Elevated zinc induces endothelial apoptosis via disruption of glutathione metabolism: role of the ADP translocator

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Zinc is the second-most abundant transition metal within cells and an essential micronutrient. Although adequate zinc is essential for cellular function, intracellular free zinc (Zn2+) is tightly controlled, as sustained increases in free Zn2+ levels
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  Elevated zinc induces endothelial apoptosis via disruption of glutathione metabolism: role of the ADP translocator  Dean A. Wiseman , Shruti Sharma , and Stephen M. Black Vascular Biology Center, Medical College of Georgia, 1459 Laney Walker Blvd CB-3210, Augusta,GA 30912-2500, USA Stephen M. Black: sblack@mcg.edu  Abstract Zinc is the second-most abundant transition metal within cells and an essential micronutrient.Although adequate zinc is essential for cellular function, intracellular free zinc (Zn 2+ ) is tightlycontrolled, as sustained increases in free Zn 2+  levels can directly contribute to apoptotic endothelialcell death. Moreover, exposure of endothelial cells to acute nitrosative and/or oxidative stress inducesa rapid rise of Zn 2+  with mitochondrial dysfunction and the initiation of apoptosis. This apoptoticinduction can be mimicked through addition of exogenous ZnCl 2  and mitigated by zinc-chelationstrategies, indicating Zn 2+ -dependent mechanisms in this process. However, the molecular mechanisms of Zn 2+ -mediated mitochondrial dysfunction are unknown. Here we report that freeZn 2+  disrupts cellular redox status through inhibition of glutathione reductase, and induces apoptosis by redox-mediated inhibition of the mitochondrial adenine nucleotide transporter (ANT). Inhibitionof ANT causes increased mitochondrial oxidation, loss of ADP uptake, mitochondrial translocationof bax, and apoptosis. Interestingly, pre-incubation with glutathione ethyl ester protects endothelialcells from these observed effects. We conclude that key mechanisms of Zn 2+ -mediated apoptoticinduction include disruption of cellular glutathione homeostasis leading to ANT inhibition and decreases in mitochondrial ATP synthesis. These pathways could represent novel therapeutic targetsduring acute oxidative or nitrosative stress in cells and tissues. Keywords Mitochondrial dysfunction; Apoptosis; Redox status Introduction Understanding the nature of how cells transduce, interpret, and respond to stress signals, bothin terms of extracellular perception as well as signals from within, is fundamental for discovering and developing new therapies and interventions for situations where cells, tissues,and organs lose the ability to maintain homeostatic balance. It is now apparent reactive oxygenand nitrogen species (ROS, RNS, respectively) serve as signal transduction molecules in biological systems. ROS such as hydrogen peroxide (H 2 O 2 ) and superoxide (O 2• − ) are alsoincreasingly understood to have complex and highly nuanced signal transduction roles in cellsand tissues (Forman and Torres 2001). However, these same chemical species can also initiatesignaling pathways that result in apoptosis (Bauer 2000). We, and others, have previouslyreported that exposure of pulmonary artery endothelial cells to elevated levels of ROS and/or  © Springer Science+Business Media, LLC. 2009Correspondence to: Stephen M. Black, sblack@mcg.edu .  NIH Public Access Author Manuscript  Biometals . Author manuscript; available in PMC 2011 February 1. Published in final edited form as:  Biometals . 