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INTRODUCTION |
Sphingosine 1-phosphate
(S1P),1 generated from
sphingosine by sphingosine kinase, is a recently described lipid
mediator regulating cellular responses, including cell proliferation
(1), cell motility (2, 3), and morphological changes (2, 4) in
endothelial cells. In addition, S1P was also demonstrated to protect
endothelial cells from apoptosis induced by C2-ceramide, TNF
, and anti-Fas antibody (1, 4, 5). All these effects of S1P on
endothelial functions are thought to correlate with the intracellular
signaling pathway linked to the EDG-1 (endothelial differentiation gene-1), EDG-3, and EDG-5
subtypes of G protein-coupled receptors (1-4, 6). Of these receptors,
EDG-1 is exclusively expressed in HUVECs (2, 4), and S1P induces a
robust calcium response mainly via the Gi-coupled S1P
receptors EDG-1 and -3 in HUVECs (4). Consistent with this view, we
have recently observed that intracellular Ca2+ mobilization
evoked by the Gi-PLC system is responsible for the signaling pathway of mitogen-activated protein kinases, focal adhesion kinase (p125FAK), and chemotaxis in response to
S1P in HUVECs (6). It is likely that intracellular signaling events
of S1P in endothelial cells may be directly associated with increase in
[Ca2+]i.
Endogenous nitric oxide (NO) is synthesized from L-arginine
by catalytic reaction of three isotypes of NO synthases (NOS), the
neuronal or type I isoform (nNOS), the inducible or type II isoform
(iNOS), and the endothelial or type III isoform (eNOS) (7, 8).
Endothelium-derived NO is formed by eNOS, which is constitutively
expressed in endothelial cells and localized in plasmalemmal caveolae
through association with the caveolae integral membrane structural
protein caveolin-1 (9). Binding of eNOS with caveolin-1 inhibits the
eNOS catalytic activity, and the inhibitory effect of caveolin-1 on the
eNOS activity can be reversed by Ca2+-calmodulin (10).
Thus, increase in [Ca2+]i elicited by diverse
extracellular signals (stimuli), including shear stress (11),
bradykinin (12), and Na+/Ca2+ exchange (12,
13), leads to activation of eNOS (8), resulting in increased NO production.
Numerous previous studies have demonstrated the role of eNOS in
controlling blood pressure, vascular remodeling, angiogenesis, and
apoptosis (14-17). In particular, NO is implicated as a cytoprotective effector molecule, protecting some cell types from apoptotic cell death
induced by TNF
, anti-Fas antibody, LPS, and trophic factor withdrawal. These apogenic stimuli activate a series of tightly controlled intracellular signaling events that induced mitochondrial cytochrome c release into cytosol and the activation of
cysteine proteases known as caspases (18). Inappropriate endothelial cell apoptosis may be linked to several cardiovascular diseases. For
example, eNOS-deficient mice exhibit delayed angiogenesis thought to be
due to decreases in endothelial cell migration, proliferation, and
differentiation, compared with wild type controls (19). In addition, we
have shown that transduction with NOS gene into aortic allografts
suppresses the development of allograft arteriosclerosis (20), possibly
in part via suppression of endothelial apoptosis (21). The
antiapoptotic effects of NO are associated with direct inhibition of
caspase activity by S-nitrosylation of the catalytic
cysteine residue of the enzymes, as well as through cGMP-dependent mechanism (22). Therefore, eNOS-activating
agents are capable of protecting endothelial cells from apoptosis. We hypothesized that S1P would protect endothelial cells by
Ca2+-dependent activation of eNOS. Here we show
that S1P increases NO production in HUVECs by elevating
[Ca2+]i through the EDG-1 and
-3/Gi-PLC pathway. This increased NO production prevents
serum-deprived apoptosis by inhibiting mitochondrial cytochrome
c and release caspase-3 activation/activity.
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MATERIALS AND METHODS |
Chemicals and Reagents--
S1P, U73122, and BAPTA-AM were
purchased from Biomol (Plymouth Meeting, PA). Pertussis toxin was from
Research Biochemical International. M199, penicillin, streptomycin,
L-glutamate, and heparin were obtained from Life
Technologies, Inc. Basic fibroblast growth factor was from
Upstate Biotechnology (Lake Placid, NY). N-Monomethyl-L-arginine (NMA),
N-acetyl-Asp-Glu-Val-Asp-p-nitroanilide (Ac-DEVD-pNA) and z-VAD-fmk were obtained form Alexis Corp. (San Diego,
CA). Antibodies against caspase-3 and eNOS were purchased from
Transduction Laboratories (Lexington, KY), and a mouse monoclonal anti-cytochrome c oxidase antibody was from Molecular
Probes. 1H-(1,2,4)oxadiazolo[4,3-a]-quinoxaline-1-one (ODQ)
was purchased from Promega (Madison, WI).
