From the Sidney Kimmel Cancer Center, Division of Vascular Biology and Angiogenesis, San Diego, California 92121
Received for publication, October 9, 2002, and in revised form, December 2, 2002
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The vascular endothelium acutely autoregulates
blood flow in vivo in part through unknown mechanosensing
mechanisms. Here, we report the discovery of a new acute
mechanotransduction pathway. Hemodynamic stressors from increased
vascular flow and pressure in situ rapidly and transiently
induce the activity of neutral sphingomyelinase but not that acid
sphingomyelinase in a time- and flow rate-dependent manner,
followed by the generation of ceramides. This acute mechanoactivation
occurs directly at the luminal endothelial cell surface primarily in
caveolae enriched in sphingomyelin and neutral sphingomyelinase, but
not acid sphingomyelinase. Scyphostatin, which specifically blocks
neutral but not acid sphingomyelinase, inhibits mechano-induced neutral
sphingomyelinase activity as well as downstream activation of
extracellular signal-regulated kinase 1 and 2 (ERK1 and ERK2) by
increased flow in situ. We postulate a novel physiological
function for neutral sphingomyelinase as a new mechanosensor initiating
the ERK cascade and possibly other mechanotransduction pathways.
Acute vasoregulation of blood flow through the mechanosensitivity
of vascular endothelial cells to hemodynamic stressors sets blood
vessel tone, maintains normal tissue homeostasis, and influences the
pathogenesis of vascular disease (1, 2). Increased hemodynamic forces
such as shear stress and pressure from fluid flowing over the luminal
surface of vascular endothelial cells lining blood vessels very rapidly
induce various cell surface signaling events, including protein
phosphorylation, the Ras/Raf/mitogen-activated protein
(MAP)1 kinase pathway (3),
and, most notably, endothelial nitric oxide synthase (eNOS) to generate
the critical compensatory vasodilator nitric oxide (NO) (4, 5).
Increasing fluid shear over cultured endothelial cell monolayers
stimulates heterotrimeric G protein-coupled signaling (6-8), MAP
kinases (9), NO production (5, 7), focal adhesion kinase (FAK), and ion
channels (10). This diversity of the mechanoresponse may be categorized
temporally (11). Acute responses, occurring in seconds to minutes, have
been detected in most cases both in vivo/situ and
in culture and include the activation of ion channels, eNOS,
plasmalemmal tyrosine kinases, G proteins and, minutes later, the
cytosolic MAP kinases (3, 4, 6-10). After 15 min to 1 h, FAK and
c-Jun N-terminal kinase (JNK) are activated, at least in culture
systems (12). Later, transcription factors will be activated to alter
gene expression (2, 13, 14) and, with time, to modulate endothelial
cell phenotype, cell adhesion, and even atherogenesis (2, 14). The
mechanisms mediating these effects, especially the key initiating mechanosensing molecule, remain largely unknown.
Caveolae are small, flask-shaped, plasmalemmal invaginations that are
found on the surface of many cell types. In the endothelium, caveolae
are thought to function primarily as vesicular carriers transporting
molecules into and across the cell (15). Our laboratory has discovered
that endothelial caveolae can also act as mechanosensing organelles
that respond to mechanical stressors (3, 4, 11, 16). Caveolae are
highly concentrated in tyrosine kinase activity, and increased vascular
flow and pressure can rapidly induce the tyrosine phosphorylation of
cell surface proteins located primarily in caveolae (3). Disassembly of
caveolae by the depletion of cholesterol disperses the molecular
constituents of this specialized compartment and prevents acute flow
activation of cell surface tyrosine kinases as well as downstream
cytosolic kinases such as ERK, but not JNK (3, 4, 17). Increased flow
also rapidly stimulates eNOS, which is concentrated at the endothelial
cell surface in caveolae (4). Many of the newly implicated cell surface
molecules (tyrosine kinases (11, 18), caveolin (3, 4, 11), eNOS (4),
and G-proteins (19)) exist in caveolae on the cytoplasmic side of the
plasma membrane (4, 20, 21) and may respond to an unidentified upstream mechanoregulator.
Caveolae may have a distinct lipid composition consisting of
cholesterol, sphingolipids (sphingomyelin and glycosphingolipids), and
phosphatidylinositols (20, 22). Depending on the cell type, caveolae
may contain up to 95% of total cell sphingomyelin. In addition,
ligand-induced sphingomyelin hydrolysis occurs within caveolin-rich
membranes (23, 24). The sphingomyelin pathway is initiated by
activation of sphingomyelinase (SMase), which hydrolyzes sphingomyelin
to ceramide, a second messenger stimulating multiple signaling cascades
(for review, see Refs. 25 and 26). Here, we report a novel
physiological function for neutral sphingomyelinase (N-SMase), namely
the discovery that N-SMase, an enzyme likely to be exposed externally
on the plasma membrane and thus directly to fluid stressors, appears to
initiate acute mechanotransduction, leading to activation of downstream
MAP kinases.
Materials--
Reagents and other supplies were obtained
from the following sources:
[N-methyl-14C]sphingomyelin (55 mCi/mmol) and Percoll from Amersham Biosciences; imidazole,
ceramide, DETAPAC, octyl- Rat Lung Perfusion--
As described previously (3, 4), the lung
vasculature of anesthetized Sprague-Dawley rats (Harlan Sprague-Dawley,
150-170 g) was first flushed via the pulmonary artery at
6-8 mm Hg for 5 min with mammalian Ringer's solution using a syringe
infusion pump and then perfused at pressure ranging from 6 to 20 mm Hg for 0-10 min at 37 °C.
Subfractionation to Isolate Luminal Endothelial Cell Plasma
Membrane and Caveolae--
After the perfusion described above, the
luminal cell plasma membranes and caveolae of the endothelium were
purified using the in situ silica coating procedure as
described in our past work (28, 29). Briefly, the luminal surface of
the endothelium of the rat lung vasculature was coated with an ice-cold
colloidal silica solution in situ. After tissue
homogenization, the silica-coated endothelial cell plasma membrane (P)
was isolated from the tissue homogenate (H) by density centrifugation.
