Departments of 1 Internal Medicine and 2 Biomedical Engineering, University of Iowa, Iowa City, Iowa 52242; and 3 Department of Biology and Physics, School of Science, Rensselaer Polytechnic Institute, Troy, New York 12180
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ABSTRACT |
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To determine how histamine regulates endothelial barrier function through an integrative cytoskeletal network, we mathematically modeled the resistance across an endothelial cell-covered electrode as a function of cell-cell, cell-matrix, and transcellular resistances. Based on this approach, histamine initiated a rapid decrease in transendothelial resistance predominantly through decreases in cell-cell resistance in confluent cultured human umbilical vein endothelial cells (HUVECs). Restoration of resistance was characterized by initially increasing cell-matrix resistance, with later increases in cell-cell resistance. Thus histamine disrupts barrier function by specifically disrupting cell-cell adhesion and restores barrier function in part through direct effects on cell-matrix adhesion. To validate the precision of our technique, histamine increased the resistance in subconfluent HUVECs in which there was no cell-cell contact. Exposure of confluent monolayers to an antibody against cadherin-5 caused a predominant decrease in cell-cell resistance, whereas the resistance was unaffected by the antibody to cadherin-5 in subconfluent cells. Furthermore, we observed an increase predominantly in cell-cell resistance in ECV304 cells that were transfected with a plasmid containing a glucocorticoid-inducible promoter controlling expression of E-cadherin. Transmission electron microscopy confirmed tens of nanometer displacements between adjacent cells at a time point in which histamine maximally decreased cell-cell resistance.
electrical resistance; cadherin; electron microscopy; modeling; transfection
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INTRODUCTION |
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HISTAMINE TRANSIENTLY increases microvascular permeability in vivo (12, 27). The transient changes in permeability are associated with a decrease followed by a restoration of cell adhesion in endothelial cells (3, 11, 14, 15, 17). Although these rapidly resolving changes in cell adhesion are clearly described, the mechanisms by which they occur are uncertain.
Changes in cell adhesion require changes in cell shape. These changes in cell shape imply a change in the balance of forces that round cells up and forces that keep cells in a spread configuration (10, 25). For example, when extracellular calcium is chelated, sites of cell adhesion dependent on calcium for binding are weakened. The resting centripetally directed tension in the cells is less opposed, and adjacent cells retract from each other (20, 22, 23).
Histamine causes a small and transient increase in myosin light chain phosphorylation in endothelial cells (17, 18), but the level of phosphorylation does not reach those that increase the activity of the actomyosin ATPase (19). Consistent with the level of myosin light chain phosphorylation, histamine decreased endothelial cell adhesion without causing a detectable increase in centripetal tension (18). These data imply that histamine changes barrier function by altering sites of adhesion, similar to the effects of chelating extracellular calcium rather than through contraction of the actin cytoskeleton. If histamine mediates direct effects on cell adhesion, then further identification of the sites at which histamine affects cell adhesion is necessary to better understand how inflammatory edema is regulated.
Identification of the adhesion sites and the mechanisms by which histamine disrupts and restores barrier function is complex because the cytoskeleton is an integrated three-dimensional network of filaments and adhesion proteins in which mechanical forces at cell-cell regions are mechanically coupled to cell-matrix regions (25). If the cytoskeleton is viewed as an integrative structure, histamine could disrupt cell-cell adhesion through two basic mechanisms. Activation of signal transduction events could decrease adhesion at cell-matrix sites and cause cell rounding, which in turn could result in a secondary or reactive loss in cell-cell adhesion. Alternatively, activation of signal transduction pathways may directly target cell-cell sites and cause a direct loss in cell-cell adhesion with a reactive loss in cell-matrix adhesion. Because there are distinct adhesion proteins at cell-cell and cell-matrix sites that could be affected differently by signaling pathways, it is important to identify the specific sites that histamine directly affects.
To understand how histamine directly affects endothelial cell-cell and cell-matrix adhesion, we modeled the electrical resistance across a confluent endothelial monolayer as a function of cell-cell resistance, cell-matrix resistance, and transcellular resistance. This approach was first introduced by Giaever and Keese (8) and Lo et al. (13) to model the resistance of unstimulated cultured fibroblasts and epithelial cells. We have now extended this approach to cultured endothelial cells to derive a real-time evaluation of cell-cell and cell-matrix adhesion in response to a physiological stimulus. We validated the precision of this approach by testing the model with specific biological interventions. We have correlated cell-cell resistance with ultrastructural details of cell-cell strain using electron microscopy. Using this model, we were able to demonstrate that histamine transiently disrupts barrier function by first disrupting cell-cell adhesion and later by increasing cell-matrix adhesion.
