Acute hypoxia increases intracellular [Ca2+] in pulmonary arterial smooth muscle by enhancing capacitative Ca2+ entry

Jian Wang, Larissa A. Shimoda, Letitia Weigand, Wenqian Wang, Dejun Sun, and J. T. Sylvester

Division of Pulmonary and Critical Care Medicine, Johns Hopkins University School of Medicine, Baltimore, Maryland

Submitted 4 December 2004 ; accepted in final form 19 January 2005


    ABSTRACT
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
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Hypoxic pulmonary vasoconstriction (HPV) requires influx of extracellular Ca2+ in pulmonary arterial smooth muscle cells (PASMCs). To determine whether capacitative Ca2+ entry (CCE) through store-operated Ca2+ channels (SOCCs) contributes to this influx, we used fluorescent microscopy and the Ca2+-sensitive dye fura-2 to measure effects of 4% O2 on intracellular [Ca2+] ([Ca2+]i) and CCE in primary cultures of PASMCs from rat distal pulmonary arteries. In PASMCs perfused with Ca2+-free Krebs Ringer bicarbonate solution (KRBS) containing cyclopiazonic acid to deplete Ca2+ stores in sarcoplasmic reticulum and nifedipine to prevent Ca2+ entry through L-type voltage-operated Ca2+ channels (VOCCs), hypoxia markedly enhanced both the increase in [Ca2+]i caused by restoration of extracellular [Ca2+] and the rate at which extracellular Mn2+ quenched fura-2 fluorescence. These effects, as well as the increased [Ca2+]i caused by hypoxia in PASMCs perfused with normal salt solutions, were blocked by the SOCC antagonists SKF-96365, NiCl2, and LaCl3 at concentrations that inhibited CCE >80% but did not alter [Ca2+]i responses to 60 mM KCl. In contrast, the VOCC antagonist nifedipine inhibited [Ca2+]i responses to hypoxia by only 50% at concentrations that completely blocked responses to KCl. The increased [Ca2+]i caused by hypoxia was completely reversed by perfusion with Ca2+-free KRBS. LaCl3 increased basal [Ca2+]i during normoxia, indicating effects other than inhibition of SOCCs. Our results suggest that acute hypoxia enhances CCE through SOCCs in distal PASMCs, leading to depolarization, secondary activation of VOCCs, and increased [Ca2+]i. SOCCs and CCE may play important roles in HPV.

vascular smooth muscle; calcium; ion channels; pulmonary artery; rat


HYPOXIC PULMONARY VASOCONSTRICTION (HPV) plays important roles in health and disease, but its mechanisms remain unclear (60). Although pulmonary arterial endothelial cells exert major influence (26, 49, 68), the essential sensor, transducer, and effector mechanisms of HPV are thought to be contained within pulmonary arterial smooth muscle cells (PASMCs). Considerable evidence suggests that hypoxia inhibits sarcolemmal voltage-dependent potassium (KV) channels in these cells, leading to depolarization, activation of voltage-operated Ca2+ channels (VOCCs), Ca2+ influx, Ca2+/calmodulin-dependent activation of myosin light chain kinase, phosphorylation of myosin light chains, actin-myosin interaction, and contraction. This evidence includes demonstration that hypoxia reduced KV currents (40, 41, 49, 71), caused depolarization (1, 15, 27, 40, 41, 71), increased intracellular Ca2+ concentration ([Ca2+]i) (2, 3, 7, 8, 40, 43, 45, 49, 63, 64, 72), and induced contraction (19, 23, 26, 28, 34) in isolated pulmonary arteries (PAs) and PASMCs. Moreover, VOCC antagonists or removal of extracellular Ca2+ inhibited hypoxic responses in isolated PAs (19, 23, 26) and PASMCs (3, 7, 63), as well as HPV in isolated lungs (31, 42, 56) and intact animals (35, 51, 53, 62).

