Bacterial endotoxin produces an inflammatory response that precipitates substantial changes in vascular regulation and cell injury. Although severe endotoxemia produces overt injury to blood vessels resulting in increased permeability, loss of vascular tone and, ultimately, shock and death, milder endotoxic stress produces alterations in vascular reactivity to specific mediators via changes in cell signaling pathways. One such mediator is endothelin-1 (ET-1). ET-1 is a potent peptide constrictor whose production in increased in a variety of inflammatory and oxidative stress conditions (1). Our work in the liver microcirculation has shown that sensitivity to the constrictor action of ET-1 is significantly increased in endotoxemia as well as in many other stresses such as ischemia/reperfusion and remote trauma (2–7). ET-1 acts on the microcirculation through the binding to two distinct receptor subtypes: ETA receptors are expressed on hepatic stellate cells and vascular smooth muscle cells and always mediate constriction while ETB receptors are also expressed on the endothelial cells where they are coupled to endothelial nitric oxide synthase (eNOS) activation thus mediating dilation (8–11).
Recent work from our group has shown that endotoxemia effectively uncouples ET-1 binding to ETB receptors on hepatic sinusoidal endothelial cells from activation of eNOS, thus decreasing the compensatory dilation. This decrease in eNOS activation is associated with increased binding of eNOS to caveolin-1 and altered patterns of phosphorylation at key regulatory sites (12–14) as well as from the effects of hydrogen sulfide production (15). These changes would result in lowered overall activity of the enzyme resulting in decreased NO production. eNOS can also be regulated by availability of substrate and cofactor (16–18). In order for eNOS to effectively produce NO, it requires not only activation by Ca-Calmodulin and appropriate phosphorylation but also adequate supplies of L-arginine and tetrahydrobiopterin (16–21). When these are limited, the superoxide formed by donation of an electron from NADPH to O2 can be directly released rather than combined with L-arginine to complete the formation of NO and L-citrulline. Given that such uncoupling of the enzyme from NO production is not likely to be complete, the same enzyme would be producing NO and superoxide providing the perfect environment for generation of peroxynitrite (22). Thus an uncoupling of the eNOS enzyme, in addition to decreasing NO availability, would also contribute to vascular dysregulation via the production of reactive oxygen and reactive nitrogen species (ROS and RNS, respectively). Recent studies on pulmonary hypertension have shown that ET-1 can activate NADPH oxidase resulting in increased superoxide that can then react with NO, but it is not known whether eNOS itself produces reactive oxygen in endothelial cells in endotoxemia. Therefore, in this study we tested whether eNOS might contribute to ROS production in vascular endothelial cells in response to endotoxin and whether this can be ameliorated by increased availability of either substrate (L-arginine) or cofactor (BH4).
MATERIALS AND METHODS
First passage Human Umbilical Vein Endothelial Cells (HUVECs) were purchased from Cambrex (Walkersville, MD). RPMI 1640 and heparin were purchased from Invitrogen (Carlsbad, CA). Endothelial cell growth supplement was obtained from BD Biosciences (Bedford, MA). ET-1 was purchased from American Peptide (Sunnyvale, CA). All antibodies were acquired from BD Transduction Laboratories (San Diego, CA). 5-(and-6)-carboxy-2′7′-dichlorodihydrofluorescein diacetate (DCF), MitoSOX, and Hoechst were purchased from Molecular Probes Inc (Eugene, OR). DCF, MitoSOX, and Hoechst were dissolved in dimethyl sulfoxide (DMSO) as a stock solution and kept frozen at −20°. For cell loading, fluorescent probes were mixed with Hank's Balanced Salt Solution (HBSS, Life Technologies, Carlsbad, CA) warmed to 37°C. Unless otherwise noted, all other reagents were purchased from Sigma Aldrich (St. Louis, MO).
