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Am J Physiol Lung Cell Mol Physiol 290: L375-L384, 2006. First published October 7, 2005; doi:10.1152/ajplung.00307.2005
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Hypoxia induces hypersensitivity and hyperreactivity to thromboxane receptor agonist in neonatal pulmonary arterial myocytes

M. Hinton,1,3 L. Mellow,3 A. J. Halayko,1,2,3 A. Gutsol,3 and S. Dakshinamurti1,2,3

Departments of 1Physiology and 2Pediatrics, University of Manitoba; and 3Biology of Breathing Group, Manitoba Institute of Child Health, Winnipeg, Canada

Submitted 14 July 2005 ; accepted in final form 30 September 2005


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
PPHN, caused by perinatal hypoxia or inflammation, is characterized by an increased thromboxane-prostacyclin ratio and pulmonary vasoconstriction. We examined effects of hypoxia on myocyte thromboxane responsiveness. Myocytes from 3rd–6th generation pulmonary arteries of newborn piglets were grown to confluence and synchronized in contractile phenotype by serum deprivation. On the final 3 days of culture, myocytes were exposed to 10% O2 for 3 days; control myocytes from normoxic piglets were cultured in 21% O2. PPHN was induced in newborn piglets by 3-day hypoxic exposure (FIO2 0.10); pulmonary arterial myocytes from these animals were maintained in normoxia. Ca2+ mobilization to thromboxane mimetic U-46619 and ATP was quantified using fura-2 AM. Three-day hypoxic exposure in vitro results in increased basal [Ca2+]i, faster and heightened peak Ca2+ response, and decreased U-46619 EC50. These functional changes persist in myocytes exposed to hypoxia in vivo but cultured in 21% O2. Blockade of Ca2+ entry and store refilling do not alter peak U-46619 Ca2+ responses in hypoxic or normoxic myocytes. Blockade of ryanodine-sensitive or IP3-gated intracellular Ca2+ channels inhibits hypoxic augmentation of peak U-46619 response. Ca2+ response to ryanodine alone is undetectable; ATP-induced Ca2+ mobilization is unaltered by hypoxia, suggesting no independent increase in ryanodine-sensitive or IP3-linked intracellular Ca2+ pool mobilization. We conclude hypoxia has a priming effect on neonatal pulmonary arterial myocytes, resulting in increased resting Ca2+, thromboxane hypersensitivity, and hyperreactivity. We postulate that hypoxia increases agonist-induced TP-R-linked IP3 pathway activation. Myocyte thromboxane hyperresponsiveness persists in culture after removal from the initiating hypoxic stimulus, suggesting altered gene expression.

smooth muscle; pulmonary hypertension; persistent pulmonary hypertension of the newborn

PERSISTENT PULMONARY HYPERTENSION of the newborn (PPHN), defined as a failure of the normal fall in pulmonary vascular resistance at or shortly after birth (36), has an incidence of between 0.4 and 6.8 per 1,000 live births (57). It represents a common pathway of injury response following a multiplicity of perinatal stresses, including hypoxia, inflammation, and direct lung injury such as meconium aspiration (9). The initial clinical picture of PPHN is of dynamic pulmonary vasospasm, with labile flow through the pulmonary circuit and right-to-left shunting of blood across the ductus arteriosus. Vasodilator response progressively diminishes (18), giving way to increased extracellular matrix deposition, medial smooth muscle proliferation (39), and impaired vascular distensibility (54), resulting in an increase in pulmonary vascular resistance no longer amenable to therapeutic intervention. Although the primary defect in PPHN has been held to be of endothelial origin, up to one-third of patients treated with exogenous nitric oxide do not respond (23), suggesting a downstream alteration in pulmonary vascular smooth muscle function.

In neonatal sepsis or direct lung injury, an immediate increase in circulating inflammatory cytokines, including TNF-{alpha}, IL-1{beta}, IL-6, and IL-10 (11, 37, 49) triggers second messenger pathways favoring contraction and smooth muscle proliferation (8). Cyclooxygenase (COX) pathway metabolites are implicated in increased pulmonary vascular tone. Arachidonic acid metabolites contribute to the early pulmonary hypertensive response in meconium aspiration (51) and sepsis (27). Thromboxane A2, a prostanoid with potent vasoconstrictive and mitogenic properties, contracts pulmonary vascular smooth muscle by binding to Gq/11-coupled sarcolemmal receptors, leading to increased intracellular [Ca2+] ([Ca2+]i), force generation, and sensitization of the contractile apparatus to Ca2+ (7). Thromboxane is known to be crucial in mediating septic pulmonary hypertension in the neonate (16, 27). Thromboxane also underlies the pulmonary arterial constrictor response to acetylcholine in hypoxic neonatal piglets; early development of thromboxane-mediated constriction contributes to the pathogenesis of chronic hypoxic pulmonary hypertension in newborns (20). Moreover, in an animal model of perinatal hypoxia, diminished COX-1 and prostacyclin synthase activity are shown to cause a shift in production of arachidonic acid metabolites toward an increased thromboxane-to-prostacyclin ratio; the relative increase in thromboxane may contribute to development of increased pulmonary arterial tone in hypoxic pulmonary hypertension (19). Acutely altered arachidonic acid metabolism in the hypoxic pulmonary circuit is localized to the arterial endothelium and adventitia, with smooth muscle acting as the effector (2). In infants with congenital diaphragmatic hernia, postductal hypoxemia and alveolar-to-arterial oxygen ratio are inversely correlated with thromboxane-to-prostacyclin metabolite ratio, suggesting an increased influence of thromboxane (42).

