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Originally published In Press as doi:10.1074/jbc.M413674200 on January 24, 2005

J. Biol. Chem., Vol. 280, Issue 14, 13349-13354, April 8, 2005
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Hypoxic Modulation of Ca2+ Signaling in Human Venous Endothelial Cells

MULTIPLE ROLES FOR REACTIVE OXYGEN SPECIES*

Parvinder K. Aley{ddagger}, Karen E. Porter{ddagger}, John P. Boyle{ddagger}, Paul J. Kemp§, and Chris Peers{ddagger}

From the {ddagger}School of Medicine, University of Leeds, Leeds LS2 9JT and §School of Biosciences, University of Cardiff, Cardiff CF10 3US, United Kingdom

Received for publication, December 6, 2004


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The effects of hypoxia (pO2 ~25 mm Hg) on Ca2+ signaling stimulated by extracellular ATP in human saphenous vein endothelial cells were investigated using fluorimetric recordings from Fura-2 loaded cells. In the absence of extracellular Ca2+, ATP-evoked rises of cytosolic Ca2+ concentration ([Ca2+]i) because of mobilization from the endoplasmic reticulum (ER). These responses were reduced by prior exposure to hypoxia but potentiated during hypoxia. Hypoxia itself liberated Ca2+ from the ER, but unlike the effects of ATP this effect was not inhibited by blockade of the inositol trisphosphate receptor. By contrast, ryanodine blocked the effects of hypoxia but not those of ATP. Antioxidants abolished the effects of hypoxia but potentiated the effects of ATP. Inhibition of NADPH oxidase also augmented ATP-evoked responses but was without effect on hypoxia-evoked rises of [Ca2+]i. However, either uncoupling mitochondrial electron transport or inhibiting complex I markedly suppressed the actions of hypoxia yet exerted only small inhibitory effects on ATP-evoked rises of [Ca2+]i. Both hypoxia and ATP were able to activate capacitative Ca2+ entry. Our results indicate that hypoxia regulates intracellular Ca2+ signaling via two distinct pathways. First, it modulates agonist-evoked liberation of Ca2+ from the ER primarily through regulation of reactive oxygen species generation from NADPH oxidase. Second, it liberates Ca2+ from the ER via ryanodine receptors, an effect requiring mitochondrial reactive oxygen species generation. These findings suggest that local O2 tension is a major determinant of Ca2+ signaling in the vascular endothelium, a finding that is likely to be of both physiological and pathophysiological importance.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The vascular endothelium plays a central role in the control of vascular function, exerting important influences on vital functions as diverse as coagulation, inflammation, vessel permeability, angiogenesis, and vascular tone (reviewed by Refs. 1–4). Many of these functions, such as production of vasoactive agents (5, 6), rely on regulated changes of intracellular Ca2+ concentration ([Ca2+]i). As in other non-excitable cells, Ca2+ homeostasis in endothelial cells involves uptake and release of Ca2+ into intracellular organelles (particularly the endoplasmic reticulum (ER)1) as well as controlled influx from the extracellular environment (1, 2, 7–10). This process of Ca2+ influx is linked to depletion of intracellular stores such that store depletion triggers capacitative Ca2+ entry (7, 11–13), which has been linked specifically to the activation of nitricoxide synthase (6).

Although the vascular endothelium can be considered a syncytium, it clearly experiences different environments in different regions of the vasculature. The most striking difference is between arterial and venous environments; clearly, venous endothelial cells experience an environment which, as compared with those of the arterial vessels, is of much lower pressure and is also relatively hypoxic and hypercapnic. Of these parameters, we have focused on the effects of hypoxia on Ca2+ signaling in venous endothelial cells. This is a poorly studied area that deserves investigation for several reasons. First, most in vitro studies of Ca2+ signaling in endothelial (and other) cells have been conducted using perfusate equilibrated with room air (~150 mm Hg), which is hyperoxic even for the arterial endothelium. Second, where any effects of hypoxia have been studied, they have usually been studied in combination with other altered parameters such as glucose removal (to mimic ischemia/reperfusion conditions) and/or have been studied over very prolonged time courses (14–16). Third, whereas the effects of acute hypoxia on ion channel function have been studied in depth in a variety of cell types (17–19), their actions on other ion flux mechanisms, such as those responsible for Ca2+ homeostasis in non-excitable cells, are poorly understood. Finally, although local venous O2 levels clearly influence physiological functions (such as release of nitric oxide (20)), hypoxia may be important in the development of specific vascular disease states, such as varicoses. It has been proposed that hypoxic "activation" of endothelial cells (which can occur via blood stasis, particularly in leg veins) requires mobilization of Ca2+ and initiates a cascade of events leading to varicose formation (21, 22). Thus, we have investigated the effects of acute hypoxia on Ca2+ homeostasis in primary cultures of human saphenous vein endothelial cells and compared responses to those of the well characterized Ca2+ mobilizing agent, extracellular ATP (23).


