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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 93 No. 12 (June 15), 1999:
pp. 4293-4299
Induction and Suppression of Endothelial Cell Apoptosis by
Sphingolipids: A Possible In Vitro Model for Cell-Cell Interactions
Between Platelets and Endothelial Cells
By
Nobuo Hisano,
Yutaka Yatomi,
Kaneo Satoh,
Shigeo Akimoto,
Masako Mitsumata,
Masayuki A. Fujino, and
Yukio Ozaki
From the Department of Laboratory Medicine, First Department of
Pathology and First Department of Internal Medicine, Yamanashi Medical
University, Yamanashi, Japan.
 |
ABSTRACT |
Because sphingosine (Sph) is actively incorporated into platelets
and rapidly converted to sphingosine 1-phosphate (Sph-1-P), which is
then released extracellularly, it is important to study the effects of
Sph and Sph-1-P on endothelial cells from the viewpoint of
platelet-endothelial cell interaction. In this study, we found that
Sph, as well as ceramide, induces apoptosis in human umbilical vein
endothelial cells (HUVECs). In contrast, Sph-1-P acts as a HUVEC
survival factor; this bioactive lipid was shown to protect HUVECs from
apoptosis induced by the withdrawal of growth factors and to stimulate
HUVEC DNA synthesis. In metabolic studies, [3H]Sph,
incorporated into HUVECs, was converted to [3H]Cer and
further to [3H]sphingomyelin in a time-dependent manner,
whereas [3H]Sph-1-P formation from [3H]Sph
was weak and transient. These findings in HUVECs are very different
from those of platelets, which possess a highly active Sph kinase but
lack Sph-1-P lyase. As a result, platelets abundantly store Sph-1-P,
whereas HUVECs contain much less Sph-1-P. Finally, HUVECs, in contrast
to platelets, failed to release Sph-1-P extracellularly, indicating
that HUVECs themselves are not able to supply the survival factor Sph-1-P, but receive it from activated platelets. Our results suggest that platelets may maintain the integrity of endothelial cells
by incorporating Sph and releasing Sph-1-P.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
SIGNAL TRANSDUCTION pathways that use
glycerophospholipid metabolites have been very well
characterized.1-3 Recently, sphingolipids, another major
class of membrane lipids, have also emerged as signal-transducing
lipids.4-7 Functionally, a distinguishing characteristic of
the sphingolipids is their apparent participation in pro- or
anti-proliferative cell regulation pathways. For example, ceramide
(Cer)4,5,8 and sphingosine (Sph)7,9,10 are important regulatory participants in programmed cell death (apoptosis), whereas sphingosine 1-phosphate (Sph-1-P) induces mitogenesis and has
been implicated as a second messenger in cellular proliferation induced
by platelet-derived growth factor and serum.11 It has been
reported that the balance between the intracellular levels of Cer and
Sph-1-P and their regulatory effects on different family members of
mitogen-activated protein kinase may determine cell fate.12
To clarify the involvement of sphingolipids in hemostasis, thrombosis,
and vascular biology, we have studied the metabolism and functional
effects of Sph derivatives in human platelets. We found that, in
platelets, Sph-1-P is rapidly formed from Sph by Sph kinase, abundantly
stored intracellularly, and released into the extracellular environment
upon stimulation.13,14 Furthermore, exogenously added
Sph-1-P induces platelet aggregation,13 suggesting that
Sph-1-P acts as an autocrine platelet stimulator.
Because exogenous Sph is actively incorporated into platelets and
rapidly converted to Sph-1-P, which is then released extracellularly, it is important to study the effects of Sph and Sph-1-P on endothelial cells from the viewpoint of platelet-endothelial cell interaction. In
this study, we investigated the effects of sphingolipids, including Sph
and Sph-1-P, on the cell fate of human umbilical vein endothelial cells
(HUVECs). Furthermore, we also examined the metabolism of HUVEC
sphingolipids in detail.
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MATERIALS AND METHODS |
Materials.