Hypoxia is known to alter perinatal pulmonary vascular agonist responses, blunting acetylcholine-mediated vasorelaxation, while engendering a constrictor response to cholinergic stimulation, mediated by thromboxane (19). The early development of thromboxane-mediated constriction is postulated to contribute to the pathogenesis of PPHN. The role of hypoxia in potentiating the prostanoid vasoconstrictor response in neonatal pulmonary artery and the effects of hypoxia on myocyte thromboxane sensitivity have not been elucidated to date. In this study, we examine pathways of thromboxane-induced calcium mobilization in primary cultured neonatal pulmonary arterial myocytes following in vivo and in vitro exposures to moderate hypoxia; our hypotheses were that the calcium response of hypoxic myocytes to thromboxane would be enhanced and that in vivo hypoxic thromboxane sensitization would not be extinguished by subsequent normoxic cell culture.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Induction of hypoxic PPHN. All experiments were carried out in accordance with the guidelines of the Canadian Council on Animal Care and approved by the institutional review board. Animals were housed in a thermoregulated isolette with appropriate diurnal cycling. Newborn piglets (<24 h old; n = 10) were killed on the day of arrival from a pathogen-free farm supplier. For the in vivo hypoxic model, newborn piglets (n = 4) were placed in a normobaric hypoxic chamber (FIO2 0.10, achieved by a mixture of room air with N2) for 3 days. The chamber was opened for no more than 1 h a day for feeding and cleaning; this protocol has been rationalized by other groups (20). Age-matched control piglets were also obtained at 3 days of age (n = 10). All piglets were killed by pentobarbital overdose and exsanguination. Heart and lungs were removed en bloc and placed in oxygenated cold (4°C) Ca2+-free Krebs-Henseleit physiological buffer containing (in mM): 112.6 NaCl, 25 NaHCO3, 1.38 NaH2PO4, 4.7 KCl, 2.46 MgSO4·7 H2O, and 5.56 Dextrose; pH 7.4. Relative cardiac weight ratio (blotted tissue weight, right ventricle to left ventricle plus septum) was measured to establish the diagnosis of PPHN by estimating right ventricular afterloading.

Primary pulmonary arterial smooth muscle culture. Pulmonary arteries were cultured by a dispersed cell method selective for myocytes (50). In brief, 2nd–6th generation pulmonary arteries were obtained by microdissection into Ca2+-free Krebs-Henseleit physiological buffer. Arteries were allowed to recover in cold HEPES-buffered saline solution (HBS; composition in mM: 130 NaCl, 5 KCl, 1.2 MgCl2, 1.5 CaCl2, 10 HEPES, and 10 glucose; pH 7.4) supplemented with an antibiotic-antimycotic mixture and gentamicin. Arteries were then washed twice with Ca2+-reduced HBS (20 µM CaCl2) and finely minced. Arterial tissue was transferred to a digestion medium containing Ca2+-reduced HBS, type I collagenase (1,750 U/ml), dithiothreitol (1 mM), bovine serum albumin (BSA, 2 mg/ml), and papain (9.5 U/ml) for 15 min at 37°C with gentle agitation. Dispersed pulmonary arterial smooth muscle cells (PASMCs) were collected by centrifugation at 1,200 rpm for 5 min, washed in Ca2+-free HBS to remove digestion solution, and then resuspended in culture medium. The cells were plated on four-chambered cover-glass plates at a density of 4.4 x 104 cells/cm2, in Ham's F-12 medium with L-glutamine supplemented with 10% fetal calf serum, 1% penicillin, and 1% streptomycin.

Once cells were grown to confluence in 21% O2 (~10 days), they were serum-deprived for 2 days (in Ham's F-12 medium with L-glutamine-penicillin-streptomycin and 1% insulin-transferrin-selenium) to synchronize in a contractile phenotype. Hypoxic cultures were then maintained serum-free in a sealed hypoxic environment (10% O2, 5% CO2) for 3 days (equivalent to the exposure time for in vivo hypoxia); control cultures were maintained serum-free in normoxia (21% O2, 5% CO2). PASMCs from hypoxic piglets and from their age-matched controls were also grown to confluence in normoxia. The four treatment groups are delineated in Fig. 1.



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  		Fig. 1. In vivo and in vitro hypoxic exposure protocols. Treatment groups consist of: pulmonary arterial myocytes from piglets exposed to in vivo hypoxia for the first 3 days of life [hypoxic animals (HA)], myocytes from age-matched normoxic control piglets at 3 days of age [normoxic animals (NA)], myocytes from newborn (<24 h old) piglets cultured in in vitro hypoxia for 3 days [hypoxic myocytes (HM)] or cultured in normoxia [normoxic myocytes (NM)]. All cells are synchronized in contractile phenotype by serum deprivation for a total of 5 days before agonist challenge.