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Isolation and Culture of Saphenous Vein Endothelial Cells—The isolation of primary cultures of saphenous vein endothelial cells was adapted from methods described previously (24). Saphenous vein samples were collected from patients undergoing coronary bypass grafting following local ethical permission and informed, written patient consent. Tissue from a total of 36 patients was used, 10 female (28%) and 26 male (72%). The age range was 44–79, the median age was 67, and the mean age was 65.7 ± 1.5 years. All patients were undergoing elective coronary artery bypass surgery, and any patients with potentially confounding conditions (e.g. diabetes) were excluded.

Individual samples ranging from 10–30 mm in length were opened longitudinally and pinned, lumen uppermost, onto silicone elastomer-coated 60-mm Petri dishes using A1 Minuten pins. The tissue sample was then incubated in 1 mg/ml Type II collagenase (Worthington) dissolved in Medium 199 (37 °C, 15 min). The collagenase solution was collected along with 2 x 10 ml of wash solution (minimal essential medium supplemented with 5% fetal calf serum and 1% antibiotic/antimycotic), which was used to detach any residual endothelial cells from the tissue. The suspension was centrifuged for 6 min at 600 x g, the supernatant was removed, and the pellet was resuspended in 25 ml of wash solution and recentrifuged. The supernatant was once again removed, and the final pellet was resuspended in 4 ml of complete endothelial culture medium (M199) supplemented with 20% fetal calf serum, 1% penicillin-streptomycin, 1% glucose, 1 M HEPES (Invitrogen), heparin (5 units/ml, Leo Laboratories), endothelial growth factor (15 µg/ml), and pyruvate (1 µM, Sigma-Aldrich, Poole, Dorset, UK). This mixture was then plated into a 25 cm2 flask and maintained in a humidified incubator at 37 °C (95% air, 5% CO2). 2 days following plating, cells received a full medium change to remove non-adherent cells. Culture medium was then half changed every 2–3 days, resulting in a confluent flask within 2–3 weeks. This was designated passage 0; cells were subcultured using trypsin and used for experiments up to passage 3.

Measurement of [Ca2+]i—Cells were plated onto glass coverslips in 24-well culture plates and grown to ~80% confluence. Coverslips onto which cells had grown were incubated in 2 ml of culture medium containing 4µM Fura-2AM (Molecular Probes, Cambridge, UK) for 40 min. at 37 °C in the dark and then left to de-esterify for 15 min in control solution. Fragments of coverslips were then transferred into an 80-µl recording chamber mounted on the stage of an inverted microscope where cells were continuously perfused under gravity at a rate of ~5 ml/min. Control perfusate was composed of: 135 mM NaCl, 5 mM KCl, 1.2 mM MgSO4, 2.5 mM CaCl2, 5 mM HEPES, and 10 mM glucose (pH 7.4, osmolarity adjusted to 300 mosM with sucrose, 21–24 °C). Solutions were made hypoxic where indicated by continuous bubbling with N2 for at least 30 min prior to perfusion of cells, which produced no shift in pH. pO2 was measured (at the cell) using a polarized carbon fiber microelectrode (25) and ranged from 20–25 mm Hg. Ca2+-free perfusate (used in all experiments except part of those in Fig. 6) contained 1 mM EGTA and no added Ca2+. [Ca2+]i was determined ratiometrically using an Improvision monochromator-based imaging system (Openlab Image Processing & Vision Company Ltd, Coventry, UK) with alternating excitation 340 and 380 nm (0.2 Hz), emission 510 nm. Regions of interest were used to restrict data collection to individual cells. All imaging was controlled by Improvision software, including Openlab 2.2.5. Drugs and agonists were applied to cells via the gravity-fed perfusion system, and all experiments were conducted at 21–24 °C. Fluorescence signals were calibrated according to the formulae of Ref. 26. Changes in [Ca2+]i were calculated by determining the rise of [Ca2+]i relative to basal levels measured immediately before that particular experimental maneuver. Where relevant, results are expressed as means ± S.E. together with example traces, and statistical comparisons were made using unpaired Student's t tests. All mean data were obtained from the number of individual cells indicated. In each case these were collected from cells of at least 3 (and usually 4–6) different patients.



