From the Departments of Biochemistry and
§ Cell Biology, Weill Medical College of Cornell
University, New York, New York 10021
Received for publication, December 5, 2002
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In response to chemoattractants neutrophils
extend an actin-rich pseudopod, which imparts morphological polarity
and is required for migration. Even when stimulated by an isotropic
bath of chemoattractant, neutrophils exhibit persistent polarization
and continued lamellipod formation at the front, suggesting that the
cells establish an internal polarity. In this report, we show that
perturbing lipid organization by depleting plasma membrane cholesterol
levels reversibly inhibits cell polarization and migration. Among other
receptor-mediated responses, Over the past decade, compelling evidence has emerged that
challenges the notion of the plasma membrane lipid bilayer as a homogeneous passive entity that merely provides a scaffold for protein-mediated signaling (1, 2). This evidence suggests that certain
lipids preferentially associate and form lateral heterogeneities in the
membrane (3-6). One type of membrane domain has been described as
rafts, glycolipid-enriched membrane domains (GEMs), or
detergent-resistant membrane domains (DRMs) because they are enriched
in glycosphingolipids, sphingomyelin, and cholesterol, which make them
resistant to solubilization by cold non-ionic detergents (7). These
lipid domains have been postulated to serve as centers for some signal
transduction processes by virtue of their copurification with signaling
molecules following cell lysis with cold nonionic detergents and
floatation on sucrose density gradients (3). Because the direct
visualization of lipid domains has been elusive, it is thought that
they typically exist as dynamic submicron-sized regions within the
plasma membrane (6, 8-10). However, most of the surface area of cells
such as fibroblasts and neutrophils is resistant to extraction by cold Triton X-100 (11-13), suggesting that lipid organization may be more
complex than depicted in a simple version of the raft model. In fact,
several studies now indicate that multiple types of membrane microdomains co-exist within the plasma membranes of cells (14-16) and
may coalesce into dense assemblies to form larger domain structures (1,
2, 11, 15).
Cholesterol is the most abundant lipid component of the plasma
membrane, and it plays an important role in lipid organization. Extraction of cholesterol from the plasma membrane using the synthetic cholesterol chelator methyl- Migration of neutrophils and other immune cells is contingent upon
their ability to adopt a polarized morphology in response to
chemotactic stimulation. In the presence of chemoattractants, neutrophils rapidly transform from roughly spherical resting cells to
migratory ones with distinctive leading and trailing edges. Actin
polymerization occurs almost exclusively at the leading edge, resulting
in a dramatic accumulation of F-actin at just one end of the cells
(21). Notably, this remarkable asymmetry occurs even when the external
chemotactic signal is uniform, suggesting that at least one signaling
step leads to an internal polarization of the cell. Given that
chemoattractant receptors are typically distributed uniformly across
the cell surface even after polarization (22), this internal signal
occurs somewhere between receptor occupancy and actin polymerization.
The exact point at which polarization is induced is not known. However,
the possibilities have been narrowed by an elegant study in
neutrophil-differentiated HL-60 cells using the recruitment of a
GFP-tagged plekstrin homology domain to the plasma membrane as a
readout for polarization (22). This study showed that amplification of
polarization signals depends on one or more of the Rho GTPases, which
anchor to the plasma membrane via lipid tails, and is downstream of the
lipid products of phosphatidylinositol 3-kinase (22). In other words,
amplification of polarization occurs at or near the plasma membrane.
This, together with our finding that the plasma membrane of neutrophils
segregates into distinct lipid domains that comprise either pole of
migrating neutrophils (11), led us to hypothesize that signal
amplification is dependent on plasma membrane domains. To test this, we
used M Neutrophil Preparation and Stimulation--
Polymorphonuclear
neutrophils (PMNs) were isolated from whole blood donated by
healthy volunteers by centrifugation through Polymorphprep
(Invitrogen). After lysis of contaminating erythrocytes by a 30-s
hypotonic shock, cells were washed with phosphate-buffered saline (pH
7.4), and then resuspended in incubation buffer (150 mM
NaCl, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, 10 mM glucose, 20 mM HEPES, pH 7.4). For all experiments, PMNs were plated
onto fibronectin-coated coverslip dishes for 5 min at 37 °C, and
then stimulated with 10 nM formyl-Met-Leu-Phe (fMLF) for
the indicated times.
Labeling and Detergent Extraction--
To visualize membrane
domains, PMNs were plated and stimulated as above, and then extracted
with ice-cold detergent. Specifically, cells were bathed in ice-cold
cytoskeleton stabilizing buffer (CSB; 138 mM KCl, 3 mM MgCl2, 2 mM EGTA, 0.32 M sucrose, 10 mM MES, pH 6) containing 0.5%
Triton X-100 and 1× protease inhibitor mixture (BD PharMingen) for 30 min, and then washed with ice-cold CSB. Samples were fixed with 3.3%
paraformaldehyde and 0.05% glutaraldehyde in phosphate-buffered saline
for 10 min on ice, and then incubated sequentially with a monoclonal
anti-CD44 (clone HERMES-3, American Type Culture Collection, Manassas,
VA) and AlexaFluor 488-goat anti-mouse secondary antibody (Molecular
Probes, Eugene, OR). Anti-CD44-labeled cells were imaged immediately
after completion of the labeling protocol. Images were acquired with a
Zeiss LSM510 laser scanning confocal microscope (Jena, Germany) or a
Leica DMIRB widefield microscope, and then analyzed with MetaMorph
image analysis software (Universal Imaging Corporation, Downington, PA).
Cholesterol Depletion and Repletion--
Plasma membrane
cholesterol was depleted by incubating PMNs (1.2 × 107/ml) in incubation buffer containing 10 mM
M Immunofluorescence--
After stimulation with fMLF, cells were
fixed with 3.3% paraformaldehyde in the presence of 0.25 mg/ml saponin
for 5 min at room temperature. When it was important to preserve the
labile pools of F-actin, excess fluorescently conjugated phalloidin
(Molecular Probes, Eugene, OR) was included during fixation and
throughout the remaining labeling steps. Rac was visualized with a
mouse monoclonal antibody against Rac1 and Rac2 (clone 23A8, Upstate Biotechnology Inc., Lake Placid, NY), followed by an AlexaFluor 546-conjugated goat anti-mouse secondary antibody (Molecular Probes). Cells labeled for Rac were imaged by either confocal or wide-field fluorescence microscopy. Wide-field fluorescence images of all samples
within one experiment were acquired under identical conditions and
quantified using Metamorph image analysis software. Following background correction, the average fluorescence intensity per cell was
measured for more than 100 cells per condition, and these measurements
were normalized to the level of the unstimulated control cells.
Scanning Electron Microscopy--
Cells were fixed in 2%
buffered glutaraldehyde, washed with 0.1 M sodium
cacodylate buffer pH 7.3, then postfixed in 1% osmium tetroxide in
sodium cacodylate buffer and washed. Samples were then dehydrated
through a graded ethanol series, freeze-dried, and sputter-coated with
gold-palladium. Samples were examined at 20 kV using a JEOL 100 CX-II electron microscope fitted with an ASID-scanning unit, and
photographs were recorded on Polaroid Type 55 P/N film.