 
Calcium imaging. Cells were rinsed free of media in Hanks' balanced salt solution (HBSS; containing in mM: 1.26 CaCl2, 0.493 MgCl2-6 H2O, 0.407 MgSO4-7 H2O, 5.33 KCl, 0.441 KH2PO4, 4.17 NaHCO3, 137.93 NaCl, and 0.338 NaHPO2) with 0.1% BSA. Myocytes were loaded with the Ca2+-sensitive fluorescent dye fura 2-acetoxymethyl ester (fura-2 AM) dissolved in DMSO, as 5 µM in an HBSS/0.1% BSA solution, with 1.0 µg/ml pluronic acid (used for AM ester solubilization), for 1 h at 37°C. Extracellular fura-2 AM was washed off with HBSS/0.1% BSA. The cells were then allowed to recover for 30 min at room temperature, allowing for complete cleavage of intracellular AM esters. Incubation and calcium imaging were carried out in 21% O2. For studies done in a Ca2+-free environment, immediately before recording, cells were rinsed twice and then allowed to recover in Ca2+-free HBSS/0.1% BSA, in which CaCl2 was replaced with equimolar MgCl2 for maintenance of cell adherence (17). Ca2+-free HBSS was thereafter used for agonist vehicle and washes. The Ca2+-free studies were utilized for within-treatment group comparisons only, in view of sarcolemmal ion current variables introduced by the difference in external Mg2+ concentration.

Coverglass plates were secured on an inverted microscope (Olympus) in room air and studied at x20 magnification. Real-time ratiometric imaging of intracellular calcium concentration used excitation wavelengths of 340 and 380 nm and an emission wavelength of 510 nm; data were captured via a charge-coupled device camera and Perkin Elmer software.

Responses to U-46619 and KCl. Cells were equilibrated in HBSS/0.1% BSA, and a stable 50-s baseline was recorded at the start of each recording. Agonist stock solutions in HBSS/0.1% BSA were diluted to desired concentration upon addition to the culture well. Thromboxane mimetic U-46619 [(1,5,5,)-hydroxy-11-{alpha}, 9 {alpha}-epoxymethano prosta 5Z, 13E-dienoic acid; Sigma, St. Louis, MO] was used in final concentrations ranging between 10–10 and 10–4 M (29). KCl concentrations ranged from 25 to 200 mM. A single agonist was added for each culture well. After agonist addition, intracellular calcium concentration was recorded for 150 s. Agonist was removed by washing twice with 400 µl of HBSS +0.1% BSA. A postwash stable baseline was also recorded to ensure no shift occurred.

Calcium channel and thromboxane receptor blockade. Those pathways known to regulate calcium mobilization in response to agonist were examined by incubation of pulmonary arterial myocytes with individual calcium channel blockers for 20 min at room temperature following fura-2 AM loading and before addition of U-46619. Nifedipine (1 µM) was used to selectively block sarcolemmal L-type calcium channels. Ryanodine (100 µM) was used to block sarcoplasmic reticulum (SR) ryanodine-sensitive channels, and xestospongin (20 µM) was used to block 1,4,5-trisphosphate (IP3)-gated channels (1). Cells were challenged with 10–6 M U-46619, and calcium mobilization responses were recorded. In some experiments, to confirm specificity of receptor activation, the thromboxane A2-selective prostanoid receptor (TP-R) was blocked by incubation with 10–5 M SQ-29548 for 20 min at room temperature before addition of 10–6 M U-46619.

Mobilization of specific intracellular calcium pools. Non-TP-R-linked SR calcium mobilization was quantified with 20 µM ATP to stimulate P2X and P2Y receptors. Maximal calcium mobilization via P2Y was considered an estimate of the SR IP3-gated calcium pool. P2Y response was isolated by removal of extracellular calcium; P2X response was isolated by blockade of IP3-gated channels with 20 µM xestospongin. The SR ryanodine-sensitive calcium pool was quantified by mobilization with 5 µM ryanodine and/or 40 mM caffeine.

Calcium mobilization data analysis. Background fluorescence was measured from cell-free areas and subtracted from total fluorescence before analysis. No more than eight equally sized, square regions containing three to five cells were selected for the presence of minimal cell-free areas from each microscope field, before quantification of calcium responses. Whole cell Ca2+ mobilization was analyzed en bloc for each region. Ca2+ traces were discarded only if there was not a stable baseline before agonist addition or if the 340-nm excitation value dropped at the same time as the 380-nm excitation value, as this would give erroneous peak [Ca2+]i values with no physiological merit. Peak [Ca2+]i response was defined as the maximal point of displacement of the rapid calcium transient; as no sustained plateau was apparent in cultured myocyte calcium responses, all subsequent calculations reference peak [Ca2+]i mobilization. Maximum change in [Ca2+]i was calculated as the average baseline value subtracted from the peak [Ca2+]i response to agonist. The time to peak [Ca2+]i from the point of agonist addition was also documented. The 340/380 nm excitation ratio values were converted to nanomolar [Ca2+]i values on a standard curve where F is fluorescence intensity at specified excitation wavelength and R represents fluorescence intensity ratio of F340nm/F380nm. Rmin is the minimum ratio, Rmax is maximum ratio, and Ka is the calcium association constant of the indicator; these three values were predetermined from an in vitro calibration curve linearly plotting log Ca2+ concentration vs. the log of the F340nm/F380nm fluorescence ratio after subtraction of background at each wavelength, generated using free fura-2 and calcium standards and calculated by the method of Grynkiewiez (24):

Data from individual microscope regions were analyzed by unpaired t-test or one-way ANOVA, with individual comparisons performed by Tukey's honestly significant difference test. Data are expressed as means ± SE; P < 0.05 was considered significant.

