Pulmonary microvascular and macrovascular endothelial cells: differential regulation of Ca2+ and permeability

J. J. Kelly1, T. M. Moore1, P. Babal2, A. H. Diwan2, T. Stevens1, and W. J. Thompson1

Departments of 1 Pharmacology and 2 Pathology, University of South Alabama College of Medicine, Mobile, Alabama 36688

    ABSTRACT
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Cytosolic Ca2+ concentration ([Ca2+]i) plays an important role in control of pulmonary vascular endothelial cell (ECs) barrier function. In this study, we investigated whether thapsigargin- and ionomycin-induced changes in cytosolic Ca2+ induce permeability in rat pulmonary microvascular (RPMV) versus macrovascular (RPA) ECs. In Transwell cultures, RPMVECs formed a tighter, more restrictive barrier than RPAECs to 12,000-, 72,000-, and 150,000-molecular-weight FITC-labeled dextrans. Thapsigargin (1 µM) produced higher [Ca2+]i levels in RPAECs than in RPMVECs and increased permeability in RPAEC but not in RPMVEC monolayers. Due to the attenuated [Ca2+]i response in RPMVECs, we investigated whether reduced activation of store-operated Ca2+ entry was responsible for the insensitivity to thapsigargin. Addition of the drug in media containing 100 nM extracellular Ca2+ followed by readdition media with 2 mM extracellular Ca2+ increased RPMVEC [Ca2+]i to a level higher than that in RPAECs. Under these conditions, RPMVEC permeability was not increased, suggesting that [Ca2+]i in RPMVECs does not initiate barrier disruption. Also, ionomycin (1.4 µM) did not alter RPMVEC permeability, but the protein phosphatase inhibitor calyculin A (100 nM) induced permeability in RPMVECs. These data indicate that, whereas increased [Ca2+]i promotes permeability in RPAECs, it is not sufficient in RPMVECs, which show an apparent uncoupling of [Ca2+]i signaling pathways or dominant Ca2+-independent mechanisms from controlling cellular gap formation and permeability.

microvasculature; fluorescein isothiocyanate-labeled dextran; gap formation; thapsigargin; ionomycin

    INTRODUCTION
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

THE PULMONARY MICROVASCULAR endothelium is a dynamic barrier that is critical for lung gas exchange and regulation of fluid and solute passage between the blood and interstitial compartments in the lung. Derangement of this barrier increases vascular permeability and contributes to the hypoxemia associated with adult respiratory distress syndrome and noncardiogenic pulmonary edema. Experimental models using whole animal and isolated lung preparations have shown that multiple inflammatory mediators cause increased vascular permeability and edema. Furthermore, studies of cultured conduit endothelial cell (EC) monolayers have implicated various inflammatory mediators in the control of EC barrier function. However, little is known about responses of microvascular ECs (MVECs) from the pulmonary circulation to these inflammatory mediators due to the difficulty and technical limitations in harvesting and maintaining MVECs in culture. Recent data on pulmonary MVECs show improved monolayer permeability (10) compared with previous results using conduit cell isolates (4, 33, 34). Moreover, comparative studies from bovine endothelium have shown that pulmonary MVECs form a more restrictive barrier to macromolecules than to pulmonary artery or vein ECs, suggesting phenotypic differences (9, 35, 43).

The relationship between cytosolic Ca2+ concentration ([Ca2+]i) and barrier dysfunction is well studied in the literature. Typically, agents that elevate [Ca2+]i cause increased vascular permeability in intact lungs (6, 19) and cultured cells (14, 21, 22). Moreover, elevations in [Ca2+]i result in activation of several cellular events, including activation of Ca2+/calmodulin-dependent myosin light chain kinase (MLCK; see Refs. 14, 16, 24, 39, 49), F-actin reorganization (16, 32), and decreased cell-matrix and cell-cell tethering (16, 22, 25, 26, 36), all of which may ultimately lead to EC gap formation and increased permeability.

Despite the collective evidence that [Ca2+]i promotes development of intercellular gaps, recent studies have suggested that increases in [Ca2+]i alone may not induce permeability changes (5, 17). We have previously shown that the MLCK pathway is not required in rat pulmonary (RP) MVEC permeability (10). These data suggest that the Ca2+-dependent processes recognized in control of pulmonary artery EC (PAEC) permeability may not play a major role in pulmonary MVEC (PMVEC) permeability, providing a possible explanation for the tighter barrier that is associated with MVECs. These observations may in part be attributable to phenotypic differences in the ECs studied. Therefore, the purpose of this study was to investigate critical determinants of RPAEC and RPMVEC barrier function. Specifically, experiments were designed to investigate whether activation of store-operated Ca2+ entry, previously shown to increase permeability in isolated perfused rat lungs, increases [Ca2+]i in RPAECs and RPMVECs sufficient to increase macromolecular permeability. Our data demonstrate that, compared with RPAECs, RPMVECs 1) generate a more restrictive barrier to macromolecules and 2) do not generate intercellular gaps and increased permeability in response to increases in [Ca2+]i.