Other data, however, suggest that mechanisms of HPV are more complicated. KV channel antagonists did not prevent hypoxic responses in PASMCs (49), PAs (1), or isolated lungs (5, 16). In addition, antagonists of VOCCs did not completely block [Ca2+]i responses to hypoxia in PASMCs (7, 20, 45) or HPV in isolated lungs (31). Furthermore, contractile and [Ca2+]i responses to hypoxia persisted in PAs after depolarization and inhibition of VOCCs (44), and the increase in [Ca2+]i caused by hypoxia in PASMCs occurred before depolarization (40). Finally, hypoxic responses in PASMCs (64), PAs (18, 26), and isolated lungs (33) were inhibited by ryanodine, which blocks Ca2+ release from sarcoplasmic reticulum (SR). These data suggest that HPV requires Ca2+ release from SR and Ca2+ influx through pathways other than VOCCs.

One such pathway could be capacitative Ca2+ entry (CCE). Although CCE has long been recognized as a major pathway for Ca2+ influx in nonexcitable cells, there is growing appreciation that CCE is also important in excitable cells, such as vascular smooth muscle (12, 30, 48, 54). CCE is triggered by depletion of SR Ca2+ stores, which can result from either enhanced release of SR Ca2+ through ryanodine and inositol 1,4,5-trisphosphate receptors in the SR membrane or depressed SR Ca2+ uptake by sarcoplasmic-endoplasmic reticulum Ca2+ ATPases. The pathway for CCE is probably sarcolemmal cation channels composed of proteins homologous to the transient receptor potential (TRP) proteins that provide Ca2+ entry in photoreceptor cells of Drosophila melanogaster (32, 38). CCE is thought to replete SR Ca2+ stores and signal cellular responses (38).

Previously, we (65) and others (13, 20, 21, 29, 36, 52, 58) found that CCE and TRP proteins were expressed in PASMCs. Here, we test the hypothesis that the increase in [Ca2+]i caused by acute hypoxia in PASMCs requires enhanced CCE. A preliminary report of our findings has appeared (66).


    METHODS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell isolation and culture. Distal (>4th generation) intrapulmonary arteries and descending thoracic aortas were dissected from male Wistar rats (body wt = 300–500 g). Endothelium was removed with a cotton swab. As previously described (65), myocytes were obtained by enzymatic digestion and cultured for 3–6 days in smooth muscle growth medium 2 (Clonetics, Walkersville, MD) containing 5% serum in a humidified atmosphere of 5% CO2-95% air at 37°C. Twenty-four hours before an experiment, serum concentration in culture media was decreased to 0.3% to stop cell growth. Cellular purity of cultures, as assessed by morphological appearance under phase-contrast microscopy and immunofluorescence staining for {alpha}-actin (65), was >95%. On the day of the experiment, we also confirmed that 60 mM KCl caused [Ca2+]i to increase, indicating the presence of functional VOCCs.

Measurement of intracellular Ca2+. As previously described (65), coverslips with myocytes were incubated with 7.5 µmol/l fura-2 AM (Molecular Probes, Eugene, OR) for 60 min at 37°C under an atmosphere of 5% CO2-95% air, mounted in a closed polycarbonate chamber clamped in a heated aluminum platform (PH-2; Warner Instrument, Hamden, CT) on the stage of a Nikon TSE 100 Ellipse inverted microscope (Nikon, Melville, NY), and perfused at 0.5 ml/min with Krebs Ringer bicarbonate solution (KRBS), which contained (in mM) 118 NaCl, 4.7 KCl, 2.5 CaCl2, 0.57 MgSO4, 1.18 KH2PO4, 25 NaHCO3, and 10 glucose. Perfusate was equilibrated with 5% CO2 and the desired O2 concentration in heated reservoirs and led to the chamber through stainless steel tubing. PO2 in the chamber was 112 ± 2.0 and 34 ± 1.8 mmHg when reservoir O2 concentrations were 16 and 4%, respectively. Chamber temperature was maintained at 37°C with an in-line heat exchanger and dual channel heater controller (SF-28 and TC-344B, Warner Instrument). In experiments with LaCl3, we used a HEPES-buffered salt solution (HBSS), which did not contain bicarbonate, phosphate, or EGTA to avoid precipitation and chelation of La3+ (44, 52, 65). This perfusate contained (in mM) 130 NaCl, 5 KCl, 2.5 CaCl2, 1.2 MgCl2, 10 HEPES, and 10 glucose and was equilibrated with 16, 4, or 2% O2-balance N2.

