Shear stress induces a time- and position-dependent increase in
endothelial cell membrane fluidity
Peter J.
Butler1,2,
Gerard
Norwich1,
Sheldon
Weinbaum2, and
Shu
Chien1
1 The Whitaker Institute of Biomedical Engineering and
Department of Bioengineering, University of California, San Diego,
La Jolla, California 92093-0427; 2 Center for Biomedical
Engineering and Department of Mechanical Engineering, City College
of New York, New York, New York 10031
 |
ABSTRACT |
Blood flow-associated
shear stress may modulate cellular processes through its action on the
plasma membrane. We quantified the spatial and temporal aspects of the
effects of shear stress (
) on the lipid fluidity of
1,1'-dihexadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate [DiIC16(13)]-stained plasma membranes
of bovine aortic endothelial cells in a flow chamber. A confocal
microscope was used to determine the DiI diffusion coefficient
(D) by fluorescence recovery after photobleaching on cells
under static conditions, after a step-
of 10 or 20 dyn/cm2, and after the cessation of
. The method
allowed the measurements of D on the upstream and downstream
sides of the cell taken midway between the respective cell borders and
the nucleus. In <10 s after a step-
of 10 dyn/cm2,
D showed an upstream increase and a downstream decrease, and both changes disappeared rapidly. There was a secondary, larger increase in upstream D, which reached a peak at 7 min and decreased thereafter, despite the maintenance of
.
D returned to near control values within 5 s after
cessation of
. Downstream D showed little secondary
changes throughout the 10-min shearing, as well as after its cessation.
Further investigations into the early phase, with simultaneous
measurements of upstream and downstream D, confirmed that a
step-
of 10 dyn/cm2 elicited a rapid (5-s) but transient
increase in upstream D and a concurrent decrease in
downstream D, yielding a significant difference between the
two sites. A step-
of 20 dyn/cm2 caused D to
increase at both sites at 5 s, but by 30 s and 1 min the
upstream D became significantly higher than the downstream D. These results demonstrate shear-induced changes in
membrane fluidity that are time dependent and spatially heterogeneous. These changes in membrane fluidity may have important implications in
shear-induced membrane protein modulation.
mechanotransduction; membrane fluidity; fluorescence recovery after
photobleaching; cholesterol; alcohol
 |
INTRODUCTION |
BLOOD FLOW IMPARTS ON
VASCULAR endothelium a tangential shear stress, which initiates
cellular processes related to vessel wall homeostasis and
pathophysiology. While many cellular structures, including the cell
membrane (6, 7, 13), the cytoskeleton (33),
focal adhesions (10), integrins (31), and
glycocalyx (18), have been proposed to respond to shear
stress and initiate the cellular signaling processes, there have been
few experimental attempts to quantify the direct effects of shear
stress on these structures. Recent studies have suggested that at least
one site of mechanotransduction may be the cell membrane via
force-induced changes in fluidity (15, 18, 22).
We developed a new technique to determine the shear-induced changes in
membrane fluidity at specific locations on the cell at various time
intervals by measuring fluorescence recovery after photobleaching
(FRAP) with a Bio-Rad 1024 confocal microscope and Time-Course
software. FRAP was chosen over other methods of fluidity measurements,
e.g., fluorescence polarization, electron spin resonance, and
fluorescence correlation spectroscopy, because rapid measurements
(temporal resolution of 1-2 s) with high spatial resolution (~2
µm) can be made. A simplified one-dimensional (1-D) model was
developed to derive diffusion coefficients from the FRAP curves. The
diffusion coefficient (D) of a lipophilic fluorophore is a
quantitative measurement of membrane lipid fluidity (29). This novel approach of performing FRAP analysis of cells in a flow
chamber led to measurements of the temporal and spatial variations of
shear-induced changes in membrane-lipid fluidity, thus providing insights into mechanotransduction of shear stress by vascular endothelium.
 |
METHODS |
Cell cultures.
