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?JCB, Volume 170, Number 7, 1079-1090! Y2 w6 d/ |# S; i, F4 [, W
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Selective role for superoxide in InsP3 receptor–mediated mitochondrial dysfunction and endothelial apoptosis

Muniswamy Madesh1,2, Brian J. Hawkins1, Tatyana Milovanova1, Cunnigaiper D. Bhanumathy3, Suresh K. Joseph3, Satish P. RamachandraRao4, Kumar Sharma4, Tomohiro Kurosaki5, and Aron B. Fisher1: i2 y3 S( d) R! H; P$ i& s5 Y
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1 Institute for Environmental Medicine, University of Pennsylvania, Philadelphia, PA 19104( U8 o5 U) c1 U7 X. b- U1 p% v
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2 Department of Cancer Biology, University of Pennsylvania, Philadelphia, PA 191046 t1 _5 t9 N8 z% q5 Q8 p& ^

3 Department of Pathology, Anatomy, and Cell Biology

4 Dorrance Hamilton Research Laboratories, Thomas Jefferson University, Philadelphia, PA 19107- b, x2 ?# Z! b, S& I0 H4 d0 }

5 Laboratory for Lymphocyte Differentiation, Institute of Physical and Chemical Research, Research Center for Allergy and Immunology, Turumi-ku, Yokohama 230-0045, Japan
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Correspondence to Muniswamy Madesh: madeshm@mail.med.upenn.edu' W. H; O( D- t/ {: a
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Reactive oxygen species (ROS) play a divergent role in both cell survival and cell death during ischemia/reperfusion (I/R) injury and associated inflammation. In this study, ROS generation by activated macrophages evoked an intracellular Ca2  (i) transient in endothelial cells that was ablated by a combination of superoxide dismutase and an anion channel blocker. i store depletion, but not extracellular Ca2  chelation, prevented i elevation in response to O2.– that was inositol 1,4,5-trisphosphate (InsP3) dependent, and cells lacking the three InsP3 receptor (InsP3R) isoforms failed to display the i transient. Importantly, the O2.–-triggered Ca2  mobilization preceded a loss in mitochondrial membrane potential that was independent of other oxidants and mitochondrially derived ROS. Activation of apoptosis occurred selectively in response to O2.– and could be prevented by i buffering. This study provides evidence that O2.– facilitates an InsP3R-linked apoptotic cascade and may serve a critical function in I/R injury and inflammation.

Abbreviations used in this paper: 2-APB, 2-aminoethoxydiphenyl borate; BAPTA, 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetracetate; i, intracellular calcium; m, mitochondrial membrane potential; DCF, dichlorofluorescein; DPI, diphenyleneiodonium; ECM, extracellular medium; GPCR, G protein–coupled receptor; InsP3, inositol 1,4,5-trisphosphate; InsP3R, InsP3 receptor; I/R, ischemia/reperfusion; KO, knockout; LPS, lipopolysaccharide; MPTP, mitochondrial permeability transition pore; PI, propidium iodide; PMVEC, pulmonary microvascular endothelial cell; ROS, reactive oxygen species; SOD, superoxide dismutase; t-BuOOH, tert-butyl hydroperoxide; Tg, thapsigargin; TKO, triple KO; TMRE, tetramethylrhodamine, ethyl ester, perchlorate.
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Introduction/ T8 v2 K6 G1 W2 @/ Q3 w
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Receptor-mediated generation of reactive oxygen species (ROS) is necessary for signal transduction, gene expression, and cell proliferation in smooth muscle cells, T and B lymphocytes, and fibroblasts (Devadas et al., 2002). Conversely, ROS produced under pathological conditions such as ischemia/reperfusion (I/R) or inflammation are associated with cellular dysfunction and apoptosis (Davies, 1995). Endothelial cells respond to numerous external stimuli by producing the superoxide anion (O2.–). In physiological conditions, mitochondrial respiratory chain proteins produce O2.–, which can be dismutated into hydrogen peroxide (H2O2) or react with nitric oxide to produce peroxynitrite. In addition, reaction of H2O2 with iron leads to hydroxyl radical formation via Fenton chemistry. During I/R injury, O2.– production in the vasculature is substantially increased (Wei et al., 1999) and is accompanied by endothelial cytotoxicity (for review see Li and Shah, 2004). However, the molecular mechanisms by which ROS lead to organ damage are poorly understood.
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In pathological conditions, cell death is facilitated by an elevation in intracellular calcium (i; Hajnoczky et al., 2003; Orrenius et al., 2003) via inositol 1,4,5-trisphosphate (InsP3). InsP3 is a second messenger produced by the hydrolysis of phosphatidylinositol-4,5-bisphosphate by PLC. InsP3 receptor (InsP3R)–mediated i changes are associated with a rapid, transient Ca2  release from Ca2  stores in the ER followed by Ca2  entry through slow-activating plasma membrane store-operated channels (Putney and Bird, 1993; Parekh and Penner, 1997; Berridge et al., 1998). InsP3 i signals control a wide range of cellular functions, including cell proliferation and apoptosis (Berridge et al., 2000; Orrenius et al., 2003). Apoptosis is reduced in cells lacking all three InsP3R isoforms (DT40 avian B cells) and after selective suppression of InsP3R-3 (Jayaraman and Marks, 1997; Sugawara et al., 1997), indicating the important role of InsP3 in cell death mechanisms (Pan et al., 2001). Alterations in i after oxidative stress facilitate activation of the mitochondrial permeability transition pore (MPTP), which releases cytochrome c from the mitochondrial intermembrane space, leading to mitochondrial membrane potential (m) loss, assembly of the apoptosome, and activation of downstream caspases (Crompton, 1999). Recent evidence suggested that cytochrome c transiently released from mitochondria interacts with InsP3R and amplifies Ca2 -mediated apoptosis (Boehning et al., 2003).
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Endothelial cells subjected to oxidative stress undergo apoptosis (Warren et al., 2000). Although there is evidence that perturbations of cellular Ca2  homeostasis (including i elevation, ER Ca2  depletion, and mitochondrial Ca2  increases) occur, the mechanisms by which oxidative stress mediates endothelial apoptosis remain unclear. Events in the early stages of stress signaling include the mobilization of i (Patterson et al., 2004), the generation of ROS, and the formation of lipid peroxides. However, it is unclear whether radical formation is a consequence of Ca2  mobilization or a parallel event in early stress signaling. The proximity between mitochondria and the ER facilitates a higher Ca2  exposure in mitochondria relative to the cytosol when released from the ER (Rizzuto et al., 1998). During pathological situations, excess ER-released Ca2  may be detrimental to mitochondrial function and may trigger mitochondrial fragmentation and apoptosis. Previously, Bcl-2 family proteins have been implicated in apoptosis by affecting cellular Ca2  homeostasis (Pinton et al., 2000; Pan et al., 2001; Li et al., 2002). A recent study reported that a functional interaction of Bcl-2 with InsP3R attenuated InsP3R activation, which in turn controlled InsP3-evoked Ca2  release (Chen et al., 2004), in contrast to our findings that Bcl-XL activates InsP3R (White et al., 2005). In addition, ER-localized Bax and Bak can either interfere with ER Ca2  homeostasis or initiate apoptosis by activating caspase 12 (Zong et al., 2003).7 [; t3 L2 F6 k3 }1 \+ }

