Multiple organ failure after sepsis remains an important clinical problem and is the leading cause of mortality in intensive care units (1). Although these patients do not typically die specifically from liver failure, hepatic dysfunction contributes to metabolic dysregulation, and as the home of approximately 70% of the body's fixed tissue macrophages, it serves as a site for amplification of the systemic inflammatory response and may be responsible for the development of lung failure. There is now substantial evidence indicating that dysregulation of the hepatic microcirculation is a major factor in the development of liver dysfunction in sepsis. Work from our laboratory, as well as others, has demonstrated that a significant contributor to the development of microvascular dysfunction in endotoxemia and other stresses is an enhanced constrictor response to the peptide vasoregulator endothelin 1 (ET-1) (2-4). This enhanced constrictor response is mediated primarily by a loss of the normal ability of ET-1 to activate eNOS in liver sinusoidal endothelial cells (LSECs), thus abrogating the compensatory dilation resulting from NO production.
Regulation of eNOS in vascular endothelial cells is complex, involving both calcium-/calmodulin (CaM)-dependent and phosphorylation-dependent mechanisms. One mechanism that has been identified for regulation of eNOS in injury models via competition for CaM binding is the overexpression of caveolin 1 (Cav-1). Caveolin 1 is a membrane scaffolding protein associated with lipid raft domains called caveolae, and its expression levels are increased in injury models ranging from cirrhosis to endotoxemia. This would suggest that overexpression leads to impaired activation of eNOS in these stress models; however, it is not clear whether the relationship is associative or causal. Therefore, to further investigate the role of Cav-1 in the development of vascular dysregulation in the liver during endotoxemia, we studied the regulation of eNOS after LPS treatment in liver sinusoidal endothelial cells from Cav-1 knockout (KO) mice.
MATERIALS AND METHODS
Endothelin 1 was obtained from American Peptide Company (Palo Alto, Calif). Methyl-β-cyclodextrin and LPS (Escherichia coli O26:B6) were acquired from Sigma-Aldrich (St. Louis, Mo). Rabbit polyclonal antibodies for phospho-eNOS-Ser1177, and polyclonal phospho-eNOS-threonine-495 (Thr495) were purchased from Cell Signaling Technology, Inc. (Beverly, Mass). Mouse monoclonal eNOS antibody was acquired from BD Transduction Laboratories (San Diego, Calif). Rabbit Cav-1 antibody was purchased from Sigma-Aldrich. 4,5-Diaminofluorescein diacetate (DAF-2 DA) was acquired from Calbiochem (La Jolla, Calif). All other chemicals and regents used in the described experiments were purchased from Sigma-Aldrich unless otherwise specified.
Male Cav1tm1Mls/J and C57BL/6 mice (Charles River Laboratories, Fayetteville, NC) weighing 20 to 25 g were housed in a temperature-controlled animal facility with alternating 12-h/12-h light/dark cycles and were fed standard laboratory chow with free access to water.
Hepatic Sinusoidal Endothelial Cell Isolation
All studies were performed under a protocol approved by the Institutional Animal Care and Use Committee of the University of North Carolina at Charlotte and adhere to the Guide for the Care and Use of Laboratory Animals (NIH publication 86-23, revised 1985). Liver sinusoidal endothelial cells were isolated using a protocol as previously described (5). Each of the wells on a 12-well plate was precoated with type VI collagen and was then loaded with 8 × 106 freshly isolated LSECs. After overnight incubation, LSECs (100% confluent) were either treated with culture media or with 100 ng/mL LPS dissolved in the culture media for 6 h.
eNOS Activity Assay
Rate of NO production was evaluated by eNOS activity assay as previously described (5). Briefly, 1 μCi/μL [3H]-l-arginine was added to each well after pretreatments with LPS. The LSECs were then treated with vehicle, ET-1, or ionomycin (positive control) for 30 min. The cells were washed twice with 1 mL arginine-free HEPES media on ice. After the reaction was terminated by the addition of ice-cold phosphate-buffered saline (PBS), the LSECs were lysed, and cell lysates were collected. After centrifugation at 10,000 × g for 5 min, samples were run through glass columns filled with a cation exchange chromatography resin (Bio-Rad) that was previously equilibrated with a stop buffer. The flow-through of each sample was collected in a glass vial, and a scintillation cocktail fluid (Fisher Scientific) was then added. Radioactivity of each sample was quantified in a liquid scintillation counter (Beckman-Coulter). Negative control LSECs were treated with an arginine-free HEPES media containing 10 μmol N(G)-nitro-l-arginine methyl ester, 10 mmol EDTA, and 10 mmol EGTA, and from which background radiation was evaluated. To convert radioactivity (disintegration per minute) into the actual rate of NO production in each well, we designed this equation:
where x is the measured radioactivity data in disintegration per minute.
