Mechanical stress-induced
Ca2+ entry and
Cl
current in cultured
human aortic endothelial cells
Miki
Nakao,
Kyoichi
Ono,
Susumu
Fujisawa, and
Toshihiko
Iijima
Department of Pharmacology, Akita University School of Medicine,
Akita 010-8543, Japan
 |
ABSTRACT |
A fluid stream
through a microtube was applied to cultured human aortic endothelial
cells to investigate the endothelial responses of both the ionic
currents and intracellular Ca2+
concentration
([Ca2+]i)
to mechanical stimulation. The fluid stream induced an increase in
[Ca2+]i
that was dependent on both the flow rate and the extracellular Ca2+ concentration.
Gd3+ and niflumic acid inhibited
the fluid stream-induced increase in
[Ca2+]i,
whereas Ba2+ and
tetraethylammonium ion exhibited no effect. The fluid stream-induced [Ca2+]i
increase was accompanied by the activation of an inward current at
52.8 mV. The reversal potential of the fluid stream-induced current shifted to positive potentials when the external
Cl
concentration was
reduced but was not affected by variation of the external
Na+ concentration. During the
exposure to the fluid stream,
[Ca2+]i
was voltage dependent, i.e., depolarization decreased
[Ca2+]i.
We therefore conclude that the fluid stream-induced current is largely
carried by Cl
and that the
Cl
current may thus play a
role in modulating the Ca2+ influx
by altering the membrane potential of endothelial cells.
shear stress; stretch; calcium signaling
 |
INTRODUCTION |
VASCULAR ENDOTHELIAL CELLS lining the surface of blood
vessels are constantly exposed to various mechanical forces induced by
the blood pressure and pulsatile flow. The endothelial cell membrane
exhibits certain ion channel responses to mechanical forces, and the
resulting changes in membrane potential and/or intracellular
Ca2+ concentration
([Ca2+]i)
are involved in modulating a number of second messenger pathways such
as the production of endothelium-derived relaxing factor and
PGI2, the synthesis
of various proteins, and the regulation of gene expression (1, 11, 24,
30).
Forces acting on endothelial cells in vivo are generally classified
into at least three different categories (2, 17): shear stress acting
parallel to the surface of endothelial cells, transmural force due to
blood pressure, and membrane stretch as a consequence of pulsatile
vessel expansion. The endothelial responses to these mechanical forces,
particularly those of
[Ca2+]i
and ion channels, have been investigated based on the above classification. For example, a shear stress of only 0.7 dyn/cm2 has been reported to
activate a K+ current in bovine
aortic endothelial cells (BAEC) grown on the inner surface of glass
capillary tubes (27). The activation of the
K+ current results in
hyperpolarization that may facilitate
Ca2+ entry (12, 27). It remains
controversial, however, as to whether or not shear stress causes an
increase in
[Ca2+]i;
several groups have reported an obvious
[Ca2+]i
elevation using a parallel-plate flow chamber, whereas others have been
unable to obtain such responses (4, 31, 34). When a physiological
solution was blown onto endothelial cells, a
Ca2+-permeable cation channel was
activated and thus increased
[Ca2+]i
(33). On the other hand, membrane stretch, applied as negative pressure
from the patch pipette, activated a cation channel in the pig aorta
(15) and porcine cerebral capillaries (29). Furthermore, the stretching
of cellular membranes increased
[Ca2+]i
in endothelial cells cultured on silicon membranes (21). The responses
of ion channels and
[Ca2+]i
are thus considered to differ depending on how mechanical forces are
applied to endothelial cells. These findings indicate that endothelial
responses may vary in the physiological environment, where endothelial
cells are subjected to complex mechanical forces, depending on the
regions of vasculature and hemodynamic conditions.
In this study, a stream of fluid was used to apply mechanical stress to
cultured human aortic endothelial cells (HAEC). Although the method may
not cause pure shear stress and also likely includes other mechanical
forces, it enables us to monitor
[Ca2+]i
and the concomitant changes in ionic currents simultaneously. It is
thus a useful method for examining the cellular mechanisms underlying
functional linkage between Ca2+
transient and ionic currents. We show herein that the fluid stream facilitates the Ca2+ influx and
activates the Cl
current.
In addition, the possible role of the
Cl
current in regulating
the Ca2+ entry is also discussed.
 |
MATERIALS AND METHODS |
Culture of endothelial cells.
HAEC at passage
3 or
4 were purchased from Clonetics (San
Diego, CA) and Kurabo (Osaka, Japan). The cells from Clonetics were grown in culture medium (endothelial basal medium, Clonetics) supplemented with 2% fetal bovine serum, bovine brain extract protein
contents (12 µg/ml), human recombinant epidermal growth factor (10 ng/ml), 1 µg/ml hydrocortisone, 50 µg/ml gentamicin, and 50 ng/ml
amphotericin B in 5%
CO2-containing air at 37°C. The culture medium used for the cells from Kurabo (Humedia-EB2, Kurabo)
was similar to that described above, except that 5 ng/ml human
recombinant fibroblast growth factor B and 10 µg/ml heparin were
added instead of bovine brain extract protein contents. The culture
medium was exchanged every 48 h until a subconfluent growth stage was
obtained. The cells were detached by exposure to 0.025% trypsin in a
Ca2+- and
Mg2+-free solution containing
0.01% EDTA for ~180 s, diluted in the culture medium, and then
reseeded on coverslips (9 × 9 mm) coated with fibronectin
(Biomedical Technologies, Stoughton, MA) with a cell density of
~2,500 cells/cm2. We kept the
cells in culture for 1-4 days before use. No differences were
found in the experimental results obtained by using these two different
sources and types of culture media.
Solution and drugs.