Caveolae were separated from P by homogenization in the presence or
absence of 1% Triton X-100 at 4 °C and then isolated by flotation
in sucrose gradient (V and V' fraction, respectively).
Isolation of Plasma Membrane and Caveolin-enriched Membrane
Fractions from Cultured Cells--
Bovine lung microvascular
endothelial cells (BLMVECs) were grown and used to isolate a
plasmalemmal (PM) and caveolin-rich membrane fractions (AC) as
described (29-31). Briefly, confluent BLMVEC cells (ten 150 mm dishes)
were washed, scraped and homogenized before centrifugation (1000 × g) for 10 min at 4 °C. The post nuclear supernatant
(PNS) was overlaid on 30% Percoll before centrifugation at 84,000 × g, for 1 h at 4 °C. A membranous band visible
about two-thirds from the bottom of the tube was collected (PM). AC was
separated from PM by sonication and Optiprep density centrifugation. AC
is enriched in caveolin but also contains lipid rafts, nuclear, Golgi,
and possibly other membranes (31).
Immuno-affinity Isolation of Caveolae--
Magnetic
immunoisolations were performed as described (31). Briefly, M450
Dynal beads conjugated to the caveolin monoclonal antibody (clone 2234)
(2 × 107 M450 beads and 25 µg of IgG) were
incubated for 1 h at 4 °C with the starting membrane fraction
and then washed and magnetically separated to isolate two fractions,
i.e. material bound to the beads and unbound material.
Assay for N- and A-SMase Activity--
The activity of N- and
A-SMase was estimated by the method of Wiegmann et al. (32).
Briefly, selected fractions (H, P, V or V', T or S, and P-V or P-V'; 10 µg protein) were incubated at 37 °C for 1 h in 20 mM HEPES buffer (pH 7.4) containing 1 mM MgCl2 and 3.4 µM
[N-methyl-14C]sphingomyelin (55 mCi/mmol). The reaction was terminated by adding 800 µl of
CHCl3/CH3OH (1:1; v/v) and 200 µl of
H2O. [14C]Phosphorylcholine released from
[N-methyl-14C]sphingomyelin
in the aqueous phase was collected and counted by liquid scintillation
counting. To measure the activity of A-SMase, perfused rat lungs were
homogenized in 20 mM Hepes (pH 7.4) containing 1 mM EDTA and 0.05% Nonidet P-40, and the subcellular
fractions (10 µg protein) were incubated at 37 °C for 1 h
with 250 mM sodium acetate (pH 5.0) containing 1 mM EDTA and 3.4 µM
[N-methyl-14C]sphingomyelin.
[14C]Phosphorylcholine was collected and measured as
described above.
Lipid Assay--
Ceramide was measured by diacylglycerol kinase
labeling (22, 33). Rat lung P (100 µg) was extracted with
CHCl3/CH3OH/1 N HCl (100:100:1;
v/v), and lipids in organic phase were collected and dried under
N2. The reaction was initiated by mixing the lipid film
with reaction buffer containing 50 mM imidazole (pH 6.5), 50 mM NaCl, 12.5 mM MgCl2, 1 mM EDTA, 5 mM cardiolipin, and 1.5% octyl- Protein Assays--
Protein concentrations were determined by
Micro BCA (Pierce) and DC (Bio-Rad) protein assays accordingly to each
manufacturer's instructions using bovine serum albumin as a standard.
Western analysis was performed as in past work (3).
Mechanoactivation of N-SMase but Not A-SMase to Generate Ceramide
at the Endothelial Cell Surface--
To determine whether elevated
hemodynamic stressors in situ could activate SMase activity,
we measured the activity of N- and A-SMase in rat lungs under elevated
pulmonary artery pressure. The blood was flushed at 37 °C from the
lung vasculature at pulmonary artery pressures of 6-8 mm Hg for 5 min.
Then the vascular pressure was either maintained at 6-8 mm Hg (basal
flow) or elevated to 14-16 mm Hg, which more than doubles the flow
rate (high flow). The lungs were subfractionated in order to measure
the activity of both N- and A-SMase in the total lung H and in the
isolated P. P relative to H is enriched 15-100-fold in endothelial
cell surface markers (i.e. caveolin, 5'-nucleotidase
(5'-NT), and angiotensin converting enzyme) while being markedly
depleted of proteins localized elsewhere in the cell or tissue
(i.e. fibroblast surface antigen,
We next focused on the mechanoactivation of N-SMase at the endothelial
cell surface in greater detail. Fig.
2A shows that the increase in
the specific activity of N-SMase in P began to occur within 30 s
of elevated perfusion pressure/flow. The activity reached a maximum of
2.2-fold over baseline at 2 min (1.53 + 0.1 nmol/mg/h
(n = 4) versus 0.680 + 0.017 nmol/mg/h
(n = 4)). The N-SMase activity began to decrease after
3 min and had returned to baseline levels by 10 min. The N-SMase
activity appeared constant in controls subjected only to the baseline
pressure/flow. We also tested whether this transient mechanoresponse
could be repetitively induced. Lungs perfused at high pressures for 2 min and then at baseline pressure for 3 min followed by a second high
pressure/flow rate perfusion for 2 min showed a second rapid increase
in N-SMase activity in the P (data not shown). Thus, the activity of N-
but not A-SMase was acutely responsive to, and rapidly activated by, increased hemodynamic stressor in situ. This induction was
transient but not refractory after only 3 min.
Because SMase catalyzes the hydrolysis of sphingomyelin to produce
ceramide, the activation of this enzyme should result in the transient
accumulation of the product in the membranes. We measured ceramide
levels in P under similar perfusion conditions and found that ceramide
levels closely followed N-SMase activity. Just 1 min of vascular
perfusion at the elevated pressure/flow rate increased ceramide level
detected in P by 30% over the control (Fig. 2B). This
response reached a maximum of 80% after 3 min of perfusion, maintained
a plateau through 5 min, and decreased significantly by 10 min. In the
controls where the flow rate/pressure was kept constant over the same
time, the ceramide levels did not change.
