene T-25 flasks were incubated for up to 4 h with either medium (control) or the indicated concentration of DCVCS. Cellular contents of ATP were measured in perchloric acid extracts of cells by reversed-phase HPLC. Results are means ± S.E.M. of measurements from four separate cell cultures. Significantly different (P < 0.05) from corresponding control: 0.5 h, 50 µM and higher; and 1, 2, and 4 h, 10 µM and higher.

Effects of DCVCS on GSH Concentrations of hPT Cells. Although DCVC does not markedly deplete cellular GSH concentrations in rPT cells (Lash and Anders, 1986), DCVCS is more chemically reactive than its precursor and reacts directly with GSH (Sausen and Elfarra, 1991). Incubation of hPT cells with up to 200 µM DCVCS produced relatively modest decreases in cellular GSH content (Fig. 10). In each case, the modest depletion of GSH, which was maximal after 2 or 4 h of incubation, was followed by a rebound by 8 h to near initial levels. Only in cells incubated with 500 µM DCVCS was more than 50% depletion of GSH observed. Additionally, cells incubated with 500 µM DCVCS did not exhibit a rebound in cellular GSH content by the 8-h incubation time.

View larger version (19K): [in this window] [in a new window]   Fig. 10. GSH depletion in hPT cells incubated with DCVCS. Confluent hPT cells grown on collagen-coated, polystyrene T-25 flasks were incubated for up to 8 h with either medium (control) or the indicated concentration of DCVCS. Cellular contents of GSH were measured in perchloric acid extracts of cells after derivatization with iodoacetate and 1-fluoro-2,4-dinitrobenzene by reversed-phase HPLC. Results are means ± S.E.M. of measurements from four separate cell cultures. Significantly different (P < 0.05) from control: 1 h, 50, 200, and 500 µM;2h,100 µM and higher; 4 h, 100 µM and higher; and 8 h, 500 µM.

	Discussion

Top Abstract Materials and Methods Results Discussion References

 
We show here that DCVCS is a potent cytotoxicant in primary cultures of hPT cells, as judged by effects on several parameters of cell function and toxicity, including morphology, LDH release, apoptosis, mitochondrial function, and GSH status. These findings agree with previous studies in rats and in isolated rPT cells (Lash et al., 1994). Three critical issues on which the present studies shed some light are the 1) temporal relationship between the various effects of DCVCS, 2) comparative toxicity of DCVCS and DCVC, and 3) role of DCVCS in TRI- and DCVC-induced nephrotoxicity in human kidney.

DCVCS produced both necrosis and apoptosis. Similar to the case with DCVC (Lash et al., 2001a), apoptosis induced by DCVCS was primarily a low-dose, early-incubation time response, whereas necrosis, as indicated by LDH release, was primarily a high-dose, late-incubation time response. Maintenance of mitochondrial function, as assessed by , respiration, and ATP concentrations, was also associated with the capability to undergo apoptosis, although the relationship was not always obvious.

With respect to membrane potential, hPT cells incubated for 2 h with up to 200 µM DCVCS still exhibited red-yellow JC-1 fluorescence, indicating maintenance of some degree of mitochondrial integrity. In contrast, cells incubated for 4 h with 50 µM DCVCS exhibited little to no red-yellow JC-1 fluorescence, indicating an inability to generate a membrane potential. Thus, although a statistically significant increase in the induction of apoptosis by 10 µM DCVCS occurred as early as 1 h, maximal induction of apoptosis did not occur until 4 to 8 h. Although the JC-1 data suggested little change in mitochondrial membrane potential by 10 µM DCVCS at 4 h, this is a relatively qualitative assay. It is conceivable that enough of a decrease in membrane potential occurred in the overall cell population or that membrane potential decreased in enough individual cells to cause the observed small, but statistically significant increase in apoptosis.

