Direct ANP inhibition of hypoxia-induced inflammatory pathways in pulmonary microvascular and macrovascular endothelial monolayers

D. C. Irwin,1 M. C. Tissot van Patot,2 A. Tucker,1 and R. Bowen1

1Department of Biomedical Sciences, College of Veterinary and Biomedical Sciences, Colorado State University, Fort Collins, and 2Department of Anesthesiology, University of Colorado Health Science Center, Denver, Colorado

Submitted 5 August 2004 ; accepted in final form 16 December 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Atrial natriuretic peptide (ANP) has been shown to reduce hypoxia-induced pulmonary vascular leak in vivo, but no explanation of a mechanism has been offered other than its vasodilatory and natriuretic actions. Recently, data have shown that ANP can protect endothelial barrier functions in TNF-{alpha}-stimulated human umbilical vein endothelial cells. Therefore, we hypothesized that ANP actions would inhibit pulmonary vascular leak by inhibition of TNF-{alpha} secretion and F-actin formation. Bovine pulmonary microvascular (MVEC) and macrovascular endothelial cell (LEC) monolayers were stimulated with hypoxia, TNF-{alpha}, or bacterial endotoxin (LPS) in the presence or absence of ANP, and albumin flux, NF-{kappa}B activation, TNF-{alpha} secretion, p38 mitogen-activated protein kinase (MAPK), and F-actin (stress fiber) formation were assessed. In Transwell cultures, ANP reduced hypoxia-induced permeability in MVEC and TNF-{alpha}-induced permeability in MVEC and LEC. ANP inhibited hypoxia and LPS increased NF-{kappa}B activation and TNF-{alpha} synthesis in MVEC and LEC. Hypoxia decreased activation of p38 MAPK in MVEC but increased activation of p38 MAPK and stress fiber formation in LEC; TNF-{alpha} had the opposite effect. ANP inhibited an activation of p38 MAPK in MVEC or LEC. These data indicate that in endothelial cell monolayers, hypoxia activates a signal cascade analogous to that initiated by inflammatory agents, and ANP has a direct cytoprotective effect on the pulmonary endothelium other than its vasodilatory and natriuretic properties. Furthermore, our data show that MVEC and LEC respond differently to hypoxia, TNF-{alpha}-stimulation, and ANP treatment.

altitude; atrial natriuretic peptide; endothelial cell permeability; tumor necrosis factor-{alpha}; lipopolysaccharide


ACUTE HYPOXIA OR INFLAMMATORY AGENTS increase vascular permeability and contribute to forms of noncardiogenic pulmonary edema such as high-altitude pulmonary edema (HAPE) and acute respiratory distress syndrome (3, 6, 9, 10, 18, 44). Despite extensive research into the effects of acute hypoxia or inflammation on the pulmonary circulation, the mechanisms underlying noncardiogenic pulmonary edema remain unclear (3, 12). In vivo models of lung injury have revealed that atrial natriuretic peptide (ANP) can protect endothelial barrier function (1416, 43), but apart from its vasodilatory and natriuretic actions, no other mechanisms have been identified to explain the cytoprotective benefits of ANP. Recently, ANP was shown to inhibit TNF-{alpha} synthesis in murine macrophages stimulated with bacterial endotoxin (LPS) (19). ANP also has been shown to inhibit NF-{kappa}B and p38 mitogen-activated protein kinase (MAPK) pathways in TNF-{alpha}-stimulated human umbilical vein endothelial cells (HUVEC) in vitro (25, 26). ANP actions are highly cell type and organ specific (31, 41, 45), and no experimental work has addressed ANP actions on cultured pulmonary microvascular (MVEC) and macrovascular endothelial cells (LEC) exposed to acute hypoxia or inflammatory agents.

We hypothesized that ANP could preserve pulmonary endothelial barrier integrity during hypoxic or inflammatory stress by inhibiting the cytokines that initiate a signal cascade, or the proteins within the signal transduction pathway, to alter cell morphology, cause gap formation, and increase permeability. Our approach was to examine the effect of ANP on TNF-{alpha} secretion, NF-{kappa}B activity, and activation of p38 MAPK in cultured MVEC and LEC exposed to hypoxia, TNF-{alpha}, or LPS. The goal was to determine whether ANP protects cells from stress fiber formation, preserving cell morphology, barrier function, and endothelial cell permeability. Our study was designed to determine the effects of ANP on hypoxia-, TNF-{alpha}-, and LPS-treated pulmonary endothelial cells and to further our knowledge of endothelial permeability in the pulmonary vasculature.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials

ANP was obtained from Calbiochem (King of Prussia, PA), collagenase II (CLS2) was obtained from Worthington Biomedical (Lakewood, NJ), retina-derived growth factor was obtained from Vec Tec (Rensselaer, NY), nylon mesh was obtained from Sefar America (Depew, NY), and 1,1'-dioctadecly-3,3,3',3'-tetramethylindocarbocyanine (DiI-acetylated LDL) was obtained from Biomedical Technologies (Stoughton, MA). FITC-phalloidin (Alexa Fluor 488 phalloidin) was obtained from Molecular Probes (Eugene, OR). All other materials were purchased from Sigma (St. Louis, MO).