Next, we studied the sensitivity of N-SMase and ceramide formation to
different flow rates and vascular pressures. Because maximal activation
was observed after 2 min stimulation (Fig. 2A), we perfused
the vasculature for 2 min at pressures varying from 6 to 18 mm Hg,
which produced flow rates in this system varying from 4 to 14 ml/min,
respectively. As shown in Fig. 2C, even a small increase to
8 mm Hg (6 ml/min) from a baseline of 6 mm Hg increased N-SMase
activity in P by 30%. A maximum of nearly 2-fold over baseline
activity was reached at 12-14 mm Hg (10-12 ml/min). Under the same
conditions, ceramide levels in P also increased progressively to an
apparent maximum of 1.7-fold over baseline at 12-14 mm Hg (10-12
ml/min) (Fig. 2D). These cumulative results demonstrated
that the acute increase in vascular pressure/flow rapidly stimulates
N-SMase activity to induce formation and accumulation of ceramide in
the endothelial cell plasma membrane in a time- and flow
rate-dependent manner.
N-SMase and Its Activity in Caveolae--
The mechanosensitive
SMase existing at the luminal endothelial cell surface may be
associated with caveolae, because endothelial caveolae are primary
sites for rapid mechano-induced tyrosine phosphorylation of proteins
(3) and may be mechanosensing organelles (3, 4, 11, 16) containing many
signaling molecules including sphingomyelin and nonreceptor tyrosine
kinases (20, 22). Enzyme activities were measured in caveolae isolated
from P in the presence or absence of Triton X-100 (V and V' fraction,
respectively). With detergent, the major activity was detected in the
Triton-soluble fractions at levels nearly 5-fold enriched over P (Fig.
3A). This is consistent with
the known detergent solubility of N-SMase as well as sphingomyelin
extractability (34). Without detergent, substantial N-SMase activity
was found in caveolae with nearly 6-fold enrichment over P (Fig.
3B). By contrast, the major activity of A-SMase was found in
lung homogenates with >10-fold depletion in P and even more so in
caveolae (Fig. 3C). As shown in Fig. 3D and in
past studies (4, 20, 28, 31), the isolated caveolae were enriched
>15-fold in caveolar markers, such as caveolin, while being markedly
depleted (>15-fold) in noncaveolar markers such as
To determine the protein localization of N- and A-SMase, we examined
their distribution in plasma membranes and a caveolae fraction isolated
from BLMVEC using our immunoisolation procedure (31). Western analysis
detected significantly more N-SMase in AC over the crude PM, whereas
A-SMase resided primarily in the PNS (Fig.
4). Because AC contains nuclear and other
membrane microdomains in addition to cell surface lipid rafts and
caveolae (31, 36), we immunoisolated caveolae from AC using
anti-caveolin-1 antibodies (31). The bound fraction of caveolae
contained readily detectable N-SMase not found in the unbound fraction.
We used bovine cells because the N-SMase antibodies were generated
against a peptide derived from bovine enzyme (27) and did not
cross-react with N-SMase from rat tissues (data not shown). Western
analysis of H and P from BLMVEC detected a single immunoreactive band
of a 95-100-kDa protein, which was not immunoreactive to the preimmune serum (data not shown). Western analysis using a commercially available
A-SMase antibody showed enrichment in PNS but not in PM or AC (Fig. 4).
Thus, N-SMase resides primarily in caveolae.
Increased Pressure/Flow Stimulates Caveolar N-SMase--
To
determine whether increased pressure/flow stimulates N-SMase located in
caveolae, we subfractionated rat lungs subjected to baseline
versus high pressure/flow conditions before measuring N-SMase activity. Consistent with the experiments described above (Fig.
2A), the N-SMase activity in P nearly doubled when the
pressure/flow rate was elevated. This effect was even more pronounced
in V' (Fig. 5), with a 2.6-fold increase
over baseline conditions. With increased pressure/flow, the
N-Smase-specific activity in caveolae was 6.4-fold higher than that in
P fraction. Under high flow conditions, a modest but significant (56%
over the control) increase in the N-SMase activity in plasma membranes
stripped of caveolae (P-V') was observed. This effect can be explained
by some residual caveolae left attached to P-V' as detected by Western
analysis for caveolin-1 (Fig. 3D). Because the caveolar
membrane constitutes about 50% of the lung luminal endothelial plasma
membrane and assuming that V' is indeed representative of those
caveolae, then >85% of the baseline as well as mechano-induced
N-SMase activity at the plasma membrane occurs in the caveolae. These
results cumulatively indicate that the N-SMase activity associated with
caveolae is sensitive to mechanical shear forces and can be rapidly
activated by increased vascular pressure/flow in situ.
N-SMase Activity is Required for MAP Kinase
Mechanoactivation--
One of the best described signaling events
caused by mechanical stressors is activation of cytosolic MAP kinases,
i.e. ERK1 and 2 (3, 9). To assess whether N-SMase activity
is upstream and required for MAP kinase mechanoactivation, we used
scyphostatin, a specific N-SMase inhibitor (27, 37). Using a monoclonal antibody specific for activated ERK1 and 2, we found that (as in our
past work; see Ref. 3) high pressure/flow dramatically activate both
kinases (Fig. 6). Preexposure to
scyphostatin (50 µM) totally blocks this MAP kinase to
mechanoactivation. We also confirmed past work (27, 37) in in
vitro experiments, showing that scyphostatin, at 50 µM, inhibited N-SMase activity in P by 75% ± 2, whereas
A-SMase activity was not affected (0% ± 5). We also found that
scyphostatin does not in itself inhibit kinase activity as assessed
using in vitro assays (Ref. 20, and data not shown). Because
our in situ system does not discriminate between pressure
and shear effects, we subjected bovine aortic endothelial cell
monolayers to increased shear stress and found scyphostatin inhibited
shear stress-activation of MAP kinases (data not shown). Thus,
this inhibition of N-SMase prevents the downstream mechanoactivation of
MAP kinase signaling pathways, further supporting the key role of
N-SMase and ceramides as initiators of mechanotransduction.
