| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
1 Department of Pharmacology, University of Nevada School of Medicine, Reno, NV 89557, USA
 |
Abstract |
|---|
 Top Abstract IntroductionMethodsResultsDiscussionReferences |
|---|
(Received 29 October 2004;
accepted after revision 20 December 2004;
first published online 21 December 2004)
Corresponding author J. R. Hume: Department of Pharmacology/318, University of Nevada School of Medicine, Reno, NV 89557, USA. Email: joeh{at}med.unr.edu
|
Introduction |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
In the case of hypoxic-induced release of Ca2+ from intracellular Ca2+ stores, Dipp et al. (2001) suggest that released Ca2+ may be the trigger that activates constriction, and that Ca2+ influx across the plasmalemma is not essential for HPV. On the other hand, depletion of intracellular stores, but not the released Ca2+ per se, has been suggested to mediate HPV through activation of a separate voltage-independent extracellular Ca2+ entry pathway (Jabr et al. 1997; Robertson et al. 2000), so-called capacitative Ca2+ entry (CCE) (Putney and McKay, 1999). The reason for these inconsistencies is unclear, but it could be explained by the variations in animal species, degree of vessel wall pretone and experimental protocols used (see Weissmann et al. 2001 for review).
Several recent studies have confirmed the existence of CCE in PASMCs, and confirmed the expression of several homologues of transient receptor potential proteins, putative candidates for store-operated channels, in PASMCs (Ng and Gurney, 2001; Walker et al. 2001; Wilson et al. 2002; Snetkov et al. 2003; Wang et al. 2003). In cultured rabbit PASMCs, Kang et al. (2003) demonstrated the presence of CCE and hypoxia enhanced the depletion-activated rise in [Ca2+]i. They also found that hypoxia caused Ca2+ release from the SR Ca2+ stores and a nifedipine-insensitive rise in [Ca2+]i (Kang et al. 2002). These observations led to the suggestion that hypoxia may activate CCE by releasing Ca2+ from the SR. However, no direct evidence was provided on the nature of the pathway involved. Thus, the possibility that acute hypoxia causes activation of CCE remains to be demonstrated. More recently, chronic hypoxic exposure of cultured rat PASMCs was found to up-regulate store-operated channels, which may contribute to the enhanced vascular tone in HPV (Lin et al. 2004). This study suggested that HPV may activate CCE, but the question of whether acute hypoxia may activate CCE, and whether this is a direct action of hypoxia on CCE channels or is mediated indirectly by depletion of intracellular stores, has yet to be directly tested. We have recently characterized the properties of CCE activated by intracellular Ca2+ store depletion in freshly isolated canine PASMCs by measuring changes in cytosolic Ca2+ concentration, and by direct measurement of activation of a store-operated Ca2+ permeability pathway by monitoring the rate of Mn2+ quench of fura-2 fluorescence (Wilson et al. 2002). The purpose of the present study was to utilize these same experimental approaches to directly test the hypothesis that acute hypoxia, by causing Ca2+ release from the SR Ca2+ stores, indirectly leads to activation of CCE in freshly isolated canine PASMCs.
|
Methods |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
PASMCs were isolated from canine pulmonary artery as previously described (Wilson et al. 2002). Cross-breed dogs of either sex were killed with pentobarbital sodium (45 mg kg–1 I.V.) and ketamine (15 mg kg–1 I.V.), as conforming with the requirements of the University of Nevada at Reno Institutional Care and Use Committee. The heart and lungs were removed en bloc, and the third and fourth branches of the pulmonary artery were dissected at 5°C to decrease cellular metabolic activity. The arteries were flushed with a low-Ca2+ physiological salt solution (PSS) composed of the following (mM): NaCl 125, KCl 5.36, Na2HPO4 0.34, K2HPO4 0.44, MgCl2 1.2, Hepes 11, glucose 10 and CaCl2 0.05 (pH 7.4 adjusted with Tris). Arteries were cleaned of connective tissue, cut into small pieces and placed in a tube containing fresh low-Ca2+ PSS. Tissue was immediately digested or cold stored at 5°C for up to 24 h. To disperse cells, pulmonary arterial tissue was incubated with the low-Ca2+ PSS containing (mg ml–1): collagenase type XI 0.5, elastase type IV 0.04 and bovine serum albumin 0.5 (fat-free) for 16–18 h at 5°C. The tissue was then transferred to an enzyme-free, low-Ca2+ PSS, and triturated with a fire-polished Pasteur pipette. The resulting dispersed PASMCs were cold-stored in the refrigerator at 5°C until use, where they remained viable for 8–10 h.