Protein Extraction and Western Blots
After each corresponding treatment (100 ng/mL LPS for 6 h and/or 10 nmol ET-1 for 30 min), LSECs (8 × 106 cells per sample) were rinsed twice with PBS and were lysed with a lysis master mix. Cell lysates were collected and were centrifuged at 13,000 rpm for 10 min to remove insoluble fractions. Protein concentration of each sample was determined by Micro bicinchoninic acid protein assay kit (Pierce Biotechnology), and the volume of protein loading in Western blot was adjusted accordingly. Each protein sample (5 μg) was mixed with Laemmli loading buffer (Bio-Rad) and was then boiled for 7 min at 100°C. Standard Western blotting procedures were then followed as previously described (5). Precision Plus Protein Standards were used as a molecular weight marker, and β-actin was used as a housekeeper in each experiment (data not shown).
Visualization of NO With DAF-2 DA
Liver sinusoidal endothelial cells (1 × 106 cells per well) were cultured on #1 chamber coverglass (Lab-Tek). After the LPS pretreatment, 5 μmol DAF-2 DA, and 10 nmol ET-1 were added. 4,5-Diaminofluorescein diacetate penetrates cell membrane and is hydrolyzed by intracellular esterase to membrane-impermeable DAF-2, which then reacts with NO to form a fluorescent triazolofluorescein. After incubation with DAF-2 DA and ET-1 for 30 min, the cells were rinsed twice with PBS and fixed with 2% paraformaldehyde for 10 min. Cell nuclei were stained with 2.5 μg/mL 4′,6-diamidino-2-phenylindole for 5 min at room temperature. Confocal images were acquired at excitation/emission wavelengths of 495/515 nm with the FluoView confocal laser scanning microscope (Olympus).
Data from five independent experiments are presented as mean ± SEM. Statistical differences between the media control and treatment groups were determined using one-way ANOVA, followed by post-hoc Dunnett test. Statistical significance was set at P < 0.05. All statistical significances were calculated with SigmaStat (version 2.03; SPSS Inc., Chicago, Ill), and bar graphs were plotted with Microsoft Excel (version XP; Redmond, Wash).
LPS Increases Cav-1 Expression in Wild-Type LSECs
To determine if LPS inhibits eNOS activation through the induction of Cav-1 in primary LSECs, we evaluated Cav-1 protein expression after treating the cells with 100 ng/mL LPS for 6 h. As expected, Cav-1 was not expressed in Cav-1 KO mice (Fig. 1A). However, LPS increased Cav-1 expression in wild-type (WT) LSECs (Fig. 1, A and B; +88%; P < 0.05), but β-actin expression remained relatively constant (Fig. 1A).
Basal eNOS Activity Is Higher in Cav-1 KO Than in WT LSECs
To investigate if the deletion of Cav-1 alters eNOS activity, we evaluated the rate of NO production in the WT and Cav-1 KO LSECs (8 × 106 cells per well) using a [3H]-l-arginine conversion assay. In control media, WT LSECs produced 0.15 ± 0.01 fmol of NO/min per well; in contrast, the rate of NO production in Cav-1 KO LSECs was 0.40 ± 0.04 fmol/min per well (Fig. 2), which was +262% higher than in the WT during the same period (P < 0.05).
LPS Inhibits ET-1-Mediated eNOS Activity in WT LSECs
Endothelin 1 induced eNOS activity to 0.20 ± 0.01 fmol/min per well in WT LSECs (Fig. 2; +31.9% as compared with vehicle; P < 0.05). Treatment with LPS alone has no effect on eNOS activity (Fig. 2; 0.15 ± 0.01 fmol/min per well). However, the ET-1-stimulated eNOS activity was completely abolished after LPS pretreatment (Fig. 2; 0.16 ± 0.01 fmol/min per well).