The HEPES-buffered saline (HBS) used as the standard bath solution
contained (in mM) 136.9 NaCl, 5.4 KCl, 1.0 CaCl2, 1.0 MgCl2, 11.1 glucose, and 5.0 HEPES. The pH was adjusted to 7.4 with NaOH. In some experiments,
MgSO4 was used instead of
MgCl2, although no significant
difference was observed compared with the experiments using
MgCl2 in HBS. When the
extracellular Ca2+ concentration
([Ca2+]o)
varied, CaCl2 was either added to
HBS or simply omitted from HBS. The
low-Cl
solutions (17.4, 35, and 70 mM; see Figs. 5B, 9, and 10)
were prepared by replacing NaCl with equimolar NaOH plus
methanesulfonic acid. In some experiments, sodium isethionate was used
as a substitute for NaCl. The external solution containing
tetraethylammonium ion (TEA) was prepared by adding 20 mM TEA chloride
to HBS in place of NaCl (see Fig.
5D). The
high-K+ solution (142.3 mM; Fig.
5E) was prepared by replacing NaCl
with KCl in HBS (pH adjusted to 7.4 with KOH). The
K+-free solution was prepared by
replacing KCl with equimolar CsCl in HBS (see Figs. 8-10). In the
Na+-free solution, NaCl in HBS was
totally replaced with equimolar N-methyl-D-glucamine
plus HCl (see Fig. 8B).
The pipette solution for the nystatin perforated patch recording
contained (in mM) 120 potassium gluconate, 30 KCl, and 10 HEPES, and pH
was adjusted to 7.2 with KOH. Nystatin (Sigma Chemical) was dissolved
in methanol as a 10 mg/ml stock solution and added to external
solutions to obtain a final concentration of ~400 µg/ml. The
pipette solution for the conventional whole cell mode contained (in mM)
20 CsCl, 95 cesium aspartate, 5 Na2ATP, 5 MgCl2, 5 EGTA, and 5 HEPES
titrated to pH 7.2 with CsOH.
Fura 2-AM (Dojindo Laboratories, Kumamoto, Japan) and fura PE3-AM
(Texas Fluorescence Laboratory) were dissolved in DMSO as 1 µg/µl
stock solutions, respectively, and diluted in 1 ml HBS with 2 µl 10%
cremophor EL (Sigma Chemical). The final concentration of the
Ca2+ indicators was 5 µM.
Niflumic acid (Sigma Chemical) was dissolved in DMSO as a 100 mM stock
solution and diluted in HBS to a given concentration described in the
text (see Fig. 5A).
Apparatus for applying mechanical stress.
The method for applying the fluid stream was essentially the same as
that described by Schwarz et al. (32, 33) (see Fig. 1). In brief, a
polyethylene tube with an opening of 300 µm was placed parallel to
the bottom of the chamber at a distance of 300 µm from a single
endothelial cell. The pipette position was adjusted to obtain a
nonturbulent stream homogeneously distributed over the whole cell
surface. The flow of the solution was driven by gravity, and the flow
rate was controlled from 0 to 1.10 ml/min by varying the
hydrostatic pressure between 0 and 50 cmH2O.
We developed a semi-closed circuit apparatus to apply hydrostatic
pressure to the endothelial cells (see Fig.
3A). The cells on a coverslip were
mounted in an acrylic chamber (1.5 × 1.1 × 0.7 mm) filled
with HBS. The chamber was sealed tightly with an acrylic cover using
four screws made of stainless steel. The inlet of the chamber was
connected to a reservoir of HBS via a polyethylene tube, and the
internal pressure of the chamber was raised by compressed oxygen that
was applied to the reservoir. The outlet of the chamber was connected
both to a mercury barometer for monitoring the hydrostatic pressure
inside the chamber and to an effluent bottle via a valve for
controlling the flow rate of the perfusate. The flow rate was
constantly maintained at 2-3 ml/min during the experiments. Using
this apparatus, the hydrostatic pressure inside the chamber could be
increased up to 200 mmHg.
Fluorescence measurement of
[Ca2+]i.
The cells on coverslips were incubated either in HBS containing 5 µM
fura 2-AM for 30 min at 37°C or in HBS containing 5 µM fura PE3
for 60 min at 37°C. After treatment, the coverslips were rinsed
several times with HBS and incubated in HBS for 30 min at 37°C. The
coverslip was mounted in a chamber placed on an inverted microscope
(TMD-1SJ, Nikon, Tokyo, Japan) equipped with a ×40 fluorite
objective. The fluorescence measurements of
[Ca2+]i
from a single cell were performed using a spectrophotofluorometer (CAM-230, Japan Spectroscopic, Tokyo, Japan). The excitation wavelength alternated at 400 Hz between 340 and 380 nm during recording of emission fluorescence at 500 nm. The emitted light was directed through
a rectangular iris that limited the light collected to a single cell
under observation. The emission intensities at 340 and 380 nm
excitation (F340 and
F380) were monitored on a chart recorder (Thermal Arraycorder WR7700, Graphtec, Tokyo, Japan). F340 and
F380, together with the current
and voltage signals, were sampled every 1 s onto the hard disk of a
computer running Superscope software (version 1.1, GW Instruments). At
the end of every experiment, the background fluorescence was measured from the cell-free part of the coverslip, and these values were subtracted from the raw data. The
F340 and
F380 values, thus corrected, were
used to calculate the fluorescence ratio
(F340/F380).
Fura PE3, a derivative of fura 2, is known to be more resistant to
rapid leakage and compartmentalization than fura 2 and remained
responsive to changes in
[Ca2+]i
for hours. In the present study, however, all experiments were completed within 15 min, and no significant difference could be detected between the two indicators. It has been reported that the
absolute values of
[Ca2+]i
may be calculated based on the dissociation constants for
Ca2+ binding (224 nM for fura 2 and 290 nM for fura PE3 at 37°C; Refs. 6, 36) using the following
equation (6)
|
(1)
|
where
Kd is the
dissociation constant, R is
F340/F380
(the ratio of relative fluorescence),
Rmin and
Rmax are the
F340/F380 values measured in Ca2+-free (4 mM
EGTA) HBS and by the addition of 10 µM ionomycin to HBS,
respectively, and
Sf2/Sb2
is the ratio of fluorescence intensities measured at excitation
wavelength 380 nm in the Ca2+-free
and ionomycin-containing solutions. The basal
[Ca2+]i
thus calculated in either fura 2- or fura PE3-loaded HAEC ranged from
50 to 100 nM in normal HBS under our experimental conditions. However,
the absolute values of
[Ca2+]i
could not be calculated because the dissociation constant of the
indicators and Ca2+ in the cytosol
might be different from those measured in the absence of protein (13).