S-Nitroso-N-acetyl-D,L-penicillamine (SNAP) was synthesized, as described previously (23). The following 18-mer phosphothiate oligonucleotides were synthesized to block the
expression of EDG-1 and EDG-3: antisense EDG-1, 5'-GAC GCT GGT GGG CCC
CAT-3'; sense EDG-1, 5'-ATG GGG CCC ACC AGC GTC-3'; antisense EDG-3,
5'-CGG GAG GGC AGT TGC CAT-3'; sense EDG-3, 5'-ATG GCA ACT GCC CTC
CCG-3'. All other reagents were purchased from Sigma, unless indicated otherwise.
Cell Culture--
HUVECs were isolated as described previously
(6). The cells were grown onto a gelatin-coated 75-cm2
flask in M199 with 20% fetal bovine serum, 100 units/ml penicillin, 100 mg/ml streptomycin, 3 ng/ml basic fibroblast growth factor, and 5 units/ml heparin at 37 °C under 5% CO2, 95% air. To
induce apoptosis, serum-free medium without basic fibroblast
growth factor was used. The cells used in this study were between
passages 2 and 7.
Nitrite and Nitrate Measurement--
Production of nitrite plus
nitrate (NOx) was measured by ozone-chemiluminescence method. Culture
media from HUVECs were collected and assayed for NOx in the
chemiluminescent NO analyzer NO 2000 analyzer (Ki-Woo Biotech, Seoul,
Korea) and quantified with sodium nitrate as a standard.
Cell Viability--
HUVECs (1 × 105
cells/well) were plated onto six-well plates in 1 ml of M199 containing
20% fetal bovine serum. The next day, the cells were switched to
serum-free M199 with or without 20% fetal bovine serum and S1P. After
24 h, cell viability was assessed by trypan blue exclusion. Counts
were performed on triplicate wells.
Western Blot Analysis--
Cell pellets were suspended in
ice-cold sterilized water and kept on ice for 1 min. The suspension was
mixed with an equal volume of buffer A (20 mM HEPES, pH
7.5, 10 mM KCl, 1.5 mM MgCl2, 1 mM EGTA, 1 mM EDTA, 0.5 mM
phenylmethanesulfonyl fluoride, 5 µg/ml aprotinin, 5 µg/ml
pepstatin, and 10 µg/ml leupeptin) containing 500 mM
sucrose solution and carefully homogenized in a Dounce tissue grinder
with a loose pestle. Cytosols were obtained for measuring cytochrome
c release by centrifugation at 100,000 × g
for 1 h. For Western blot of caspase-3, cells (2 × 105 cells) were suspended in 100 mM Tris-HCl
buffer (pH 7.4) and lysed by three freeze-thaw cycles. Cytosolic
proteins were obtained by centrifugation at 12,000 × g
for 20 min at 4 °C. Proteins (40 µg) were separated on 14%
SDS-PAGE for caspase-3 and then transferred to nitrocellulose. The
membranes were hybridized with antibodies against caspase-3 and
cytochrome c, and protein bands were visualized by exposing
to x-ray film, as described previously (23).
Assay for DEVDase Activity--
The cell pellets were washed
with ice-cold phosphate-buffered saline and resuspended in 100 mM HEPES buffer, pH 7.4, containing protease inhibitors (5 µg/ml aprotinin and pepstatin, 10 µg/ml leupeptin, and 0.5 mM phenylmethanesulfonyl fluoride). The cell suspension was
lysed by three freeze-thaw cycles, and the cytosolic fraction was
obtained by centrifugation at 12,000 × g for 20 min at
4 °C. DEVDase activity was assayed in the presence or absence of 20 mM DTT by measuring the increased absorbency at 405 nm
after cleavage of 150 µM Ac-DEVD-pNA (22).
In Vitro Cleavage of
PARP--
[35S]Methionine-labeled PARP was synthesized
using a transcription/translation-coupled transcription and translation
system (Promega). Aliquots (4 µl) of in vitro translated
35S-labeled PARP were incubated with 4 µg of cytosolic
protein in 10 µl of the total reaction volume at 37 °C for 1 h. The reaction was stopped by mixing with an equal volume of 2 × SDS sample buffer and heating the mixture for 2 min. Cleavage profiles
of PARP were examined by electrophoresis on 10% SDS-PAGE and protein
visualized by fluorography.