Targeted Mutation of Cav-1 Reverses LPS Inhibition of ET-1-Mediated eNOS Activation
In Cav-1 KO LSECs, ET-1 induced eNOS activity to 0.52 ± 0.02 fmol/min per well (Fig. 2; +31.1% as compared with media control; P < 0.05). Treatment with LPS alone had minimal effect on eNOS activity, which was at 0.44 ± 0.01 fmol/min per well (Fig. 2). In the WT LSECs, the ET-1-induced eNOS activity was inhibited by LPS pretreatment. However, this LPS-suppressed ET-1 activation of eNOS was reversed in Cav-1 KO LSECs. Endothelin 1 effectively stimulated eNOS activity to 0.50 ± 0.01 fmol/min per well after LPS treatment in Cav-1 KO LSECs (Fig. 2; +25.7% as compared with the control; P < 0.05).
LPS Inhibits ET-1-Mediated eNOS-Ser1177 Phosphorylation in WT LSECs
Phosphorylation of eNOS at the serine-1177 residue increases eNOS activity (6). Treatment with ET-1 stimulated eNOS-Ser1177 phosphorylation in WT LSECs (Fig. 3A; +43.7% as compared with media control; P < 0.05). Treatment with LPS had no effect on eNOS-Ser1177 phosphorylation (Fig. 3A; −2.2%); however, it prevented the ET-1-mediated eNOS-Ser1177 phosphorylation in WT LSECs (Fig. 3A; −6.8%).
Targeted Mutation of Cav-1 Reverses LPS Inhibition of ET-1-Mediated eNOS-Ser1177 Phosphorylation
Treatment with ET-1 stimulated eNOS-Ser1177 phosphorylation in Cav-1 KO LSECs (Fig. 3B; +37.0% as compared with media control; P < 0.05). Similar to the treatment in WT LSECs, LPS also had no effect on eNOS-Ser1177 phosphorylation in Cav-1 KO LSECs (Fig. 3B; −7.1%). In contrast, ET-1 induced eNOS-Ser1177 phosphorylation after LPS pretreatment in Cav-1 KO LSECs (Fig. 3B; +40.3%).
Phosphorylation of eNOS-Thr495 Is Induced in WT LSECs but Reduced in Cav-1 KO LSECs After ET-1 Treatment in the Presence of LPS
Phosphorylation of eNOS at the Thr495 residue decreases eNOS activity (7). Endothelin 1 treatment did not alter the phosphorylation of eNOS-Thr495 in WT LSECs (Fig. 3C; +3.9%) or in Cav-1 KO LSECs (Fig. 3D; +5.2%). LPS also did not significantly change eNOS-Thr495 phosphorylation in WT LSECs (Fig. 3C; +7.4%) or in Cav-1 KO LSECs (Fig. 3D; −0.3%). However, treatment with ET-1 increased eNOS-Thr495 phosphorylation in WT LSECs (Fig. 3C; +24.5%; P < 0.05) but decreased the phosphorylation in Cav-1 KO LSECs (Fig. 3D; −8.8%; P < 0.05). Treatments with ET-1 and/or LPS did not change eNOS protein expression in WT LSECs (Fig. 3E) and Cav-1 KO LSECs (Fig. 3F).
LPS Inhibits eNOS Translocation to the Plasma Membrane in WT LSECs but not Cav-1 KO LSECs
In control media, eNOS is localized in the perinuclear region of both WT (Fig. 4A) and Cav-1 KO LSECs (Fig. 4E). Treatment with ET-1 induced the translocation of eNOS from the perinuclear region to the plasma membrane in both WT (Fig. 4B) and Cav-1 KO LSECs (Fig. 4F). The perinuclear localization of eNOS was maintained after LPS treatment in both WT (Fig. 4C) and Cav-1 KO LSECs (Fig. 4G). Endothelin 1 failed to induce the translocation of eNOS in WT LSECs after LPS pretreatment (Fig. 4D). In contrast, ET-1 stimulated eNOS translocation to the plasma membrane after LPS pretreatment in Cav-1 KO LSECs (Fig. 4H).
LPS Inhibits NO Production in WT LSECs but not Cav-1 KO LSECs
NO production was negligible in WT LSECs cultured in control media (Fig. 5A). Endothelin 1 induced NO production in the perinuclear region and in the plasma membrane of WT LSECs (Fig. 5B). A minimal amount of NO production was observed after LPS treatment in WT LSECs (Fig. 5C). Endothelin 1-induced NO production was abrogated in WT LSECs after LPS pretreatment (Fig. 5D).
