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The John B. Pierce Laboratory and Department of Cellular and Molecular Physiology, Yale University School of Medicine, New Haven, Connecticut 06519
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ABSTRACT |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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We performed intracellular recording with
Lucifer yellow dye microinjection to investigate the cellular
pathway(s) by which constriction and dilation are conducted along the
wall of arterioles (diameter 47 ± 1 µm,
n = 63) supplying blood flow to the
cheek pouch of anesthetized hamsters. At rest, membrane potential
(Em) of
endothelial (
36 ± 1 mV) and smooth muscle (35 ± 1 mV) cells was not different. Micropipette delivery of norepinephrine
(NE) or phenylephrine (PE) produced smooth muscle cell depolarization (5-41 mV) and vasoconstriction (7-49 µm) at the site of
release and along the arteriole with no effect on
Em of endothelial
cells. KCl produced conduction of depolarization and
vasoconstriction with similar electrical kinetics in endothelial and
smooth muscle cells. Acetylcholine triggered conduction of vasodilation
(2-25 µm) and hyperpolarization (3-33 mV) along both cell
layers; in smooth muscle, this change in
Em was prolonged
and followed by a transient depolarization. These cell-specific
electrophysiological recordings uniquely illustrate that depolarization
and constriction are initiated and conducted along smooth muscle,
independent of the endothelium. Furthermore, conduction of vasodilation
is explained by the spread of hyperpolarization along homologously
coupled endothelial and smooth muscle cells, with distinctive responses between cell layers. The discontinuity between endothelium and smooth
muscle indicates that these respective pathways are not electrically
coupled during blood flow control.
gap junctions; blood flow control; cell-to-cell communication; cell coupling; membrane potential; endothelium-derived hyperpolarizing factor; microcirculation; resistance vessels
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INTRODUCTION |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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CONDUCTION OF VASOMOTOR RESPONSES enables the coordination of blood flow magnitude and distribution within resistance networks (27, 36). The conduction of vasodilation and vasoconstriction have been taken to reflect the electrotonic spread of electrical signals through gap junctions between cells of the arteriole wall (18, 27, 29, 38). Early studies of electrical conduction in arterioles assumed that the coupling between smooth muscle cells provided the signaling pathway (13). Subsequently, both electrical and dye coupling between endothelial cells were found to be prevalent (16, 17, 26). Whereas such homologous coupling within these respective cell types is now established, the nature of heterologous coupling between smooth muscle and endothelial cells has remained controversial (1, 2, 19, 26). In part, such controversy may reflect differences between measurements performed using cultured cells (16, 17), excised arteries (1, 2), and microvessel preparations (13, 19, 26, 29, 38).
To date, electrophysiological studies of conduction in arterioles have utilized isolated preparations studied in vitro (13, 29, 37, 38), in which transmural pressure and luminal flow had been eliminated. A variety of second messengers, which are able to regulate the conductance of gap junctions [e.g., cyclic nucleotides, Ca2+, and nitric oxide (20, 30)], are produced in response to both pressure and flow. Therefore, in addition to the physical trauma induced by dissection, the elimination of tangential wall stress and luminal shear stress may alter gap junctional coupling relative to in vivo conditions.
The goal of this study was to define the cellular pathway(s) by which
electrical responses are initiated and conducted along arterioles in
vivo. We tested the hypothesis that each cell layer (endothelial,
smooth muscle) could provide a pathway for conducting vasomotor
responses along the arteriolar wall. In the hamster cheek pouch (27)
-adrenoceptor and muscarinic agonists were applied via micropipettes
onto arterioles to initiate responses on smooth muscle and endothelial
cells, respectively (14, 37); membrane potential
(Em) and
diameter were monitored at the site of application or at a remote site
along the vessel. Our data from cells identified by dye labeling
indicate that homologous gap junctional coupling enables current to
spread between endothelial cells or between smooth muscle cells;
however, heterologous electrical coupling was not prevalent between the
two cell types. Thus smooth muscle and endothelium each provide a
signaling pathway, with the nature of the stimulus determining which
cell layer conducts the signal along the vessel wall.