Determination of NOS Activity--
Cells were cultured in 60-mm
culture dishes with S1P for different time periods, washed twice, and
equilibrated for 20 min with HEPES buffer, pH 7.4, containing: 10 mM HEPES, 145 mM NaCl, 5 mM KCl, 1 mM MgSO4, 1 mM CaCl2,
and 1 mM glucose. Cells were incubated for 40 min with 1 µCi of L-[3H]arginine plus 8 µM L-arginine. To assess the calcium
dependence of NOS activation, experiments were performed in
calcium-free HEPES buffer containing 20 µM BAPTA-AM. The
reaction mixture was removed, and ice-cold ethanol (0.5 ml) was added.
After allowing for evaporation of ethanol, 2 ml of 10 mM
HEPES-Na, pH 5.5, was added and kept for 20 min. The supernatant was
collected and applied onto a 2-ml cation-exchange column (50W-X8 resin
(converted to Na+ form), Bio-Rad).
[3H]Citrulline product was collected in the eluate by
washing the column with 4 ml of H2O and then quantified by
scintillation counting. Enzyme activity was normalized to protein
concentration. NOS activity was expressed as pmol/mg protein.
DNA Fragmentation--
DNA fragmentation was demonstrated by
harvesting total cellular DNA and agarose gel electrophoresis. After
the indicated treatment, both adherent and detached cells were
harvested, washed with phosphate-buffered saline, and lysed in 50 mM Tris-EDTA, pH 7.4, 1% SDS, and 0.5 mg/ml proteinase K
for 3 h at 50 °C. Samples were then extracted with
phenol/chloroform and precipitated with ethanol. The pellet was
resuspended in 20 mM Tris-EDTA, pH 8.0. After digesting RNA with RNase (0.1 mg/ml) at 37 °C for 1 h, DNA was separated by electrophoresis on a 1.2% agarose gel stained with ethidium bromide.
Other Analysis--
Protein concentration was determined with
the BCA assay (Pierce). Data are presented as mean ± S.D. of at
least three separate experiments, except where results of blots are
shown in which case a representative experiment is depicted in the
figures. Comparisons between two values analyzed using Student's
t test. Differences were considered significant when
p < 0.05.
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RESULTS |
S1P Protects HUVECs from Serum-deprived Apoptosis--
Previous
studies showed that trophic factor deprivation induces apoptotic cell
death in several cell types, including endothelial cells (24), and that
S1P protects endothelial cells from C2-ceramide- and
TNF
-induced apoptotic cell death (25, 26). To investigate whether
S1P protects HUVECs from serum deprivation-induced cell death, HUVECs
were cultured in serum-free M199 medium with or without S1P, and
cell viability was assayed by trypan blue exclusion after 24 h.
Serum deprivation decreased cell viability to ~30% of control cells
grown in the presence of serum. The addition of S1P (0-5
µM) reversed the serum-deprived effects associated with
cell death in a dose-dependent manner (Fig.
1). Cells can die by either necrosis or
apoptosis, and the inhibition of caspase activation/activity can
protect cells from apoptotic cell death, but not from necrotic cell
death (27). HUVECs were cultured in serum-free medium with or
without the caspase inhibitor z-VAD-fmk. z-VAD-fmk prevented
serum-deprived cell death (Fig. 1), indicating that serum deprivation
causes caspase-dependent apoptosis.

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Fig. 1.
S1P protects HUVECs from serum
deprivation-induced cell death. HUVECs (1 × 105
cells/well) were plated onto six-well plates in 1 ml of M199 containing
20% fetal bovine serum. The next day, the cells were switched to
serum-free M199 with or without 20% fetal bovine serum or with 0-5
µM S1P or 100 µM z-VAD-fmk. After
24 h, cell viability was determined by trypan blue exclusion. Data
represent mean ± S.D. (n = 4).
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NMA Suppresses the Inhibitory Effects of S1P on Serum-deprived
Apoptosis--
It has been shown that NO production from eNOS or
exogenous NO donor protects endothelial cells from apoptosis induced by TNF
(22) and LPS (21). To investigate the involvement of NO in the
protective effect of S1P, HUVECs were treated with S1P in the presence
of the specific inhibitor of NOS NMA, and cell viability was measured.