Thromboxane receptor (TP-R) localization. PASMCs from newborn piglets were grown to semiconfluence on glass coverslips in 12-well dishes at a plating density of 4.4 x 104 cells/cm2. Cells were rinsed free of culture media with cytoskeletal buffer (containing in mM: 10 MES, 150 NaCl, 5 EGTA, 5 MgCl2, and 5 glucose) and then fixed with either methanol for 10 min to study cell surface TP-R or 3% paraformaldehyde for 15 min at room temperature followed by permeabilization with 0.3% Triton X-100 for 5 min for study of intracellular TP-R. Cells were rinsed twice with CB buffer and stored in Cyto-TBS (composition in mM; 20 Tris, 154 NaCl, 2 EGTA, and 2 MgCl2) at 4°C. Nonspecific binding was blocked by incubation with 10% normal donkey serum in Cyto-TBS + 1% BSA for 20 min at room temperature. PASMCs were then incubated with TP-R rabbit polyclonal antibody (1:50 in Cyto-TBS + 1% BSA, Cayman Chemical) overnight at 4°C, followed by incubation with FITC-conjugated donkey anti-rabbit antibody (1:50 in Cyto-TBS + 1% BSA, Jackson ImmunoResearch Lab) for 2 h at room temperature. Nuclei were counterstained with Hoechst 33342. Fluorescence immunocytochemistry images were acquired using an Olympus 1X 70 microscope with wavelengths of 494 nm excitation/518 nm emission for TP-R, and 346 nm excitation/460 nm emission for nuclei. Images were then analyzed with an UltraPix FSI digital camera and UltraView software (Perkin Elmer). Total TP-R protein abundance was also assayed by Western blot in whole cell lysates from confluent myocytes in hypoxic and normoxic culture. Membrane fractions were obtained by whole cell lysate ultracentrifugation at 150,000 g for 30 min at 4°C. Samples of 20 µg of protein separated by SDS-PAGE and blotted onto nitrocellulose membrane were then probed for TP-R. Equality of protein loading was ensured by reprobing blots for {beta}-actin. Visualization of protein bands was by enhanced chemiluminescence (ECL; Amersham, Piscataway, NJ). The band representing native TP-R protein was selected on the basis of migrating molecular weight; two additional adjacent bands were selected based on known weights of glycosylated TP-R (25, 52). We quantified data with a digital imaging densitometer, under nonsaturating conditions, with background subtraction achieved by reading the absorbance of an equal sized region directly adjacent to the band. Summated glycosylated and unglycosylated TP-R band densities were normalized to {beta}-actin band density and represented as proportional to the control value. Data are presented graphically as means ± SE for a minimum of three replicate samples.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
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Hypoxic exposure and induction of PPHN. The in vivo hypoxic environment for PPHN animals (HA) was maintained at FIO2 9.96 ± 1.37%. Control piglets (NA) exhibited a developmentally appropriate decrease in right ventricle-to-left ventricle plus septum weight ratio over the first 3 days of life. The development of early pulmonary hypertension in hypoxic piglets was determined by an increased right-to-left cardiac ventricular weight ratio (P < 0.01), predominantly owing to increased right ventricular weight due to right heart afterloading (Fig. 2).



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  		Fig. 2. Hypoxic pulmonary hypertension diagnosed by relative increase in right-to-left ventricular weight ratio. Pulmonary hypertension is diagnosed by increased cardiac weight ratio of right ventricle to left ventricle plus septum (RV/LV+S; blotted tissue) following 3 days exposure to moderate normobaric hypoxia, in day 3 hypoxic pigs (HA, n = 11) compared with day 3 control pigs (NA, n = 9). Day 0 control pigs (birth, n = 11) presented as maturational control. #P < 0.05 compared with birth; *P < 0.05 compared with NA.

 
Calcium mobilization in pulmonary arterial myocytes to U-46619. We used the following four cell culture models: 1) NA were PASMCs derived from normal 3-day-old pigs, in primary culture at 21% O2; 2) HA were PASMCs derived from 3-day-old piglets with hypoxic PPHN, in primary culture at 21% O2; 3) HM were PASMCs derived from newborn pigs, in primary culture with incubation in 10.76 ± 0.14% O2 for the final 3 days of culture; and 4) NM were control PASMCs derived from newborn pigs, in primary culture at 21% O2.