HUVECs were cultured in 10% fetal calf serum (FCS) supplemented endothelial cell growth media (RPMI 1640, 0.03 mg/mL EC growth supplement, 0.059 mg/mL heparin, 0.72 μg/mL insulin, 10 μM dexamethasone, 25 μg/ml gentamycin, 1X P/S/F) and seeded in 12 of 24 well plates at 1 × 105/0.5 × 104 cells per well respectively and allowed to attach overnight in 10% FCS media. Cells were quiesced the day following seeding in 0.1% FBS media for 24 h. HMECs were a gift from Dr Vijay Kumar Kalra (University of Southern California, Los Angeles, CA). They were cultured in MCDB 131 growth medium without l-glutamine, 10% heat-inactivated fetal bovine serum (FBS), 10% antibiotic antimycotic, 10% GlutaMAX (Life Technologies), 10 ng/mL endothelial growth factor, and 1 μg/mL hydrocortisone. The cells were maintained on collagen-coated 24-well plates in 5% CO2/95% air at 37°C.
The conversion of [3H]L-arginine to [3H]L-citrulline was used to assess NOS activity in intact HUVECs, as we previously described with a few modifications (12). All experiments were conducted in duplicates using Ionomycin (Io), a calcium ionophore as a positive control for NOS activity and L-Iminoethyly Ornithine (L-NIO) as an NOS inhibitor to assess basal NOS activity Following the 24 h the cells were washed ×2 in DPBS and treated with 250 ng/mL LPS or 250 ng/mL LPS and 100 μM Sepiapterin for 6 h in 1% FBS supplemented media. After 6 h the cell monolayers were washed once with Krebs Hepes Buffer (KHB) (NaCl 99 mM, KCl 4.7 mM, MgSO4 1.2 mM, KH2PO4 1.0 mM, CaCl2 1.9 mM, NaHCO3 25 mM, HEPES 20 mM) and incubated at 37°C for 30 min with KHB containing 100 μM (L-NIO for 20 min. One μCi/μL [3H]L-Arginine (10 μM) was added to each well followed by stimulation with ET-1 (10 nM) for 30 min at 37°C. The reaction buffer was aspirated and the cell monolayers were washed ×1 with KHB to remove extracellular radioactivity. The plate was then placed on ice and 150 μL of lysis buffer (5 mM Tris (pH 7.4), 20 mM EDTA, 0.5%v/v Triton X-100) was added to each well and the reaction was terminated by the addition of cold PBS (L-Arginine 5 mM, EDTA 4 mM, EGTA 4 mM). The wells were scraped to completely lyse and detach the cells and the lysate was centrifuged at 10,000 rpm for 5 min to precipitate the cellular debris. A 150-μL aliquot of the centrifuged lysate was passed through a cation exchange chromatography resin (Dowex AG 50W-X8, Molecular Biology Grade, 200–400 mesh sodium form, BioRad) equilibrated in stop buffer (20 mM HEPES, 5 mM EDTA, 5mMEGTA, pH 5.5) to separate [3H]L-arginine from the [3H]L-citrulline. The resin was washed with stop buffer to elute the samples. Eight hundred fifty microliter of liquid containing [3H]L-citrulline was collected in scintillation vials. Sixml of scintillation fluid (SX0-5 Scinti-safe Econo-1, Fisher Scientific, Pittsburgh, PA) was added and the samples were counted on a liquid scintillation counter (Beckman LS6000 series, Beckman Instruments Inc, Fullerton, CA).
Ferricytochrome C reduction was used to determine ROS production in HUVECs (23). Cells were seeded in 24-well plates at an initial density of 5 × 104 cells/well and allowed to attach overnight in 10% FBS supplemented growth media. Cells were quiesced the day following seeding in 0.1% FBS supplemented media for 24 h. Following the 24 h the cells were washed ×2 in DPBS and treated with 250 ng/mL LPS or LPS (250 ng/mL) and Sepiapterin (100 μM) for 6 h in 1% FBS supplemented media. In order to determine if eNOS was the source of the reactive oxygen radicals, NOS inhibitor L-NIO was used to target eNOS as the putative source. The cells were washed once with Krebs Hepes Buffer (KHB) to remove all remaining media and incubated at 37°C for 30 min with KHB containing 50 μM cytochrome c in the presence or absence of SOD (350 units/mL), or in the presence of 100 μM L-NIO. Following the 30-min incubation that allows for reduction of cytochrome C, absorbance of the medium was read spectrophotometrically (μQuant, BIO-TEK Instruments Inc, Winooski, Vt) at 550 nM. Reduction of cytochrome c in the presence of SOD was subtracted from the values without SOD.
