Correspondence to Muniswamy Madesh: madeshm{at}mail.med.upenn.edu
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Introduction |
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In pathological conditions, cell death is facilitated by an elevation in intracellular calcium ([Ca2+]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 [Ca2+]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 [Ca2+]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 [Ca2+]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).
Endothelial cells subjected to oxidative stress undergo apoptosis (Warren et al., 2000). Although there is evidence that perturbations of cellular Ca2+ homeostasis (including [Ca2+]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 [Ca2+]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).
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 [Ca2+]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.
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Results |
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O2. mediates coupling of [Ca2+]i elevation and mitochondrial uptake
It is believed that agonist-induced [Ca2+]i rise can be buffered by mitochondria (Bernardi and Petronilli, 1996). To determine if the O2.-triggered [Ca2+]i spike is delivered to mitochondria, rhod-2 (mitochondrial Ca2+ indicator) and Fluo-4loaded PMVECs were subjected to O2.. Exposure of PMVECs to O2. induced an [Ca2+]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 [Ca2+]i rise followed by mitochondrial [Ca2+] 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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Discussion |
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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 [Ca2+]i elevation in adjacent cells. In comparison to the macrophage data, the observed [Ca2+]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 [Ca2+]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 [Ca2+]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 [Ca2+]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 [Ca2+]i signal was prevented by the combination of SOD and the anion channel blocker DIDS. The O2.-induced transient rise of [Ca2+]i was propagated to mitochondria, where a sustained Ca2+ elevation was observed. In contrast, the [Ca2+]i response to the physiological stimulus ATP triggered a transient mitochondrial Ca2+ elevation. The O2.-induced [Ca2+]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.
Another important finding is that Tg, but not EGTA, pretreatment eliminated the O2.-induced increase in [Ca2+]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 [Ca2+]i rise after O2. application, unless InsP3R was reintroduced by transient transfection. Reintroduction of InsP3R type 1 restored the [Ca2+]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-ß isoforms (Rhee, 2001) but lack the GPCRs necessary for PLC-ß activation (Venkatachalam et al., 2001; Patterson et al., 2002). Surprisingly, we observed that PLC-
2 KO cells displayed a rapid [Ca2+]i store release in response to O2., suggesting the activation of PLC-ßmediated Ca2+ release by O2.. PLC inhibition in these PLC-
2 KO cells by U-73122 indicates activation of PLC and suggests that O2.-induced [Ca2+]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-ß (Liao et al., 1989) was investigated using the G proteincoupled 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-ß 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..
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 [Ca2+]i changes or a
m change. In contrast, the complex III inhibitor antimycin A caused a sharp decline in the
m without concomitant [Ca2+]i mobilization. This finding suggests that O2. generation by complex III directly facilitates
m loss independent of [Ca2+]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 [Ca2+]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.
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 [Ca2+]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 [Ca2+]i chelator BAPTA-AM prevents O2.- but not H2O2-mediated endothelial
m loss. Thus, the O2.-initiated
m loss is dependent on an [Ca2+]i rise and independent of mitochondrial ROS generation. These findings suggest that extracellularly generated O2. rapidly evokes the observed [Ca2+]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 channeldependent 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-3truncated InsP3R type I may elicit a prolonged [Ca2+]i elevation during apoptosis (Assefa et al., 2004). Our model indicates that caspase-3 activation is downstream of [Ca2+]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.
In conclusion, activated macrophage-derived O2. acts as an important signaling molecule that mediates InsP3R-linked [Ca2+]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.
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Materials and methods |
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Visualization of ROS generation
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.
[Ca2+]i measurement
Measurement of [Ca2+]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 [Ca2+]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
24 h before experiments, cells (106/ml) were transferred to myo-inositolfree DME and incubated in the presence of myo-[2-3H]inositol (2 µCi/ml; 20 Ci/mmol; MP Biomedical, Inc.). After washing with myo-inositolfree DME, cells were incubated for 30 min in myo-inositolfree 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 (200400 mesh; Bio-Rad Laboratories). These columns were washed as previously described (Takata et al., 1995). Elution was performed with increasing concentrations of ammonium formate (0.10.7 M).
Simultaneous confocal imaging of cytosolic and mitochondrial Ca2+ in PMVECs
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.
Kinetics of [Ca2+]i elevation and mitochondrial membrane depolarization
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.
Detection of caspase-3, -8, and -9 activity
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 [Roche], 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).
Confocal imaging analysis of apoptotic markers in PMVECs
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 analysis
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.
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.
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Acknowledgments |
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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.
Submitted: 4 May 2005
Accepted: 16 August 2005
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