Bovine aortic endothelial cells (BAECs) were cultured in DMEM
supplemented with 2 mM L-glutamine, 50 U/ml penicillin, 50 mg/ml streptomycin, 1 mM sodium pyruvate, and 10% FCS. BAEC cultures were maintained at 37°C in a gas mixture of 95% air-5%
CO2. Cells used were from passages 6-17.
Serum-free DMEM containing 100 µM of the lipophilic probe,
1,1'-dihexadecyl-3,3,3',3'-tetramethylindocarbocyanine
perchlorate [DiIC16(3);
Molecular Probes, Eugene, OR] was used to stain the cell membrane. The
endothelial cells were washed with warm, serum-free medium, incubated
at 37°C in the staining solution for 10 min, and then washed three
times with serum-free medium and twice with complete medium. This
procedure resulted in a uniform staining of the membrane surface as
assessed by confocal microscopy (Fig. 1B, top). Fluidity
measurements were obtained within the first 20 min after
staining.

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Fig. 1.
A: confocal fluorescence recovery after photobleaching
(FRAP) experimental setup. A temperature-controlled flow chamber is
placed on the stage of a Bio-Rad 1024 confocal microscope. The
argon-ion laser (AL) performs the bleaching event and the krypton/argon
laser (KL) is used for monitoring. UR, upstream reservoir; OBJ,
microscope objective; FS, fast shutter; NF, neutral density filter; and
PMT, photomultiplier tube. B:
1,1-dihexadecyl-3,3,3'3'-tetramethylindocarbocyanine perchlorate
(DiI)-stained monolayer. Top: DiI-stained live cells;
bar = 10 µm. Bottom: immobile fluorophores (described
in text) with a representative bleach line; bar = 3 µm. The
x-coordinate originates at the midpoint of the bleached line
and is perpendicular to the line. Flow direction is parallel to the
bleached line.
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Shear stress apparatus.
Immediately after staining, the glass coverslip was assembled into a
parallel-plate flow chamber with dimensions of 2 mm × 5 cm × 100 µm. An infusion-withdrawal pump with adjustable rate (Harvard
Apparatus, Holliston, MA) was used to induce flow through the chamber
by withdrawing fluid from the downstream side of the flow chamber, with
the upstream side of the flow chamber connected to a syringe barrel
open to atmospheric pressure (Fig. 1A). A flow rate was
chosen to yield a shear stress of 10 or 20 dyn/cm2 using
the equation
= 6Qµ/wh2, where Q is
flow rate, µ is medium viscosity (0.007 poise), w is
channel width, and h is channel height. Temperature was
maintained at 37°C by a control loop consisting of a thermocouple, a
temperature controller (Omega, Stamford, CT ), and a heat gun (Master
Appliance, Racine, WI ) in the ultraviolet protection hood of the
confocal microscope. The medium was preequilibrated with 95% air-5%
CO2 overnight in the incubator. The upstream reservoir of
the flow apparatus was covered with mineral oil to maintain gas tension.
Confocal-FRAP system.
BAECs were sheared at 10 or 20 dyn/cm2 at 37°C using
complete medium (DMEM with FCS). FRAP analysis was performed using a
Bio-Rad 1024 confocal microscope. A thin line (0.902 µm wide)
parallel to the flow direction was bleached in the otherwise uniformly fluorescent membrane midway between the nuclear region and the cell
border by using repeated scans with all visible wavelengths from an
argon-ion laser at 10% power (5 scans at 2 ms/scan; Fig. 1B,
bottom). Diffusion-mediated recovery of DiI fluorescence into the
bleached region was monitored using the krypton/argon laser (1% power,
excitation = 568 nm,
emission = 585 nm) using line scans at 2 ms/scan for up to 2 s. Diffusion
coefficients were calculated by using the theory for 1-D diffusion with
a Gaussian initial condition for the bleach profile and a Gaussian
profile for the monitoring beam (21). The resulting
equation used to fit the recovery curves was
|
(1)
|
where t is time, f(t) is
fluorescence recovery over time t , fi = f(t < 0) is fluorescence before bleaching,
f0 = f(t = 0) is fluorescence
immediately after bleaching, f
= f(t
) is the asymptotic value of fluorescence recovery reached in infinite time, Lb is initial Gaussian
(e
2) half-width of the bleached line,
Lm is Gaussian half-width of the monitoring beam
(krypton/argon), and D is diffusion coefficient. The details
of the mathematical derivation of Eq. 1 are given in the
APPENDIX. From f0, f
, and
fi we calculated the fraction of fluorophore available for
diffusion,
, where
= (f
f0)/(fi
f0).