In the present experiments,
F340/F380 was thus used as a relative measurement of
[Ca2+]i.
Electrophysiology.
Single endothelial cells on coverslips were voltage clamped using
either the conventional whole cell technique (7) or the nystatin
perforated patch-clamp technique (9). The electrode resistance ranged
from 4 to 6 M
when electrodes were filled with the pipette solution.
The current-voltage
(I-V)
relationship was measured by voltage ramp pulses that were applied from
a holding potential of
52.8 mV to +60 and
140 mV with
change in voltage over time
(dV/dt)
of either ±0.2 or ±0.1 V/s. The current and voltage signals were digitized online at 500 Hz using pCLAMP software (version 6.0.3, Axon Instruments). The reference electrode was usually
HBS agar with an integral Hg-HgCl wire. In the experiments in which the
extracellular Cl
concentration
([Cl
]o)
was varied, a flowing 3 M KCl solution was used instead of HBS agar to
minimize any possible changes of the junction potential at the tip of
the reference electrode (18). The liquid junction potential between the
pipette solution and HBS was directly measured to be
12.8 mV on
the basis of the assumption that the junction potential at the
tip of a 3 M KCl electrode in these solutions was negligible. The
voltage values shown in the text were corrected accordingly.
The bath solution was warmed by the use of a water jacket around the
perfusing tube, so that the temperature of the chamber was kept at 35.5 ± 1.0°C. The flow rate of the bath perfusion was 2-3
ml/min.
Data analysis.
The Gd3+-induced suppression of
the fluid stream-induced
[Ca2+]i
increase was analyzed by the following equation
|
(2)
|
where
RGd represents the magnitude of
the decrease in
F340/F380
at various extracellular concentrations of
Gd3+
([Gd3+]o)
and Rcont is the amplitude of
F340/F380
induced by the fluid stream alone.
Imax indicates the maximum
inhibition, and n is the Hill coefficient.
The results are expressed as means ± SE;
n indicates the number of cells
examined. The statistical analysis was performed using Student's
paired t-test, and values of
P < 0.05 were considered to be
statistically significant.
 |
RESULTS |
Fluid stream-induced increase of
[Ca2+]i
in HAEC.
As shown in Fig. 1, a
polyethylene tube with an opening of 300 µm (inside diameter) was
placed parallel to the bottom of the recording chamber at a distance of
300 µm from a single endothelial cell, and the cell was exposed to a
fluid stream of HBS containing 1 mM
Ca2+. When the flow rate was
increased from 0 to 0.45 ml/min, the peak of the
[Ca2+]i
increase was observed at ~1 min. During the continuous presence of
the fluid stream,
[Ca2+]i
appeared to remain elevated for ~2 min and thereafter declined. A
further increase of the flow rate to 1.1 ml/min caused an initial peak
of
[Ca2+]i,
which thereafter decreased to a plateau level. A subsequent cessation
of the flow reversibly decreased
[Ca2+]i
toward the basal level. From an average of five cells,
F340/F380 at the plateau phase was 0.67 ± 0.04 in the control, 0.77 ± 0.04 at 0.45 ml/min, and 0.93 ± 0.03 at 1.10 ml/min. Regarding the initial peak of
[Ca2+]i,
its magnitude appeared to depend on the flow rate, i.e., the transient
elevation was more marked at the flow rate of 1.1 ml/min than at 0.45 ml/min. Furthermore, the initial peak was consistently observed if the
flow rate changed instantaneously from 0 to 0.45 or 1.1 ml/min (not
illustrated). However, such an instantaneous change of the flow rate
often caused a detachment of the cells from the coverslip, and the
experiments were thus interrupted. To avoid a loss of cells, we changed
the flow rate by manually opening or closing the valve of the perfusing
tube while confirming the attachment of the cells. As a result, a few
seconds were required to reach a new flow rate level, which may thus
have resulted in varying degrees of transient
[Ca2+]i
elevation among the different endothelial cells examined (Figs. 1, 2,
4-6, and 11). The transient nature of the
[Ca2+]i
increase could not be analyzed extensively in the present study. The
following experiments focused on the sustained elevation of [Ca2+]i.

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Fig. 1.
Top: experimental device for
application of fluid stream. Bottom:
fluid stream-induced increase in intracellular
Ca2+ concentration
([Ca2+]i)
in human aortic endothelial cells (HAEC). Time course of
[Ca2+]i
changes in response to fluid stream is shown. Cell was loaded with fura
PE3, and fluorescence ratio of emitted fluorescence at 340- and 380-nm
excitation wavelengths
(F340/F380)
is plotted against experimental time. Time course of application of
fluid stream (FS) of 0.45 and 1.10 ml/min is indicated above trace.
External solution contained 1 mM
Ca2+. Trace shown is
representative of 5 different experiments.
|
|
The effects of the fluid stream were examined in different
[Ca2+]o
(Fig.
2A). In
the absence of external Ca2+, no
significant change in
[Ca2+]i
was detected in response to the fluid stream. However, the flow-dependent increase of
[Ca2+]i
became evident when
[Ca2+]o
increased to 5 and 10 mM. The relationship between
[Ca2+]i
and the flow rate was measured at
[Ca2+]o
of 0, 1, 5, and 10 mM. At each
[Ca2+]o,
five cells were subjected to an increasing flow from 0 to 0.45 and 1.1 ml/min in a manner similar to that for Fig.
2A. The results are summarized in Fig.