Culture and Isolation of Bovine Pulmonary MVEC

MVEC were isolated and cultured in our laboratory. Briefly, bovine lung tissue was harvested locally, transported (4°C) to our laboratory, and harvested within 4 h. Pleura was removed from the periphery of an external lobe, and small pulmonary vessels (diameter ~200 µm) were dissected, minced, and digested with collagenase (10 ml of 1,000 U/ml collagenase II, 5% BSA; rocker, 37°C, 30 min). The digested tissue was then filtered (160-µm nylon mesh) and centrifuged (200 g, 5 min). The pellet was resuspended in culture medium (RPMI containing 20% FBS, 70 µg/ml porcine intestinal heparin, 6 µg/ml retina-derived growth factor, 0.1% gentamicin, and 0.1% fungizone), washed three times, and resuspended in plating medium (3 parts RPMI + 1 part endothelial cell conditioned medium). Cells were divided equally into six-well gelatin-coated (1%) plates (~104 cells/well) and placed in a standard cell culture incubator (37°C, 5% CO2). After 1 wk, cell populations exhibiting cobblestone morphology were removed and seeded onto new 6-cm gelatin-coated plates and grown to confluence. Cell populations appearing clear of contaminating fibroblast and smooth muscle cells were sorted by standard fluorescence-activated cell sorting (FACS) analysis by using the uptake of DiI-acetylated LDL to obtain MVEC populations >98% pure (18, 37).

Culture and Isolation of Bovine Pulmonary Artery LEC

Endothelial cells were isolated from bovine conduit pulmonary arteries (diameter ~3 cm) obtained locally as previously described (37). Briefly, vessels were cut longitudinally, and the internal surface was digested with collagenase (0.1% collagenase type 1A; 30 min, 37°C). Endothelial cells were gently removed with a cell scraper and cultured in standard cell culture conditions (medium: D-valine MEM supplemented with 20% FBS, 70 µg/ml porcine intestinal heparin, 0.1% gentamicin, and 0.1% amphotericin B). Primary cultures were grown to confluence, and populations appearing clear of contaminating fibroblasts and smooth muscle cells were sorted using FACS analysis as previously described to obtain LEC populations >98% pure (18, 37).

Characterization of MVEC vs. LEC

Lectin binding. As previously described (27), the lectin glycine max preferentially binds MVEC vs. LEC. Therefore, we performed an agglutination test with the lectin glycine max. Briefly, glycine max (1:1,000) was added to confluent MVEC and LEC in 6-mm dishes and incubated for 10 min. The cells were trypsinized and titrated to ensure single-cell suspension and were then resuspended in PBS. A drop from each tube was applied to a glass microscope slide and viewed under a microscope. Glycine max preferentially bound to MVEC (data not shown).

Cell growth. As previously described (27), MVEC have a faster rate of growth compared with LEC. Therefore, to further characterize our endothelial cell phenotypes, we determined the growth rate of our MVEC and LEC. Briefly, endothelial cells were seeded at 1 x 105 cells/well in 6-cm culture dishes as described above (n = 2) and counted every 24 h for 4 days after the seeding date; cells were resuspended using trypsin and counted using a hemocytometer. Both LEC and MVEC exhibited a lag phase followed by exponential growth. MVEC displayed a faster rate of growth (5.75 ± 0.25 x 105 vs. 4.2 ± 0.25 x 105 cells at day 4, respectively).

Experimental Design

For all experiments, MVEC and LEC were used from the same passages (510), grown to confluence over the same time frame, and cultured under standard cell culture conditions unless otherwise noted. Cell preparations were untreated or treated with ANP (10–8, 10–7, 10–6 M) 30 min before endothelial cell stimulation with hypoxia, TNF-{alpha}, or LPS, unless noted.

In Vitro Hypoxia Model

Cell preparations were placed into a special humidified hypoxic chamber equipped with a thermostat set at 37°C and an antechamber to pass supplies in and out. Use of the antechamber ensured that once cells became hypoxic, they were never exposed to a normoxic environment. The chamber utilized a positive pressure system and was supplied with a gas mixture of 3% O2, 5%CO2, and a balance of nitrogen. All culture media used in permeability tests were allowed to equilibrate to 3% O2 by being placed in the hypoxic chamber ~2 h before initiation of the test. All relevant permeability tests and monolayer fixations were conducted within the hypoxic chamber to avoid the introduction of normoxia during tissue preparation.

