Vol. 281, Issue 3, H1397-H1406, September 2001
Lipopolysaccharide reduces intercellular coupling in vitro and
arteriolar conducted response in vivo
Karel
Tyml1,3,
Xiaowei
Wang1,3,
Darcy
Lidington2,3, and
Yves
Ouellette2
1 A. C. Burton Laboratory and 2 Child Health
Research Institute, Lawson Health Research Institute, and
3 Department of Medical Biophysics, University of Western
Ontario, London, Ontario, Canada N6A 5C1
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ABSTRACT |
Our recent in vitro study
(Lidington et al. J Cell Physiol 185: 117-125,
2000) suggested that lipopolysaccharide (LPS) reduces communication
along blood vessels. The present investigation extended this study to
determine whether any effect of LPS and/or inflammatory cytokines
[tumor necrosis factor-
, interleukin (IL)-1
, and IL-6] on
endothelial cell coupling in vitro could also be demonstrated for an
arteriolar conducted response in vivo. Using an electrophysiological approach in monolayers of microvascular endothelial cells, we found
that LPS (10 µg/ml) but not these cytokines reduced intercellular conductance (ci) (an index of cell
communication) and that LPS together with these cytokines did not
further reduce ci. Also, ci was restored after LPS washout, and the
LPS-induced reduction was prevented by protein tyrosine kinase (PTK)
inhibitors (1.5 µM Tyr A9 and 10 nM PP-2). In our in vivo experiments
in arterioles of the mouse cremaster muscle, local electrical
stimulation evoked vasoconstriction that conducted along arterioles.
LPS in the muscle superfusate did not alter local vasoconstriction but
reduced the conducted response. Washout of LPS restored the conducted
response, whereas PTK inhibitors prevented the effect of LPS. On the
basis of a newly developed mathematical model, the LPS-induced
reduction in conducted response was predicted to reduce the arteriolar
ability to increase resistance to blood flow. We conclude that LPS can reduce communication in in vitro and in vivo systems comparably in a
reversible and tyrosine kinase-dependent manner. Based on literature
and present results, we suggest that LPS may compromise microvascular
hemodynamics at both the arteriolar responsiveness and the conduction levels.
endothelial cell monolayer; mouse cremaster muscle; tyrosine
kinase; mathematical model
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INTRODUCTION |
THE ABILITY OF THE
ARTERIOLE to change its diameter is a key determinant in
microvascular blood flow control. However, the response of arteriolar
wall cells to stimuli impinging directly on these cells and the
subsequent local diameter change may not necessarily alter the blood
flow. Kurjiaka and Segal (19) showed that a stimulus
causing only a local arteriolar dilation yielded no increase in blood
flow in the microvascular network fed by the stimulated arteriole.
However, a stimulus causing both local and conducted dilation did
elevate blood flow. Thus both the local responsiveness of blood vessel
wall cells and their ability to conduct responses along the blood
vessel length may be required for microvascular blood flow control
(19).
Sepsis, a systemic inflammatory response to a local infectious insult,
impairs vascular responsiveness and may lead to maldistribution of
blood flow in organs and eventually to organ dysfunction
(22). Although it is well known that sepsis reduces the
vasoconstrictive (17) and vasodilative (35)
responsiveness of arterioles, it is not known whether sepsis also
compromises arteriolar ability to conduct responses along the
arteriolar length. Our recent study demonstrated that
lipopolysaccharide (LPS), an initiating factor in sepsis, increased
intercellular resistance (ri) in cultured microvascular endothelial cell monolayers by about 60%
(24). This finding suggested that LPS could reduce
conduction and/or communication along blood vessels and implied that
sepsis could compromise microvascular blood flow control at the
arteriolar responsiveness as well as the conduction level.
Our recent finding of increased ri after
exposure to LPS could be questioned for its applicability to an in vivo
model; for example, it is not clear what reduction in conducted
arteriolar response may correspond to the measured 60% increase in
ri in the cell monolayer. Although a number of
reports have characterized the arteriolar-conducted response in several
animal models (8, 14, 18, 31, 32) and have addressed the
mechanism of conduction (3, 6, 9, 11, 29, 39), little is
known about the effect of any particular disease process on the
conducted response. Thus in the context of these reports, the main
objective of the present study was to determine whether the conducted
response can be modulated by agent(s) of the inflammatory process. We
tested whether any effect of LPS and/or inflammatory cytokines [tumor necrosis factor- (TNF-), interleukin (IL)-1, IL-6, or a
combination thereof] on endothelial cell coupling in vitro could also
be demonstrated for a conducted arteriolar response in vivo.
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METHODS |
Isolation and Culture of Microvascular Endothelial Cells
Rat microvascular endothelial cells (RMEC) were harvested from
the extensor digitorum longus muscle as we described previously (24, 37). Briefly, the muscle was enzymatically digested
and RMECs were separated from other cells by using Griffonia
simplicifolia lectin-coated magnetic beads and a magnetic
concentrator. Harvested RMECs were plated and grown on 12-mm diameter
glass coverslips in culture medium in a standard incubator. The culture
medium consisted of medium 199 (M199, GIBCO, Mississauga, ON, Canada) supplemented with fetal bovine serum (10%, GIBCO), endothelial growth
supplement (50 µg/ml, Collaborative Research; Bedford, MA), heparin
(5 U/ml, Leo Laboratories; Ajax, ON, Canada), L-glutamine (0.1 mg/ml, GIBCO) and antimycotic-antibiotic solution (10 µl/ml, GIBCO). Cells were periodically tested for markers of endothelial phenotype as previously described (37) and were used
between passages 5 and 15.
Measurement of ri and Intercellular Conductance
in Cell Monolayers
In RMEC monolayers, ri (
) was
determined by an electrophysiological approach and a Bessel function
mathematical model as detailed in our recent study (24).