2B. It is clearly shown that the fluid
stream increases
[Ca2+]i
in a manner that is both flow rate dependent and
[Ca2+]o
dependent. In the absence of external
Ca2+,
F340/F380
during the fluid stream of either 0.45 or 1.1 ml/min was not
significantly different from control (0.59 ± 0.03 in control, 0.63 ± 0.04 at 0.45 ml/min, and 0.63 ± 0.04 at 1.1 ml/min;
P > 0.1). These findings suggest
that Ca2+ entry across the plasma
membrane is involved in the fluid stream-induced rise in
[Ca2+]i.

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Fig. 2.
Flow rate-dependent and extracellular
Ca2+ concentration
([Ca2+]o)-dependent
changes of
[Ca2+]i
during fluid stream. A: time course of
change of
[Ca2+]i
during exposure to fluid stream. Cells were loaded with fura PE3. Each
trace was obtained from different cells and is representative of 5 cells at 0 mM (top), 5 mM
(middle), and 10 mM
(bottom)
[Ca2+]o.
Time course of application of fluid stream is indicated above each
record. B: relationship between flow
rate and
F340/F380.
Data are means ± SE (n = 5), and
no error bars are shown when SE is smaller than symbol.
|
|
For subsequent experiments conducted to examine the mechanism
underlying the fluid stream-induced increase in
[Ca2+]i,
it was necessary to obtain a substantial response with little interference from any background noise. In the following experiments, we therefore recorded the response to the fluid stream in HBS containing 5 mM Ca2+ at 1.1 ml/min, with which almost a maximal response was expected.
Effect of hydrostatic pressure on HAEC.
Mechanical stress acting on the endothelial cells in a fluid-stream
method may include not only shear stress but also transmural pressure
and membrane stretch. To address whether transmural pressure by itself
causes an increase in
[Ca2+]i,
we intentionally changed the hydrostatic pressure in the recording chamber, using a semi-closed circuit apparatus (Fig.
3A).
Cells on a coverslip were mounted in an acrylic chamber filled with HBS, and then the chamber was sealed tightly. After the control [Ca2+]i
had been measured, the internal pressure of the chamber was raised to
80 mmHg with compressed oxygen that was applied to the reservoir for 3 min. Subsequently, the internal pressure was further increased to 140 mmHg for 3 min, and thereafter the pressure was released. Two
representative records with 1 mM (Fig.
3B,
top) and 10 mM (Fig.
3B,
bottom)
[Ca2+]o
are shown. With
[Ca2+]o
of 1 mM, hydrostatic pressure ranging from 0 to 140 mmHg produced virtually no effect on
[Ca2+]i.
F340/F380
was 0.63 ± 0.03 in the control, 0.63 ± 0.03 at 80 mmHg, and
0.62 ± 0.03 at 140 mmHg
(n = 4; not significant). When the
external solution contained 10 mM
Ca2+, however,
[Ca2+]i
was only slightly affected by increasing the hydrostatic pressure. F340/F380
was 0.64 ± 0.03 in the control, 0.68 ± 0.03 at 80 mmHg, and
0.69 ± 0.02 at 140 mmHg (n = 9).
The differences between the value in the control and those obtained at
80 or 140 mmHg were statistically significant
(P < 0.014). These findings suggest that the transmural pressure by itself may be able to increase [Ca2+]i.
However, the transmural pressure alone is not sufficient to account for
the fluid stream-induced rise in
[Ca2+]i,
and other types of mechanical stress must be involved.

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Fig. 3.
Effect of hydrostatic pressure on HAEC. A: schematic
diagram of chamber used to evaluate effects of hydrostatic pressure.
For explanation, see MATERIALS AND METHODS. B:
effects of hydrostatic pressure on
[Ca2+]i. Time course of
application of hydrostatic pressure is indicated above each record.
[Ca2+]o
was 1 mM (top) or 10 mM
(bottom). Traces are representative
of 4 and 9 different experiments with
[Ca2+]o
of 1 and 10 mM, respectively. Cells were loaded with fura PE3.
|
|
Inhibition of the
[Ca2+]i
response to fluid stream by various ion channel blockers.
To characterize the Ca2+ influx
pathway, the effects of various channel blockers were examined. Fig.
4A shows
the effect of Gd3+, known as a
nonselective cation channel blocker (21), on the sustained elevation of
[Ca2+]i.
The
[Ca2+]i
level was increased by the fluid stream at a rate of 1.1 ml/min with
[Ca2+]o
of 5 mM. During the maintained elevation of
[Ca2+]i,
10 µM Gd3+ was applied and
resulted in a suppression of the fluid stream-induced rise in
[Ca2+]i
in a reversible manner. To quantitatively analyze the inhibitory effect
of Gd3+, the magnitude of the
decrease in
F340/F380
was measured at various [Gd3+]o
and normalized to that of the control response in the absence of
Gd3+. The results in 18 cells are
summarized in Fig. 4B. The
IC50 value obtained using
Eq. 2
was 0.43 µM, with maximum inhibition of ~80%. It should be noted
that
F340/F380
does not linearly reflect [Ca2+]i
or the influx that is responsible for the fluid stream-induced [Ca2+]i
elevation. Therefore, the concentration-response curve cannot indicate
the stoichiometry for Gd3+ and its
binding sites. Nevertheless, the findings indicate that Ca2+ entry, which is sensitive to
micromolar concentrations of Gd3+,
contributes to the fluid stream-induced rise in
[Ca2+]i.

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Fig. 4.
Effects of Gd3+ on fluid
stream-induced change in
[Ca2+]i.
A: time course of changes in
F340/F380
in response to Gd3+. External
solution contained 5 mM Ca2+. Time
course of application of fluid stream and
Gd3+ is indicated above trace.
Cell was loaded with fura PE3. B:
concentration-dependent inhibition by
Gd3+ of fluid stream-stimulated
[Ca2+]i
elevation. Of 18 cells examined, 4 cells were loaded with fura PE3
( ) and 14 cells were subjected to fura 2 ( ). Smooth curve is best
fit of all data to Eq.