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  		FIG. 6.
Hypoxia evokes capacitative Ca2+ entry. A, example recording of [Ca2+]i demonstrating a lack of effect of application of Ca2+ (2.5 mM) to the extracellular perfusate. B, superimposed traces taken from individual endothelial cells previously treated for 20 min with 1 µM thapsigargin. Recordings commenced during perfusion with Ca2+-free solution. For the period indicated by the solid horizontal bar Ca2+ (2.5 mM) was readmitted to the perfusate either in the absence or presence of the concentrations of Gd3+ indicated. C, as in B except that the cells were exposed to a range of concentrations of La3+, as indicated. D, three superimposed example traces illustrating the transient rise of [Ca2+]i evoked by ATP (applied for the period indicated by open bar) in the absence of extracellular Ca2+. Following the removal of ATP, cells were exposed to a perfusate containing 2.5 mM Ca2+ (as indicated by the solid bar) either in the absence (solid line) or presence of 1 mM Gd3+ (dashed line) or 10 µM La3+ (dotted line). E, as in D except that the cells were exposed to hypoxia rather than ATP for the period indicated by the open bar. Scale bars apply to all traces (A–E). F, bar graph illustrating the mean peak magnitude (with S.E. bar) of [Ca2+]i rise caused by Ca2+ entry in cells treated with thapsigargin (TG, 1 µM), ATP (10 µM), or hypoxia, as indicated, either in the absence or presence (+ Gd3+) of 1 mM Gd3+ or 10 µM La3+ (+ La3+). Values determined from between 10 and 42 cells in each case. p values indicate statistically significant differences.

 

   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Hypoxia Mobilizes Intracellular Ca2+ and Modulates ATP-evoked Ca2+ Signaling—Agonists evoke rises of [Ca2+]i in non-excitable cells via Ca2+ mobilization from intracellular stores and Ca2+ influx. To delineate these pathways, we examined the ability of ATP to evoke rises of [Ca2+]i while cells were perfused with a Ca2+-free solution (replaced with 1 mM EGTA) to prevent Ca2+ influx. Fig. 1A shows an example transient rise of [Ca2+]i caused by the application of 10 µM ATP, a well defined mobilizer of Ca2+i in endothelial cells, acting via P2Y receptors to generate inositol trisphosphate (IP3) (23). Exposure of cells to hypoxia (pO2 ~25mmHg) caused a small but discernible, transient rise of [Ca2+]i (Fig. 1B) in 96 of 113 cells (85%) examined. Whenever cells were exposed to acute hypoxia followed by ATP (10 µM) in a normoxic solution, this agonist was still able to elicit a transient rise of [Ca2+]i (e.g. Fig. 1B). However, responses were significantly smaller than those observed in cells not previously exposed to hypoxia (Fig. 1E). By contrast, exposure of cells to hypoxia after exposure to ATP consistently failed to evoke a rise of [Ca2+]i (Fig. 1C, representative of 29 recordings). Perhaps surprisingly, when ATP was applied during hypoxia responses were greater in magnitude (Fig. 1, D and E) than those observed in normoxia, despite the fact that during normoxia this concentration of ATP was supramaximal (data not shown). These findings are summarized in the bar chart of Fig. 1E.



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  		FIG. 1.
Hypoxia stimulates Ca2+ release from an intracellular, ATP-sensitive pool. A, example recording of [Ca2+]i from an endothelial cell during perfusion with a Ca2+-free solution. Note the transient rise evoked by ATP (10 µM). B, as in A, except the cell was first exposed to a hypoxic solution (pO2 ~25 mm Hg) before application of 10 µM ATP. Inset shows a magnified region indicated by dashed lines to illustrate the transient rise of [Ca2+]i caused by hypoxia. C, as in B, except that hypoxia was applied after the cell was exposed to 10 µM ATP. D, as in B, except that ATP was applied during continued exposure of cell to hypoxia. In each panel (A–D), the period of application of ATP is indicated by open bars, and exposure to hypoxia is indicated by solid bars. Ca2+-free perfusate was present throughout. Scale bars apply to all main panels. E, bar graph showing mean peak responses under the experimental conditions illustrated in A–D. Each bar is the mean with vertical S.E. bar taken from 29–100 cells. Significant differences are indicated by p values above bars.