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MATERIALS AND METHODS |
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Materials. For these studies, we used a commercial
anti-cadherin-5 antibody (Transduction Laboratories; Lexington, KY)
that was prepared in a carrier buffer that did not contain glycerol and
sodium azide. The antibody recognizes an epitope at amino acids 26194
of the extracellular domain of cadherin-5.
Cell cultures. Cultured human umbilical vein endothelial cells (HUVECs) were prepared by collagenase treatment of freshly obtained human umbilical veins as described (3). Harvested primary cultures designated for cell adhesion assays were plated on 60-mm tissue culture plates that were coated with 100 µg/ml of fibronectin (Collaborative Research; Bedford, MA). All cells were cultured in medium 199 and supplemented with 20% heat-inactivated fetal calf serum, basal medium Eagle vitamins and amino acids, glucose (5 mM), glutamine (2 mM), penicillin (100 U/ml), and streptomycin (100 µg/ml). Cultures were identified as endothelial cells by their characteristic uniform morphology, uptake of acetylated low-density lipoprotein, and indirect immunofluorescent staining for factor Vlll.
Cell adhesion assay. Cell adhesion was measured using a
previously reported technique (6-8, 18, 24). In this system, referred to as electric cell-substrate impedance sensing, cells were
cultured on a small gold electrode (5 × 104 cm2) using culture
medium as the electrolyte, and barrier function was measured
dynamically by determining the electrical impedance of a cell-covered
electrode. The total impedance of the monolayer is composed of the
impedance between the ventral surface of the cell and the electrode,
the impedance between the cells, and the impedance of the cell
membranes dominated by the membrane capacitance (8). Membrane impedance
is very large, and thus most of the current flows under and between the
cells. Furthermore, membrane impedance is not expected to change on
addition of histamine. Thus measured changes in impedance represent
alterations primarily in cell-cell adhesion and/or cell-matrix adhesion.
A 1-V, 4,000-Hz alternating current signal was supplied through a
1-M resistor to approximate a constant-current source. Voltage and
phase data were measured with a SRS830 lock-in amplifier (Stanford
Research Systems) stored and processed with a personal computer. The
same computer also controlled the output of the amplifier and relay
switches to different electrodes. Critical features of the setup are
the current frequency of 4,000 Hz and the small area of the active
electrode (a surface area of 10
4
cm2).
For experiments, electrodes were coated with adsorbed fibronectin by exposure to a 100 µg/ml solution for 30 min. HUVECs were inoculated on electrodes at a confluent density of 105 cells/cm2. The in-phase voltage (proportional to the resistance) and the out-of-phase voltage (proportional to the capacitive reactance) were measured. We chose to express barrier integrity as a function of resistance, normalized to the initial value and expressed as a fractional change, because there were greater changes in resistance than impedance or reactance. Thus a 10% decline in resistance, for example, would represent a fractional resistance of 0.9. Electrical resistance increased after cells attached and covered the electrodes, and the resistance achieved a steady-state level by 24 h.
In some studies, we directly measured changes in cell-matrix adhesion by measuring the resistance in subconfluent cells that had not formed cell-cell contact. Microelectrodes were inoculated at a density of 2.5 × 104 cells/cm2. Microelectrodes were viewed under a microscope 24 h later, and experiments were conducted in wells in which cell-cell contact was not observed. The signal was amplified and typically reflected the change in resistance in less than 10 cells on the active electrode, which approximately represented one-tenth of the electrode area.
Mathematical model to resolve experimental resistance into changes
in cell-cell and cell-matrix adhesion. We used a previously derived
mathematical model to calculate specific cell-cell and cell-substrate
adhesion (5, 8, 13). The model assumes that current flows radially from
under the ventral surface of the cell and the electrode and escapes
between cells, with a minor amount going directly through the cell
membrane by capacitive coupling. In this model (Fig.
1, inset) the total impedance
across a cell-covered electrode is composed of the impedance between the ventral surface of the cell and the electrode (related to ), the
impedance between cells (indicated by Rb), the
transcellular impedance (Zm), and the impedance of
a naked electrode (Zn). For these calculations, the
cells are regarded as circular disks and Zm is
inversely related to membrane capacitance (Cm).
can also be defined as
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Because Zn and the cell-covered impedance
(Zc) are measured and Zm is the
impedance of the two cell membranes in series, , Rb and Cm are the only
adjustable parameters in the model. We first identified the values of
, Rb and Cm before the
addition of test agents. The
and Rb cannot be
explicitly solved because they are dependent on modified Bessel
functions and thus have to be derived by curve fitting. First,
Zc and Zn are measured as a
function of current frequency (
) in untreated cells. By iteratively choosing values for Cm,
Rb, and
we can arrive at the best fit to the
experimental impedance. Because resistance is also dependent on the
same values for Cm, Rb, and
as they are for impedance or reactance, Cm,
Rb, and
were measured by finding the best fit
of the calculated resistance to the experimental resistance. The
resistance was measured at 13 separate frequencies between 22 and
90,000 Hz and is expressed as the ratio of normalized resistance of
cell-covered to cell-free electrodes.