PASMC basal (unstimulated) intracellular calcium levels were significantly elevated following in vitro hypoxic exposures, compared with normoxic controls (Fig. 3A). Intracellular Ca2+ levels were increased in myocytes from NA and HA. Pulmonary arterial myocytes exposed to hypoxic conditions, both in vivo (HA) and in vitro (HM), exhibited significantly higher peak Ca2+ mobilization response to 10–6 M U-46619 than did control normoxic myocytes from newborn (NM) and 3-day pigs (NA) (Figs. 3B and 2C), despite the very different protocols used for hypoxic exposure in vivo compared with in vitro. Peak calcium mobilization was enhanced in HM cells over a wide range of U-46619 doses, compared with NM cells (Fig. 3D). The myocyte response to U-46619 in all conditions was completely blocked by preincubation with the TP-R inhibitor SQ-29548 (Fig. 3E), indicating receptor specificity of the U-46619 response. The time from agonist addition to peak [Ca2+]i, in cells exposed to hypoxia in vitro or in cells cultured from animals exposed to hypoxia in vivo, was significantly faster (P < 0.01) than in their age-matched normoxic control cells at lower doses of U-46619 (10–7 M) (Fig. 4A). However, this difference between treatment groups was not apparent at higher U-46619 doses (10–6 M), suggesting saturability (Fig. 4B).



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  		Fig. 3. Myocyte Ca2+ mobilization response to U-46619. A: pulmonary artery smooth muscle cells (PASMCs) exposed to hypoxia for 3 days (HM, n = 120) have significantly increased resting [Ca2+]i compared with normoxic controls (NM, n = 120). Baseline [Ca2+]i in PASMC from 3-day normoxic pigs (NA, n = 65) and from animals exposed to 3-day in vivo hypoxia (HA) are also significantly increased compared with normoxic control (n = 120); *P < 0.01. B: representative traces of Ca2+ mobilization (in µM) in response to 10–6 M U-46619 stimulation (arrow) of myocytes derived from neonatal pigs raised in hypoxia for 3 days in vivo and then cultured in normoxia (HA), myocytes derived from 3-day normoxic pigs and grown in normoxia (NA), myocytes derived from newborn pigs and exposed to in vitro hypoxia for 3 days (HM), and control myocytes derived from newborn pigs and cultured in normoxia (NM). C: peak [Ca2+]i responses of HA cells (n = 30) and HM cells (n = 45) to 10–6 M U-46619 are significantly higher than in NM cells (n = 37) and NA cells (n = 41); *P < 0.01. D: peak [Ca2+]i in HM and NM myocytes in response to a range of U-46619 doses. E: representative traces of NM and HM responses to 10–6 M U-46619 after preincubation with thromboxane A2-selective prostanoid receptor (TP-R) inhibitor SQ-29548 (arrow), demonstrating receptor specificity of agonist. Numerical data quantified in microscopic regions containing 3–5 PASMCs loaded with fura-2 AM (n = number of microscope regions).

 


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  		Fig. 4. Time to peak Ca2+ mobilization response after agonist addition. A: low-dose U-46619 (10–7 M) results in significantly slower peak response time in normoxic control myocytes (NM, n = 24) compared with myocytes exposed to hypoxia in vitro (HM, n = 24). PASMCs from 3-day normoxic piglets grown in normoxia (NA, n = 26) also have a significantly slower peak response time than myocytes exposed to hypoxia in vivo but cultured in normoxia (HA, n = 38) (*P < 0.01). B: at higher dose (10–6M), no difference in peak U-46619 response time between NM (n = 29), HM (n = 24), NA (n = 40), and HA (n = 32) groups (n = number of microscope regions).

 
Sensitivity of myocyte responsiveness to U-46619 was determined by dose-response curves normalized to maximal [Ca2+]i peak within each treatment group. Hypoxia induced sensitization of myocytes to U-46619 (Fig. 5A). Dose-response curves for U-46619 were significantly left shifted in hypoxia, resulting in a lower EC50 for agonist-induced calcium mobilization (2.1683 x 10–6 ± 0.07751 M in HM cells, compared with NM cells EC50 of 1.5422 x 10–5 ± 0.06030 M, P < 0.0001). In contrast, the EC50 of calcium mobilization in response to membrane depolarization by KCl was not altered by hypoxic exposure in vivo or in vitro (Fig. 5B), and peak calcium responses to KCl were not significantly altered.



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  		Fig. 5. Dose-response curves for peak Ca2+ mobilization to KCl and U-46619. A: U-46619 dose response curve for in vitro hypoxic myocytes (HM, n = 3) is significantly left shifted compared with the curve for normoxic control cells (NM, n = 3); P < 0.0001. Peak Ca2+ mobilization is normalized within each group as percent of maximal agonist response. B: KCl dose response curve for NM cells (n = 3) is not significantly different from the curve for HM (n = 3); EC50 = 60.66 mM vs. 79.23 mM, P = not significant (n = number of animals).

 
Pathway of calcium mobilization. HM PASMCs incubated in a Ca2+-free environment (Fig. 6A) maintained an increased peak Ca2+ response to 10–6 M U-46619 compared with NM PASMCs, as did HM PASMCs preincubated with nifedipine (Fig. 6B) before agonist challenge. However, a significant decrease in HM peak Ca2+ mobilization (P < 0.01) occurred with both ryanodine (Fig. 6C) and xestospongin (Fig. 6D), dropping the agonist response to a level similar to that of NM cells.