The following materials were obtained from the indicated suppliers:
D-erythro-Sph, 12-O-tetradecanoylphorbol 13-acetate (TPA), lysophosphatidic acid (LPA), sphingomyelin, and sphingomyelinase (from
Staphylococcus aureus) (Sigma, St Louis, MO); Sph-1-P,
N,N-dimethylsphingosine (DMS), and C2-Cer (Biomol, Plymouth
Meeting, PA); thrombin (Mochida Pharmaceutical, Tokyo, Japan);
interleukin-1 (IL-1; Boehringer Mannheim Biochemica, Mannheim,
Germany); tumor necrosis factor- (TNF-; Genzyme, Cambridge, MA);
angiotensin II (Bachem California, Torrance, CA);
[Arg8]-vasopressin (AVP; Seikagaku Corp, Tokyo, Japan);
platelet-activating factor (PAF; Avanti Polar Lipids, Alabaster, AL);
staurosporine (Kyowa Medex, Tokyo, Japan); and
D-erythro-[3-3H]Sph (22.0 Ci/mmol),
[3-3H]C6-Cer (22.3 Ci/mmol), and
[methyl-3H]thymidine (20.0 Ci/mmol) (Du-Pont NEN, Boston, MA).
Cell preparation.
HUVECs were isolated from human umbilical cords with trypsin treatment,
plated onto 0.2% gelatin-coated dishes, and cultured in Dulbecco's
modified Eagle's medium (DMEM) with 20% fetal calf serum (FCS; ICN
Biomedicals, Aurora, OH), 10 ng/L of recombinant human basic fibroblast
growth factor (bFGF; Becton Dickinson Labware, Lincoln Park, NJ),
penicillin G (100 U/mL), and streptomycin sulfate (100 µg/mL) at
37°C under an atmosphere of 5% CO2 and 95% room air.
FCS and bFGF were removed to deprive HUVEC growth factors. Human
platelets were prepared as described previously.13
Fluorescence microscopy.
After the reactions indicated, cells (floating cells and adhesive
cells) were collected and washed twice with phosphate-buffered saline
(PBS). They were then fixed overnight in 1% glutaraldehyde (Nacalai,
Kyoto, Japan) and washed in PBS. Cells were stained with bisbenzimide
trihydrochloride (1 mmol/L in 30% glycerol/PBS; Hoechst 33258;
Calbiochem, San Diego, CA) in darkness for 10 minutes and visualized
using fluorescence microscopy. Apoptotic cells were identified by the
findings of condensation and fragmentation of chromatin. A total of 20 random microscope fields were examined under each of the experimental
conditions. The total number of cells and the number of apoptotic cells
were counted in each field.
Analysis of DNA fragmentation by electrophoresis.
After the reactions indicated, cells were harvested, washed,
resuspended in TTE buffer (1 mol/L Tris HCl, 0.5 mol/L EDTA, 10%
Triton X-100), and treated with 400 µg/mL of RNase A for 1 hour at
37°C. Then, 400 µg/mL of proteinase K was added and the incubation was continued for 2 hours at 37°C. DNA was extracted with phenol/chloroform/isoamyl alcohol, washed in ethanol, resuspended in TE buffer, and separated by electrophoresis in a 2% agarose gel.
Then, the DNA was stained with ethidium bromide. The gels were
photographed with UV transillumination.
Measurement of DNA synthesis in HUVECs.
HUVECs were cultured in 35-mm dishes until the cells were confluent and
quiescent in 10% FCS plus bFGF. Cells were washed and preincubated in
10% FCS for 20 minutes and then treated with Sph-1-P for 24 hours;
[3H]thymidine (2 µCi/mL) was present for the last 2 hours of the incubation. This time period was chosen because it gave
maximum incorporation of the radiolabel at 10 µmol/L Sph-1-P in
preliminary experiments (data not shown). After the incubation, HUVEC
morphology was checked; cells firmly adhered to dish and were of normal
size and shape. HUVECs were washed twice with Hanks' balanced salt solution and then with 5% trichloroacetic acid. The acid-insoluble material was then redissolved in 0.1 N sodium hydroxide, and an aliquot
was taken to measure the radioactivity levels by liquid scintillation counting.
Metabolism of [3H]Sph and
[3H]C6-Cer in HUVECs.