Briefly, two cells of the monolayer were impaled by two
microelectrodes, each connected to an electrometer (Intra 767, WPI).
After reading a stable membrane potential (Em)
in each cell (Fig. 1), four to five
hyperpolarizing pulses of current (50 nA, 100- ms duration) were
injected into one cell and a change in Em
(
Em) was noted in the other cell. This
process was repeated three times for other cell pairs of the monolayer
at different microelectrode distances (d) (range 50-400
µm). The plot of Em versus log
(d) was fitted by a zero order Bessel function to determine
the ri of the monolayer (24). The
intercellular conductance, ci = 1/ri (Siemens), was used in the present study as
a measure of the spread of the injected current (an index of
coupling/communication) between the cells of the monolayer.
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Fig. 1.
Design of electrophysiological experiments in rat microvascular
endothelial cell (RMEC) monolayers. Two cells of the monolayer were
impaled by two microelectrodes separated by 50 µm, aiming to obtain a
stable membrane potential (Em) in each cell
(left, Em1 in one cell;
right, Em2 in the other cell). Then,
4-5 hyperpolarizing pulses were injected into the first cell and
the resulting hyperpolarization was noted in the other cell (change in
Em2). Then, electrode Em1
was lifted from the cell and inserted into another cell at a distance
of 100 µm from electrode Em2 (this electrode
remained inserted in its cell), pulses were injected into the new cell,
and the resulting hyperpolarization in Em2 was
noted. The same procedure was repeated for interelectrode distances of
200 and 400 µm. Changes in hyperpolarization in
Em2 vs. the interelectrode distance were used to
compute the intercellular resistance, ri, as
detailed by Lidington et al. (24). (Adapted from Ref.
24).
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Mouse Cremaster Muscle Preparation for Intravital Microscopy
To study the conducted response, we used a mouse preparation
rather than a rat preparation to permit future mechanistic studies in
genetically altered mice. The procedure for preparation of mouse
cremaster and all experimental protocols were approved by the Council
on Animal Care at the University of Western Ontario. Male mice
(C57BL/6, 20-30 g) were anesthetized with a mixture of ketamine
(80 mg/kg) and xylazine (4 mg/kg) injected intraperitoneally. Supplementary injections of this mixture (30% of initial dose) were
given as needed. In some mice, the left carotid artery was cannulated
to permit measurement of the arterial pressure. The mouse was placed in
a supine position on a styrofoam board and kept warm by radiant heat to
keep the rectal temperature at 37°C. The cremaster muscle preparation
was similar to that described for the rat (34). The skin
of the scrotum and its underlying connective tissue were cut
longitudinally along the anterior midline, exposing the cremaster
muscle sac. The sac was isolated from the scrotum, placed on a
histological glass slide, and irrigated with physiological saline
solution (PSS), pH 7.4, composed of (in mM) 131.9 NaCl, 4.7 KCl, 2.2 CaCl2, 1.2 MgSO4, and 20.0 NaHCO3
bubbled continuously with 95% N2-5% CO2 gas.
A longitudinal cut was made in the ventral surface of the muscle to
open the sac. Sutures were stitched at the muscle edge, pulled, and
pinned to the styrofoam board to gently spread the muscle over the
glass slide. The testicular mesentery and the vessels attached to it
were cauterized and cut to separate the contents of the sac from the
muscular sac itself. The contents were pushed into the abdominal
cavity; the cremaster muscle was then continuously superfused with PSS
(33-34°C) at a rate of 3-4 ml/min. The muscle was
epi-illuminated by means of a fiber-optic light guide and visualized
with an intravital microscope (ELR, Leitz) with a long-working distance
objective (×20/0.32 numerical aperture) and an eyepiece (×6.3). The
resultant field of view (0.56 × 0.38 mm) was video recorded by a
closed circuit system (MTI camera, Panasonic WV5410 monitor, and
Mitsubishi U82 sVHS tape recorder). Inner luminal arteriolar diameters
were measured off-line from the video screen with resolution of
about ±1 µm.
Experimental Protocols in Vitro
We have previously shown (24) that exposure of
RMECs to LPS for 1-24 h at a concentration of 10 µg/ml
increases ri from 3.3 to 5.3 M (a maximal
ri already occurs at 1 h). In terms of LPS
concentration dependency, increased ri was
already seen at 10 ng/ml (24). In the present study, we
aimed to determine whether 1) exposure of RMECs to a
combination of LPS and inflammatory cytokines further elevates
ri using TNF-, IL-1, and IL-6, agents known to be elevated in septic plasma (1, 10);
2) the effect of LPS + cytokines is reversible; and
3) the effect can be prevented by pretreatment of RMECs with
protein tyrosine kinase (PTK) inhibitors. The outcome of these in vitro
experiments dictated the design of subsequent in vivo experiments. The
concentrations of LPS (10 µg/ml, isolated from Escherichia
coli serotype 055:B5), TNF- (100 ng/ml), IL-1 (50 ng/ml)
(all from Sigma), and IL-6 (50 ng/ml, R&D Systems; Minneapolis, MN)
were chosen at the upper end of the concentration range reported in the
literature (2, 30) and were expected to produce a maximum
effect (30). We used PTK inhibitors tyrphostin (Tyr) A9
(1.5 µM, ICN Radiochemicals; Costa Mesa, CA) and PP-2 (10 nM,
Calbiochem; La Jolla, CA). The reason for choosing these two
structurally different agents was to minimize the possibility of their
nonspecific effect. To this end, we also used the inactive PTK
inhibitor Tyr A1 (0.1 mM, ICN Radiochemicals) as an "isoform
control." The concentrations of PP-2 and Tyr A1 and A9 were based on
previous reports (4, 7, 16) and the IC50
values reported by the manufacturer. Stock solutions of inhibitors were
dissolved in 100% DMSO; the final concentration of DMSO in the culture
medium was 0.01% unless otherwise stated.
