Lysophosphatidylcholine increases endothelial permeability: role of PKC{alpha} and RhoA cross talk

Fei Huang,1 Papasani V. Subbaiah,2 Oksana Holian,3 Jihang Zhang,1 Arnold Johnson,4 Nancy Gertzberg,4 and Hazel Lum1

1Department of Pharmacology, Rush University Medical Center, Chicago; 2Department of Medicine, University of Illinois, Chicago; 3Department of Medicine, John H. Stroger Hospital of Cook County, Chicago, Illinois; and 4Research Service, Stratton Veterans Affairs Medical Center, Albany, New York

Submitted 4 January 2005 ; accepted in final form 4 March 2005


    ABSTRACT
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Lysophosphatidylcholine (LPC) is a bioactive proinflammatory lipid that can be generated by pathological activities. We investigated the hypothesis that LPC signals increase in endothelial permeability. Stimulation of human dermal microvascular endothelial cells and bovine pulmonary microvascular endothelial cells with LPC (10–50 µM) induced decreases (within minutes) in transendothelial electrical resistance and increase of endothelial permeability. LPC activated (within 5 min) membrane-associated PKC phosphotransferase activity in the absence of translocation. Affinity-binding analysis indicated that LPC induced increases (also by 5 min) of GTP-bound RhoA, but not Rac1 or Cdc42. By 60 min, both signaling pathways decreased toward baseline. Inhibition of RhoA with C3 transferase inhibited ~50% of LPC-induced resistance decrease. Pretreatment with PKC inhibitor Gö-6983 (concentrations selective for classic PKC), PMA-induced depletion of PKC{alpha}, and transfection of antisense PKC{alpha} oligonucleotide each prevented 40–50% of the LPC-induced resistance decrease. Furthermore, these three PKC inhibition strategies inhibited 60–80% of the LPC-induced GTP-bound RhoA. These results show that LPC directly impairs the endothelial barrier function that was dependent, at least in part, on cross talk of PKC{alpha} and RhoA signals. The evidence indicates that elevated LPC levels can contribute to the activation of a proinflammatory endothelial phenotype.

protein kinase C; signal transduction


LYSOPHOSPHATIDYLCHOLINE (LPC) belongs to a group of bioactive glycerol- or sphingosine-based lysophospholipids [i.e., lysophosphatidic acid, sphingosine-1-phosphate, and sphingosylphosphorylcholine (SPC)] generated from membrane phospholipids as part of normal physiological activities or disease processes. It has been found to accumulate in pathological tissues such as in the ischemic myocardium, atherosclerotic aortas, and other inflammatory lesions of blood vessels (5, 33). LPC is also a major phospholipid component (40–50%) of oxidized LDL (17) and is implicated as a critical atherogenic factor of oxidized LDL. Several diseases such as endometriosis (22), asthma (18), and ovarian cancer (26) are associated with two- to threefold increased circulating levels of LPC. It is believed that a primary source of pathological levels of LPC is through the action of phospholipase A2 (PLA2) on membrane phosphatidylcholine, generating LPC concomitantly with arachidonic acid (2, 16, 32, 35). Despite these documented elevations of LPC in association with pathological conditions, the pathophysiological role of LPC in diseases remains to be established.

There is clear evidence that the bioactive activities of LPC include activation of vascular endothelium. For example, extracellular LPC (10–100 µM) is reported to upregulate expression of adhesion molecules (8, 21, 40), production of cytokines (23), secretion of O2 (7), and DNA-binding activity of NF-{kappa}B (34). In vivo studies show that direct LPC injection either subcutaneously (31) or into the spinal cord (27) causes inflammatory cell infiltration accompanied by vascular leakage. Overall, evidence indicates that LPC induces a significant proinflammatory endothelial phenotype. Yet, to date, the effects of LPC on vascular endothelial permeability have not been systematically investigated, and the mechanisms are unknown. Therefore, the goals of this study are to determine whether LPC induces endothelial barrier dysfunction and to identify the signaling mechanisms responsible for this regulation.

The critical findings are that LPC 1) impaired endothelial barrier function that was sensitive to albumin concentration, 2) increased PKC phosphotransferase activity in the absence of translocation of either PKC{alpha} or PKC{beta}, 3) activated RhoA, but not Rac1 or Cdc42, and 4) induced barrier dysfunction that was dependent in part on PKC{alpha} and RhoA cross talk. These findings provide the first report that proinflammatory LPC directly increases endothelial permeability through activation of signaling mechanisms.