 
Results presented in Fig. 1, A–C, suggested that hypoxia might stimulate release of Ca2+ from the same intracellular pool as ATP, presumably the ER. To investigate this, we pretreated cells for 20 min with the ER Ca2+-ATPase inhibitor thapsigargin (1 µM). In cells treated this way, neither ATP (n = 99) nor hypoxia (n = 29) evoked rises of [Ca2+]i during perfusion with Ca2+-free solution (e.g. Fig. 2A). We also investigated the effects of 2-aminoethoxydiphenyl borate (2-APB), originally considered an antagonist of IP3 receptors (27) but now recognized as a modulator of Ca2+ signaling via additional mechanisms (e.g. Ref. 28). Bath application of 500 µM 2-APB in Ca2+-free perfusate caused a rise of [Ca2+]i, but consistent with its ability to inhibit IP3 receptors, it fully blocked rises of [Ca2+]i evoked by ATP (Fig. 2, B and C). In contrast, hypoxia was still able to evoke rises of [Ca2+]i because it appeared additive with the 2-APB responses (Fig. 2, B and C). Thus, hypoxia appeared to mobilize Ca2+ from the endothelial ER via a mechanism that did not involve IP3 receptors. The involvement of ryanodine receptors (RyRs) was investigated by exposing cells to either ATP or hypoxia in the continued presence of 100 µM ryanodine. As illustrated in Fig. 2C (upper trace), responses to ATP were unaffected (representative of 12 cells), but hypoxic responses were significantly attenuated (Fig. 2, C (lower trace) and E). Responses to hypoxia were also markedly suppressed by pre-exposure for 20 min to the cyclic ADP-ribose (cADPR) antagonist, 8-Br-cADPR (29) (30 µM, Fig. 2, D (upper trace) and E) or by pre-exposure for 30 min to nicotinamide, which prevents cADPR formation (30). Collectively, these data indicate that hypoxia evokes Ca2+ release from the ER via cADPR-mediated activation of RyRs.



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  		FIG. 2.
Hypoxia and ATP mobilize intracellular Ca2+ via different mechanisms. A, two separate recordings of [Ca2+]i made in cells pretreated for 20 min with 1 µM thapsigargin. Cells were exposed to 10 µM ATP (upper trace) or hypoxia (lower trace) for the periods indicated by the horizontal bars. Scale bars apply to both traces. B, example traces indicating that exposure of cells to 2-APB (for the period indicated by the horizontal bar) alone (control) evoked a rise of [Ca2+]i. This was unaffected by 10 µM ATP (+ ATP) but enhanced by hypoxia (+ hypoxia). C, upper trace, ATP-evoked rise of [Ca2+]i during exposure to ryanodine (agents were applied for the periods indicated by horizontal bars. Lower trace, lack of ability of hypoxia to alter [Ca2+]i during exposure to ryanodine. Scale bars apply to both traces. D, illustrative lack of ability of hypoxia to evoke rises of [Ca2+]i in cells pretreated with either 30 µM 8-Br-cADPR (upper trace) or 6 mM nicotinamide (lower trace). Scale bars apply to both traces. In all traces (A–D), Ca2+ was omitted from the extracellular solution. E, mean (±S.E.) effects of hypoxia alone or in the presence of ryanodine, 8-Br-cADPR, or nicotinamide, as indicated. Values are taken from between 23 and 60 cells. p values indicate statistically significant differences.