We previously reported that cells exposed to O2.– induced a rapid and large cytochrome c release (Madesh and Hajnoczky, 2001). We now provide evidence that O2.– evokes a large, transient i pool release from the ER, causing mitochondrial Ca2  elevation and rapid depolarization. Remarkably, the observed InsP3-linked mitochondrial phase of apoptosis was specific to O2.– and not other oxidant species. The O2.–-induced mitochondrial depolarization and downstream apoptotic cascades are independent of mitochondrial ROS production. Overall, this evidence provides a mechanism by which O2.– is a key signaling molecule that coordinates multiple processes that lead to mitochondrial apoptotic events and endothelial dysfunction.7 V6 v/ u+ g& D2 N2 f
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Results# M' g- U! S' X; [
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Lipopolysaccharide (LPS)-stimulated macrophages evoke Ca2  transients in endothelial cells
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Activated macrophages are known to generate ROS and may be involved in organ damage during I/R (Droge, 2002). To test the significance of the selective role of macrophage-derived ROS during pathophysiological conditions, LPS-stimulated murine macrophages were used as a O2.–-generating source. We determined whether O2.– released from macrophages could evoke Ca2  mobilization in two cell types, endothelial and HepG2 cells. ROS production in LPS-stimulated mouse macrophages was measured via H2DCF-DA, which is a nonfluorescent dye that produces the fluorescent compound dichlorofluorescein (DCF) when oxidized by ROS. DCF fluorescence was measured in untreated macrophages and those stimulated with LPS (1 μg/ml) or a combination of LPS and the NADPH oxidase inhibitor diphenyleneiodonium (DPI; 30 μM). LPS stimulation was associated with a pronounced increase in DCF fluorescence that was attenuated by DPI treatment, suggesting that LPS stimulated ROS production through activation of oxidative burst reactions (Fig. 1 A). The activation of macrophage NADPH oxidase generates O2.– extracellularly without altering intracellular production of ROS by mitochondria (Lambeth, 2004). To elucidate whether a paracrine ROS signal can be transduced to adjacent cells in pathological conditions, LPS-stimulated macrophages were added onto pulmonary microvascular endothelial cells (PMVECs; Fig. 1 B) that had been previously loaded with the i indicator dye Fluo-4 (Fig. 1 C). Application of LPS-activated macrophages evoked a i rise in PMVECs that was attenuated by DPI pretreatment (Fig. 1 C). To exclude the contribution of autocrine extracellular ROS production, a similar experiment was performed using HepG2 parenchymal cells, as these cells generate minimal O2.– (Kikuchi et al., 2000). HepG2 cells displayed an i elevation after LPS-stimulated macrophage exposure, whereas no i transient was noted after application of nonstimulated macrophages (Fig. 1 D). In contrast, exposure of HepG2 cells to macrophages that had been stimulated by LPS plus DPI triggered only an extremely small i rise (Fig. 1 D). The oscillatory i transient pattern observed in individual HepG2 cells but not PMVECs is notable, indicating a potential difference in Ca2  handling between cell types (unpublished data). Overall, this result suggests that O2.– is specifically required for elevation of i in endothelial cells.

Figure 1. Endothelial cell Ca2  mobilization in response to activated macrophage-derived ROS. (A) J774A.1 murine macrophages were activated by 1 μg/ml LPS for 3 h in the presence or absence of the flavoprotein inhibitor DPI for 1 h. Cells were incubated with 10 μM of the ROS-sensitive dye H2DCF-DA and visualized using confocal microscopy. (B) Macrophages (M) were applied to PMVECs (E) to assess paracrine O2.– signaling. Ca2  indicator Fluo-4/AM–loaded PMVECs (C; n = 3) and HepG2 cells (D) were exposed to nonactivated, LPS-treated, and LPS DPI-treated macrophages Fluo-4 fluorescence change was recorded every 3 s for 10 min (n = 2). Ca2  mobilization was measured as described in Materials and methods.

O2.– evokes endothelial Ca2  transients through InsP3 signaling, b" t. h! I) p3 K0 O0 s) F

To identify the mechanisms by which O2.– triggers i signals in PMVECs, we extended our studies to examine the effects of O2.– on basal i. To exclude the possible contribution of other macrophage factors, the xanthine xanthine oxidase (X XO) system was used to generate O2.– externally. Cells exposed to O2.– demonstrated a rapid increase in i followed by a slightly delayed return to baseline (Fig. 2 A). Similarly, the physiological stimulus ATP generated a marked i transient (Fig. S1, available at http://www.jcb.org/cgi/content/full/jcb.200505022/DC1). The O2.–-evoked i increase was abolished by pretreatment with the XO inhibitor allopurinol (1 mM; Fig. 2 B) or by a combination of the antioxidant superoxide dismutase (SOD; 2000 U/ml) and the anion channel blocker DIDS (100 μM; Fig. 2 C). Treatment with either xanthine or allopurinol did not alter basal i in control cells (unpublished data). These findings suggest that acute exposure of PMVECs to extracellular O2.– results in a rapid i rise. We next sought to determine the source of the elevated i. Thapsigargin (Tg) inhibits the SERCA Ca2 ATPase, causing Ca2  depletion from the ER (Ma et al., 2000, 2001). Pretreatment with 2 μM Tg virtually abolished O2.–-induced Ca2  transients (Fig. 2 D). Conversely, removal of Ca2  from the external medium was without effect on i (Fig. 2 E). Together, these results indicate that O2.– induces a release of Ca2  from internal stores. ER Ca2  stores in endothelial cells can be modulated by production of the second messenger InsP3 by PLC and subsequent binding to receptors on the ER (InsP3R). To characterize the release of Ca2  from intracellular stores, PMVECs were pretreated for 10 min with either the PLC inhibitor U-73122 or its inactive analogue U-73343. U-73122, but not U-73343 (both 100 μM), inhibited the O2.–-induced Ca2  release (Fig. 2, F and G). This result suggests that the O2.–-induced i transient was mediated by InsP3. To further characterize O2.–-induced Ca2  release, cells were incubated with 2-aminoethoxydiphenyl borate (2-APB; 75 μM) before O2.– stimulation. 2-APB has widely been used as an inhibitor of InsP3-sensitive Ca2  release and store-operated Ca2  channels in intact cells (Ma et al., 2001; Bootman et al., 2002). In agreement with our PLC data, O2.–-induced Ca2  transients were abolished in cells pretreated with 2-APB (Fig. 2, H and I). Thus, the O2.–-induced i rise in PMVECs was due to the InsP3-dependent release of Ca2  from internal stores.4 p8 z7 C) x7 ]5 e: J1 N2 n* U' }* @7 }