    METHODS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Preparation of LPC

LPC (1-palmitoyl-2-hydroxy-sn-glycero-3-phosphocholine) was purchased from Avanti Polar Lipids (Alabaster, AL), checked for fatty acid composition by gas-liquid chromatography, and found to be at least 96% pure. It was dissolved in chloroform:methanol (2:1) and stored at –20°C. Aliquots were evaporated under nitrogen in glass tubes and resuspended in sufficient volume of Hanks’ balanced salt solution to give a final concentration of 1 mM. The samples were vortexed at room temperature for 1 min (2x) to yield a clear dispersion, and the final concentration was confirmed by analysis of lipid phosphorus by the modified Bartlett procedure (15). The phospholipid dispersions were stored at 4°C and were used within 30 days of the preparation.

Cell Culture

Human dermal microvascular endothelial cells (HMEC) (1) were maintained in culture in MCDB-131 medium (GIBCO BRL, Gaithersburg, MD) and supplemented with 5% fetal bovine serum (FBS; Hyclone, Logan, UT), 10 µg/l human epidermal growth factor (Sigma Chemical, St. Louis, MO), 1 mg/l hydrocortisone, 1% penicillin-streptomycin, and 1% L-glutamine. HMEC is a well-characterized cell line that exhibits the expected morphological and functional endothelial phenotypes, expresses and secretes von Willebrand factor, takes up acetylated LDL, forms tubes when grown in Matrigel, and expresses CD31, CD36, ICAM-1, and CD44 (1). HMEC were passaged 3–4 days when confluent and used for studies at passages 25–35. These cells form a highly restrictive barrier when evaluated by albumin permeability and resistance (11, 12).

Bovine pulmonary microvessel endothelial cells (BPMEC) derived from fresh calf lungs were obtained at passage 4 (VecTechnologies, Rensselaer, NY) and serially cultured from 4 to 12 passages for studies (4, 11). The culture medium contained DMEM (GIBCO BRL; Grand Island, NY) supplemented with 20% FBS, 15 µg/ml endothelial cell growth supplement (Upstate Biotechnology, Lake Placid, NY), and 1% nonessential amino acids (GIBCO BRL). The BPMEC were identified as endothelial monolayers by 1) the characteristic "cobblestone" appearance using contrast microscopy, 2) the presence of factor VIII-related antigen, 3) the uptake of acylated LDL, and 4) the absence of smooth muscle actin.

Transendothelial Electrical Resistance

The transendothelial electrical resistance was determined in real time using the electric cell-substrate impedance sensor system (Applied BioPhysics, Troy, NY) that measures electrical current flow through cells grown onto gold-plated electrodes (5 x 10–4 cm2) with a 500-µl well fitted above each electrode. For study, endothelial cells (2.5 x 105 cells/cm2) were grown to confluence on sterile, fibronectin-coated electrodes, and resistance was measured as previously described by us (11) and others (20). Initial baseline resistance typically showed resistance >7,000 {Omega} (11), and monolayers with lower resistances were rejected from study. We used the minimum baseline electrical resistance of 7,000 {Omega} (corresponds to the calculated value of 3.5 {Omega} cm2) to screen for relatively restrictive monolayers since this resistance is within the reported range of values of 3–6 {Omega} cm2 for endothelial cell monolayers. The endothelial cells were challenged with reagents according to experimental protocol, and resistance was recorded continuously for up to 4 h. Reported values were normalized to initial baseline resistance of each monolayer.

Endothelial Permeability Assay

The permeability assay system measures the transendothelial clearance rate of albumin in the absence of hydrostatic and oncotic pressure gradients and has been described in previous studies (4, 12). In brief, the system consisted of two compartments separated by a sterile gelatinized microporous polycarbonate filter (13-mm diameter, 0.8-µm pore size; Corning Costar; Cambridge, MA) as previously described (12). BPMEC (2 x 105) were seeded onto the filters and cultured to confluence (3–5 days). The luminal (upper) compartment (0.7 ml) was suspended in the abluminal (lower) compartment (25 ml), both of which contained Hanks’ balanced salt solution, 0.5% BSA, and 20 mM HEPES buffer. The lower chamber was stirred continuously for complete mixing, and the entire system was kept in a water bath at 37°C. Evans blue-labeled albumin (20 mg/ml) was added to the upper chamber, 300-µl samples were taken from the lower compartment at 5-min intervals for 60 min, and absorbance was read at 620 nm. At the beginning of each study, an upper chamber sample was diluted 1:100 to determine the initial absorbance of the Evans blue-albumin. The absorbance of free Evans blue in the luminal and abluminal compartments was always <1%, indicating that most of albumin was bound by the dye. The clearance rate of Evans blue-albumin was determined by least-squares linear regression between 10 and 60 min for the control and experimental groups.