Permeability Assay

MVEC and LEC cell preparations were either left untreated or stimulated with hypoxia or TNF-{alpha} (25 ng/ml) in the presence or absence of ANP or the p38 MAPK inhibitor SB-202190 (10–6 M). After 24 h, the permeability rate was determined. Briefly, MVEC and LEC were seeded (~100,000 cells/insert) on gelatin-coated (1%) polystyrene filters (Costar Transwell, no. 3470, 6.5-mm diameter, 0.4-um pore size; Cambridge, MA). Endothelial cells were grown to confluence on Transwell inserts over 3 days. After cells were washed three times in serum-free medium, 50 µM FITC-labeled albumin suspended in serum-free medium was added to the endothelial cell monolayers (100 µl). The insert was placed in a new well of a 24-well plate containing serum-free medium (0.6 ml to ensure that the fluid volume on either side of the inserts was equalized to avoid a hydrostatic gradient that might alter the rate of albumin flux). The transfer rate of albumin across the monolayer was assessed by measuring the rise in FITC-albumin in the lower well after 60 min. FITC-albumin was quantified in a fluorescence spectrofluorophotometer (Mithras LB940; Berthold Technologies, Oak Ridge, TN). As previously described (13), albumin flux F across the monolayer was expressed as F = (d[A]2/dt x V)/S (in mol·s–1·cm–2), where V is volume in the abluminal well, d[A]2 is rise of albumin concentration in the bottom well during the time interval dt, and S is monolayer surface area. To facilitate the comparison of data obtained from our study with those of others, the permeability coefficient (P, measured in cm/s) of the combined system of monolayer and filter support can be calculated from F according to Fick's law of diffusion as follows: P = F/([A]1 – [A]2), where [A]1 and [A]2 are tracer concentrations in the luminal and abluminal compartments, respectively (13). The permeability coefficient P under standard culture conditions was 1.71 x 10–6 and 1.49 x 10–6 cm/s for MVEC and LEC, respectively. Data for mean P are expressed as percentages of a defined control situation (i.e., LEC under standard culture conditions).

TNF-{alpha} Secretion

MVEC and LEC monolayers were pretreated with ANP (10–7 M, 30 min), and then cells were stimulated with hypoxia for 24 h or LPS (1 µg/ml) for 6 h. Supernatants were collected and frozen at –70°C for TNF-{alpha} bioassay analyses.

TNF-{alpha} Bioassay

This assay is based on quantification of the cytotoxic activity of TNF-{alpha} on L929 cells in the presence of actinomycin D (19, 36). The mitochondrial reduction of 3,4,5-(dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) to formazan was determined as an indicator of L929 cell viability (19, 36). Briefly, the supernatants described in TNF-{alpha} Secretion were added to confluent L929 cells and incubated for 24 h at 37°C. MTT (5 mg/ml) was added to each well and incubated (4 h at 37°C). The formazan crystals were solubilized in DMSO overnight, and absorbency was measured at 595 nm on a spectrophotometer (Labsystems Multiscan; Helsinki, Finland). TNF-{alpha}-soluble receptor II (TNF-{alpha} sRII) was used to as a positive control. Serial dilutions (400–0.04 ng/ml) of TNF-{alpha} sRII indicated that the ED50 dose was 4 ng/ml in the presence of 500 pg of recombinant TNF-{alpha}. TNF-{alpha} sRII abolished the TNF-{alpha} activity in supernatants from hypoxic and LPS-stimulated cells at a concentration of 4 ng/ml, indicating that the assay was specific for TNF-{alpha} activity.

NF-{kappa}B and Activated (Phospho)-p38 MAPK

MVEC and LEC were treated as described in TNF-{alpha} Secretion; however, cells were stimulated with TNF-{alpha} (10 ng/ml; 4 h) rather than LPS. Nuclear extracts were prepared as previously described (38). Briefly, cells were washed with ice-cold PBS, scraped, and resuspended in hypotonic buffer (10 mM HEPES, pH 7.9, 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM DTT, and 0.5 mM PMSF for 15 min on ice). Nonidet P-40 (10%, 25 µl) was added to the solution, and cells were vortexed and centrifuged (12,000 g for 30 s). Supernatant was removed, and the nuclear pellet was extracted with hypertonic buffer B (20 mM HEPES, pH 7.9, 0.4 M NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM DTT, and 1 mM PMSF) by shaking at 4°C for 15 min and then centrifuged at 12,000 g. Supernatant was frozen (–70°C) before NF-{kappa}B analyses were performed.

Cytosolic protein was prepared as previously described (26). Briefly, MVEC and LEC were washed with ice-cold PBS, scraped, and resuspended in cell lysis buffer (1 ml: 50 mM {beta}-glycerophosphate, 10 mM HEPES, pH 7.4, 70 mM NaCl, 2 mM EDTA, and 1% SDS; no. 80-0943; Assay Designs, Ann Arbor, MI). Cells were incubated on ice (20 min), vortexed, and frozen (–70°C) before activated p38 MAPK analyses were performed.

Protein Assay

Protein concentrations were determined using the method of Bradford (Coomassie assay kit no. 23200; Pierce, Rockford, IL). Briefly, standard or unknown samples (5 µl) were added to Coomassie reagent (250 µl) in microplate wells and incubated for 10 min at room temperature (RT). Absorbency was read on a spectrophotometer at 595 nm.