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  		Fig. 6. Pathway of hypoxic myocyte hyperreactivity to U-46619 determined by serial calcium channel blockade. Ca2+ mobilization responses to 10–6 M U-46619 in normoxic controls (NM, n = 84) and in vitro hypoxic myocytes (HM, n = 62). A: stimulation with U-46619 after removal of extracellular calcium increases Ca2+ mobilization globally, but the difference between treatment groups is unaffected (*P < 0.01); note this histogram has a different y-axis scale than the ones following. [Ca2+]ext, extracellular calcium concentration. B: stimulation with U-46619 after preincubation with nifedipine (L-type voltage-operated calcium channel blockade) results in unaltered calcium mobilization in hypoxia and normoxia (*P < 0.01). C: stimulation with U-46619 after preincubation with 100 µM ryanodine [blockade of sarcoplasmic reticulum (SR) ryanodine-sensitive Ca2+ channel] decreases Ca2+ mobilization in hypoxic myocytes to normoxic levels and obliterates difference between treatment groups. D: stimulation with U-46619 after preincubation with xestospongin [SR 1,4,5-trisphosphate (IP3)-gated channel blockade] also decreases Ca2+ mobilization in hypoxic myocytes to normoxic levels, obliterating difference between treatment groups (n = number of microscope regions; *P < 0.01).

 
Intracellular calcium pools in hypoxia. Ca2+ mobilization via ATP-stimulated release from P2Y/IP3-linked stores was unaltered by in vitro hypoxic exposure, suggesting no increase in the intracellular IP3-gated Ca2+ pool (Fig. 7B). Total ATP-induced Ca2+ mobilization was also unaltered in hypoxic vs. normoxic myocytes (Fig. 7, A and C). Direct stimulation of Ca2+ release from ryanodine-sensitive stores was negligible in cultured PASMCs (Fig. 7D).



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  		Fig. 7. Thromboxane receptor-independent calcium pool mobilization from intracellular SR stores. Peak Ca2+ mobilization responses in normoxic control (NM, n = 32) and in vitro hypoxic PASMCs (HM, n = 40). A: 20 µM ATP stimulation of both P2X and P2Y receptors. B: ATP stimulation of P2Y receptors alone, achieved by removing [Ca2+]ext. C: ATP stimulation of P2X receptors alone, achieved by preincubation with 20 µM xestospongin blocking IP3-gated SR channels. No differences in maximal Ca2+ release to ATP in HM vs. NM, P = NS. D: representative trace: responses to 5 µM ryanodine (Ry, left arrow) are absent in both NM and HM myocytes (similar responses obtained to 40 µM caffeine, not shown). Subsequent stimulation with 10–6 M U-46619 (right arrow) confirms viability.

 
TP-R localization. Normoxic myocytes have ample cell surface TP-R (Fig. 8A), as well as internalized TP-R distributed uniformly in the cytoplasm (Fig. 8C). In HM, there is a significant decrease in cell surface TP-R signal (Fig. 8B). Permeabilization by paraformaldehyde (PFA) fixation reveals decreased TP-R immunostaining, with translocation of the receptor protein to the perinuclear region in HM (Fig. 8D). Total protein abundance of TP-R is stable in hypoxic whole cell lysates compared with the normoxic group, as demonstrated by Western blot (Fig. 8E). TP-R abundance in the membrane fraction is decreased in HM compared with NM controls; however, this is not significant statistically (Fig. 8F).



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  		Fig. 8. Thromboxane receptor localization is altered by hypoxic exposure. Representative photomicrographs using a polyclonal thromboxane receptor antibody in NM (N = 3) and HM (N = 3) PASMCs. Intact cell immunocytochemistry using methanol fixation reveals cell surface TP-R is more abundant in normoxic myocytes (A) than in hypoxic myocytes (B). Immunostaining after cell permeabilization by paraformaldehyde (PFA) fixation indicates cytoplasmic TP-R is widely distributed in normoxic cells (C), while translocating to the perinuclear region in hypoxic PASMCs (D). E: total cell TP-R protein abundance, normalized to {beta}-actin remains constant in hypoxia (HM, n = 9; NM, n = 9). Membrane (memb) TP-R protein content, normalized to whole cell TP-R, is diminished in HM (n = 8) compared with NM (n = 8) (F), but this change does not achieve statistical significance (N = number of pigs, n = number of samples).