Figure 2. Extracellular O2.– induces i mobilization through an InsP3-dependent pathway. (A) Fluo-4/AM–loaded PMVECs exposed to the O2.–-generating system (100 μM X  5 mU/ml XO) display a Ca2  transient (n = 15). (B and C) Inhibition of XO by 1 mM allopurinol and combination scavenge (2000 U/ml SOD) and entry inhibition (100 μM DIDS) attenuated Ca2  mobilization (n = 3). Intracellular store depletion (2 μM Tg; D; n = 4) but not extracellular Ca2  chelation (10 mM EGTA; E) prevented O2.–-evoked i mobilization (n = 3). (F and G) The PLC inhibitor U-73122 (100 μM) abolishes the i rise, whereas the analogue U-73343 (100 μM) fails to inhibit the O2.– effect (n = 4). (H) Pretreatment with 75 μM of the InsP3R antagonist 2-APB eliminated the O2.– effect (n = 4). (I) Relative Fluo-4 fluorescence change was quantified. Data are means ± SEM.
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O2.–-triggered i release is abolished in InsP3R triple knockout (TKO) cells
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To examine the specific role of InsP3R in the O2.–-triggered  rise, the InsP3R-deficient DT40 chicken B-lymphocyte cell line (DT40 InsP3R TKO) was used. Wild-type cells demonstrated a significant i increase after O2.– exposure. After i returned to basal levels, 2 μM Tg was added to the medium to induce a transient increase in i as a consequence of passive depletion of endogenous stores upon ER Ca2 /Mg2 -ATPase blockade (Fig. 3, A and B). Similar to PMVECs, pretreatment with 2 μM Tg eliminated the O2.–-induced Ca2  transients in wild-type DT40 cells (unpublished data). In contrast, addition of a O2.– pulse failed to elicit Ca2  release from intracellular stores in DT40 InsP3R TKO cells, whereas subsequent addition of 2 μM Tg triggered a complete depletion of Ca2  stores (Fig. 3, A and B). These data suggest that Ca2  release through the InsP3R underlies the O2.–-evoked rise of i. To confirm that DT40 InsP3R TKO cells retain the machinery necessary for the O2.–-mediated i transient, we transfected the rat InsP3R type I into TKO cells. This procedure restored the responsiveness of TKO cells to O2.– (Fig. 3 C). This result indicates that in TKO cells, a O2.–-mediated signal activates InsP3R type I and causes Ca2  release from ER store.

Figure 3. Elimination of O2.–-evoked i mobilization in InsP3R TKO cells. (A) Fluo-4/AM–loaded DT40 chicken B lymphocytes display a normal Ca2  response to extracellular O2.– (100 μM X 5 mU/ml XO; n = 7). In DT40 InsP3R TKO cells, O2.– failed to elicit i rise, whereas the subsequent addition of 2 μM Tg caused a large store depletion (n = 3). O2.–-evoked Ca2  mobilization is not dependent on PLC-2–mediated InsP3 production (n = 3). (B) Quantitation of i transient in wild-type DT40, TKO, and PLC-2 KO cells. (C) Measurement of i transient in DT40 InsP3R TKOs after reexpression of InsP3R Type I (n = 4). 38 out of 369 cells displayed i transient after O2.– exposure, as demonstrated by the single cell tracing. (D) PLC-2 KO of DT40 cells pretreated with U-73122 but not U-73343 attenuated Ca2  release from stores after exposure to O2.– (n = 3).

In PMVECs, inhibition of PLC with U-73122 prevented the rise of i induced by exposure to O2.–. We therefore further investigated the role of PLC in O2.–-triggered Ca2  mobilization using PLC-2–deficient DT40 cells. O2.– exposure triggered a substantial rise of i in PLC-2–deficient DT40 cells (Fig. 3, A and B). In wild-type DT40 cells, B cell receptor agonist IgM (2 μg/ml) induced a series of rapid i oscillations representing i release and reuptake. In contrast, anti-IgM failed to elicit Ca2  mobilization in both InsP3R TKO and PLC-2 knockout (KO) cells (unpublished data). These data indicate that the nonreceptor tyrosine kinase–linked cascade, to which PLC-2 is coupled, is dispensable for the O2.–-triggered i rise. In agreement with our findings, G protein–coupled receptor (GPCR)–mediated Ca2  oscillations were previously abolished by U-73122, which inhibits all PLC-? isoforms (Zeng et al., 2003). To further understand the role of InsP3, PLC-2 KO cells were pretreated with either PLC inhibitor U-73122 or U-73343 as described in Fig. 2 (F and G). U-73122, but not U-73433, attenuated the O2.–-evoked i rise (Fig. 3 D). To ensure that the O2.– elicits InsP3 accumulation, InsP3 was assessed in wild-type DT40, DT40 InsP3R TKO, and DT40 PLC-2 KO cells. Direct measurement of InsP3 production indicated that O2.– markedly activated InsP3 formation in wild-type DT40, DT40 InsP3R TKO, and DT40 PLC-2 KO cells. In contrast, pretreatment of DT40 PLC-2 KO cells with U-73122 significantly attenuated this response (Fig. S2 A, available at http://www.jcb.org/cgi/content/full/jcb.200505022/DC1). Similarly, PMVECs exposed to O2.– exhibited markedly greater InsP3 production than the physiological stimulus ATP (Fig. S2 B). Collectively, these findings suggest that extracellular O2.– causes Ca2  release via a PLC-mediated increase in InsP3.( \9 v' D* p. ^7 V* Y7 U. C