At basal condition, NO production in the Cav-1 KO LSECs was qualitatively higher than in WT LSECs (Fig. 5A and 5E). 4,5-Diaminofluorescein fluorescence was visualized in the perinuclear region of Cav-1 KO LSECs cultured in the control media (Fig. 5E). An increasing amount of NO production was localized in the plasma membrane after the treatment of Cav-1 KO LSECs with ET-1 (Fig. 5F). NO production after LPS treatment in Cav-1 KO LSECs (Fig. 5G) was similar to the cells cultured in the media (Fig. 5E). Unlike the WT LSECs, ET-1 increased NO production in the plasma membrane of Cav-1 KO LSECs (Fig. 5H).
After endotoxemia, portal resistance is increased, and the impaired hepatic microcirculation leads to areas of ischemia and subsequently causes hepatocellular damage (8). However, the mechanisms of endotoxin-mediated inhibition of hepatic sinusoidal blood perfusion were unclear. Under inflammatory and oxidative stress conditions, ET-1 gene expression is increased (9), and the contractile effect of ET-1 in the hepatic sinusoids is exacerbated (3). However, the hyper-responsiveness to the vasoconstrictive effects of ET-1 is not related to an increase in ETA receptors. Expression of ETA receptors remains relatively constant after inflammatory and oxidative stress events (10, 11). In contrast, inflammatory and oxidative stress conditions induce an overexpression of ETB receptors (10, 11), which normally mediate vasodilation in the hepatic vasculature. This raises a paradoxical question of how an increase in ETB receptors that normally mediates vasodilation would lead to a hyperconstrictor effect in the liver after the stress conditions. Using a Cav-1 KO model in the present study, we showed that Cav-1 at least partly mediates the effects of endotoxin to disrupt the coupling between ETB receptor and eNOS that results in a hyperconstrictor effect in the liver.
Independent models of liver diseases have been shown to increase hepatic Cav-1 protein expression. For example, an increase in Cav-1 expression is detected after bile duct ligation in rats (12). In this model, the increase in Cav-1 protein expression within hepatic sinusoids and venules impairs eNOS activity and has been implicated in the development of portal hypertension (12). Other experimental models, including carbon tetrachloride-induced model of liver cirrhosis (13), chronic alcoholic liver disease model (14), and Niemann-Pick disease type C model (15), also detected an increase in Cav-1 protein level. In the present study, LPS induces Cav-1 overexpression in WT mice LSECs in conjunction with diminished eNOS activity. A causative role for Cav-1 in the process of eNOS inhibition is also supported by the demonstration that targeted mutation of Cav-1 partially reverses the deficient eNOS activity in the LPS-treated cells.
The factors within the liver that increase caveolin expression after stress conditions are not well defined. In humans, Cav-1 and Cav-2 genes are colocalized in the q31.1 region of human chromosome 7 (16). The first and second exons of these genes are embedded within CpG islands, which may suggest that the caveolin gene expression may be regulated, in part, by the methylation of these genes (17). In the promoter region, the Cav-1 and Cav-2 genes contain SP1 binding sites (18). After oxidative stress generated by hydrogen peroxide, the binding of SP1 is enhanced, and Cav-1 gene transcription is increased (19). These results showed that the signaling machinery links oxidative stress to Cav-1 gene transcription through an SP1-mediated mechanism. However, the inflammatory stress response elements within the Cav-1 promoter and the molecular mechanism that LPS up-regulates Cav-1 expression remain to be determined.
Cellular cholesterol levels may also directly link to Cav expression and signaling in the liver. This concept was introduced by the demonstration of an up-regulation of Cav-1 protein expression in the liver from mice with Niemann-Pick disease type C, which is characterized by prominent accumulation of cholesterol in the liver (15). Marked increases in serum cholesterol are also associated with a significant increase in Cav-1 expression in liver lysates after bile duct ligation in rats (12). These results were supported by studies in cultured cells showing that cholesterol influx up-regulates Cav-1 expression in bovine aortic endothelial cells (20) and in Madin-Darby canine kidney cells (21). Using methyl-β-cyclodextrin to deplete cholesterol content from the plasma membrane and disrupt the caveolae, we recently showed that the pharmacological approach reduces Cav-1/eNOS colocalization and results in an increase in eNOS activity and NO production (revised manuscript submitted).