    MATERIALS AND METHODS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Materials. Heparin, thapsigargin (TG), and ionomycin were purchased from Calbiochem. FBS was from Intergen. Rat tail collagen IV and ECs growth supplement were obtained from Collaborative Biomedical Products. Chambered glass coverslips were from Lab Tek. Microspheres (50 µm) were obtained from NEN. RPMI 1640, serum-free Aim V medium, gentamicin, and viokase were purchased from GIBCO-BRL. Rat epidermal growth factor was from Biomedical Technologies. DMEM was obtained from Cellgro Mediatech. Fura 2-AM and pluronic acid were from Molecular Probes. HEPES was from Research Organics. Hyaluronidase I, Krebs-Henseleit buffer, FITC-dextran (average mol wt 12,000, 72,000, and 150,000), bovine serum albumin (BSA), and all other analytical reagents were purchased from Sigma.

Culture and isolation of RPMVECs. RPMVECs were isolated and cultured in our laboratory (11) using a modified method of Ryan et al. (33). Briefly, rat lungs (male Sprague-Dawley, 350-400 g) were initially perfused without recirculation with 300 ml of Krebs-Henseleit buffer containing 8% BSA (19). The perfusate was changed to serum-free Aim V medium containing 8% BSA, 50 mg of hyaluronidase I, and collagen IV-coated 50-µm microspheres in a volume of 100 ml. The direction of perfusate flow was alternated from anterograde to retrograde (0.03 ml · min-1 · g body wt-1), and perfusate containing the EC-bound microbeads was collected on ice. The microbeads with cells were washed three times with RPMI containing 25% FBS and resuspended in RPMI containing 20% rat serum, 0.1% gentamicin, and EC-conditioned medium (2:1). After pure primary cultures were established, cells were grown in 20% FBS-DMEM containing 100 U/ml penicillin, 0.1 mg/ml streptomycin, 25 µg/ml fungizone, and 10 µg/ml gentamicin until passage 10 and with 10% FBS thereafter. The cell stocks were harvested by scraping without proteases for passage until used for experimentation. Cells were studied between passage 4 and passage 20.

RPMVECs grow initially forming strings and characteristic capillary-like structures before assuming typical cobblestone morphology at confluence. Cells were further characterized as MVECs through collective criteria of factor VIII antigen antibody staining, uptake of 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine-labeled low-density lipoprotein (DiI-acetylated LDL) and intense binding of fluorescent Ulex europaeus (UEA-I), Ricinus communis (RCA-I), and Arachis hypogaea (PNA) lectins (9, 11, 35).

Isolation and culture of RPAEC. Male Sprague-Dawley rats (350-400 g) were killed with a pentobarbital sodium overdose. The heart and lungs were removed, and the pulmonary arterial segment between the heart and lung hili was dissected, split, and fixed onto a 35-mm petri dish. ECs were obtained by gentle intimal scraping with a plastic cell lifter and were seeded on a 100-mm petri dish containing 10 ml of medium (F-12 nutrient mixture and DMEM ~1:1 supplemented with 10% FBS, 100 U/ml penicillin, and 0.1 mg/ml streptomycin). After 1 wk, contaminating smooth muscle cell colonies were marked and removed by pipette aspiration. Primary cells were allowed to grow and were passaged at confluency by trypsin digestion into T75 culture flasks. EC phenotype was characterized by factor VIII antigen antibody staining, uptake of DiI-acetylated LDL, and negativity for smooth muscle alpha -actin. Standard tissue culture techniques were performed until the cells were used for experimentation. Cells were studied between passage 4 and passage 20.

Permeability assay. RPMVECs and RPAECs at preconfluence were washed two times in Ca2+-free PBS and treated with viokase (1.0 g/l) for 3 (RPMVECs) or 5 min (RPAECs) at 37°C. Upon resuspension in fresh medium (10% FBS in DMEM/DMEM + F-12), cells were seeded (65-80,000/insert) on Transwell insert polycarbonate filters (6.5-mm diameter, 0.4-µm pore size). Insert filters containing ECs formed the base of a top chamber suspended over a bottom well (2 cm2). Fresh medium (1 ml) was added into the top wells every 2 days after careful removal of medium. Monolayers were studied 4-5 days postseeding.

Permeability assays to assess barrier function of monolayers were performed using a modified protocol to that described by Diwan et al. (10). Briefly, RPMVEC and RPAEC monolayers were incubated for 15 min at 37°C with 1 ml of 25 mM HEPES-DMEM (pH 7.4) added to the top well. The HEPES-DMEM was then carefully pipetted from the top well, and the transport was initiated by the addition of 1 mg/ml (in a volume of 100 µl) of FITC-labeled dextran in 25 mM HEPES-DMEM to the top well. The top well was removed to a fresh bottom well (in a 24-well plate) containing 0.7 ml of 25 mM HEPES-DMEM. This procedure ensures cell contact with media above and below the monolayer and equivalent media levels on the top and bottom. Three different molecular-weight ranges of FITC-labeled dextran [~12,000, 72,000, and 150,000 molecular weight (Mr)] were used throughout the study. Aliquots (50 µl) were removed from the bottom well at the times specified for each experiment and collected in white 96-well plates for fluorescence analysis. Drugs or vehicles were either mixed directly before the experiment in the dextran-media mixture or added directly to the Transwell.