In separate experiments we measured the profiles of the bleached line
(using immobile fluorophores) and the monitoring beam [using a
variation of the point-scan technique (30)]. The Gaussian (e
2) half-width of the bleached line was 0.451 µm for a bleach depth similar to that induced on live cells, and the
Gaussian (e
2) half-width of the monitoring
beam was 0.316 µm. The bleached line extended in length beyond the
region of interest which was 1.7 µm in length. The slope of the
membrane at the bleach location ( ~0.3 µm/µm) was estimated from
three-dimensional reconstruction of confocal slices of DiI-stained
cells (Fig. 2B). The error in the FRAP measurement associated with this slope is evaluated in the APPENDIX.

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Fig. 2.
Experimental protocol. A: FRAP measurements were made
periodically on the upstream or the downstream side of the cell.
B: the cell shape was estimated from serial reconstruction
of confocal slices of the DiI-stained endothelial cells by assigning
height-specific pixel values to each serial slice and then
reconstructing the image (using LabVIEW image analysis software). The
result is a gray-scale image in which the pixel intensities are
proportional to the height above the coverslip of that part of the
cell. An example is seen in the inset (maximum height ~5
µm). The slope of the membrane is obtained from image analysis using
NIH image software and agrees with that from Barbee et al.
(4). , shear stress.
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Experimental protocols.
As positive controls, we tested the effects of a membrane fluidizing
agent, benzyl alcohol (BA), and a membrane rigidifying agent,
cholesterol, on endothelial cell membrane fluidity. Cells were cultured
to confluence in 2 × 2-cm wells. For BA experiments, DiI-stained
cells were incubated for 10 min in complete medium containing 30 mM BA
(Sigma Chemicals). For cholesterol experiments, cells were incubated
for 3 h in 0.1 mM cholesterol (cholesterol in
methyl-
-cyclodextrin; Sigma) in complete medium and stained with
DiI. For both experiments, up to three FRAP measurements were taken per
cell on multiple cells in multiple wells, all done within 3 min after staining.
To assess long-term changes in fluidity due to shear stress, periodic
FRAP measurements were taken at the same spot on the cell surface over
the course of the experiment (Fig. 2A). To assess spatial
variation in shear-induced membrane fluidity, FRAP measurements were
taken on the upstream side (as defined by the flow direction) or on the
downstream side of the cell. The upstream and downstream positions were
in the central portion of the membrane between the nucleus and the
upstream or downstream cell border, respectively.
Membrane fluidity measurements were made before the step-shear of 10 dyn/cm2, within 5 s after the step-shear, at 1-min
intervals while the shear stress was maintained, immediately after the
shear stress was stopped, and at 1-min intervals for 5 min thereafter.
For the very-low-shear control experiments, the same measurement
protocol was used but the shear stress was 0.1 dyn/cm2.
In additional experiments to assess early fluidity changes,
simultaneous measurements of upstream and downstream membrane fluidity
were taken at early time points (5 s, 10 s, 30 s, 1 min, and
2 min) on cells subjected to a step-shear of 10 or 20 dyn/cm2. Because the line scan was rapid (2 ms/scan),
virtually simultaneous upstream and downstream measurements could be
accomplished by bringing both regions into the field of view and
selecting two regions of interest along the scan line for fluorescence monitoring.