2 with an
IC50 of 0.43 µM.
|
|
The
[Ca2+]i
elevation was also inhibited by niflumic acid, which is known to block
not only various Cl
channels (38) but also cation channels (5) (Fig.
5A).
Niflumic acid (100 µM) decreased
F340/F380
by 73.4 ± 10.7% (n = 4). In Fig.
5B, the effect of reducing
[Cl
]o
was examined. The sustained elevation of
[Ca2+]i
was reversibly decreased by reducing
[Cl
]o
from 154.3 to 17.4 mM and substituting sulfonate ions. The inhibition
was 52.1 ± 3.9% (n = 5). When
isethionate ions were used for
Cl
substitution, the
average inhibition was 37.5 ± 4.8%
(n = 5).

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Fig. 5.
Effects of ion channel blockers on fluid stream-induced change in
[Ca2+]i.
Time course of application of fluid stream of 1.10 ml/min is indicated
by single bars. Double bars indicate time course of application of 100 µM niflumic acid (A), low external
Cl concentration
([Cl ]o;
17.4 mM; B), 0.5 mM Ba2+
(C), 20 mM tetraethylammonium
(TEA+; D), and high
K+ (142.3 mM;
E). Traces are representative of 4 (A,
C, and
D), 5 (B), or 8 (E) different experiments. External
solution contained 5 mM Ca2+.
Fluorescence indicator used is fura PE3 for
A and fura 2 for
B-E.
|
|
In a separate series of experiments, the possible contribution of
K+ conductance to regulation of
Ca2+ entry was examined. As shown
in Fig. 5C, 0.5 mM
Ba2+ did not appreciably inhibit
the elevation of
[Ca2+]i
(n = 4). Essentially similar
results were obtained in three of four cells when another
K+ channel blocker, 20 mM TEA, was
applied on top of the response to the fluid stream (Fig.
5D). In one of four cells, however, 20 mM TEA further increased
[Ca2+]i
by 43%, and the effect was reversible after washout of TEA (not
illustrated). On the other hand,
[Ca2+]i
was reversibly decreased by increasing the extracellular
K+ concentration
([K+]o;
Fig. 5E). The inhibition was 38.8 ± 7.1% at
[K+]o
of 142.3 mM (n = 8).
Fluid stream-induced membrane current and increase in
[Ca2+]i.
It is well known that in endothelial cells
[Ca2+]i
increases in response to hyperpolarization and decreases upon
depolarization (10, 16, 37). It is thus possible that the fluid
stream-induced change in
[Ca2+]i
or its inhibition by the various blockers described so far might be
caused by a secondary change in the driving force for Ca2+ entry. To investigate the
mechanisms underlying the fluid stream-induced Ca2+ entry, changes in the
membrane current associated with the increase of
[Ca2+]i
were recorded by the nystatin perforated patch-clamp technique. After
membrane perforation was established, the membrane potential was held
at
52.8 mV and the voltage ramp of ±0.2 V/s was applied every 20 s to obtain the
I-V
relationship (Fig. 6). In response to a
fluid stream (1.1 ml/min) containing 5 mM
Ca2+, the membrane current shifted
to a downward direction, accompanied by a gradual increase in
[Ca2+]i
(Fig. 6A). It was a consistent
finding that the activation of the current preceded the increase in
[Ca2+]i,
i.e., the current reached a peak within several tens of seconds and
[Ca2+]i
increased thereafter. It was also noted that the inward current gradually decreased after its activation despite the continuous presence of the fluid stream, thus indicating a desensitization of the
response. However, the extent of the decay varied from cell to cell. In
some cells, the current decreased as shown in Fig.
6A, whereas the current amplitude was
maintained at almost a constant level in others (see Fig. 9). In Fig.
6B, the
I-V
curves recorded before and during the application of the fluid stream are illustrated. In the control (Fig.
6B, trace
a), the
I-V
curve showed a marked inward rectification at potentials more negative than
80 mV, thus indicating the presence of inwardly rectifying K+ channels. The resting membrane
potential was
77.7 mV, as indicated by the zero-current
potential. During the exposure to the fluid stream (Fig.
6B, trace
b), the current deflection in response to ramp pulse
became larger and the membrane potential was depolarized to
36.6
mV. The fluid stream-induced current (trace
b
trace a)
was obtained by digitally subtracting the control current from the
current in the presence of the fluid stream and is plotted in Fig.
6C. The
I-V
curve of the fluid stream-induced current showed an outward
rectification with a reversal potential
(Erev) of
24.8 mV.

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Fig. 6.
Simultaneous recording of
[Ca2+]i
and membrane current
(Im) on fluid
stream. A: time course of changes in
Im
(top) and
[Ca2+]i
(bottom) measured by nystatin
perforated patch-clamp method with standard external and
K+-rich internal solution.
External solution contained 5 mM
Ca2+. Ramp pulses between
140 and +60 mV
(dV/dt = ±0.2 V/s) were applied every 20 s from holding potential of
52.8 mV.
Vm, membrane
potential. Bar indicates application of fluid stream with a flow rate
of 1.1 ml/min. Dashed line, 0 current level. Cell was loaded with fura
2. B: current-voltage
(I-V)
relationship in control (a) and
during fluid stream (b) measured at
times indicated by corresponding letters in
A. Arrows, zero-current potential
before and during application of fluid stream.
C:
I-V
curve of difference current obtained by digitally subtracting control
current (a in
B) from current recorded during
fluid stream (b in
B).
|
|
The same protocol was applied to nine cells, and
Erev of the fluid
stream-induced current averaged
28.3 ± 1.9 mV. In Fig. 7, the change in membrane potential in
these nine cells is shown. Under unstimulated conditions (in the
control), the membrane potential varied from
7.9 to
87.2
mV and thus correlated with previous findings (3, 8, 19). During the
exposure to the fluid stream, however, the membrane potential averaged
28.7 ± 2.6 mV and became close to the
Erev of the fluid
stream-induced current.

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Fig. 7.
Changes in Vm
induced by a fluid stream.