PKC Phosphotransferase Activity

In vitro PKC activity was determined from cell fractions as previously reported (13). In brief, HMEC were grown in 60-mm culture dishes and treated according to experimental protocol. All subsequent steps were carried out on ice using ice-cold reagents. The cells were washed with PBS, collected in extraction buffer (0.02 M Tris, 0.5 mM EDTA, 0.5 mM EGTA) containing protease inhibitors (2.5 mM PMSF, 25 µg/ml pepstatin A, 25 µg/ml leupeptin, 25 µg/ml aprotinin) and 10 mM 2-mercaptoethanol, and cytosolic and membrane fractions were prepared. The phosphotransferase activity of cell fractions was determined by incorporation of 32P into the PKC consensus peptide substrate in a reaction mixture of 33 µM unlabeled ATP plus [{gamma}32P]ATP, PKC peptide substrate [Ser25]PKC 19-31 (0.1 mg/ml), 0.01% phosphatidylserine, 0.01% diacylglycerol, 5.4 mM MgCl2, 20 mM Tris base, and 5 mM CaCl2. The mixture was incubated for 5 min at 30°C before being quenched with 75 mM ice-cold H3PO4, vacuum filtered through ion-exchange cellulose discs, and counted in a scintillation counter. Values were calculated as specific PKC activity (pmol 32P incorporated·min–1·mg protein–1).

Rho GTPase Affinity-Binding Assay

The activation of Rho was determined by affinity binding of the target protein rhotekin (binds RhoA-GTP) or CD-PAK (binds Rac1 and Cdc42-GTP) as previously described (30). HMEC or BPMEC were grown in six-well dishes to confluence, treated according to experimental protocol, and collected in glutathione-S-transferase (GST)-FISH buffer [50 mM Tris (pH 7.4), 10% glycerol, 100 mM NaCl, 1% Nonidet P-40, 2 mM MgCl2, 25 mM NaF, and 1 mM EDTA] plus protease inhibitor cocktail (10 µg/ml of pepstatin A, 10 µg/ml each of aprotinin and leupeptin, and 1 mM PMSF). Cell lysates were pelleted by centrifugation at 10,000 g at 4°C for 5 min, and equal volumes of supernatant were incubated with purified GST-rhotekin or GST-CD-PAK coupled to glutathione Sepharose 4B beads (Amersham Pharmacia Biotech, Piscataway, NJ) at 4°C for 1 h. The GTP form of Rho bound to the rhotekin- or CD-PAK-sepharose beads was eluted by being boiled in 2.5x Laemmli sample buffer. The eluted sample and total cell lysate were electrophoresed on 12.5% SDS-PAGE, and Western blot analysis was made with affinity-purified antibodies directed against RhoA, Cdc42 (Santa Cruz Biotechnology), or Rac1 (BD Transduction Laboratories, San Jose, CA). The blots were quantified by scanning densitometry.

PKC Translocation

HMEC were plated on glass coverslips coated with 1 µg/ml of fibronectin and grown overnight. After being treated according to experimental protocol, the cells were washed with PBS twice, fixed with 3.7% formaldehyde in PBS for 20 min, and permeabilized with 0.5% Triton X-100 in PBS for 5 min at room temperature. After being blocked with 5% donkey serum for 30 min, anti-PKC{alpha} (Transduction Laboratories, Lexington, KY) or anti-PKC{beta}1 antibody (Sigma) was added for incubation overnight at 4°C. The cells were washed with PBS three times, incubated with Cy2-conjugated anti-mouse IgG antibody (Jackson Immunoresearch Laboratories, West Grove, PA) in PBS for 1 h at room temperature, and viewed on an Olympus AH3-RFCA fluorescent microscope (Olympus, Tokyo, Japan). Images were recorded with a Nikon digital camera (DXM 1200; Nikon, Kanagawa, Japan).