Real-time changes in cell-cell adhesion (Rb) and
cell-matrix adhesion () in response to histamine or other
interventions were determined at a single frequency of 4,000 Hz.
Zn and Zm were presumed
constant at 4,000 Hz, and Rb and
were assumed
to be the only variables that change during the course of the
experiment. Even if Rb(t) and
(t) changed significantly over time, by using repeated
linearization, the error using this procedure could be kept low. We
herein refer to this method as a dynamic optimization procedure.
Custom instrumentation software to perform the impedance measurements and the modeling analyses were first developed by Applied Biophysics. Custom software written in the LabVIEW programming language (National Instruments) was later developed to perform the same operations, and it achieved equivalent results.
Scanning electron microscopy. Confluent monolayers were exposed to test agents for the prescribed duration. The medium was rapidly removed, and the cells were fixed with 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer for 5 min at room temperature. Cells were subsequently rinsed for three 5-min intervals in 0.1 M sodium cacodylate. Cultured cells were then secondarily fixed with a solution containing 1% osmium tetroxide and 1% potassium ferrocyanide in 0.1 M sodium cacodylate for 30 min at room temperature. Monolayers were then subsequently washed with 0.1 M sodium cacodylate, followed by double-distilled water. Next, monolayers were subjected to serial dehydration in 25, 50, 75, 95, and finally 100% ethanol for 15 min each. Cells were then exposed to fresh 100% ethanol for 1 h. The tissue dish was cut into 2 × 2-cm sections with a hot scalpel under 100% ethanol and air-dried under hexamethyldisilazane-100% ethanol at a 1:1 volume ratio for 30 min and then repeated for 1 h. Dried specimens were mounted onto aluminum stubs that were painted with graphite. Stubs were coated with gold-palladium using an argon-based sputter. Images were taken with a Hitachi S-4000 electron microscope with an accelerating voltage from 1 to 5 kV at a magnification of ×1,000. Final prints were made on Polaroid 55 electron microscopy grade film.
Transmission electron microscopy. Studies were conducted on cultured cells grown on plastic dishes coated with 0.01% gelatin for 30 min, followed by 30 µg/ml of fibronectin for 30 min. Control monolayers and monolayers exposed to 10 µM histamine for 1 and 20 min were fixed with 2.5% glutaraldehyde in 0.1 M sodium cacodylate at 37°C for 10 min. Monolayers were washed with 0.1 M sodium cacodylate. Cells were then secondarily fixed for 30 min in a solution consisting of two parts 0.1 M sodium cacodylate, one part 4% osmium tetroxide in water and one part 6% potassium ferrocyanide in water. Cells were washed three times with 0.1 M sodium cacodylate buffer for 10 min each and once with distilled water for 10 min at room temperature. Cells were then exposed to 2.5% uranyl acetate at room temperature for 20 min. After the cells were rinsed with distilled water, monolayers were sequentially dehydrated in 25, 50, 75, 95, and 100% ethanol at room temperature for 30 min. Cells were embedded in resin consisting of one part eponate resin and one part 100% ethanol at room temperature for 1 h. The latter step was repeated. Finally, cells were subjected twice to 100% eponate for 2 h at room temperature. The resins were baked overnight at 70°C. Specimens were pivoted 90 degrees and cut into sections with a diamond knife followed by poststaining with Reynolds lead citrate. The images were then acquired at 75 kV in a Hitachi H7000 electron microscope. Prints were then developed on multigrade deluxe type 4 Ilford paper.
To measure the average separation distance between adjacent cells, we performed the following procedure. Electron micrographs were scanned with an Apple One Scanner using Ofoto Software, and image analysis was accomplished with National Institutes of Health Image. The entire length of the gap between two adjacent cells was subdivided into 10 equidistant regions. After pixels were calibrated into a nanometer scale, a plot of radiance intensities and distance measurements was obtained across a gap at a regional point of interest. The separation distance was determined by measuring the distance between the two radiance maxima.
Plasmid transfection. Immortalized cultured HUVECs, ECV304 cells, were transfected with a pLKneo vector containing the cDNA for E-cadherin (graciously provided by James Nelson) using Lipofectamine (as per the manufacturer's protocol) (1). The pLKneo vector contains a glucocorticoid-inducible promoter (9). Transfected cells were selected with G-418 (1 mg/ml) in medium 199 with 10% fetal bovine serum. Surviving cells were isolated, and clones were expanded in the presence of G-418. Clones were tested for expression of E-cadherin after stimulation with dexamethasone for 18 h using either Western blotting or fluorescent-activated cell sorting (FACS; University of Iowa Core FACS Facility) of cells prepared to examine surface expression of the E-cadherin.