Experimental Protocols in Cremaster Muscle in Vivo
Arterioles in the mouse cremaster muscle have been shown to
exhibit conducted responses after application of local dilatory and
constriction stimuli (18, 20). We used the approach of Gustafsson and Holstein-Rathlou (14) to stimulate
arterioles locally with a train of unipolar depolarization pulses. The
rationale for using this approach rather than that of local agonist
application (18, 20) was our wish to exclude possible
effect(s) of LPS/cytokines on the initiation of conducted response
(agonist-induced local diameter change). Briefly, glass micropipettes
of outer tip diameter 6-10 µm were backfilled with 2 M NaCl
solution and connected to a Grass stimulator (model S88) via an Ag-AgCl
wire. A reference electrode was inserted into the mouse tail. Each
pipette was mounted on a micromanipulator and the tip was positioned as
close to an arteriole as possible to deliver a 40-s train of pulses at
10 Hz frequency, 2 ms duration, and 40-80 V amplitude. In each
experiment, after the surgical exposure of the cremaster muscle and a
30-min stabilization period, a site in the middle portion of a 1.5- to 3-mm-long unbranched arteriole (~50 µm diameter, 1A or 2A branching order) was chosen for local electrical stimulation. A key criterion for
the arteriole selection was clear visibility of the arteriolar wall at
the local stimulation site and at a site 500 µm upstream from the
pipette tip. At the beginning of the experiment (one arteriole per
mouse), the amplitude was adjusted to produce ~50% reduction in the
local arteriolar diameter near the pipette tip. For any subsequent
treatments of the arteriole/cremaster muscle, we have stimulated
exactly the same local site with the same amplitude determined at the
beginning of the experiment. In the majority of experiments, we
simultaneously video recorded diameter changes occurring locally and at
the 500-µm upstream site. On the basis of prestimulation diameter
measurements (Dlocal,pre and
D500,pre) and the minimal diameter measurements
during the 40-s stimulation period (Dlocal,min
and D500,min), we defined the relative local and
upstream diameter changes as Dlocal(%) = 100% × (Dlocal,min 
Dlocal,pre)/Dlocal,pre
and D500(%) = 100% × (D500,min D500,pre)/D500,pre, respectively. We used the communication ratio at the 500-µm site, computed as CR500(%) = 100% × D500(%)/Dlocal(%),
as an index of the conducted response.
Effect of LPS on conducted response.
The design of the present in vivo experiments was driven by our
preceding cell culture work. Because cytokines had no effect on
ci (see RESULTS for details), the
present in vivo work dealt with LPS only. On the basis of LPS dose,
time dependencies determined by us in vitro (24), and
limited stability of the cremaster muscle preparation (3-4 h), the
in vivo protocol included 10 µg/ml LPS application for 1 h to
ensure maximal effect. The protocol consisted of the 30-min
stabilization period and cremaster muscle superfusion with PSS + 0.01% DMSO, followed by two to three repeated stimulations (~3 min
between stimulations) of the same local arteriolar site with the
predetermined amplitude. The outcome of these repeated stimulations
served to verify the reproducibility of the response. The muscle was
then superfused for 1 h with PSS containing 0.01% DMSO and 10 µg/ml LPS; the same local arteriolar site was again stimulated
2-3 times. Finally, the muscle was superfused for 1 h with
PSS + 0.01% DMSO only (washout period), and the same
arteriole was again stimulated 2-3 times. For each set of the
2-3 stimulations, Dlocal,pre,
D500,pre, Dlocal,min, and
D500,min were measured and then averaged. On the
basis of these averages, Dlocal(%),
D500(%), and CR500(%) were computed.
Effect of PTK inhibitors and LPS on conducted response.
Following the stabilization period, an arteriole was initially
stimulated 2-3 times and then subjected to one of the following three protocols. The first protocol included 1-h superfusion with PSS + 0.01% DMSO, followed by 1-h superfusion with 10 µg/ml LPS in PSS + 0.01% DMSO followed by 2-3 repeated stimulations.
The second protocol consisted of 1-h superfusion with a PTK inhibitor (1.5 µM Tyr A9 or 10 nM PP-2) in PSS + 0.01% DMSO or inactive Tyr A1 (0.1 mM) in PSS + 0.1% DMSO, followed by 1-h superfusion with PTK inhibitor plus 10 µg/ml LPS in PSS + 0.01/0.1% DMSO, and then followed by 2-3 repeated stimulations. The third protocol consisted of 2-h superfusion with PSS + 0.01% DMSO followed by 2-3 repeated stimulations (yielding the time-matched control response).
Control Experiments
To assess the direct effect of electrical stimulation on the
diameter response at the 500-µm upstream site, we positioned the
microelectrode tip at a site 100 µm away from the original local site
(the tip to upstream site distance was maintained at 500 µm) and
observed responses at the local and upstream sites. To assess the role
of arteriolar innervation in the presently studied conducted responses,
we evaluated CR500 and CR1000 (computed similarly to CR500) after tetrodotoxin (TTX, 10 µM final
concentration) was added to the PSS superfusate and allowed to
equilibrate for 15-20 min. The positive test for TTX included
surgical exposure of the right peroneal nerve and the right extensor
digitorum longus muscle of the mouse, supramaximal stimulation of the
nerve with a pair of Ag electrodes (5-10 V, 6 Hz), and the
subsequent observation of muscle twitching. The nerve was then
superfused with the 10 µM TTX solution for 15-20 min, and the
response to supramaximal stimulation (no twitching) was noted. Finally,
in separate mice, we assessed the vasomotor tone (or vasodilative
potential) of the approximately 50-µm arterioles used in the present
study. Similar to Hungerford et al. (18), a maximal
dilation was obtained by adding ACh (0.1 mM final concentration) to the
PSS superfusate and by allowing 10- to 15-min equilibration.