Measurement of intracellular Ca2+
The cytosolic Ca2+ concentration was estimated in PASMCs loaded with fura-2 acetoxymethyl ester (fura-2 AM) (Molecular Probes, Eugene, OR, USA) using a dual excitation digital Ca2+ imaging system (IonOptix, Inc., Milton, MA, USA) equipped with an intensified CCD camera as previously described (Wilson et al. 2002). PASMCs were loaded with 10 µM Fura-2 AM for 30 min in the dark at room temperature, and placed on the coverslip in a 0.2 ml perfusion chamber mounted on an epifluorescence microscope (Nikon). To remove extracellular fura-2 AM, cells were washed several times with 2 mM Ca2-PSS composed of the following (mM): NaCl 113, KCl 5, MgCl 1, CaCl2 2, NaH2PO4 0.5, NaHCO3 24 and glucose 10 (gassed continuously with 21% O2/5% CO2/74% N2, pH 7.4). Cells were illuminated with xenon arc lamp at 340 ± 15 and 380 ± 12 nm (Omega Optical, Brattleboro, VT, USA), and from regions that encompassed single cells, emitted light was collected with a CCD camera at 510 nm (Nikon). All experiments were performed at 35–37°C, and images were acquired at 1 Hz, and stored on the compact disk for later analysis. Background fluorescence was collected automatically and subtracted from the acquired fluorescence video images during each experiment. The ratio of fluorescence (R) excited at the two excitation wavelengths was used to estimate intracellular Ca2+ concentration ([Ca2+]i), as described by Grynkiewicz et al. (1985):
|
|
In control experiments, PASMCs were superfused continuously with normoxic PSS in an open air recording chamber. Normoxic PSS was prepared by continuous gassing with certified gas mixture containing 21% O2, 5% CO2 and 74% N2 (Air Liquide, Santa Fe Springs, CA, USA). In experiments where the effect of hypoxia was investigated, hypoxia was induced by switching normoxic PSS to hypoxic PSS, which continuously superfused the cells in the recording chamber. Hypoxic PSS was prepared by continuous gassing with uncertified gas mixture containing 95% N2 and 5% CO2 (Sierra Welding, Sparks, NV, USA). The uncertified gas mixture contained minimal amount of oxygen, which equilibrated with PSS to avoid exposure of cells to anoxic condition. All solutions were placed in a water bath at 37°C and saturated with either normoxic or hypoxic gas mixtures for at least 30 min before the start of perfusion, and maintained at pH 7.4. The PO2 was measured in preliminary experiments with an oxygen-sensitive electrode (MI-730; Microelectrodes, Inc., Bedford, NH, USA) to be 145 ± 1 mmHg during normoxic PSS perfusion, and falling to 15 ± 1 mmHg within 79 ± 2 s of hypoxic exposure. The PO2 of hypoxic solutions was measured at the end of each experiment, and was found to be 15–18 mmHg, which ensured that the PO2 did not approach anoxia during the recording of each experiment. The effect of hypoxia was examined in cells incubated in Ca2+-free PSS for 10 min, followed by the re-exposure of cells with 2 mM Ca2+ PSS for another 10–15 min. Ca2+-free PSS was identical to 2 mM Ca2+ PSS, but with CaCl2 omitted and 1 mM EGTA added. An elevation in [Ca2+]i above basal levels during 2 mM Ca2+ re-addition was used as a marker of hypoxia-induced extracellular Ca2+ entry. In experiments where the Ca2+ influx pathway was studied, the rate of Mn2+-induced quenching of fura-2 fluorescence was recorded during excitation at 360 nm in nominally Ca2+-free PSS containing 10 µM nisoldipine (Wilson et al. 2002). Nominally Ca2+-free solutions were similar to 0 Ca2+ PSS, but with EGTA omitted. In experiments where CCE was investigated, a store-depletion protocol previously described by Wilson et al. (2002) was used. The SR Ca2+ stores were maximally depleted by exposure of cells to a cocktail containing 10 µM cyclopiazonic acid (CPA) and 10 µM ryanodine, followed by a 30 s exposure to 10 µM 5-HT and 10 mM caffeine in Ca2+-free solutions. All experiments were performed on viable isolated PASMCs. Cell viability was assessed by visual examination to ensure that cells remained relaxed under resting unstimulated conditions, and Mn2+ quench experiments were only performed on cells in which the rate of Mn2+ quench of the fura-2 signal in the presence of ionomycin was at least fourfold greater than the basal rate, as previously described (Wilson et al. 2002).
Drug solutions and data analysis
CPA, nisoldipine and ionomycin were dissolved in dimethylsulphoxide (DMSO). Other drugs were dissolved in deionized water. Data are expressed as means ± S.E.M. of n cells isolated from at least three dogs. Statistical comparisons employed Student's paired t tests or one-way analysis of variance (ANOVA) with Tukey's pairwise comparison. A value of P < 0.05 was considered significant.