O2.– mediates coupling of i elevation and mitochondrial uptake8 B3 A0 P% |+ ]7 d2 ^. m; O
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It is believed that agonist-induced i rise can be buffered by mitochondria (Bernardi and Petronilli, 1996). To determine if the O2.–-triggered i spike is delivered to mitochondria, rhod-2– (mitochondrial Ca2  indicator) and Fluo-4–loaded PMVECs were subjected to O2.–. Exposure of PMVECs to O2.– induced an i increase as evidenced by an increase in Fluo-4 fluorescence, as shown earlier (Fig. 2 A), followed by an elevation of mitochondrial Ca2  fluorescence (Fig. 4, A and B). Similarly, ATP induced a i rise followed by mitochondrial  elevation (Fig. 4, C and D). These results indicate Ca2  signal propagation from the cytosol to the mitochondria in both physiological (purinergic receptor agonist) and pathological conditions (oxidative stress). Notably, O2.–-evoked mitochondrial Ca2  elevation was increased and sustained compared with the transient pattern observed in response to ATP. These results suggest that O2.–-induced intracellular pool Ca2  release evokes elevated mitochondrial Ca2  uptake during oxidative stress.
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Figure 4. Coupling of O2.–-evoked cytosolic Ca2  elevation and mitochondrial Ca2  signaling in endothelial cells. Simultaneous imaging of O2.– (100 μM X  5 mU/ml XO) or ATP (100 μM) induced changes in cytosolic and mitochondrial Ca2  using Fluo-4/AM and compartmentalized rhod-2/AM. (A and C, left) Images show the i response to addition of O2.– and ATP. (A and C, right) Confocal images of endothelial cells loaded with the mitochondrial Ca2  indicator rhod-2 display Ca2  accumulation (n = 4). (B and D) Synchronized measurements of O2.–- or ATP-induced changes in cytosolic and mitochondrial Ca2 . O2.–, but not ATP, triggered cytosolic Ca2  mobilization and sustained mitochondrial Ca2  elevation. The experimental data indicate that cytosolic Ca2  elevation precedes mitochondrial Ca2  uptake.

O2.–-induced Ca2  transients evoke rapid mitochondrial depolarization* J7 L. a, h# u/ }  u* m

Reversible depolarization of m occurs as a consequence of electrogenic uptake of Ca2  by mitochondria in response to transient i (Duchen, 1992). However, ROS may also promote MPTP opening (Huser et al., 1998). Because mitochondrial Ca2  elevation is a common pathway in both normal physiological and pathological stimuli, we examined whether the observed mitochondrial Ca2  uptake after O2.– exposure is associated with mitochondrial depolarization. Simultaneous fluorescence measurements of i and m were conducted in PMVECs during O2.– exposure (Fig. 5, A and B). In response to ATP, an i rise was observed similar to that in cells after O2.– exposure. However, in contrast to O2.–, PMVECs exposed to ATP exhibited only a nominal change in m (Fig. 5 C), possibly due to transient Ca2  uptake (Fig. 4, B and D). Application of O2.– evoked a rapid and transient rise in i that preceded a decrease in tetramethylrhodamine, ethyl ester, perchlorate (TMRE) fluorescence, indicating that mitochondrial depolarization is associated with the onset of the i rise (Fig. 5 B). Because O2.– is rapidly dismutated into H2O2, we sought to determine which oxidants are involved in the observed m loss. Cells incubated with H2O2 (1 mM) displayed no rapid i transient. Rather, H2O2 induced a slight increase in i (Fig. 5 D) and a delayed loss of m. Tg pretreatment did not affect the H2O2-facilitated slow i rise (unpublished data). These findings suggest that H2O2 may not affect the intracellular store, but instead facilitates Ca2  entry from the extracellular milieu independent of mitochondrial depolarization. Oxidized phospholipid byproducts are involved in cell death during oxidative stress (Ran et al., 2004). However, the lipid-oxidizing agent t-butyl hydroperoxide (t-BuOOH; 200 μM) did not evoke either an i rise or m loss (Fig. 5 E). This finding suggests the selective role of O2.–, and not other oxidants, in eliciting an i rise and mitochondrial depolarization.& @/ R# L' K. q" N
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Figure 5. Selective O2.– induction of Ca2  signaling evokes mitochondrial depolarization. (A) PMVECs loaded with Fluo-4/AM (30 min) and stained with TMRE (15 min) were exposed to O2.– as indicated (n = 8). (B) Relative brightness of Fluo-4 fluorescence and punctate–diffuse index of TMRE was calculated and plotted over time. (C) Change in i and m in response to 100 μM ATP (n = 3). i level and mitochondrial m were recorded in response to 1 mM H2O2 (D; n = 4) and 200 μM t-BuOOH (E; n = 4).! \" K8 D* P0 [

Extracellular O2.–-mediated signaling functions independent of mitochondrially derived ROS