Since cytochrome C may largely reflect extracellular ROS, we confirmed the synergistic effect of ET-1 and LPS using DCF. On the day of the experiment, medium was removed and replaced with 1% FBS medium and incubated with or without LPS for 6 h. Medium was removed and cells were incubated with a 4 μM Hoechst + 25 μM DCF-DA staining solution for 20 min. The cells were washed three times and HBSS warmed to 37°C was placed in each well. MitoSOX staining solution was added to each well to a final concentration of 0.5 μM. Loaded cells were placed in a multiwell fluorescence plate reader with temperature maintained at 37°C. The excitation filter was set at 360/40 and the emission filter was set at 460/40. The fluorescence from each well was captured to normalize for total cell count prior to the addition of MitoSOX. The excitation filters were then set at 485/20 and 525/20 and the emission filters were set at 530/25 and 590/35, respectively, for simultaneous measurement or DCF and MitoSOX fluorescence. The fluorescence of the cells from each well was measured and recorded for 30 min.
Western blot analysis
Cells were cultured, plated, and treated in the same manner as for other experiments. Following the 6-h treatment with LPS and 30-min incubation with ET-1 cells were washed with KHB to remove all media and the cells were lysed in 30-μL RIPA lysis buffer (20 mM Tris-HCl (pH 7.5), 150 mM NaCl, 2 mM EDTA, 1%NP-40, 0.5% deoxycholate, 0.1% SDS, 50 μM PMSF, 50 μM Na3VO4, 2 μg/mL aprotinin). The lysate was then centrifuged at 15,000 rpm for 10 min to remove debris. The samples were boiled for 5 min in a 1:1 mixture with Laemmlli loading dye (10%BME) prior to separation on a 10% for resolution of eNOS and 15% gel to resolve the smaller 17Kda Calmodulin. Proteins were transferred to nitrocellulose membrane and then stained with Ponceau S to confirm equal loading. The membranes were washed with TBS and blocked for 1 h in 1% blocking buffer (Roche Diagnostic, Indianapolis, IN). Membranes were then incubated with primary antibodies overnight. Following recognition of protein by primary antibody the membranes were washed four times in TBS with 0.1% Tween 20 and incubated for 1 h with a horseradish peroxidase conjugated secondary antibody (Jackson ImmunoResearch Laboratories Inc, West Grove, PA). The membranes were washed with TBS-Tween to remove nonspecific binding and exposed to enhanced chemiluminescence reagent (Roche) for 2 min and exposed to Biomax film (Fisher Scientific). To reprobe with different primary antibodies, the membranes were stripped with Blot Restore (Chemicon, Temecula, CA), blocked with the blocking buffer, and incubated with alternate primary antibody.
eNOS dimerization by LT-SDS page
Western blot analysis was performed as described previously (24). Briefly, cells were lysed in a non-denaturing lysis solution containing 50 mM Tris–HCl pH 8, containing 0.2% Nonidet P-40, 180 mM NaCl, 0.5 mM EDTA, 100 mM phenylmethylsulforyl fluoride, 1 M DTT, and protease inhibitors. Equal amounts of cellular proteins were resolved by 6% SDS-PAGE and transferred to nitrocellulose membranes. To investigate eNOS homodimer formation in endothelial cells, low-temperature SDS-PAGE was employed as described previously (16). Membranes were incubated with a 1:1,000 dilution of mouse anti-eNOS monoclonal antibody and 1:5000 dilution of anti-mouse IgG secondary antibody.