Vm was measured
as zero-current potential in
I-V
curve obtained by same protocol as for Fig. 6. Each symbol indicates a
different cell (n = 9). All cells were
loaded with fura 2.
|
|
Fluid stream-induced current.
Erev of about
28 mV cannot be explained by a change in
K+ selective conductances, but it
does suggest that Cl
and/or a nonselective cation current is responsible for the
fluid stream-induced current. The ion selectivity of the inward current was examined by the conventional whole cell patch-clamp method using
Cs+-rich internal and
K+-free external solutions (Fig.
8A).
Under the blockade of K+ currents,
the control
I-V
curve (Fig. 8A,
middle,
trace
a) showed a slightly outward
rectification with a zero-current potential of about
30 mV. The
fluid stream at 1.1 ml/min increased the current deflection in response
to the voltage ramp (Fig. 8A,
middle, trace
b). The configuration of the
I-V
curve of the fluid stream-induced current (Fig.
8A,
right), obtained by subtracting the
control current from the current obtained in the presence of fluid
stream, was qualitatively similar to that recorded in the presence of intracellular and extracellular K+
(see Fig. 6). The
Erev of the
difference current was
36.3 ± 0.9 mV
(n = 5). The fluid stream-induced
current could be recorded even when external
Na+ was completely removed (Fig.
8B). The average
Erev was
31.5 ± 2.5 mV (n = 6). The
value was slightly more positive than that obtained in the presence of
Na+.

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Fig. 8.
Fluid stream-induced current. Whole cell currents were recorded under
blockade of K+ currents using
Cs+-rich internal solution and
K+-free external solution.
A: results obtained in
Na+-containing external solution.
Left: original current traces.
Vertical deflections indicate response to ramp pulses, and horizontal
bar indicates application of fluid stream. Dashed lines, zero-current
level. Middle:
I-V
relationships measured at times indicated by corresponding letters in
traces at left.
Right:
I-V
curve of difference current obtained by subtracting control current
(a) from current obtained in
presence of fluid stream (b).
B: experiment was carried out with
Na+- and
K+-free external solutions. Same
explanation as for A. Traces shown in
A and
B are representative of 5 and 6 different experiments, respectively. In all experiments, external
solution contained 5 mM Ca2+ and
flow rate was 1.1 ml/min.
|
|
In contrast to the results of the
Na+ replacement study,
Erev of the fluid
stream-induced current was markedly influenced by varying
[Cl
]o.
As shown in Fig. 9, the current response to
the fluid stream was recorded with 70 (A), 35 (B), and 17.4 (C) mM
Cl
. The flow rate was 1.1 ml/min, and the bathing solution contained 5 mM
Ca2+. The
I-V
curves recorded before and during the application of the fluid stream
are plotted in Fig. 9,
A-C,
middle. The difference currents
between the control and the current in the presence of the fluid stream
(Fig. 9,
A-C,
bottom) show that the outward
component became smaller as
[Cl
]o
decreased and that
Erev shifted to
positive potentials.
Erev was
14.4 ± 1.4 mV with 70 mM
Cl
(n = 5),
7.8 ± 1.6 mV with
35 mM Cl
(n = 5), and
3.3 ± 1.3 mV with 17.4 mM Cl
(n = 5).

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Fig. 9.
Effect of Cl substitution
on fluid stream-induced current. Fluid stream-induced current was
recorded with
[Cl ]o
of 70 (A), 35 (B), and 17.4 (C) mM. In each
case, top shows original current trace
on chart recorder, middle shows
I-V
curves recorded before (a in
A, c
in B,
e in
C) and during
(b in
A, d
in B,
f in
C) application of fluid stream, and
bottom shows
I-V
curve of difference current (b a in
A, d c in
B, f e in
C). In all experiments, external
solutions contained 5 mM Ca2+ and
flow rate was 1.1 ml/min.
|
|
In Fig. 10,
Erev of the fluid
stream-induced current, obtained with an internal
Cl
concentration of 30 mM,
is plotted against
[Cl
]o
on a semi-logarithmic scale. Although the data for
[Cl
]o
of 17.4 and 35 mM obviously deviated from the linearity expected for a
Cl
-selective electrode from
the Nernst equation, these results indicate that the fluid
stream-induced current is carried mainly by
Cl
at physiological
[Cl
]o.

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Fig. 10.
Relationship between reversal potential
(Erev) and
[Cl ]o
(means ± SE of 5 cells). No error bars are shown when SE is smaller
than symbol. Dashed line, theoretical linear relation expected for a
Cl -selective electrode
calculated from Nernst equation
Erev = 60 · log([Cl ]o/30).
Solid line was drawn by eye.
|
|
Voltage-dependent
[Ca2+]i
change induced by fluid stream.
The simultaneous measurement of
[Ca2+]i
and the membrane currents was carried out by the nystatin perforated
patch-clamp method to examine the voltage-dependent change of
[Ca2+]i.
In the absence of the fluid stream,
[Ca2+]i
was affected neither by depolarization nor by hyperpolarization of the
membrane potential (Fig.
11A).
In contrast, during exposure to the fluid stream,
[Ca2+]i
decreased in response to depolarization from
52.8 to
2.8 mV in a reversible manner, although hyperpolarization to
102.8 mV only slightly increased
[Ca2+]i
(Fig. 11B). To quantitatively
analyze the voltage-dependent change in
[Ca2+]i,
the magnitude of the fluid stream-induced rise in
F340/F380 was measured at various membrane potentials and was normalized to that
obtained at
52.8 mV in the same cell. The data obtained from 14 cells are summarized in Fig. 11C. It
was demonstrated that [Ca2+]i
increased in response to hyperpolarization. Furthermore,
[Ca2+]i
appeared to be saturated at potentials more negative than
60 mV.

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Fig. 11.
Voltage-dependent
[Ca2+]i
change induced by fluid stream. A and
B:
Im and
[Ca2+]i
were measured by nystatin perforated patch-clamp method before
(A) and during
(B) application of fluid stream.
Top and
middle: command potential
(Vm) and
Im, respectively.