Sphingomyelin is a structural lipid component of the plasma
membrane, and its metabolites play important roles in signal
transduction. The sphingomyelin pathway is initiated by the activation
of SMase by cytokines and cellular stressors such as UV and ionizing
radiation and heat shock, leading to the generation of ceramide, which
acts as a second messenger in activating a variety of cellular
functions (26, 38, 39). It is most likely that different isoenzymes, localized to different subcellular compartments, contribute to sphingomyelin turnover and sphingolipid signaling (26). Here, we report
the discovery that N-SMase functional activity responds to important
cardiovascular mechanical stressors, specifically from elevated
intravascular pressure and fluid flow shearing. These hemodynamic
stressors rapidly (within less then a minute) but transiently induced
N-SMase, but not A-SMase, activity at the luminal endothelial cell
surface to generate a transient increase in membrane ceramides.
These stressors represent a novel physiological pathway for N-SMase
stimulation. Although hemodynamic activation of N-SMase could occur
generally over the plasma membrane, it appears to be concentrated and
more greatly stimulated in the caveolae. It remains to be seen whether
some N-SMase resides at or near the neck region of caveolae for more
direct exposure to fluid shear and/or whether more than one pool of
N-SMase can be activated to a different extent. The availability of the
substrate, sphingomyelin (20, 22, 40), may also contribute to the high activity and rapid stimulation observed in caveolae.
There is no consensus yet on the number, localization, and function of
N-SMase isoforms. To date, several N-SMase isoforms have been cloned
(41-45) or purified (34, 46-50) from various mammalian tissues, and
these isoforms may exhibit distinct tissue and intracellular
distribution (43, 51, 52). Ligand-induced sphingomyelin hydrolysis and
the generation of ceramide within caveolin-rich fractions prepared by a
nondetergent method was reported previously in fibroblasts (23) and
PC12 cells (24). More recently, two independent groups have reported
N-SMase activity in low buoyant density, detergent-resistant,
caveolin-rich membrane fractions (35, 53). These N-SMase activities,
however, were Triton X-100 insoluble, whereas the mechano-induced
N-SMase activity detected in our rat lung endothelial caveolae isolates
was Triton soluble (detected in V' and T but not in V; see Fig.
3A), suggesting that the N-SMase activities in the two
systems correspond to different forms. Furthermore, whether the
previously noted SMase activities reside in caveolae remains to be
established because of the heterogeneity of the isolated membrane
fractions (35, 53).
The detection of distinct isoforms may be due to differences in cell
type (cultured murine endothelial cells (35), human fibroblasts (53) or
rat lung endothelial cells in situ (this study)) and
isolation methods used in the present and past (35, 53) studies. Note
that similarly prepared caveolin-enriched membrane fractions from whole
cell lysates (35, 53) have been shown to contain membrane microdomains
from plasmalemmal intercellular junction complexes and various
intracellular organelles (Golgi, nucleus) in addition to caveolae,
caveolin-coated vesicles, and lipid rafts rich in GPI-anchored proteins
(28, 31, 54). Here, we used the silica-coating procedure in rat lungs
in situ as well as the immunoisolation of caveolae from
caveolin-enriched bovine endothelial cell membranes to obtain
highly pure preparations of plasma membrane caveolae essentially free
of intracellular membranes. Although various isoforms of N-SMase may
exist in caveolae or lipid rafts, the caveolar isoform identified here
by using specific antibodies corresponds to the isoform cloned
by Bernardo et al. (27), which may be the Triton-soluble
isoenzyme that responds to mechanostimulation in the rat lung.
In this report, we describe the localization of N-SMase activity in rat
lung endothelial cells to caveolae, which are putative "signaling
centers" and "mechanosensing organelles" (16). We also
demonstrate by Western analysis the presence of one N-SMase isoform
(27) in caveolae isolated from bovine endothelial cells. Sphingomyelin,
the major sphingolipid of mammalian plasma membrane, is localized to
the outer leaflet of lipid bilayer, which is oriented externally on the
plasma membrane (55-57). Interestingly, the N-SMase isoform described
by Veldman et al. (53) can be inhibited by interaction with
a peptide corresponding to the scaffolding domain of caveolin-1,
suggesting an intracellular orientation, contrary to past reports (55,
58). N-SMase also localizes to the plasma membrane and likely exhibits
extracellular orientation (55, 58). Thus, the enzyme might be directly
exposed to hemodynamic stressors, including fluid shear and vascular
pressure, and serve as a mechanosensor on the luminal endothelial cell
surface to initiate signal transduction into the cell. If N-SMase is a
cell surface mechanosensor acutely initiating the transduction of
hemodynamic forces through its direct exposure to these forces
that increase activity, then the product of its increased
activity, ceramide, will stimulate diverse downstream signaling
pathways. Here, we show that inhibition of the N-SMase activity by
scyphostatin leads to the inhibition of downstream cytosolic
elements of mechanosignaling, namely ERK1 and ERK2 kinases.
The elucidation of upstream and downstream molecules and the mechanisms
of their involvement in endothelial mechanoresponsiveness is currently
underway in our laboratory. Ceramide can activate several intracellular
signaling molecules, including JNK (59) through unknown mechanisms and
MAP kinase (60) through the interaction of ceramide-activated protein
kinase with Raf (61). Ceramide can also regulate the activity of ion
channels (62, 63), NRTK and Ras (64), eNOS (65), and, ultimately,
transcription factors, such as NF Caveolae may play a central role in acute mechanotransduction, acting
as mechanosensing organelles by forming a distinct microdomain or
compartment that concentrates N-SMase and its substrate, sphingomyelin, as well as key downstream effectors such as eNOS, NRTK, and PI-3-kinase (20), which respond to the localized generation of ceramide. It appears
likely that this specialized compartment is critical for the efficient
propagation into the cell of one or more signals induced mechanically.