2010 February ; 23(1): 19–30. doi:10.1007/s10534-009-9263-y. NI  H-P A A  u t  h  or M an u s  c r i   p t  NI  H-P A A  u t  h  or M an u s  c r i   p t  NI  H-P A A  u t  h  or M an u s  c r i   p t    RNS leads to significant elevation in intracellular free zinc (Zn 2+ ) (Tang et al. 2001; Wisemanet al. 2006, 2007; Bernal et al. 2008). Zinc is the second-most common transition metal in cells(Atwood and Steed 2004), and zinc ions function in multiple different intracellular processes.It is estimated that at least 3% of the known human genes encode proteins containing zinc-finger domains (Maret 2003). However, the concentration of unbound “free” intracellular zincis typically found at picomolar levels (Simons 1991). Thus, although Zn 2+  is classified as“redox inert,” if not adequately sequestered it can also function as a potent disruptive moleculewithin cells. Here, we present data that demonstrate that the effect of Zn 2+  on mitochondrialdysfunction is indirect. Further, we show that a key mechanism for this indirect effect isdisruption of glutathione homeostasis. Interestingly, we find that a direct consequence of Zn 2+ -mediated depletion of reduced cellular glutathione is the inhibition of mitochondrialadenine nucleotide transport. Together, these data illustrate a cell-death mechanism wherebymitochondrial dysfunction is initiated through a Zn + -induced loss of cellular antioxidantcapacity, resulting in mitochondria deprived of adequate metabolites for normal function. Materials and methods Cell culture Primary cultures of ovine fetal pulmonary artery endothelial cells (PAECs) were isolated and identified as described previously (Wedgwood et al. 2003). Cells between passages 3–10 weremaintained in DMEM supplemented with 10% fetal bovine serum (Hyclone, Logan, UT),antibiotics and antimycotics (MediaTech, Herndon, VA) at 37°C with 5% CO 2  –95% air. Cellswere seeded at ~50% confluence, and at ~90% confluence. Before experimental treatment,except as noted, cells were trypsinized, counted, re-plated in 6-, 24-, or 96-well plates (Costar,Corning, NY) at a density of 5 × 10 5  cells/cm 2 , and incubated 18 h. Cells were serum starved in serum-free, phenol red-free DMEM (SF/PRF-DMEM, GIB-CO-BRL, Gaithersburg, MD)and incubated overnight. During serum deprivation, no exogenous sources of Zn 2+  are availableto the cells. Cells were pretreated for 2 h with SF/PRF-DMEM ± glutathione ethyl ester (GSH- EE, 2 mM, Sigma, St. Louis, MO), and then exposed 0–4 h with 0–1 mM ZnCl 2  (200 mM inPBS, diluted in SF/PRF-DMEM), 0–1 mM  N  -[2-aminoethyl]-  N  -[2-hydroxy-2-nitroso-hydrazino]-1,2-ethylenediam-ine (spermine NONO-ate; Calbiochem, San Diego, CA) in SF/PRF-DMEM. Doses were selected to examine the entire range of potential response in PAECsas we (Wiseman et al. 2006, 2007), and others (Koh and Choi 1994) previously described. Atdesignated time points, cells were subjected to immediate analysis, unless noted. Mitochondrial isolation PAECs were scraped, washed PBS, and pelleted by centrifugation 850× g  at 4°C for 2 min.Mitochondria were isolated via the manufacturer’s protocol (Pierce Biotechnology, Rockford,IL). Mitochondrial fractions were resuspended in 0.25 M sucrose, 10 mM Tris-C1, pH 7.4, and 0.5 mM EDTA. In order to avoid potential issues with chelation of Zn 2+  ions by EDTA,immediately prior to the beginning of each experiment, mitochondrial fractions werecentrifuged and resuspended in EDTA-free buffer, and assayed as described below. Fluorescence microscopy A PC-based imaging system consisting of: an Olympus IX51 microscope equipped with acharge-coupled camera (Hamamatsu Photonics, Hamamatsu City, Japan) was used for acquisition of fluorescent images. Fluorescent-stained cells were observed using appropriateexcitation and emission, measuring at least 300 cells per sample, and the average fluorescentintensities (to correct for differences in cell number) were quantified using ImagePro Plus v5.0software (Media Cybernetics, Silver Spring, MD). High-resolution cellular images wereobtained using an Applied Precision, Inc. (Issaquah, WA) DeltaVision™ imaging system, Wiseman et al.Page 2  Biometals . Author manuscript; available in PMC 2011 February 1. NI  H-P A A  u t  h  or M an u s  c r i   p t  NI  H-P A A  u t  h  or M an u s  c r i   p t  NI  H-P A A  u t  h  or M an u s  c r i   p t    fitted with an environmental control chamber. Data obtained were quantified usingDeltaVision™ software and compared by statistical analysis. Detection of bax translocation Ten hours following treatment, mitochondria were isolated and resuspended