FITC-labeled dextran was measured in a luminescence spectrometer (LS 50B; Perkin-Elmer), using 480 and 530 nm as the excitation and emission wavelengths, respectively. Calculations were made for molecular-weight dextran polymers to accommodate the FITC content per mole of glucose that varied between batches. Relative permeability is expressed as FITC fluorescence of the sample separated into aliquots minus background fluorescence. A fluorescence value of 200 is ~8, 7, and 5 µg/ml for the low, medium, and high Mr dextrans. Reflection coefficients for control, TG-treated, and calyculin-treated monolayers were calculated for 12,000- and 72,000-molecular-weight dextrans according to Suttorp et al. (46) without the use of tritiated water, since a minimal hydrostatic pressure is generated by the Transwell chamber.

Scanning electron microscopy. Transwells with cultured ECs were washed with PBS, fixed in 3% glutaraldehyde in 0.2 M cacodylate buffer for 2 h, washed in the same buffer, and dehydrated in a series of ethyl alcohol washes. The membranes were critical-point dried in CO2, covered with a 15-nm-thick gold-palladium layer, and evaluated in a Philips XL-20 scanning electron microscope operating at 10 kW. Specimens were observed at 15° inclination.

[Ca2+]i measurement by fura 2 fluorescence. RPAECs and RPMVECs were seeded onto four-chambered glass coverslips (Nunc) at ~100-150 × 103 cells and were grown to confluency (2 days). [Ca2+]i was estimated using the Ca2+-sensitive fluorophore fura 2-AM according to methods previously reported (43). Briefly, ECs were washed with 2 ml HEPES-Krebs, pH 7.4. Loading was for 20 min using a solution of 1 ml of buffer containing 3 µM fura 2-AM, 0.03% pluronic acid, and 2 mM CaCl2 in a CO2 incubator at 37°C followed by wash with 2 ml of buffer with 2 mM nM CaCl2 for 20 min. [Ca2+]i was measured using an Olympus IX70 inverted microscope at ×400 with a xenon arc lamp photomultiplier system (Photon Technologies). Data were calculated and analyzed with PTI Felix software. Epifluorescence was measured from three to four ECs in a confluent monolayer, and the changes in [Ca2+]i were expressed as the fluorescence ratio of the Ca2+-bound (340 nm) to Ca2+-unbound (380 nm) excitation wavelengths emitted at 510 nm. [Ca2+]i were extrapolated from calibration curves plotted with known Ca2+ concentrations.

Ca2+ add-back experiments employed a similar protocol except cells in chambered coverslips were preincubated in HEPES-Krebs buffer containing 100 nM Ca2+ for 15 min. Buffer was replaced, and fluorescence tracing was initiated. TG was added at 4 min, and HEPES-Krebs buffer containing 2 mM Ca2+ was added ~6 min later.

Extracellular Ca2+ effect on monolayer permeability. RPMVECs and RPAECs were grown on Transwell inserts as described above and preincubated in 25 mM HEPES-Krebs buffer (pH 7.4) for 15 min. The experiment was initiated after buffer removal, addition of 0.5 mg/ml FITC-dextran (Mr ~12,000) in HEPES-Krebs containing either 2 mM (control) or 100 nM CaCl2 to the top well (100 µl), and transfer to a fresh bottom well containing HEPES-Krebs buffer (0.7 ml). At 15 min, TG (1 µM) mixed with FITC-dextran in buffer containing either 2 mM (control) or 100 nM (100 µl) CaCl2 was added to the upper chamber. At 45 min, CaCl2 (2 mM) was added back to wells initially containing 100 nM Ca2+. Aliquots (50 µl) were removed from the bottom well at 15, 30, 45, 60, and 75 min. The passage of dextrans across the cell monolayer was measured by fluorescence, as described above, except in black 96-well plates.

    RESULTS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Morphology of the endothelial monolayers. Scanning electron microscopy of the pulmonary artery and microvascular endothelial monolayers grown on Transwells showed striking differences in surface morphology. RPMVECs expressed a high density of surface projections. RPAECss displayed a much smoother surface with a scatter of the fingerlike projections. Furthermore, the MVECs showed tight intercellular connections, whereas the macrovascular endothelium formed membrane associations with visible gaps between the cells (Fig. 1).


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Fig. 1.   Scanning electron microscopic photographs of rat pulmonary artery (RPA; A) and rat pulmonary microvascular (RPMV; B) endothelial cells (ECs) in culture on semipermeable membranes. Immediately after termination of permeability experiments (see Fig. 2), Transwells with cultured ECs were washed in PBS and fixed in 3% glutaraldehyde, as described in MATERIALS AND METHODS. Cell junctions (arrowheads) of RPAECs formed visible gaps unlike the RPMVEC tight intercellular junctions. MVECs showed a greater number of dense hairy fingerlike cytoplasmic protrusions on their surface compared with RPAECs. Bar = 10 µm.