NF-{kappa}B Quantification

NF-{kappa}B activity was determined in nuclear extracts (50 µl, 200 µg/ml protein) by using a commercially available enzyme-linked immunoassay (ELISA) kit (no. TF 01; Oxford Biomedical Research, Oxford, MI). This chemiluminescent ELISA employs an oligonucleotide containing the DNA-binding NF-{kappa}B consensus sequence bound to a 96-well ELISA plate. The DNA-bound NF-{kappa}B is selectively recognized by the primary antibody (p50 and p105 specific), which in turn is detected by the secondary antibody alkaline phosphatase conjugate. Luminescence was measured at 540 nm in a chemiluminometer plate reader (Mithras LB940; Berthold Technologies).

Activated (Phospho)-p38 MAPK Quantification

The amount of activated p38 MAPK in cell lysates (100 µl, 100 µg/ml protein) was determined by using a commercially available phospho-p38 MAPK enzyme immunometric assay kit (no. 900-101; Assay Designs). The color generated by the enzyme reaction was read at 450 nm on a spectrophotometer, as previously described.

F-Actin Staining

MVEC and LEC were prepared on either 6-cm culture dishes or glass chamber slides for F-actin quantification and visualization, respectively, and were treated as described above. Briefly, MVEC and LEC cells were fixed (4% formaldehyde, 10 min, RT), permeabilized (5 min, 0.1% Triton X-100), washed (PBS), and incubated (1% BSA, 30 min) before being stained with FITC-phalloidin (6.6 µM, 20 min, 4°C, in the dark). Stained F-actin was visualized using a Nikon Eclipse E800 microscope equipped with epifluorescence illumination at x100–400 magnification.

F-Actin Quantification

F-actin staining was performed, and the bound dye was extracted with methanol (1 ml, 1 h, 4°C, in the dark). The extracted dye was measured in a chemiluminescence plate reader with excitation at 490 nm and emission at 520 nm. Data are expressed as mean relative light units (RLU) and compared between groups.

Morphological Investigation

Immediately after the permeability tests were completed, Transwells with endothelial cells were washed, fixed (3% glutaraldehyde, 1 h, RT) and stained (crystal violet, 1 h, RT) for microscopy analyses. The stained inserts were removed from the Transwell, placed on slides, and visualized with a Nikon Eclipse microscope at x100–400 magnification.

Statistical Analysis

All experiments followed a randomized block design with the use of cells from at least three different cell preparations. All assays were completed in duplicate or triplicate. Data are expressed as means ± SE, and significance between groups was determined by one-way analysis of variance (ANOVA) using the statistical software package JMP (v. 5; SAS Institute, Cary, NC) on a personal computer. Statistical significance was set at P ≤ 0.05.


    RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Microvascular vs. Macrovascular Endothelial Cell Permeability

Microvascular and macrovascular endothelial cells respond differently to various stimuli (1, 18, 33, 39), so we sought to determine the permeability of bovine pulmonary MVEC and bovine pulmonary artery LEC cultured monolayers exposed to standard cell culture conditions (21%O2, 5%CO2, N2 balance; control) in response to hypoxia (3% O2) or TNF-{alpha} (25 ng/ml) for 24 h. Hypoxia and TNF-{alpha} treatment increased endothelial permeability in MVEC and LEC (Fig. 1); however, MVEC had the greatest permeability in all conditions tested (P = 0.045, MVEC vs. LEC). When permeability rates were normalized to the control conditions within a similar cell type, MVEC and LEC had similar changes in permeability (Tables 1 and 2).



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Fig. 1. Atrial natriuretic peptide (ANP) effect on permeability. Confluent microvascular endothelial (MVE) and macrovascular endothelial (LE) cells in Transwell chambers were unstimulated (ctrl) or stimulated with hypoxia (Hyp; 3% O2, 5% CO2, and 92% N2) or TNF-{alpha} (25 ng/ml) for 24 h in the presence or absence of ANP (10–8, 10–7, 10–6 M). Data are means ± SE of 4 separate experiments done in triplicate (n = 12). *P ≤ 0.002, MVE vs. MVE + ANP during hypoxia or TNF-{alpha} stimulation. **P ≤ 0.02, LE vs. LE + ANP during TNF-{alpha} stimulation. ***P = 0.045, MVE vs. LEC. {ddagger}P ≤ 0.02, ANP effect on MVE vs. LE.

 

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Table 1. Summary of ANP effects

 

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Table 2. Summary of similarities and differences of each cell type in response to hypoxia or TNF-{alpha} stimulation

 
ANP Inhibits Hypoxia- and TNF-{alpha}-Induced Endothelial Monolayer Permeability

To test the hypothesis that ANP can prevent an increase in pulmonary vascular permeability, we pretreated MVEC and LEC with ANP (10–8, 10–7, and 10–6 M) and then exposed them to hypoxia or treated them with TNF-{alpha}. ANP elicited a reduction in permeability in MVEC during hypoxia and TNF-{alpha} treatment and in LEC during TNF-{alpha} treatment (Fig. 1 and Table 1).