in lysis buffer (20mM Tris base, pH 7, 2.5 mM EDTA, 1% Triton-X, 1:100 protease inhibitor cocktail [Sigma]).Lysates were assayed for protein concentration by BCA protein assay (Pierce Biotechnology,Rockford, IL) and normalized for concentration. Sixty microgram of mitochondrial protein per sample was subjected to SDS–PAGE and analyzed by standard Western blot analysis asdescribed previously (Brennan et al. 2003) using anti-bax antibody (Cell SignalingTechnology, Boston, MA). Detection of apoptotic events PAECs in 96-well plates were treated as described. Caspase activation was visualized by co-treating cells with 1 µM CaspACE FITC-vad-FMK (Promega, Madison, WI). This fluorescentanalog of the pancaspase inhibitor  N  -benzyloxycarbonyl-Val-Ala-Aspfluor-omethylketone(Z-vad-FMK) readily enters cells and binds irreversibly to activated caspases. After treatment,cells were washed with ZnCl 2 -free DMEM and incubated overnight in PRF-DMEM mediawith 1% FBS. Eighteen hours post-exposure, cells were incubated in media with 10 µM FITC-conjugated caspase inhibitor (FITC-vad-FMK). After 20 min incubation at 37°C in dark conditions, cells were washed with fresh media and visualized using fluorescence microscopy.A second method involved TUNEL analysis. PAECs were incubated for ~18 h in PRF-DMEMsupplemented with 1% FBS. Following incubation, analysis was performed as we have previously described (Wedgwood and Black 2003). EPR detection of mitochondrial O 2• −  levels Mitochondrial-specific O 2• −  production was measured using electron paramagnetic resonance(EPR) spectroscopy with spin probe, 1-hydroxy-3-methox-ycarbonyl-2,2,5,5-tetramethyl- pyrrolidine•HCl (CMH, Alexis Biochemicals, San Diego, CA) as described (Wiseman et al.2007). Following exposure, mitochondrial isolation was performed as described above.Mitochondrial fractions were incubated for 1 h in the presence of CMH (PBS, pH 7.4 + 25 µMdeferrioxamine mesylate [Calbiochem], CMH 5 mg/ml). Thirty five microliter of sample wasloaded into a 50 µl capillary tube and analyzed with a MiniScope MS200 EPR (Magnettech,Berlin, Germany) at a microwave power of 40 mW, modulation amplitude of 3,000 mG, and modulation frequency of 100 kHz. EPR spectra were analyzed measured for amplitude usingANALYSIS software (v2.02; Magnettech). Detection of cellular and mitochondrial redox status PAECs were plated into 10 cm dishes at a density of ~15,000 cells/cm 2  (~1.25 × 10 6  cells per dish) and transfected with expression constructs of roGFP, redox-sensitive GFP analogs wheresurface-exposed residues are substituted with serine, allowing the ability to monitor the cellular thiol redox status of cells. These GFP analogs demonstrate a redox-specific shift in fluorescencespectra directly correlated to cellular oxidation status (Hanson et al. 2004). Redox-sensitiveconstructs (generously provided by S. James Remington, University of Oregon) delivered either general cytosolic roGFP expression (“cRo,” Fig. 3a), or expression of roGFP specificallywithin mitochondria (“mRo,” Fig. 3a). Transfections were performed via Effectene® lipid- based delivery protocols (Qiagen). Following transfection with either cRo or mRo constructs,cells were subdivided into three groups and re-plated onto cover glass. One group was treated as described above to assess cellular redox status for up to 4 h post-onset of exposure, whilethe other two groups were used to calibrate maximal reduction (using 1 mM dithiothreitol,Sigma) and maximal oxidation (using 1 mM t  -butyl hydroperoxide, Sigma) (Farrow et al. Wiseman et al.Page 3  Biometals . Author manuscript; available in PMC 2011 February 1. NI  H-P A A  u t  h  or M an u s  c r i   p t  NI  H-P A A  u t  h  or M an u s  c r i   p t  NI  H-P A A  u t  h  or M an u s  c r i   p t    2008), and cellular fluorescence at 470 and 360 nm excitation wavelengths in order todetermine the ratio of reduced versus oxidized GFP for up to 4 h. In addition, an untransfected sample of cells was quantified to determine nonspecific background. Each experimental groupwas evaluated as a percentage