Evidence indicates that external ROS may evoke mitochondrial O2.– production (Zorov et al., 2000; Aon et al., 2003). Because the O2.–-evoked i rise is a prerequisite for m loss, we aimed to exclude the involvement of intracellular ROS production by mitochondrial electron transport proteins in m loss. Antimycin A inhibits the normal electron flow through complex III, but triggers O2.– production through the accumulation of ubisemiquinone. Antimycin A triggered an immediate m loss without an apparent change in i (Fig. 6 A). Rotenone inhibits electron transfer from complex I (NADH dehydrogenase) to ubiquinone and diminishes O2.– production from complex III (Turrens et al., 1985). In contrast to antimycin A, rotenone affected neither i nor m. However, subsequent addition of O2.– triggered an i rise followed by m loss (Fig. 6 B). Oligomycin, which inhibits the mitochondrial FoF1-ATPase by binding to ATP synthase, was used to exclude possible mitochondrial ATP-dependent ROS production. Treatment with oligomycin failed to trigger either i mobilization or m loss. Subsequent addition of O2.– established both events (Fig. 6 C). This result indicates that complex III is the major site of mitochondrial ROS production during oxidative stress. It has been reported that mitochondrial Ca2  uptake requires an intact m and that dissipation by a mitochondrial uncoupler abolishes mitochondrial Ca2  uptake and delays i clearance (Boitier et al., 1999). Close examination of PMVECs exposed to the mitochondrial uncoupler FCCP revealed that a rapid m loss was associated with i elevation (Fig. 6 D). This i rise most likely reflects Ca2  release from the mitochondria as a consequence of mitochondrial depolarization. Surprisingly, subsequent application of O2.– evoked a transient rise in cytosolic Fluo-4 fluorescence followed by a rapid recovery of m. The m recovered after O2.–treatment is almost identical to the initial potential observed before FCCP addition. Collectively, these results suggest that the mitochondrial ROS-evoked m loss is independent of InsP3R-linked m changes by O2.–.
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Figure 6. Contribution of mitochondrially derived ROS on the Ca2  transient and mitochondrial depolarization. (A) 20 μM antimycin A immediately induced m without an apparent elevation of i. m was further exaggerated by subsequent application of O2.– in Fluo-4/AM– and TMRE-loaded PMVECs (n = 4). (B) 20 μM rotenone failed to demonstrate either a Ca2  transient or mitochondrial depolarization. Subsequent addition of O2.– facilitated i mobilization followed by m (n = 3). (C) 10 μg/ml oligomycin did not affect either i levels or m, and successive addition of O2.– facilitated i pool depletion and m (n = 3). (D) Exposure to 3 μM of the mitochondrial uncoupler FCCP before addition of O2.– caused a rapid m dissipation and i elevation that was reestablished by O2.– (n = 4).! Y- T& R9 `" ^6 ?
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Ca2  buffering protects against O2.–-triggered mitochondrial depolarization
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To assess whether the O2.–-induced rise of i is required for the O2.–-evoked m loss, PMVECs were loaded with the Ca2  chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetate (BAPTA) by incubation with the permeant acetoxymethyl ester (25 μM for 30 min) before application of the O2.–. BAPTA loading significantly inhibited O2.–-induced m loss (Fig. 7, A and B). In contrast, the H2O2-induced m loss was unaffected by pretreatment with BAPTA (Fig. 7 C). These experimental data provide evidence that m loss induced specifically by O2.– requires a rise of i. Other oxidants such as H2O2 are deleterious to mitochondrial function but appear to affect m through a Ca2 -independent pathway.

Figure 7. Buffering of O2.–-evoked i elevation with BAPTA prevents Ca2 -induced m in PMVECs. (A and B) O2.–-induced i elevation and m was prevented by pretreatment of cells with 25 μM of the membrane-permeable Ca2  chelator (BAPTA-AM; n = 4). (C) Chelation of intracellular Ca2  using BAPTA failed to attenuate H2O2-induced m (n = 4).: I2 {% o0 U8 e( M3 a, T$ G% x

O2.–-mediated signaling triggers caspase activation- v% D% j  i& r5 q: w3 D+ b2 B6 r
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Caspase cysteine proteases augment mitochondrial dysfunction by both activating proapoptotic Bcl-2 family proteins such as Bax, Bak, and Bid and inactivating antiapoptotic proteins such as Bcl-2 (Wei et al., 2001). To determine the dose and time course of receptor-mediated and mitochondrially dependent caspase activation in PMVECs after oxidant exposure, cytosolic extracts were collected after treatment with O2.–, H2O2, and t-BuOOH. Remarkably, when cells were exposed to O2.–, robust caspase-3 activity was observed in a dose-dependent manner (Fig. 8, A and D). Interestingly, even a low dose (1 mU X XO) was able to induce caspase-3 activity, indicating that O2.– may activate downstream caspases through a mitochondrially dependent pathway. Similarly, prominent caspase-9 activity was observed after O2.– treatment (Fig. 8, C and F). H2O2 elicited some caspase-3 and -9 activity, but at a level severalfold less than O2.–. In contrast, t-BuOOH did not activate either caspase-3 or -9. During apoptotic conditions, caspase-8 can activate caspase-3 directly through an extrinsic pathway. As shown in Fig. 8 (B and E), treatment of PMVECs with O2.– induced caspase-8 activity that was sevenfold higher than control and other oxidants. Inhibition of m loss by i buffering prevented caspase-3 and -9 activation (Fig. 8, D and F). Collectively, these results provide evidence that O2.– activates both extrinsic and intrinsic caspase pathways.
6 I9 i& ?2 E* X  |( D( D
Figure 8. O2.– -dependent activation of caspases in PMVECs. Cells were exposed to various concentrations of O2.–, H2O2, and t-BuOOH. After 3 h of treatment, lysates were assessed for caspase-3 (n = 3; A), -8 (n = 3; B), and -9 (n = 3; C) activity. Time-dependent experiments were also performed for caspase-3 (n = 3; D), -8 (n = 3; E), and -9 (n = 3; F). Pretreatment with 25 μM BAPTA-AM for 30 min attenuated caspase-3, -8, and -9 (n = 3) activation in response to O2.– as indicated in D, E, and F. Control cells were treated with 25 μM BAPTA-AM alone. Data are means ± SEM.5 o$ a6 a4 y( }( }
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O2.–-evoked i overload executes the cell death machinery
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Our results reveal that O2.– stimulates i mobilization that triggers subsequent mitochondrial events, leading to caspase activation in PMVECs. To directly demonstrate that O2.– induces apoptosis, we treated PMVECs with various oxidants at different doses, and then stained them for the early apoptotic marker annexin V and the late stage apoptotic (or necrotic) marker propidium iodide (PI). Cells treated with O2.– for 5 h displayed positive annexin V staining with no detectable PI labeling, indicating cells in the early stages of apoptosis (Fig. 9 A). Strikingly, cells exposed to a high concentration of O2.– demonstrated a dose-dependent elevation of both apoptotic and necrotic cell death as displayed in Fig. 9 B. Cells treated with 500 μM H2O2 also revealed an apoptotic phenotype, although at a lower level than observed in response to O2.–. In contrast, t-BuOOH (200 μM) treatment primarily led to necrosis, as evidenced by positive annexin V and PI staining. Control conditions resulted in nominal levels of apoptotic- or necrotic-positive cells. Previously, our results provided evidence that buffering of O2.– evoked i rise by BAPTA-AM and markedly prevented PMVEC m loss. Therefore, we tested whether i buffering inhibits O2.–-induced apoptosis. BAPTA-AM pretreatment (25 μM) attenuated apoptosis in PMVECs (Fig. 9 C), providing evidence that O2.–-induced i elevation is essential for mitochondrially dependent apoptosis. Conversely, BAPTA-AM treatment was ineffective 20 min after application of the O2.– (unpublished data). DT40 B-cells lacking all forms of InsP3R display reduced apoptotic cell death in response to anti-IgM (Sugawara et al., 1997). Because O2.–-induced i elevation is ablated in DT40 InsP3R TKO cells, we next investigated apoptosis in DT40 cells. DT40 InsP3R TKO cells, but not wild-type cells, display increased resistance to apoptosis after O2.– application (Fig. S3, available at http://www.jcb.org/cgi/content/full/jcb.200505022/DC1). These results suggest that O2.– selectively alters ER Ca2  homeostasis resulting in caspase activation, which in turn leads to apoptosis.2 C% G& d9 J) ^* F4 h1 Q6 C4 \