HUVECs were incubated with 1% FCS containing 1 µmol/L (0.2 µCi)
[3H]Sph or [3H]C6-Cer. At the
indicated time points, the reaction was terminated by the addition of 1 mL of ice-cold methanol, and lipids were extracted from the cell and
medium separately by the method of Bligh and Dyer15 and
then analyzed for [3H]Sph or
[3H]C6-Cer metabolism as described
previously.13 Portions of lipids obtained from the lower
chloroform phase were applied to silica gel high-performance thin layer
chromatography (TLC) plates (Merck, Darmstadt, Germany), and the plates
were then developed in butanol/acetic acid/water (3:1:1), followed by
autoradiography. Each autoradiogram shown is a typical one from at
least three experiments. When indicated, silica gel areas containing
radiolabeled sphingolipids were scraped off and counted by liquid
scintillation counting. The radioactivity counts were corrected by
recovery rates of the sphingolipids into the lower phase. Under our
conditions, recovery rates in the lower phase of Sph, Sph-1-P, Cer, and
sphingomyelin were 78%, 47%, 73%, and 71%, respectively.
Sphingomyelinase treatment of the extracted lipids.
The lower phase samples of the lipid extract were dried completely and
recovered in a solution of 0.1 mol/L Tris-HCl (pH 7.4), 0.01% Triton
X-100, and 40 mmol/L MgCl2, with sonication.
Sphingomyelinase (5 U/mL) was added to the reaction mixture (200 µL
in all). After 1 hour at 37°C, the reaction was terminated by the
addition of 800 µL of ice-cold chloroform/methanol/concentrated HCl
(100:200:1), and the lipids were extracted and the phases separated by
the method of Bligh and Dyer.15 The resultant lower
chloroform phase samples were analyzed for sphingomyelin degradation by
TLC developed in butanol/acetic acid/water (3:1:1), followed by autoradiography.
Quantitative measurement of Sph-1-P.
Sph-1-P was extracted from platelets and HUVECs and quantitated by
N-acylation with [3H]acetic anhydride into
[3H]C2-Cer-1-P
(N-[3H]acetylated Sph-1-P), as described
previously.16 The phospholipid amounts were also measured,
as described previously,16,17 to normalize the sample
amounts for Sph-1-P quantitation.
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RESULTS |
Induction of HUVEC apoptosis by sphingolipids.
Of sphingolipids, Cer, formed via the sphingomyelin cycle, has been
shown to be a lipid second messenger or biomodulator of stress-related
responses, including apoptosis, in a variety of systems.4,5,8,18 In contrast, the effects of Sph on cell fates seem to be cell-type specific. Sph shows strong mitogenic effects
in some cells,6,19,20 whereas the role of Sph in apoptosis
induction has been reported in others.7,9,10 To examine the
possible role(s) of sphingolipids in HUVEC fate, we first examined the
effects of various Sph derivatives on HUVEC apoptosis. In HUVECs
challenged with Sph, phase-contrast microscopy showed morphological
changes of apoptosis; when exposed to 20 µmol/L Sph for 4 hours,
HUVECs shrank and retracted from neighboring cells, and floating
apoptotic cells appeared in the culture medium (data not shown). When
HUVECs were stained with Hoechst 33258 and assessed by fluorescence
microscopy, cells with condensed chromatin or fragmented nuclei and
blebbing of the plasma membrane were clearly visualized
(Fig 1A). The induction of apoptosis by Sph
was confirmed by demonstrating DNA fragmentation through agarose gel
electrophoresis (Fig 1B). Sph-induced HUVEC apoptosis was a
concentration-dependent process (Fig 2).
Under identical conditions, DMS, a methylated derivative of
Sph,7 induced apoptosis in a stronger manner than Sph (Fig
2).
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| Fig 1.
Induction of HUVEC apoptosis by Sph. (A) Morphological
features of HUVEC apoptosis induced by Sph. HUVECs were treated without
(a and c) or with (b and d) 20 µmol/L Sph for 4 hours in the presence
(a and b) or absence (c and d) of growth factors. Apoptotic cells were
detected using Hoechst 33258 staining and visualized with fluorescence
microscopy. (B) DNA fragmentation in Sph-treated HUVECs.
HUVECs were treated without (lanes a and c) or with (lanes b and d) 20 µmol/L Sph for 4 hours in the presence (a and b) and absence (c and
d) of growth factors. Fragmented DNA was isolated and electrophoresed
on a 2% agarose gel. DNA was then visualized with ethidium bromide
staining. Lane m is a 200-bp DNA ladder.
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| Fig 2.
Dose-dependent induction of HUVEC apoptosis by Sph or
DMS. HUVECs were treated with various concentrations of Sph ( ,  )
or DMS ( ,  ) for 4 hours in the presence (, ) or absence
(, ) of growth factors. Apoptotic cells were detected using
Hoechst 33258 staining and visualized with fluorescence microscopy.