Regarding mitochondrial respiration, hPT cells needed to be incubated with at least 100 µM DCVCS for at least 4 h to elicit >30% inhibition of cellular oxygen consumption with succinate as respiratory substrate, again indicating maintenance of mitochondrial function at lower toxicant concentrations and at early incubation times. While significant inhibition of site I (i.e., NADH dehydrogenase)-coupled respiration was also observed, site II (i.e., succinate/ubiquinone oxidoreductase)-coupled respiration was clearly the most sensitive and site III (i.e., ubiquinol/cytochrome c oxidoreductase) respiration was insensitive to inhibition by DCVCS.

Although ATP depletion was a highly sensitive indicator of mitochondrial dysfunction, hPT cells required at least a 2-h incubation with 50 µM DCVCS to elicit >70% ATP depletion. However, the observations that cellular ATP concentrations were <2 nmol/mg of protein with DCVCS concentrations 50 µM after 2 or 4 h and apoptosis was not maximal until 4 to 8 h could suggest a temporal discrepancy between apoptosis and ATP supply. Although a minimum level of intracellular ATP is needed for cells to remain competent to undergo apoptosis (Kroemer et al., 1998), it is unclear what precisely this value is, although an ADP/ATP ratio of 0.2 has been suggested as the critical level (Richter et al., 1996). It would seem that cellular ATP concentrations of 1 to 2 nmol/mg protein are still sufficient to allow apoptosis to occur in hPT cells.

Cellular responses to DCVCS differed from those to DCVC in several important ways. Experiments with DCVCS reported in this study were performed on different cell cultures from different kidney donors than those with DCVC that were previously published (Lash et al., 2001a). This is potentially important because of possible interindividual differences in bioactivation and susceptibility, although there have been no documented interindividual differences in rates of DCVC metabolism. We previously noted that hPT cells from males were more sensitive to DCVC than were hPT cells from females but that there were no observable gender-dependent differences in apoptosis (Lash et al., 2001a). In terms of acute cellular necrosis, hPT cells from male donors seemed to be slightly less sensitive to DCVCS than to DCVC, particularly at concentrations <100 µM. Specifically, the time course for cell death was slower in hPT cells incubated with DCVCS. hPT cells also exhibited significantly less apoptosis when incubated with DCVCS than with DCVC. Thus, whereas the maximal extent of apoptosis, as indicated by FACS analysis, was only about 12% in hPT cells incubated with DCVCS, the maximal extent of apoptosis due to DCVC was approximately 30%. Similarly, whereas the maximal amount of apoptosis as indicated by annexin V-FITC staining was about 12% above control cells for DCVCS, it was about 30% above control cells for DCVC. Although effects of DCVCS on mitochondrial and respiration rates were generally comparable with those of DCVC, DCVCS caused a much greater extent of ATP depletion than DCVC (>90% for DCVCS and only about 60% for DCVC; L. H. Lash, D. A. Putt, and S. E. Hueni, unpublished observations). The more rapid and extensive ATP depletion caused by DCVCS is consistent with its diminished ability relative to DCVC to cause hPT cells to undergo apoptosis.

Finally, although DCVCS only produced marked GSH depletion at the highest concentration tested (i.e., 500 µM), the amounts of GSH depletion observed with DCVCS were still markedly higher than those produced by DCVC (L. H. Lash, D. A. Putt, and S. E. Hueni, unpublished observations). This is likely due to the chemical reactivity of DCVCS with soft nucleophiles such as GSH (Sausen and Elfarra, 1991). DCVC selectively depletes mitochondrial GSH in rat kidney by oxidation and does not have a measurable impact on total renal cellular GSH content (Lash and Anders, 1986, 1987).

Based on the present work and previous studies, we conclude that sulfoxidation of DCVC does indeed play a significant role in the mechanism of toxicity in human kidney. Previous demonstrations in both freshly isolated and primary cultures of hPT cells (Cummings and Lash, 2000; Lash et al., 2001a) showing marked protection from DCVC-induced necrosis or apoptosis by preincubation of cells with the FMO substrate methimazole directly implicate DCVCS formation in the mechanism of toxicity. Moreover, the relatively low amount of total -lyase activity in the human kidney (Lash et al., 1990) and the lack of or marginal ability of aminooxyacetic acid to protect hPT cells from DCVC-induced necrosis or apoptosis, suggest further that FMO-dependent bioactivation is important. We also recently demonstrated expression of FMO isozymes in human kidney (Krause et al., 2003). These studies, although derived from a limited number of samples, also provided data suggesting that humans exhibit a wide range of levels of expression of FMOs in the kidneys. This may indicate a correspondingly wide range of susceptibility in the general population to nephrotoxicity from chemicals that undergo FMO-dependent bioactivation. Additional studies with higher sample numbers and specific genotyping of individuals are needed to confirm this finding.