|
Results |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
R
= 0.08 ± 0.01) above basal levels (Fig. 1C, n
= 148, P < 0.01). The response to hypoxia was sustained and reversed slowly upon reoxygenation. In control experiments, cells were exposed to normoxic solutions throughout the protocol (Fig. 1B). Removal of extracellular Ca2+ caused a mean 104 ± 12 nM (R
= 0.14 ± 0.01) decrease in [Ca2+]i, from a basal value of 153 ± 22 nM (R
= 0.83 ± 0.02) to 49 ± 13 nM (R
= 0.69 ± 0.02, n
= 66, P < 0.01). Subsequent addition of 2 mM Ca2+ caused an increase in [Ca2+]i to 160 ± 20 nM (R
= 0.85 ± 0.02, n
= 66), only 7 ± 6 nM (Fig. 1B; R
= 0.02 ± 0.01, n
= 66) higher than the basal values (not significant). Figure 1D shows that the increase in [Ca2+]i induced by hypoxia was significantly reduced by approximately 35% (n
= 119, P < 0.01) by 10 µM nisoldipine, a dihydropyridine Ca2+ antagonist. Therefore, the hypoxic induced rise in [Ca2+]i involves both dihydropyridine-sensitive and -insensitive Ca2+ entry pathways. This contrasts with the Ca2+ store depletion-induced rise in [Ca2+]i in these cells, which was previously shown to only involve a dihydropyridine-insensitive Ca2+ entry pathway (Wilson et al. 2002).
|
R
= 0.06 ± 0.01) to 20 ± 7 nM (R
= 0.02 ± 0.01) (Fig. 2A and C; n
= 39, P < 0.05), and 500 µM Ni2+ reduced it by about 94%, from 67 ± 11 nM (R
= 0.08 ± 0.01) to 4 ± 8 nM (R
= 0.005 ± 0.007) (Fig. 2B and C; n
= 28, P < 0.01). La3+ (100 µM) (Fig. 2C; n
= 11) and Gd3+ (100 µM) (Fig. 2C; n
= 20) did not significantly reduce the increase in [Ca2+]i.
|
|
We next investigated the effect of hypoxia on Mn2+ quench of fura-2 fluorescence in PASMCs using a similar experimental approach. Figure 4A shows in a single PASMC that removal of extracellular Ca2+ alone did not cause any decline in fluorescence. Addition of MnCl2 (in the presence of 10 µM nisoldipine to inhibit dihydropyridine-sensitive Ca2+ entry) reduced fluorescence intensity at a rate of 0.033 a.u. s–1. Subsequent exposure to hypoxia resulted in a 97% increase in the rate of decline of fluorescence intensity to 0.065 a.u. s–1, indicating hypoxic activation of a nisoldipine-insensitive Ca2+ entry pathway. To determine if CCE contributes to hypoxia-activated Ca2+ entry, the pharmacology of hypoxia-activated Ca2+ entry was examined by using blockers of store-operated channels. Figure 4B and C shows that
|
If CCE contributes to the hypoxic-induced rise in [Ca2+]i, then depletion of the SR Ca2+ stores may be a prerequisite for hypoxia to activate nisoldipine-insensitive Ca2+ entry. Thus predepletion of the stores might prevent the hypoxic induced rise in [Ca2+]i. In accordance with this, pre-exposure of cells to hypoxia might also prevent the store-depletion activated rise in [Ca2+]i. To test these possibilities, SR Ca2+ stores were maximally depleted by exposure of cells to a cocktail including CPA (10 µM) and ryanodine (10 µM), followed by a 30 s exposures to 5-HT (10 µM) and caffeine (10 mM) in Ca2+-free solutions (Wilson et al. 2002). Figure 5A shows that following the store depletion, 2 mM Ca2+ re-addition caused a 102 ± 9 nM (Fig. 5C; R
= 0.041 ± 0.005, n
= 41, P < 0.01) increase in [Ca2+]i above the basal values. Subsequent exposure of cells to hypoxia caused a small, but not significant increase in the depletion-activated rise in [Ca2+]i (Fig. 5C, n
= 41). Similarly, pre-exposure of cells with hypoxia prevented the store-depletion-activated rise in [Ca2+]i. Figure 5B illustrates that hypoxia caused a 175 ± 14 nM increase in [Ca2+]i in the absence of nisoldipine (Fig. 5C; R
= 0.059 ± 0.004, n
= 48, P < 0.01). Subsequent depletion of the intracellular stores did not enhance the sustained rise in [Ca2+]i induced by hypoxia (Fig. 5C; n
= 48). Thus, predepletion of the stores prevented hypoxic induced rise in [Ca2+]i, and hypoxia did not affect the rise in [Ca2+]i induced by store-depletion (Fig. 5A). On the other hand, pre-exposure of cells with hypoxia prevented the store-depletion activated rise in [Ca2+]i, and store-depletion did not affect the rise in [Ca2+]i induced by hypoxia (Fig. 5B). It is noteworthy that the increase in [Ca2+]i induced by hypoxia was significantly higher than that induced by store-depletion (Fig. 5C; P < 0.01). This may be due to an additional component of Ca2+ influx through voltage-gated Ca2+ channels in the absence of nisoldipine.