of oxidation, relative to fluorescence values obtained for fullyreduced and fully oxidized conditions. Measurement of glutathione reductase activity The assay for glutathione reductase activity is based on oxidation of NADPH to NADP+catalyzed by a limiting concentration of glutathione reductase. One GR activity unit is defined as the amount of enzyme catalyzing reduction of one micromole of GSSG per minute at pH7.6 and 25°C. Given that one molecule of NADPH is consumed for each molecule of GSSGreduced, reduction of GSSG is determined indirectly by measurement of NADPHconsumption, observed as a decrease in absorbance at 340 nm (A 340 ) over time. Briefly, PAECswere treated and harvested by scraping and centrifugation at 1,000× g  for 10 min at 4°C. Cell pellets were homogenized in 200 µl of ice-cold KPO 4  buffer (100 mM, 0.1% BSA, 5 mMEDTA, pH 7.5) and centrifuged at 10,000× g  for 15 min at 4°C, and supernatants kept on icefor immediate assay. To each sample, a known amount of NADPH was added, and A 340  wascontinuously monitored 60 s. Extinction coefficients were calculated and activity determined via comparison to known standards of purified glutathione reductase. To determine if the effectof Zn 2+  is acting directly upon the glutathione reductase enzyme, samples of purified glutathione reductase was exposed or not to 50 µM ZnCl 2  and analyzed as described. Determination of cellular GSH:GSSG ratio This assay used is based on reaction of GSH with DTNB (Ellman’s reagent, [5,5 ′ -dithiobis-2-nitrobenzoic acid]), which produces a detectable product with a maximal absorbance at 412nm. The rate of product formation is proportional to the concentration of GSH. As GSHtypically far exceeds the amount of oxidized GSSG within cells, this assay uses a thiol-scavenging reagent, 1-methyl-2-vinylpyridinium trifluoromethanesulf-onate1 (M2VP) toscavenge GSH. Samples are then incubated with a known amount of purified glutathionereductase in the presence of DTNB, and ratios calculated according to the manufacturer’s protocol (Oxis International). For both GSSG and GSH, absorbance curves were generated and concentrations were extrapolated via comparison to known GSH standards. Determination of mitochondrial ATP production Following exposure to Zn 2+  (500 µM, 4 h), isolated mitochondria were incubated in physiological conditions with exogenous ADP (100 µM, Sigma, St. Louis, MO) in the presenceof ATP-dependent luciferase, according to the manufacturer’s protocol (Sigma, St. Louis,MO). Mitochondrial isolates were volume adjusted to 800 µl, pH 7.8. Hundred microliter of ATP Assay Mix solution was added to isolates, and allowed to incubate at room temperaturefor ~3 min, during which any endogenous ATP is hydrolyzed, decreasing background signal.The assay was performed by rapidly adding 100 µl of either ATP standard or ADP, with mixing,and measured with a luminometer. Negative controls were performed using mitochondrial-freereactions and mitochondrial fractions absent of exogenous ADP.In addition, we directly measured mitochondrial adenine uptake. [ 3 H]ADP uptake wasmeasured by carboxyatractyloside inhibitor stop-technique described previously (Chan and Barhour 1979). Approximately 50 µg of mitochondria were suspended in 200 µl buffer containing 116 mM KC1, 21 mM Tris/HCI, pH 7.4, 1.05 mM EDTA (KCl buffer), 5.26 mM2-oxoglutarate, and 5.26 µM  p -ruthenium red. Following 5-min incubation at RT, 10 pgoligomycin was added to the suspension and chilled to 2°C. Various concentrations (5 µl) of [ 3 H]ADP were added to 45 µ aliquots of mitochondrial suspension under constant mixing.After 12 s, reactions were stopped by 10 µl injection of 200 µM  p -carboxy-atractyloside, and  Wiseman et al.Page 4  Biometals . Author manuscript; available in PMC 2011 February 1. NI  H-P A A  u t  h  or M an u s  c r i   p t  NI  H-P A A  u t  h  or M an u s  c r i   p t  NI  H-P A A  u t  h  or M an u s  c r i   p t    mitochondria were pelleted at 12,000× g  for 4 min. Supernatant was removed and mitochondriawere washed with 200 µl KC1 buffer containing 10 nM