All data are presented as mean ± SEM of six separate experiments. Two-way analysis of variance (ANOVA) was employed to test for statistical difference between the various groups. A P < 0.05 was considered to be significant and all analysis was performed using Sigma-Stat.
Effect of treatments on cell viability
Preliminary trypan blue exclusion studies were done to evaluate the effect of the different treatments on cell viability. Cells were treated with concentrations of LPS ranging from 100 ng/mL to 1 μg/mL and it was found that concentrations above 250 ng/mL induced cell death (data not shown). Hence, 250 ng/mL was used in this model to characterize the effect of LPS and ET-1 on ROS production in HUVECs and HMECs.
LPS disrupts ET-1-induced NOS activity
Initial tests were done to examine the effects of LPS on ET-1-stimulated NO production in HUVEC. Confluent HUVECs were treated with LPS (250 ng/mL) for 6 h prior to stimulation with ET-1 (10 nM) for 30 min and NOS activity was assessed. The NOS inhibitor L-NIO abolished basal NOS activity and was subtracted as background from the other experimental groups. ET-1 significantly increased NOS activity in control cells by 1.4-fold (Fig. 1). HUVECs exposed to a 6-h LPS treatment did not show significantly altered basal NOS activity. LPS pretreatment impaired ET-1-induced NOS activity as indicated by the decreased conversion of [3H]L-arginine to [3H]L-citrulline. ET-1 is an activator of eNOS, other free radical production was investigated as an alternative to NO production.
ET-1 stimulates ROS production following LPS pretreatment
There are multiple generators of ROS in the vasculature including xanthine oxidases and NADH reductase. eNOS has been shown to be uncoupled when exposed to various stresses. Reactive oxygen generation by eNOS was quantified by SOD inhibitable ferricytochrome C reduction following 6-h LPS treatment. To identify NOS as a source of ROS, L-NIO, the NOS inhibitor was used to block NOS activity. ET-1 in control cells did not have a significant effect on ROS generation (Fig. 2). A 6-h LPS treatment increased ROS production, but this increase was not significant. ET-1 stimulation of LPS-treated cells resulted in an exacerbated release of ROS from eNOS. This response was abrogated by pretreating the cells with L-NIO for 30 min. This concomitant release of ROS with the decreased NO production from eNOS is likely to be related to various factors that have been shown to result in increased ROS generation from eNOS (19, 25–27). Since cyt C is largely extracellular, we tested whether the ROS was produced intracellularly. DCF-DA is taken up by cells and de-esterfied, thus trapping it intracellularly. Figure 3 shows that ET-1 increased ROS intracellularly, although not significantly. LPS plus ET-1, however, resulted in a significant increase in ROS as indicated by increased DCF fluorescence. On the other hand, there was no difference in any group in Mitosox fluorescence suggesting that mitochondria were not a significant contributor to the ROS produced. Western Blot showed that there was no altered calmodulin or eNOS protein expression (Fig. 4), although oxidized calmodulin was not examined. Tetrahydrobiopterin and L-arginine, the critical factors in eNOS stability, were examined as regulators of ROS generation from eNOS.
Arginine supplementation does not decrease ET-1-induced SO
Decreased L-arginine availability to eNOS can cause the heme-oxy complex to dissociate and generate oxygen-free radicals. HUVECs were treated with LPS for 6 h and then ET-1-induced SO production was measured in the presence of 1 mM L-arginine in the extracellular medium. ROS generated through ET-1 stimulation of LPS-treated HUVECs was not altered by increasing availability of L-arginine in the extracellular medium. LPS-induced ROS was not altered by the excess L-arginine either (Fig. 5). This indicates that there are other factors that play a role in the production of ROS from eNOS.