The protective effect of S1P was significantly reduced by the addition
of NMA (Fig. 2A). DNA
fragmentation typical of apoptosis was identified in serum-deprived
HUVECs (Fig. 2B). S1P prevented DNA fragmentation, and the
protective effect of S1P was blocked by the addition of NMA. These
results suggest that the cytoprotective effect of S1P on serum-deprived
apoptosis is mediated by NO production from HUVECs.

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Fig. 2.
NMA blocks S1P-mediated HUVECs survival.
HUVECs were incubated in serum-free medium containing S1P (5 µM) in the absence or presence of NMA. Cell viability was
measured by trypan blue after 24 h (A), and DNA
fragmentation was determined by isolation of DNA and agarose gel
electrophoresis after 18 h (B). Data represent
mean ± S.D. of three experiments. *, p < 0.01 versus treatment with S1P alone.
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Antisense Oligonucleotides of EDG-1 and -3 Suppress S1P-mediated
Survival and NO Production--
Exogenous S1P regulates cellular
responses such as angiogenesis (2), morphogenesis (2, 4), cell
migration (2, 3), and cell survival (28) through the activation of
EDG-1 and -3 receptors or through the activation of sphingosine kinase, which synthesizes endogenous formation of S1P. We next examined the
involvement of these signaling pathways in the S1P-mediated increase in
cell survival and NO production by treating the HUVECs with antisense
oligonucleotides for EDG-1 and -3 or the sphingosine kinase inhibitor
dihydrosphingosine. Pretreatment of HUVECs with the antisense, but not
sense, oligonucleotide to EDG-1 significantly suppressed S1P-mediated
cell survival and S1P-induced NO production (Fig.
3, A and B). In
contrast, pretreatment with antisense, but not sense, oligonucleotide
to EDG-3 only partially suppressed the effects of S1P (Fig. 3,
C and D). A combination of antisense oligonucleotides to EDG-1 and -3 suppressed the effects of S1P to about
90% (data not shown). Treatment with the sphingosine kinase inhibitor
dihydrosphingosine did not inhibit S1P-mediated cell survival or NO
production (data not shown). These results suggest that S1P requires
both EDG-1 and -3 for full activation of the cellular signal needed to
promote cell survival and NO production in HUVECs.

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Fig. 3.
Antisense oligonucleotides to EDG-1 and -3 block S1P-mediated cell survival and NO production. HUVECs were
pretreated with antisense or sense oligonucleotide (5 µM) to EDG-1 or -3 for 12 h. The medium was replaced
with serum-free fresh medium containing 5 µM S1P
and 5 µM oligonucleotide. After 24 h, cell viability
and NO production were measured. Antisense oligonucleotide of EDG-1
decreases HUVEC survival (A) and NO production
(B). Antisense oligonucleotide of EDG-3 decreases HUVEC
survival (C) and NO production (D). Data
represent mean ± S.D. of three experiments.
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S1P Inhibits Caspase Activity/Activation and Cytochrome c Release
in a NO-dependent Manner--
Since serum deprivation
induces apoptosis by activating the caspase signal cascade and
mitochondrial cytochrome c release (29), we examined the
effect of S1P on caspase activation/activity and mitochondrial
cytochrome c release in serum-deprived HUVECs. HUVECs were
treated with or without S1P in serum-free medium, and DEVDase
(caspase-3-like protease) activity was measured in the cytosol by a
colorimetric assay using the tetrapeptide substrate Ac-DEVD-pNA and a
PARP cleavage assay. DEVDase activity was significantly increased in
the serum-deprived HUVECs compared with that of the untreated
control cells (Fig. 4A).
Treatment with S1P or the caspase inhibitor z-VAD-fmk blocked the
increase in DEVDase activity. The effect of S1P was almost completely
reversed by NMA. NO produced by endothelial cells and interleukin-1
plus interferon
-stimulated hepatocytes inhibits DEVDase
activity by S-nitrosylation of the caspase enzymes. This
inhibition can be reversed by the reducing agent DTT, which removed the
NO from the active site cysteine (17, 22). Preincubation of the cytosol
with DTT partially reversed the suppressed DEVDase activity by S1P.
PARP, a well known biosubstrate of caspase-3, was cleaved to an 85-kDa
fragment by the cytosol from serum-deprived HUVECs, while cytosols
from z-VAD-fmk and S1P-treated cells did not cleave PARP (Fig.
4B). This effect of S1P was also reversed by NMA.

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Fig. 4.