After perfusion for 10 min to remove extracellular dye, we measured fura-2 fluorescence emitted at 510 nm after excitation at 340 and 380 nm (F340, F380) at 30- to 60-s intervals in 15–30 cells using a xenon arc lamp, interference filters, electronic shutter, x20 fluorescence objective, and cooled charge-coupled device imaging camera, as previously described (65). Protocols were executed and data were collected on-line with InCyte software (Intracellular Imaging, Cincinnati, OH). Fura-2 fluorescence ratios (R = F340/F380) allowed estimation of [Ca2+]i from R measured in calibration solutions with [Ca2+] between 0 and 1,350 nM (Molecular Probes). To determine whether hypoxia altered behavior of fura-2 in PASMCs, as reported for endothelial cells (55), we perfused PASMCs at 37°C with KRBS without added Ca2+ after equilibration with 5% CO2 and either 16 or 4% O2. After 10–15 min, we depleted SR Ca2+ stores by adding cyclopiazonic acid (CPA, 10 µM), an inhibitor of SR Ca2+-ATPase, to the perfusate. After an additional 10 min, we perfused cells with one of three KRB calibrating solutions containing 10 µM CPA, 10 µM 4-bromo-A23187 (a nonfluorescent Ca2+ ionophore), and either 1) 2.5 mM [Ca2+], 2) no added Ca2+ and 5 mM EGTA, or 3) the Ca2+ buffer dibromo-BAPTA (1 mM) and sufficient Ca2+ to yield a free [Ca2+] of 897 nM, as determined by a computer program (http://www.stanford.edu/~cpatton/maxc.html) that takes pH, temperature, and ionic strength into account (39). Stable levels of F340 and F380 were achieved within 5 min and used to calculate maximum, minimum, and intermediate values of R (Rmax, Rmin, and Rmed, respectively) after subtraction of background fluorescence, which was estimated as the minimal fluorescence achieved during a subsequent 10-min perfusion with Ca2+-free KRBS containing CPA, 4-bromo-A23187, and 0.5 mM MnCl2. Under normoxic conditions, Rmin, Rmed, and Rmax averaged 0.535 (SD 0.088), 1.988 (SD 0.279), and 3.257 (SD 0.433), respectively (n = 5, 5, and 6). These values did not differ significantly from those measured under hypoxic conditions [0.562 (SD 0.030), 2.096 (SD 0.313), and 3.519 (SD 0.155), respectively; n = 5 in each group], indicating that hypoxia did not alter behavior of fura-2 in our cells.

To assess CCE, we perfused PASMCs for 10–15 min with normoxic or hypoxic Ca2+-free KRBS containing 0.5 mM EGTA to chelate residual Ca2+, 5 µM nifedipine to prevent Ca2+ entry through L-type VOCCs, and 10 µmol/l CPA to deplete SR Ca2+ stores. EGTA was not included in the Ca2+-free HBSS used in La3+ experiments. When used, CCE antagonists (SKF-96365, NiCl2, and LaCl3) were added to the perfusate at the same time as CPA. We then estimated CCE as previously described (65) from 1) the increase in [Ca2+]i caused by restoration of perfusate [Ca2+] to 2.5 mM and 2) the rate at which fura-2 fluorescence excited at 360 nm (F360) was quenched after addition of MnCl2 (200 µM).

Drugs and materials. Fura-2 AM (Molecular Probes) was prepared on the day of the experiment as a 2.5 mM stock solution in dimethyl sulfoxide (DMSO). Stock solutions of SKF-96365 (10 mM), LaCl3 (100 mM), and NiCl2 (500 mM) in water and CPA (30 mM), nifedipine (30 mM), and 4-bromo-A23187 (10 mM) in DMSO were made daily. All of these agents were obtained from Sigma Chemical, St. Louis, MO, except 4-bromo-A23187 (CalBiochem, La Jolla, CA). The KRB intermediate Ca2+ calibration solution (see above) was made by adding appropriate amounts of dibromo-BAPTA (Molecular Probes), CaCl2 standard solution (0.5 mol/l, Sigma Chemical), and highly purified NaCl, NaHCO3, KCl, and KH2PO4 (Biochemika Ultra, Sigma Chemical) to deionized water equilibrated with 16% O2-5% CO2 at 37°C.