Activation of p38 MAPK is known to alter F-actin (stress fiber formation) and increase endothelial permeability (25). Therefore, to determine a possible mechanism by which ANP action inhibits monolayer permeability, we pretreated MVEC and LEC with SB-202190 (10–6 M), a p38 MAPK inhibitor. p38 MAPK inhibition decreased permeability in MVEC and LEC during hypoxia and TNF-{alpha} treatment (Fig. 2).



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Fig. 2. Effect of p38 inhibition on endothelial cell permeability. Confluent MVE and LE cells in Transwell chambers were unstimulated or stimulated with hypoxia (3% O2, 5% CO2, and 92% N2) or TNF-{alpha} (25 ng/ml) for 24 h in the presence or absence of SB-202190 (10–6 M), a p38 mitogen-activated protein kinase inhibitor. Data are means ± SE of 3 separate experiments done in triplicate (n = 9). *P ≤ 0.03, MVE vs. MVE + SB-202190 during hypoxia or TNF-{alpha} stimulation. **P ≤ 0.04, LE vs. LE + SB-202190 during hypoxia or TNF-{alpha} stimulation.

 
ANP Inhibits Endothelial Secretion of TNF-{alpha}

Because lung permeability may be enhanced by TNF-{alpha} at high altitude (11, 28, 29), and because ANP inhibits synthesis and secretion of TNF-{alpha} (24), we determined TNF-{alpha} secretion from MVEC and LEC in response to hypoxia. LPS treatment was used as a positive control. Hypoxia induced TNF-{alpha} secretion from MVEC and LEC (P ≤ 0.05) but was twofold greater in MVEC vs. LEC (P ≤ 0.006) (Table 1). LPS increased TNF-{alpha} secretion by the MVEC and LEC three- and fivefold more than did hypoxia and was similar in MVEC and LEC (Fig. 3 and Table 1). ANP reduced hypoxia- and LPS-stimulated TNF-{alpha} secretion in MVEC and LEC (Fig. 3).



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Fig. 3. ANP inhibition of TNF-{alpha} secretion in MVE and LE cells. MVE and LE cells were grown to confluence in a 6-cm culture dish and left unstimulated or stimulated with hypoxia (3% O2, 5% CO2, and 92% N2; 24 h) or LPS (1 µg/ml; 6 h) in the presence or absence of ANP (10–7 M). Culture supernatants were assayed for TNF-{alpha} secretion by using L929 cells in a bioassay. Data are means ± SE of 3 separate experiments done in duplicate (n = 6) *P ≤ 0.001, MVE vs. MVE control. **P ≤ 0.001, LE vs. LE control. ***P ≤ 0.006, MVE vs. LE.

 
ANP Inhibits Activated NF-{kappa}B

NF-{kappa}B promotes TNF-{alpha} transcription during hypoxia and inflammation (32). Therefore, to establish a mechanism for ANP inhibition of endothelial TNF-{alpha} secretion, we determined the effect of ANP on the activity of transcription factor NF-{kappa}B. Hypoxia and TNF-{alpha} increased MVEC and LEC NF-{kappa}B activity (Table 1). Hypoxia activated NF-{kappa}B twofold higher in MVEC vs. LEC (P ≤ 0.0005) (Fig. 4). ANP decreased hypoxia- and TNF-{alpha}-stimulated NF-{kappa}B activity in MVEC and LEC (Fig. 4).



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Fig. 4. ANP inhibition of NF-{kappa}B activity. MVE and LE cells were grown to confluence in a 10-cm culture dishes and left unstimulated or stimulated with hypoxia (3% O2, 5% CO2, and 92% N2; 24 h) or TNF-{alpha} (10 ng/ml; 4 h) in the presence or absence of ANP (10–7 M). Nuclear protein was assayed for activated NF-{kappa}B by using a chemiluminescence-based sandwich-type ELISA. Data are means ± SE of 3 separate experiments assayed in duplicate (n = 3) *P ≤ 0.0015, MVE vs. MVE control. **P ≤ 0.03, LE vs. LE control. ***P ≤ 0.0015, MVE vs. LE during hypoxia.

 
Differential Activation of p38 MAPK in MVEC and LEC

Others have demonstrated that TNF-{alpha} can increase activation of p38 MAPK, which is known to alter cell morphology and increase endothelial permeability (30, 32). Therefore, we sought to determine whether hypoxia or TNF-{alpha} increases activation of p38 MAPK in MVEC and LEC and whether ANP inhibits that activation. Hypoxia decreased activated p38 MAPK (P ≤ 0.05) in MVEC but caused an increase in LEC (P = 0.002). Conversely, TNF-{alpha} stimulation increased activation of p38 MAPK in MVEC but not in LEC (Table 1). Activation of p38 MAPK was greater in MVEC vs. LEC in standard cell culture conditions and with TNF-{alpha} treatment, but during hypoxia, activation of p38 MAPK was less in MVEC vs. LEC (Fig. 5). In standard culture conditions, ANP decreased activation of p38 MAPK in MVEC but had no effect on LEC (Fig. 5). ANP had no effect on activation of p38 in hypoxia-stimulated MVEC but inhibited activation of p38 MAPK in LEC. ANP treatment reduced activation of p38 MAPK in TNF-{alpha}-stimulated MVEC but had no effect on LEC (Table 1).