Sepiapterin supplementation decreases LPS-induced SO production
Sepiapterin is a precursor of tetrahydrobiopterin and is converted intracellularly by sepiapterin reductase to tetrahydrobiopterin. HUVECs were treated with LPS in the presence or absence of sepiapterin for 6 h and then ROS generation was assessed in the presence and the absence of ET-1. As observed in prior experiments, LPS treatment showed increased ROS production with ET-1 further exacerbating this response. The groups that were incubated with sepiapterin and LPS for 6 h showed little or no SOD inhibitable ferricytochrome C reduction in the presence or more importantly in the absence of ET-1 (Fig. 6). Sepiapterin, which is converted to tetrahydrobiopterin intracellularly, inhibits the ET-1-stimulated production of ROS from eNOS. Consequence of LPS-induced reduction in ET-1-stimulated NO production is increased SO production. Conversely, it was examined if decreased SO production by supplementation of sepiapterin restored the ET-1-induced eNOS activity that was disrupted by LPS treatment.
Sepiapterin does not restore ET-1 induced NO production
If tetrahydrobiopterin was the crucial factor in the LPS-mediated disruption of ET-1-induced NO production then augmenting levels of intracellular BH4 should restore normal NOS activity. Following a 6-h treatment with LPS and sepiapterin, [3H]L-arginine and ET-1 were added to the cells in an arginine-free KHB buffer and 30 min later the cells were lysed and NOS activity was assessed as described in the methods. Augmenting BH4 did not restore ET-1-induced NOS activity as predicted (Fig. 7). Sepiapterin had no effect on NO production from LPS treated cells.
Effect of LPS and ET1 on eNOS dimerization
eNOS is active as a homodimer. So if LPS disrupts the dimer ET-1 will not activate the enzyme to produce NO. Thus, we looked at the dimer to monomer ratio following LPS and ET-1 treatments. The cells were lysed in a non-denaturing lysis buffer and low-temperature SDS PAGE (LT-SDS PAGE) was run as described. Treatment with ET-1 for 30 min following a 6-h LPS treatment resulted in significantly lower levels of the protein dimer (Fig. 8). LPS by itself does not seem to significantly alter the dimer to monomer ratio. Sepiapterin loading in LPS-treated cells abolishes the ET-1-induced monomerization that is seen when the cells are treated with LPS alone.
The vascular endothelium is now recognized to be a major site of vascular regulation under both unstressed and stressed conditions. This includes regulation of adhesion of inflammatory cells as well as production of endothelium-dependent dilators such as nitric oxide and endothelium-dependent constrictors such as endothelin-1. Although reactive oxygen signaling has long been recognized to contribute to vascular injury in inflammatory stresses such as endotoxemia, the major source of the radicals has traditionally been considered to be neutrophils and macrophages. In contrast, NO produced by eNOS in the endothelial cells is generally considered to have an anti-inflammatory effect in part due to its ability to scavenge superoxide. Our results indicate that, following a relatively mild endotoxin stress endothelial cells not only fail to synthesize NO in response to endothelin-1 stimulation, but also increase the production of ROS. Moreover, we show that the shift to superoxide production is caused largely by a functional deficiency of BH4.
Recent studies from our laboratory have shown that endotoxin treatment either in vivo or in culture results in a functional inhibition of ET-1-stimulated eNOS activity (12, 15, 28). This functional inhibition is at least in part due to overexpression of caveolin-1 (12, 13, 28) and the influence of increased H2S production (29). This partly explains the increased constrictor response of the hepatic vasculature to ET-1 following stresses such as bacterial endotoxin. Our present results show a very similar response in HUVECs, suggesting that this uncoupling of NO production from ET-1 binding is not peculiar to hepatic sinusoidal endothelial cells. HUVECs are more easily maintained in culture and can thus provide a useful model for studying this response although results from cell culture studies must always be applied carefully to in vivo conditions. Along with the increased constrictor response, we observed and increase in cellular injury following ET-1 infusion in livers pretreated with LPS but not in controls given ET-1 without LPS pretreatment. This increased injury is in excess of what would be expected from the decreased blood flow alone suggesting that other mechanisms are also important.