N-SMase and caveolae may be fruitful targets for future investigations
for many reasons, including the importance of endothelial dysfunction
and disrupted vasoregulation in the pathogenesis of vascular disease.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-D-glucopyranoside,
sphingomyelinase (Staphylococcus aureus), glycerophosphate,
anti-diphosphorylated ERK1 and 2 monoclonal antibody, and anti-pan ERK
antibody from Sigma; cardiolipin from Avanti Polar Lipids
(Alabaster, AL); diacylglycerol kinase from Calbiochem (San
Diego, CA); [
-32P]ATP (3000 Ci/mmol) from PerkinElmer
Life Sciences; the bicinchoninic acid (BCA) protein assay kit
from Pierce; anti-acid sphingomyelinase (A-SMase) antibodies from Santa
Cruz Biotechnology; anti-caveolin-1 monoclonal antibody (2234) from BD
Biosciences; M-450 Dynabeads from Dynal (New Hyde Park, NY); and
Optiprep from Invitrogen. Preimmune serum and serum raised
against bovine N-SMase were generously provided by Dr. Martin Kroenke
(Cologne University, Germany) (27). Scyphostatin was a generous gift
from Dr. Takeshi Ogita (Sankyo Co., Tokyo, Japan).
-D-glucoside, 0.2 mM DETAPAC, 2 mM dithiothreitol, 4 µg of diacylglycerol kinase, and 1 mM [
-32P]ATP (3000 Ci/mmol). After
incubation at 25 °C for 30 min, the reaction was terminated by
CHCl3/CH3OH/1 N HCl (100:100:1;
v/v). Ceramide-1-phosphate was resolved by thin layer chromatography using CHCl3/CH3OH/acetic acid at the ratio
65:15:5 (v/v). The lipid was identified by
-32P-labeled
ceramide standard and quantified by liquid scintillation counting.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-COP, ERK1/2) (3, 4,
20, 29) (Fig 3D). Our previous studies using this perfusion
model and isolation system detected rapid activation of eNOS and
tyrosine kinases at the luminal endothelial cell plasma membrane
(3, 4). Here, N-SMase activity in P increased rapidly with greater
vascular pressure/flow rate (Fig. 1).
This response was not detected in H, consistent with the small proportion of luminal endothelial cell plasma membranes in the tissue.
A-SMase activity was detected, but did not change in either fraction
over the 5 min of increased vascular pressure/flow.
View larger version (18K):
[in a new window]
Fig. 1.
Acute mechanoactivation of N- but not A-SMase
in endothelial cell plasma membranes. Rat lung vasculature was
perfused in situ first for 5 min at 6-8 mm Hg and then at a
higher pressure of 14-15 mm Hg for the time indicated. The activity of
either N-SMase (filled symbols) or A-SMase (open
symbols) in the rat lung homogenates (H,
squares) or the isolated luminal endothelial cell plasma
membranes (P, circles) was measured as described
under "Experimental Procedures." The percentage of the enzyme
activity compared with time zero is presented. Data are the average of
two experiments (n = 2).
View larger version (24K):
[in a new window]
Fig. 2.
Mechanoactivation of N-SMase and subsequent
ceramide formation. A and B, rat lung vasculature
perfusion was maintained either under control conditions (6-8 mm Hg,
open circles) or elevated to high pressure/flow conditions
(14-15 mmHg, filled circles) for the time indicated.
C and D, the perfusions were continued for 2 min
at pressures ranging from 6-18 mm Hg to give the indicated flow rates.
The luminal endothelial plasma membranes were isolated and used to
measure N-SMase activity (panels A and
C) and ceramide levels (panels B and
D). Data are expressed as the mean ± S.D.
(n = 4).
-actin, 5'-NT,
and
-COP. These results indicated that the luminal endothelial cell
plasma membranes and their caveolae are enriched in functional N- but
not A-SMase and that this N-SMase is at least partially
detergent-soluble. These results are consistent with past data showing
A-SMase to be an intracellular enzyme primarily localized to the
cytoplasm, lysosomes, and endosomes (26). Although one report found
A-SMase activity in caveolin-rich membrane subfractions (35), a more
detailed examination of these fractions by Western analysis and
immunoisolation with caveolin antibodies demonstrated that these
isolates contain not only caveolae but also lipid rafts as well as
possible microdomains of endosomal, Golgi, nuclear, and possibly other
cell membranes (28, 31, 36)
View larger version (18K):
[in a new window]
Fig. 3.
Enrichment of N- but not A-SMase in the
endothelial luminal surface plasma membranes and their caveolae.
The activity of N-SMase (panels A and
B) or A-SMase (panel C) was measured in 10 µg
of rat lung homogenate (H), purified silica-coated luminal
endothelial cell plasma membranes (P), caveolae isolated
either in the presence (V in panels A and
C) or absence (V' in panel B) of
Triton X-100, the Triton-soluble phase (T), the 40% sucrose
fraction (S, in the absence of detergent), or resedimented
silica-coated plasma membranes stripped of caveolae (P-V in
panels A and C or P-V' in panel B).
Data are expressed as the mean ± S.D. (n = 3)
(panel B) and the mean from two experiments
(panels A and C). D,
Western analysis of the indicated subfractions using antibodies
specific for caveolin, 5'-NT, and -actin.
View larger version (34K):
[in a new window]
Fig. 4.