Basal permeability of RPAEC and RPMVEC monolayers to macromolecules. To examine differences in barrier properties between macrovascular ECs and MVECs in culture, the passage of different molecular-weight dextran polymers was monitored in RPAEC and RPMVEC monolayers. Both cell types formed monolayers that demonstrated a time- and size-dependent flux of the different dextran polymers (Fig. 2) from upper to lower chambers. Under basal conditions, RPMVEC monolayers allowed passage of fewer 12,000 (Fig. 2A)-, 72,000 (Fig. 2B)-, and 150,000 (Fig. 2C)-molecular-weight FITC-dextrans compared with RPAEC (P < 0.05). RPAEC monolayers exhibited the same increased flux of dextrans even with extended time in Transwell culture (data not shown). The permeability ratio for the different-size dextran molecules (12,000:72,000:150,000) was calculated for RPMVECs (8:2:1) and RPAECs (4:2:1), suggesting that RPMVECs are less permeable to larger (>72,000 mol wt) molecules relative to smaller (12,000 mol wt) molecules than RPAECs, as shown previously for rat (10) and bovine cells (9, 35).


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Fig. 2.   Comparison of RPAEC and RPMVEC monolayer permeability to a range of FITC-labeled dextrans [12,000 (A), 72,000 (B), and 150,000 (C) mol wt; 1 mg/ml] applied to the luminal aspect of confluent monolayers of RPAECs and RPMVECs. Barrier function ("basal leak") was determined by the accumulation of different-size dextrans subluminally in the lower chamber at 30, 60, and 120 min. Relative permeability is defined as the fluorescence of FITC-labeled dextrans measured at 480-530 nm. See MATERIALS AND METHODS for further details. RPAEC and RPMVEC monolayers used in these experiments were grown for 6 days. Data are expressed as means ± SE of 3 separate experiments performed in triplicate (n = 9).

Effect of TG on RPAEC and RPMVEC permeability. Due to the superior barrier properties displayed by RPMVECs, experiments were performed to measure the functional significance of this barrier to compounds known to alter ECs shape and increase permeability. TG, a plant alkaloid that inhibits microsomal Ca2+-ATPase activity to increase [Ca2+]i and activates store-operated Ca2+ entry, has been reported to increase vascular permeability in the intact lung (6, 18) and change RPAECs shape in cultured cells (data not shown). Treatment of RPAECs monolayers with TG (1 µM) had a profound effect on barrier function (Fig. 3A), increasing monolayer permeability to both 12,000- and 72,000-molecular-weight dextran polymers in a time-dependent manner (P < 0.05). The calculated reflection coefficients were 0.85, 0.71, and 0.36 versus 0.97, 0.92, and 0.82 at 30, 60, and 120 min for TG-treated and control RPAECs, respectively, or similarly 0.98, 0.95, and 0.88 versus 0.99, 0.99, and 0.97 for 12,000- and 72,000-molecular-weight dextrans, respectively. Conversely, treatment of RPMVECs monolayers with TG (1 µM) had no effect on macromolecular permeability (Fig. 3B). Reflection coefficients for 12,000-molecular-weight dextrans were 0.99, 0.98, and 0.94 for TG-treated and control RPMVECs at 30, 60, and 120 min, respectively. Reflection coefficients of 0.99 were calculated at each time point for 72,000-molecular-weight dextrans regardless of treatment. Furthermore, increasing the concentration of TG to 15 µM did not significantly alter RPMVEC permeability to FITC-dextrans (data not shown). RPMVECs appeared resistant to changes in [Ca2+]i.


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Fig. 3.   Effect of thapsigargin on RPAEC (A) and RPMVEC (B) monolayer barrier function. FITC-labeled dextran polymers (12,000 and 72,000; 1 mg/ml) were applied to the luminal aspect of confluent monolayers of RPAECs and RPMVECs in the presence (black-square, bullet ) and absence (square , open circle ) of 1 µM thapsigargin. Relative permeability was determined at 30, 60, and 120 min by sampling of the lower chamber and measuring the appearance of dextran by fluorescence at 480-530 nm. See MATERIALS AND METHODS for further details. RPAEC and RPMVEC monolayers used in these experiments were grown for 4 and 5 days, respectively. Data are expressed as means ± SE of 3 (RPAECs) and 6 (RPMVECs) separate experiments performed in triplicate (n = 9 and n = 18, respectively).

Effect of TG on intracellular Ca2+ measurements. Because TG did not increase permeability in RPMVECs, we determined whether TG was capable of increasing [Ca2+]i in these cells. As shown in Fig. 4A, under normal physiological Ca2+ levels (2 mM), baseline ratios of 340- to 380-nm wavelengths in fura 2-loaded cells were the same in RPAECs (0.9 ± 0.02) and RPMVECs (0.9 ± 0.02). TG produced a smaller increase in [Ca2+]i levels in RPMVECs compared with RPAECs, with peak ratios of 2.7 ± 0.2 and 4.3 ± 0.2, respectively (Fig. 4B). Furthermore, the increase in [Ca2+]i in RPMVECs was more transient, returning toward baseline levels by the end of the experiment (15 min), whereas the TG-induced increase in RPAEC [Ca2+]i was maintained at a plateau of 3.0 ± 0.2 (Fig. 4B).