PKC{alpha} Antisense Oligonucleotides

Translation of PKC{alpha} RNA was inhibited using a phosphorothioated antisense oligonucleotide complementary to a region two bases upstream from the initiation codon of bovine PKC{alpha} [5'-GTC CCT CGC CGC CTC CTG-3' (38)]. The control oligonucleotide was a scrambled nonsense nucleotide (5'-TGC CTC CGC GCC TCC CGT-3'). Specificity of the sequence to bovine PKC{alpha} and scrambled nonsense was verified by using the Entrez-GenBank data base of the National Institutes of Health. The oligonucleotides were transfected with Lipofectin (GIBCO BRL) using a protocol previously described (4). BPMEC were incubated with either antisense or nonsense oligonucleotide at a final concentration of 0.2 nmol/ml in Lipofectin (6.6 µl/ml) at 37°C for 4 h, a period over which the PKC{alpha} mRNA was reduced 75–85% below baseline (4). The cells were then used for experimental protocol as needed.

Cell Viability Assay

The assay is based on the reduction of the tetrazolium salt 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT; Sigma) by mitochondrial dehydrogenases in living cells. HMEC were plated at 80,000 cells/well in a 96-well tissue culture dish, grown to confluence, and treated with LPC (0–50 µM) for 30 min. The cells were washed and incubated with 0.5 mg/ml of MTT for 3 h at 37°C, and 0.04 M HCl in {beta}-isopropanol was added for reading of the optical density (OD) at 570 nm in a plate reader (Molecular Devices, Sunnyvale, CA). The OD units provided an index of enzymatic activity of living cells.

Statistics

Single-sample data were analyzed by the two-tailed t-test; a multiple range test (Scheffé’s test) was used for comparisons of experimental groups with a single control group. Two-way ANOVA was used for analysis to determine significant differences between groups.


    RESULTS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
LPC Decreases Transendothelial Resistance

We determined the direct effects of LPC on the in vitro barrier function of endothelial cells. Stimulation of HMEC with LPC (10–50 µM) caused rapid decreases (within minutes) of resistance from baseline (Fig. 1A). The inset shows an expanded x-axis of the 25 µM LPC response, indicating a resistance decrease to near maximal by ~15 min (Fig. 1A, inset). The resistance decrease in response to the lower concentrations of 10–25 µM recovered to near baseline within 30–60 min. However, at the higher LPC concentration of 50 µM, the resistance drop was sustained and did not return to baseline within the 3-h period of study. The extent of the resistance drop increased with increasing LPC concentrations, reaching a maximum decrease from baseline at 50 µM LPC (Fig. 1, A and B). Evaluation of the effects of LPC on cell viability indicated a 5–8% decrease of the mitochondrial dehydrogenase activity at 20–50 µM LPC and was not statistically significant from control (results not shown).



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Fig. 1. Dose response of lysophosphatidylcholine (LPC) on transendothelial electrical resistance. Human dermal microvascular endothelial cells (HMEC) were grown onto electric cell-substrate impedance sensor (ECIS) electrodes until confluent and treated with LPC (10–50 µM; arrow), and resistance was determined for up to 3 h. A: representative graph of resistance normalized to baseline (Norm. Resist.) in response to LPC challenge (arrow); n = 3–5; C, control buffer. Inset: expanded x-axis of the time course of the decrease and recovery in response to 25 µM LPC. B: summary of the maximal resistance decrease from baseline in response to LPC; n = 3–5.

 
Because LPC is known to normally bind serum proteins, particularly to albumin, we also determined the effects of altering the relative proportion of albumin concentration with that of LPC concentration on the resistance decrease. Results indicated that decreased albumin levels exacerbated the LPC-induced resistance decrease (Fig. 2). In the presence of 8 mg/ml of albumin, 25 µM LPC did not decrease resistance (Fig. 2A). Under these experimental conditions, the ratio of [LPC] to [albumin] was ~3, approximating the ratio in circulation. Reducing the albumin concentration 10-fold to 0.8 mg/ml (thereby, the ratio is increased to 31), 25 µM LPC significantly decreased resistance from baseline (Fig. 2A). However, in the absence of albumin, only 2 µM LPC was needed to reduce resistance (Fig. 2B) to a similar extent as 25 µM in the presence of 0.8 mg/ml albumin. These results suggest that the bioactivity of LPC is inversely related to binding by albumin or other serum proteins.



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Fig. 2. Albumin prevents the biological activity of LPC. A: HMEC grown to confluence on ECIS electrodes were stimulated with 25 µM LPC (arrow) in the presence of 0.8 mg/ml or 8.0 mg/ml of albumin. B: HMEC were stimulated with 2 µM LPC (arrow) in the absence of albumin. Each is a representative graph showing resistance changes in real time; n = 3.

 
To determine whether LPC impairs barrier function of endothelial cells from other vascular beds, we evaluated the effects of LPC on the transendothelial electrical resistance of BPMEC. LPC stimulation resulted in a similar pattern of resistance decrease as HMEC. That is, LPC induced a rapid dose-dependent decrease of the resistance that was reversible at only the lower concentrations (Fig. 3).