Figure 9. O2.– signaling selectively evokes apoptosis in PMVECs. After a 5-h exposure to O2.–, H2O2, or t-BuOOH, cells were labeled with annexin V Alexa Fluor-488 conjugate and PI for 15 min. (A) Cells exposed to O2.– (X XO; 5 mU/ml; n = 3) demonstrated positive annexin V staining, indicating early apoptosis. H2O2 (500 μM; n = 3) treatment demonstrated considerably less positive annexin V staining. Cells treated with 200 μM t-BuOOH (n = 3) stained positive for both annexin V and PI, indicating membrane permeabilization and necrotic cell death. (B) Quantitation of annexin V– and PI-positive cells after exposure to ROS. (C) i chelation (BAPTA-AM; 25 μM) attenuated O2.–-induced cell death (n = 3). Data are means ± SEM.
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Discussion4 |9 A) L, Q$ a( M! @& S9 \7 Z
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The mechanisms that contribute to apoptosis during I/R injury remain unclear, but it is generally believed that the release and/or activation of various bioactive molecules, such as ROS (Zhao, 2004) and inflammatory cytokines (Haimovitz-Friedman et al., 1997), are responsible for cell death. During these conditions, xanthine (Malis and Bonventre, 1986) and NADPH oxidases play a key role in O2.– production (Wei et al., 1999) and trigger pathological signaling. Reperfusion of ischemic cells generates oxidative stress and alters mitochondrial function (Hausenloy et al., 2004). Coordination of mitochondrial function during injury is an essential component of cell physiology and survival, yet little is known about the factors that contribute to cell death during oxidative stress. This study demonstrates that O2.– facilitates a transient i elevation followed by mitochondrial Ca2  uptake and depolarization that ultimately induces apoptotic cascades in endothelial cells.

Macrophage activation by endotoxin elicits O2.– generation via NADPH oxidase and autocrine production of ROS (Johnston et al., 1978). However, whether released O2.– has a potential paracrine signaling role in nearby cells is unknown. This study provides direct evidence that activated macrophages initiate an ROS-induced i elevation in adjacent cells. In comparison to the macrophage data, the observed i transient using the X XO was larger and less sustained. The enzymatic X XO system generates only O2.–, whereas activated macrophages may release other factors that could alter the amplitude of i in PMVECs. In addition, xanthine oxidase has been shown to interact with the vascular endothelium during inflammatory conditions (Houston et al., 1999). Because of the short half-life of the O2.– radical, close association between endothelial cells and the O2.– source may facilitate a greater response. A single pulse of O2.– evoked an i rise in PMVECs that caused m loss. These results suggest a potential mechanism by which macrophage-mediated oxidative stress perpetuates endothelial dysfunction. This O2.–-mediated response has several features. The i signals were observed in adherent PMVECs, HepG2, and DT40 suspension cell types, indicating a common mechanism in the cellular response to O2.–. The O2.–-evoked i signal was prevented by the combination of SOD and the anion channel blocker DIDS. The O2.–-induced transient rise of i was propagated to mitochondria, where a sustained Ca2  elevation was observed. In contrast, the i response to the physiological stimulus ATP triggered a transient mitochondrial Ca2  elevation. The O2.–-induced i transient subsequently evoked mitochondrial depolarization independent of mitochondrially derived ROS. In addition to this novel observation, our results suggest that O2.– selectively evokes Ca2 -dependent m loss independent of other oxidants.$ Z: y3 `0 i2 \
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Another important finding is that Tg, but not EGTA, pretreatment eliminated the O2.–-induced increase in i, indicating release from the ER. We therefore conclude that Ca2  store release in response to O2.– may be PLC dependent and mediated by InsP3R on the ER. This conclusion was supported by the observation that DT40 cells lacking all three InsP3R isoforms failed to show an i rise after O2.– application, unless InsP3R was reintroduced by transient transfection. Reintroduction of InsP3R type 1 restored the i transient, indicating the existence of the Ca2  signaling machinery in TKO cells. Furthermore, we found that the PLC inhibitor U-73122 blocked the O2.– response in endothelial cells. PLC normally presents as a key enzyme in cellular metabolism and signaling in response to extracellular agonists by coupling with GTP-binding proteins. DT40 cells express PLC-2 and PLC-? isoforms (Rhee, 2001) but lack the GPCRs necessary for PLC-? activation (Venkatachalam et al., 2001; Patterson et al., 2002). Surprisingly, we observed that PLC-2 KO cells displayed a rapid i store release in response to O2.–, suggesting the activation of PLC-?–mediated Ca2  release by O2.–. PLC inhibition in these PLC-2 KO cells by U-73122 indicates activation of PLC and suggests that O2.–-induced i rise requires InsP3. Because InsP3 levels were greatly elevated by O2.– in all three DT40 cell lines, it is apparent that generation of InsP3 by PLC is the essential signal in response to O2.– for InsP3R activation. Ca2  release via PLC-? (Liao et al., 1989) was investigated using the G protein–coupled muscarinic M5 receptor agonist carbachol. No detectable Ca2  signals were observed in response to carbachol (500 μM), indicating that DT40 cells lack the GPCR machinery necessary for PLC-? activation (unpublished data). However, we cannot exclude that O2.– may directly activate signaling upstream of PLC or regulate InsP3R. Earlier, we demonstrated the activation of mitochondrial PLA2 by O2.– (Madesh and Balasubramanian, 1997), lending support to our findings on the activation of signaling enzymes by O2.–.) g: L- ~1 h8 K6 U
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Our findings suggest that m loss in response to O2.– is dependent on ER stores and not extracellular Ca2 . However, it is unclear whether mitochondrially derived ROS exacerbate Ca2  release from ER stores during oxidative stress. Rotenone and other distal complex I inhibitors generate O2.– on the matrix side of the inner membrane (Brookes et al., 2004). Our data indicate that cells pretreated with rotenone alone did not trigger either i changes or a m change. In contrast, the complex III inhibitor antimycin A caused a sharp decline in the m without concomitant i mobilization. This finding suggests that O2.– generation by complex III directly facilitates m loss independent of i levels. Cell death can be initiated by mitochondrial inhibitors through a reduction in ATP levels in a process known as necrosis. Specifically, oligomycin is known to reduce available ATP through inhibition of mitochondrial FoF1-ATPase and to elicit cell death through a switch from apoptosis to necrosis. In our system, endothelial cells pretreated with oligomycin did not experience either a rapid i change or m decay. However, subsequent delivery of O2.– perturbed the ER Ca2  level and subsequent m loss. Experiments using the mitochondrial uncoupler FCCP indicate that mitochondrial Ca2  efflux precedes m dissipation. Apparently, mitochondrial depolarization evoked by paracrine O2.– differs from m alterations induced by mitochondrially derived ROS.
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The question arises whether extracellular O2.– generation evokes selective signaling during endothelial dysfunction. Previously, cells exposed to O2.– but not H2O2 elicited a rapid and large cytochrome c release from the mitochondria, followed by m loss (Madesh and Hajnoczky, 2001). Cell death has been associated with elevation of Ca2  through various means. Moreover, elevation of i has been implicated in the induction of apoptosis by ROS (Orrenius et al., 2003). It is suggested that H2O2 facilitates Ca2  entry from the extracellular milieu or from the intracellular pools (Zhao, 2004), and H2O2-induced apoptosis in I/R injury has also been proposed (Inserte et al., 2000). This study suggests that O2.–, but not H2O2, evoked an intracellular store Ca2  release that regulates the m. Strikingly, pretreatment with the i chelator BAPTA-AM prevents O2.–- but not H2O2-mediated endothelial m loss. Thus, the O2.–-initiated m loss is dependent on an i rise and independent of mitochondrial ROS generation. These findings suggest that extracellularly generated O2.– rapidly evokes the observed i elevation and pathological m loss. Interestingly, we illustrate that externally delivered O2.–, and not other oxidants, triggers a cytosolic signal that initiates the mitochondrial phase of apoptosis.