Data and statistical analysis.
D, f0 , and f
were evaluated by
fitting Eq. 1 to the raw recovery data using a
Levenberg-Marquardt nonlinear least-squares regression with the aid of
SlideWrite software (Advanced Graphics Software, Encinitas, CA) or a
custom program written in LabVIEW programming language (Fig.
3; National Instruments, Austin, TX). The
diffusion coefficients for shear experiments were normalized by using
the preshear value (Dinit). These ratios
(D/Dinit) were averaged and expressed as
means ± SE. For statistical analysis that involved multiple
pairwise comparisons, ANOVA was performed using SigmaStat software
(SPSS, Chicago, IL). For those groups showing significance among
groups, the significance of differences between each experimental group
was assessed using Tukey's post hoc test. For analysis of differences
between simultaneous upstream and downstream fluidity measurements,
upstream and downstream measurements of D/Dinit
for each cell were paired and plotted as means ± SE, and a paired
t-test was used to assess significant differences at each
time point. P
0.05 was considered to be significant. In
addition, 95% confidence intervals were computed for
D/Dinit for each group and each time point to
assess differences from initial D/Dinit values.
All values presented in the text and figures are means ± SE.

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Fig. 3.
Representative FRAP curve with curve fit. Data were
normalized to the average prebleach fluorescence, and postbleach
recovery was fitted with Eq. 1. Diffusion coefficient
(D) = 5.34 × 10 9 cm2/s.
f, fluorescence.
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RESULTS |
Positive controls.
Table 1 shows the means ± SE values
of the DiI diffusion coefficients for control cells (no treatment),
BA-treated cells, and cholesterol-treated cells (all without shear).
Incubation of cells in 30 mM BA for 10 min resulted in a significant
increase in lateral mobility of DiI in the apical membrane of BAECs
over the control values. Conversely, incubation of cells in the
rigidifying agent, cholesterol (0.1 mM for 3 h), resulted in a
significant reduction in the DiI diffusion coefficient relative to
controls. The mobile fraction
was nearly 100% for control cells,
and
decreased slightly to 90% for BA-treated cells and to a
greater extent (70%) for cholesterol-treated cells.
Effects of shear stress on membrane fluidity.
For control experiments, repeated FRAP measurements resulted in a
gradual decrease in D/Dinit (
38% over 15 min;
Fig. 4). Following the application of a
step-shear stress of 10 dyn/cm2,
D/Dinit measurements on the downstream side of
the cell resulted in an abrupt decrease in fluidity followed by
essentially the same time course as in the control experiments.
Measurements on the upstream side of the cell, however, showed that
D/Dinit rapidly increased after step-shear,
returned immediately to control values, and then began to rise by 3 min
and became significantly higher than control at 7 min
(D/Dinit = 2.01 ± 0.47) after the
application of step-shear. Thereafter, the
D/Dinit value decreased with time, despite the
maintenance of shear stress, but remained significantly elevated over
control for the 10-min duration of shear. On cessation of shear stress,
D/Dinit returned to near control values. While the earliest changes in upstream and downstream
D/Dinit were not significantly different
from control values, they were significantly different from each other
as assessed by ANOVA and a Tukey's post hoc pairwise comparison. The
mobile fraction of cells subjected to shear, as for the control cells,
was ~100% in all cases.

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Fig. 4.
Long-term shear-induced changes in fluidity. Increases in
fluidity due to the application of a step-shear of 10 dyn/cm2 at time 0 occurred only on the upstream
side of the cell. Measurements on upstream and downstream parts of
cells were performed on different cells from different experiments.
*Significant difference between upstream and control values.
#Significant difference between upstream and downstream
values (P 0.05).