Results were expressed as the percentage of apoptotic cells (apoptotic
cells/total cells × 100).
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It is well known that growth factor deprivation of HUVEC induces
apoptosis with characteristic morphological and biochemical features.21,22 We confirmed this under our conditions (Figs 1 and 2). Furthermore, the percentage of apoptotic cells increased when
HUVECs were challenged with Sph or DMS in the absence of growth
factors, although the effects of these sphingolipids themselves became
less clear (Figs 2 and 3).
C2-Cer is a synthetic cell-permeable Cer and has been shown
to induce apoptosis in endothelial cells, which was confirmed under our
conditions (Fig 3). When the induction of
apoptosis was compared among sphingolipids, the order of potency was
DMS > Sph = C2-Cer; DMS induced statistically significant
apoptosis in the absence of growth factors as well as in their
presence, whereas Sph and C2-Cer did not (Fig 3). Under all
conditions, Sph-1-P did not induce apoptosis at all (data not shown).
Staurosporine is known to induce apoptosis possibly through its protein
kinase C inhibition. This inhibitor, used as a control in our
experiments, was found to induce marked HUVEC apoptosis (Fig 3).
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| Fig 3.
Induction of HUVEC apoptosis by various sphingolipids and
staurosporine. HUVECs were treated without (C) or with 20 µmol/L Sph,
DMS, or C2-Cer or 1 µmol/L staurosporine (Ssp) for 4 hours in the presence (left panel) or absence (right panel) of growth
factors. Apoptotic cells were detected using Hoechst 33258 staining and
visualized with fluorescence microscopy. Results were expressed as the
percentage of apoptotic cells (apoptotic cells/total cells × 100).
Columns and error bars represent the mean ± SD (n = 3).
*Statistically significant (t-test, P < .05) compared
with the control cells (without treatment).
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DNA synthesis and suppression of apoptosis by Sph-1-P in HUVECs.
It is established that, in some systems, Sph-1-P, formed from Sph by
Sph kinase-catalyzed phosphorylation, is involved in cell
survival.6,11,12 To investigate the possible role of Sph-1-P in HUVEC proliferation, we examined the effect of Sph-1-P on
DNA synthesis in quiescent HUVECs cultured in a 10% FCS medium. Sph-1-P was found to stimulate proliferation of quiescent HUVECs as
measured by [3H]thymidine incorporation
(Fig 4, left panel). A marked mitogenic effect was observed at 1 to 10 µmol/L. However, Sph-1-P at 20 µmol/L did not induce significant [3H]thymidine uptake,
possibly because DNA synthesis was complete before the
[3H]thymidine addition.
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| Fig 4.
Stimulation of DNA synthesis and suppression of apoptosis
by Sph-1-P. (Left panel) Incorporation of [3H]thymidine
into HUVEC DNA. HUVECs were treated with the indicated concentrations
of Sph-1-P for 22 hours in the presence of 10% FCS, followed by
incubation in the presence of [3H]thymidine for 2 hours.
DNA-associated radioactivity was measured using a liquid scintillation
counter. Columns and error bars represent the mean ± SD (n = 3).
(Right panel) Inhibition of apoptosis induced by HUVEC growth factor
deprivation. HUVECs were treated with indicated concentrations of
Sph-1-P for 4 hours in the absence of growth factors. Apoptotic cells
were detected using Hoechst 33258 staining and visualized with
fluorescence microscopy. The protection percentage from apoptosis was
calculated as ([apoptotic cells in the absence of growth factors]  [apoptotic cells in the presence of Sph-1-P])/ ([apoptotic cells in
the absence of growth factors] [apoptotic cells in the presence
of growth factors]) × 100.
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As described above, deprivation of growth factor induced apoptosis in
HUVECs. Sph-1-P was found to counteract the apoptosis in a
concentration-dependent manner (Fig 4, right panel). The percentage of
apoptosis suppression by 20 µmol/L Sph-1-P was as high as 75%.
Accordingly, Sph-1-P plays an important role in HUVEC survival, not
only by stimulation of cellular proliferation, but also by protection
from apoptosis.
Metabolism of sphingolipids in HUVECs.