Quantification of F-actin Content--
PMNs in suspension were
stimulated with fMLF (10 nM) for 5 min at 37 °C, then
simultaneously fixed, permeabilized, and labeled for F-actin by the
addition of an equal volume of phosphate-buffered saline containing
6.6% paraformaldehyde, 0.1% glutaraldehyde, 0.5 mg/ml saponin, and 2 units/ml AlexaFluor 488 phalloidin. Cell-associated fluorescence was
measured by flow cytometry (Beckman-Coulter XL, Fullerton, CA) for
2,000 cells per condition and from three separate experiments. Aliquots
from each sample were removed and imaged by confocal microscopy. To
verify the results in adherent cells, PMNs were plated, stimulated,
fixed, and labeled with TRITC-conjugated phalloidin as described above.
Wide-field images of fluorescent-labeled cells were obtained using a
Leica DMIRB equipped with a 63 × 1.32 numerical aperture
objective and a Princeton Instruments (Princeton, NJ) cooled CCD camera
driven by MetaMorph Imaging System software. Following background
correction, the average fluorescence intensity per cell was measured
for over 150 cells per condition from three experiments on different
days. The average fluorescence intensity per cell for
cholesterol-depleted cells was 46 ± 8% (S.D.) of the value for
control cells.
Functional Assays--
Motility assays were performed as
described (23). Briefly, motility of plated and stimulated PMNs was
monitored for 4 min using a Leica DMIRB (Leica Mikroscopie und Systeme
GmbH, Germany) set up for differential interference contrast
microscopy. Time-lapse images were acquired with a cooled CCD camera
driven by MetaMorph Imaging System software. Migrating PMNs were
defined as those whose tail and leading lamella moved at least 7 µm
from their initial starting position within 240 s. Separate dishes
were used for each treatment, and at least three dishes were monitored
for each treatment on at least three different days.
To monitor changes in intracellular free calcium levels
([Ca2+]i), PMNs were loaded with
the ratiometric fluorescent indicator, fura-2 as described (24).
Briefly, cells were incubated in a 5 µM solution of
acetoxymethyl ester-derivatized fura-2 (Fura-2/AM, Molecular Probes)
for 40 min at room temperature. Cells were then washed and incubated
for an additional 10 min at room temperature to allow cleavage of the
ester. Cells were treated with M Inhibition of Actin-Myosin Contraction--
During plating,
neutrophils were incubated with 10 µM myosin light chain
kinase inhibitor, ML-7 (26). Cells were then stimulated in the
continued presence of ML-7 for the indicated times.
In response to the chemoattractant, fMLF neutrophils undergo
polarized morphological changes with actin-driven protrusion of a
lamellipod at just one end of the cell. Because of the dramatic morphological and functional asymmetry exhibited by migrating cells, it
has been suggested that the front and back of polarized cells may
manifest different lipid environments (11, 15, 17, 27). Indeed, we have
shown that cold Triton X-100 preferentially removes a lipid marker of
rafts, DiIC16, from the leading lamellae of polarized PMNs
but extracts other non-raft lipid probes uniformly (11). Likewise,
several protein constituents of rafts, including the GPI-linked CD16
and the transmembrane CD44, are retained preferentially at the rear of
polarized PMNs following extraction with cold Triton X-100 (Fig.
1 and Ref. 11). These findings, coupled
with those from other laboratories (15, 17), provide compelling
evidence that the lipid environments at the front and back of polarized PMNs are dissimilar and may represent micron-scale membrane
domains.
2 integrin
up-regulation was unaffected, and initial calcium mobilization was only
partially reduced by cholesterol depletion, indicating that this
treatment did not abrogate initial receptor-mediated signal
transduction. Interestingly, cholesterol depletion did not prevent
initial activation of the GTPase Rac or an initial burst of actin
polymerization, but rather it inhibited prolonged activation of
Rac and sustained actin polymerization. Collectively, these
findings support a model in which the plasma membrane is organized into
domains that aid in amplifying the chemoattractant gradient and
maintaining cell polarization.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
Discussion
REFERENCES
-cyclodextrin
(M
CD)1 causes
reorganization of lipids with preferences for both ordered and
disordered domains (13). While the effects of cholesterol depletion on
ordered, raft-like domains have been emphasized, additional effects on
the overall organization of lipids also occur (13). Cholesterol
depletion has been used to examine the role of lipid domains in a
multitude of cellular processes, including cell migration (15, 17) and
phagocytosis (18-20). Disruption of lipid domain organization by
M
CD treatment resulted in inhibition of motility of breast
cancer-derived cells (17) and T cells (15), and abrogation of the
phagocytosis of Escherichia coli by mast cells (18) and
mycobacteria by macrophages (20) and neutrophils (19). Because cell
motility and phagocytosis are mechanistically related, with both
processes involving directional extension of actin-rich membranes, it
seems likely that their inhibition following domain disruption may have
the same underlying molecular basis.
CD to disrupt membrane domain organization in human
neutrophils and studied its effects on polarization and signaling.
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
Discussion
REFERENCES
CD (Sigma) for 15 min at 37 °C. At the end of this 15-min
treatment, cells were diluted to 2.4 × 106/ml with
incubation buffer (depleted cells). The extent of cholesterol depletion
was determined by an enzymatic method using a commercially available
kit (Free cholesterol C; Wako Biochemicals, Osaka). The cholesterol
content of M
CD-treated cells was measured to be 20.9 ± 3.6%
(mean ± s.d., n = 5) less than that of control cells. To replete membrane cholesterol, cholesterol-depleted cells were
incubated with 5 mM Chol-M
CD for 2.5 min at 37 °C
(repleted cells).
CD or not, and then placed on ice
until use. 2-ml suspensions of fura-2-loaded PMNs (1-1.5 × 106 cells/ml) were maintained at 37 °C in cuvettes with
continual stirring throughout each experiment.
[Ca2+]i was monitored with a SLM
8000C spectrofluorometer (Aminco, Urbana, IL) operated in dual
excitation mode (excitation 340 and 380 nm, emission 510 nm). In the
absence of stimulation, there was a negligible upward drift of the
signal due to leakage of dye from the cells (not shown). To quantify
and compare peak responses, fractional responses (F) for each sample
were calculated as the change in the ratio value induced by addition of
fMLF (i.e. the difference between the peak ratio value
attained and the basal ratio value: Ipeak
Ibasal) divided by the maximum range of ratio values (25). The maximum range of ratio values was determined for each
sample by addition of 0.5% Triton X-100 at the end of each experiment
to obtain the maximum possible ratio value
(Imax), followed by addition of 4 mM
EGTA to obtain the minimum ratio value (Imin).
In summary, fractional responses were calculated using the formula in
Equation 1.
To verify the ability of PMNs to upregulate their integrins in
response to fMLF, PMNs were incubated at 37 °C with Fab fragments of
an anti-
(Eq. 1)
2 integrin antibody (IB4) that was directly
conjugated to AlexaFluor 488 in the presence or absence of fMLF, as
indicated. At various time points after the start of the incubation
period, aliquots of cells were removed and diluted into ice-cold
incubation buffer to stop exocytosis. Cell-associated fluorescence was
monitored using a Beckman-Coulter XL analytical flow cytometer. To
account for any nonspecific binding of the IB4 Fab, fluorescence
intensity of PMNs incubated with an irrelevant control AlexaFluor
488-conjugated Fab was set to an arbitrary value, and all other samples
were measured relative to this value. Each data point represents
measurements from 2,000 cells, and the data as a whole are
representative of experiments from three different days.
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
Discussion
REFERENCES
View larger version (89K):
[in a new window]
Fig. 1.