Statistical analysis. Unless otherwise indicated, data are expressed as means ± SE, and n = number of experiments performed, where the number of cells in each experiment = 15–30, as indicated in figure legends. Statistical comparisons were performed using Student's t-test or analysis of variance. In these analyses, mean data across 15–30 cells in each experiment were entered as an n of 1. Differences were considered significant when P < 0.05.


    RESULTS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
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In PASMCs, acute hypoxia increased [Ca2+]i in a concentration-dependent manner (Fig. 1, A and C). The [Ca2+]i response to hypoxia was consistent among cells in individual experiments (Fig. 2) and half-maximum at a PO2 of 39 mmHg (Fig. 1D). PASMCs and aortic smooth muscle cells (ASMCs), which were isolated and cultured in the same manner as PASMCs, increased [Ca2+]i in response to 60 mM KCl (Fig. 1B), but ASMCs did not increase [Ca2+]i in response to hypoxia (Fig. 1A).



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Fig. 1. Mean changes in intracellular calcium concentration ([Ca2+]i) caused by 4% O2 (A) or 60 mM KCl (B) in rat distal pulmonary arterial smooth muscle cells (PASMCs, n = 12 experiments in 335 cells) and aortic smooth muscle cells (ASMCs, n = 9 experiments in 228 cells). Responses to hypoxia, but not KCl, were different between PASMCs and ASMCs (P = 0.0005 and 0.96, respectively). Effects of hypoxia were concentration dependent (C) with half-maximum response occurring at a PO2 (P50) of 39 mmHg (D, n = 33 experiments in 825 cells). Sigmoid line in D shows least-squares fit of the Hill equation to average peak {Delta}[Ca2+]i measured at each PO2, expressed as a fraction of average {Delta}[Ca2+]i at 0% O2 ({Delta}max).

 


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Fig. 2. Changes in [Ca2+]i caused by 4% O2 in a single experiment. Changes in each of the 16 PASMCs in this experiment are shown on the left. Means and SDs across cells are shown on the right.

 
Perfusion of PASMCs with Ca2+-free KRBS did not alter [Ca2+]i under normoxic conditions (data not shown) but rapidly reversed the increase in [Ca2+]i caused by 4% O2 (Fig. 3A). In contrast, the L-type VOCC antagonist nifedipine, given in a concentration (5 µM) that completely blocked the [Ca2+]i response to KCl (Fig. 3B), only partially prevented (Fig. 3C) or reversed (Fig. 3D) the [Ca2+]i response to hypoxia.



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Fig. 3. Effects of perfusion with Ca2+-free Krebs Ringer bicarbonate solution (KRBS, 0 Ca2+) on [Ca2+]i in rat distal PASMCs during hypoxia (A, n = 4 experiments in 99 cells). Concentration-dependent effects of nifedipine on [Ca2+]i responses of rat distal PASMCs to KCl (B, n = 14 experiments in 390 cells). Effects of 5 µM nifedipine on the [Ca2+]i response to 4% O2 when given before (C, n = 15 experiments in 387 cells) or during hypoxic exposure (D, n = 8 experiments in 218 cells).

 
We assessed CCE by measuring the peak increase in [Ca2+]i caused by restoration of extracellular [Ca2+] to 2.5 mM in PASMCs perfused with Ca2+-free KRBS containing CPA and nifedipine. As shown in Fig. 4, A and B, peak {Delta}[Ca2+]i was threefold greater in hypoxic than normoxic cells. Similar results were obtained in PASMCs perfused with bicarbonate-, phosphate- and EGTA-free HBSS (Fig. 4, C and D).



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Fig. 4. Effects of restoration of extracellular [Ca2+] to 2.5 mM in PASMCs perfused with Ca2+-free KRBS (0 Ca2+) containing cyclopiazonic acid (CPA, 10 µM) and nifedipine (Nifed, 5 µM) during normoxia (n = 5 experiments in 146 cells) and hypoxia (n = 4 experiments in 118 cells, A). Maximum increase in [Ca2+]i after restoration was ~3-fold greater during hypoxia (B). Similar results were obtained in PASMCs perfused with Ca2+-free HBSS containing CPA and nifedipine (C), where maximum increase in [Ca2+]i after extracellular Ca2+ restoration (D) was ~2-fold greater during hypoxia (n = 4 experiments in 112 cells) than normoxia (n = 7 experiments in 193 cells).