carboxyatractyloside and re-pelleted.The mitochondrial pellet was dissolved in 100 µl of 2% SDS, transferred to a scintillation vialcontaining 3.0 ml of counting fluid, and counted by liquid scintillation. Statistical analyses Statistical calculations were performed using Graph-Pad Prism V. 4.01 software. The mean ±SD was calculated for all samples and significance determined by either by the unpaired t   testor ANOVA. For ANOVA, Neuman-Kuehls post-hoc testing was utilized. A value of P  < 0.05was considered significant. Results Our previous findings indicate that pulmonary artery endothelial cells (PAECs) undergomitochondrial dysfunction and apoptosis when exposed to either oxidative or nitrosative stress,and this effect can be mimicked if cells are exposed in the same fashion to exogenous Zn 2+ .Initially, we determined if elevated intracellular Zn 2+  induces mitochondrial dysfunction viaa direct or indirect mechanism. Isolated mitochondrial fractions (from ~2 × 10 7  cells) weresuspended in metabolic buffer (free from metal chelators, such as EDTA) and acutely exposed to either ZnCl 2  (Fig. 1a) or spermine NONOate (Fig. 1b, as a positive control) or for 1 h and superoxide levels determined using the spin-trap, CMH, a cyclic hydroxylamine with relativelyspecific affinity for O 2• −  (Fink and Dikalov 2002). CMH-O 2• −  products were detected by EPR analysis as we have described (Wiseman et al. 2006). Our data indicate that although NOincreases mitochondrial-derived O 2• −  (Fig. 1b), Zn 2+  alone does not (Fig. 1a), indicating thatZn 2+ -mediated mitochondrial dysfunction occurs indirectly.Previous studies show an important role for reduced GSH in maintaining mitochondrialfunction (Garcia-Ruiz et al. 1995). Thus, we next determined if the indirect effect of freeZn 2+  on mitochondrial function could be due to alterations in cellular GSH homeostasis. Wefound that Zn 2+  exposure caused a decrease in the GSH-to-GSSG ratio (Fig. 2a). As the rate-limiting enzyme in the homeostatic maintenance of cellular GSH-to-GSSG ratio is glutathionereductase (GR), we next ascertained if the mechanism by which increased free Zn 2+  produceselevated GGSG is mediated through inhibition of glutathione reductase (GR) enzymaticactivity. Our d ata indicate that elevated free Zn 2+  leads to the inhibition of GR activity inPAECs (Fig. 2b). Furthermore, to verify that Zn 2+  is a direct biochemical inhibitor of GR, wedetermined if Zn 2+  inhibits GR in vitro. We found the presence of Zn 2+  directly inhibits theability of GR to reduce GSSG to GSH (Fig. 2c).In order to examine the effect of the decreased GSH-to-GSSG ratio induced by elevated freeZn 2+  on cellular redox status, we utilized redox-sensitive GFP constructs targeted to either cytosol (“cRo”) or to mitochondria (“mRo”) (Fig. 3a). Following transfection with either cRoor mRo redox-sensitive GFP, we exposed PAECs to Zn 2+  (0–1 mM ZnCl 2 ) and monitored average cellular fluorescence for 4 h. Firstly, we observed a dose-dependent, significant shiftin mitochondrial roGFP to a more oxidized state (Fig. 3b). Secondly, we observed a shift in both cytosolic and mitochondrial redox status to a more oxidized state, but mitochondrialoxidation occurred earlier (Fig. 3c). Lastly, in cells treated with 1 mM glutathione ethyl ester  prior and during onset of exposure to Zn 2+ , we found significant protection of mitochondrialroGFP from Zn 2+ -induced oxidation (Fig. 3d).GSH is known to be important in preventing oxidation and subsequent inhibition of the adeninenucleotide translocator (ANT) protein in neuronal cells (Vesce et al. 2005). Thus, we next wedetermined if the shift of the mitochondria to a more oxidized environment had an effect onANT function. We utilized two assays to measure of ANT function: Initially we examined the Wiseman et al.Page 5  Biometals . Author manuscript; available in PMC 2011 February 1. NI  H-P A A  u t  h  or M an u s  c r i   p t  NI  H-P A A  u t  h  or M an u s  c r i   p t  NI  H-P A A  u t  h  or M an u s  c r i   p t  
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