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Fig. 4.   Store-operated Ca2+ entry in RPAECs and RPMVECs. [Ca2+]i was estimated in confluent monolayers using fura 2 ratios as described in MATERIALS AND METHODS. A: typical trace showing 340/380 fluorescence obtained from 4-5 cells within a confluent monolayer. Resting Ca2+ in both ECs was 65 nM. RPAEC peak Ca2+ was 727 nM, and RPMVEC peak Ca2+ was 323 nM. RPAEC Ca2+ showed a plateau at 382 nM, and RPMVEC Ca2+ returned to basal at later time points. B: data are expressed as means ± SE of 5 separate experiments performed as described in A. RPAEC and RPMVEC monolayers used in these experiments were grown for 4 and 3 days, respectively.

To determine whether the lower [Ca2+]i levels in RPMVECs after TG treatment were responsible for the lack of a permeability response, experiments were designed to maximally activate extracellular Ca2+ entry. In low extracellular Ca2+ concentration ([Ca2+]e) the application of TG to cells diminishes the rise in [Ca2+]i but facilitates Ca2+ entry upon readdition of [Ca2+]e. One explanation of the differences in Ca2+ transients of Fig. 4 is reduced store-operated channel activity in RPMVECs. Therefore, in an attempt to elevate RPMVEC [Ca2+]i to levels comparable to those measured in RPAECs that increase EC permeability, we added TG to cells in buffer containing 100 nM [Ca2+]e. In these conditions, baseline ratios were reduced to 0.67 ± 0.03 (RPAECs) and 0.73 ± 0.02 (RPMVECs), and TG (1 µM) had little initial effect on RPAEC and RPMVEC [Ca2+]i (0.9 ± 0.2 and 1.0 ± 0.1, respectively; Fig. 5). Subsequent readdition of [Ca2+]e to RPAECs increased [Ca2+]i to a peak ratio of 4.6 ± 0.3 and plateau 3.3 ± 0.2. Readdition of [Ca2+]e to RPMVECs increased [Ca2+]i to a peak ratio of 7.0 ± 0.9 and plateau of 3.8 ± 0.4, 2.5-fold higher than the response measured in 2 mM extracellular Ca2+ (Fig. 4A). Thus, under these experimental conditions, RPMVEC [Ca2+]i achieved a level that, in RPAECs, increased permeability.


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Fig. 5.   Effect of extracellular Ca2+ concentration on store-operated Ca2+ entry in RPAECs and RPMVECs. [Ca2+]i was estimated in confluent monolayers using fura 2 ratios, as described in MATERIALS AND METHODS. A: typical traces comparing the [Ca2+]i response in RPAECs and RPMVECs to thapsigargin (TG; 1 µM) in the absence of [Ca2+]o (100 nM) followed by the addition of extracellular Ca2+ (2 mM). Resting Ca2+ in both ECs was 55 nM. RPAEC peak Ca2+ after TG was 59 nM, and RPMVEC peak Ca2+ was unchanged. After readdition of 2 mM medium Ca2+, RPAECs showed a peak value of 836 nM internal Ca2+. RPMVECs peaked off scale at >1 µM internal Ca2+. RPAECs showed a plateau at 448 nM after stimulation, whereas RPMVECs decreased linearly to 574 nM at termination. RPAEC and RPMVEC monolayers used in these experiments were grown for 4 and 3 days, respectively. B: data from the tracings shown in A are expressed as means ± SE of 5 separate experiments.

Role of extracellular Ca2+ on permeability. Compared with RPAECs, RPMVECs have a significantly reduced influx of Ca2+ to TG, evidenced by the magnitude of both the peak and plateau [Ca2+]i responses (Fig. 4, A and B). However, under the experimental conditions of Fig. 5, the influx of Ca2+ in response to TG in RPMVECs was increased to levels higher than in RPAECs. Therefore, under similar conditions, permeability was studied with TG to determine whether reduced store-operated Ca2+ entry prevented TG-induced permeability in RPMVECs. TG (1 µM) did not increase permeability in RPAECs in the absence of [Ca2+]e but increased permeability significantly upon readdition of 2 mM CaCl2 (Fig. 6A). Likewise, TG (1 µM) did not increase RPMVEC monolayer permeability in the absence of [Ca2+]e. However, permeability was not stimulated with the readdition of [Ca2+]e (Fig. 6B) even though [Ca2+]i levels were shown to be higher than in RPAECs (Fig. 5). The data suggest the involvement of store-operated Ca2+ entry in TG-induced permeability in RPAECs and also suggest that RPMVECs regulate cellular Ca2+ entry and/or extrusion differently.


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Fig. 6.   Effect of store-operated Ca2+ entry on RPAEC (A) and RPMVEC (B) barrier function. FITC-dextran polymers (12,000 molecular weight; 1 mg/ml) were applied to the luminal surface of confluent monolayers, and the passage of dextran was monitored by sampling of the lower chamber at the times indicated. TG (1 µM) or vehicle was added in the absence (~100 nM) of extracellular Ca2+. Thirty minutes later, extracellular Ca2+ (2 mM) was readded to monolayers. Relative permeability is defined as the fluorescence of FITC-labeled dextrans measured at 480-530 nm. See MATERIALS AND METHODS for more details. RPAEC and RPMVEC monolayers used in these experiments were grown for 5 and 4 days, respectively. Data are expressed as means ± SE of 3 separate experiments performed in triplicate (n = 9).