Statistics
All data were expressed as means ± SE. Parameters were
analyzed using an analysis of variance, followed by t-test
with Bonferroni correction for multiple comparisons when applicable. A
level of P < 0.05 was considered significant.
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RESULTS |
Effect of LPS and cytokines on ci in in vitro
experiments.
Figure 1 exemplifies the electrophysiological approach we used to
determine the ri (or ci)
of the RMEC monolayer. Figure 1A demonstrates that the size
of the current-induced hyperpolarization (Em2)
decreased with increasing interelectrode distance. Using the Bessel
function mathematical model, the rate of decrease with distance was
used to estimate ri (details in Ref.
24). The average resting Em was
29 ± 1 mV (n = 316 impaled cells); any of the subsequent treatments of the monolayer had no significant effect on
resting Em. For untreated control monolayers,
the values were ri = 3.5 ± 0.1 M,
ci = 0.28 ± 0.01 µS (Fig.
2). LPS significantly reduced
ci but LPS applied together with all of the
three cytokines did not result in a further reduction in
ci (Fig. 2). None of these cytokines applied
alone had a significant effect on ci. Based on
these findings, all of our subsequent in vivo experiments used LPS
alone to mimic the effect of sepsis on conducted arteriolar response.
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Fig. 2.
Effect of lipopolysaccharide (LPS) and cytokines on
intercellular conductance (ci). LPS (10 µg/ml)
and the cytokines tumor necrosis factor (TNF)- (100 ng/ml),
interleukin (IL)-1 (50 ng/ml), and IL-6 (50 ng/ml) were tested alone
and in combination for an effect on intercellular conductance
(ci = 1/ri). After a
2-h exposure, LPS alone and the combination of three cytokines plus LPS
caused a 33% reduction in ci. Cytokines alone
had no effect. Similar results were obtained for a 24-h exposure,
instead of 2-h exposure (data not shown). The control
ci value was based on 36 measurements of
ci from 12 monolayers; each of the remaining
values was based on 12 measurements of ci from 3 monolayers. *Significant difference from control.
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Baseline measurements and control experiments in in vivo model.
The mean arterial pressure was typically 85-100 mmHg during
experiments (average 93 ± 3 mmHg, subset of 17 mice). On the
basis of visual assessment of the microvascular flow in the cremaster muscle and on blood pressure measurement, preparations were stable during the 3- to 4-h experimental protocol. In the subgroup of 1A and
2A arterioles used to determine the vasodilative potential (average
diameter (D) 44 ± 7 µm, n = 10), ACh
dilated arterioles by D = 4.1 ± 5 µm
(13 ± 4% diameter increase). Figure
3A exemplifies the time course
of arteriolar diameter changes measured at the tip of the electrode
(local) and 500 µm upstream during a 40-s electrical stimulation
period in PSS superfused muscle. At the local site, the diameter
reached a minimum quickly (within the first 10-s period after the onset
of the stimulus), and then it tended to recover before the end of the
stimulus. At the 500-µm site, the arteriole also constricted quickly,
but the minimum diameter was reached somewhat later than the minimum at
the local site. Figure 3B underscores these features, based
on the average local and 500-µm site responses normalized with
respect to the prestimulation diameter. The size of the standard error
bars in Fig. 3B reflects an appreciable variability observed
in the time course of local and upstream diameter responses among
different arterioles. Figure
4A summarizes observed
diameter changes at the local, 500-µm, and 1,000-µm upstream sites
during control PSS superfusion and after TTX application. Figure
4B shows the communication ratios (CR500 and
CR1000) for these two protocols. Because our control
experiments with peroneal nerve stimulation showed that our TTX
solution was effective, we show in Fig. 4 that the nerves did not
participate in the conducted response measured under the present
experimental conditions. Finally, repositioning of the
electrode tip 100 µm away from the arteriolar wall but delivering a
comparable stimulus (63 ± 4 V, n = 6) as in Fig. 4 abolished diameter responses at the local and upstream sites (data
not shown). Thus responses at the upstream sites required the presence
of local diameter responses.
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Fig. 3.
A: Example of time course of diameter
(D) changes in a mouse cremaster muscle arteriole. Changes
were induced by a 40-s local electrical stimulation of the arteriole
and were measured at the microelectrode tip (local) and 500 µm
upstream from the tip (500 µm). B: time course of relative
diameter [D(%)] changes caused by electrical
stimulation. To compute D(%), the stimulation-induced
reduction in diameter (in µm) was divided by the prestimulation
diameter. Data are based on measurements from 6 arterioles in 6 mice.
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Fig. 4.
A: Effect of tetrodotoxin (TTX, 10 µM) on
the relative arteriolar diameter change measured at the tip of the
stimulation pipette (local) 500-µm and 1,000-µm upstream sites. In
5 arterioles, diameter changes were measured at local and 500-µm
sites. In 3 of these arterioles, diameter changes were also measured at
the 1,000-µm site. Local prestimulation diameter before application
of TTX (58.1 ± 8.2 µm, n = 5 arterioles in 5 mice) was not altered by the TTX treatment. Average amplitude of the
local stimulus was 68 ± 6 V. B: communication ratio,
CR, determined from the data in A for the 500-µm and
1,000-µm sites [CR500(%) = 100% × D500(%)/ Dlocal(%),
CR1000(%) = 100% × D1000(%)/Dlocal(%)].
CR was used as an index of the conducted response. There was no effect
of TTX on the conducted response.
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Effects of LPS, wash, and PTK inhibitors on ci in vitro
and conducted response in vivo.
Although our in vitro experiments were completed before in vivo work
began, data on LPS, wash, and PTK inhibitors are presented together to
permit comparison between our in vitro and in vivo models. Figure
5A demonstrates that 10 µg/ml LPS was significantly reduced, but a subsequent 1-h wash
restored, the ci in monolayers in vitro (raw
electrophysiological data for 5A were also used in Ref.