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Fig. 5. ANP inhibition of activated p38 MAPK. MVE and LE cells were grown to confluence in a 10-cm culture dishes and left unstimulated or stimulated with hypoxia (3% O2, 5% CO2, and 92% N2; 24 h) or TNF-{alpha} (10 ng/ml; 4 h) in the presence or absence of ANP (10–7 M). Cytosolic protein was assayed for phospho-p38 MAPK by using an enzyme immunometric assay kit. Data are means ± SE of 3 separate experiments assayed in duplicate (n = 3) *P ≤ 0.05, MVE vs. MVE control. **P ≤ 0.002, LE vs. LE control. ***P ≤ 0.001, MVE vs. LE.

 
Differential Actin Polymerization in MVEC and LEC

Activation of p38 MAPK increases endothelial permeability by stimulating a signal cascade that causes polymerization of G-actin into F-actin (2, 25, 30, 35), also known as stress fibers. Therefore, we sought to determine whether hypoxia or TNF-{alpha} increases stress fiber formation in MVEC and LEC and whether ANP inhibits that activation. Hypoxia did not affect stress fiber formation in MVEC but did increase stress fibers in LEC (Fig. 6). TNF-{alpha} stimulation increased stress fiber formation approximately twofold in MVEC, but no change was evident in LEC. ANP reduced stress fiber formation in hypoxic LEC and TNF-{alpha} -stimulated MVEC (Fig. 7). To further assess stress fiber formation by p38 MAPK activation, we pretreated MVEC and LEC with SB-202190, a p38 MAPK inhibitor. Treatment with SB-202190 decreased F-actin formation in hypoxia- and TNF-{alpha}-treated MVEC and LEC (Fig. 8).



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Fig. 6. F-actin formation. MVE and LE cells were grown to confluence in 6-cm culture dishes and left unstimulated or stimulated with hypoxia (3% O2, 5% CO2, and 92 %N2; 24 h) or TNF-{alpha} (10 ng/ml; 24 h). F-actin was quantified by staining with FITC-phalloidin followed by fluorescence photometry. Values are expressed as relative light units (RLU/100). Data are means ± SE of 2 separate experiments done in duplicate (n = 6). *P ≤ 0.05, MVE vs. MVE control. **P ≤ 0.05, LE vs. LE control. ***P ≤ 0.05, MVE vs. LE.

 


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Fig. 7. ANP inhibition of F-actin formation. MVE and LE cells were grown to confluence in 24-well culture dishes and left unstimulated or stimulated with hypoxia (3% O2, 5% CO2, and 92% N2; 24 h) or TNF-{alpha} (10 ng/ml; 24 h) in the presence or absence of ANP (10–8, 10–7, 10–6 M). F-actin was quantified by staining with FITC-phalloidin followed by fluorescence photometry. Values are expressed as RLU/100. Data are means ± SE of 2 separate experiments done in triplicate (n = 6). *P ≤ 0.05, MVE vs. MVE control. **P ≤ 0.05, LE vs. LE control. ***P ≤ 0.05, MVE vs. LE.

 


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Fig. 8. Inhibition of activated p38 MAPK on F-actin formation. MVE and LE cells were grown to confluence in 24-well culture dishes and left unstimulated or stimulated with hypoxia (3% O2, 5% CO2, and 92% N2; 24 h) or TNF-{alpha} (10 ng/ml; 24 h) in the presence or absence of SB-202190 (10–6 M). F-actin was quantified by staining with FITC-phalloidin followed by fluorescence photometry. Values are expressed as RLU/100. Data are means ± SE of 2 separate experiments done in quintuplicate (n = 10). *P ≤ 0.05, MVE vs. MVE during hypoxia or TNF-{alpha} stimulation. **P ≤ 0.05, LE vs. LE during hypoxia or TNF-{alpha} stimulation.

 
Morphological Differences Between MVEC and LEC

MVEC and LEC presented typical cobblestone morphology. Hypoxia-stimulated MVEC and LEV started to retract from one another and form intercellular gaps (Figs. 9 and 10). TNF-{alpha}-stimulated MVEC and LEC showed more pronounced retraction and formation of intercellular gaps with a noticeable change in architecture from their typical cobblestone appearance to more elongated cells (Figs. 9 and 10). ANP abrogated hypoxia- and TNF-{alpha}-stimulated changes in MVEC and LEC, respectively (Figs. 9 and 10).



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Fig. 9. ANP inhibits hypoxic, and TNF-{alpha}-stimulated morphological changes in MVE (A) and LE cells (B), respectively. Confluent MVE and LE cells in Transwell chambers were cultured under control, hypoxic (3% O2, 5% CO2, and 92% N2; 24 h), or TNF-{alpha}-stimulated (25 ng/ml; 24 h) conditions in the presence or absence of ANP (10–8, 10–7, 10–6 M). Cells were stained with crystal violate immediately after the experiment. Images are representative photographs from 4 independent experiments, with ANP effect shown at 10–7 M. Original magnification, x10.