Mitochondrial membrane permeabilization evoked by apoptotic stimuli facilitate apoptogenic protein release from the intermembrane space and can lead to the downstream activation of both caspase-dependent and -independent apoptotic cascades. Our previous observation proposed that O2.–, but not H2O2, elicited cytochrome c release via a voltage-dependent anion channel–dependent mitochondrial membrane permeabilization (Madesh and Hajnoczky, 2001). Cytochrome c release is regulated by the Bcl-2 family of proteins, and the target of these proteins in the cell is the MPTP (Kroemer and Reed, 2000; Mattson and, Kroemer, 2003). This study shows the activation of initiator and effector caspases by O2.– specifically, and to some extent, by high doses of H2O2. Recent evidence has indicated that a caspase-3–truncated InsP3R type I may elicit a prolonged i elevation during apoptosis (Assefa et al., 2004). Our model indicates that caspase-3 activation is downstream of i elevation and m loss. However, we cannot rule out modification of InsP3R type I in the late stages of O2.–-triggered apoptosis. Collectively, these findings establish that ER Ca2  mobilization is upstream of mitochondrial events evoked by O2.– in endothelial apoptosis.! W7 J! L5 f# \6 P3 k. P
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In conclusion, activated macrophage-derived O2.– acts as an important signaling molecule that mediates InsP3R-linked i elevation and mitochondrial dysfunction in endothelial cells and provides a novel signaling link between inflammatory and endothelial cells under pathological conditions. We therefore propose that paracrine O2.– signaling is critical to endothelial cell death.! Y. k2 E% W& |' s
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Materials and methods# }/ a9 \9 I2 B& ]

Cell culture

Primary rat PMVECs (provided by T. Stevens, University of South Alabama, Mobile, AL) were cultured in DME supplemented with 10% FBS, nonessential amino acids, and antibiotics. Cells of wild-type DT40 chicken B cell line, triple InsP3R KO cell line (DT40 InsP3R KO), and PLC-2 KO (provided by A. August, Pennsylvania State University, Philadelphia, PA) cell line were cultured in RPMI 1640 supplemented with 10% FCS, 1% chicken serum, 50 μM 2-mercaptoethanol, 4 mM L-glutamine, and antibiotics. J774A.1 monocyte-derived mouse macrophages were cultured in Hank's F12 (supplemented with 10% FBS) and antibiotics. Heptocellular carcinoma cell line (HepG2) was cultured in MEM with 10% FBS, 2 mM L-glutamine, 0.50 mM sodium pyruvate, 0.1 mM nonessential amino acids, and antibiotics. Cells between passages 5 and 10 were used for experiments.

Visualization of ROS generation" K- u" ~- A9 \
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J774.1 mouse monocyte-derived macrophages (106 cells/ml) were cultured on glass bottom 35-mm dishes (Harvard Apparatus) for 48 h. Cells were challenged with 1 μg/ml LPS for 3 h at 37°C. For DPI treatment, 2.5 h LPS-treated macrophages were incubated with 30 μM DPI for 30 min. The oxidation-sensitive dye H2DCF-DA (10 μM; Invitrogen) was added separately to dishes 20 min before visualization under confocal microscopy. Macrophage cells treated under similar conditions were used for co-culture model Ca2  mobilization.
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i measurement

Measurement of i changes was performed using the Ca2 -sensitive fluorescent dye Fluo-4/AM (Invitrogen). Cells adherent to 25-mm-diam glass coverslips were incubated at RT in extracellular membrane (ECM) containing 5 μM Fluo-4/AM for 30 min, followed by an additional 10-min incubation in a dye-free medium. Coverslips were affixed to a chamber and mounted in a PDMI-2 open perfusion microincubator (Harvard Apparatus) and maintained at 37°C on an inverted microscope (model TE300; Nikon). Confocal imaging was performed using the Radiance 2000 imaging system (Bio-Rad Laboratories) equipped with a Kr/Ar-ion laser source at 488-nm excitation using a 60x oil objective. Images were collected using LaserSharp software (Bio-Rad Laboratories) every 3 s for i changes. Mobilization was induced by the application of 100 μM and 5 mU/ml, respectively, of the xanthine/xanthine oxidase O2.–-generating system. Whole cell masking was used to quantitate individual cell responses (Spectralyzer, custom software; provided by Paul Anderson, Thomas Jefferson University, Philadelphia, PA).