Plasma membrane cholesterol depletion
inhibits stimulation-induced membrane domain coalescence. Control
(a-c) or cholesterol-depleted (d-f) PMNs were
labeled with a fluorescently conjugated monoclonal antibody against the
membrane domain marker, CD44, and then stimulated for 0 (a,
d), 30 (b, e) or 60 (c,
f) s with fMLF. At the end of the stimulation period, cells
were extracted with cold Triton X-100 to reveal membrane domain
organization. Differential interference contrast (DIC, a-f)
and fluorescence (CD44, a'-f') images are shown.
Asterisks in b and c indicate cell
uropods. Bar, 10 µm.
MCD, a chelator of cholesterol, has been used to selectively deplete
plasma membrane cholesterol and thereby alter lipid organization in
several cell types (8, 13, 17, 28). Another common method of depleting
membrane cholesterol is long term treatment of cells with HMG-CoA
reductase inhibitors (i.e. statins), which work by blocking
cholesterol biosynthesis. Because the levels of mevalonate and other
intermediates of cholesterol synthesis (including isoprenoids) are
altered by treatment with statins, and because these intermediates are
necessary for cell proliferation and other important cell functions,
statins are likely to have pleiotropic effects on cells that may be
explained by reasons other than cholesterol reduction (29-35). To
avoid the non-cholesterol related effects of long-term cholesterol
depleting strategies, we used M
CD to acutely deplete cholesterol
from PMNs. To determine if cholesterol depletion by M
CD would also
disrupt the large-scale lipid organization found in PMNs, we visualized
domain organization in control (Fig. 1, a-c) and
M
CD-treated (Fig. 1, d-f) cells. Domain
organization was revealed by extracting cells with cold Triton X-100
and then labeling the cells by indirect immunofluorescence for the
transmembrane protein, CD44, which localizes to raft-like domains in
PMNs (11). Fig. 1 (a'-c') shows the progression
of domain organization as control cells polarize in response to fMLF. As we reported previously, rafts are evenly distributed around the
periphery of unstimulated cells (Fig. 1a'), form larger
patches after 15-30 s of stimulation (Fig. 1b'), and
finally coalesce into a cap toward the cell rear (Fig. 1c')
(11).
After the plasma membranes of cells have been cholesterol-depleted with
MCD (Fig. 1, d-f), the raft component CD44 is
less well retained following detergent extraction; the average
fluorescence intensity per cholesterol-depleted cell was decreased by
40.8% ± 3.2% (mean ± S.D.) compared with control cells. (The
intensities of panels d'-f' have been
enhanced relative to panels a'-c' to allow visualization of
the CD44 distribution). In contrast to control cells, fMLF stimulation
does not induce capping of CD44 in cholesterol-depleted cells; rather,
the CD44 that is retained remains relatively uniformly distributed
around the cell periphery (Fig. 1,
d'-f'). That is, cholesterol depletion
inhibits the redistribution of detergent-resistant raft components into
a cap at the cell rear, indicating that M
CD treatment inhibits large
scale lipid organization in addition to disrupting microdomains.
Acute depletion of cholesterol by treatment with MCD has dramatic
effects on neutrophil polarization and migration (Fig. 2). When cellular cholesterol is depleted
by just ~21% (see "Materials and Methods"), PMN migration is
inhibited by >90% (Fig. 2a), consistent with the report by
Manes et al. (17) that depletion of membrane cholesterol
inhibits migration of MCF-7 cells. However, in contrast to the
interpretation reported in Manes et al., we find that
cholesterol depletion abolishes lamellipod formation (Fig.
2b, compare center to left panel).
Inhibition of migration and polarization can be attributed to effects
of cholesterol modulation as opposed to nonspecific effects of M
CD
treatment since the ability to polarize and migrate is restored to
previously cholesterol-depleted cells upon replenishing membrane
cholesterol with cholesterol-chelated M
CD (chol-M
CD, Fig. 2,
a and b, repleted).
|
Because it has been reported, based on observation of cells by light
microscopy, that cholesterol depletion does not affect membrane
extension and ruffling (17, 20), we further investigated whether
lamellipod extension in PMNs was affected by changes in cholesterol
levels. Scanning electron microscopy reveals that cholesterol-depleted
cells are unable to form membrane extensions or ruffles in response to
fMLF (compare Fig. 3, b to
a). The lack of membrane ruffling in cholesterol-depleted
cells is accompanied by an inhibition of stimulated actin
polymerization. Following cholesterol depletion and stimulation with
fMLF, cells were simultaneously fixed, permeabilized, and stained for
F-actin with fluorescently conjugated phalloidin. Projections of
confocal images show that control cells exhibited dramatic F-actin-rich
ruffles within their lamellae (Fig. 3c), whereas
cholesterol-depleted cells had no F-actin-rich projections (Fig.
3d). Using flow cytometry, we quantified the fluorescent
phalloidin binding to F-actin in parallel samples of suspended
cholesterol-depleted and control cells, which had been stimulated with
fMLF, or left unstimulated, for 5 min at 37 °C. Typically in
response to fMLF PMNs increased their F-actin content by 2.5-3-fold
(Fig. 3e, control). In contrast, fMLF stimulation of cholesterol-depleted cells led to only a 1.5-fold increase in
F-actin content over unstimulated cholesterol-depleted cells (Fig.
3e, depleted). In these experiments, the final
F-actin content of fMLF-stimulated cholesterol-depleted cells was 49%
of the value for fMLF-stimulated control cells. We obtained similar
results for adherent cells in which we measured the F-actin content of fMLF-stimulated cholesterol-depleted cells to be 46% that of
stimulated control cells (data not shown). Aliquots of the flow
cytometry samples of unstimulated PMNs were plated on a slide, and then imaged by confocal microscopy to examine the effects of MCD
treatment on F-actin structure. Fig. 3, f and g
show the F-actin structure in unstimulated control and
cholesterol-depleted cells, respectively. M
CD treatment causes a
slight disruption in the cortical F-actin structure in unstimulated
PMNs, with only an insignificant effect on the total F-actin content of
unstimulated cells (Fig. 3e, compare gray
bars).
|
Accumulation of F-actin at the leading edge of human neutrophils and
subsequent lamellipodia formation and membrane ruffling, is at least
partially due to activation of the small GTPases Rac1 and Rac2
(henceforth, collectively referred to as Rac, albeit Rac2 is the
predominant isoform (>96%) in neutrophils) (36-40). In light of
reports implicating an important role for Rac in stimulated actin
polymerization in neutrophils (40, 41), and because the Rho GTPases
associate with membranes via C-terminal lipid modifications, we tested
whether Rac targeting was affected by changes in cholesterol levels.
F-actin within newly formed lamellae is structurally and spatially
distinct from cortical F-actin within the cell body (42) and is
observed as short spikes or fingers when labeled with
fluorophore-conjugated phalloidin (Fig.
4c'). It has been shown that
establishment and maintenance of this asymmetrical distribution of
F-actin spikes involves the Arp2/3 complex, and it was proposed that
recruitment of Arp2/3 might be achieved through a Rho family member
(43). Indeed, actin nucleation in neutrophils in response to fMLF has
been shown to occur via both Cdc42- and Rac-dependent
pathways (40), but spatial control of these signaling molecules was not
shown. Using indirect immunofluorescence we visualized endogenous Rac
in unstimulated and fMLF-stimulated PMNs. In unstimulated PMNs (Fig.