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METHODS |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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Hamster cheek pouch preparation. All procedures were approved by the Animal Care and Use Committee of The John B. Pierce Laboratory and were performed in accordance with the Guide for the Care and Use of Laboratory Animals of the National Research Council (revised 1996). Male Golden hamsters (80-110 g, n = 35; Charles River Breeding Laboratories) were anesthetized with pentobarbital sodium (60 mg/kg ip) and tracheotomized (PE-190 tubing) to ensure airway patency. A cannula (PE-50) secured in the left femoral vein enabled continuous replacement of fluids and maintenance of anesthesia throughout experiments (10 mg pentobarbital sodium/ml isotonic saline, infused at 0.41 ml/h). Esophageal temperature was maintained at 37-38°C with conductive heating.
With the use of a stereomicroscope, the cheek pouch was exteriorized onto a Plexiglas board and superficial connective tissue was removed for observation of microvessels (27). The preparation was superfused continuously with a bicarbonate-buffered Ringer solution at 37°C (pH 7.2-7.4) of the following composition (in mM): 137 NaCl, 4.7 KCl, 1.2 MgSO4, 2.0 CaCl2, and 18 NaHCO3 and equilibrated with 5% O2-5% CO2-90% N2. These reagents were obtained from J. T. Baker or Sigma; others were obtained from Sigma unless noted. The preparation was placed on the stage of an intravital microscope (model 20T, Zeiss) and transilluminated with Köhler illumination using a Leitz UM 32 objective (NA = 0.20), which provided a 15-mm working distance to facilitate microelectrode penetration. The optical image was coupled to a video camera (model NC-70X, Dage-MTI); total magnification on the monitor face (model PVM 133, Sony) was ×470.Membrane potential, intracellular dye injection, and diameter
recordings.
Recording microelectrodes were pulled (model P-87, Sutter Instruments)
from borosilicate glass tubes (no. GC120F-10, Warner Instruments) to
produce a flexible tip that could move with the vessel wall during
vasomotor activity. Tips were filled with Lucifer yellow dye (4%
solution in deionized water); the remainder of the electrode
(resistance 300-400 M
) was backfilled with 150 mM
LiCl2. The reference electrode
(Ag/AgCl pellet) was positioned in the effluent solution, and
Em was monitored
using an electrometer (model 773, World Precision Instruments).
Microiontophoresis and pressure microejection of vasoactive stimuli. Norepinephrine (NE; 1 M), phenylephrine (PE; 0.5 M), and acetylcholine (ACh; 1 M) were prepared daily in deionized H2O and micropipettes using microiontophoresis (model 260, World Precision Instruments). Ejection current was held at 500 nA, and stimulus duration varied from 150 to 2,000 ms; retain currents were <200 nA. KCl (0.5 M) was ejected from micropipettes using pressure (20-40 psi, 250-2,000 ms; model PLI-100, Medical Systems). Stimulus pipettes (tip inner diameter: 1 µm for microiontophoresis and 4-5 µm for pressure microejection) were fabricated with the same capillary tubes and puller used for the recording electrodes.
Experimental protocol. Suitable sites for study were identified during the period of equilibration (~60 min) of the preparation on the intravital microscope. Hydraulic micromanipulators were used to minimize mechanical disturbance. A stimulus pipette was secured in one manipulator (model MO-102, Narishige) and positioned with its tip adjacent to a second- or third-order arteriole. With reference to a calibrated eyepiece reticule, the recording microelectrode was positioned parallel to the vessel axis with its tip located at a distance of 530 µm upstream from the stimulus pipette; this distance nearly spanned the field of view. The tip of the microelectrode was lowered carefully (model MX510, SOMA) onto the edge of the arteriole, and a cell was penetrated. After a stable Em and vessel diameter were attained (~1 min), a stimulus was applied; 1-2 min of recovery were allowed between successive stimuli. At the upstream site, changes in Em and vessel diameter were recorded as "conducted" responses. Once a range of conducted responses were obtained (3-4 stimuli), the stimulus pipette was repositioned with its tip adjacent to the recording site. Identical stimuli were applied, and the changes in Em and vessel diameter were recorded as "local" responses. In such a manner, both conducted and local responses were obtained from the same cell within a given experiment. If a particular stimulus elicited vasomotor responses without a corresponding change in Em (see RESULTS), KCl was added to the 50-ml reservoir of superfusion solution to produce an external K+ concentration of 55 mM; depolarization confirmed successful impalement and intracellular recording.