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RESULTS |
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Modeling transendothelial resistance across endothelial
cell-covered electrode. Giaever and Keese (8) modeled the
resistance across a monolayer of fibroblasts as a circuit with three
components, the membrane capacitance (Cm), the
resistance created by the apposition of the ventral surface of the cell
to the matrix (), and the resistance created by the apposition of
the lateral surfaces of adjacent cells to each other
(Rb). We had to determine if these same components
contributed to the resistance across an endothelial cell monolayer, and
if so, how much each component contributed in an unperturbed state.
A model based on the assumption that there was no resistance due to
cell-cell and cell-matrix adhesion (Fig. 1A;
Rb and approaches 0, Cm = 1 µF/cm2, the Cm of a theoretical
smooth membrane) was a poor fit to the experimental resistance. Under
these circumstances, the calculated resistance, expressed as the
normalized ratio of the resistance of the cell-covered electrode to a
naked electrode, did not change between frequencies of 22 and 90,000 Hz. In contrast, the experimental normalized ratio was maximum at 4,000 Hz, consistent with previous reports (6-8).
The experimental results also did not fit a model based on the
assumption that the resistance was due to cell-matrix adhesion and not
to cell-cell adhesion (Fig. 1B; Rb
approaches 0, = 5.4
1/2 · cm, and
Cm = 1 µF/cm2).
Similarly, the experimental results also did not precisely fit a model
based on the assumption that resistance was predominantly due to
cell-cell adhesion (Fig. 1C; Rb = 2.2
· cm2,
approaches 0, Cm = 1 µF/cm2).
In contrast to these three models, a model in which resistance was
created by cell-cell adhesion and cell-matrix adhesion and membrane
capacitance (Fig. 1D; Rb = 0.9 · cm2,
= 4
1/2 · cm, and Cm
=1 µF/cm2) was a much better fit to the experimental
data. This model was further improved by increasing the
Cm to >3 µF/cm2 (Fig.
1E). This increase in cell Cm
is consistent with a folded or convoluted membrane with a greater
surface area than a simple smooth membrane.
Characterizing Rb, , and
Cm in monolayers of different ages of
confluency. We next compared the values of Rb,
, and Cm used to fit data from young and older
postconfluent cultured monolayers. Rb increased
from 1.205 ± 0.17
· cm2 in cells 1 day postconfluent to 2.28 ± 0.17
· cm2 in cells 5 days postconfluent.
decreased from 3.62 ± 0.15
1/2 · cm in cells 1 day postconfluent
to 3.15 ± 0.10
1/2 · cm in cells 5 days postconfluent. Cm also decreased from 3.57 ± 0.14 µF/cm2 in cells 1 day postconfluent to 3.09 ± 0.15 µF/cm2 in cells 5 days postconfluent. These data suggest
that mechanical forces were directed toward increased horizontal
tethering with older postconfluent monolayers.
Identifying dynamic changes in cell-cell and cell-matrix resistance
to predictable perturbations. To validate the ability of the model
to predict real-time changes in cell-cell and cell-matrix adhesion, we
measured dynamic changes in and Rb in response to a predictable perturbation in transendothelial resistance. We added
an antibody to cadherin-5 to cause a predictable decrease in cell-cell
adhesion. An antibody against cadherin-5, which disrupts homeotypic
cadherin binding, decreased the resistance by 15-20% across the
cell-covered electrode, whereas neither control serum nor the special
vehicle for the antibody altered the resistance (Fig.
2A). As anticipated, the
anti-cadherin-5 antibody decreased Rb, which
preceded decreases in the experimental resistance or in
(Fig.
2B). The anti-cadherin-5 antibody caused a small (10-15%) secondary decline in
, which lagged behind the change in
experimental resistance and which is consistent with a paradigm in
which a primary loss in cell-cell tethering induced a reactive stress that in turn strained cell-matrix adhesion sites. Thus the optimization procedure detected the expected response for
and
Rb.
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Because the model indicated that the antibody to cadherin-5 mediated a
primary decrease in cell-cell adhesion, we further validated the
analysis by measuring the resistance of subconfluent cells that had not
established cell-cell contact. Under these conditions
Rb is negligible, and thus changes in
transendothelial resistance would reflect primary changes in . As
expected, when we added the antibody to cadherin-5 to subconfluent
cells, we did not detect a decline in resistance (Fig. 2C).
However, we did detect a decline in resistance in these same
subconfluent cells in response to 2 mM EGTA, which would affect sites
of cell-matrix adhesion. The final resistance on exposure to EGTA
approached the resistance of a naked electrode (1,700
), which is
consistent with the notion that EGTA mediated a loss in cell-matrix
adhesion. Hence, the failure of the anti-cadherin-5 antibody to
decrease the resistance of subconfluent cells was not due to a
sensitivity limit of the system.