It is well known that generation of ROS contributes to injury following endotoxemia; however, these ROS are considered to be produced primarily by infiltrating neutrophils and macrophages or by NADPH oxidase in both inflammatory and other cells (30). In the liver, the Kupffer cells are a particularly important source of reactive oxygen. However, given the short half life of these reactive species, even moderate amounts of reactive oxygen produced in critical locations can be of great functional importance. Our work has shown that reactive oxygen has a significant impact on regulation of local circulation by endothelial cells. Pretreatment with H2O2 for 6 h resulted in a significant upregulation of caveolin-1, alteration in phosphorylation pattern of eNOS toward one that favors inactivation of the enzyme and, ultimately, attenuated activation of eNOS by ET-1. The results of our present experiment demonstrate that after LPS treatment, reactive oxygen is produced within the endothelial cells in response to ET-1 stimulation. Although the exact target of the ROS that results in altered gene expression and phosphorylation patterns are not known, production of ROS in a cell such as vascular endothelium which has such a sparse cytoplasm would allow availability of effective concentrations throughout the cell.
Uncoupling of the eNOS enzyme so that it produces superoxide rather than nitric oxide can result from either inadequate substrate or inadequate cofactor. Inflammatory stresses such as endotoxemia have significant effects on expression of iNOS in vivo. The high activity of iNOS can lead to excessive consumption of L-arginine. In addition, LPS is known to cause downregulation of amino acid transporters including the one for L-arginine. Therefore, we tested whether L-arginine supplementation might reverse the inhibition of NO production and increase in ROS production. Our results showed that L-arginine supplementation had no effect on either parameter. From these results, we conclude that either L-arginine availability is not limiting in the effective production of NO by eNOS or that supplementation with exogenous L-arginine is not effective in overcoming the limitation. Considering the several reports showing that L-arginine supplementation can overcome L-arginine deficiency, we would conclude that L-arginine is probably not limiting following endotoxin treatment of endothelial cells. This conclusion is further supported by the almost complete abrogation of superoxide production in response to ET-1 stimulation in LPS-treated endothelial cells by sepiaterin. Sepiaterin is a precursor of BH4 that has been shown to effectively enter the cell and replete BH4 levels. A major mechanism by which BH4 functions in regulating eNOS activity is via the enhancement of dimerization of eNOS.
Vascular pathological complications have been linked to endothelial dysfunction by a variety of different studies (31, 32). The causative molecular mechanisms of the dysregulation following an inflammatory stress have not been clearly elucidated to date although oxidative stress has been widely implicated. LPS is directly associated with inflammation, immune system activation, and increased formation of reactive oxygen species. All three are key factors in the pathogenesis of heart disease, atherosclerosis, diabetes, and other vascular conditions. LPS primes the system for ROS release and in turn for cellular proliferation, migration as well induction of the NF-kB and AP-1 pathways. Other results of the increase in oxidative stress are oxidation and nitrosylation of critical cellular components which includes structural proteins, enzymes, and even immunoglobulins. Decreased NO bioavailability has been linked to a vast number of causes including decreased L-arginine availability (33), modifications of upstream signaling molecules including Akt and PI3Kinase (34), as well as altered calcium (35), downregulation of GTP-cyclohydrolase leading to decreased intracellular tetrahydrobiopterin (24, 36). NO can also be scavenged by superoxide anions as it is being produced in the absence of a normal redox system (37). This can also lead to diminished NO bioavailability. As a paracrine nitrogen radical, NO diffuses from the endothelial cells into the smooth muscle and causes relaxation of the smooth muscle through a cyclic GMP and PKG-dependent pathway.