4a), Rac (Fig. 4a") is found throughout the
cytoplasm and bounded within the cortical F-actin ring (Fig.
4a'). This observation is consistent with biochemical analyses of membrane and cytosol fractions from human neutrophils that
have shown that Rac in unstimulated cells is nearly entirely cytosolic,
with no detectable Rac associated with membrane fractions (44). Within
15-30 s of stimulation, PMNs form multiple lamellipodia (Fig.
4b), which protrude in all directions and are rich in
F-actin spikes (Fig. 4b'). Strikingly, each of the
F-actin-containing membrane extensions stains brightly for Rac (Fig.
4b"). The large increase in fluorescence intensity for the
Rac staining in stimulated compared with resting cells is likely
because activated membrane-bound Rac is better retained than cytosolic
Rac after cell permeabilization. After 60 s of stimulation, when
PMNs have become fully polarized (Fig. 4c), endogenous Rac
is asymmetrically distributed to a band at the leading edge (Fig.
4c") that localizes to the same region of the cell as the
F-actin spikes (Fig. 4c'). The overlay images show that Rac
and F-actin spikes colocalize to similar regions of the cells. Rac
localizes to the plasma membrane, while F-actin is intracellular, so
these molecules are not coincident on the molecular level, and thus
they will not necessarily appear as yellow in the overlay images. The
localization of Rac shown in Fig. 4, b" and c" is
consistent with the idea that Rac is locally active, as seen in motile
Swiss 3T3 fibroblasts (45) and mediates actin polymerization in those
regions.
|
Localization of Rac in cholesterol-depleted cells suggests an explanation for the lack of membrane ruffling and actin polymerization in these cells (Fig. 4, d-f). As for control cells (Fig. 4, a-c), cholesterol-depleted cells undergo a burst of actin polymerization and Rac recruitment to the membrane within 15-30 s of stimulation (compare Fig. 4, d' and e' to d" and e"). However, unlike control cells, distinct lamellipodia are not observed in these cells, and the F-actin appears circumferentially distributed (Fig. 4e'). Also, there is little correlation between Rac localization (Fig. 4e") and F-actin (Fig. 4e' and overlay); where there are brighter regions of F-actin staining in depleted cells, there are not necessarily bright regions of Rac staining, whereas there is a one-to-one correlation in control cells. After 60 s of stimulation (Fig. 4f), F-actin in cholesterol-depleted cells is still localized around the cell perimeter (Fig. 4f') and is much less intense than that found in corresponding control cells (Fig. 4c'). Interestingly, in contrast to control cells, the Rac staining (Fig. 4f") has diminished in intensity compared with cells that have been stimulated for just 15-30 s (Fig. 4e"), suggesting that Rac in these cells was only transiently activated.
Quantifying wide-field fluorescence images of cells labeled for Rac provides a means to assess the effect of cholesterol depletion on stimulated Rac recruitment to the membrane (Fig. 4g). Consistent with the images shown in Fig. 4(a-c), stimulation of control cells with fMLF causes a marked and sustained increase in the intensity of Rac staining compared with unstimulated cells. The intensity of Rac labeling in control cells remained elevated until at least 120 s after stimulation. In contrast, fMLF stimulation of cholesterol-depleted cells caused only a transient increase in Rac labeling intensity, with Rac intensity diminishing to close to the starting level by 120 s after stimulation.
The dramatic inhibition of fMLF-stimulated actin polymerization in
cholesterol-depleted PMNs could result from a general inhibition of
receptor-mediated signaling. To investigate this, and to further investigate the observed effect of MCD on cell morphology, we examined the effect of M
CD treatment on fMLF-stimulated
2 integrin up-regulation.
2 integrins are
the most abundant integrins in PMNs, and their up-regulation at the
surface and activation are required for cell spreading and migration
(46). Consequently, another explanation for the absence of shape change
of cholesterol-depleted PMNs could be inhibition of
2
integrin transport to the surface. To exclude this possibility, as well
as to test the effects of M
CD on fMLF-stimulated secretion, we used
flow cytometry to monitor the surface expression of
2
integrins in control and cholesterol-depleted cells over time. In these
experiments, fMLF-stimulated integrin up-regulation was detected with
fluorescently conjugated Fab fragments of a monoclonal antibody against
2 integrins. Fab fragments were used as opposed to whole
antibody because cross-linking of
2 integrins can induce
secretion in the absence of fMLF stimulation. We verified that the
fluorescently conjugated anti-
2 Fab fragments did not
induce secretion on their own (Fig.
5a, open symbols). Despite the profound effects on spreading and migration, cholesterol depletion had no effect on the extent or kinetics of fMLF-stimulated up-regulation of
2 integrins (Fig. 5a,
filled symbols), indicating that cholesterol-depleted cells
are able to carry out regulated secretion in response to fMLF. To
visually verify the flow cytometry data, aliquots of cells
were washed, fixed, and imaged by fluorescence microscopy. Fig.
5b shows images of PMNs before and after 15 min of
stimulation with fMLF. As demonstrated by flow cytometry (Fig. 5a), control and cholesterol-depleted PMNs show a comparable
increase in expression of
2 integrins following fMLF
stimulation. Activation of integrins is also apparently unaffected by
cholesterol depletion as there was no obvious difference in adhesion
between cholesterol-depleted and control cells (not shown). These
results show that several consequences of stimulation by fMLF are not
affected significantly by the cholesterol depletion protocol we
used.
|
To further examine whether MCD inhibits responses other than actin
polymerization and cell polarization, we tested the effects of M
CD
treatment on an early fMLF-mediated signaling event, elevation of
intracellular free calcium levels
([Ca2+]i). Fig.
6 shows representative
experiments in which we monitored fMLF-induced changes in
[Ca2+]i over time for both control
and cholesterol-depleted cells. For these studies, PMNs were loaded
with the ratiometric indicator, fura-2, treated with M
CD (or not),
and then monitored by spectrofluorometry. As shown in Fig.
6a, M
CD-treated cells are able to mobilize
[Ca2+]i in response to 10 nM fMLF, but the response is diminished compared with
control cells. The initial rise of cholesterol-depleted cells was
78.7 ± 7.8% (S.E., n = 9) of control cells.
Interestingly, although the peak calcium response in
cholesterol-depleted cells was only partially reduced compared with
control cells, the sustained response was almost completely inhibited.
These findings are generally similar to those reported by Barabe
et al. (47) but they are not identical. In that study,
cholesterol depletion with M
CD was likewise shown to inhibit the
sustained calcium response induced by fMLF in PMNs; however, in
contrast to our findings, it had no effect on the peak response. In
those studies, PMNs were stimulated with 100 nM fMLF as
opposed to 10 nM fMLF, as used in Fig. 6a. To
determine if this difference in dose could account for the difference
in our findings, we performed the same experiment using 100 nM rather than 10 nM fMLF. Fig. 6b
shows that the peak response of cholesterol-depleted cells is not
diminished when the cells are stimulated with this higher dose of fMLF,
even though the sustained phase of the response is inhibited as
observed for stimulation with 10 nM fMLF.
|
The sustained phase of calcium elevation following fMLF stimulation includes contributions from both capacitative calcium entry across the plasma membrane and repeated transient releases from internal stores (48). To determine if the effect of cholesterol depletion on the sustained calcium response was due to an effect on calcium entry across the plasma membrane or an effect on the release from internal stores, cholesterol-depleted and control PMNs were stimulated in the absence of extracellular calcium (Fig. 6, a and b, dotted lines). Both the peak and sustained responses of control cells are diminished when extracellular calcium is removed (compare dotted to solid black lines). Similarly, the peak response of cholesterol-depleted cells in the absence of extracellular calcium was less than that of cholesterol-depleted cells in calcium containing medium (compare dotted to solid gray lines), and less than control cells in calcium-free medium (compare dotted gray to dotted black line, Fig. 6a). Our results with 100 nM fMLF are similar to those reported by Barabe et al. (47) However, it should be noted that in all of our experiments, the differences between the calcium responses of control and cholesterol-depleted cells were always greater when the cells were stimulated with 10 nM fMLF than when cells were stimulated with 100 nM fMLF, and we found that the peak response of cholesterol-depleted cells was almost always smaller than that of control cells when the cells were stimulated with 10 nM fMLF in the presence of EGTA.