To validate our assumption that Cm does not significantly change, we measured Cm before and 15 min after the addition of the anti-cadherin-5 antibody at frequencies between 22 and 90,000 Hz (as described for Fig. 1). In a separate series of experiments, the Cm was 2.85 ± 0.05 µF/cm2 in untreated cells and 2.87 ± 0.35 µF/cm2 in cells exposed to the antibody.
To further validate the precision of our model, we also measured the
change in Rb and in confluent human endothelial
cells that were transfected with an E-cadherin expression vector that was under the control of a glucocorticoid-inducible promoter. We
observed an increase in resistance on exposure to dexamethasone in
cultured ECV304 cells that were transfected with a pLKneo vector that
contained the cDNA insert for E-cadherin (Fig.
3). The resistance increased from 8,396 ± 876
in control monolayers to 13,014 ± 582
after 18 h of
exposure to 1 µM dexamethasone. In contrast, the resistance did not
increase in response to dexamethasone when cultured monolayers were
transfected with the pLKneo vector without the E-cadherin insert (26).
As expected, exposure of dexamethasone to cultured ECV304 cells
mediated a greater effect on Rb than on
(Fig.
3).
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Localizing the effects of histamine to cell-cell and cell-matrix
sites. We next measured dynamic changes in and
Rb in response to histamine by measuring the
resistance and reactance at a single frequency. Figure
4 illustrates the effect of 10 µM
histamine on transendothelial resistance, reactance, and impedance.
Histamine caused a rapid but transitory decrease in
transendothelial resistance. Histamine decreased the transendothelial
resistance by 20-30% within 30-60 s. The resistance
recovered to basal levels within 3-5 min and subsequently
increased to above initial basal levels. In contrast, histamine
mediated smaller changes in transendothelial impedance and even smaller
changes in transendothelial reactance.
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When we used our dynamic optimization procedure to arrive at measured
values of and Rb, the decrease in resistance
occurred primarily at sites of cell-cell adhesion, especially in more
mature monolayers. Histamine initiated decreases in
and
Rb, with much greater fractional decreases in
Rb, which paralleled the decline in the
experimental resistance in monolayers 1 day postconfluent (Fig.
5A). In monolayers 5 days
postconfluent, histamine caused a very little decrease in
(Fig.
5B), with predominant decreases in Rb.
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When changes in Rb and , which occurred within
1 min after exposure to histamine, were examined with finer
time resolution, the decline in experimental resistance temporally
correlated with a decline in Rb (50-60%
change) (Fig. 5C). The decline in Rb
preceded the decline in experimental resistance. In contrast, the
decline in
(20-30% change) lagged behind the experimental
resistance, similar to the reactive decline in
observed in cells
exposed to the cadherin-5 antibody.
Although histamine decreased the resistance predominantly through
effects on cell-cell adhesion, histamine initially restored the
resistance at cell-matrix sites in cells 1 day postconfluent. We
detected increases in before increases in the experimental resistance (Fig. 5A). In contrast, the initial increases in
Rb lagged behind increases in the experimental
resistance. Hence, during the initial restoration phase, histamine
engaged cell-matrix sites and increased resistance, whereas early
restoration of cell-cell adhesion occurred later.
Whereas histamine engaged cell-matrix sites during the initial part of
the restoration phase, later increases in the resistance to above basal
levels were due primarily to increases in cell-cell adhesion. After 5 min, remained constant, whereas the experimental resistance
continued to increase in monolayers 1 day postconfluent (Fig.
5A). In contrast, changes in Rb paralleled
the increase in the experimental resistance after this time (Fig.
5A).
There was a closer association between the restoration phase of the
experimental resistance and increases in Rb in
cells 5 days postconfluent (Fig. 5B). There was very little
increase in .
To further support the idea that the primary effects of histamine on
the endothelium were to decrease cell-cell adhesion and to increase
cell-matrix adhesion, we measured the resistance in subconfluent cells
that had not established cell-cell contact. Because the contribution of
Rb would be negligible, the change in resistance
would represent the direct effect of histamine on cell-matrix adhesion.
An additional advantage of this approach is that it removes the
reactive effects of histamine on cell-matrix adhesion by eliminating
the mechanical coupling between cell-cell and cell-matrix sites. As
predicted from the model, we observed no significant decline in
resistance in response to histamine, which validates the detected
direct effect of histamine on cell-cell adhesion. Histamine did not
decrease resistance but instead increased the resistance (Fig.
6). Taken together, these data demonstrate that histamine disrupts barrier function by directly decreasing cell-cell tethering and inducing a reactive loss in cell-matrix adhesion. In contrast, histamine restores barrier function by first
increasing cell-matrix adhesion.