To study metabolism of sphingolipids in HUVECs, we first examined the
metabolic fate of Sph in HUVECs by exogenous addition of
[3H]Sph into the cell culture. The added
[3H]Sph was incorporated into the HUVECs and rapidly
(within 20 minutes) converted to [3H]Sph-1-P
(Fig 5, upper panel). However, this
conversion was transient and the [3H]Sph-1-P band could
not be observed 4 hours after the label addition (Fig 5, upper panel),
possibly due to degradation by Sph-1-P lyase. This is completely
different from [3H]Sph-1-P formation in platelets labeled
with [3H]Sph; we previously reported that the high
percentage of the [3H]Sph originally added remains as
[3H]Sph-1-P, even after a long incubation, indicating the
stability of Sph-1-P in platelets, which lack Sph-1-P lyase
activity.14
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| Fig 5.
Metabolism of [3H]Sph in HUVECs. (Upper
panel) HUVECs were incubated with [3H]Sph for various
durations. Lipids were then extracted from cells or media and analyzed
for [3H]sphingolipids by TLC autoradiography. Locations
of standard lipids are indicated on the left. SM, sphingomyelin; O,
origin. (Lower panel) HUVECs and human platelets were incubated with
[3H]Sph for 4 hours and, after the autoradiography,
silica gel areas containing radiolabeled sphingolipids were scraped off
and counted using a liquid scintillation counter. Radioactivity was
expressed as a percentage of the value of [3H]Sph at time
0. Columns and error bars represent the mean ± SD (n = 3).
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In sharp contrast to the weak and transient formation of
[3H]Sph-1-P, the formation of
[3H]sphingomyelin, possibly through
[3H]Cer, was active and time-dependent (Fig 5, upper
panel). We also examined whether [3H]DMS could be formed,
because DMS was found to induce marked apoptosis in HUVECs (see above)
and because Sph N-methyltransferase activity exists in some tissues and
cells.23,24 However, DMS was not formed in HUVECs labeled
with [3H]Sph both in the resting (Fig 5) and activated
(see Fig 7) states.
Quantitative comparison was made between [3H]Sph
metabolism in HUVECs and platelets (Fig 5, lower panel). In HUVECs,
[3H]Sph was mainly converted to [3H]Cer by
N-acylation and further to [3H]sphingomyelin
through the action of sphingomyelin synthase.25 On the
other hand, in platelets, [3H]Sph-1-P is overwhelmingly
the main product of [3H]Sph, which can be best explained
by the fact that platelets possess very active Sph kinase and
practically no lyase activity for degradation of Sph-1-P
to a fatty aldehyde and ethanolamine phosphate.14
We previously developed an assay for quantification of Sph-1-P by its
N-acylation with [3H]acetic anhydride into
[3H]C2-Cer-1-P (N-acetylated
Sph-1-P).16 When the platelet extract containing Sph-1-P
was N-acylated with [3H]acetic anhydride, a highly
radioactive C2-Cer-1-P was formed (Fig 6, left lane). Under identical
conditions, only a trace amount of
[3H]C2-Cer-1-P was formed from the HUVEC
extract (Fig 6, right panel). These results confirm our previous
finding that platelets abundantly store Sph-1-P16,26 and
further indicate that HUVECs contain much less Sph-1-P.
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| Fig 6.
Detection of Sph-1-P in the extracts from human platelets
and HUVECs. The extracts from platelets (Plt) and HUVECs were
N-acylated with [3H]acetic anhydride into
[3H]C2-Cer-1-P to quantify Sph-1-P. The
extracts analyzed were those obtained from the cells containing 5 µmol phospholipid.
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As described above, [3H]Sph-1-P was transiently formed in
HUVECs incubated with [3H]Sph (Fig 5). This conversion
into [3H]Sph-1-P was not affected by IL-1, thrombin,
or TPA (Fig 7). Furthermore,
[3H]Sph-1-P was not detected in the medium under these
conditions (Fig 7), indicating that stimulation-dependent Sph-1-P
release does not occur in HUVECs. This is in contrast with platelets, which release Sph-1-P extracellularly, possibly in a protein kinase C-dependent manner.14
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| Fig 7.
Effects of various agents on [3H]Sph
metabolism in HUVECs. HUVECs were incubated with
[3H]Sph for 1 minute and then challenged without (C) or
with 100 U/mL of IL-1 (IL-1), 1 U/mL of thrombin (Thr), or 1 µmol/L TPA for 20 minutes in the presence of a 1% FCS medium. Lipids
were extracted from cells or media and analyzed for
[3H]Sph metabolism. Locations of standard lipids are
indicated on the left.