|
|
Discussion |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
50–100 nM (Wilson et al. 2002). This rise in [Ca2+]i was insensitive to nisoldipine, and inhibited by 500 µM Ni2+ but not by 100 µM Gd3+. Experiments in intact vessels revealed a potentiated contractile response to hypoxia in the presence of CPA or thapsigargin (Jabr et al. 1997). This response was partially sensitive to nisoldipine, but abolished in the absence of extracellular Ca2+, suggesting a recruitment of a novel Ca2+ entry pathway. This pathway was similar to CCE described in rat PASMCs (Ng and Gurney, 2001), since it was also blocked by 50 µM SKF 96365. In the present study, hypoxia caused intracellular Ca2+ release as indicated by a transient rise in [Ca2+]i detected by fura-2 in Ca2+-free solution. Subsequent addition of 2 mM Ca2+ elicited a nisoldipine-sensitive and nisoldipine-insensitive rise in [Ca2+]i. Interestingly, the pharmacological properties of the nisoldipine-insensitive rise in [Ca2+]i (50 nM) are similar to CCE. More importantly, hypoxia accelerated Mn2+ quench of fura-2 fluorescence at a rate similar to that found for store-depletion activated Mn2+ entry. Furthermore, the increase in fura-2 quench rates induced by store-depletion and hypoxia were both inhibited by SKF 96365 and Ni2+, but not affected by Gd3+ and La3+, confirming that hypoxia stimulates Ca2+ entry through a pathway similar to CCE. These data provide the first direct evidence in freshly isolated PASMCs that acute hypoxia, by releasing Ca2+ from the intracellular stores, activates a Ca2+ entry pathway resembling CCE.
In the present study, a rapid fall (100 nM) in cytosolic Ca2+ concentration was observed in response to removal of extracellular Ca2+. This is consistent with our previous published data in canine PASMCs (Wilson et al. 2002). In most of the cells studied in Ca2+-free solution, [Ca2+]i was well maintained at concentration between 20 and 50 nM, and not approaching zero. This observation is not uncommon but in fact consistent with many studies employing similar protocol, where removal of extracellular Ca2+ resulted in a rapid fall in cytosolic Ca2+ (e.g. Doi et al. 2000; Morales et al. 2004). None of these papers explain why removing extracellular Ca2+ causes a rapid fall in [Ca2+]i. However, the nonlinearity of the relationship between fluorescence and Ca2+ binding would predict this phenomenon. It is unlikely that the rapid fall in [Ca2+]i seen in the present study is related to cell viability because re-addition of Ca2+ resulted in a rapid rise in [Ca2+]i to the basal level in the control experiment (Fig. 1B), or above the basal level during hypoxic challenge (Figs 1A, and 2A and B), suggestive of good responsiveness of cells to external Ca2+.
The possibility that hypoxia causes membrane depolarization leading to Ca2+ influx through the activation of L-type Ca2+ channels has long been considered a component of HPV in many pulmonary preparations (Madden et al. 1984; Harder et al. 1985; Archer et al. 1985; Woodmansey et al. 1995; Cornfield et al. 1994; Jabr et al. 1997; Robertson et al. 2000). However, the mechanism(s) underlying these events remains unclear. Our previous electrophysiological study in canine PASMCs showing that hypoxia inhibited K+ currents (Post et al. 1992), together with the present study that hypoxia causes intracellular Ca2+ release and activates a nisoldipine-sensitive rise in [Ca2+]i, is consistent with the possibility that hypoxic release of Ca2+ from intracellular stores may inhibit Kv channels, leading to membrane depolarization and subsequent activation of L-type Ca2+ channels (Post et al. 1995). It is also possible that the Ca2+ release from the intracellular stores may activate Ca2+-dependent Cl– channels, leading to membrane depolarization and hence activation of L-type Ca2+ channels (Clapp et al. 1996; Ng and Gurney, 2001), or that hypoxia may directly activate L-type Ca2+ channels (Franco-Obregón and López-Barneo, 1996).
In various vascular smooth muscle preparations, it has been reported that L-type Ca2+ channel blockers may inhibit Ca2+ release from intracellular Ca2+ stores at concentrations well above those use to block membrane L-type Ca2+ channels (Saida and van Breemen, 1983; Kanaide et al. 1988; Kalsner, 1997). However, the inhibitory effects on Ca2+ release vary among different blockers and vascular preparations. In rabbit mesenteric artery, diltiazem inhibited noradrenaline-induced contractions in Ca2+-free medium at concentrations between 1 and 100 µM, but nisoldipine only partially inhibited the contractions at the same concentrations (Saida and van Breemen, 1983). In cultured rat aorta smooth muscle cells, verapamil and diltiazem partially inhibited noradrenaline-induced Ca2+ release at concentrations up to 100 µM (Kanaide et al., 1988). In contrast, verapamil (10 µM) failed to inihibit Ca2+ release or contraction induced by noradrenaline in Ca2+-free medium in rat aortas (Hagiwara et al. 1993). In the present study, 10 µM nisoldipine was used to inhibit voltage-gated Ca2+ entry activated by hypoxia to study CCE. Although we did not directly test if nisoldipine also inhibited Ca2+ release, in our experiments nisoldipine may have little or no effect on hypoxia-induced Ca2+ release because it was applied after re-addition of external Ca2+, where the stores would have been depleted by hypoxia in Ca2+-free solution (Fig. 2). Furthermore, in cells pretreated with nisoldipine, hypoxia increased the rate of Mn2+ quench of fura-2 fluorescence (Fig. 4). If nisoldipine prevented hypoxia from causing the SR Ca2+ release, the increase in the Mn2+ quench of fura-2 induced by hypoxia may not be observed because activation of CCE required depletion of the intracellular Ca2+ stores.