 
Because increases in peak {Delta}[Ca2+]i induced by restoration of extracellular [Ca2+] could be due to decreased efflux and intracellular sequestration of Ca2+ as well as increased influx, we also assessed CCE from the rate at which Mn2+ quenched fura-2 fluorescence, which is thought to be a specific index of Ca2+ influx. In normoxic PASMCs perfused with Ca2+-free KRBS containing nifedipine, but not CPA, the fall of fluorescence in cells exposed to Mn2+ was not different from that occurring spontaneously in cells not exposed to Mn2+ (Fig. 5, A and B), indicating that Ca2+ influx was negligible under these conditions. Hypoxia in the absence of CPA increased Mn2+ quenching, indicating hypoxic activation of Ca2+ entry through channels other than VOCCs (Fig. 5, A and B). Depletion of SR Ca2+ stores with CPA markedly increased Mn2+ quenching in normoxic cells, and this increase was further enhanced by hypoxia (Fig. 5, C and D). Similar results were obtained in PASMCs perfused with Ca2+-free HBSS containing CPA and nifedipine (Fig. 5, E and F). The curvilinearity of the average fluorescence-time relationship after administration of Mn2+ during hypoxia (Fig. 5, C and E) was due to achievement of minimal background fluorescence before 10 min in some cells.



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Fig. 5. Quenching of fura-2 fluorescence by 200 µM Mn2+ in PASMCs perfused with Ca2+-free KRBS (0 Ca2+) containing nifedipine (5 µM) but not CPA during normoxia (n = 4 experiments in 76 cells) and hypoxia (n = 5 experiments in 93 cells, A). Magnitude of quenching 10 min after Mn2+ administration during normoxia was not different from the spontaneous decrease in fluorescence measured in normoxic cells not exposed to Mn2+ (n = 4 experiments in 83 cells), but hypoxia increased this rate 2-fold (B). Mn2+ quenching was markedly increased in normoxic PASMCs perfused with Ca2+-free KRBS (0 Ca2+) containing both nifedipine (5 µM) and CPA (10 µM) (n = 6 experiments in 163 cells) and increased further by hypoxia (n = 8 experiments in 216 cells, C and D). Similar results were obtained in PASMCs perfused with Ca2+-free HBSS containing CPA and nifedipine (E and F; n = 4 experiments in 111 cells exposed to 16% O2, n = 6 experiments in 157 cells exposed to 4% O2). Note that scales of A and B are different from C–F. F, fluorescence intensity; F0, baseline fluorescence intensity at time O.

 
Because 50 µM SKF-96365, 500 µM Ni2+, and 100 µM La3+ inhibited CCE by >80% in rat distal PASMCs during normoxia (65), we determined whether these concentrations would also prevent enhancement of CCE by hypoxia. As shown in Fig. 6, A and B, SKF-96365 and Ni2+ markedly reduced peak {Delta}[Ca2+]i resulting from restoration of extracellular [Ca2+] in hypoxic PASMCs. La3+ had similar effects in hypoxic PASMCs perfused with bicarbonate-, phosphate-, and EGTA-free HBSS (Fig. 6, C and D). These antagonists also reversed increases in Mn2+ quenching caused by hypoxia alone (Fig. 7, A and B) or hypoxia + CPA (Fig. 7, C–F).



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Fig. 6. Effects of 50 µM SKF-96365 (SKF, n = 4 experiments in 116 cells) and 500 µM NiCl2 (n = 4 experiments in 112 cells) on [Ca2+]i responses to restoration of extracellular [Ca2+] in hypoxic PASMCs perfused with Ca2+-free KRBS (0 Ca2+) containing CPA (10 µM) and nifedipine (5 µM) (A). Compared with untreated hypoxic PASMCs (n = 4 experiments in 91 cells), the antagonists reduced the maximum increase in [Ca2+]i after restoration by >80% (B). LaCl3 (100 µM) had similar effects in hypoxic PASMCs perfused with Ca2+-free HBSS containing CPA and nifedipine (C and D; n = 4 experiments in 103 control cells, n = 4 experiments in 112 cells treated with La3+).