Vertical current deflection indicates response to ramp pulses.
Bottom:
F340/F380.
Cells were loaded with fura 2. External solution contained 5 mM
Ca2+.
C: voltage-dependent change in
F340/F380.
Amplitude of fluid stream-induced increase in
F340/F380
was measured and normalized to that obtained at 52.8 mV in same
cell. Number of experiments is indicated in parentheses (means ± SE, except that only mean is shown for 82.8 mV). All cells were
loaded with fura 2.
|
|
 |
DISCUSSION |
The major finding reported herein is that the fluid stream induces an
increase in
[Ca2+]i,
accompanied by the activation of a
Cl
current in HAEC. The
fluid stream as a method to provide mechanical stimuli, used in the
present study, can be a useful method for recording changes in both the
ionic currents and
[Ca2+]i
(33) but does not necessarily provide a well-defined shear stress (4,
34). Even though the tip of the perfusing tube was placed very close to
a single endothelial cell, the flow of the solution was observed to
radiate from the tip under the microscope. Thus the results must have
been influenced by a disturbed laminar flow. However, we consider that
the responses to the fluid stream may reflect those in certain
physiological hemodynamic environments in which endothelial cells are
subjected to complex mechanical forces, including shear stress,
membrane stretch, and transmural pressure.
The present experiments also provide some insights into the possible
role of the Cl
current in
regulating
[Ca2+]i
during the exposure to the fluid stream. First, the fluid
stream-stimulated [Ca2+]i
was not affected by K+ channel
blockers, TEA and Ba2+. It is thus
unlikely that the observed
[Ca2+]i
increase is driven by the activation of the
K+ current and the resulting
hyperpolarization. Second,
[Ca2+]i
was not affected by varying the membrane potential alone before the
application of the fluid stream. During exposure to the fluid stream,
however,
[Ca2+]i
decreased upon depolarization under the voltage-clamp conditions, thus
indicating that the fluid stream-induced
[Ca2+]i
increase is caused by the activation of
Ca2+ entry through the plasma
membrane. Third, the fluid stream caused either depolarization or
hyperpolarization depending on the membrane potentials in the control
(unstimulated) condition, i.e., depolarization occurred in those cells
that had a negative membrane potential in the control, whereas the
cells having less negative resting potentials were hyperpolarized in
response to the fluid stream (Fig. 7). As a result, the membrane
potential became stable near
30 mV. Fourth, the fluid
stream-induced increase of
[Ca2+]i
was suppressed by reducing
[Cl
]o.
This should be the case if the membrane potential is determined largely
by Cl
during the
stimulation of the fluid stream. A reduction of
[Cl
]o
would depolarize the membrane, thereby decreasing the driving force for
Ca2+ entry. Although the
inhibition of the fluid stream-induced
[Ca2+]i
changes due to a high-K+ solution
can be explained by depolarization, this finding does not necessarily
indicate that the membrane potential is determined by
K+ at physiological
[K+]o.
On the other hand, the observation of little or no effect of
K+ channel blockers suggests that,
at physiological
[K+]o,
the contribution of K+ conductance
is not larger than that of the
Cl
current.
On the basis of the above observations, it appears that the
Cl
current may play a
functional role in the fluid stream-induced [Ca2+]i
elevation. If the fluid stream facilitated
Ca2+ entry with little change in
the membrane conductance, then
[Ca2+]i
would have varied from cell to cell, depending on the membrane potential of the individual cells. In the
[Ca2+]i
measurements, however, we observed a consistent, quantitatively similar
increase in
[Ca2+]i
in response to the fluid stream in the various cells examined (for
example, see Fig. 5). We thus consider that the activation of the
Cl
current stabilizes the
membrane potential near the equilibrium potential for
Cl
(ECl)
and provides a constant driving force for
Ca2+ entry. Ono et al. (28) showed
the physiological
ECl to be about
28 mV when measured by the gramicidin perforated patch-clamp method. At this potential, a moderate increase of
[Ca2+]i
would occur, according to the
[Ca2+]i-membrane
potential relationship shown in Fig. 11. It should be noted that the
Cl
current decayed more or
less despite the continuous presence of the fluid stream. The degree of
the decay varied among the individual cells. Some cells showed a clear
decay (Fig. 6), whereas no decay was detected in others (Figs. 8 and
9). Nevertheless, because the input impedance of endothelial cells is
relatively high at a potential range between
70 and 0 mV (3,
35), the opening of only a small number of the channels might be
sufficient to cause a substantial change in the membrane potential.
Mechanical stress-induced
Ca2+ transient.
A fluid stream induced a transient increase in
[Ca2+]i,
followed by a sustained elevation. It is very unlikely that the fluid stream-induced rise in
[Ca2+]i
is due to transmural pressure, since the hydrostatic pressure in the
fluid stream method might be negligibly small and since an increase in
the hydrostatic pressure alone increased
[Ca2+]i
only slightly. Thus the fluid stream-induced
[Ca2+]i
appears to be caused by shear stress and/or membrane stretch, according to the usual classification of mechanical forces (see introduction). It remains, however, controversial as to whether or not
pure shear stress by itself induces an increase in
[Ca2+]i.
For example, the well-defined shear stress from 0.08 to 8 dyn/cm2 produced by the use of a
parallel-plate flow chamber caused only a transient increase in
[Ca2+]i
in BAEC, probably due to a release from the intracellular store sites
(34). On the other hand, Geiger et al. (4) reported both a transient
increase and a sustained elevation of
[Ca2+]i.
Schilling et al. (31) observed no change in
[Ca2+]i
in response to shear stress in the range between 2.4 and 25 dyn/cm2 in calf pulmonary artery
endothelial cells. Because of the inconsistency of the experimental
results among the various investigators, we cannot definitively
conclude that the increase in
[Ca2+]i
observed in the present study is caused by shear stress. Alternatively, shear stress of a much greater amplitude than that tried in the previous studies might be required for an increase in
[Ca2+]i.