 
Reactive Oxygen Species (ROS) Differentially Modulate Ca2+ Signaling Evoked by Hypoxia and ATP—ROS modulate Ca2+ signaling in endothelial cells (31, 32). Much evidence suggests that mitochondria provide one important intracellular source of ROS, and recent evidence suggests that ROS production is modulated by hypoxia (33, 34). Therefore we explored the possibility of ROS acting as a signal between mitochondria and the ER by investigating first the ability of antioxidants to modulate Ca2+ signaling in endothelial cells. To do this, we employed two mechanistically distinct antioxidants: trolox (6-hydroxy-2,5,7,8-tetramethylchroman 2-carboxylic acid, a cell-permeable, water-soluble derivative of vitamin E (see Ref. 35)) and TEMPO (4-hydroxy-2,2,6,6-tetramethylpiperidine 1-oxyl), applied together with catalase. TEMPO catalyzes the conversion of superoxide to H2O2, which is then removed by catalase (see Ref. 36). Cells were pretreated with 500 µM trolox for 30 min or 500 µM TEMPO plus 250 units/ml catalase for 45 min before exposure to either hypoxia or ATP. The antioxidants were also present during recordings. Trolox itself had no effect on basal [Ca2+]i, but TEMPO/catalase caused a small, significant increase (p < 0.01) of ~30% (data not shown). Fig. 3A (the mean data are plotted in Fig. 3C) indicates that ATP-evoked rises of [Ca2+]i measured in Ca2+-free perfusate, were significantly enhanced in the presence of antioxidants. However, hypoxiaevoked rises (again measured in Ca2+-free perfusate) were completely inhibited, and indeed small decreases of [Ca2+]i were evident (Fig. 3, B and D). These data strongly suggest an important role for ROS in endothelial Ca2+ signaling during both normoxia and hypoxia and further support the idea that the mechanisms underlying ATP- and hypoxia-evoked release of Ca2+ from the ER are distinct.



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  		FIG. 3.
A, three superimposed example changes in cytosolic [Ca2+]ievoked by 10 µM ATP (applied for the period indicated by the open bar) in the absence of antioxidant (control, solid line) or in the presence of trolox (Tr, dotted line) or TEMPO with catalase (T/C, dashed line). B, as in A except that cells were exposed to hypoxia rather than ATP. Ca2+-free perfusate was employed in all experiments (A and B). C, mean (±S.E.) changes in [Ca2+]i evoked by 10 µM ATP in the absence of antioxidant (control, solid bar, n = 88) or in the presence of trolox (n = 10) or TEMPO with catalase (n = 21). D, mean (±S.E.) changes in [Ca2+]i evoked by hypoxia in the absence of antioxidant (control, solid bar, n = 53) or in the presence of trolox (n = 8) or TEMPO with catalase (n = 18). p values indicate statistically significant differences.

 
Distinct Sources of ROS Differentially Modulate Ca2+ Signaling—ROS are produced at various distinct sites within cells, and of these mitochondria and NADPH oxidase are of particular importance in endothelial cells (31, 32, 37). To investigate NADPH oxidase as a ROS source, we examined the effects of hypoxia and ATP during inhibition of the oxidase by two distinct pharmacological inhibitors. Again, these experiments were conducted during perfusion of cells with Ca2+-free solution. Both phenylarsine oxide (2 µM) and diphenylene iodonium (5 µM) significantly augmented the magnitude of Ca2+ transients evoked by ATP (Fig. 4, A and B; mean data plotted in E). In contrast, neither agent significantly altered rises of [Ca2+]i evoked by hypoxia (Fig. 4, B and D; mean data plotted in F). It is also noteworthy that both agents produced a small but sustained rise of basal Ca2+ levels (seen most clearly in Fig. 4, C and D).



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  		FIG. 4.
NADPH oxidase inhibitors augment ATP-evoked Ca2+ signaling. A and B, example changes in cytosolic [Ca2+]i evoked by 10 µM ATP in the presence of the NADPH oxidase inhibitors phenylarsine oxide (PAO) or diphenylene iodonium (DPI). ATP was applied for the periods indicated by the solid bars; oxidase inhibitor applications are indicated by open bars. C and D, as A and B except that cells were exposed to hypoxia rather than ATP for the periods indicated by the solid bars. E, mean (±S.E.) changes in [Ca2+]i evoked by ATP in the absence of oxidase inhibitor (ATP, solid bar, n = 88) or in the presence of phenylarsine oxide (n = 23) or diphenylene iodonium (n = 13). F, mean (±S.E.) changes in [Ca2+]i evoked by hypoxia in the absence of oxidase inhibitor (hypoxia, solid bar, n = 53) or in the presence of phenylarsine oxide (n = 19) or DPI (n = 17). p values indicate statistically significant differences.