Measurement of inositol phosphates
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24 h before experiments, cells (106/ml) were transferred to myo-inositol–free DME and incubated in the presence of myo-inositol (2 μCi/ml; 20 Ci/mmol; MP Biomedical, Inc.). After washing with myo-inositol–free DME, cells were incubated for 30 min in myo-inositol–free DME supplemented with 10 mM LiCl and then exposed to either ATP (100 μM) or X XO (100 μM xanthine and 5 mU/ml XO) for 20 min at 37°C. The medium was subsequently removed and cells were scraped into 1 ml of 10% (wt/vol) TCA for the extraction of soluble inositol phosphates. After centrifugation of the cell lysates, the supernatant was applied to AG 1-X8 (formate form) ion exchange columns (200–400 mesh; Bio-Rad Laboratories). These columns were washed as previously described (Takata et al., 1995). Elution was performed with increasing concentrations of ammonium formate (0.1–0.7 M).7 ?5 f7 z0 @1 s5 P: V2 v& C

Simultaneous confocal imaging of cytosolic and mitochondrial Ca2  in PMVECs1 c8 E& @2 r; d

Endothelial cells were loaded with 2 μM rhod-2/AM in ECM containing 2.0% BSA in the presence of 0.003% pluronic acid at 37°C for 50 min. Cells loaded with rhod-2 dye were washed and then reloaded with Fluo-4/AM for an additional 30 min at RT. Cells were placed on a temperature-controlled stage and images were recorded using the Radiance 2000 imaging system with excitation at 488 and 568 nm for Fluo-4 and rhod-2, respectively.
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Kinetics of i elevation and mitochondrial membrane depolarization  s0 P0 M- n- v' ?3 S/ k0 o! V5 M
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Cells cultured on 25-mm-diam glass coverslips were loaded for 30 min with 5 μM Fluo-4/AM at RT. The cationic potentiometric fluorescent dye TMRE (100 nM) was added to the loading medium and allowed to equilibrate for at least 15 min. Under these conditions, TMRE fluorescence was largely localized to the mitochondrial matrix space. After dye loading, the cells were washed and resuspended in the experimental imaging solution (ECM containing 0.25% BSA). Intracellular esterase action then resulted in loading of both the cytoplasmic and mitochondrial compartments of the cell. Experiments were performed in ECM containing 0.25% BSA at 37°C. Images were recorded using the Radiance 2000 imaging system with excitation at 488 and 568 nm for Fluo-4 and TMRE, respectively. Fluo-4 and TMRE fluorescent changes were determined by background subtraction followed by masking of total cell area or intracellular regions. During m loss, the exit of TMRE from mitochondria into the cytoplasm leads to quenching of the dye. The rapid redistribution of TMRE into the cytoplasm after depolarization of m can be transiently detected in the nucleus.
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Detection of caspase-3, -8, and -9 activity" ~, w3 r/ @: f' `2 ]- H# r; ]

The assay is based on the ability of the active enzymes to cleave the fluorogenic substrates Ac-DEVD-AFC (caspase-3), Ac-IETD-AFC (caspase-8), or Ac-LEHD-AFC (caspase-9; Calbiochem). Cells treated with various oxidants were harvested via trypsinization and washed with PBS. The cell pellet was gently resuspended in lysis buffer (25 mM Hepes, pH 7.4, 2 mM EDTA, 0.1% CHAPS, 5 mM DTT, 1 mM PMSF, and protease inhibitor cocktail , lysed, and centrifuged; the supernatant was used as the assay. Caspase substrates were added to a final concentration of 50 μM and the samples were incubated at 37°C for 45 min in caspase assay buffer. Incubated samples were measured at an excitation of 400 nm and an emission of 505 nm in a multiwavelength-excitation dual wavelength-emission fluorimeter (Delta RAM; Photon Technology International).1 X  |' L+ [! \- t5 l5 ?

Confocal imaging analysis of apoptotic markers in PMVECs
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To determine cellular outcome in response to oxidative stress, cells were exposed to the O2.–-generating system, H2O2, and t-BuOOH for 5 h. To assess the externalization of phosphatidylserine in the plasma membrane, as occurs in the early stage of apoptosis, cells were incubated with the conjugate annexin V Alexa Fluor-488 (Invitrogen) and PI (0.5 μg/ml) for 15 min. After treatment, annexin V– and PI-stained cells were visualized and counted. In normal cells, impermeable PI is internalized as the plasma membrane loses integrity. Thus, positive PI staining indicates either late stage of apoptosis or necrosis.

Data analysis4 B, i$ o" t2 g5 S5 J
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Tracings are representative of the mean fluorescence value of all cells in one field and are indicative of n independent experiments. Data given are representative of duplicate analysis of n independent experiments as mean ± SEM.$ v: `: E6 X8 P9 N, D0 x+ |: D
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Online supplemental material

Fig. S1 shows the Ca2  response to the physiological and pathological stimuli ATP and O2.–, respectively, in PMVECs. Fig. S2 details the measurement of InsP3 generation in both DT40 and PMVECs. Fig. S3 shows the analysis of apoptosis in DT40 cells in response to O2.–. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200505022/DC1.8 \7 z- W9 I  I0 [; {. D8 D* L5 y
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Acknowledgments8 M3 z5 A: t( }4 ]! u
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We thank Drs. Troy Stevens and Avery August for providing endothelial and PLC-2 KO cells, respectively. We are grateful to Dr. Kevin Foskett for critical manuscript review. We also thank Drs. Craig Thompson and Sheldon Feinstein for helpful suggestions and Paul Anderson for Spectralyzer image analysis software.

This work was supported by startup funds from the Institute for Environmental Medicine to M. Madesh. A.B. Fisher is funded by National Institutes of Health (NIH) grant HL-60290. S.K. Joseph is supported by NIH grant DK-34804.* O5 L. Y* w9 @0 `' Z3 M( Z' i
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