One possible explanation for the diminution of the peak calcium
response of cholesterol-depleted cells compared with control cells
under both calcium-free and replete conditions is that cholesterol depletion affects the endoplasmic reticulum (ER) membrane as well as
the plasma membrane, and thus affects internal calcium stores. To test
this possibility, cholesterol-depleted and control PMNs were treated
with 100 nM thapsigargin, which depletes ER calcium stores
by preventing calcium reuptake. For the first several minutes, the
increase in intracellular calcium induced by thapsigargin is identical
in control and cholesterol-depleted cells, but then the responses
diverge, with the calcium level in control cells continuing to rise,
while that in depleted cells plateaus (compare solid black
to gray traces in Fig. 6c). The deviation in
responses likely results from differences in calcium entry across the
plasma membrane, and, in fact, the responses of cholesterol-depleted and control cells to thapsigargin are indistinguishable when the cells
are monitored in calcium-free medium (Fig. 6c, dotted
lines). These results are consistent with the findings by Barabe
et al. (47) that Mn2+ influx is blocked by
MCD treatment, and they support the idea that M
CD treatment
reduces capacitative calcium entry across the plasma membrane.
Since cholesterol depletion altered some calcium responses of PMNs, we
explored whether the lack of shape changes in cholesterol-depleted cells is a consequence of weakened calcium responses in these cells.
First, we tested the ability of cholesterol-depleted cells to undergo
shape changes in response to stimulation with100 nM fMLF.
Fig. 6d shows that 100 nM fMLF does not induce
shape changes in cholesterol-depleted PMNs, even though the magnitudes
of the initial calcium response to this dose of fMLF are comparable in control and MCD-treated cells (see Fig. 6b).
As a second test, we considered whether inhibition of calcium entry and depletion of intracellular calcium stores would produce effects similar to cholesterol depletion. We had observed previously that PMNs polarize and extend lamellae in the absence of external calcium (47). Pretreatment of PMNs with 100 nM thapsigargin for 3 min was used to deplete intracellular calcium stores (47), and extracellular EGTA was used to prevent influx of calcium across the plasma membrane. Fig. 6d shows that thapsigargin-treated cells in the absence of extracellular calcium readily spread and undergo shape changes within 60 s of stimulation with 10 nM fMLF. At later times after stimulation, these cells extended lamellae and became polarized in response to fMLF, although the process was somewhat slower than in control cells (not shown). This result, which is consistent with previous observations of [Ca2+]i- buffered PMNs (49), suggests that the effects of cholesterol depletion on lamellar extension are not due to depletion of intracellular calcium stores or to inhibition of calcium influx.
The results obtained from our cholesterol depletion studies suggest
that membrane domain organization is important for cell polarization.
To further test this hypothesis, we wanted a second distinct method of
disrupting domain organization. We have shown previously that when
myosin was inhibited by treatment with ML-7, CD44-containing membrane
domains did not redistribute upon stimulation of PMNs (11). That is,
CD44-containing domains did not form caps, but rather remained
distributed in patches around the cells and interspersed with
CD44-negative domains. Using this independent method to disrupt
large-scale domain organization, we looked at the effects on F-actin
and Rac localization (Fig. 7). In
striking contrast to the effect of depleting plasma membrane
cholesterol, inhibiting actin-myosin contraction by treatment of PMNs
with ML-7 leads to the circumferential extension of a single large lamella following fMLF stimulation (26) (Fig. 7, b and
c). Unlike fMLF-stimulated control cells (Fig.
4c'), F-actin in stimulated ML-7-treated cells (Fig. 7,
b' and c') is distributed evenly around the
cells, without a vectorial bias. Fig. 7, b" and c" show that Rac, similar to the F-actin, is distributed around the circumference of
the cells, demonstrating that Rac activation is not inhibited by ML-7
treatment, but its polarized localization is disrupted.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
When neutrophils and other leukocytes polarize, the lipids at the
front and rear of the cells take on different properties (11). This can
be most clearly seen by the difference in sensitivity to solubilization
by cold Triton X-100, with the rear of the cells being largely
resistant to extraction (11). Although it has been recognized for some
time that the plasma membrane can form localized submicron scale
regions with different compositions and properties, the co-existence of
many types of membrane domains and their ability to coalesce into
larger structures is just beginning to be appreciated (1, 2, 15). Even
detergent-resistant ordered domains (i.e. rafts) have been
found to be heterogeneous in their composition and properties,
including detergent sensitivity (14-16, 50, 51). For migratory cells,
both the anterior (lamellipod) and posterior (uropod) of cells have
been reported to be composed of subtypes of rafts that apparently
coalesce to form micron-sized domains (see Fig.
8 and Refs. 11, 15, 17, 27, 52). It is
likely that there are numerous types of submicron scale domains within
these larger regions at the front and rear of polarized neutrophils,
but the exact nature (size, composition, dynamics, etc.) of these
domains is unknown (2). Evidence to date supports a model in which cell
uropods are composed of CD44-rich microdomains that are tightly
anchored to the underlying cortical actin cytoskeleton, while lamellae
are composed of different types of domains, which mediate dynamic actin
reorganization, are on average more disordered, and are apparently
enriched in PIP2 (Fig. 8 and Refs. 1, 2, 11, 15, 17, 27,
50, 52-55).
|
We have used MCD depletion of cholesterol to alter the lipid
properties in neutrophils responding to fMLF. We have shown that
depleting plasma membrane cholesterol prevents
chemoattractant-stimulated actin polymerization, cell polarization, and
formation of large-scale domains (Figs. 1-3). Restoring cellular
cholesterol resulted in a significant restoration of these responses.
While M
CD treatment is often used with the intention of disrupting
cholesterol-rich rafts, it actually has profound effects on the
distribution and order of lipid probes that partition into both ordered
and disordered domains (13). For this reason, it is important to
interpret results with M
CD in the context of a more complex model of
membrane organization. For example, considering that raft-like domains are small but occupy a large fraction of the surface (11-13), it follows that a large fraction of the cell surface is in or near raft
boundaries (2). This view of membrane organization has implications for
interpreting the effects of cholesterol depletion; cholesterol
depletion reduces the raft boundary area and/or changes the properties
of the boundaries, as seen in model membrane studies and in studies of
cells labeled with fluorescent lipid analogues (13, 56). If boundary
regions are important for the recruitment/retention of certain
signaling molecules to non-raft regions of the plasma membrane, then
cholesterol depletion would affect these non-raft membrane proteins as
well as raft membrane proteins.