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We finally examined the metabolism of the radiolabeled
C6-Cer. One radioactive band was detected in the extract of
HUVECs incorporating [3H]C6-Cer
(Fig 8A). This band was found to be located
just below sphingomyelin on TLC and was specifically eliminated by
treatment with sphingomyelinase (data not shown). These results
indicate that [3H]C6-sphingomyelin is formed
from [3H]C6-Cer in HUVECs. This is consistent
with the fact that [3H]sphingomyelin was formed later
than [3H]Cer when HUVECs were incubated with
[3H]Sph (Fig 5).
[3H]C6-sphingomyelin production from
[3H]C6-Cer was not affected by treatment with
thrombin, TPA, LPA, angiotensin II, AVP, or PAF (Fig 8B).
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| Fig 8.
Metabolism of [3H]C6-Cer in
HUVECs. (A) HUVECs were incubated with
[3H]C6-Cer for various durations. Lipids were
then extracted from cells and analyzed for
[3H]C6-Cer metabolism. Locations of standard
lipids are indicated on the left. (B) HUVECs were incubated with
[3H]C6-Cer for 1 minute and then challenged
without (C) or with 1 U/mL of thrombin (Thr), 1 µmol/L TPA, 10 µmol/L LPA, 1 µmol/L angiotensin II (A II), 1 µmol/L AVP, or 1 µmol/L PAF for 20 minutes in the presence of a 1% FCS medium.
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DISCUSSION |
Regulation by sphingolipids of HUVEC apoptosis and growth.
It is now acknowledged that the branching pathways of sphingolipid
metabolism may determine whether a cell survives or dies. Whereas Cer
has been shown to be an important regulatory component of
apoptosis,4,5,8 Sph-1-P is reportedly involved in cellular survival as a signaling molecule.6,11,12 In contrast, Sph has been reported to mediate either apoptotic or mitogenic responses, depending on the cell type.6,7,9-11,19,20 In this study, we
first investigated the ability of various sphingolipids to induce
apoptosis in HUVECs. We found that not only Cer, but also Sph and its
methylated derivative DMS, induced HUVEC apoptosis. The effect of DMS
on apoptosis induction was more potent than that of Sph, which may be
explained by the fact that DMS, but not Sph, is metabolically stable
when incubated with intact cells.27
Both Sph and DMS have an inhibitory effect on protein kinase
C28,29; a pharmacological protein kinase C inhibitor,
staurosporine,30 also induced apoptosis. These observations
suggest that the induction of apoptosis by Sph and DMS may be related
to protein kinase C inhibition. However, recent reports have shown that
the mechanism(s) by which Sph and DMS induces apoptosis is complex and
cannot be explained only by protein kinase C inhibition.7
Induction of apoptosis by Sph is strongly correlated with inhibition of
mitogen-activated protein kinase, and this is unrelated to protein
kinase C inhibition.31 Furthermore, Sph downregulates the
expression of the antiapoptotic proteins, Bcl-2 and
Bcl-XL.32,33 Recently, inhibition of DNA primase by Sph/DMS was also reported.34 There is also the
possibility that a Sph/DMS-dependent protein kinase may be involved in
induction of apoptosis.35 The mechanism by which Sph (and
DMS) induces apoptosis remains to be determined.
It is well known that deprivation of growth factor induced apoptosis in
HUVECs.21,22 In this study, we demonstrated that Sph-1-P
protects HUVECs from apoptosis induced by withdrawal of growth factors.
Furthermore, Sph-1-P stimulates DNA synthesis in HUVECs, indicating
that this bioactive lipid acts as a HUVEC survival factor both by
protection from apoptosis and by stimulation of proliferation. The
results were as we expected, because Sph-1-P reportedly enhances cell
survival in some cells6,11,12 and because HUVECs express
the cell surface Sph-1-P receptor, Edg-1.36
Metabolism of sphingolipids in HUVECs.
Although the physiological roles of sphingolipids have been suggested,
current evidence for the involvement of sphingolipids in endothelial
cell function(s) consists largely of data on the cellular effects
caused by the exogenous addition of sphingolipids. Few studies have
reported the metabolic analysis of sphingolipids in endothelial cells.