24). Figure 5C shows that the same
concentration of LPS in the superfusate reduced the conducted response
in the cremaster muscle in vivo, and that 1-h wash restored it
[changes in D500(%), Fig. 5B,
showed comparable reduction and restoration]. LPS did not affect the
local diameter response [Dlocal(%), Fig.
5B] nor the time delays of the measured diameter minima
(time after the stimulus onset 11 ± 1 and 28 ± 3 s for
pre-LPS stimulation, n = 6; and 14 ± 2 and
28 ± 3 s for post-LPS stimulation for local and upstream
minima, respectively). Thus based on comparison of Fig. 5,
A and C, LPS caused qualitatively similar
effects on communication in our in vitro and in vivo models.
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Fig. 5.
A: Washout of LPS. In endothelial cell
monolayers, a 2-h exposure of LPS (10 µg/ml) was followed by
replacement of the exposure medium by control medium. After 1 h of
wash, ci returned to the control level. Control
values are based on 12 measurements of ci from 4 monolayers; LPS and washout (1-2 h) values are each based on 20 measurements of ci from 5 monolayers.
B: effect of LPS (10 µg/ml, 1 h) in the cremaster
muscle superfusate and of wash (1 h) on the relative arteriolar
diameter change measured at local and 500-µm sites. Local
prestimulation diameter (55.6 ± 6.3 µm, n = 6 arterioles in 6 mice) was not altered by the LPS or wash treatments.
Average amplitude of the local stimulus was 59 ± 6 V. C: communication ratio, CR500, was determined
from the data in B. *Significant difference from control.
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Figure 6, A and B,
summarizes the effect of PTK inhibitors. In both models, pretreatment
with the same concentration of Tyr A9 and PP-2 prevented the
LPS-induced reduction in communication (raw data for LPS + PP-2
bar in Fig. 6A were also used in Ref. 24). As
well, pretreatment with the inactive inhibitor Tyr A1 had no effect on
the LPS-induced reduction. Thus comparison of Fig. 6, A and
B, revealed that PTK inhibitors caused a qualitatively similar effect on LPS-induced reduction in communication in both of our
models. In our in vivo model, the application of Tyr A1, A9, or PP-2
alone had no effect either on the local diameter response to the
electrical stimulus or on D500(%); in our in
vitro model, these agents alone had no effect on
ci (data not shown).
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Fig. 6.
Effect of tyrosine kinase inhibitors. A: in
endothelial cell monolayers, a 1-h pretreatment with tyrphostin (Tyr)
A9 (1.5 µM in 0.01% DMSO) but not the inactive control Tyr A1 (100 µM in 0.1% DMSO) prevented the reduction in intercellular
conductance after 24-h exposure to 10 µg/ml LPS. The inhibitor PP-2
(10 nM in 0.01% DMSO) also prevented the LPS response. Values of
ci in control and LPS groups are based on 44 measurements from 11 monolayers; all other values are based on
12-16 measurements of ci from 3-4
monolayers. B: CR500 for arterioles exposed to
PSS + 0.01% DMSO (control group), LPS, Tyr A9 + LPS,
PP-2 + LPS, or Tyr A1 + LPS at superfusate concentrations
identical to those used in vitro experiments of A. Separate
arterioles (one arteriole/mouse) were used in each of the 5 groups
shown. Among the 5 groups, the number of arterioles, the average local
prestimulation diameter, and the average stimulus amplitude ranged from
5 to 8, from 42.0 ± 2.5 to 64.4 ± 6.6 µm, and from
64 ± 5 to 75 ± 2 V, respectively. None of the treatments
affected the prestimulation diameter, whereas there was no effect of
DMSO (either 0.1 or 0.01%) on ci or
CR500 (data not shown). *Significant difference from
control.
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DISCUSSION |
The objectives of this study were to determine whether
communication data obtained from our in vitro monolayer model could provide a suitable framework for our in vivo animal experiments, and
whether LPS can modulate conducted arteriolar response in the mouse
cremaster muscle. On the basis of qualitatively comparable data in both
in vitro and in vivo models, our results indicate that LPS attenuates
microvascular cell-to-cell communication in a reversible and tyrosine
kinase-dependent manner.
Experimental models.
A considerable amount of work has been done in the area of vascular
cell communication, based on in vitro (see review in Ref. 5) and in vivo approaches (see review in Ref.
15). However, to our knowledge, there are no studies
exploiting both approaches simultaneously. In the present study, we
took advantage of our recent in vitro work where LPS was shown to
increase ri in RMEC monolayers
(24). In our preliminary experiments (unpublished observation) using the same electrophysiological approach as that of
our RMEC study, LPS also increased ri in
monolayers derived from mouse skeletal muscle endothelial cells. Thus
based on this background work and on the mechanistic potential of
genetically engineered mice, we chose the mouse cremaster muscle as an
in vivo model to complement our RMEC monolayer model. In general, the
presently measured Em and the intercellular
communication ability in the RMEC monolayer were consistent with
previous reports (27, 28). Similarly, the baseline
hemodynamic parameters of the cremaster muscle preparation (e.g., mean
arterial blood pressure, stability of perfusion) and the ability to
conduct local diameter changes along the arteriole length agreed with
published reports of conducted response in mice (6, 18,
20).
In the present study, we used the electrical depolarization protocol of
Gustafsson and Holstein-Rathlou (14) to elicit conducted vasoconstriction. There were two reasons for choosing this protocol. First, because the vasodilative potential of our arterioles was modest
(13%), the detectability of conducted vasodilation and its modulation
by LPS could have been compromised by the limited resolution of our
diameter measurement technique (±1 µm). Thus conducted
vasoconstriction would provide a better "signal-to-noise" ratio.