In intact arteries, agonist preconstriction or pretone is usually a prerequisite to demonstrate hypoxic pulmonary constriction (e.g. Rodman et al. 1989; Ogata et al. 1992; Demiryurek et al. 1993; Hoshino et al. 1994; Vandier et al. 1997; Robertson et al. 2000; Liu et al. 2001). On the other hand, hypoxia was found to induce pulmonary artery contraction and a rise in [Ca2+]i in the absence of pretone (Dipp et al. 2001; Kang et al. 2002). The nature of this priming event is unresolved. It may involve increased myofilament Ca2+ sensitivity, partial depolarization of resting membrane potential or Ca2+ release (see Ward & Aaronson, 1999; Sylvester, 2001 for reviews). How the pretone helps to potentiate the effect of hypoxia is unclear, but the fact that it is needed in some preparations implies that there may be synergy between the pathways activated by agonists and hypoxia (Gurney, 2002). In canine pulmonary artery, hypoxia elicited a significant amount of contraction in vessels preconstricted with phenylephrine (Jabr et al. 1997). Surprisingly, pretreatment with agonist is not necessary for hypoxia to cause an increase in [Ca2+]i in isolated PASMCs. This could be explained by the experimental protocol used in the present study in which cells were exposed to hypoxia in Ca2+-free solution followed by readmission of external Ca2+. When external Ca2+ was removed, a slow release of Ca2+ from the SR may occur (Morales et al. 2004), but this was not sufficient to activate CCE in canine PASMCs because no significant rise in [Ca2+]i above basal level was observed after readmission of 2 mM Ca2+ in the control condition. In addition, removal of external Ca2+ may cause a partial depolarization due to increased Na+ entry (Gonzalez-Martinez, 2003). Such depolarization, however, was not sufficient to activate L-type Ca2+ channels in canine PASMCs because we have found that nisoldipine had no effect on the Ca2+ entry upon re-addition of Ca2+ in unstimulated cells (Wilson et al. 2002). Nevertheless, the small amounts of Ca2+ release from the stores and the partial depolarization of the resting membrane potential may serve as a priming condition necessary for hypoxia to cause a significant rise in [Ca2+]i. Therefore, perhaps it is not the pretone per se that is important, but the activation of particular cellular pathways that acts in synergy with the mechanisms activated in smooth muscle by hypoxia (Gurney, 2002).
We previously demonstrated that ryanodine pretreatment with brief caffeine exposures attenuated the hypoxia-induced contraction in canine pulmonary arteries, suggesting that Ca2+ release from the ryanodine-sensitive stores plays an important early role in HPV (Jabr et al. 1997). While these results have been confirmed by other laboratories (Robertson et al. 2000; Dipp et al. 2001), it is unlikely that hypoxia depletes only the ryanodine-sensitive stores in the present study because hypoxia was found to activate CCE, and the activation of CCE required simultaneous depletion of IP3-sensitive and ryanodine-sensitive stores in canine PASMCs (Wilson et al. 2002). We have previously found that depletion of IP3-sensitive stores alone by CPA and angiotensin II failed to activate CCE in canine PASMCs (Wilson et al. 2002), but an enhanced hypoxia contraction was observed in pulmonary arteries pretreated with CPA or thapsigargin (Jabr et al. 1997). Therefore, it is possible that apart from causing Ca2+ release from the ryanodine-sensitive stores, hypoxia may also inhibit Ca2+-ATPase pump of the SR, thereby depleting the IP3-sensitive stores by preventing Ca2+ uptake (Evans and Dipp, 2002). Although we have no evidence to support this, our findings in canine PASMCs that IP3-sensitive and ryanodine-sensitive stores are two separate entities in the SR (Janiak et al. 2001), and that simultaneous depletion of these stores attenuates the hypoxia-induced rise in [Ca2+]i, suggests that Ca2+ release from both stores may be an early event in the cascade activated by hypoxia, which leads to CCE activation.