In this study, the data from mice deficient in Cav-1 strongly support the negative regulatory effect of Cav-1 on eNOS function. We observed a 3-fold increase in NO production from Cav-1 KO mice hepatic LSECs. Removing Cav-1 from the cells increases basal eNOS activity and results in higher concentration of plasma NO. It has been estimated that plasma NO levels in Cav-1-null mice are five times higher than those in WT mice (22). These data were consistent with a Cav-1 peptide transfection model demonstrating that overexpression of the Cav-1 scaffolding domain attenuates NO production (23). However, a more mechanistic understanding is necessary to fully appreciate how Cav-1 inhibits eNOS given the complex regulation of eNOS activity by protein-protein interaction, subcellular localization, and phosphorylation.
The interaction of eNOS with CaM increases eNOS activity, whereas its interaction with Cav-1 inhibits its activity. In cirrhotic livers, the expressions of Cav-1 and CaM are up-regulated. However, the overexpression of Cav-1 not only increases its association with eNOS but also decreases the interaction of eNOS with CaM. This molecular response of Cav-1 leads to the inhibition of NO production and the increase in vascular resistance in the liver. Similarly, transduction of a constitutively active form of eNOS fails to overcome the inhibition imposed by Cav-1 up-regulation (24). These results suggest that Cav-1 overexpression in livers serves to functionally impair eNOS activity.
The caveolin-signaling hypothesis focuses on the interaction between the caveolin scaffolding domain and signaling proteins. This model would predict that the distribution of signaling proteins will be changed in the absence of Cav-1; however, this may not be the case. It has been shown that in an siRNA-mediated Cav-1 knockdown model, the abundance and subcellular fractionation of a number of signaling proteins are unaffected, including the insulin receptor, eNOS, Gαq, c-Src, and Rac (25). In this study, we showed that the ET-1-induced translocation of eNOS was not altered in Cav-1 KO LSECs and supported the idea that caveolae are not required for eNOS localization and activation. Instead, the membrane targeting of eNOS is mostly controlled by cotranslational N-myristoylation (26) and post-translational palmitoylation (27). However, our results suggested that Cav-1 is necessary to mediate the LPS inhibition of eNOS activity and showed that LPS reduced ET-1-mediated eNOS translocation and NO production in WT but not in Cav-1 KO mice LSECs. Localization of eNOS is of special interest because it may indicate the mechanism by which the eNOS is activated. For example, eNOS is predominantly activated by calcium/CaM in the caveolae; in contrast, it is mainly activated by Akt in the Golgi (28). We have recently showed that the translocation of eNOS is cyclodextrin-sensitive, and eNOS is targeted to the Golgi when the cells are treated with methyl-β-cyclodextrin (revised manuscript submitted).
The knockdown of Cav-1 enhances Akt phosphorylation without affecting the abundance and phosphorylation of phosphatase and tensin homolog, phosphoinositide-dependent protein kinase 1, and extracellular signal-regulated kinases 1/2 (25). Our data showed that ET-1 reverses the LPS inhibition of eNOS activity in Cav-1 KO mice, at least in part, by stimulating the phosphorylation of eNOS-Ser1177 and the dephosphorylation of eNOS-Thr495 residues. The increase in eNOS-Ser1177 phosphorylation may be related to the ET-1-mediated Akt activation (6). However, the mechanism of Cav-1 regulation of eNOS-Thr495 dephosphorylation is less defined (29). Because serine/threonine protein phosphatases PP1 and PP2B/calcineurin have been shown to mediate the dephosphorylation of eNOS-Thr495 (7) and the overexpression of Cav-1 inhibits PP1 and PP2A activity (29), activity of PP1 may be controlled by Cav-1 in the regulation of eNOS-Thr495 dephosphorylation.
In conclusion, we hypothesized that Cav-1 overexpression is causally linked to decreased eNOS activity in LSECs after the LPS treatment. Our results support this hypothesis and showed that LPS induced Cav-1 overexpression in WT LSECs, and the KO of Cav-1 increased basal eNOS activity and at least partially restored ET-1-mediated eNOS stimulation, eNOS translocation to the plasma membrane, and NO production in the LSECs after the LPS treatment.
The authors appreciate the editorial comments from Jacqueline J. Dienemann, MD, Didier Dréau, MD, Charles Y. Lee, MD, and Jian X. Zhang, MD, and the superior technical assistance and support of Cathy Culberson, David L. Gray, Katarzyna Korneszczuk, and Amaya Oregui from the University of North Carolina at Charlotte.
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