Second, we wanted to exclude the possible dependence of the size of the
conducted response on the size of the initial local response
(38). Thus we used the protocol to set the size of the
local response to a desired level by adjusting the amplitude of the
stimulus. In general, the maximal local diameter reduction elicited by
a particular voltage was highly variable among arterioles (reflecting
variability in the muscle surface-to-arteriole distance, pipette tip
diameter, or responsiveness of the vessel itself) (20).
The range of amplitudes (40-80 V) used to achieve the desired
local constriction (~50%) was comparable to the voltage range
reported for the rat mesentery (14).
Local and conducted responses under control experimental
conditions.
The size of local and conducted responses at 500-µm and 1,000-µm
sites and the tendency of the conducted response to lag behind the
local response (Figs. 3 and 4) were comparable to those reported for
the rat mesentery (14). The time course of local
vasoconstriction during the 40-s stimulation period (Fig.
3B) agreed with the time course of vasoconstriction in mouse
cremaster muscle arterioles (20). The rapid onset of
constriction followed by a gradual reduction in constriction during the
stimulation (Fig. 3B) may be accounted for by coordinated
smooth muscle and endothelial cell function (calcium signaling-induced
nitric oxide release) (20, 40). Figure 4 shows that the
present local and conducted responses were independent of perivascular
nerves as TTX (a fast sodium channel antagonist) had no effect on these
responses. This finding is consistent with that of Kumer et al.
(20), but it appears to disagree with the results of
electrical depolarization protocol reported by Hungerford et al.
(18). The disagreement could be due to differences in
stimulation parameters (1 ms pulse at 32 Hz, 60-100 V)
(18) and the size of the local constriction employed
(80%).
Effect of LPS on vascular cell communication.
Although the effect of LPS on conducted response can be studied in mice
injected with LPS, the experimental outcome could be difficult to
interpret because it may depend on the direct effect of LPS on the
microvasculature as well as on the systemic response to LPS. For this
reason, the present study involved adding LPS to the cremaster muscle
superfusate to keep LPS as local as possible to minimize systemic effects.
Data from cell culture experiments (Figs. 2, 5A, and
6A) extended the findings of our recent study
(24) where LPS increased ri in RMEC
monolayers. The data (Fig. 2) show for the first time that cytokines
TNF-, IL-1, and IL-6 applied alone had no effect on conductance
and that LPS applied together with all of the three cytokines did not
further reduce conductance. Data in Fig. 6A agree with our
report that PTK inhibitors prevent the effect of LPS on cell-to-cell
communication (24).
Data from our experiments in mice (Figs. 5C and
6B) demonstrate for the first time that 1) LPS
reduced the conducted arteriolar response, 2) a wash
restored the response, and 3) PTK inhibitors prevented the
effect of LPS. Furthermore, data in Figs. 5C and 6B permit comparison with our cell culture work (Figs.
5A and 6A). Clearly, our in vitro and in vivo
models differed. The spread of signal(s) in two dimensions of
endothelial cell monolayer might not be comparable to the prevalently
one-dimensional spread of signal(s) in endothelial and smooth muscle
cells along the arteriolar wall (40). Yet, despite these
differences, LPS, LPS washout, and PTK inhibitors had strikingly
similar effects in the two models. Although it is possible that
the parallel outcome for the five treatments (LPS, LPS washout,
LPS + TYR A9, LPS + PP-2, and LPS + TYR A1) was
coincidental, it is also possible that the outcome reflected a
fundamental mechanism common to both models. One such mechanism could
be the spread of signal(s) in both models via gap junctional (GJ)
communication (24, 25). LPS could modulate GJ function in
both models similarly, including activation of the PTK pathway.
Although the mechanism of this modulation has not been clarified, PTK
inhibitors in the present experiments could have prevented LPS
receptor-mediated phosphorylation of cytosolic kinases or GJ proteins.
Cytosolic Src kinases (e.g., pp60src) have been shown to
phosphorylate tyrosine residues in GJ proteins and reduce cell-to-cell
communication (26) or, in turn, activate other tyrosine
kinases (e.g., p125Fak) which could also phosphorylate GJ
tyrosine residues (21, 23).
Conducted response in arterioles in vivo has been shown to be reduced
by GJ uncouplers (31) and enhanced by angiotensin II
(14). To our knowledge, the physiological impact of
modulation of conducted response has not been addressed. Referring to
Figs. 5 and 6, it is difficult to estimate this impact, based on the LPS-induced 40-50% reduction in ci in our
in vitro model. However, this task may be easier considering the
reduction in the conducted response (40-50% reduction in
CR500). To this end, we have developed a mathematical model
and used the present data to estimate the modulatory effect of LPS on
the resistance to blood flow (R) in an unbranched arteriole
(APPENDIX). Assuming that the conducted response in our
mice was of the same exponential character as that reported for the rat
mesenteric arteriole (employing the same stimulation protocol)
(14), our local 50% constriction was predicted to
increase R in control arterioles by a factor of 4.31 (Fig.
8). The same local constriction during exposure to LPS was predicted to
increase R by a factor of 2.83 (Fig. 8). Thus LPS could
reduce the arteriolar ability to control resistance by ~30%. Thus it
is possible that during an LPS-induced inflammatory response the effect
of the documented reduced vasoconstrictive ability (13,
33) could be aggravated by the reduced ability to conduct
constriction along the blood vessel length. To this end, agents aimed
at restoring/enhancing communication in blood vessel wall (enhancing
the ability to increase peripheral resistance) could possibly be
beneficial against LPS-induced hypotension. Clearly, given the multiple
effects of LPS on the vessel wall, further studies are needed to
address this possibility.
The present approach of cell culture-driven in vivo work in mice may
provide a framework for such future studies. Recently, Giepmans et al.