 


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Fig. 7. Quenching of fura-2 fluorescence by 200 µM Mn2+ in hypoxic PASMCs perfused with Ca2+-free KRBS (0 Ca2+) containing nifedipine (5 µM) but not CPA and either 50 µM SKF-96365 (n = 4 experiments in 91 cells) or 500 µM NiCl2 (n = 4 experiments in 86 cells, A). Compared with hypoxic control PASMCs (n = 5 experiments in 93 cells), these antagonists reduced Mn2+ quenching by >40% (B). Despite increased Mn2+ quenching in hypoxic PASMCs perfused with Ca2+-free KRBS (0 Ca2+) containing both CPA (10 µM) and nifedipine (5 µM) (n = 8 experiments in 195 cells), 50 µM SKF-96365 (n = 5 experiments in 123 cells) or 500 µM NiCl2 (n = 5 experiments in 131 cells) still reduced quenching by >60% (C and D). LaCl3 (100 µM) had similar effects in hypoxic PASMCs perfused with Ca2+-free HBSS containing CPA and nifedipine (E and F, n = 4 experiments in 115 control cells and 4 experiments in 118 La3+-treated cells). Note that scales of A and B differ from C–F.

 
Finally, we determined the effect of CCE antagonists on the [Ca2+]i response to hypoxia. In rat distal PASMCs perfused with normal KRBS, SKF-96365 and Ni2+ caused concentration-dependent reversal of the increase in [Ca2+]i caused by 4% O2 (Fig. 8, A and B). To estimate IC50, we divided the average maximum {Delta}[Ca2+]i caused by hypoxia before and after antagonist administration into the minimum value achieved during antagonist administration. The baseline [Ca2+]i used for these calculations was the lowest value achieved during the protocol. Fitting the mean data to the Hill equation yielded IC50 estimates of 3.2 µM for SKF-96365 and 36.7 µM for NiCl2 (Fig. 8C). These measurements were not performed for LaCl3 because this antagonist caused large increases in normoxic baseline [Ca2+]i in PASMCs perfused with normal HBSS {{Delta}[Ca2+]i = 43 (SD 12.2), 57 (SD 5.3), and 129 (SD 44.1) nM at [LaCl3] = 1, 30, and 100 µM, respectively}. Instead, we exposed cells first to LaCl3 (100 µM) and then to 2% O2 for 10 min during the middle of the 30-min La3+ exposure. PASMCs exposed to only LaCl3 or 2% O2 served as controls. As shown in Fig. 9, LaCl3 increased [Ca2+]i during normoxia and prevented the increase in [Ca2+]i caused by hypoxia.



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Fig. 8. Concentration-dependent reversal of [Ca2+]i response to 4% O2 by SKF-96365 (A, n = 24 experiments in 643 cells) or NiCl2 (B, n = 26 experiments in 706 cells). IC50s were estimated by fitting the Hill equation to the data (C), where minimum responses during exposure to antagonists were expressed as percentages of average maximum response before and after exposure to antagonists.

 


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Fig. 9. A: effect of hypoxia alone (2% O2, n = 6 experiments in 141 cells), LaCl3 alone (100 µmol/l, n = 5 experiments in 135 cells), and hypoxia + LaCl3 (n = 5 experiments in 146 cells) on time course of change in [Ca2+]i from baseline value at time of onset of LaCl3 exposure in rat distal PASMCs. For each group, the relevant exposures to hypoxia and/or LaCl3 are indicated by the bars at the top. B: maximum increase in [Ca2+]i from its value at onset of hypoxia in PASMCs exposed to hypoxia alone, LaCl3 alone, or hypoxia + LaCl3.

 

    DISCUSSION
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Hypoxia increased [Ca2+]i in PASMCs, but not ASMCs (Fig. 1A). This response required extracellular Ca2+ (Fig. 3A) and was half-maximal at a PO2 of 39 mmHg (Fig. 2, C and D). These findings are consistent with previous observations in PASMCs (3, 8, 37, 45, 49, 63, 64, 72), PAs (15, 17, 26, 47), and lungs (4, 11, 57, 59) and confirm the physiological relevance of our preparation.