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Correlation between changes in Rb and
separation distance between adjacent cells. Because a decrease in
Rb is dependent on increased separation distance or
strain between adjacent cells, we asked whether functional changes in
Rb correlated with structural changes in cell-cell
separation. At ×1,000 magnification using scanning electron
microscopy (SEM), we did not observe the formation of discrete gaps
between adjacent cells in monolayers exposed to histamine when viewed
from the cell surface (Fig. 7). Cell-cell contact was preserved between control cells (Fig. 7A), cells
exposed to histamine for 1 min (Fig. 7B), and cells exposed to
histamine for 20 min (7C). In contrast, we observed large
generalized gaps between adjacent endothelial cells exposed to 2 mM
EGTA (Fig. 7D). As expected, we observed the formation of focal
gaps between adjacent endothelial cells on neutralization of cadherin-5
(Fig. 7E).
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Histamine-induced gap formation between adjacent cells could not be
detected when viewed from the surface using SEM because the cell
membranes of adjacent cells overlapped. To resolve gap formations,
cross sections were prepared by cutting the specimen in the y-z
axis and were subsequently viewed under transmission electron
microscopy (TEM; Fig. 8A).
Interestingly, we did not observe a complete full-thickness separation
between adjacent cells in monolayers exposed to histamine for 60 s
(Fig. 8B). Instead, we observed only focal and nonspecific
areas of separation between adjacent cells. Within 20 min of exposure
to histamine, cell-cell contact was restored toward its basal state
(Fig. 8C).
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The average separation distance increased from 18.9 ± 0.93 nm in control cells to 103 ± 14.9 nm in cells treated with histamine for 60 s (Fig. 9). In cells exposed to
histamine for 20 min, the average separation distance recovered to 24.8 ± 1.18 nm. Whereas the initial loss in Rb
correlated with the increase in the average separation distance, the
final separation distance approached but did not fully explain the
final increase in Rb (Fig. 9).
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DISCUSSION |
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Histamine initiates transitory increases in endothelial permeability in situ and in vitro (3, 18, 27). Under in situ conditions, increased permeability is associated with the development of small gaps between adjacent endothelial cells, and restored barrier function is associated with the reapposition of adjacent cells (12, 27). In recent years, several investigators have hypothesized that these gaps might develop as the result of an increase in centripetal tension developed by endothelial cells. However, we found that histamine decreases barrier function in confluent human endothelial cells without increasing centripetally directed tension (18). Rather than tearing the cells apart by an increase in centripetal tension, histamine appears to act by reducing tethering forces and utilizes the resting tension to create the development of small gaps. This paradigm is consistent in part with the gaps that develop between adjacent endothelial and epithelial cells when calcium-dependent adhesion sites are severed by chelating extracellular calcium (22, 23).
Resolving the sites at which histamine remodels tethering in cultured
endothelial cells is particularly complex for two reasons. First,
cell-cell and cell-matrix sites are mechanically coupled by an
intervening series of filamentous cytoskeleton that can transfer local
mechanical forces to distal sites (16, 25). This makes it difficult to
resolve primary stresses, which are consequent to the specific
targeting of signal transduction, from reactive forces, which are
consequent to expressed primary stresses. Second, changes in tethering
in response to edemagenic stimuli are rapid. We previously reported
rapid decreases in transendothelial resistance in response to histamine
that occur over a period of only 60 s (18). Thus to dynamically and
quantitatively localize primary stresses in response to edemagenic
stimuli, changes in cell-cell and cell-matrix adhesion need to be
separately measured but simultaneously compared. To begin to localize
the effects of histamine, we modeled the resistance across a
cell-covered electrode as a resistor and capacitor in series (5, 8). In
this model, the transendothelial resistance is a function of the
resistance created by cell-matrix adhesion (), the resistance created by cell-cell adhesion (Rb), and
Cm.
The hypothesis that the total resistance is dependent on ,
Rb and Cm in cultured
endothelial cells is supported by several observations. First, the
experimental resistance did not fit a model based on the assumption
that there was no cell-cell and cell-matrix adhesion (
and
Rb approaching zero). Second, the experimental
resistance did not fit models that were dependent on
or
Rb alone. Instead, the experimental resistance best
fit a model that was dependent on Rb,
, and a
cell Cm greater than 3 µF/cm2. The
higher Cm suggests that the plasma membrane was not
smooth but rather convoluted with great surface area. This is
consistent with the observations of Schmid-Schonbein et al. (21), who
reported numerous surface membrane folds in endothelial cells using
electron microscopy.
We observed quantitative differences in , Rb,
and Cm in confluent monolayers of different ages.
We observed higher values of Rb in cells 5 days
postconfluent compared with cells 1 day postconfluent. In contrast,
and Cm decreased in older cultures. Consistent with
our observation, Davies et al. (4) reported a decrease in focal
contacts in older confluent monolayers based on interference reflection
microscopy. Our data suggest that with increased age, there is
increased horizontal tethering and less vertical tethering.