NMA blocks the inhibitory effects of S1P on
caspase-3 activity/activation and mitochondrial cytochrome c
release in serum-deprived HUVECs. HUVECs were cultured in
serum-free medium with or without S1P (5 µM) in
the absence or presence of 100 µM z-VAD-fmk or 1 mM NMA. A, DEVDase activity was measured in
cytosol by chromogenic assay using synthetic substrate Ac-DEVD-pNA
after incubation of lysate with or without 20 mM DTT. *,
p < 0.01 versus without DTT. B,
in vitro PARP cleaving activity. 35S-Labeled
PARP was incubated for 40 min at 37 °C with the cytosolic S-100
fraction prepared from HUVECs. PARP cleavage was analyzed by SDS-PAGE.
C, mitochondrial cytochrome c release and
proteolytic activation of caspase-3. Cytosolic extract was obtained by
centrifugation at 100,000 × g for 1 h at 4 °C
after homogenization of cells in buffer A containing 250 mM
sucrose. Whole cell lysates were prepared by three freeze-thaw cycles.
Proteins (40 µg) were separated on 14% SDS-PAGE and transferred into
nitrocellulose membrane. Protein bands were visualized by Western blot
using antibodies for cytochrome c (Cyt c),
cytochrome c oxidase (Cyt c OX), and caspase-3.
D, relationship between mitochondrial cytochrome
c release and DEVDase activation. Cytosolic proteins were
resolved by 14% SDS-PAGE, transferred to nitrocellulose, and detected
by Western blot using antibodies for cytochrome c (Cyt
c), cytochrome c oxidase (Cyt c OX), and
caspase-3. DEVDase activity was measured in cytosol by chromogenic
assay using synthetic substrate Ac-DEVD-pNA. Data represent mean ± S.D. (n = 4).
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To further determine whether the release of mitochondrial cytochrome
c into cytosol and the activation of caspase-3 by serum deprivation are affected by treatment with S1P, we measured appearance of the cytosolic cytochrome c and the active fragment (p17)
of caspase-3 by Western blot analyses. As shown in Fig. 4C,
serum deprivation induced the release of cytochrome c to
cytosol and the cleavage of pro-caspase-3 (p32) into a 17-kDa fragment.
These events were suppressed by the addition of S1P or z-VAD-fmk. The S1P effects were reversed by NMA. However, cytochrome c
oxidase as an indicator of mitochondrial contamination was not detected in the cell lysates. Studies in several cell-free systems of apoptosis have revealed that cytosolic cytochrome c interacts with
Apaf-1 and procaspase-9 and forms an apoptosome in the presence of
dATP. This results in the activation of caspase-9, which leads to
activation of caspase-3 (18). Therefore, we next determined
whether mitochondrial cytochrome c release precedes
increases in DEVDase activity (Fig. 4D). Mitochondrial
cytochrome c release into cytosol and DEVDase activity
increased 12 and 16 h following serum withdrawal, respectively. Cytochrome c release was maximal at 16 h, and DEVDase
activity peaked at 20 h. This is consistent with the view that
cytochrome c release is upstream of caspase-3 activation in
the apoptotic signal cascade of serum-deprived HUVECs. These results
suggest that the antiapoptotic effect of S1P in serum-deprived
HUVECs includes the suppression of cytochrome c release
and the inhibition of caspase-3 activation/activity.
NO-mediated Protective Effect of S1P--
The antiapoptotic action
of NO is mediated at least in part through cGMP in hepatocytes (22),
PC12 cells (29), and cultured neuronal cells (31). To examine the role
of NO/cGMP pathway in S1P-mediated protection, the effect of S1P on
cell viability was examined in the presence of the specific inhibitor
of soluble guanylate cyclase ODQ (Fig.
5A). ODQ did not inhibit the
protective effect of S1P, while NMA did. Furthermore, treatment with an
exogenous NO donor SNAP (100 µM) protected cells from
serum-deprived cell death, and this protection would not be blocked by
ODQ. As expected a NO scavenger hemoglobin (400 µM as
heme concentration) prevent the protective effects of SNAP (Fig.
5B). In addition, the membrane-permeable cGMP analog
8-Br-cGMP did not inhibit serum-deprived apoptotic cell death. These
results indicate that the protective effect of S1P is
NO-dependent, but cGMP-independent.

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Fig. 5.
cGMP does not prevent serum-deprived
apoptosis. A, HUVECs were cultured in serum-free
medium containing 5 µM S1P in the absence or
presence of NMA (1 mM) or ODQ (40 µM).
B, cells were cultured in serum-free medium
containing SNAP (100 µM) with or without hemoglobin
(Hb, 400 µM as heme concentration), ODQ (40 µM) or 8-bromo-cGMP (8-Br-cGMP) (400 µM). After 24 h of incubation, cell viability was
determined by trypan blue exclusion. Data represent mean ± S.D.