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Fig. 3. LPC decreases resistance in bovine pulmonary microvascular endothelial cells (BPMEC). BPMEC were grown onto ECIS electrodes until confluent and treated with 10–50 µM LPC, and resistance was determined for up to 2 h. Shown is a representative graph of resistance normalized to baseline in response to LPC challenge (arrow); n = 4.

 
Effects of LPC on PKC Activation

HMEC stimulated with LPC (25 µM) significantly increased PKC phosphotransferase activity of the membrane fraction within 5 min, which slightly decayed with time but remained significantly elevated for up to 60 min (Fig. 4A). Surprisingly, LPC stimulation did not produce a corresponding decreased PKC activity of the cytosolic fraction (Fig. 4B), which suggests that the increased PKC activity of the membrane fraction occurred in the absence of translocation of the enzyme. We further investigated this possibility by evaluating the immunofluorescent localization of the PKC{alpha} and PKC{beta} isoforms in HMEC. We focused on these classic PKC isoforms since we (13, 24) and others (4, 19) have shown that PKC{alpha} and PKC{beta} provide critical regulation of endothelial barrier dysfunction induced by a wide range of mediators. The immunofluorescent localization results showed that HMEC stimulated with LPC (25 µM for 15 min) did not translocate PKC{alpha} (Fig. 5A) or PKC{beta} (data not shown) to the membrane, consistent with the PKC activity data. The positive control group in which HMEC were treated with PMA (100 nM, 15 min) showed decreased PKC{alpha} cytoplasmic localization and increased localization at the cell membrane (Fig. 5A). Further analysis by confocal generation of Z-sections from apical toward basal endothelial regions indicated absence of PKC{alpha} redistribution in response to LPC stimulation (Fig. 5B).



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Fig. 4. LPC increases PKC phosphotransferase activity. HMEC were treated with LPC (25 µM) for 1, 5, 15, and 60 min, and incorporation of 32P into the PKC consensus peptide substrate by the cell membrane (A) and cytosolic fractions (B) were determined in duplicates. Specific PKC activity, pmol·min–1·mg–1 protein, is reported as mean experimented/control % (Exp/C %) ± SE; n = 4; *P < 0.05 compared with control (0) group.

 


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Fig. 5. Immunofluorescent intracellular localization of PKC{alpha}. HMEC were untreated (control) and challenged with 25 µM LPC for 15 min or as positive control (100 nM PMA for 15 min). A: confocal laser microscopy showing the cellular distribution of PKC{alpha}; arrows indicate cell periphery; original magnification is x400; n = 2. B: confocal serial Z-sections showing distribution of PKC{alpha} (control or stimulated with 25 µM LPC for 15 min). The cells were scanned from apical to basal regions of the cells (a–d) to capture Z-sections at 1-µm intervals; original magnification is x600; n = 2.

 
LPC Activates RhoA, but not Rac-1 or Cdc42

We determined whether LPC activates RhoA, Rac-1, and Cdc42 in endothelial cells by measuring the active GTP-bound form of Rho using the affinity-binding assay (see METHODS). Results indicated that in HMEC, LPC (25 µM) induced a twofold increase in RhoA-GTP within 5 min that was sustained at 15 min and decreased by 60 min (Fig. 6A). Total RhoA in whole cell lysates (GTP-bound + GDP-bound forms) was similar among experimental groups, indicating that LPC did not increase de novo generation of RhoA (results not shown). However, stimulation with LPC did not increase Rac1-GTP and Cdc42-GTP (Fig. 6B).



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Fig. 6. Effects of LPC on activation of Rho GTPases. Rho activation was determined by affinity binding of Rho target fusion proteins as described in METHODS. A: confluent HMEC were treated with 25 µM LPC at 0, 1, 5, 15, and 60 min. A representative Western blot shows the affinity-binding results of RhoA. Densitometric analysis is shown as Exp/C % as means ± SE of 5–10 independent experiments; *P < 0.01 compared with no LPC treatment. B: representative Western blot showed that LPC (25 µM, 5 min) did not induce increase of Rac1-GTP and Cdc42-GTP.