We have previously shown in sinusoidal endothelial cells that ET-1-induced NO production is disrupted by LPS pretreatment (12, 28, 29). We hypothesized that endotoxin uncouples eNOS in endothelial cells leading to a decrease in ET-1-stimulated NO production with a concomitant increase in the generation of oxygen radicals and induction of the subsequent oxidative stress. Hence, this study was designed to investigate the mechanisms by which LPS affects NOS function and in HUVECs. ET-1 can act both as a vasodilator and a vasoconstrictor and hence any imbalance in its effect can theoretically induce an exacerbated vasodilatory or vasoconstrictive response depending on which side of the scale is tipped. This study showed that LPS suppressed ET-1-induced NO production by eNOS. Concomitantly, there is an increased production of reactive oxygen species in response to ET-1 following the LPS stress. This increase is not seen when the cells are pretreated with the irreversible eNOS blocker L-NIO, indicating that eNOS is a major contributor to LPS-induced reactive oxygen production in response to ET1 in this particular model. ROS can oxidize intracellular cofactors of eNOS (tetrahydrobiopterin, CaM, L-arginine, and NADPH) leading to decreased eNOS efficiency. Calcium–CaM complexes bind to the eNOS homodimer allowing for efficient electron flow from the reductase domain to the oxygenase domain. A western blot using CaM antibodies did not detect any significant changes in CaM levels (data not shown), indicating that the increased ROS production was not due to decreased electron channeling from the reductase domain.
Sepiapterin increases intracellular BH4 via the salvage pathway. When HUVECs were simultaneously incubated with sepiapterin and LPS, the ET-1-induced release of reactive oxygen was abrogated. L-arginine supplementation did not reduce reactive oxygen release. LT-SDS PAGE showed that there was a significant induction of monomerization of the eNOS homodimer following LPS and ET-1 treatments. Simultaneous LPS and sepiapterin loading seemed to stabilize the protein homodimer and abolished the monomerization that is seen when the cells are incubated with LPS in the absence of sepiapterin. Intriguingly, suppression of ET-1-induced NO production by LPS was not rescued by sepiapterin loading. SO production on the other hand is reduced when the cells are treated with sepiapterin at least in part due to the restoration of the homodimer.
These results suggest that LPS decreases NO production by eNOS through a mechanism that seems to be partially independent of ROS production as the treatment that rescues ET-1-induced ROS production does not have any significant effect on restoring the disrupted ET-1-induced NO production. Another possible explanation is the multifactorial effect on eNOS by the chronic LPS stress as previously described in our laboratory (38). Sepiapterin loading rescues the LPS primed ET-1-induced ROS production. This result indicates that LPS decreases intracellular BH4 bioavailability. BH4 is critical for the stability of the eNOS homodimer and particularly the stable conformation of the oxygenase domain (39). Thus, a decrease in BH4 destabilizes the eNOS homodimer, which is then susceptible to monomerization upon ET-1 treatment. Once monomerized an activated eNOS oxygenase domain uses the electrons flowing from its own reductase domain to reduce the heme-oxy complex in the active site to produce superoxide radicals. Another mechanism of generating superoxide is by the spontaneous dissociation of the heme-oxy complex in the absence of enzyme activity (39). These superoxide radicals can scavenge NO in the endothelium leading to an even further decrease in NO availability to the smooth muscle resulting in a hyperconstricted state of the vasculature. Free radicals also lead to smooth muscle proliferation and have severe artherogenic effects among other deleterious effects to the local environment (40). LPS-induced disruption of ET-1-induced NO production is not averted by the sepiapterin loading leading us to believe that the decreased ET-1-induced enzyme activity following a chronic endotoxin stress is not only due to the lack of pterin cofactor availability but due to disruptions in other cellular mechanisms as well.
In the overall context of the hepatic dysfunction during sepsis, our in vitro studies add mechanistic insights into the causes of vascular dysregulation. Our previous reports show that microvascular dysfunction as evidenced by enhanced vasoconstriction in response to endothelin occurs in polymicrobial sepsis (41) as well as in endotoxemia. Although overexpression of caveolin-1 by sinusoidal endothelial cells accounts for a significant portion of the impaired eNOS activation, the exact mechanism by which caveolin-1 is overexpressed is not well understood. We have shown that oxidative stress independently produces caveolin-1 overexpression (42) and Kalivendi et al. (43) have shown that ROS can also affect BH4 levels. This suggests that production of ROS and upregulation of caveolin-1 act together, and possibly in a positive feedback manner, to cause impaired hepatic blood flow regulation in sepsis-related conditions via impaired activation of eNOS. Complete elucidation of the contribution of these mechanisms will require further in vivo studies.
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