In conclusion, hypoxia activates CCE involving a store depletion-induced mechanism, which contributes to the development of HPV. CCE has been shown to associate with cell proliferation, being greater in proliferating bronchial smooth muscle cells (Sweeney et al. 2002) and human PASMCs (Golovina, 1999; Golovina et al. 2001) compared to those that are growth-arrested. Furthermore, hypoxic-induced increases in AP-1 binding activity enhances CCE in human pulmonary artery endothelial cells, which could be implicated in stimulating PASMC proliferation, leading to the pulmonary vascular remodelling in patients with hypoxia-mediated pulmonary hypertension (Fantozzi et al. 2003). Although the mechanism(s) linking hypoxia to CCE activation remain to be elucidated, our finding that hypoxia activates CCE in PASMCs suggests that the molecular signal that links store-depletion to the activation of CCE may play an important role in hypoxia, which may be a useful target for the development of new vasodilators to treat pulmonary hypertension.
|
References |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Clapp
L, Turner
JL
&
Kozlowski
RZ (1996). Ca2+-activated Cl– currents in pulmonary arterial myocytes. Am J Physiol Heart Circ Physiol
270, H1577–1584.
Cornfield
DN, Steven
T, McMurtry
IF, Abman
SH
&
Rodman
DM (1994). Acute hypoxia causes membrane depolarization and calcium influx in fetal pulmonary artery smooth muscle cells. Am J Physiol Lung Cell Mol Physiol
266, L469–475.
Demiryurek AT, Wadsworth RM, Kane KA & Peacock AJ (1993). The role of endothelium in hypoxic constriction of human pulmonary artery rings. Am Rev Respir Dis 147, 283–290.[Medline]
Dipp
M, Nye
PC
&
Evans
AM (2001). Hypoxic release of calcium from sarcoplasmic reticulum of pulmonary artery smooth muscle. Am J Physiol Lung Cell Mol Physiol
281, L318–325.
Doi
S, Damron
DS, Horibe
M
&
Murray
PA (2000). Capacitative Ca2+ entry and tyrosine kinase activation in canine pulmonary arterial smooth muscle cells. Am J Physiol Lung Cell Mol Physiol
278, L118–130.
Evans AM & Dipp M (2002). Hypoxic pulmonary vasoconstriction: cyclic adenosine diphosphate-ribose, smooth muscle Ca2+ stores and the endothelium. Respir Physiol Neurobiol 132, 3–15.[CrossRef][Medline]
Fantozzi
I, Zhang
S, Platoshyn
O, Remillard
CV, Cowling
RT
&
Yuan
JX (2003). Hypoxia increases AP-1 binding activity by enhancing capacitative Ca2+ entry in human pulmonary artery endothelial cells. Am J Physiol Lung Cell Mol Physiol
285, L1233–1245.
Franco-Obregón A & López-Barneo J (1996). Differential oxygen sensitivity of calcium channels in rabbit smooth muscle cells of conduit and resistance pulmonary arteries. J Physiol 491, 511–518.[Medline]
Golovina
VA (1999). Cell proliferation is associated with enhanced capacitative Ca2+ entry in human arterial myocytes. Am J Physiol Cell Physiol
277, C343–349.
Golovina
VA, Platoshyn
O, Bailey
CL, Wang
J, Limsuwan
A, Sweeney
M, Rubin
LJ
&
Yuan
JX (2001). Upregulated TRP and enhanced capacitative Ca2+ entry in human pulmonary artery myocytes during proliferation. Am J Physiol Heart Circ Physiol
280, H746–755.
Gonzalez-Martinez
MT (2003). Induction of sodium dependent depolarization by external calcium removal in human sperm. J Biol Chem
278, 36304–36310.
Grynkiewicz
G, Poenie
M
&
Tsien
RY (1985). A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem
260, 3440–3450.
Gurney AM (2002). Multiple sites of oxygen sensing and their contributions to hypoxic pulmonary vasoconstriction. Respir Physiol Neurobiol 132, 43–53.[CrossRef][Medline]
Hagiwara S, Mitsui M & Karaki H (1993). Effects of felodipine, nifedipine and verapamil on cytosolic Ca2+ and contraction in vascular smooth muscle. Eur J Pharmacol 234, 1–7.[CrossRef][Medline]
Harder
DR, Madden
JA
&
Dawson
C (1985). Hypoxic induction of Ca2+-dependent action potentials in small pulmonary arteries of the cat. J Appl Physiol
59, 1389–1393.
Hoshino
Y, Morrison
KJ
&
Vanhoutte
PM (1994). Mechanisms of hypoxic vasoconstriction in the canine isolated pulmonary artery: role of endothelium and sodium pump. Am J Physiol Lung Cell Mol Physiol
267, L120–127.
Jabr RI, Toland H, Gelband CH, Wang XX & Hume JR (1997). Prominent role of intracellular Ca2+ release in hypoxic vasoconstriction of canine pulmonary artery. Br J Pharmacol 122, 21–30.[CrossRef][Medline]
Janiak
R, Wilson
SM, Montague
S
&
Hume
JR (2001). Heterogeneity of calcium stores and elementary release events in canine pulmonary arterial smooth muscle cells. Am J Physiol Cell Physiol
280, C22–33.