 
Barrier Dysfunction Is Regulated by PKC and RhoA Cross Talk

Dependency on RhoA signaling. The effects of inhibition of RhoA on resistance were determined by using C3 transferase toxin (Clostridium botulinum; Biomol, Plymouth Meeting, PA), which inhibits RhoA, RhoB, and RhoC by ADP-ribosylation of Rho at Asn41 (9). Results indicated that in HMEC pretreated with C3 toxin (5 µg/ml) for 24 h, the LPC-induced resistance decrease was significantly inhibited (~50%; Fig. 7, A and B). C3 transferase alone did not decrease resistance from baseline. Higher concentrations of C3 transferase (10 µg/ml) did not further inhibit the resistance decrease (results not shown).



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Fig. 7. Rho inhibitor blocks LPC-induced resistance drop. HMEC grown onto ECIS electrodes until confluent were pretreated with 5 µg/ml of C3 transferase for 24 h and challenged with 25 µM LPC (arrow), and resistance was measured as described. A: representative graph of resistance changes. B: summary of resistance decrease from baseline in response to LPC; n = 5; *P < 0.01 compared with LPC alone.

 
Dependency on PKC signaling. We used multiple approaches to test the dependency of PKC signaling in the LPC-induced barrier dysfunction. Pharmacological pretreatment with the broad spectrum PKC inhibitor Gö-6983 was made at selected concentrations to approximate its IC50 values for PKC isoforms (IC50 for PKC{alpha}, PKC{beta}, and PKC{gamma} = 6–7 nM; IC50 for PKC{delta} = 10 nM). Results indicated that pretreatment of HMEC with 8 and 20 nM resulted in 40–50% inhibition of the LPC-induced resistance decrease (Fig. 8A). Higher concentrations of Gö-6983 (up to 70 nM) did not further inhibit the resistance decrease (IC50 for PKC{zeta} = 60 nM; results not shown). Gö-6983 pretreatment at all these concentrations alone did not alter baseline resistance. Pretreatment with 10 µM rottlerin, a selective PKC{delta} inhibitor (IC50 3–6 µM), also did not prevent the LPC-induced resistance drop (results not shown). These results suggest that LPC-induced barrier dysfunction is mediated by classic PKC isoforms.



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Fig. 8. Effects of PKC inhibition on LPC-mediated resistance. HMEC were grown to confluence on ECIS electrodes and pretreated. A: cells were pretreated with the PKC inhibitor Gö-6983 (GÖ) from 8 to 20 nM and challenged with 25 µM LPC, and resistance was determined for up to 3 h. Shown is a representative graph of the LPC-induced resistance change following pretreatment with GÖ. Control (C) indicates absence of GÖ and LPC; open arrow indicates time of inhibitor pretreatment; solid arrow indicates challenge with LPC; n = 5. B: HMEC were treated with 1 µM PMA for up to 16 h to downregulate PKC. Control indicates absence of PMA. Western blot shows PKC{alpha}, but not PKC{beta}1, was downregulated by 16 h (top). The downregulated HMEC were stimulated with 25 µM LPC (arrow), and resistance was measured. A representative graph shows the effects of downregulated PKC{alpha} on resistance decrease in response to LPC challenge; n = 5.

 
To further investigate this possibility, HMEC were treated with PMA (1 µM) for 16 h to deplete PKC. Western blot analysis showed selective loss of PKC{alpha}, but not PKC{beta} (Fig. 8B, top). This downregulation was associated with ~50% inhibition of the LPC-induced resistance decrease (Fig. 8B, bottom), suggesting that PKC{alpha} may be the more important isoform in the regulation.

The specific role of PKC{alpha} was tested by use of antisense oligonucleotides. In these studies, the antisense PKC{alpha} oligonucleotides inhibited the LPC-induced resistance decrease (Fig. 9, A and B) as well as the LPC-induced increase in albumin clearance rate of BPMEC (Fig. 9C). These observations provide strong support that PKC{alpha} is critical in the regulation of LPC-induced endothelial barrier dysfunction.



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Fig. 9. PKC{alpha} antisense (AS) oligonucleotides inhibit endothelial barrier dysfunction. A: BPMEC were grown on ECIS electrodes to confluence, transfected with PKC{alpha} AS oligonucleotide (0.2 nM/ml) for 4 h, and then challenged with 25 µM LPC. The negative control, PKC{alpha}-nonsense (NS) oligonucleotide (0.2 nM/ml), was similarly transfected into separate groups of BPMEC, and resistance was measured as described. Shown is a representative graph of resistance changes. B: summary of resistance decrease from baseline in response to LPC; n = 5; *P < 0.01 compared with LPC alone. C: BPMEC grown to confluence on gelatinized micropore filters were transfected with PKC{alpha}-AS oligonucleotide as described for resistance studies, and albumin clearance rate was determined; n = 4; *P < 0.05 compared with C (nontreated) group; **P < 0.05 compared with LPC alone.