MCD treatment has been used to investigate the role of lipid
organization in signaling through G-protein-coupled receptors, including the bradykinin receptor (54) and the high-affinity receptor
for IgE, Fc
RI (28). We found that delivery of
2
integrins to the cell surface following stimulation of the
G-protein-coupled fMLF receptors was not inhibited significantly by
cholesterol depletion. We also found that cholesterol depletion had
only a partial effect on the initial rise in
[Ca2+]i induced by 10 nM fMLF stimulation, and it had no significant effect on
the initial [Ca2+]i response when
100 nM fMLF was used. The sustained calcium response
induced by either dose of fMLF was strongly inhibited by cholesterol
depletion. Our findings using 100 nM fMLF are in complete
agreement with those reported by Barabe et al. (47). Barabe
et al. (47) showed that abrogation of the sustained response can be attributed to inhibition of capacitative calcium entry, while
release of calcium from intracellular stores is largely unaffected by
cholesterol depletion. We confirmed these findings (Fig. 6,
a-c) and showed that the effects of M
CD on cell
polarization cannot be attributed to the altered calcium response in
these cells (Fig. 6d). Instead, we suggest that
cholesterol-depleted neutrophils are unable to polarize because Rac is
not retained at the membrane.
Rac was transiently recruited to the plasma membrane in MCD treated
cells, and there was a brief period of stimulated actin polymerization
(Fig. 4, d"-f"). However, in the
cholesterol-depleted cells, actin polymerization did not persist and
Rac did not remain membrane associated. In agreement with this finding,
it has been reported that cholesterol depletion inhibits phorbol
ester-stimulated membrane recruitment of Rac, but not Rac activation,
in A431 cells (57). It remains to be resolved if interruption of Rac
retention caused inhibition of actin polymerization or vice
versa. In support of the former scenario, the Rho
GTPase-dependent asymmetric recruitment of the plekstrin
homology domain of AKT (protein kinase B) to the plasma membrane of
neutrophil-like HL60 cells can occur in the absence of actin
polymerization (22). In either case, the effects of cholesterol
depletion on stimulated actin polymerization and Rac retention provide
a mechanistic explanation for effects on both phagocytosis (18-20) and
migration (this report and Ref. 17).
There is little information on how the organization of outer leaflet
membrane components into microdomains might cause compartmentalization of lipids and proteins associated with the inner leaflet of the plasma
membrane. However, the functional coupling of inner and outer leaflet
membrane domain components has been well documented for the immune
receptors, FcRI and the T cell receptor (58-61). Indirect evidence
that microdomains extend over both leaflets of the plasma membrane
consists of co-patching experiments in whole cells. Aggregation of
outer leaflet-anchored (e.g. GPI-anchored proteins) or
transmembrane (e.g. CD44, Fc
RI) raft components causes
co-aggregation of inner membrane-associated signaling molecules (e.g. Lyn, Lck, Fyn, annexin II), and vice versa
in the case of CD44 and annexin II (10, 28, 50, 51, 62, 63).
Cholesterol depletion inhibits the coupling between transmembrane or
GPI-anchored receptors and membrane domain components at the
cytoplasmic face of the plasma membrane, apparently because it disrupts
both inner and outer leaflet membrane domain integrity; neither inner
leaflet (e.g. Lyn, Lck) nor transmembrane (e.g.
CD44) domain components partition into the low density fraction of
sucrose gradients after cholesterol depletion, and inner and outer
leaflet domain components no longer co-cluster on intact cells (11, 28,
50, 51).
Our data show that during polarization and migration Rac is recruited
stably to membrane regions that are excluded from CD44-containing raft-like domains. Stable incorporation of Rac to more disordered lipid
domains is consistent with model membrane and whole cell studies, which
showed that peptides with various types of isoprenyl-based lipid
anchors preferentially partition into liquid-disordered domains as
compared with co-existing liquid-ordered domains (64-66). MCD
treatment disrupts both raft and non-raft lipids (presumably on both
sides of the membrane), and it has profound effects on the extent of
boundary regions in the membrane (13). This change in lipid order
appears to alter the ability of Rac to be incorporated stably into the membrane.
We also show here that Rac is recruited stably to the membrane in the ML-7-treated cells (Fig. 7, b" and c"). In these cells Rac is found all around the cell, and, as noted previously, a circumferential lamellipod is formed (26). One interpretation of this is that coalescence of CD44-rich rafts at the rear of the cell acts to exclude disordered domains (and thus minimize boundaries), and in this way stable recruitment of Rac and lamellipod formation is prevented. In the ML-7 treated cells, raft-like regions that contain CD44 are kept dispersed and interspersed with detergent-soluble regions all around the cell (11). It is interesting to compare the phenotypes of ML-7-treated and cholesterol-depleted cells: neither can polarize, but for very different reasons. In the cholesterol-depleted cells, stable Rac recruitment and actin assembly do not occur, while in ML-7-treated cells they occur in an unrestrained fashion. In the cholesterol-depleted cells, membrane domains become coalesced into larger domains, whereas in ML-7-treated cells membrane domains are dispersed (11, 13, 56). These findings suggest that overall lipid organization (not simply the existence of rafts and non-rafts) is important for cell polarization. A description of the mechanism by which cholesterol depletion and ML-7 treatment affect Rac retention in the membrane will require a better overall understanding of lipid organization.
We cannot determine precisely which step in polarization is primarily
affected by cholesterol depletion. In fact, the ordering of steps in
the development of cell polarization is an unresolved issue, and there
is evidence that there are feedback cycles in this process that
complicate attempts at ordering the steps (67). Stable Rac recruitment
is the earliest step for which our experiments show a defect in
cholesterol-depleted cells. Activation of Rac has been placed both
upstream (22, 40) and downstream (68) of polyphosphoinositide
production, and it is possible that both effects are part of an
amplification loop. MCD inhibited stimulated phosphatidylinositol
turnover and Rac recruitment to membranes in A431 cells (54, 57).
Altering cholesterol levels in PMNs may similarly disrupt the
production and/or distribution of polyphosphoinositides, which have
been implicated in several aspects of cell polarization (22, 69-74)
and Rac-mediated actin polymerization (40, 68). Spatially restricting
polyphosphoinositide production to the front of polarized cells may be
necessary to achieve polarized actin polymerization and lamellipod
protrusion. In one model, Rac and polyphosphoinositides are part of a
signal amplification loop at the front of migrating cells (67), and a
hypothetical long-range inhibitor was proposed to prevent
chemoattractant receptor signaling and polyphosphoinositide build-up in
the trailing regions of motile cells. As an alternative, we suggest
that the large scale CD44-rich lipid domain that is established as
cells polarize can prevent lamellipod formation, perhaps by separating
out PIP2 microdomains and/or boundary regions that mediate
actin remodeling (55).
There is increasing evidence that compartmentalization of the plasma
membrane is important for establishing and maintaining cell polarity
(1, 11, 15, 17). In this report we provide the beginning of a
mechanistic explanation for the effects of cholesterol depletion on
cell polarization. Namely, we showed that cholesterol depletion
inhibits actin polymerization possibly because Rac retention in the
membrane is inhibited. At each stage of cell polarization, Rac
recruitment to the membrane occurs precisely in regions of actin
polymerization, which are always located outside of raft-like domains.