Consistent with previous observations in PASMCs and isolated lungs (7, 20, 31, 45), the L-type VOCC antagonist nifedipine (5 µM) only partially prevented (Fig. 3C) or reversed (Fig. 3D) the effects of hypoxia on [Ca2+]i. Because 5 µM nifedipine completely blocked the [Ca2+]i response to KCl-induced depolarization (Fig. 3B) and removal of extracellular Ca2+ completely reversed the [Ca2+]i response to hypoxia (Fig. 3A), this result indicates that hypoxia activated influx of Ca2+ through L-type VOCCs and another independent pathway.

CCE is the primary pathway for Ca2+ influx in nonexcitable cells but may also be important in excitable cells. PAs or PASMCs treated with thapsigargin (TG) or CPA in Ca2+-free media exhibited increased influx of the Ca2+ surrogate, Mn2+, and contractions or increased [Ca2+]i upon restoration of extracellular Ca2+ (10, 14, 21, 29, 36, 44, 52, 65). In PASMCs subjected to patch clamping under conditions designed to isolate Ca2+ currents, CPA or TG elicited currents with reversal potentials near 0 mV (13, 29, 36, 52, 58, 70). These responses were blocked by SOCC antagonists such as La3+, Cd2+, Ni2+, and SKF-96365, but not by VOCC antagonists such as nifedipine.

To determine whether hypoxia increased CCE, we first measured the effects of restoring extracellular Ca2+ in PASMCs perfused with Ca2+-free salt solutions containing CPA to deplete SR Ca2+ stores and activate SOCCs, and nifedipine to block Ca2+ entry through L-type VOCCs. As shown in Fig. 4, hypoxia caused two- to threefold enhancement of the increase in [Ca2+]i elicited by Ca2+ restoration. This result is consistent with enhanced Ca2+ influx through SOCCs; however, it is also consistent with decreased Ca2+ efflux and decreased intracellular Ca2+ sequestration. Indeed, hypoxia may decrease Ca2+ efflux in PASMCs by inhibiting Na+-Ca2+ exchange (46, 67). Furthermore, decreased mitochondrial uptake of Ca2+ was thought to explain why anoxia in the absence of glucose increased [Ca2+]i during the decay phase of [Ca2+]i transients induced by restoration of extracellular Ca2+ in PASMCs treated with CPA and nifedipine (21). Because of these possibilities, we next determined the effects of hypoxia on Ca2+ influx more directly by measuring the rate at which Mn2+ quenched fura-2 fluorescence in PASMCs perfused with Ca2+-free salt solutions containing CPA and nifedipine. As shown in Fig. 5, C–F, hypoxia increased the rate of Mn2+ quenching twofold. Because Mn2+ is thought to use the same entry pathways as Ca2+, these results indicate that hypoxia increased CCE through SOCCs.

SOCCs can be inhibited by inorganic antagonists such as La3+, Cd2+, and Ni2+ and organic antagonists such as SKF-96365 (25, 36). In PASMCs, we (65) previously found that Ca2+ influx through SOCCs was blocked >80% by SKF-96365, La3+, and Ni2+ at concentrations that did not inhibit VOCCs (50, 100, and 500 µM, respectively). In the present study, these concentrations blocked hypoxic enhancement of both the [Ca2+]i response to extracellular Ca2+ restoration (Fig. 6) and the rate of Mn2+ quenching (Fig. 7, C and D). These results provide additional evidence that hypoxia potentiated CCE through SOCCs.

In normoxic PASMCs treated with nifedipine but not CPA, Mn2+ quenching did not differ from the spontaneous decline of fluorescence in cells not exposed to Mn2+ (Fig. 5, A and B), indicating that Ca2+ entry through pathways other than VOCCs was negligible during normoxia. During hypoxia, however, the rate of Mn2+ quenching in the absence of CPA increased, and this increase was blocked by SKF-96365 and NiCl2 (Fig. 7, A and B). These results suggest that hypoxia was able to evoke CCE on its own, as well as amplify CCE triggered by CPA. These effects of hypoxia could result from either depletion of SR Ca2+ stores or facilitation of signal transduction pathways linking store depletion to SOCC activation. The former is consistent with proposals that hypoxia causes release of SR Ca2+ in PASMCs (9, 18, 20, 28, 45, 64); however, if CPA depleted SR Ca2+ stores completely, the latter may also have occurred.