Kalsner
S (1997). Vasodilator action of calcium antagonists in coronary arteries in vitro. J Pharmacol Exp Ther
281, 634–642.
Kanaide
H, Kobayashi
S, Nishimura
J, Hasegawa
M, Shogakiuchi
Y, Matsumoto
T
&
Nakamura
M (1988). Quin 2 microfluorometry and effects of verapamil and diltiazem on calcium release from rat aorta smooth muscle cells in primary culture. Circ Res
63, 16–26.
Kang TM, Park MK & Uhm DY (2002). Characterization of hypoxia-induced [Ca2+]i rise in rabbit pulmonary arterial smooth muscle cells. Life Sci 70, 2321–2333.[CrossRef][Medline]
Kang TM, Park MK & Uhm DY (2003). Effects of hypoxia and mitochondrial inhibition on the capacitative calcium entry in rabbit pulmonary arterial smooth muscle cells. Life Sci 72, 1467–1479.[CrossRef][Medline]
Lin
MJ, Leung
GPH, Zhang
WM, Yang
XR, Yip
KP, Tse
CM
&
Sham
JSK (2004). Chronic hypoxia-induced upregulation of store-operated and receptor operated Ca2+ channels in pulmonary arterial smooth muscle cells. Circ Res
95, 496–505.
Liu
Q, Sham
JS, Shimoda
LA
&
Sylvester
JT (2001). Hypoxic constriction of porcine distal pulmonary arteries: endothelium and endothelin dependence. Am J Physiol Lung Cell Mol Physiol
280, L856–865.
Madden JA, Harder DR & Dawson C (1984). Hypoxia-induced activation in small isolated pulmonary arteries from the cat. J Appl Physiol 59, 113–118.
Morales
S, Camello
PJ, Alcon
S, Salido
GM, Mawe
G
&
Pozo
MJ (2004). Coactivation of capacitative calcium entry and L-type calcium channels in guinea pig gallbladder. Am J Physiol Gastrointest Liver Physiol
286, G1090–1100.
Morio
Y
&
McMurtry
IF (2002). Ca2+ release from ryanodine-sensitive store contributes to mechanism of hypoxic pulmonary vasoconstriction in rat lungs. J Appl Physiol
92, 527–534.
Ng
LC
&
Gurney
AM (2001). Store-operated channels mediate Ca2+ influx and contraction in rat pulmonary artery. Circ Res
89, 923–929.
Ogata M, Ohe M, Katayose D & Takishima T (1992). Modulatory role of EDRF in hypoxic contraction of isolated porcine pulmonary arteries. Br J Pharmacol 114, 1165–1170.
Osipenko
ON, Tate
RJ
&
Gurney
AM (2000). Potential role for Kv3.1b channels as oxygen sensors. Circ Res
86, 534–540.
Parekh
AB
&
Penner
R (1997). Store depletion and calcium influx. Physiol Rev
77, 901–930.
Post
JM, Gelband
CH
&
Hume
JR (1995). [Ca2+]i inhibition of K+ channels in canine pulmonary artery. Novel mechanism for hypoxia-induced membrane depolarization. Circ Res
77, 131–139.
Post
JM, Hume
JR, Archer
SL
&
Weir
EK (1992). Direct role for K+ channel inhibition in hypoxic pulmonary vasoconstriction. Am J Physiol Cell Physiol
262, C882–890.
Putney JW & McKay RR (1999). Capacitative calcium entry channels. Bioessays 21, 38–46.[CrossRef][Medline]
Robertson
TP, Hague
D, Aaronson
PI
&
Ward
JP (2000). Voltage-independent calcium entry in hypoxic pulmonary vasoconstriction of intrapulmonary arteries of the rat. J Physiol
525, 669–680.
Rodman
DM, Yamaguchi
T, O'Brien
RF
&
McMurtry
IF (1989). Hypoxic contraction of isolated rat pulmonary artery. J Pharmacol Exp Ther
248, 952–959.
Saida
K
&
van Breemen
C (1983). Mechanism of Ca2+ antagonist-induced vasodilation. Intracellular actions. Circ Res
52, 137–142.
Salvaterra
CG
&
Goldman
WF (1993). Acute hypoxia increases cytosolic calcium in cultured pulmonary arterial myocytes. Am J Physiol Lung Cell Mol Physiol
264, L323–328.
Snetkov VA, Aaronson PI, Ward JP, Knock GA & Robertson TP (2003). Capacitative calcium entry as pulmonary specific vasoconstrictor mechanism in small arteries of the rat. Br J Pharmacol 140, 97–106.[CrossRef][Medline]
Sweeney
M, McDaniel
SS, Platoshyn
O, Zhang
SYuY, Lapp
BR, Zhao
Y, Thistlethwaite
PA
&
Yuan
JX (2002). Role of capacitative Ca2+ entry in bronchial contraction and remodeling. J Appl Physiol
92, 1594–1602.