Effect of Ca2+ sensitivity on RPMVEC permeability. The ineffectiveness of TG to induce permeability in RPMVECs suggested two other possibilities as follows: 1) desensitization of the molecular components of permeability to the [Ca2+]i response or 2) the TG-induced increased [Ca2+]i is compartmentalized (or extruded) separate from molecular targets that regulate permeability. To assess whether the TG-induced increase in [Ca2+]i is compartmentalized in RPMVECs, we performed permeability experiments with the Ca2+ ionophore ionomycin, which nonselectively elevates [Ca2+]i. In RPMVECs, 1.4 µM ionomycin increased internal Ca2+ to >1 µM in <30 s, and Ca2+ remained elevated for 16 min (unpublished observations). Ionomycin (1.4 µM) did not significantly increase permeability of RPMVECs (Fig. 7), emphasizing the possible uncoupling of [Ca2+]i from MVEC gap formation and permeability.


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Fig. 7.   Effect of ionomycin on RPMVEC monolayer barrier function. FITC-labeled dextran polymers (12,000 and 72,000 mol wt; 1 mg/ml) were applied to the luminal aspect of confluent monolayers of RPMVECs in the presence (square , open circle ) and absence (black-square, bullet ) of 1.4 µM ionomycin. Relative permeability was estimated at 30, 60, and 120 min by sampling of the lower chamber and measuring the appearance of dextran by fluorescence at 480-530 nm. See MATERIALS AND METHODS for further details. Monolayers used in these experiments were grown for 4 days. Data are expressed as means ± SE of 8 (12,000 mol wt) or 3 (72,000 mol wt) separate experiments performed in triplicate (n = 24 and n = 9, respectively).

As previously reported, the protein phosphatase inhibitor calyculin A (Cal A) had marked effects on RPMVEC monolayer barrier function through a Ca2+-independent mechanism involving the phosphorylation of proteins (10) other than myosin light chain (MLC). Consistent with these data, Cal A (100 nM) increased permeability significantly after 60 min to both 12,000- and 72,000-molecular-weight FITC-dextran polymers (Fig. 8). Because protein phosphatase inhibitors did not initiate Ca2+ transients (data not shown), these results suggest that Ca2+-independent mechanisms can regulate gap formation and permeability in MVECs. Reflection coefficients for transport of 12,000-molecular-weight dextrans were 0.98, 0.80, and 0.11 for Cal A treated at 30, 60, and 120 min versus 0.99, 0.98, and 0.96 for untreated RPMVEC monolayers. For 72,000-molecular-weight dextrans, the coefficients were 0.99, 0.95, and 0.68 and 0.99, 0.99, and 0.99 for Cal A treated and control cells, respectively.


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Fig. 8.   Effect of calyculin A on RPMVEC monolayer barrier function. FITC-labeled dextran polymers (12,000 and 72,000 mol wt; 1 mg/ml) were applied to the luminal aspect of confluent monolayers of RPMVECs in the presence (square , open circle ) and absence (black-square, bullet ) of 100 nM calyculin A. Relative permeability was estimated at 30, 60, and 120 min by sampling of the lower chamber and measuring the appearance of dextran by fluorescence at 480-530 nm. See MATERIALS AND METHODS for further details. Monolayers used in these experiments were grown for 4 days. Data are expressed as means ± SE of 3 separate experiments performed in triplicate (n = 9).

    DISCUSSION
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

In vivo models of clinical noncardiogenic pulmonary edema suggest that most fluid filtration occurs in the microcirculation. However, mostly from technical limitations in the isolation and culture of MVECs, the majority of in vitro data on mechanisms controlling pulmonary EC barrier properties has been derived from conduit ECs. These data indicate that increased [Ca2+]i and, in particular, store-operated Ca2+ entry promote barrier disruption (6). Over the years, experiments designed utilizing cell-cultured ECs have contributed enormously to the current understanding of endothelium barrier properties by enabling studies to be performed that were impossible in the intact lung. The present studies were designed to study permeability responses and Ca2+ signaling pathways in MVECs in relation to PAECs. These studies were made possible using a technique to isolate RPMVECs modified from Ryan et al. (11, 33). The results show that, unlike ECs cultured from conduit vessels, elevated [Ca2+]i by three different processes (store-operated entry, intracellular store release, and ionomycin-induced extracellular influx) is uncoupled from cellular events governing gap formation and permeability in pulmonary MVECs. This conclusion is consistent with published in vivo and in vitro evidence showing that pulmonary MVECs (bovine) differ phenotypically from PAECs (9, 34, 35). Earlier studies have also shown differences in pulmonary MVEC permeability from ECs derived from other sources (4).