 


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Fig. 10. ANP reduces F-actin formation in hypoxic and TNF-{alpha}-stimulated MVE (A) and LE cells (B), respectively. Confluent MVE and LE cells in 8-well chamber slides were cultured under control, hypoxic (3% O2, 5% CO2 and 92% N2; 24 h), or TNF-{alpha}-stimulated (25 ng/ml; 24 h) conditions in the presence or absence of ANP (10–7 M). Cells were stained with FITC-phalloidin. Images are representative photographs from 3 independent experiments, with ANP effect shown at 10–7 M. Original magnification, x20.

 
Compared with quiescent MVEC, there were slightly fewer stress fibers in hypoxia-stimulated MVEC in the central region and peripheral band around the margin of the cell. However, hypoxic LEC had more stress fibers transversing the central region of the cell and a thickening of the dense peripheral band around the margin of the cell. TNF-{alpha}-stimulated MVEC showed increased central region stress fibers in a dense meshwork of thickened filament bands compared with unstimulated cells, but TNF-{alpha}-stimulated LEC showed only a small change in central region and peripheral band actin filaments compared with quiescent cells. ANP reduced hypoxia- and TNF-{alpha}-stimulated changes in stress fiber formation in LEC and MVEC, respectively (Figs. 9 and 10).


    DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Our data provide evidence of a previously undescribed mechanism for a hypoxia-induced increase in pulmonary vascular permeability: enhanced NF-{kappa}B activity, TNF-{alpha} secretion, activation of 38 MAPK, activation of stress fiber formation, and, ultimately, altered morphology leading to increased permeability. These are the first data to provide evidence that during hypoxia, ANP not only has vasodilatory and natriuretic properties but also may have directly reduced pulmonary endothelial leak. Our data also indicate that microvascular and macrovascular pulmonary endothelia respond differently to hypoxia and ANP treatment.

Acute hypoxia increases cardiac output (11, 12, 46), causes pulmonary hypertension (3, 15), and releases a web of vasoactive agents resulting in increased pulmonary vascular permeability (3, 13). Caution must be applied when interpreting data from our in vitro experiments on barrier function properties in endothelial cells, because they fail to take into account shear stress, hydrostatic pressure, and other fluid flow properties that are present in vivo. Cell culture preparations are exposed to only a few of the myriad of interacting vasoactive peptides that tip the balance in favor of pulmonary edema and, therefore, may yield different results than in vivo models. It also has been noted that preparation of MVEC and LEC cultures may alter phenotypes from that occurring in vivo (18). To reduce the risk of inducing phenotypic changes, we used cells from a low passage number (510) and used the same passage within each experiment, and cells were seeded and grown to monolayers within the same time frame. Although MVEC appear distinct from LEC in culture, MVEC cannot be identified as arteriole, capillary, or venous.

Our data suggesting that hypoxia increases permeability, activates NF-{kappa}B, increases secretion of TNF-{alpha}, activates p38 activation, and changes F-actin concentration in MVEC and LEC are consistent with those of other studies investigating endothelium activated by hypoxia or TNF-{alpha} in other organs such as the brain, liver, and umbilical cord (7, 8, 20, 25, 26). However, as far as we know, this is the first study exploring the relationship between the effects of hypoxia and inflammatory mediators in cultured pulmonary endothelial cells. Our data further indicate that when MVEC and LEC become hypoxic, a pathway is activated analogous to the inflammatory response induced by TNF-{alpha} or bacterial endotoxin (LPS). Similar data were previously reported in TNF-{alpha}-stimulated HUVEC (19, 20, 2426) but not in pulmonary endothelium.

MVEC had a higher permeability rate compared with LEC in all conditions. Data from previous studies provided evidence that under standard culture conditions, MVEC form a more restrictive barrier to macromolecules (molecular mass > 77,000 Da) and have a lower permeability rate compared with LEC (18, 39). The differing results may be due to differences between the Transwell matrices on which cells were cultured (34). Also, we used FITC-labeled-albumin, whereas others used dextran as a permeability tracer (18). MVEC have a transcytotic pathway selective for albumin (42), which may not be inherent in LEC. Although it has been reported that MVEC form a tighter barrier compared with LEC (18, 39), our data suggest that this may not be true for all molecules. Further investigations testing the permeability rates of a variety of compounds passing through MVEC and LEC monolayers are needed to clarify this issue.