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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In this study we examined the distinct phenomena of hypoxia-mediated sensitization (lower threshold response to agonist) and hyperreactivity (increased response elicited by a given dose of agonist or elevation of the plateau or maximal level of response) (21) to thromboxane receptor stimulation. Smooth muscle hyperresponsiveness has been ascribed to altered states of myocyte calcium handling, and our data are consistent with that paradigm (44). Exposure of neonatal pulmonary arterial myocytes to moderate hypoxia for 3 days in vitro results in both sensitization to the thromboxane mimetic U-46619 and hyperreactivity as measured by elevated calcium mobilization. We have found: 1) 3 days of hypoxic exposure in vitro results in an increased basal Ca2+ level, heightened peak Ca2+ response, faster peak Ca2+ response, and a decreased EC50 for U-46619; 2) these increases persist when PASMCs are exposed to hypoxia in vivo and then cultured in 21% O2; 3) the phenomenon is not ubiquitous but is specific for thromboxane; and 4) hypoxia-induced myocyte thromboxane hyperreactivity occurs in the face of decreased cell surface thromboxane receptor expression, suggesting a mechanism of increased receptor sensitivity. Mechanisms of altered calcium mobilization were studied primarily in primary pulmonary arterial myocyte cultures using an in vitro environmental manipulation model. An in vivo model of hypoxic PPHN followed by myocyte culture in normoxia was also employed to look specifically at extinction of hypoxic alterations in thromboxane response, upon removal of the hypoxic stimulus and proliferation under control conditions. Myocyte monoculture is reliable for study of intracellular calcium mobilization in hypoxia, which correlates with the accrual of early phase tension, and is unaffected by absence of endothelium (48). Whereas others have reported agonist-induced pulmonary arterial calcium mobilization to be diminished following chronic hypoxic exposure of adult rats, it is likely that myocyte agonist insensitivity and loss of calcium oscillations following prolonged hypoxic exposure may relate more to phenotypic alteration associated with pulmonary arterial remodeling (4).

Our data indicate that the elevated peak calcium response in hypoxic pulmonary arterial myocytes is mediated by increased calcium release via SR Ca2+ stores (14, 35); other calcium sources may contribute to the overall response, but their relative contribution is not increased in hypoxia, and inhibition of SR calcium mobilization decreases thromboxane-induced calcium transients in hypoxic myocytes to normoxic levels. Thromboxane mimetic U-46619-induced calcium response is known to be redundant, eliciting both intracellular and extracellular calcium mobilization (15, 38, 53). Thromboxane inhibits Kv channel current, leading to myocyte depolarization, activation of L-type Ca2+ channels, and vasoconstriction of pulmonary arteries (7); simultaneously, pharmacomechanical coupling occurs via a G protein-coupled mechanism (30), as well as calcium sensitization of smooth muscle mediated by Rho-kinase (22). It is probable that augmented SR Ca2+ release is the sole source of the elevated calcium response to U-46619 observed in hypoxia. In vascular smooth muscle cells, IP3-gated and ryanodine-sensitive SR Ca2+ channels may access separate intracellular Ca2+ pools that alter with phenotype, resulting in altering pathway of Ca2+ mobilization with phenotypic modulation (56). Unchanged Ca2+ mobilization to ATP in hypoxic myocytes attests that the PASMC IP3-gated pool is not independently increased in hypoxia. The increased hypoxic PASMC response to U-46619 may therefore be due to altered receptor-mediated IP3 generation upstream. Ryanodine-sensitive Ca2+ pools are not amenable to direct stimulation in our model and therefore likely contribute only to calcium-induced calcium release after primary IP3-gated channel activation. Enhanced IP3 generation, as reported in hypoxic pulmonary, but not systemic, arterial fibroblasts (45), may contribute toward this mechanism.

A limitation in interpretation of this study may lie in the U-46619 EC50 value ranges. The concentrations of U-46619 and SQ-29548 used in these experiments are derived from dose-response curves cited elsewhere in the literature (34, 41). EC50 data can be dependent upon the preparation type and the specific outcome being measured. U-46619 EC50 values in the nanomolar range have been reported in radioligand assays using cell membrane preparations. Much higher values are also reported (in the micromolar range) for U-46619 EC50 when contraction of an isolated arterial ring preparation is examined (3) and near micromolar range for myocyte calcium response (7). A low-affinity TP-R has been described in cultured vascular myocytes, with EC50 in the micromolar range (34). The dose-response and EC50 values we have reported here are robust with repeated measurement, and we can merely conclude they may be characteristic of our cultured myocyte preparation.

Intracellular calcium measurement is employed in this study as a proxy for myocyte contraction. The relationship has been well established between [Ca2+]i, myosin light chain (LC20) phosphorylation, and isometric force development to thromboxane analog stimulation. In guinea pig aortic strips, steady-state force for contractions stimulated by U-46619 assume a similar dependence on LC20 phosphorylation to potassium depolarization, both in the presence and absence of extracellular calcium. A [Ca2+]i/force relation indicates U-46619 stimulates greater isometric force at lower [Ca2+]i than does KCl depolarization, suggesting increased calcium sensitivity, but the myocyte LC20 phosphorylation/force relationship is unchanged by U-46619 compared with potassium-induced depolarization (32). As calcium sensitivity as well as muscle preload determine developed force at a given degree of [Ca2+]i elevation, calcium mobilization responses to U-46619 cannot be compared directly to those induced by potassium depolarization (40); however, comparisons of myocyte calcium transients generated in response to any one specific agonist may be proportionally related (13, 14).