Both RPMVEC and RPAEC cultures showed many similarities, including typical EC biochemical characteristics, cobblestone morphology at confluence, and monolayer formations on Transwells. However, MVEC cultures also showed remarkable differences in surface morphology at the scanning electron microscopy level (7). Furthermore, RPMVECs formed tighter intercellular junctions than RPAECs, consistent with the findings in bovine macrovascular ECs and MVECs (35), suggesting that RPMVECs and RPAECs are functionally and phenotypically distinct EC populations. It is possible that the isolation and culture procedures themselves may have created different phenotypes because it is not possible to prepare each culture absolutely identically. Although the earliest passages are treated with trypsin for harvesting, the serum required for growth and harvesting is different. Nevertheless, both types of EC cultures are harvested with viokase and seeded and grown to monolayers on Transwells with identical procedures for 3-4 days, which decreases the likelihood of culture-induced phenotypic artifacts. Although RPMVECs appear distinct from RPAECs in culture, the origin of the cells cannot be ensured as venous, arteriole, or capillary but are likely similar to MVECs in the intact lung. Because intact lungs show osmotic reflection coefficients of near 0.95 using isotope techniques (47), the calculated values of 0.99 for the different dextrans in both RPAECs and RPMVECs are supportive of this conclusion.

The functional correlate to these structural differences was that RPMVECs formed a tighter and a more restrictive barrier compared with RPAECs. The difference was evident even with extended time in culture after confluency. Furthermore, at any given time point, the permeability ratios for both EC types for the three sizes of dextrans (8:2:1 vs. 4:2:1 for microvascular vs. pulmonary artery, respectively) revealed that, while RPAECs transported more of each size dextran bead, RPMVECs appear to form smaller intercellular gaps that under basal conditions are less permeable to larger molecules and therefore relatively more permeable to smaller molecules. Similar comparative observations were reported for bovine lung microvessel, vein, and artery ECs (34, 35) and calf retina and aortic ECs (50). These data are consistent with our previous studies that showed our MVEC cultures formed barriers with superior integrity (10, 44), as defined by a full spectrum of hydroxyl ethyl starch particle sizes. Our permeability comparisons contribute to an increasing body of data, suggesting that MVECs from various vascular origins form tighter, more restrictive barriers to macromolecules than do macrovascular ECs.

An increase in vascular permeability in response to inflammatory agents may result from different general mechanisms, including 1) Ca2+-driven active contraction mediated by actin-myosin interaction dependent on MLC [20-kDa MLC (MLC20)] phosphorylation (13, 20, 39), 2) loss of cell-cell junctional integrity (26), 3) loss of cell-matrix tethering (36), or 4) Ca2+-independent pathways (5, 17, 42). In our present studies, TG was used as one of three protocols to address whether elevated Ca2+ could produce increased macromolecular permeability in ECs cultured from the RP microcirculation compared with RPAECs that are capable of changing shape and forming intercellular gaps in response to TG (data not shown). In addition, TG has been shown to increase solvent permeability in isolated rat (6) and dog lungs (18) as indicated by an increased microvascular filtration coefficient (Kfc). The studies reported here show that, despite increased intracellular Ca2+ induced by TG in both types of EC monolayers, the drug increased dextran permeability in RPAEC monolayers but not macromolecular permeability in similarly treated RPMVEC monolayers. The findings with the microvascular cells, but not with the macrovascular cells, appear to conflict with data published with isolated and perfused whole lung preparations in which Kfc is thought to measure microvascular permeability. Possible explanations for the apparent discrepancies are 1) Transwell permeability assays did not measure fluid flux and do not include factors of flow and pressure that would be present in intact lung permeability experiments. Thus, our experiments do not exclude an increase in solvent permeability in either type of EC monolayer to TG treatment that could be analogous to the intact lung solvent permeability detected with Kfc; 2) increased protein permeability in response to TG has not been measured in intact lung preparations, precluding direct comparisons with RPMVEC or RPAEC dextran permeabilities; and 3) as indicated above, both EC monolayers are cultured similarly, but the cells are not isolated by identical procedures. Therefore, RPMVECs, but not RPAECs, may respond differently in culture than in vivo. Although our data identify permeability differences between macrovascular ECs and MVECs, a direct extrapolation of our findings in vitro to observations from whole lung studies requires further study.

EC [Ca2+]i response to neurohumoral inflammatory mediators is generally characterized by two interrelated phases (3, 30): Ca2+ release from intracellular stores and subsequent Ca2+ entry across the cell membrane. ECs have no voltage-sensitive Ca2+ channels, and TG-induced EC permeability requires store-operated Ca2+ entry across the cell membrane. The TG-induced [Ca2+]i responses in both types of ECs showed an initial increase followed by a more sustained transient (1, 2, 26, 31). The Ca2+ transients obtained with the fura 2 ratio method differed in amplitude, as the MVEC response was less than the RPAEC response. Similar differences in cytosolic Ca2+ responses to TG were found with the indo 1 Ca2+ fluoroprobe using bovine pulmonary MVECs compared with bovine conduit artery ECs (44), but permeability was not determined in that study. Therefore, although numerous investigators have shown that a rise in [Ca2+]i in response to various stimuli initiates vascular permeability in conduit ECs (12, 23, 32, 37), the lack of TG-induced permeability in RPMVECs could be due to Ca2+ regulation differences reflected as lower [Ca2+]i. Regulation could include increased Na+/Ca2+ exchanger activity, increased membrane Ca2+-ATPase reuptake, Ca2+-mediated inhibition of Ca2+ entry channels, or unidentified Ca2+ seqestration or compartmentalization. Identification of mechanisms modifying store-operated Ca2+ entry and the possibility of an uncoupling of store-operated Ca2+ entry channels in RPMVECs await future studies.