When segregation of raft-like domains to the cell rear is prevented by
inhibition of actin-myosin contraction, polarized recruitment of Rac to
the membrane and polarized actin polymerization are also prevented. In
our model, the plasma membranes of PMNs contain several types of
membrane microdomains, some of which are anchored to the actin
cytoskeleton via transmembrane proteins such as CD44 (1, 11, 52).
CD44-containing membrane domains would redistribute to the cell rear
during polarization and act to exclude certain signaling molecules
(e.g. Rac) or recruit others (1, 11). At the same time lipid
structures, possibly PIP2-enriched microdomains (55) and/or
boundary regions, that are favorable for Rac recruitment are
established at the front of the cell, and these mediate signals leading
to actin polymerization.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. David Holowka for many helpful discussions, and we thank Drs. William Muller, David Holowka, Melanie Brazil, and Robert J. Vasquez for critical reading of the manuscript. We also thank Leona Cohen-Gould for technical expertise and assistance with scanning electron microscopy. Flow cytometry, scanning electron microscopy, and confocal microscopy were performed within core facilities provided by Weill Medical College of Cornell University.
![]() |
FOOTNOTES |
---|
* This work was supported by National Institutes of Health Grants DK 27083 (to F. R. M.), GM34770 (to F. R. M.), and GM19078 (to L. M. P.) and The Atorvastatin Research Award from Pfizer/Parke Davis (to L. M. P.).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.
¶ Supported by the Minority Access to Research Careers Program.
To whom correspondence should be addressed: 1300 York Ave.,
New York, NY 10021. Tel.: 212-746-6405; Fax: 212-746-8875; E-mail: FRMaxfie@med.cornell.edu.
Published, JBC Papers in Press, January 8, 2003, DOI 10.1074/jbc.M212386200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
MCD, methyl-
-cyclodextrin;
fMLF, formyl Met-Leu-Phe;
MES, 4-morpholineethanesulfonic acid;
PIP2, phosphatidylinositol
bisphosphate;
PMN, polymorphonuclear neutrophil;
mAb, monoclonal
antibody;
TRITC, tetramethylrhodamine isothiocyanate.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. |
Pierini, L.,
and Maxfield, F.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
9471-9473 |
2. | Maxfield, F. R. (2002) Curr. Opin. Cell Biol. 14, 483-487[CrossRef][Medline] [Order article via Infotrieve] |
3. | Brown, D., and London, E. (1998) Annu. Rev. Cell Dev. Biol. 14, 111-136[CrossRef][Medline] [Order article via Infotrieve] |
4. | Brown, D., and London, E. (1998) J. Membr. Biol. 164, 103-114[CrossRef][Medline] [Order article via Infotrieve] |
5. | Simons, K., and Ikonen, E. (1997) Nature 387, 569-572[CrossRef][Medline] [Order article via Infotrieve] |
6. | Kurzchalia, T., and Parton, R. (1999) Curr. Opin. Cell Biol. 11, 424-431[CrossRef][Medline] [Order article via Infotrieve] |
7. | Brown, D., and Rose, J. (1992) Cell 68, 533-544[Medline] [Order article via Infotrieve] |
8. | Varma, R., and Mayor, S. (1998) Nature 394, 798-801[CrossRef][Medline] [Order article via Infotrieve] |
9. | Friedrichson, T., and Kurzchalia, T. (1998) Nature 394, 802-805[CrossRef][Medline] [Order article via Infotrieve] |
10. |
Harder, T.,
Scheiffele, P.,
Verkade, P.,
and Simons, K.
(1998)
J. Cell Biol.
141,
929-942 |
11. |
Seveau, S.,
Eddy, R.,
Maxfield, F.,
and Pierini, L.
(2001)
Mol. Biol. Cell
12,
3550-3562 |
12. | Mayor, S., and Maxfield, F. (1995) Mol. Biol. Cell 6, 929-944[Abstract] |
13. |
Hao, M.,
Mukherjee, S.,
and Maxfield, F.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
13072-13077 |
14. | Roper, K., Corbeil, D., and Huttner, W. (2000) Nat. Cell Biol. 2, 582-592[CrossRef][Medline] [Order article via Infotrieve] |
15. |
Gomez-Mouton, C.,
Abad, J.,
Mira, E.,
Lacalle, R.,
Gallardo, E.,
Jimenez-Baranda, S.,
Illa, I.,
Bernad, A.,
Manes, S.,
and Martinez-A, C.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
9642-9647 |
16. |
Madore, N.,
Smith, K.,
Graham, C.,
Jen, A.,
Brady, K.,
Hall, S.,
and Morris, R.
(1999)
EMBO J.
18,
6917-6926 |
17. |
Manes, S.,
Mira, E.,
Gomez-Mouton, C.,
Lacalle, R.,
Keller, P.,
Labrador, J.,
and Martinez-A, C.
(1999)
EMBO J.
18,
6211-6220 |
18. |
Shin, J.,
Gao, Z.,
and Abraham, S.
(2000)
Science
289,
785-788 |
19. |
Peyron, P.,
Bordier, C.,
N'Diaye, E.,
and Maridonneau-Parini, I.
(2000)
J. Immunol.
165,
5186-5191 |
20. |
Gatfield, J.,
and Pieters, J.
(2000)
Science
288,
1647-1650 |
21. | Fechheimer, M., and Zigmond, S. (1983) Cell Motil. 3, 349-361[Medline] [Order article via Infotrieve] |
22. |
Servant, G.,
Weiner, O.,
Herzmark, P.,
Balla, T.,
Sedat, J.,
and Bourne, H.
(2000)
Science
287,
1037-1040 |
23. |
Pierini, L.,
Lawson, M.,
Eddy, R.,
Hendey, B.,
and Maxfield, F.
(2000)
Blood
95,
2471-2480 |
24. | Pierini, L. M., and Maxfield, F. R. (1999) in Signaling Through Cell Adhesion Molecules, CRC Methods in Signal Transduction Series (Guan, J.-L., ed) , pp. 279-301, CRC Press, Boca Raton |
25. | Grynkiewicz, G., Poenie, M., and Tsien, R. (1985) J. Biol. Chem. 260, 3440-3450[Abstract] |
26. |
Eddy, R.,
Pierini, L.,
Matsumura, F.,
and Maxfield, F.
(2000)
J. Cell Sci.
113,
1287-1298 |
27. |
Sanchez-Madrid, F.,
and del Pozo, M. A.
(1999)
EMBO J.
18,
501-511 |
28. |
Sheets, E.,
Holowka, D.,
and Baird, B.
(1999)
J. Cell Biol.
145,
877-887 |
29. | Blanco-Colio, L., Villa, A., Ortego, M., Hernandez-Presa, M., Pascual, A., Plaza, J., and Egido, J. (2002) Atherosclerosis 161, 17-26[CrossRef][Medline] [Order article via Infotrieve] |
30. |
Danesh, F.,
Sadeghi, M.,
Amro, N.,
Philips, C.,
Zeng, L.,
Lin, S.,
Sahai, A.,
and Kanwar, Y.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
8301-8305 |
31. | Laufs, U., Kilter, H., Konkol, C., Wassmann, S., Bohm, M., and Nickenig, G. (2002) Cardiovasc. Res. 53, 911-920[CrossRef][Medline] [Order article via Infotrieve] |
32. |
Ledoux, S.,
Laouari, D.,
Essig, M.,
Runembert, I.,
Trugnan, G.,
Michel, J. B.,
and Friedlander, G.