Of note, pulmonary arterial myocytes in our preparation responded to agonist challenge with a sharp, rapid calcium transient consisting of a peak, a rapidly sloping shoulder without defined plateau, and a clear return to baseline free calcium values within the 4–5 min of observation. A more prolonged period of observation did not reveal any alteration in this pattern of response; while the slope of the shoulder occasionally varied, no plateau or sustained calcium elevation was evident, and all traces returned to the preagonist baseline. As such, we focused our attention on the peak calcium transient as the primary indicator of myocyte responsiveness as this constituted virtually the entire response. This early calcium spike (<20 s) is known to correlate with peak myosin phosphorylation, maximal shortening velocity, and the majority of isotonic shortening (28) and hence is of physiological significance for change in vascular diameter. There is reasonable precedent for the measurement of peak calcium mobilization alone in determining pulmonary arterial smooth muscle agonist response (31, 46), and we have analyzed our calcium response data accordingly.

We observed elevated resting [Ca2+]i in hypoxic myocytes, as has been reported by others and ascribed to both SR release and capacitative calcium entry (12, 33). Altered membrane polarity and voltage-gated calcium channel activity do not contribute to increased calcium mobilization at this oxygen tension, as evidenced by an unchanged EC50 of KCl in agonist-naïve myocytes. These data are consistent with the finding of graded inhibition of outward K+ current and depolarization of resting membrane potential only at hypoxia <5% O2 (43).

We also observed increased sensitivity and reactivity in cells derived from neonatal pigs exposed to hypoxia in vivo but then cultured for 15 days under normoxic conditions, indicating a persistent alteration in myocyte agonist response that is not extinguished after 2 wk removal from the sensitizing stimulus. This is in contrast to reports of prompt reversibility of hypoxia-induced alterations in electrophysiological properties on return to normoxia (5, 6). The persistence of altered calcium mobilization in cultured myocytes exposed to moderate in vivo hypoxia remotely in time suggests a durable change in thromboxane receptor function and pharmacomechanical coupling that is not extinguished by subsequent normoxic exposure. This finding may carry grave implications for pulmonary circuit responsiveness to inflammatory mediators after the resolution of perinatal hypoxia. Altered gene expression driving persistent functional changes during recovery from hypoxia is not a novel concept. Expression of hypoxia-susceptible genes, including hypoxia-inducible factor, is key to pulmonary vascular remodeling (9, 47). It is known that smooth muscle recovery from hypoxia lags behind endothelial recovery (55) and that pulmonary arterial smooth muscle proliferation, apoptosis, and protein synthesis remain altered during recovery from hypoxia (26).

We postulate that hypoxic sensitization to thromboxane agonist occurs at the level of the thromboxane receptor and its associated IP3-linked pathway, in view of the left shift in agonist dose response and the increased rapidity of response at lower agonist doses; the loss of this difference at higher doses may indicate receptor saturability. The data presented here do not yet establish whether altered thromboxane receptor density or avidity may be responsible for the observed myocyte agonist sensitization. However, we demonstrate evidence of thromboxane receptor translocation from the cell surface to a perinuclear localization in hypoxia; this may represent a negative-feedback mechanism governing receptor cycling in the context of agonist hyperreactivity. An increase in receptor-ligand binding and, therefore, in receptor avidity may be implied. It would be pertinent to further evaluate hypoxia-induced sensitization with receptor-binding studies; these fall beyond the scope of this paper.

Hypoxia-induced thromboxane agonist hypersensitivity and hyperreactivity of neonatal pulmonary arterial myocytes may have a synergistic effect on intracellular calcium transients, contributing to the pulmonary vasoconstrictor response to arachidonic acid observed in perinatal hypoxia (20). In addition, we have previously reported calcium sensitization of pulmonary arterial smooth muscle in hypoxic PPHN, due to downregulation of myosin light chain phosphatase activity (10), which would be markedly potentiated by augmented agonist-induced calcium mobilization.

The relationship of hypoxia and inflammation is crucial in the time course and severity of PPHN. We conclude that hypoxia has a priming effect upon the neonatal pulmonary arterial myocyte: hypoxic exposure results in an increased basal Ca2+ level, heightened peak Ca2+ response, faster peak Ca2+ response, and a decreased EC50 for thromboxane. These increases are mediated by altered pharmacomechanical coupling specific to this agonist, despite a reduction in the cell surface TP-R population. Myocyte thromboxane hyperresponsiveness persists in culture after removal from the initiating hypoxic stimulus, suggesting an alteration at the level of gene expression. We infer that pulmonary thromboxane production, in response hypoxia, or to inflammatory stimuli including ventilator-induced trauma, may further potentiate neonatal hypoxic vasoconstriction and cause pulmonary circuit vasospasm even after resolution of hypoxia in PPHN.


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This research was funded in part by The Winnipeg Rh Foundation Institute and Manitoba Medical Services Foundation.


   FOOTNOTES
 

Address for reprint requests and other correspondence: S. Dakshinamurti, Sect. of Neonatology, WS012 Women's Hospital, 735 Notre Dame Ave., Winnipeg, Canada R3A 1R9 (e-mail: dakshina{at}cc.umanitoba.ca)

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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M. Hinton, A. Gutsol, and S. Dakshinamurti
Thromboxane hypersensitivity in hypoxic pulmonary artery myocytes: altered TP receptor localization and kinetics
Am J Physiol Lung Cell Mol Physiol, March 1, 2007; 292(3): L654 - L663.
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