Provided that a laminar flow is maintained at the tip of the polyethylene tube in the present study, flow rates of 1.10 ml/min would
have elicited fluid shear stress of 53.0 dyn/cm2 on the cell surface,
according to the geometry of the perfusing system (27).
On the other hand, Gd3+, a blocker
of nonselective cation channels, suppressed the fluid stream-simulated
rise in
[Ca2+]i.
This is in line with the previous finding that the stretch-induced increase of
[Ca2+]i
was inhibited with micromolar concentrations of
Gd3+ in human umbilical vein
endothelial cells (HUVEC) cultured on silicon membranes (21). Plausible
nonselective cation channels were found to be activated when the cell
membrane was directly distended by negative pressure in the
cell-attached patch-clamp condition (15, 29). It is thus possible that
the fluid stream causes, more or less, a deformation of the endothelial
cell membrane, which may lead to the activation of
Ca2+-permeable cation channels by
a common intracellular mechanism on the stretch-induced channel (14).
Although Ca2+-permeable channels
were expected to be active in the fluorescence measurements in the
present study, no such conductance could be detected by the patch-clamp
method. This finding is in sharp contrast to a clear activation of the
nonselective cation current observed in cultured HUVEC, despite the use
of an essentially similar method (33); the amplitude was ~300 pA at
60 mV. We consider that the amplitude of the
Ca2+-permeable current might be
too small to be resolved by the patch-clamp method and that the current
could have been masked by an apparent activation of the
Cl
current in the present
study. This view might be supported by the finding that the
Erev-[Cl
]o
relationship deviated from linearity with decreasing
[Cl
]o.
Nevertheless, such an undetectable current seems sufficient to produce
an increase in
[Ca2+]i.
For example, Oike et al. (25) reported that the
Ca2+ influx caused by depletion of
the store site was only 1 pA at
80 mV in cultured HUVEC.
Fluid stream-induced ionic currents.
The present results strongly suggest that the fluid stream-induced
current is carried by Cl
,
since the Erev of
about
30 mV was not largely affected by changes in either
[K+]o
or the external Na+ concentration
but was dependent on
[Cl
]o.
However, the measured
Erev of the fluid
stream-induced current was shown to be more positive than the
ECl (about
40 mV). In addition, the value was more positive in the
K+-containing solutions (about
28 mV) than in K+-free
solutions (about
36 mV in
K+-free,
Na+-containing solutions and about
32 mV in K+-free,
Na+-free solutions). The slightly
more positive values in the
K+-free,
Na+-free solutions than in the
K+-free,
Na+-containing solutions might be
explained by the junction potential at the tip of the reference
electrode (~4.5 mV when we measured the junction potential between
HBS and the Na+-free solution).
However, the difference between the
K+-containing solutions and
K+-free solutions cannot be
explained by any possible changes in K+ currents, since the
configuration of the
I-V
curves of the fluid stream-induced current was quite similar
irrespective of whether the solutions contained
K+. The possibility of including
an unknown junction potential in our measurements is not probable
because we obtained an
Erev of the
inwardly rectifying K+ current
using K+-containing solutions of
about
85 mV (Nakao, Ono, and Iijima, unpublished data). We have
no ready explanation for these results at present.
In BAEC grown in microcapillary tubes, fluid shear stress activated an
inwardly rectifying K+ channel
(12, 27). The K+ current developed
as a function of shear stress, with half-maximal activation of only 0.7 dyn/cm2 (27). In the present
study, we failed to detect either an increase in
K+ conductance or
hyperpolarization in response to the fluid stream, probably because a
well-defined shear stress was not achieved by the present method.
Although Cl
current is a
major component of fluid stream-induced current in HAEC, the precise
mechanisms linking the applied mechanical force to the activation of
the Cl
current are still
unknown. The fluid stream-induced
Cl
current could be
recorded even though the pipette solution contained 5 mM EGTA.
Furthermore, the bathing solutions in all experiments were isotonic.
These findings may indicate that the
Ca2+-activated
Cl
channels and/or
volume-sensitive Cl
channels, both of which were found to be present in vascular endothelial cells (22, 23), are not responsible for the fluid stream-stimulated Cl
current. However, we cannot completely rule out the possibility that
the Cl
current was
activated by a rise in
[Ca2+]i.
It might be argued that 5 mM EGTA is not sufficient to completely prevent an increase in
[Ca2+]i
and that a local Ca2+ increase
would have triggered the activation of a
Cl
current. If this were
the case, a delay between the onset of the
Cl
current and the
Ca2+ rise (Fig. 6) would thus be
expected because fura 2 measurements provide only an estimation
averaged over the entire cell. On the other hand, it might be possible
that deformation of the endothelial cell membrane, caused by the fluid
stream, activates the Cl
channels via a change in either the intracellular cytoskeleton or
related structures. Indeed, some interaction between the cytoskeleton and the channel was suggested to be involved in the activation of
swelling- or stretch-activated
Cl
channels in epithelial
cells (20), although this was not the case for the volume-sensitive
Cl
current of endothelial
cells (26).
In conclusion, we have, for the first time, demonstrated the
involvement of the Cl
current in the endothelial response to a fluid stream. We propose that
the activation of the Cl
current plays a role, at least partly, in modulating the
Ca2+ influx by altering the
membrane potential of endothelial cells.
 |
ACKNOWLEDGEMENTS |
We are grateful to Dr. B. Quinn for grammatical reading of the
manuscript and to Hitomi Meguro for excellent technical support in cell
culture. We also thank Mika Shiraiwa for secretarial service.
 |
FOOTNOTES |
This work was supported by grants from the Ministry of Education,
Science, Sports, and Culture (Japan) and in part by the Japan Heart
Foundation and by an IBM Japan research grant for 1996.
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. §1734 solely to indicate this fact.
Address for reprint requests: T. Iijima, Dept. of Pharmacology, Akita
University School of Medicine, 1-1-1 Hondoh, Akita 010-8543, Japan.
Received 26 January 1998; accepted in final form 6 October 1998.
 |
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