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To investigate the possible early changes in fluidity, we performed
additional FRAP measurements at 5 s, 10 s, 30 s, 1 min, and 2 min on the upstream and downstream sides of the cell after the
application of step-shears of 10 or 20 dyn/cm2. In these
experiments, fluorescence recovery was measured simultaneously in the
upstream and downstream parts of the cell by putting both areas in the
field of view for nearly simultaneous measurements. Consistent with the
first set of experiments, a step-shear of 10 dyn/cm2 caused
a slight increase in D (D/Dinit=
1.15 ± 0.13) at 5 s, which was significantly higher than the
downstream value (D/Dinit= 0.88 ± 0.08;
Fig. 5A). These shear-induced
differences in D vanished by 10 s. A step-shear stress
of 20 dyn/cm2 elicited an early increase (5 s) in membrane
fluidity that was greater on the upstream
(D/Dinit = 1.45 ± 0.13) than
downstream side (D/Dinit = 1.16 ± 0.09; Fig. 5B), although this difference was not
significant. On the downstream side of the cell, the initial increase
rapidly disappeared to control levels by 10 s while the upstream
diffusion coefficient remained elevated for up to 1 min. The upstream
D/Dinit was significantly different from the
downstream D/Dinit at the 30-s and 1-min time
points (Fig. 5B). This upstream D/Dinit returned to initial values by 2 min
(Fig. 5B).

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Fig. 5.
Early phase shear-induced changes in fluidity.
Simultaneous upstream and downstream FRAP measurements were made and
paired for each cell. Error bars on the D/Dinit
vs. time graphs are the mean D/Dinit ± SE.
*Significant difference between upstream and downstream values as
assessed by a paired t-test (P 0.05).
#Significant difference from 1 as assessed by 95%
confidence intervals. A: upstream and downstream
D/Dinit values at 0, 5, 10, 30, 60, and 120 s after the application of a step-shear of 10 dyn/cm2
(n = 9). B: upstream and downstream
D/Dinit values at 0, 5, 10, 30, 60, and 120 s after the application of a step-shear of 20 dyn/cm2
(n = 8).
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DISCUSSION |
In the present study we demonstrate that a confocal laser scanning
microscope can be used to perform simultaneous, multipoint FRAP
measurements with three-dimensional spatial specificity. We confirmed
the utility of this system through rigorous comparisons with more
specialized FRAP systems (see discussion below) and by showing that BA
and cholesterol cause an increase and decrease, respectively, of
endothelial cell membrane fluidity in terms of DiI diffusion
coefficient D. We used this system to measure the time
course and spatial distribution of shear-induced changes in membrane
fluidity. The main findings of the present study are as follows:
1) a shear stress of 10 dyn/cm2 was sufficient
to induce both early (5 s) and delayed (>5 min) increases in
D; 2) this moderate shear of 10 dyn/cm2 elicited an increase and decrease in up- and
downstream D, respectively, with the change being rapid and
transient; 3) a higher shear stress of 20 dyn/cm2 caused increases in both up- and downstream
D, with the upstream increase being greater and more
sustained than the downstream increase; and 4) the upstream
increases in D resulting from a step-shear of 20 dyn/cm2 were sustained for a longer period than those
resulting from 10 dyn/cm2.
Shear stress initiates many cellular processes ranging from immediate
changes in ion conductance and G protein activation (seconds) to
alterations in cytoskeletal organization and gene expression (minutes
to hours) (9). The most proximal events mediating this
mechanotransduction have not been clearly established and may occur via
multiple structures. Furthermore, long-term effects of shear stress may
be transduced by signaling pathways and cellular structures different
from those mediating the immediate events. One logical candidate for a
mechanotransducer is the cell membrane because of its proximity to the
flowing blood. It was the goal of the present study to measure the
direct effects of shear stress on the cell membrane fluidity, a
property that has been shown to be strongly correlated with cell
functions [by modifying membrane protein diffusion (1)
and function (11)]. Membrane fluidity has been proposed
as a modulator of shear-related cellular processes (11,
18), and, recently, Haidekker et al. (17) published
results on an early increase in membrane fluidity due to shear stress.