We found that [3H]Sph, incorporated into HUVECs, is
converted to [3H]Cer and further to
[3H]sphingomyelin in a time-dependent manner, whereas
[3H]Sph-1-P formation from [3H]Sph is weak
and transient. The transient [3H]Sph-1-P formation can be
explained by its degradation to ethanolamine phosphate and fatty
aldehyde by Sph-1-P lyase, as is the case with most
cells.6,11,37 The findings for HUVECs are very different
from those of platelets. Platelets possess a highly active Sph kinase
but lack Sph-1-P lyase; hence, [3H]Sph-1-P formation from
[3H]Sph is very strong and long-lasting.14
Accordingly, it is not surprising that platelets abundantly store
Sph-1-P and that HUVECs contain much less Sph-1-P. Furthermore,
platelets release Sph-1-P extracellularly upon stimulation, possibly
through a mechanism dependent on protein kinase C,14 which
platelets abundantly express,38 whereas HUVECs do not. It
is unlikely that endothelial cells may be the source of plasma Sph-1-P;
these cells themselves are not able to supply the survival factor
Sph-1-P, but receive it from activated platelets.
Platelet-endothelial cell interaction.
Hemostasis, thrombosis, and atherosclerosis are considered an
integrated group of multicellular events, of which reactions between
endothelial cells and platelets are very important.39 Some
metabolic systems that regulate platelet reactivity are present in
endothelial cells: eicosanoids (cyclooxygenase metabolites), endothelium-derived relaxing factor/nitric oxide, and
ecto-nucleotidase(s).39
On the other hand, maintenance of vascular endothelium integrity can be
attributed to secreted products of platelets. During activation,
platelets release a variety of vasoactive substances, and their role in
the repair of damaged vascular intima is established.40,41 Although the repair of the thin endothelial lining of the intima requires the appearance of endothelial mitogens and the platelet would
be a logical source of such mitogen release, not much is known
regarding the endothelial mitogens or survival factors within platelets. It has been reported that platelets store the angiogenic factor platelet-derived endothelial cell growth factor, but this was
later confirmed to be identical to thymidine
phosphorylase.42,43 Furthermore, platelets were found to
store, but not to secrete, this angiogenic factor.43 Just
recently, vascular endothelial growth factor, a potent endothelial
growth factor and permeability mediator, was found to be released by
activated platelets.44,45
We previously showed that stimulated platelets release Sph-1-P in a
protein kinase C-dependent manner.14 Furthermore, Sph-1-P was found to be a normal constituent of human plasma and serum; serum
Sph-1-P levels were elevated compared with plasma, indicative of
Sph-1-P release during whole blood coagulation.26 In the present study, we found that Sph-1-P was a survival factor for endothelial cells. Accordingly, Sph-1-P should be added to the list of
endothelial survival factors released from platelets. Furthermore, we
believe that Sph-1-P is the most potent lipid mediator released from
platelets as an endothelial survival factor, at least of those
currently known.
In summary, Sph acts as an inducer of endothelial cell apoptosis,
whereas Sph-1-P acts as a survival factor. Platelets may maintain the
integrity of endothelial cells by incorporating Sph and releasing
Sph-1-P (Fig 9).
View larger version (18K):
[in this window]
[in a new window]
| Fig 9.
Platelet-endothelial cell interaction from the viewpoint
of sphingolipids. Platelets incorporate Sph and release Sph-1-P in a
protein kinase C (PKC)-dependent manner. Sph acts as an inducer of
endothelial cell apoptosis, whereas Sph-1-P acts as a survival factor.
This may be one of the mechanisms by which platelets maintain the
integrity of endothelial cells.
|
|
| |
ACKNOWLEDGMENT |
The authors thank Dr Yasuyuki Igarashi (Hokkaido University) for
helpful discussions and Drs Y. Fukada and K. Hoshi (Yamanashi Medical
University) for providing us with human umbilical cords.
| |
FOOTNOTES |
Submitted December 14, 1998; accepted February 17, 1999.
Supported by the Clinical Pathology Research Foundation of Japan,
Uehara Memorial Foundation, and Grant-in-Aid for Scientific Research
from the Ministry of Education, Science, and Culture, Japan.
The publication costs of this
article were defrayed in part by
page charge payment. This article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
Address reprint requests to Yutaka Yatomi, MD, PhD, Department of
Laboratory Medicine, Yamanashi Medical University, Nakakoma, Yamanashi
409-3898, Japan.
| |
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July 5, 2000;
97(14):
7859 - 7864.
[Abstract]
[Full Text]
[PDF]
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TOP
ABSTRACT
INTRODUCTION