Sylvester
JT (2001). Hypoxic pulmonary vasoconstriction – A radical view. Circulation Res
88, 1228–1230.
Vandier
C, Delpech
M, Rebocho
M
&
Bonnet
P (1997). Hypoxia enhances agonist-induced pulmonary arterial contraction by increasing calcium sequestration. Am J Physiol Heart Circ Physiol
273, H1075–1081.
von Euler US & Liljestrand G (1946). Observation on the pulmonary arterial blood pressure in the cat. Acta Physiol Scand 12, 301–320.
Walker
RL, Hume
JR
&
Horowitz
B (2001). Differential expression and alternative splicing of TRP channel genes in smooth muscles. Am J Physiol Cell Physiol
280, C1184–1192.
Wang J, Shimoda LA & Sylvester JT (2003). Capacitative calcium entry and TRPC channel proteins are expressed in rat distal pulmonary arterial smooth muscle. Am J Physiol Lung Cell Mol Physiol 286, L848–858.[CrossRef][Medline]
Ward JPT & Aaronson PI (1999). Mechanisms of hypoxic pulmonary vasoconstriction: can anyone be right? Respir Physiol 115, 261–271.[CrossRef][Medline]
Weir EK, Hong Z, Porter VA & Reeve HL (2002). Redox signaling in oxygen sensing by vessels. Respir Physiol Neurobiol 132, 121–130.[CrossRef][Medline]
Weissmann
N, Grimminger
F, Olschewski
A
&
Seeger
W (2001). Hypoxic pulmonary vasoconstrction: a multifactorial response?
Am J Physiol Lung Cell Mol Physiol
281, L314–317.
Wilson
SM, Mason
HS, Smith
GD, Nicholson
N, Johnston
L, Janiak
R
&
Hume
JR (2002). Comparative capacitative calcium entry mechanisms in canine pulmonary and renal arterial smooth muscle cells. J Physiol
543, 917–931.
Woodmansey PA, Zhang F, Channer KS & Morice AH (1995). Effect of the calcium antagonist amlodipine on the two phases of hypoxic pulmonary vasoconstriction in rat large and small isolated pulmonary arteries. J Cardiovasc Pharmacol 25, 324–329.[Medline]
Yuan
XJ, Goldman
WF, Tod
ML, Rubin
LJ
&
Blaustein
MP (1993). Hypoxia reduces potassium currents in cultured rat pulmonary but not mesenteric arterial myocytes. Am J Physiol Lung Cell Mol Physiol
264, L116–123.
|
Acknowledgements |
|---|
Author's present address
S. M. Wilson: Department of Pharmacology, University of Mississippi School of Pharmacy, University, MS 38677, USA.
This article has been cited by other articles:
|
|
 |
|
  W. Lu, J. Wang, L. A. Shimoda, and J. T. Sylvester Differences in STIM1 and TRPC expression in proximal and distal pulmonary arterial smooth muscle are associated with differences in Ca2+ responses to hypoxia Am J Physiol Lung Cell Mol Physiol, July 1, 2008; 295(1): L104 - L113. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. C. Ng, B. D. Kyle, A. R. Lennox, X.-M. Shen, W. J. Hatton, and J. R. Hume Cell culture alters Ca2+ entry pathways activated by store-depletion or hypoxia in canine pulmonary arterial smooth muscle cells Am J Physiol Cell Physiol, January 1, 2008; 294(1): C313 - C323. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
W. Wu, O. Platoshyn, A. L. Firth, and J. X.-J. Yuan Hypoxia divergently regulates production of reactive oxygen species in human pulmonary and coronary artery smooth muscle cells Am J Physiol Lung Cell Mol Physiol, October 1, 2007; 293(4): L952 - L959. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Wang, L. Weigand, J. Foxson, L. A. Shimoda, and J. T. Sylvester Ca2+ signaling in hypoxic pulmonary vasoconstriction: effects of myosin light chain and Rho kinase antagonists Am J Physiol Lung Cell Mol Physiol, September 1, 2007; 293(3): L674 - L685. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
O. Platoshyn, Y. Yu, E. A Ko, C. V. Remillard, and J. X.-J. Yuan Heterogeneity of hypoxia-mediated decrease in IK(V) and increase in [Ca2+]cyt in pulmonary artery smooth muscle cells Am J Physiol Lung Cell Mol Physiol, August 1, 2007; 293(2): L402 - L416. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M.-J. Lin, X.-R. Yang, Y.-N. Cao, and J. S. K. Sham Hydrogen peroxide-induced Ca2+ mobilization in pulmonary arterial smooth muscle cells Am J Physiol Lung Cell Mol Physiol, June 1, 2007; 292(6): L1598 - L1608. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. M. Evans AMP-activated protein kinase underpins hypoxic pulmonary vasoconstriction and carotid body excitation by hypoxia in mammals Exp Physiol, September 1, 2006; 91(5): 821 - 827. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