 
RhoA is downstream target of LPC-activated PKC. The similar time courses of the LPC-induced PKC activity (Fig. 4) with RhoA activation (Fig. 6) suggest possible cross talk between these two families of signaling molecules. We investigated whether PKC regulates RhoA in response to LPC activation in endothelial cells. For these studies, PKC activation was inhibited by pretreatment with 20 nM Gö-6983 or downregulation by long-term pretreatment with PMA (1 µM for 16 h), both protocols shown to inhibit LPC-induced barrier dysfunction. Results showed that Gö-6983 pretreatment prevented ~60% of the LPC-induced RhoA activation, and PMA-induced downregulation inhibited ~80% (Fig. 10A). Similarly, pretreatment with antisense PKC{alpha} oligonucleotides prevented 68% of the LPC-induced RhoA activation (Fig. 10B). The control nonsense oligonucleotide did not inhibit the LPC-induced RhoA activation (Fig. 10B).



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Fig. 10. PKC inhibition prevents LPC-induced RhoA activation. A: HMEC were pretreated with or without 1 µM PMA overnight to induce downregulation (Dn) of PKC{alpha} or 20 nM PKC inhibitor GÖ for 15 min. Cells were then stimulated with LPC (25 µM) for 5 min. Cell extracts were assayed for GTP-bound RhoA by affinity binding as described in METHODS. Shown is a representative affinity binding of RhoA-GTP. Densitometric analysis of RhoA-GTP is presented as means ± SE of 4 separate experiments; *P < 0.01 compared with no LPC treatment (C); **P < 0.05 compared with LPC treatment alone. B: BPMEC were transfected with PKC{alpha}-AS oligonucleotide (0.2 nM/ml) for 4 h and stimulated with LPC, and RhoA activation was determined as described above. The PKC{alpha}-NS oligonucleotide served as control. A representative Western blot shows the affinity-binding results of RhoA. Densitometric analysis is expressed as Exp/C % as means ± SE of 4 independent experiments; *P < 0.05 compared with no LPC treatment; **indicate P < 0.05 compared with LPC treatment alone.

 

    DISCUSSION
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The major findings in this study are that extracellular LPC 1) impaired endothelial barrier function that was sensitive to albumin concentration, 2) increased PKC phosphotransferase activity in the absence of translocation of either PKC{alpha} or PKC{beta}, 3) activated RhoA, but not Rac1 or Cdc42, and 4) induced barrier dysfunction that was dependent in part on PKC{alpha} and RhoA cross talk. These findings provide the first report that the proinflammatory LPC directly induces increase in endothelial permeability through activation of signaling mechanisms.

Our findings support in vivo studies in which intraspinal cord injection of LPC into mice LPC induces recruitment of leukocytes and widespread breakdown of the blood-brain barrier (27). Similarly, local subcutaneous injection of LPC in human subjects results in tissue edema and increased extravasation of T lymphocytes, monocytes, and neutrophils (31). Although the LPC-induced vascular endothelial barrier dysfunction observed in these in vivo studies is likely contributed by the associated activated leukocytic infiltration, our results provide evidence that LPC directly and significantly induces increased permeability, which can further exacerbate inflammatory tissue injury.

We showed that the LPC-induced increase in endothelial permeability was regulated by two critical signaling pathways, PKC and RhoA. The inhibition of classic PKC by three different approaches (Gö-6983, PMA-induced downregulation, and antisense PKC{alpha} oligonucleotides) each prevented ~50% of the LPC-mediated resistance drop. The LPC-induced barrier dysfunction occurred in the absence of translocation of PKC{alpha} to the endothelial cell membrane. The absence of translocation was further supported by the observation that the LPC-increased PKC activity of the membrane cell fraction was not accompanied by a corresponding decrease of activity in the cytosolic fraction, suggesting an increased intrinsic catalytic activity of the membrane-associated enzyme. The positive control, PMA, stimulated PKC{alpha} translocation to the membrane in these cells, indicating that the translocation was stimulus selective. Several cell types (i.e., anterior pituitary cells, fibroblasts) have been reported to show PKC enzyme activation that was temporally unrelated to translocation (10). Furthermore, PKC enzyme activity is known to be regulated through phosphorylation of specific residues on the activation loop of the PKC kinase domain (28). Therefore, it is possible that in endothelial cells, a pool of membrane-bound PKC{alpha} exists that can be activated by LPC stimulation (28). We also observed that the antisense PKC{alpha} oligonucleotides completely inhibited the LPC-induced albumin permeability. It is not clear the reasons why the extent of inhibition differed between the two barrier assays, but they may be attributed to the inherent difference of measuring ion flow and solute flux across a cell monolayer.