To investigate whether the reduced TG-induced store-operated Ca2+ entry in RPMVECs is responsible for the inability of TG to cause permeability in the cell, a different protocol was used. TG was applied to each EC culture in the presence of low [Ca2+]e to facilate Ca2+ entry upon readdition of [Ca2+]e, a technique known to highly activate store-operated Ca2+ (8, 29, 30). TG applied in the presence of 100 nM [Ca2+]e had little effect on [Ca2+]i in RPMVECs or RPAECs. However, upon the readdition of [Ca2+]e, [Ca2+]i levels in RPMVECs increased to levels comparable to those measured in RPAECs. Importantly, the levels were equal to those where Ca2+-induced permeability was observed in RPAECs. This suggests that, under these experimental conditions, if increased [Ca2+]i is coupled to gap formation in RPMVECs, TG should induce permeability. However, no increase in permeability was observed.

Because increased permeability was not observed with either protocol in RPMVECs, we tested the hypothesis that TG-induced increases in RPMVEC [Ca2+]i may be compartmentalized, thus preventing a permeability response. The Ca2+ ionophore ionomycin did not increase permeability in RPMVECs at drug concentrations known to increase Ca2+ to levels required to induce permeability in bovine PAECs (45) and canine kidney epithelial cells (>1 µM; see Ref. 36). Because equivalent [Ca2+]i levels were induced by ionomycin, the cellular events governing RPMVEC contraction and gap formation may be uncoupled from cytosolic Ca2+. Higher concentrations of ionomycin (>10 µM) did increase the passage of dextrans through RPMVEC monolayers (data not shown). Similar observations have recently been reported in bovine PMECs where ionomycin (>5 µM) inhibited tyrosine kinase (p125FAK) phosphorylation via cAMP-mediated protein kinase A attenuation (15). These data suggest that ionomycin-induced permeability in PMECs may be due to decreased cell-cell and cell-matrix tethering and not via development of increased centripetal tension.

Further studies tested whether a [Ca2+]i-independent stimulus (i.e., not via Ca2+-activated MLCK) would increase permeability in RPMVECs. We have previously demonstrated that the protein phosphatase inhibitors Cal A and okadaic acid increase RPMVEC permeability independent of MLC20 phosphorylation (10). Similar permeability changes were measured with Cal A as agonist. Macrovascular EC contraction via MLC20 phosphorylation and subsequent cytoskeletal reorganization and intercellular gap formation have been shown by others to be dependent on elevated [Ca2+]i (13, 14, 20, 39). Thrombin-induced permeability in bovine PAECs (13) and human umbilical vein ECs (13, 38) occurs via a rapid increase in [Ca2+]i followed by phosphorylation of MLC20. Thrombin at similar concentrations had no effect on RPMVEC permeability (data not shown), suggesting a fundamental difference in the way RPMVECs control permeability. Because Ca2+-independent pathways are known for PAECs, including protein kinase C- and toxin-mediated permeability (42, 28), secondary pathways that may be overridden in larger vessel ECs might be dominant in microvascular cells as a means of cytoprotection and permeability control.

To conclude, our data suggest that microvascular and macrovascular ECs are phenotypically different cells with respect to their ability to form a semipermeable barrier. More importantly, it appears that Ca2+ signaling pathways described in macrovascular ECs are either more stringently regulated or uncoupled in MVECs from the cellular events controlling gap formation and permeability. Ca2+-independent mechanisms may contribute to RPMVEC permeability. Although speculative, our data indicate that signaling mechanisms that govern conduit and MVEC permeability in vitro may differ. However, the significance of the mechanisms controlling cultured cell monolayers in relation to those occurring in intact lungs requires further investigation.

    ACKNOWLEDGEMENTS

We thank Raquel Dien and Gina Capley for tissue culture and George Brough for technical assistance and proofreading of the manuscript. We also thank Dr. Judy King and Freda McDonald for help in preparation of the scanning electron micrographs.

    FOOTNOTES

This research was supported by National Heart, Lung, and Blood Institute Grants HL-46494 (to W. J. Thompson) and HL-56050 (to T. Stevens), by a Parker B. Francis Pulmonary Fellowship (to T. Stevens), and American Heart Association Alabama Affiliate Fellowships (to J. J. Kelly and T. M. Moore).

Address for reprint requests: W. J. Thompson, Dept. of Pharmacology, Univ. of South Alabama College of Medicine, MSB 3100, Mobile, AL 36688.

Received 10 September 1997; accepted in final form 2 February 1998.

    REFERENCES
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

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