 
To investigate a role for mitochondria in mediating Ca2+ signaling, we exposed cells either to the mitochondrial uncoupler FCCP (10 µM applied with 2.5 µg/ml oligomycin to prevent ATP consumption by the F1F0-ATP synthase) or to the complex I inhibitor, rotenone (2 µM), which inhibits hypoxia-evoked increases in ROS production in umbilical vein endothelial cells (38). Both agents caused transient rises of [Ca2+]i, presumably caused by loss of Ca2+ from the mitochondria (Fig. 5, A–D). Subsequent exposure to ATP evoked rises of [Ca2+]i that were slightly smaller than those observed when mitochondria were functional (Fig. 5, A, B, and E). In contrast, the rises of [Ca2+]i evoked by hypoxia were strikingly suppressed (Fig. 5, C, D, and F). In particular FCCP/oligomycin inhibited hypoxic responses by ~75% (Fig. 5, C and F).



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  		FIG. 5.
Modulation of Ca2+ signaling by mitochondrial inhibition. A and B, example changes in cytosolic [Ca2+] evoked by 10 µM ATP in the presence of the FCCP and oligomycin (A) or rotenone (B). ATP was applied for the periods indicated by the solid bars; mitochondrial inhibitor applications are indicated by open bars. C and D, as A and B except that cells were exposed to hypoxia rather than ATP for the periods indicated by the solid bars. E, mean (±S.E.) changes in [Ca2+]i evoked by ATP in the absence of mitochondrial inhibitor (ATP, solid bar, n = 88) or in the presence of FCCP and oligomycin (n = 37) or rotenone (n = 26). F, mean (±S.E.) changes in [Ca2+]i evoked by hypoxia in the absence of mitochondrial inhibitor (hypoxia, solid bar, n = 53) or in the presence of FCCP and oligomycin (n = 27) or rotenone (n = 14). p values indicate statistically significant differences.

 
Hypoxia Stimulates Capacitative Ca2+ Entry—Store depletion mediated (or capacitative) Ca2+ entry (CCE) is an important route of Ca2+ entry in non-excitable cells. In our endothelial cells, re-admission of Ca2+ to the perfusate (without prior discharge of intracellular stores) had no effect on [Ca2+]i (e.g. Fig. 6A), but when we first exposed cells to thapsigargin (in Ca2+-free solution) to deplete intracellular stores as completely as possible and then re-admitted Ca2+ (2.5 mM) to the perfusate, this caused a marked rise of [Ca2+]i which could be inhibited by Gd3+ (Fig. 6B) and La3+ (Fig. 6C) in a concentration-dependent manner indicative of CCE. When stores were initially depleted by application of 10 µM ATP, subsequent addition of Ca2+ to the perfusate also triggered a Gd3+- and La3+-sensitive influx of Ca2+ (Fig. 6D), and the same was true when stores were initially depleted by hypoxia (Fig. 6E). Interestingly, when comparing the magnitude of capacitative Ca2+ entry evoked by these different maneuvers, thapsigargin-evoked store depletion produced the largest CCE, and this was also the most sensitive to Gd3+, which suppressed the influx by ~86% at 1 mM (Fig. 6F). ATP- and hypoxia-evoked CCE responses were remarkably similar in terms of magnitude and Gd3+ sensitivity (1 mM causing 40% inhibition of ATP-evoked CCE and 44% inhibition of hypoxia-evoked CCE, Fig. 6E). La3+ was clearly more potent, reducing thapsigargin-evoked CCE by 88%, ATP-evoked CCE by 86%, and hypoxia-evoked CCE by 96% (Fig. 6F).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The present study demonstrates that local O2 levels have marked effects on Ca2+ signaling in vascular endothelial cells and modulate Ca2+ signaling initiated by extracellular ATP. We have defined two distinct pathways by which hypoxia regulates Ca2+ release from the ER. First is the mitochondrial pathway in which hypoxia evokes an increased production of ROS at the mitochondrion to trigger release of Ca2+ from the ER via RyRs (Fig. 7, mitochondrial regulation). Second is the oxidase pathway, in which substrate limited reduction of ROS levels during hypoxia relieves tonic inhibitory influences of oxidase-derived ROS on IP3-dependent Ca2+ release from the ER (Fig. 7, oxidase regulation). Note also that there is cross-talk between these two regulatory pathways; specifically, hypoxia-induced mitochondrial ROS production augments agonist-evoked Ca2+ release.