N-SMase but not A-SMase in immuno-isolated
caveolae. BLMVEC homogenates were fractionated on a Percoll
gradient to isolate the PM from the post nuclear supernatant
(PNS) followed by sonication of PM and sucrose gradient
centrifugation to isolate AC. AC was subsequently subjected to
immuno-affinity isolation using caveolin-1 antibody-conjugated
magnetic beads to separate caveolae bound to the beads (B)
from noncaveolar unbound material (U) in the supernatant
(31). Western analysis was performed on each fraction using antibodies
to the indicated proteins.
View larger version (15K):
[in a new window]
Fig. 5.
Acute mechanoactivation of N-SMase in
caveolae. After perfusion first at 6-8 mm Hg for 5 min and then
either at 6-8 mm Hg (open bars) or 14 mm Hg (hatched
bars) for 2 min, the subcellular fractions (H, P, V', S,
P-V') were obtained in the absence of detergent. Equal
amounts of each subcellular fraction (10 µg of protein) were measured
for the activity of N-SMase. Data are expressed as the mean ± S.D. (n = 3).
View larger version (24K):
[in a new window]
Fig. 6.
Scyphostatin inhibits mechano-induced
activation of MAP kinase. Rat lung vasculature was perfused under
basal flow (6 mm Hg) in the presence or absence of 50 µM
scyphostatin for 5 min before either maintaining baseline flow
(BF) or increasing the pressure/flow rate to 14 mm Hg
(HF) for 2 min. Lung homogenates were subjected to Western
analysis with anti diphospho-ERK1 and 2 (for activated ERK) and
anti-panERK antibodies (for load control). This is representative of
three experiments.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
B (66). Interestingly, all of these
downstream targets of ceramide also respond to hemodynamic stressors.
Increased flow, pressure, and/or shear can rapidly activate eNOS (4),
MAP kinase (3, 9), K+ channels (10), and, later, JNK (12).
Increased vascular pressure and flow in situ rapidly
activates both plasmalemmal tyrosine kinases, causing protein
phosphorylation primarily in caveolae and the Ras/Raf/MAP kinase
pathway with rapid Raf translocation to caveolae (3). Disassembly of
caveolae with dispersion of their molecular constituents over the
membrane surface prevents both rapid flow-induced protein
phosphorylation and MAP kinase activation (3). Lastly, eNOS plays a
very important role modulating vascular tone and maintaining systemic
blood pressure, vascular remodeling, and angiogenesis. We have shown in
the same system used in this study that eNOS resides quite concentrated
in the luminal endothelial caveolae (4) and can be rapidly activated by
increased vascular flow/pressure (3, 4). Others have shown
calcium-independent activation of eNOS by exogenous ceramide (65) as
well as another pathway of eNOS regulation through phosphorylation by
the protein kinase Akt (67). Shear activates
phosphoinositide-3-kinase (PI-3-kinase) and ceramide can act
upstream of Akt by calcium-sensitive activation of PI-3-kinase,
possibly via Src-like kinases (68). Increased hemodynamic stressors and
exogenous ceramide can induce acute NO-dependent
vasodilation (69) as well as ERK (9) and plasmalemmal Src-like tyrosine
kinases (9, 11). Moreover, we have observed the activation of Akt in
response to both exogenous ceramide administration and high
pressure/flow in
situ.2
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. Martin Kroenke (University of Cologne, Germany) for the generous gift of preimmune and N-SMase antisera and Dr. Takeshi Ogita (Sankyo Co., Ltd, Tokyo, Japan) for the generous gift of scyphostatin. We also acknowledge Dr. Lucy Carver for help in writing this manuscript.
![]() |
FOOTNOTES |
---|
* This work was supported in part by National Institutes of Health Grant HL67386 (to J. E. S.) and the Beth Israel Foundation. This work was presented in part at the annual meeting of the Federation of American Societies for Experimental Biology in San Diego, CA, April 15-18, 2000 (Liu, J., and Schnitzer, J. E. (2000) FASEB J. 14, A457 (abstr.)).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Joint first authors who contributed equally to this work.
§ Present address: Department of Physiology and Pharmacology and MBR Cancer Center, West Virginia University School of Medicine, P. O. Box 9229, Morgantown, WV 26506.
¶ To whom correspondence should be addressed: Sidney Kimmel Cancer Center, Division of Vascular Biology and Angiogenesis, 10835 Altman Row, San Diego, CA 92121. Tel.: 619-450-5990 (ext. 320); Fax: 619-450-3251; E-mail: jschnitzer@skcc.org.
Published, JBC Papers in Press, December 6, 2002, DOI 10.1074/jbc. M210375200
2 M. Czarny and J.E. Schnitzer, unpublished observations.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: MAP, mitogen activated protein kinase; SMase, sphingomyelinase; A-SMase, acid SMase; N-SMase, neutral SMase; H, rat lung homogenate; P, luminal endothelial cell plasma membrane; PM, plasmalemmal fraction; V, caveolae separated from P in the presence of Triton X-100; V', caveolae separated from P in the absence of Triton X-100; P-V, P stripped of caveolae in the presence of Triton X-100; P-V', P stripped of caveolae in the absence of Triton X-100; T, Triton-soluble phase; S, sucrose fraction; AC, caveolin-rich membrane fraction; DETAPAC, diethylenetriamine pentaacetic acid; eNOS, endothelial nitric oxide synthase; ERK, extracellular signal-regulated kinase; FAK, focal adhesion kinase; JNK, c-Jun N-terminal kinase; BLMVEC, bovine lung microvascular endothelial cells; NO, nitric oxide; 5'-NT, 5' nucleotidase.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. |
Davies, P. F.
(1995)
Physiol. Rev.
75,
519-560 |
2. | Topper, J. N., and Gimbrone, M. A. J. (1999) Mol. Med. Today 5, 40-46[CrossRef][Medline] [Order article via Infotrieve] |
3. |
Rizzo, V.,
Sung, A., Oh, P.,
and Schnitzer, J. E.
(1998)
J. Biol. Chem.
273,
26323-26329 |
4. |
Rizzo, V.,
McIntosh, D. P., Oh, P.,
and Schnitzer, J. E.