Dynamic changes in cell-cell and cell-matrix adhesion were resolved by
the measured resistance and reactance determined at a single frequency
because resistance and reactance are dependent on the same solution of
and Rb. Inasmuch as there are an equal number
of measurements for the number of the unknown solutions for
and
Rb (resistance and reactance), solutions for
and Rb can be computed at multiple time points.
Based on this approach, the model indicated that histamine decreased
transendothelial resistance predominantly through decreases in
cell-cell adhesion. This was consistent in cells 1 and 5 days postconfluent.
We also identified a smaller decrease in in cells 1 day
postconfluent, which temporally lagged behind the experimental
resistance and Rb. These data alone suggest that
the loss in cell-matrix adhesion was mechanically coupled to the loss
in cell-cell adhesion or that the loss of cell-matrix adhesion occurred
independent of the loss of cell-cell adhesion but at a slower rate.
However, additional data support the former paradigm. Histamine did not decrease the resistance in subconfluent cells, a condition in which
cell-matrix adhesion is mechanically uncoupled from cell-cell adhesion.
Also, the cadherin antibody similarly mediated a secondary and smaller
decrease in
in confluent cells, whereas the same antibody had no
effect in subconfluent cells.
Although the model indicated that histamine decreased the resistance
primarily through a loss in cell-cell adhesion, restoration of the
resistance was initiated by cell-matrix-directed forces that increased
cell-matrix adhesion in monolayers 1 day postconfluent. The model
detected increases in before there were detectable increases in the
experimental resistance. In contrast, increases in
Rb lagged behind increases in the experimental resistance.
The lag between the initial increase in Rb and is particularly interesting because it suggests two possible paradigms
for the initial restoration of endothelial barrier function. In one paradigm, histamine independently restores cell-cell and cell-matrix sites, but it restores cell-matrix sites first. Alternatively, restoring cell-matrix sites may help to restore cell-cell adhesion. Remodeling of cell-matrix adhesion sites might alter the distribution of tensile stress on cell-cell adhesion sites, which would redirect forces to the periphery and facilitate restoration of cell-cell adhesion.
Although the optimization procedure indicated that the initial
restoration of the experimental resistance after histamine treatment
was due to increased cell-matrix adhesion, further increases in
resistance that developed after 3-5 min were due to
cell-cell-directed forces that increased cell-cell adhesion in cells 1 day postconfluent. At these later time points, was constant,
whereas changes in Rb paralleled changes in the
experimental resistance.
The relationship between and Rb to the
experimental resistance during the restoration phase was slightly
different in cells 5 days postconfluent. The experimental resistance
increased, with less of an increase in
. Instead, there was a
temporal correlation between the increases in experimental resistance
and increases in Rb. The differences between cells
1 and 5 days postconfluent suggest that the pattern of remodeling of
barrier function may be dependent on the age of the monolayer, which is
consistent with the observed differences in
,
Rb, and Cm between cells 1 and
5 days postconfluent.
We validated the ability of the model to measure dynamic changes in and Rb through a number of controlled studies.
First, we applied the optimization procedure to localize changes in
resistance in monolayers exposed to an antibody against cadherin-5.
Because the antibody disrupts homeotypic cadherin tethering, the model should detect a predominant decrease in Rb that
temporally parallels a decline in experimental resistance. This
paradigm was supported by our experimental results. Second,
dexamethasone-mediated expression of E-cadherin in ECV304 cells
resulted in predominant changes in cell-cell resistance. Third, the
primary changes in
and Rb detected in confluent
cells when exposed to histamine or the cadherin-5 antibody are
consistent with the observed responses in subconfluent cells exposed to
these same agents. Inasmuch as there was no cell-cell contact, the
resistance was dominated by
in subconfluent cells. According to our
model, histamine should increase resistance because of its effect on
and histamine should not decrease resistance because
Rb is negligible. Exposure to the cadherin-5
antibody should not affect resistance because homeotypic cadherin
binding was not established. As anticipated, these responses were
observed, further supporting the ability of the model to identify
site-specific changes in endothelial barrier function.
Although the decrease in Rb correlated with
increased separation distance between adjacent cells in response to
histamine, the measured separation distance did not fully explain the
final increase in Rb at later time points. The
separation distance only approached control levels ( ~6 nm) at
time points corresponding to values of Rb above
basal levels. There are several potential explanations for this
discrepancy. First, Rb is an approximated value of
cell-cell adhesion. The precision of Rb is
dependent on the initial assumptions of the model and the mathematical
approach used to generate the curve fit. The model assumes that the
cell is disk shaped, which departs from the typical cuboidal shape of
an endothelial cell. Our eventual goal is to use various constitutive equations to evaluate whether different geometries alter the
contribution of
and Rb to barrier function.
Also, we must assume that the Cm does not
significantly change or, if it did change, such changes would have a
minor influence on the interpretable effects of histamine on
and
Rb.