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  		FIG. 7.
Schematic representation of the two routes of modulation of Ca2+ signaling by hypoxia. Delineated by the dashed area on the left of the figure is the pathway that links hypoxia to Ca2+ release via hypoxic modulation of mitochondrial ROS production; this is termed the mitochondrial regulation pathway. In this pathway, hypoxia increases mitochondrial ROS, which triggers Ca2+ release from the ER via RyRs. Delineated by the dashed area on the right of the figure is the pathway by which hypoxia modulates ATP-evoked Ca2+ release from the ER; this is termed the oxidase regulation pathway. Substrate-limited reduction of ROS levels during hypoxia relieves the tonic inhibitory influence of oxidase-derived ROS on IP 3-dependent Ca2+ release from the ER. Note also that there is cross talk between these two regulatory pathways; specifically hypoxia-induced mitochondrial ROS production augments agonist-evoked Ca2+ release.

 
The scheme depicted in Fig. 7 evolved from the initial finding that hypoxia mobilized Ca2+ from an intracellular pool that was also sensitive to ATP. Such an effect of hypoxia is in accord with an earlier study employing bovine pulmonary endothelial cells (39). Thus, prior exposure to ATP suppressed the hypoxic rise of [Ca2+]i (Fig. 1C), and vice versa (Fig. 1B). Furthermore, thapsigargin pretreatment of cells prevented either agent from altering [Ca2+]i (Fig. 2A). However, it was also clear that hypoxia mobilizes Ca2+ from the ATP-sensitive pool via a mechanism distinct from that of ATP itself. Thus, presumed blockade of IP3 receptors with 2-APB prevented further rises of [Ca2+]i in response to ATP, yet hypoxia caused an additional rise of [Ca2+]i (Fig. 2, B and C). Although 2-APB is known to exert multiple effects on Ca2+ signaling pathways, particularly store-operated Ca2+ entry (28, 40), its application here clearly distinguishes between the ability of hypoxia and ATP to raise [Ca2+]i. Furthermore, exposure of cells to ryanodine fully prevented hypoxic rises of [Ca2+]i yet did not significantly alter responses to ATP. Thus hypoxia mobilized Ca2+ from the ATP-sensitive pool via activation of RyRs. That 8-Br-cADPR and nicotinamide prevented the effects of hypoxia strongly suggests that RyR Ca2+ release was mediated by cADPR (29, 30).

Because mitochondrial ROS have been shown to increase during hypoxia in a variety of cell types (34), we explored their possible role in mediating Ca2+ signaling by investigating the effects of two mechanistically distinct antioxidants (Fig. 3). Surprisingly, both agents had opposing effects on rises of [Ca2+]i evoked by ATP as compared with hypoxia. These observations further strengthen our hypothesis that ATP and hypoxia mobilize Ca2+ from the ER via separate mechanisms but also raise the question of how ROS inhibition could exert such diverse effects. To explore this, we attempted to inhibit selectively ROS production from two separate sources, NADPH oxidase and mitochondria. Our findings strongly suggest the ROS derived from these two sources can exert very different effects on Ca2+ signaling. During normoxia, NADPH oxidase produces ROS, causing a tonic suppression of ATP-evoked rises of [Ca2+]i (Fig. 7). Thus, pharmacological inhibition of the oxidase potentiated ATP-evoked rises of [Ca2+]i (Fig. 4), and this effect was similar to that evoked by the antioxidants (Fig. 3). In contrast, hypoxia-evoked rises of [Ca2+]i were little affected by pharmacological oxidase inhibition, which is not surprising because removal of a large proportion of the substrate (O2) of the enzyme would suppress its activity (Fig. 7, oxidase regulation pathway). This would account for the observation that ATP-evoked signals are potentiated during hypoxia (Fig. 1, D and E) despite the fact that hypoxia causes partial depletion of the ATP-sensitive pool of Ca2+ (Fig. 1B).