(1998)
J. Biol. Chem.
273,
34724-34729 |
5. |
Furchgott, R. F.,
and Vanhoutte, P. M.
(1989)
FASEB J.
3,
2007-2018 |
6. | Ohno, M. (1993) Circulation 88, 193-197[Abstract] |
7. |
Kuchan, M., J., Jo, H.,
and Frangos, J. A.
(1994)
Am. J. Physiol.
267,
C753-758 |
8. |
Gudi, S.,
Nolan, J. P.,
and Frangos, J. A.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
2515-2519 |
9. |
Takahashi, M.,
and Berk, B. C.
(1996)
J. Clin. Invest.
98,
2623-2631 |
10. | Olesen, S. P., Clapham, D. E., and Davies, P. F. (1988) Nature 331, 168-170[CrossRef][Medline] [Order article via Infotrieve] |
11. | Rizzo, V., and Schnitzer, J. E. (1999) in Vascular Endothelium: Mechanisms of Cell Signaling (Catravas, J. D. , Callow, A. D. , and Ryan, U. S., eds), Vol. 308 , pp. 97-116, IOS Press, AmsterdamNATO Science Series A |
12. |
Jo, H.,
Sipos, K., Go, Y. M.,
Law, R.,
Rong, J.,
and McDonald, J. M.
(1997)
J. Biol. Chem.
272,
1395-1401 |
13. | Shyy, Y. J., Hsieh, H. J., Usami, S., and Chien, S. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 4678-4682[Abstract] |
14. |
Gimbrone, M. A.
(1999)
Am. J. Pathol.
155,
1-5 |
15. | Schnitzer, J. E. (1997) in Vascular Endothelium: Physiology, Pathology and Therapeutic Opportunities. (Born, G. V. R. , and Schwartz, C. J., eds) , pp. 77-95, Schattauer GmbH, Stuttgart, Germany |
16. | Schnitzer, J. E. (1995) Ann. Biomed. Eng. 23, S34 |
17. |
Park, H., Go, Y. M., St.,
John, P. L.,
Maland, M. C.,
Lisanti, M. P.,
Abrahamson, D. R.,
and Jo, H.
(1998)
J. Biol. Chem.
273,
32304-32311 |
18. |
Traub, O.,
and Berk, B. C.
(1998)
Arterioscler. Thromb. Vasc. Biol.
18,
677-685 |
19. |
Gudi, S. R.,
Clark, C. B.,
and Frangos, J. A.
(1996)
Circ. Res.
79,
834-839 |
20. |
Liu, J., Oh, P.,
Horner, T.,
Rogers, R. A.,
and Schnitzer, J. E.
(1997)
J. Biol. Chem.
272,
7211-7222 |
21. |
Oh, P.,
and Schnitzer, J. E.
(2001)
Mol. Biol. Cell
12,
685-698 |
22. | Liu, J., and Schnitzer, J. E. (1999) Methods Mol. Biol. 116, 61-72[Medline] [Order article via Infotrieve] |
23. |
Liu, P.,
and Anderson, R. G.
(1995)
J. Biol. Chem.
270,
27179-27185 |
24. |
Bilderback, T. R.,
Grigsby, R. J.,
and Dobrowsky, R. T.
(1997)
J. Biol. Chem.
272,
10922-10927 |
25. | Kolesnick, R. N., and Krönke, M. (1998) Annu. Rev. Physiol. 60, 643-665[CrossRef][Medline] [Order article via Infotrieve] |
26. | Levade, T., and Jaffrezou, J. P. (1999) Biochim. Biophys. Acta 1438, 1-17[Medline] [Order article via Infotrieve] |
27. |
Bernardo, K.,
Krut, O.,
Wiegmann, K.,
Kreder, D.,
Micheli, M.,
Schafer, R.,
Sickman, A.,
Schmidt, W. E.,
Schroder, J. M.,
Meyer, H. E.,
Sandhoff, K.,
and Kronke, M.
(2000)
J. Biol. Chem.
275,
7641-7647 |
28. | Schnitzer, J. E., McIntosh, D. P., Dvorak, A. M., Liu, J., and Oh, P. (1995) Science 269, 1435-1439[Medline] [Order article via Infotrieve] |
29. | Oh, P., and Schnitzer, J. E. (1998) in Cell Biology: A laboratory Handbook (Celis, J. E., ed), Vol. 2 , pp. 34-46, Academic Press, Orlando, FL |
30. |
Schnitzer, J. E.,
and Oh, P.
(1994)
J. Biol. Chem.
269,
6072-6082 |
31. |
Oh, P.,
and Schnitzer, J. E.
(1999)
J. Biol. Chem.
274,
23144-23154 |
32. | Wiegmann, K., Schutze, S., Machleidt, T., Witte, D., and Kronke, M. (1994) Cell 78, 1005-1015[Medline] [Order article via Infotrieve] |
33. |
Preiss, J.,
Loomis, C. R.,
Bishop, W. R.,
Stein, R.,
Niedel, J. E.,
and Bell, R. M.
(1986)
J. Biol. Chem.
261,
8597-8600 |
34. |
Liu, B.,
Hassler, D. F.,
Smith, G. K.,
Weaver, K.,
and Hannun, Y. A.
(1998)
J. Biol. Chem.
273,
34472-34479 |
35. | Romiti, E., Meacci, E., Tanzi, G., Becciolini, L., Mitsutake, S., Farnararo, M., Ito, M., and Bruni, P. (2001) FEBS Lett. 506, 163-168[CrossRef][Medline] [Order article via Infotrieve] |
36. |
Razandi, M., Oh, P.,
Pedram, A.,
Schnitzer, J. E.,
and Levin, E. R.
(2002)
Mol. Endocrinol.
16,
100-115 |
37. | Tanaka, M., Nara, F., Yamasato, Y., Ono, Y., and Ogita, T. (1999) J. Antibiot. 52, 827-830[Medline] [Order article via Infotrieve] |
38. |
Hannun, Y.