In comparing DCVCS and DCVC, it is critical to note that effects due to reactive species generated from both DCVC and DCVCS contribute to the overall effects of incubations with DCVC. In contrast, one only observes effects due to reactive species generated from DCVCS when cells are incubated with DCVCS. Furthermore, the molecular targets for reactive species generated by -lyase and FMO may differ. These considerations make a direct comparison of DCVC and DCVCS and an assessment of the role of DCVCS formation in the cytotoxicity induced by DCVC difficult. The markedly higher amount of apoptosis induced by DCVC compared with that induced by DCVCS and the consistent findings that apoptosis is a relatively low-dose, early response, suggest that for DCVC bioactivation and cytotoxicity, -lyase-dependent processes may be more important at lower doses and during the early stages of exposure, whereas a role for FMO may become more prominent at higher doses and at later exposure times. This conclusion is also supported by the apparent kinetics of DCVC metabolism by the two enzymes. Whereas the -lyase from human kidney cytosol exhibits a K_m value of approximately 1.5 to 2 mM for DCVC (Lash et al., 1990), the reported K_m value for DCVCS formation from DCVC by cDNA-expressed rabbit FMO3 is >50 mM (Ripp et al., 1997). Although this kinetic difference suggests that FMO may be only important at very high concentrations, this will be modified by the apparent, higher reactivity of reactive species derived from DCVCS than those derived from -lyase-dependent metabolism of DCVC.

In conclusion, we have shown that DCVCS is a potent cytotoxicant in primary cultures of hPT cells that elicits both apoptosis and necrosis, according to a specific temporal- and dose-dependent pattern. This pattern seems to correlate with the ability of DCVCS-treated cells to maintain mitochondrial integrity, such that the ability to generate a membrane potential and to maintain some minimal level of intracellular ATP are required for the cells to be capable of undergoing apoptosis. Although a correlation between cellular GSH status and these responses was not specifically addressed, lower concentrations of DCVCS induce a modest, but reversible depletion of GSH, suggesting a transient oxidative stress. At high DCVCS concentrations, however, the oxidative stress is irreversible and cells undergo necrosis.

	Footnotes

 
Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.

DOI: 10.1124/jpet.102.046185.

This work was funded by National Institute of Environmental Health Sciences Grant R01-ES08828 (to L.H.L.) and National Institute of Diabetes and Digestive and Kidney Diseases Grant R01-DK44295 (to A.A.E.). Core facilities funded by the National Institute of Environmental Health Sciences Center for Molecular Toxicology with Human Applications (Grant P30-ES06639) at Wayne State University were used for some of this study.

ABBREVIATIONS: TRI, trichloroethylene; GSH, reduced glutathione; DCVC, S-(1,2-dichlorovinyl)-L-cysteine; FMO, flavin-containing monooxygenase; DCVCS, S-(1,2-dichlorovinyl)-L-cysteine sulfoxide; rPT, rat proximal tubular; hPT, human proximal tubular; TMPD, N,N,N',N'-tetramethyl-p-phenylenediamine; FITC, fluorescein isothiocyanate; HPLC, high-pressure liquid chromatography; PBS, phosphate-buffered saline; DMEM/F-12, Dulbecco's modified Eagle's medium/Ham's F-12; FACS, flow activated cell sorter; LDH, lactate dehydrogenase; , mitochondrial membrane potential.

Address correspondence to: Dr. Lawrence H. Lash, Department of Pharmacology, Wayne State University School of Medicine, 540 East Canfield Ave., Detroit, MI 48201. E-mail: l.h.lash{at}wayne.edu

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