Our data indicating differential regulation of p38 MAPK activation and F-actin arrangement between hypoxic and TNF-{alpha}-stimulated MVEC and LEC are consistent with previous studies demonstrating that micro- and macrovascular endothelial cells respond differently to various stimuli (1, 4, 18). Our data suggest that during hypoxia, activation of p38 MAPK and F-actin formation was decreased or unchanged in MVEC but increased in LEC. In contrast, these results were reversed with TNF-{alpha} treatment. Decreased or unchanged p38 MAPK and F-actin formation may be a reflection of decreased endothelial cell vitality. However, this was not supported by our microscopy analyses, but an absolute measure of cell vitality such as an apoptosis assay was not performed. During hypoxia, calcium is a required upstream activator of p38 MAPK (5). We did not test intracellular calcium level, and therefore we cannot determine whether it played a role in our model. Our data further indicate that hypoxia-stimulated MVEC did not change F-actin concentration but increased permeability. These data are supported by previously published data indicating that brain microvascular endothelial cells showed increased permeability, whereas F-actin concentration remained unchanged but distribution was altered (7, 8). In contrast, it is uncertain why TNF-{alpha} increased activation of p38 MAPK in MVEC but not in LEC. It is possible that activation of p38 MAPK increased and returned toward baseline values at 24 h in TNF-{alpha}-stimulated LEC compared with MVEC. This would explain why inhibition of ANP and p38 MAPK decreased TNF-stimulated LEC monolayer permeability.

In a series of recently published studies (1926), data indicated that ANP inhibited production of proinflammatory agents from immune cells (macrophages) and prevented deleterious effects on vascular endothelial cells (HUVEC) in vitro. Similarly, our data indicate that ANP inhibits vascular permeability, TNF-{alpha} secretion, NF-{kappa}B activity, and activation of p38 MAPK in hypoxia- and TNF-{alpha}-stimulated MVEC and LEC. Our study design did not allow us to investigate the mechanism by which ANP inhibits NF-{kappa}B and p38 MAPK in MVEC and LEC, but it is likely through the induction of I{kappa}B and MAPK phosphatase-1 (MKP-1), as previously reported (25, 26). Interestingly, data show that p38 MAPK can inhibit NF-{kappa}B (17). The extent to which ANP is acting through inhibition of p38 is not known. However, our study is the first to investigate the cytoprotective effect of ANP on hypoxia-stimulated pulmonary micro- and macrovascular endothelial cells in vitro.

There are some notable differences between ANP action on MVEC vs. LEC. During hypoxia, ANP reduced permeability in MVEC but not in LEC; however, ANP had no effect on the activation of p38 MAPK and subsequent F-actin formation in MVEC but inhibited this pathway in LEC (Table 2). The data suggest that during hypoxia, ANP can preserve MVEC barrier function in MVEC by means other than inhibition of p38 MAPK activation. ANP reduced permeability in TNF-{alpha}- but not hypoxia-stimulated LEC. It has been reported that C-type natriuretic peptide receptor (NPR-C) is downregulated in the lung by hypoxia (40) in vivo. However, our study design did not allow us to determine whether there was a connection between the NPR-C receptor and the inability of ANP to reduce the leak in hypoxic LEC.

Although ANP had differential effects on MVEC and LEC, our results are consistent with animal models in which ANP ameliorated HAPE (15). HAPE is a rare, noncardiogenic pulmonary edema (11) that may be caused by increased hydrostatic pressure, inflammatory agents, or a combination of the two (44). Our data provide unique insights into the mechanisms controlling pulmonary endothelial leak during acute hypoxia, unassociated with changes in vascular pressure, that are impossible to determine in the intact lung. Data indicating that hypoxia alters barrier function in pulmonary endothelial monolayers suggest that there may be a permeability component to HAPE independent of fluid dynamics. Our data further suggest that hypoxia can activate the same pathways as inflammatory agents such as TNF-{alpha} and the bacterial endotoxin LPS (Fig. 11). We propose that acute hypoxia and inflammatory stimuli may share some common pathways leading to enhanced pulmonary endothelial permeability and pulmonary edema.



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Fig. 11. Schematic proposing where ANP exerts its effect on MVE cells (MVEC) and LE cells (LEC) cultured under control, hypoxic, TNF-{alpha}-stimulated, and LPS-stimulated conditions. Hypoxia, TNF-{alpha}, or LPS stimulation increases activated NF-{kappa}B and TNF-{alpha} secretion in MVEC and LEC. Hypoxia increases activated p38 MAPK and stress fiber formation in LEC but not in MVEC. This is reversed under TNF-{alpha}-stimulated conditions. Calcium regulation or temporal differences in p38 MAPK activation may play a role in this differential activation of p38 MAPK.

 
Our data suggest that inflammatory mediators may play a role in the hypoxia-induced increase in pulmonary endothelial permeability and that ANP may have an important role in inhibiting hypoxia- and/or inflammatory agent-mediated permeability independently of its vasodilatory and natriuretic actions.


    GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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This research project was supported, in part, by the American Heart Association, Desert Mountain Affiliate Pre-doctoral Grant 02152627 (to D. C. Irwin).


    ACKNOWLEDGMENTS
 
We thank Sandy Walchak for invaluable assistance and training, allowing successful cell culture and hypoxia studies, Ivan McMurtry for reviewing this manuscript, and the Cardiovascular and Pulmonary research laboratory at the University of Colorado Health Science Center for kindly permitting the use of a cellular hypoxic chamber.


    FOOTNOTES
 

Address for reprint requests and other correspondence: D. C. Irwin, Dept. of Biomedical Sciences, College of Veterinary and Biomedical Sciences, Colorado State Univ., Fort Collins, CO 80523 (E-mail: Davidcirwin{at}earthlink.net)

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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