(n = 4).
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S1P Increases NO Synthesis in HUVECs--
If S1P protects via NO
then it should be possible to measure increases in NO formation
following S1P exposure. The formation of NOx as stable oxidized
products of NO was measured in culture medium. S1P increased NO
production in a dose-dependent manner reaching about 2-fold
over controls at a concentration of 5 µM S1P (Fig.
6A). The accumulation of NOx
in the medium was detectable 4 h following S1P treatment
(Fig. 6B). To examine mechanism possible for S1P-induced NO
production, we examined the NOS activity in response to S1P. The NOS
activity was increased to about 1.5- and 2-fold at 4 and 12 h,
respectively, following treatment with S1P (Fig. 6C). The
increased NOS activity was suppressed by addition of the
Ca2+ chelator BAPTA-AM. Western blot analysis revealed no
significant difference of eNOS protein levels between 0 and 4 h
(Fig. 6D). These data suggest that increased NO production
by S1P is due to the activation of Ca2+-sensitive eNOS.

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Fig. 6.
S1P increases NO production by HUVECs
eNOS. A, HUVECs were incubated with different
concentration of S1P for 24 h. NOx (nitrite plus nitrate) was
measured in culture medium by chemiluminescence. B,
HUVECs were incubated with or without 5 µM S1P for
different time period. NOx accumulation was analyzed in culture
medium. C, HUVECs were incubated with 5 µM S1P for 5 h, and medium was replaced by
calcium-free HEPES buffer containing 1 µCi of
L-[3H]arginine plus 8 µM
L-arginine with or without 20 µM BAPTA-AM.
After 40-min incubation, [3H]citrulline produced by the
catalytic reaction of NOS was analyzed. D, protein level of
eNOS was analyzed by Western blot analysis. Protein level of actin was
also determined as protein-loading control. Data represent mean ± S.D. (n = 4).
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S1P Induces NO via the Gi Protein-PLC Signaling
Pathway--
To examine the signaling mechanism by which S1P
stimulates NO production in HUVECs, the effects of various inhibitors
on S1P signaling were evaluated (Fig. 7).
We (2, 6) and others (4) have observed that S1P activates intracellular
signaling cascade by a Gi-coupled
receptor-dependent manner. Treatment of HUVECs with
pertussis toxin, a blocker of Gi protein, significantly reduced the S1P-mediated NO production, suggesting that S1P mediated NO
production in endothelial cells through Gi-coupled
endothelial cell surface EDG receptors (Fig. 7A). To examine
the possible involvement of PLC and Ca2+ in the downstream
signaling pathway of Gi-coupled receptor(s), HUVECs were
cultured with S1P in the presence of the specific inhibitor of PLC
U73122 and the Ca2+ chelator BAPTA-AM. Treatment of HUVECs
with U73122 suppressed NOx accumulation in culture medium by
S1P, while U73343, which is a structurally similar derivative of
U73122, failed to exert a similar effect (data not shown). The calcium
chelator BAPTA-AM significantly blocked NOx accumulation by S1P.
Furthermore, these signal blockers reversed S1P-mediated protection of
HUVECs from serum-deprived apoptotic cell death (Fig. 7B).
Together, these results indicate that S1P-mediated increase in NO
production from HUVECs may be associated with increased
[Ca2+]i by EDG-1 and -3/Gi
protein-PLC signaling pathway.

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Fig. 7.
S1P-induced NO production is dependent on the
Gi protein-PLC signaling pathway. HUVECs in serum-free
M199 were pretreated with or without pertussis toxin (PTX,
100 ng/ml), U73122 (U, 5 µM), and BAPTA-AM
(BATA, 5 µM) for 1 h and then incubated
with or without S1P (5 µM) for additional 12 h for
NOx measurement or 24 h for cell viability. NMA (1 mM)
was cotreated with S1P. A, NOx was measured in the culture
medium by chemiluminescence. B, cell viability was
determined by trypan blue exclusion. Data represent mean ± S.D. (n = 4).