(12) indicated that inhibition of GJ communication in
cultured fibroblasts was caused by c-Src-mediated phosphorylation of
residue Y265 on the COOH-terminal tail of GJ protein Cx43. If our in
vitro model would indicate that this Y265 residue also mediates the
LPS-induced reduction in communication, then a specific transgenic
mouse with a mutation at this residue could be used to examine
1) lack of effect of LPS on conducted response and 2) possible attenuation of hypotension in LPS-injected mice.
In conclusion, the present study showed that 1) LPS
reduced arteriolar conducted response in mouse cremaster muscle in a
reversible and tyrosine kinase-dependent manner, 2) in vitro
data predicted the effect of LPS in this mouse model, and 3)
the degree of reduction of conducted response had the potential to
appreciably affect the microcirculatory hemodynamics.
| |
APPENDIX |
The model.
To assess the effect of local arteriolar diameter change on the change
in resistance to blood flow (R) in the arteriole, and the
modulating effect of LPS on this relationship, the following model
incorporated these assumptions. 1) R
(mmHg · ml
1 · min) could be computed from
the Poiseuille's law, such that R = klA/r4, where
lA (mm) is the length of unbranched arteriole,
r (mm) is the inner (luminal) radius of the arteriole, and
k
(ml · min1 · mmHg1 · mm3)
is a constant that reflects the viscosity of the blood in the arteriole; 2) viscosity of blood, and k, do not
change with r; and 3) a stimulus is applied at
the midpoint of the arteriole to cause a localized radius change. This
change spreads equally toward both ends of the arteriole, and the size
of this change decreases with distance according to a simple
exponential decay (Fig. 7).
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Fig. 7.
Schematic diagram outlining the features of the mathematical model
in APPENDIX. Heavy dashed line depicts the arteriole before
local stimulation. A local stimulus introduced via a pipette at
position i = 0 constricts the arteriole locally and via
cell-to-cell communication also along the vessel length (heavy solid
line). The shape of the constricted arteriolar lumen is approximated by
a series of cocentric cylindrical sleeves (total numbers of sleeves is
N), as detailed in APPENDIX. The length of each
sleeve is x, rc is the control
radius, rsi represents the sleeve radii, and
L is the length of half of the arteriole.
|
|
Referring to Fig. 7, rc is the control,
prestimulation radius of the arteriole. A stimulus applied at position
i = 0 changed the radius to a new value
rs0 (i being a labeled position along the x-axis). To model the effect of the exponential decay
with x, one-half of the stimulated arteriole is pictured as
a series of cocentric cylindrical sleeves whose radii
(rs1, rs2, ...,
rsi, ...,
rsN) increase with x in an
exponential fashion (Fig. 7). Here the right half of the arteriole
(length of the half is L = lA/2)
is divided into N number of equal segments of length x, such that L = Nx.
We chose to label each sleeve according to its starting position along
the x-axis. Thus the sleeve starting at position
i = 0 (sleeve 0) had the largest
stimulus-induced radius change
|
(1)
|
The next sleeve had a radius change
rs1 = rc rs1, whereas the ith sleeve had a
radius change
|
(2)
|
The size of the change in radius decays exponentially with
i, such that
|
(3)
|
where 
is the mechanical length constant of the decay for the
arteriole (15) and reflects the "degree" of conducted
response along the vessel. Substituting Eqs. 1 and 2
into Eq. 3 and rearranging the radius at position
i can be computed as
|
(4)
|
According to Poiseuille's law, the resistance to flow in the
smallest cocentric sleeve is Rs0 = kx/rs04,
whereas the resistance of the ith sleeve is
Rsi = kx/rsi4.
The total resistance of all of the N sleeves together is
|
(5)
|
Substituting Eq. 4 into Eq. 5, the total
resistance to flow of the right half of the of arteriole is
then
![R<SUB>s,total</SUB><IT>=k&Dgr;x </IT><LIM><OP>∑</OP><LL><IT>i</IT><IT>=</IT>0</LL><UL><IT>N−</IT>1</UL></LIM> [<IT>r</IT><SUB>c</SUB><IT>+</IT>(<IT>r</IT><SUB>s0</SUB><IT>−r</IT><SUB>c</SUB>)<IT>e</IT><SUP>(−<IT>i</IT><IT>&Dgr;x/&lgr;</IT>)</SUP>]<SUP>−4</SUP>](/content/vol281/issue3/fulltext/H1397/img006.gif)
|
(6)
|
It is of interest to consider the degree of the conducted
response, , with respect to the arteriolar length (L) and
the minimal local radius, rs0, with respect to
the prestimulation control radius rc. Thus we
defined the normalized "degree of conduction" as C = /L, and the normalized local radius as
S = rs0/rc. To simplify the
mathematical model further, we chose L = 1 "length units" long, and rc = 1 "radius
units" wide. Recalling that we defined L = Nx (Fig. 7), x now becomes
numerically equal to 1/N, C equal to , and
S equal to rs0. Substituting these
normalized parameters into Eq. 6, we obtain
![R<SUB>s,total</SUB><IT>=kN</IT><SUP>−1</SUP> <LIM><OP>∑</OP><LL><IT>i</IT><IT>=</IT>0</LL><UL><IT>N−</IT>1</UL></LIM> [1<IT>+</IT>(<IT>S−</IT>1)<IT>e</IT><SUP>(−<IT>i</IT><IT>/NC</IT>)</SUP>]<SUP>−4</SUP>](/content/vol281/issue3/fulltext/H1397/img007.gif)
|
(7)
|
The resistance to flow in the control, prestimulated, half of
the arteriole is Rcontrol = kL/rc4 (assuming that
rc does not change with x). Using the
same simplified length and radius units as above,
Rcontrol becomes numerically equal to
k. From Eq. 7, we can now compute the normalized
resistance to flow as Rnorm = Rs,total/Rcontrol, or
Rnorm = Rs,total/k, or
![R<SUB>norm</SUB><IT>=N</IT><SUP>−1</SUP> <LIM><OP>∑</OP><LL><IT>i</IT><IT>=</IT>0</LL><UL><IT>N−</IT>1</UL></LIM> [1<IT>+</IT>(<IT>S−</IT>1)<IT>e</IT><SUP>(−<IT>i</IT><IT>/NC</IT>)</SUP>]<SUP>−4</SUP>](/content/vol281/issue3/fulltext/H1397/img008.gif)
|
(8)
|
The normalized resistance depends on the normalized local
radius, S, (e.g., dictated by the strength of the stimulus)
and on the degree of the conductive response (C). For large
enough N (N = 1,000), computation of
Rnorm is insensitive to the choice of
N (increasing N from 1,000 to 10,000 in Fig.