(1996)
Science
274,
1855-1859 |
39. | Santana, P., Pena, L. A., Haimovitz-Friedman, A., Martin, S., Green, D., McLoughlin, M., Cordon-Cardo, C., Schuchman, E. H., Fuks, Z., and Kolesnick, R. (1996) Cell 86, |
40. | Brown, D. A., and Rose, J. K. (1992) Cell 68, 533-544[Medline] [Order article via Infotrieve] |
41. |
Tomiuk, S.,
Hofmann, K.,
Nix, M.,
Zumbansen, M.,
and Stoffel, W.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
3638-3643 |
42. |
Chatterjee, S.,
Han, H.,
Rollins, S.,
and Cleveland, T.
(1999)
J. Biol. Chem.
274,
37407-37412 |
43. |
Hofmann, K.,
Tomiuk, S.,
Wolff, G.,
and Stoffel, W.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
5895-5900 |
44. | Jung, S. Y., Suh, J. H., Park, H. J., Jung, K. M., Kim, M. Y., Na, D. S., and Kim, D. K. (2000) J. Neurochem. 75, 1004-1014[CrossRef][Medline] [Order article via Infotrieve] |
45. | Mizutani, Y., Tamiya-Koizumi, K., Irie, F., Hirabayashi, Y., Miwa, M., and Yoshida, S. (2000) Biochim. Biophys. Acta 1485, 236-246[Medline] [Order article via Infotrieve] |
46. |
Chatterjee, S.,
and Ghosh, N.
(1989)
J. Biol. Chem.
264,
12554-12561 |
47. | Maruyama, E. N., and Arima, M. (1989) J. Neurochem. 52, 611-618[Medline] [Order article via Infotrieve] |
48. | Ghosh, N., Sabbadini, R., and Chatterjee, S. (1998) Mol. Cell. Biochem. 189, 161-168[CrossRef][Medline] [Order article via Infotrieve] |
49. | Liu, B., and Hannun, Y. A. (2000) Methods Enzymol. 311, 156-164[CrossRef][Medline] [Order article via Infotrieve] |
50. | Martin, S. F., Navarro, F., Forthoffer, N., Navas, P., and Villalba, J. M. (2001) J. Bioenerg. Biomembr. 33, 143-153[CrossRef][Medline] [Order article via Infotrieve] |
51. | Hostetler, K. Y., and Yazaki, P. J. (1979) J. Lipid Res. 20, 456-463[Abstract] |
52. |
Mizutani, Y.,
Tamiya-Koizumi, K.,
Nakamura, N.,
Kobayashi, M.,
Hirabayashi, Y.,
and Yoshida, S.
(2001)
J. Cell Sci.
114,
3727-3736 |
53. | Veldman, R. J., Maestre, N., Aduib, O. M., Medin, J. A., Salvayre, R., and Levade, T. (2001) Biochem. J. 355, 859-868[Medline] [Order article via Infotrieve] |
54. |
Nusrat, A.,
Parkos, C.,
Verkade, P.,
Foley, C.,
Liang, T.,
Innis-Whitehouse, W.,
Eastburn, K.,
and Madara, J.
(2000)
J. Cell Sci.
113,
1771-1781 |
55. | Barenholz, Y., and Thompson, T. E. (1980) Biochim. Biophys. Acta 604, 129-158[Medline] [Order article via Infotrieve] |
56. | Mohan Das, D. V., Cook, H. W., and Spence, M. W. (1984) Biochim. Biophys. Acta 777, 339-342[Medline] [Order article via Infotrieve] |
57. | Allan, D., and Quinn, P. (1988) Biochem. J. 254, 765-771[Medline] [Order article via Infotrieve] |
58. | Chatterjee, S. (1999) Chem. Phys. Lipids 102, 79-96[CrossRef][Medline] [Order article via Infotrieve] |
59. | Verheij, M., Bose, R., Lin, X. H., Yao, B., Jarvis, W. D., Grant, S., Birrer, M. J., Szabo, E., Zon, L. I., Kyriakis, J. M., Haimovitz-Friedman, A., Fuks, Z., and Kolesnick, R. N. (1996) Nature 380, 75-79[CrossRef][Medline] [Order article via Infotrieve] |
60. |
Raines, M. A.,
Kolesnick, R. N.,
and Golde, D. W.
(1993)
J. Biol. Chem.
268,
14572-14575 |
61. | Yao, B., Zhang, Y., Delikat, S., Mathias, S., Basu, S., and Kolesnick, R. (1995) Nature 378, 307-310[CrossRef][Medline] [Order article via Infotrieve] |
62. |
Gulbins, E.,
Szabo, I.,
Baltzer, K.,
and Lang, F.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
7661-7666 |
63. |
Hida, H.,
Takeda, M.,
and Soliven, B.
(1998)
J. Neurosci.
18,
8712-8719 |
64. |
Hanna, A. N.,
Chan, E. Y., Xu, J.,
Stone, J. C.,
and Brindley, D. N.
(1999)
J. Biol. Chem.
274,
12722-12729 |
65. |
Igarashi, J.,
Thatte, H. S.,
Prabhakar, P.,
Golan, D. E.,
and Michel, T.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
12583-12588 |
66. |
Katsuyama, K.,
Shichiri, M.,
Marumo, F.,
and Hirata, Y.
(1998)
Endocrinology
139,
4506-4512 |
67. |
Dimmeler, S.,
Assmus, B.,
Hermann, C.,
Haendeler, J.,
and Zeiher, A. M.
(1998)
Circ. Res.
83,
334-341 |
68. |
Su, X.,
Wang, P.,
Ibitayo, A.,
and Bitar, K. N.
(1999)
Am. J. Physiol.
276,
G853-G861 |
69. | Jin, J. S., Tsai, C. S., Si, X., and Webb, R. C. (1999) Chin. J. Physiol. 42, 47-51[Medline] [Order article via Infotrieve] |