(2002)
Circ. Res.
90,
420-427 |
33. |
Wong, B.,
Lumma, W.,
Smith, A.,
Sisko, J.,
Wright, S.,
and Cai, T.
(2001)
J. Leukoc. Biol.
69,
959-962 |
34. | Bellosta, S., Ferri, N., Bernini, F., Paoletti, R., and Corsini, A. (2000) Ann. Med. 32, 164-176[Medline] [Order article via Infotrieve] |
35. | Stancu, C., and Sima, A. (2001) J. Cell Mol. Med. 5, 378-387[Medline] [Order article via Infotrieve] |
36. |
Heyworth, P.,
Bohl, B.,
Bokoch, G.,
and Curnutte, J.
(1994)
J. Biol. Chem.
269,
30749-30752 |
37. |
Hall, A.
(1998)
Science
279,
509-514 |
38. | Hall, A. (1994) Annu. Rev. Cell Biol. 10, 31-54[CrossRef] |
39. | Nobes, C., and Hall, A. (1995) Cell 81, 53-62[Medline] [Order article via Infotrieve] |
40. |
Glogauer, M.,
Hartwig, J.,
and Stossel, T.
(2000)
J. Cell Biol.
150,
785-796 |
41. | Roberts, A., Kim, C., Zhen, L., Lowe, J., Kapur, R., Petryniak, B., Spaetti, A., Pollock, J., Borneo, J., Bradford, G., Atkinson, S., Dinauer, M., and Williams, D. (1999) Immunity 10, 183-196[Medline] [Order article via Infotrieve] |
42. | Cassimeris, L., McNeill, H., and Zigmond, S. (1990) J. Cell Biol. 110, 1067-1075[Abstract] |
43. | Weiner, O., Servant, G., Welch, M., Mitchison, T., Sedat, J., and Bourne, H. (1999) Nat. Cell Biol. 1, 75-81[CrossRef][Medline] [Order article via Infotrieve] |
44. |
Quinn, M.,
Evans, T.,
Loetterle, L.,
Jesaitis, A.,
and Bokoch, G.
(1993)
J. Biol. Chem.
268,
20983-20987 |
45. |
Kraynov, V.,
Chamberlain, C.,
Bokoch, G.,
Schwartz, M.,
Slabaugh, S.,
and Hahn, K.
(2000)
Science
290,
333-337 |
46. | Smith, C. (1990) Am. J. Respir. Cell Mol. Biol. 2, 487-489[Medline] [Order article via Infotrieve] |
47. |
Barabe, F.,
Pare, G.,
Fernandes, M. J. G.,
Bourgoin, S. G.,
and Naccache, P. H.
(2002)
J. Biol. Chem.
277,
13473-13478 |
48. | Marks, P. W., and Maxfield, F. R. (1990) J. Cell Biol. 110, 43-52[Abstract] |
49. | Marks, P. W., Hendey, B., and Maxfield, F. R. (1991) J. Cell Biol. 112, 149-158[Abstract] |
50. |
Oliferenko, S.,
Paiha, K.,
Harder, T.,
Gerke, V.,
Schwarzler, C.,
Schwarz, H.,
Beug, H.,
Gunthert, U.,
and Huber, L.
(1999)
J. Cell Biol.
146,
843-854 |
51. |
Schade, A.,
and Levine, A.
(2002)
J. Immunol.
168,
2233-2239 |
52. |
Millan, J.,
Montoya, M.,
Sancho, D.,
Sanchez-Madrid, F.,
and Alonso, M.
(2002)
Blood
99,
978-984 |
53. |
Laux, T.,
Fukami, K.,
Thelen, M.,
Golub, T.,
Frey, D.,
and Caroni, P.
(2000)
J. Cell Biol.
149,
1455-1472 |
54. |
Pike, L.,
and Miller, J.
(1998)
J. Biol. Chem.
273,
22298-22304 |
55. |
Caroni, P.
(2001)
EMBO J.
20,
4332-4336 |
56. |
Feigenson, G.,
and Buboltz, J.
(2001)
Biophys. J.
80,
2775-2788 |
57. |
Grimmer, S.,
Van, D. B.,
and Sandvig, K.
(2002)
J. Cell Sci.
115,
2953-2962 |
58. | Bi, K., and Altman, A. (2001) Semin. Immunol. 13, 139-146[CrossRef][Medline] [Order article via Infotrieve] |
59. | Sheets, E., Holowka, D., and Baird, B. (1999) Curr. Opin. Chem. Biol. 3, 95-99[CrossRef][Medline] [Order article via Infotrieve] |
60. | Janes, P., Ley, S., Magee, A., and Kabouridis, P. (2000) Semin. Immunol. 12, 23-34[CrossRef][Medline] [Order article via Infotrieve] |
61. | Holowka, D., and Baird, B. (2001) Semin. Immunol. 13, 99-105[CrossRef][Medline] [Order article via Infotrieve] |
62. |
Holowka, D.,
Sheets, E.,
and Baird, B.
(2000)
J. Cell Sci.
113,
1009-1019 |
63. |
Pyenta, P.,
Holowka, D.,
and Baird, B.
(2001)
Biophys. J.
80,
2120-2132 |
64. | Wang, T., Leventis, R., and Silvius, J. (2001) Biochemistry 40, 13031-13040[CrossRef][Medline] [Order article via Infotrieve] |
65. |
Melkonian, K.,
Ostermeyer, A.,
Chen, J.,
Roth, M.,
and Brown, D.
(1999)
J. Biol. Chem.
274,
3910-3917 |
66. |
Zacharias, D.,
Violin, J.,
Newton, A.,
and Tsien, R.
(2002)
Science
296,
913-916 |
67. | Rickert, P., Weiner, O., Wang, F., Bourne, H., and Servant, G. (2000) Trends Cell Biol. 10, 466-473[CrossRef][Medline] [Order article via Infotrieve] |
68. |
Ma, A.,
Metjian, A.,
Bagrodia, S.,
Taylor, S.,
and Abrams, C.
(1998)
Mol. Cell. Biol.
18,
4744-4751 |
69. |
Sasaki, T.,
Irie-Sasaki, J.,
Jones, R.,
Oliveira-dos-Santos, A.,
Stanford, W.,
Bolon, B.,
Wakeham, A.,
Itie, A.,
Bouchard, D.,
Kozieradzki, I.,
Joza, N.,
Mak, T.,
Ohashi, P.,
Suzuki, A.,
and Penninger, J.
(2000)
Science
287,
1040-1046 |
70. | Parent, C., Blacklock, B., Froehlich, W., Murphy, D., and Devreotes, P. (1998) Cell 95, 81-91[Medline] [Order article via Infotrieve] |
71. |
Parent, C.,
and Devreotes, P.
(1999)
Science
284,
765-770 |
72. |
Li, Z.,
Jiang, H.,
Xie, W.,
Zhang, Z.,
Smrcka, A.,
and Wu, D.
(2000)
Science
287,
1046-1049 |
73. |
Hirsch, E.,
Katanaev, V.,
Garlanda, C.,
Azzolino, O.,
Pirola, L.,
Silengo, L.,
Sozzani, S.,
Mantovani, A.,
Altruda, F.,
and Wymann, M.
(2000)
Science
287,
1049-1053 |
74. |
Jin, T.,
Zhang, N.,
Long, Y.,
Parent, C.,
and Devreotes, P.
(2000)
Science
287,
1034-1036 |