We find that LPC-induced increase in endothelial permeability was also dependent on RhoA. LPC stimulation of endothelial cells increased the amount of RhoA-GTP, but not Rac1-GTP or Cdc42-GTP, the other primary Rho-GTPases in cells. RhoA activation occurred within 5 min, remained significantly elevated at 15 min, and declined toward baseline by 60 min, a time period consistent with decreased resistance in response to the same concentration of LPC. Inhibition of the LPC-activated RhoA with the C3 exotoxin prevented ~50% of the resistance decrease, and increased C3 concentration (to 10 µg/ml) did not confer additional inhibition, suggesting that other additional mechanisms contribute to the overall regulation of the increased permeability. Interestingly, a similar degree of inhibition of the thrombin-induced permeability increase by the C3 toxin is reported by Van Nieuw Amerongen et al. (35a). Although definitive targets of regulation by RhoA in the LPC-mediated barrier dysfunction are not known, one likely candidate is Rho kinase, which has been proposed for the thrombin response. Rho kinase is believed to inactivate myosin light chain (MLC) phosphatase as part of a signaling network that controls MLC phosphorylation and endothelial cell contraction (3), which is a critical mechanism underlying endothelial barrier dysfunction (20, 37).

The time course of activation of PKC and RhoA signals by LPC was similar, beginning at ~5 min, remaining elevated for 15 min, and decreasing toward baseline by 60 min. Importantly, within 15 min of LPC stimulation, resistance decrease was near maximal, and a subsequent reversal occurred during the next 30–60 min. These two coincident signaling pathways displayed cross talk in which RhoA was a downstream target of PKC activation. Interestingly, thrombin stimulation of human umbilical vein endothelial cells also demonstrates cross talk between PKC and RhoA (19). In this report, Mehta et al. (19) found that the PKC-induced activation of RhoA is through phosphorylation, and, hence, inhibition of guanine nucleotide dissociation inhibitor. It remains to be determined whether a similar mechanism of regulation of RhoA by PKC occurs in the LPC-induced endothelial barrier dysfunction. The signal transduction mechanisms by which LPC activates PKC and RhoA have yet to be defined. There is emerging evidence linking LPC-activated signals to possible orphan G protein-coupled receptors, GPR4, G2A, and OGR1 (39). We recently reported that human brain and dermal microvascular endothelial cells selectively express GPR4, but not G2A (14), and that GPR4 may have a critical role in regulation of angiogenesis in response to SPC (6). Further studies are underway to definitively establish a role of GPR4 in the regulation of endothelial barrier function.

We find that the presence of albumin greatly influenced the bioactivity of LPC. Albumin has been shown to contain one to three high-affinity binding sites as well as multiple low-affinity binding sites for LPC (29). Our results show that increasing albumin concentration approximating the proportion in the circulation (relative to LPC) prevented the LPC-induced resistance decrease. On the other hand, decreasing albumin (or increasing LPC level) would induce a corresponding impairment of the endothelial barrier function. This finding is consistent with a recent report that perfusion of LPC into analbuminemic rats resulted in a greater vasoconstriction response than in control normal rats (36). Furthermore, in the same study, infusion of albumin prevented LPC-induced vasoconstriction in both analbuminemic and control rats (36). These results strongly suggest that effective proinflammatory activities of LPC can occur in tissues either in which there is elevated LPC concentration that exceeds the binding capacity of albumin (and possibly other serum proteins) or under conditions of decreased albumin levels.

In summary, we show that LPC directly impairs the endothelial barrier function that is sensitive to albumin concentration. This LPC-induced barrier dysfunction was dependent, at least in part, on cross talk of PKC{alpha} and RhoA signals.


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This work was supported by National Heart, Lung, and Blood Institute Grants HL-71081 (H. Lum), HL-52597 (P. V. Subbaiah), HL-69585 (P. V. Subbaiah), and HL-59901 (A. Johnson) and the American Heart Association Postdoctoral Fellowship Award, Midwest (F. Huang).


    FOOTNOTES
 

Address for reprint requests and other correspondence: H. Lum, Dept. of Pharmacology, Rush Univ. Medical Center, 1735 W. Harrison St., Cohn Research Bldg., Rm. 416, Chicago, IL 60612 (e-mail: hlum{at}rush.edu)

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


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