8 altered Rnorm by
~0.1%). For very large C, Rnorm
approaches S4 (Poiseuille's law). For very
small C (e.g., local response is not conducted),
Rnorm approaches unity
[Rnorm becomes independent of the
stimulus-induced radius change, an observation consistent with
experimental results of Kurjiaka and Segal (19)]. Note that, because of symmetry, S is also equal to the normalized
arteriolar diameter, S between values 0 and 1 represents
arteriolar constriction, S >1 represents dilation, and
Rnorm is also equal to the normalized resistance
of the entire arteriole.
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|
Fig. 8.
Plots of the normalized resistance to flow,
Rnorm, vs. normalized arteriolar diameter,
S, according to Eq. 8 in APPENDIX
(N in Eq. 8 is set to 1,000).
Rnorm is shown for S ranging from 0.5 to 2 (from 50% local constriction to 100% dilation) and for two
C values (Ccontrol and
CLPS) estimated from the present in vivo data.
C represents the degree of conducted response normalized to
the arteriolar length (details in APPENDIX). According to
the model in APPENDIX, LPS treatment was predicted to
attenuate the arteriolar ability to elevate (e.g., at S = 0.5) and to lower (e.g., at S = 2) the resistance to
flow offered by the arteriole.
|
|
Application of present experimental data to model.
In the following, we wanted to determine C in Eq. 8 to plot Rnorm versus S. In
particular, we aimed to estimate C under the present control
(Ccontrol) and LPS superfusion conditions
(CLPS). According to Gustafsson and
Holstein-Rathlou (15), the conducted response can be
described by the relationship Dx =
D0
e(x/
) (D is
arteriolar diameter). If our diameter responses measured locally
(x = 0) and at x = 500 µm upstream
obeyed this relationship, then
|
(9)
|
Assuming further that the prestimulation diameters at
local and 500-µm upstream locations were of the same value
(Dlocal,pre = D500,pre, Fig. 3A), and dividing both
sides of Eq. 9 by this value, the equation could be
rewritten as D500(%) = D0(%) e(500/).
Because CR500(%) was defined as 100% × D500(%)/Dlocal(%), Eq. 9 could further be rewritten as
|
(10)
|
Let us consider an arteriole at 10 s after the onset
of the stimulus (Fig. 3). At this time, the arteriole constricted to its minimum diameter locally and almost to its minimum diameter at the
500-µm upstream location (Fig. 3B), both under control and
LPS conditions. Thus the average CR500(%) values at the
10-s time point are approximately equal to the CR500(%)
values shown in Fig. 5C (based on the maximal
constrictions). Using the data in Fig. 5C
(CR500,control = 45% and CR500,LPS = 21%) and Eq. 10, we computed control = 626 µm, and LPS = 320 µm. In a subset of our
experiments we measured the average length of unbranched arterioles to
be 1,900 ± 370 µm (n = 6) giving the average
L = 950 µm. Setting 950 µm to be one "length
unit", control was then 0.66 units and
LPS was 0.34 units, whereas the normalized degree of
conduction Ccontrol = 0.66 and
CLPS = 0.34.
Figure 8 shows the plot of Rnorm versus
S in the range from 0.5 to 2 (from 50% local constriction
to 100% local dilation) with Ccontrol = 0.66 and CLPS = 0.34 (N = 1,000). Results show that for 50% constriction (level achieved by
present stimulus, Fig. 5B) resistance to flow can be
predicted to increase by a factor of 4.31 in the control arteriole but
only by a factor of 2.83 in LPS superfused arteriole. Thus LPS could
reduce the arteriolar ability to elevate resistance by 34%. A similar
impact of LPS could be observed for the arteriolar ability to reduce
resistance by dilation (for S = 2, Fig. 8).
To our knowledge, there is no report of a mathematical model
aiming to determine the effect of the conducted response and the effect
of modulation of this response on the resistance to blood flow. The
present model is the "first approximation" model, which does not
take into account many complexities of the real arteriole (e.g.,
tapering of arteriole along its length, dependence of k on
radius when radius becomes small, a possible departure of the character
of the conducted response from a simple exponential decay, etc.). We
also attempted to minimize the effect of the complexity of arteriolar
branching by placing the stimulus at the arteriolar midpoint.
Nevertheless, despite its simplicity, the model may provide the
opportunity to theoretically assess the impact of conducted response on
microcirculatory hemodynamics and help design experiments aimed at
evaluation of this impact.
| |
ACKNOWLEDGEMENTS |
The authors thank A. Bihari, M. Keet, and L. Cheng for technical
help, and Dr. H. Ladak for help with Matlab programming of the
mathematical model in APPENDIX.
| |
FOOTNOTES |
The Heart and Stroke Foundation of Ontario and the Canadian Institutes
of Health Research provided financial support.
Address for reprint requests and other correspondence: K. Tyml,
Dept. of Medical Biophysics, Univ. of Western Ontario, London, Ontario,
Canada N6A 5C1 (E-mail: ktyml{at}lhsc.on.ca).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 9 February 2001; accepted in final form 6 June 2001.
| |
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ABSTRACT
INTRODUCTION