Hypoxia also stimulated Ca2+ release from the ER, but the mechanism appears quite distinct (Fig. 7, mitochondrial regulation pathway); antioxidants fully prevented rises of [Ca2+]i evoked by hypoxia (Fig. 3), indicating that ROS are required for this effect but not from NADPH oxidase (Fig. 4). Instead, the primary source of ROS mediating the effects of hypoxia appears to be mitochondria. Numerous studies indicate that hypoxia increases mitochondrial ROS production (34), and ROS can be generated at different sites along the respiratory chain (38, 41, 42). The mechanism is unknown, but we have found hypoxia depolarizes mitochondria. In umbilical vein endothelial cells, ROS generated at complex I appear to be functionally important because rotenone prevents ROS-mediated interleukin-6 production (38). Finally, mitochondria-derived ROS may also cause a small stimulatory effect on ATP-mediated signaling (Fig. 7, cross talk pathway) because both rotenone and FCCP effected small decreases in ATP-evoked signals (Fig. 5E). However, this effect is normally masked by the inhibitory effect of NADPH oxidase-derived ROS because antioxidants (which would presumably buffer all ROS, regardless of source) potentiated ATP-evoked rises of [Ca2+]i.

Results presented in Fig. 6 indicate an additional, important effect of hypoxia on Ca2+ homeostasis. Thus, store depletion by hypoxia clearly activated a Ca2+ influx pathway because addition of Ca2+ to the perfusate following discharge of Ca2+ stores either by thapsigargin, ATP, or hypoxia led to a rise of [Ca2+]i. Regardless of the method of store depletion, the subsequent Ca2+ influx pathway was clearly sensitive to Gd3+ and La3+. However, there were differences between the influx pathway(s) activated by thapsigargin pretreatment and those activated by ATP or hypoxia. Thus, the thapsigargin-activated Ca2+ influx was much greater in magnitude than that activated by ATP or hypoxia and was inhibited >85% by 1 mM Gd3+. In contrast, the magnitudes of ATP- and hypoxia-evoked influxes were extremely similar to each other (despite these agents discharging markedly different amounts of Ca2+ from internal stores (Fig. 1)) as was the lesser sensitivity to blockade by Gd3+ (~40%). Sensitivity to the more potent inhibitor, La3+, was similar regardless of the mechanism of store depletion (Fig. 6F). Store-operated Ca2+ entry (or CCE) is believed to be mediated by transient receptor potential (TRP) channels, of which the canonical (TRPC) family are best known to be store-operated (43). Endothelial cells express six of the seven known members of the TRPC family, TRPC1–6 (43). Our data suggest that ATP and hypoxia may activate the same TRPC channels, but these may be distinct from (or possibly a subset of) those activated by thapsigargin. We cannot presently identify specific members of this family involved in the CCE recorded in this study, but the relatively weak sensitivity to blockade by Gd3+ probably excludes the involvement of TRPC3 (44).

In summary, our results indicate that hypoxia is a key determinant of intracellular Ca2+ signaling in human venous endothelial cells, modulating agonist-evoked liberation of Ca2+ from the ER through regulation of ROS generation from NADPH oxidase. Additionally, hypoxia itself liberates Ca2+ from the ER via mitochondrial ROS generation and activation of RyRs. Furthermore, because hypoxia is also capable of activating CCE, our results indicate that local O2 levels should be considered when investigating such mechanisms in these and other cell types. Such effects are likely to be of importance not only in the normal physiology of the vasculature but also in pathophysiological states induced by, or involving, hypoxia.


   FOOTNOTES
 
* This work was supported by the British Heart Foundation, Medical Research Council, Wellcome Trust, and the Biotechnology and Biological Sciences Research Council/Pfizer Central Research (through a Collaborative Awards in Science and Engineering award to P. K. A.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed: School of Medicine, University of Leeds, Leeds LS2 9JT, UK. Tel.: 113-343-4174; Fax: 113-343-4803; E-mail: c.s.peers{at}leeds.ac.uk.

1 The abbreviations used are: ER, endoplasmic reticulum; IP3, inositol trisphosphate; 2-APB, 2-aminoethoxydiphenyl borate; RyR, ryanodine receptor; cADPR, cyclic ADP-ribose; ROS, reactive oxygen species; trolox, 6-hydroxy-2,5,7,8-tetramethylchroman 2-carboxylic acid; TEMPO, 4-hydroxy-2,2,6,6-tetramethylpiperidine 1-oxyl; CCE, capacitative Ca2+ entry; TRP, transient receptor potential; TRPC, canonical TRP. Back


   ACKNOWLEDGMENTS
 
We thank D. J. O'Regan (consultant cardiothoracic surgeon, Leeds General Infirmary) for human saphenous vein samples.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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