Second, adhesion assays were performed on gold, whereas TEM studies were performed on plastic. Thus minor differences in cell motility may be attributed to differential effects from different substrates.
Third, the method used to quantitate cell-cell separation distance may underestimate cell-cell impedance. These gaps were focal and not uniform. Histamine-mediated cell-cell gaps required the resolution of TEM to detect the tens of nanometers of displacement. Changes in Rb actually reflect three-dimensional displacements in cell-cell adhesion, whereas our measured cell-cell separation distance was based on a two-dimensional measurement. The electrical current may take a convoluted course between adjacent cells, which may not be fully appreciated because of the overlapping cell membranes.
Although Cm was not included in our evaluation of
the measured impedance at 4,000 Hz, the solutions for and
Rb in confluent cells were still predictive for the
cadherin interventions, validated by the subconfluent responses, and
paralleled changes in cell-cell separation distance. To include
Cm in the evaluation, at least three different
measurements of resistance and reactance are required. One approach
would be to measure resistance and reactance in real time at multiple
frequencies, but at this point, the data acquisition among the lock-in
amplifier, relay stations, and the computer would need to be restructured.
Although is directly related to Rc (cell
radius) and inversely related to the square root of h (the
average separation distance between the substrate and the ventral
surface of the cell), changes in Rc likely had
little effect on measured changes in
. There are several reasons
that support this notion. First, the scale of h is 1,000 times
smaller than Rc. Thus the factor that includes h has a much greater effect than Rc on the
measured
. Second, if histamine decreased
exclusively through
effects on Rc, then we should have observed greater
changes in Rc. Histamine decreased cell radius by
84 nm, which compared with the initial Rc
(~10-11 µm) represents only a 1% change. The
should have
decreased by only 1% rather than the observed decrease of 25%. Thus
very large changes in Rc would have to occur, which
would have been easily detected by SEM. Third, if large changes in
Rc occurred, the monolayer would unlikely have been
able to restore resistance to its basal level in such a short period of time.
Giaever and Keese (8) reported that Rc would have
to change 400 times more than h to achieve the same unit change
in . Based on their report in cultured fibroblasts, they argued that
should be more sensitive to changes in h unless there are
very large changes in Rc.
It is important to distinguish h, which was used in our model, from other reported measurements of cell-substrate separation. Davies et al. (4) reported a quantitative method using interference reflection microscopy to define focal contact adhesion. They defined focal contact sites as radiances associated with cell-substrate separation distances of 15-30 nm. Yet, cell matrix adhesion is also dependent on nonfocal contact adhesion or close contact adhesion, which is defined by the absence of focal contact proteins and is associated with cell-substrate distances of greater than 30 nm (2). The value of h used in our model represents an integration of the entire cell-substrate interface, which consists of both focal and nonfocal contact sites.
Consistent with the results of our analysis, Winter et al. (26) reported that histamine did not decrease transcellular resistance in ECV304 cells, which do not express cadherin-5 or E-cadherin in wild-type cells. Yet, they reported that histamine decreased resistance in cells transfected with a pLKneo vector expressing E-cadherin or cadherin-5. The effect of histamine on resistance in ECV304 cells transfected with these cadherin-expression vectors was similar to its effect on resistance in cultured HUVECs. Thus this report suggests that the cadherin complex may be an important target site to decrease cell-cell apposition in response to histamine.
In summary, fitting resistance measurements with a model of cell-electrode interactions indicates that histamine rapidly and transiently altered endothelial barrier function through sequential changes at cell-cell and cell-matrix sites. These changes in response to histamine were best modeled by a sequential decrease in cell-cell adhesion, then increased cell-matrix adhesion, and finally increased cell-cell adhesion. This model now facilitates identifying the specific molecules and mechanisms that regulate inflammatory edema formation.
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ACKNOWLEDGEMENTS |
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We thank Randy Nessler for technical assistance on the electron microscopy studies and Jeff VanEngelenhoven and Matthew Lorson for technical assistance on the electric cell-substrate impedance sensing studies.
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FOOTNOTES |
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Scanning and transmission electron microscopic studies were conducted in the Central Microscopy, Central Research Facility.
This work was supported by a University of Iowa College of Medicine Grant, an AHA Grant-in-Aid (A. B. Moy), and National Heart, Lung, and Blood Institute Grant HL-33540 (D. M. Shasby).
This work was conducted during A. B. Moy's tenure as a recipient of the American Heart Association (AHA) Clinician Scientist Award and a Clinical Investigator Award from the American Lung Association.
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 and other correspondence: A. B. Moy, Dept. of Internal Medicine, C-33 GH, Univ. of Iowa College of Medicine, Iowa City, IA 52242 (E-mail: alan-moy{at}uiowa.edu).
Received 23 April 1999; accepted in final form 6 December 1999.
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