To our knowledge, the present study represents the first experimental
quantification of the temporal (short-term and long-term) and spatial
(upstream vs. downstream) aspects of shear-induced increases in
fluidity on the apical plasma membrane.
To assess diffusion of unbleached fluorophore into a bleached line, we
solved the 1-D diffusion equation (2, 20). The effects of
convection of fluorophores via lipid flow or of cell deformation in the
direction of flow on the FRAP measurements were neglected, because the
lines were made parallel to the flow direction. The membrane where the
FRAP measurement was made was considered flat and devoid of
corrugations and/or microvilli (see APPENDIX for estimation
of the error associated with the assumption of a flat membrane). To
measure the initial bleach geometry, we performed the bleaching
experiments on fluorophores made immobile by drying the stained cells
and then returning them to culture medium. We measured the beam shape
using a variation of the point scan method (30) by
scanning a laser past a small stationary (0.17-µm diameter)
fluorescent latex microsphere (Molecular Probes) and then fitting the
fluorescent profile with a Gaussian curve. Alignment of bleaching and
monitoring lasers was verified each day by projecting the beams through
a target-labeled prism mounted on the microscope nose piece. Finally,
we compared the ability and accuracy of our model to assess
D with that of Stolpen et al. (32), who
bleached with an extended elliptical beam to approximate a line bleach.
For the same D, f0/fi,
f
/fi, Lb, and
Lm, fluorescence recovery curves generated by
our model agreed with those generated by Stolpen et al. within 4%.
While 46 terms were summed in their model for this comparison, our
model does not require any summation and hence is convenient for curve
fitting using standard curve-fitting software.
We report that both physiologically moderate (10 dyn/cm2)
and high (20 dyn/cm2) shear stress elicits a significant
immediate (5 s) increase in membrane fluidity as measured by FRAP. This
increase subsided by 10 s for 10 dyn/cm2 and by 2 min
for 20 dyn/cm2. The transient nature of the response
suggests that the early increase in membrane fluidity is in response to
the sudden onset of shear and that this response is dissipated.
Recently, Haidekker et al. (17), using a molecular rotor,
9-(dicyanovinyl)-julolidine (DCVJ), to assess membrane fluidity, also
showed an increase in fluidity by 5 s after step-shear, but there
the increase in fluidity was found to be sustained. The reason for the
differences in persistence of this early change is not clear at this
time but may be due to differences in cell types (human umbilical vein
endothelial cells vs. BAECs) or perfusion medium (Hanks' balanced salt
solution vs. DMEM-FCS), or methods of measurement (DCVJ
fluorescence vs. FRAP). The DCVJ measurement represents fluidity
averaged over the entire cell culture, whereas our FRAP measurement was
localized to specific subcellular regions on the cell membrane observed under confocal microscopy. In any event, the present data support, as
did Haidekker et al., the suggestion that membrane fluidity may play a
role in modulating some of the earliest events known to be related to
the mechotransduction of shear stress (e.g., G protein hydrolysis, ion
channel conductance).
Simultaneous upstream and downstream measurements of DiI diffusion
revealed a spatial distribution of the effects of shear stress on the
membrane, and the magnitude and persistence of the change were related
to the level of shear stress. A shear stress of 10 dyn/cm2
caused a rapid (5 s) increase and decrease in DiI diffusion on the
upstream and downstream portions of the membrane, respectively, and
both returned to initial values by 10 s. A step-shear of 20 dyn/cm2, however, caused increases in DiI diffusion on both
the upstream and downstream parts of the cell, with the upstream
increase persisting for 1 min and the downstream increase falling to
initial values by 10 s. These differences in spatial distribution
and persistence of the shear-induced increases in fluidity may provide
insights into how the cell can sense differences in shear magnitude.