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DISCUSSION |
This study was undertaken to determine the molecular mechanism by
which S1P protects endothelial cells from apoptosis. We provide
evidence that S1P protects HUVECs from serum-deprived apoptosis by
increasing NO production. S1P increases NO production by
Ca2+-sensitive eNOS activation without changing eNOS
protein expression. NO production is associated with the inhibitory
effect of S1P on apoptotic cell death, DEVDase activity, and
mitochondrial cytochrome c release. Increases in cell
survival and NO production by S1P were significantly suppressed by
pretreatment with antisense oligonucleotides of EDG-1, and to a lesser
extent, by EDG-3 antisense, indicating that the cellular effects of S1P
involve the activation of EDG-1 and -3. The antiapoptotic effects of
S1P could not be inhibited by ODQ, indicating that the protective
effect does not involve cGMP production. This study also provides
evidence that the increase in NO production by S1P occurred through the
involvement of EDG-1 and -3, Gi protein, PLC, and
Ca2+. Thus, S1P increases eNOS activity via intracellular
Ca2+ mobilization, and the resulting increases in NO
protect endothelial cells from apoptosis by suppression of
apoptotic signaling cascade.
Apoptotic cell death by cytotoxic stimuli and serum or growth factor
withdrawal is induced by tightly controlled intracellular signaling
events, which require serial activation of caspase family proteases and
mitochondrial cytochrome c release. Caspase-deficient animals or cells are resistant to apoptosis induced by anti-Fas antibody (32) and DNA-damaging agents (33). Treatment with caspase
inhibitors reduces fulminant liver injury and animal mortality induced
by administration of anti-Fas antibody (34) or LPS (35). These studies
indicate that inhibition of caspase activation/activity is a potential
approach for the prevention of apoptosis. Our data show that serum
deprivation induced HUVEC apoptosis, as indicated by increased DNA
fragmentation, DEVDase activity, caspase-3 activation, and cytochrome
c release. Treatment with S1P and the caspase inhibitor z-VAD-fmk protected cells from serum-deprived apoptosis by suppressing these effects. These results raise the possibility that S1P is an
endogenous protectant against endothelial cell apoptosis through a
mechanism of suppressed cytochrome c release and
caspase-mediated signaling.
In this study, our major question was how S1P prevents serum-deprived
apoptotic cell death in HUVECs. We (21) and others (17) have previously
shown that NO protects endothelial cells from LPS- and TNF
-induced
apoptosis. These studies directed our attention toward NO as a
potential mediator of the S1P effect. Importantly, the present study
revealed that S1P increased NO production from HUVECs by enhancing
Ca2+-sensitive eNOS activity without significant increase
in the eNOS protein. eNOS is normally located at the cell membrane
within caveolae in an inactive state (36). Increases in intracellular Ca2+ promote calmodulin binding to the enzyme, leading to
transient eNOS activation and NO production. Other interaction and
modification can alter eNOS activity, including binding to caveolin-1,
which inhibits eNOS activity (37), and phosphorylation (38) and binding
of heat shock protein 90 (39) that enhance NO production. Our studies
suggest that the dominant action of S1P is to increase Ca2+
levels; however influence on these other regulatory processes cannot be excluded.
NO has been shown to regulate either induction of apoptosis in some
cells (40) or prevention of apoptosis in others (22, 29). High levels
of NO have been shown to be a cytotoxic through the inhibition of ATP
synthetic enzymes, including mitochondrial aconitase and electron
transfer complexes I and II (41) or through DNA damage (42). NO also
suppresses cell proliferation through the inhibition of ribonucleotide
reductase (43) and eukaryotic initiation factor-2
(44). NO can also
prevent apoptosis in several cell types, including hepatocytes (22,
45, 46), human B lymphocytes (47), PC12 cells (29), splenocytes (48), eosinophils (49), ovarian follicles (50), neuronal cells (31), MCF7
(51), Jurkat cells (52), and endothelial cells (17, 21). Several
mechanisms for the antiapoptotic effect of NO have been identified,
including the up-regulation of protective proteins such as heat shock
protein 70 (23), heme oxygenase (53), or Bcl-XL (54).
NO-mediated increases in cGMP levels account for the protective effects
in some cells (22, 29, 49, 50). In our studies, S1P did not increase
many of proteins listed above, as measured by Western blot analysis
(data not shown), suggesting that the antiapoptotic effect of S1P is
unlikely linked to the expression of these cytoprotective genes. We
also excluded cGMP as a mediator of the S1P-NO protective pathway using
the soluble guanylate cyclase inhibitor ODQ. Another mechanism involves
the directed inhibition of caspases by NO via
S-nitrosylation of either the inactive (55) or active forms
of the protease (17, 22). NO suppresses the proteolytic activation and
activity of multiple caspases in intact cells, including caspase-3 and
caspase-8. This can prevent the downstream amplification of the signal
by suppressing mitochondrial cytochrome c release or
activation of other downstream caspases. Our data show that S1P
suppressed mitochondrial cytochrome c release and caspase-3
activation/activity and that this suppression was reversed by NMA.