In contrast to our results, Kang et al. (21) were unable to demonstrate that hypoxia increased the rate of Mn2+ quenching in distal PASMCs exposed to Ca2+-free conditions and nifedipine. This discrepancy could be due to differences in species (rat vs. rabbit), cell culture protocol (primary culture with 1–2 serum-free days before study vs. resuspension of frozen confluent cells and culture in 10% serum for 2–3 days before study), temperature (37 vs. 33–34°C), or stimulus (4% O2 in the presence of 10 mM glucose vs. 0% O2 in the absence of glucose). Among these differences, we speculate that the last may be the most significant, because anoxia without glucose, unlike anoxia with glucose, caused profound acidosis and deterioration of energy state in pulmonary arterial smooth muscle (24). Such changes, which were similar in degree to those induced by 10 mM NaCN (24), might be severe enough to cause cell injury, disrupt normal ionic homeostasis, or alter affinity of intracellular fura-2 for Mn2+ and Ca2+. Alternatively, autofluorescence due to the marked increases in NAD(P)H that would be expected to occur under these severe conditions may have interfered with measurement of fura-2 fluorescence (22).

To confirm that the [Ca2+]i response to hypoxia (Fig. 1) required activation of SOCCs, we measured effects of SKF-96365 and NiCl2 in PASMCs perfused with normal KRBS (Fig. 8). Both agents caused concentration-dependent inhibition of the hypoxic response, with estimated IC50 = 3.2 and 36.7 µM, respectively (Fig. 8C). These values are comparable to concentrations found to inhibit CCE in other preparations (10, 13, 29, 36, 61, 69) and with our own measurements in rat distal PASMCs (6.3 and 191 µM, respectively) (65). The somewhat lower IC50 values for hypoxic responses suggests that the smaller {Delta}[Ca2+]i ({approx}100 nM) caused by hypoxia may have been blocked more readily than the larger {Delta}[Ca2+]i ({approx}500 nM) caused by restoration of extracellular Ca2+ after store depletion. In PASMCs perfused with normal HBSS, LaCl3 increased [Ca2+]i during normoxia, suggesting nonspecific actions such as inhibition of sarcolemmal Ca2+ pumps (21, 50). Nevertheless, 100 µM LaCl3, which abolished CCE in rat distal PASMCs (65), also abolished the [Ca2+]i response to hypoxia (Fig. 9). Together, our results provide strong evidence that SOCCs were essential contributors to the [Ca2+]i response to hypoxia in rat distal PASMCs. It remains possible that recently hypothesized, SKF-96365-sensitive, store- and voltage-independent Ca2+ channels may also have played a role (44).

The highest concentrations of SKF-96365, LaCl3, and NiCl2 (50, 100, and 500 µM, respectively) completely blocked the [Ca2+]i response to hypoxia (Figs. 8 and 9) but did not alter the [Ca2+]i response to KCl (65). In contrast, nifedipine (5 µM) completely blocked the response to KCl (Fig. 3B) but only partially inhibited the response to hypoxia (Fig. 3, C and D). These data suggest that the increase in [Ca2+]i caused by hypoxia was due primarily to activation of SOCCs and secondarily to activation of VOCCs. Because SOCCs are probably nonspecific cation channels with equilibrium potentials near 0 mV (38) and local increases in [Ca2+]i from CCE could alter activity of sarcolemmal K and Cl channels (40), we speculate that activation of VOCCs during hypoxia may have occurred as a result of SOCC-dependent depolarization. Moreover, if SOCCs transported Na+ in preference to Ca2+, depolarization and increasing intracellular [Na+] may have promoted Ca2+ entry via reverse-mode Na+-Ca2+ exchange (6). Further investigation is needed to evaluate these possibilities and to clarify the role of SOCCs and CCE in HPV.


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This work was supported by National Heart, Lung, and Blood Institute Grants HL-51912 and HL-75113 (J. T. Sylvester) and HL-67919 (L. A. Shimoda), a research grant from the American Lung Association of Maryland, and an American Heart Association Scientist Development Grant (J. Wang).


    FOOTNOTES
 

Address for reprint requests and other correspondence: J. T. Sylvester, Div. of Pulmonary & Critical Care Medicine, Johns Hopkins Asthma and Allergy Center, 5501 Hopkins Bayview Cir., Baltimore, MD 21224 (E-mail: jsylv{at}jhmi.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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