This novel finding that the increase in membrane fluidity is
predominantly found in the upstream side of the cell correlates well
with the location of positive shear stress gradient distributions, which were computed by Barbee et al. (5), but not with the absolute shear stress distributions. Using measured cell topography and
computational fluid dynamics, Barbee et al. showed that shear stress is
symmetrically distributed on the upstream and downstream side of the
cell, whereas positive temporal shear stress gradients are concentrated
on the upstream side of the cell. Hence our observation that shear
stress induced a differential spatial (upstream vs. downstream) change
in membrane fluidity suggests that shear stress gradients, in addition
to shear stress per se, may play an important role in modulating
membrane fluidity.
The results here, demonstrating a delayed (>5 min) increase in
membrane fluidity, also support the hypothesis that the membrane fluidity, as measured by DiI-FRAP, may play a role in modulating later
responses to shear stress. The only other study to investigate the
later perturbing effects of shear stress on the cell membrane was
performed by Berthiaume and Frangos (6), who used the
shear-induced incorporation of MC540 (a lipophilic dye) into the
endothelial cells to reflect an increase in membrane permeability. They
found a significant increase in the incorporation of the dye beginning at 5 min after shearing with medium 199 supplemented with 20% FCS. Our
results on the time course of fluidity changes with shear stress are in
excellent agreement with theirs on the shear-induced incorporation of
this lipophilic dye.
The causes of shear-induced increases in membrane fluidity remain
unclear. It is likely that fluid shear would cause a time-dependent cell deformation in the direction of flow, thus leading to temporally varying and spatially heterogeneous stresses in the cell membrane. These may, in turn, induce time- and position-dependent fluidity changes. In support of this hypothesis, Sato et al. (28)
showed that, when endothelial cells are suctioned with a small pipette, a portion of the cell exhibits an immediate elastic deformation followed by a slower viscous deformation. The time scales of these deformations agree well with the early and late increases in membrane fluidity shown in the present study. Wang et al. (34),
modeling cell deformation with shear stress, have suggested that shear causes the nuclear bulge to deform in the direction of flow. Such deformation may lead to increased tension in the upstream cell membrane. Experimental support of such cell deformation has been given
by Helmke et al. (19), who showed that intermediate
filaments close to the apical membrane are displaced in the direction
of flow within the first 3 min after the application of a step-shear stress. Finally, the link between membrane strain and membrane fluidity
is suggested by the increase in membrane fluidity of human fibroblasts
in response to hypotonic swelling (3).
Physiological processes in the cell may be caused by increases in
membrane fluidity. Prostacyclin production has been shown to be
enhanced by an increase in membrane fluidity (3). Hence, the increases in fluidity after a step-shear observed here may partially explain the prostaglandin-mediated vasodilation seen in our
recent study (8). The physiological implications of the
more sustained component of increases in membrane fluidity shown here
are suggested by a study in which cell apoptosis was caused by
agents that induced a sustained increase in membrane fluidity
(12).
The cytoskeleton may modulate membrane dynamics. There is evidence that
actin filaments remodel on the time scale of the later fluidity changes
seen in this study (23). Morita et al. (24) showed that a low shear stress of 5 dyn/cm2 could elicit
the depolymerization of F-actin to G-actin in as early as 5 min.
Although the shear-induced actin depolymerization is not expected to
alter membrane fluidity directly (29), it is likely to
alter cell deformation with flow (25). In an earlier study
from our laboratory, Galbraith et al. (14) noted that microtubule remodeling due to shear stress occurred preferentially in
the upstream side of the cell, and Hage Chahine et al.
(16) showed that microtubule disassembly increases
membrane fluidity of fibroblasts as measured with FRAP. Finally, Sato
et al. (27) noted that the cell surface was stiffer on the
upstream side of the cell after 6 h of flow and attributed this
polarization to localized stress fiber development. Together, these
studies suggest that the cytoskeleton has an intimate association with
the membrane and may modulate its functions via membrane
stabilization/destabilization cycles.
In summary, we have introduced a novel quantitative measurement of the
temporal changes in membrane fluidity and its spatial distribution in
response to shear stress. The time course and heterogeneous
distribution of the increase in fluidity with shear stress may be
related to calcium signaling, shear-induced phospholipid metabolism,
and cytoskeletal remodeling. Fluidity changes are likely to have a
significant effect on membrane proteins and their interactions
(1, 26).