Also, the inhibition of DEVDase activity was partially reversed by
preincubating the cytosol with the reducing agent dithiothreitol,
suggesting that NO produced by S1P inhibited in part DEVDase activity
by redox-sensitive S-nitrosylation of the catalytic cysteine
residue. This is consistent with the observation that
S-nitrosylation of caspase-3 by NO inhibits TNF
-induced apoptosis in endothelial cells (17). It is likely that
S-nitrosylated caspases cannot proteolytically activate
other caspases or induce mitochondrial cytochrome c release.
NO can also activate other survival signals, including Akt/PKB (56),
which prevents caspase activation and mitochondrial cytochrome
c release by phosphorylation of Bad (57) and procaspase-9
(58) as well as activation of NF-
B (59, 60). Our studies did not
exclude a role for Akt. Others have shown that Akt activation can also
protect endothelial cells through increasing eNOS activity
(38).
We also addressed the question of how S1P increases NO production in
HUVECs. Endothelial cells produce measurable levels of NO from eNOS at
basal levels of [Ca2+]i (50-140 nmol/liter).
Activity of eNOS can be enhanced by receptor-dependent
stimuli such as bradykinin (12) and by receptor-independent stimuli
such as Ca2+ ionophore (62) and shear stress (63). In these
cases, an acute increase in [Ca2+]i has been
observed before the elevation in NO production. Recent studies have
shown that S1P induces a rapid increase in [Ca2+]i in endothelial cells, CHO, and HL-60,
mainly via the Gi-coupled receptors (4, 64, 65). HUVECs
express EDG-1 and -3, which transduce signals to increase
[Ca2+]i in response to <10 nM of S1P
(4). Our results show that S1P increased cell survival and NO
production in serum-deprived HUVECs at the concentration of
µM levels. We (2, 6) and others (1, 66, 67) have also
shown that µM levels of S1P regulate ERK activation, FAK
phosphorylation, and apoptosis. Antisense oligonucleotides of EDG-1
suppressed much of the S1P-mediated cell survival and NO production,
whereas EDG-3 antisense oligonucleotide partially inhibited the effects
of S1P (Fig. 3). Furthermore, HUVECs were cotreated with S1P and
the sphingosine kinase inhibitor dihydrosphingosine, and then the
S1P-mediated cell survival and NO production were measured.
Dihydrosphingosine did not inhibit the effects of S1P (data not shown),
suggesting that exogenous S1P does not act through intracellular
generation of S1P by activation of sphingosine kinase. Recently, it has
been shown that S1P treatment targets its receptor EDG-1 to
plasmalemmal caveolae and facilitates the activation of eNOS in COS
cells cotransfected with EDG-1 and eNOS (67), suggesting the
involvement of EDG-1 in NO production. Therefore, S1P-induced NO
production is most likely to be occurred by the S1P receptors EDG-1
and-3 in HUVECs. Binding of eNOS with caveolin-1 inhibits the eNOS
catalytic activity, and the inhibitory effect of caveolin-1 on the eNOS
activity can be reversed by Ca2+-calmodulin (10). This
evidence suggests that S1P may increase NO production by enhancing
[Ca2+]i through the activation of S1P
receptor-mediated signaling cascade in HUVECs. Indeed, our data reveal
that S1P-induced NO production in HUVECs was significantly reduced by
the treatment with the Gi protein inhibitor pertussis
toxin, indicating that S1P induced NO production in endothelial cells
through the Gi-coupled S1P receptors, EDG-1 and -3. Gi protein dissociates into G
i and 
subunits upon receptor activation, and the released 
subunit is
known to stimulate PLC (61). S1P-induced NO production was markedly suppressed by the specific inhibitor of PLC U73122, suggesting
that a PLC pathway may be downstream mediator of the Gi-coupled S1P receptors. PLC is known to activate
intracellular Ca2+ mobilization by generation of inositol
triphosphate. The calcium chelator BAPTA-AM almost completely blocked
NOx accumulation in culture media of HUVECs by S1P. These observations
suggest that the increased NO production from HUVECs by S1P may be
associated with the eNOS activation by the elevation of
[Ca2+]i through the EDG-1 and
-3/Gi-PLC pathway. However, the delayed, but sustained,
increase in eNOS activity measured in our cells suggest that the
up-regulation of eNOS activity is influenced not only by rapid
increases in [Ca2+]i levels, but also by other
mechanisms such as Ca2+-independent phosphorylation of eNOS
(30).