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APPENDIX |
FRAP Model
The following model describes isotropic, 1-D diffusion into a
bleached line with a Gaussian profile initial condition. We first write
the 1-D diffusion equation
|
(A1)
|
where c is concentration of unbleached fluorophore, t
is time, x is coordinate perpendicular to the bleached line,
D is diffusion coefficient, and Lb is
Gaussian half-width of the bleach line. IC is the initial condition,
and BC is the boundary condition. The solution is (using
Fourier analysis)
|
(A2)
|
To obtain the fluorescence (f) from these concentration
profiles, we integrate the product of the concentration and the monitor beam intensity profile (Im) over the confocal
aperture distance (a). In other words
|
(A3)
|
where Im0 is the peak intensity and
Lm is the Gaussian beam radius for the
monitoring beam. The solution to Eq. A3 is
|
(A4)
|
To simplify Eq. A4 we solve for
f(t
) = f
and f(t = 0) = f0 and recognize that the error function,
erf(
), approaches 1. Because a (the confocal
aperture distance) is large, the erf term goes to 1 and Eq. A4 simplifies to
|
(A5)
|
which, when normalized to the initial fluorescence,
becomes Eq. 1 in the text.
The error associated with the assumption of a flat membrane can be
estimated by using the slope of the membrane, the length of the region
of interest for fluorescence measuring, and the depth of the confocal
field. If the membrane has a slope of 0.3 µm/µm (see Fig. 2 and
Ref. 4), and the region of interest (ROI) is 1.7 µm
long, then the left and right edges of the ROI will be out of the focal
plane by 0.25 µm. (Note that the bleached line is longer than the ROI
and, therefore, there is no diffusion of fluorophores from the upstream
and downstream edges of the ROI). The depth of the confocal field for a
×60 1.4 numerical aperture (NA) objective is ~0.61 µm
(14), or 0.3 µm above and below the center of the ROI.
The error arises from the broadening of the laser beam with distance
from the focal point and the consequent broadening of
Lb and Lm. From the
definition of numerical aperture, NA= 1.5 sin
(where 1.5 is the
refractive index of the immersion oil,
is the half angle subtended
by the laser beam, and NA = 1.4), we can estimate that the beam
will broaden by ~0.1 µm at 0.3 µm from the focal point. We then
compute the area increase for the bleaching and monitoring beams and
compute effective Lm and
Lb that yield the effective areas. These widths
are 0.476 µm for the bleaching beam and 0.340 µm for the monitoring
beam. Replacing Lb and
Lm with these values yields an ~12%
underestimation of D for control values and an ~10%
underestimation of the peak values of D. When normalized,
these errors yield an ~4% overestimation of the twofold increase
seen on the upstream side at 7 min after a step-shear stress of
10 dyn/cm2.
 |
ACKNOWLEDGEMENTS |
We thank Dr. Jeffrey H. Price, of the National Science
Foundation-Whitaker Quantitative Imaging and Confocal Microscopy
Resource at University of California San Diego for assistance in
confocal microscopy and valuable suggestions regarding FRAP, and we
thank Dr. Shunichi Usami for expert technical advice and assistance.
 |
FOOTNOTES |
This work was supported by National Heart, Lung, and Blood Institute
Grants HL-19454 and HL-43026.
P. J. Butler is a recipient of an National Institutes of Health
National Research Service Award.
Address for reprint requests and other correspondence: S. Chien, The Whitaker Institute of Biomedical Engineering and Department of Bioengineering, University of California, San Diego, 9500 Gilman Drive MC 0427, La Jolla, CA 92093-0427 (E-mail: shuchien{at}ucsd.edu; pbutler{at}be-research.ucsd.edu).
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.
Received 2 December 1999; accepted in final form 20 October 2000.
 |
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