Data analysis. Recordings were selected to illustrate the characteristic effects of each stimulus on endothelial cells and smooth muscle cells. Because each experiment involved recording from a single cell, endothelial cell and smooth muscle cell recordings within each of Figs. 2-5 are from separate experiments and are shown together for illustrative purposes. To help quantify the discordant electrical responses between endothelial cells and smooth muscle cells to ACh (see RESULTS), the total duration of an electrical response was expressed as a percentage of the total duration of the vasomotor response; each was defined as the time between the response onset and complete recovery. Summary data from all recordings are presented in Fig. 6.
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RESULTS |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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Identification of recording sites. The cell type from which a recording was obtained was ascertained by examining the labeling pattern following dye microinjection (Fig. 1). Individually labeled smooth muscle cells appeared as a narrow band that wrapped circumferentially around the arteriole; there was no evidence of dye spread into adjacent smooth muscle cells or underlying endothelial cells. In contrast, labeled endothelial cells appeared as a band parallel to the vessel axis. Typically, dye spread from the injected cell into many endothelial cells along the vessel axis, with little circumferential labeling; there was no evidence of dye coupling to surrounding smooth muscle cells.
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Resting conditions.
The diameter of arterioles at rest was 47 ± 1 µm
(n = 63); all vessels studied
exhibited brisk and reversible dilation (typical magnitude, 20-30
µm) to the addition of sodium nitroprusside (10 µM) to the
superfusate. Vessel diameters were unaltered with successful impalement
with microelectrodes. The
Em of endothelial
cells was 36 ± 1 mV (n = 40)
and that of smooth muscle was 35 ± 1 mV
(n = 23); these values are not
different from those recorded in preliminary experiments using
microelectrodes filled with 1 M KCl (data not shown). With constant
arteriole diameter at rest,
Em often
fluctuated slightly (by 2-3 mV), particularly when recording from
smooth muscle cells.
Local and conducted vasoconstriction.
Microiontophoresis of NE or PE elicited vasoconstriction that was
conducted along the arteriole wall (Figs. 2
and 3). A rapid depolarization of smooth
muscle cells preceded the vasomotor response by ~2 s; repolarization
was typically slower and corresponded with the return to resting
diameter. In contrast to these consistent effects on smooth muscle
cells, neither NE nor PE had any effect on the
Em of endothelial
cells (Figs. 2 and 3). Nevertheless, these same endothelial cells
promptly depolarized (to 13 ± 2 mV;
n = 18) when superfusate KCl was
raised to 55 mM. Such dichotomous electrical responses to NE release
were also found in preliminary recordings using KCl microelectrodes
(data not shown). When KCl was released from a stimulus pipette,
similar depolarizations were initiated in both endothelial and smooth
muscle cells, and these conducted readily along both cell layers (Fig.
4).
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Local and conducted vasodilation. ACh release triggered the conduction of vasodilation along the arteriole. Vasomotor responses to ACh were preceded (2-3 s) by hyperpolarization in endothelial cells and in smooth muscle cells (Fig. 5). Whereas the onset of hyperpolarization was rapid in both cell types, the shape and duration of electrical responses varied markedly and characteristically between cell types. Typically, endothelial cell hyperpolarization was followed by an equally rapid repolarization that occurred at or before the peak of the dilation. In contrast, smooth muscle cell repolarization was prolonged and typically (in 6 of 8 experiments) followed by an "afterdepolarization" that increased (magnitude 2-19 mV) with stimulus intensity. As a consequence, the duration of the entire electrical response (see Fig. 5) in smooth muscle cells was approximately twice that of endothelial cells (P < 0.05; t-tests) when expressed as a percentage of the duration of vasomotor response for respective cell types (local: 79 ± 8 vs. 37 ± 4%; conducted: 82 ± 5 vs. 44 ± 4%).
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Summary data. Diameter and Em responses to NE, PE, KCl, and ACh are summarized in Fig. 6. Note that electrical responses were monitored over a full range of vasomotor responses by varying the pulse duration of respective stimuli. The magnitude of the vasomotor response in an endothelial cell or a smooth muscle cell typically increased with the magnitude of the electrical response, indicating a role for electromechanical coupling in response to each stimulus tested. The notable exception was the lack of endothelial cell response to NE and PE, in which no change in electrical activity was recorded throughout a wide range of vasoconstriction.
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DISCUSSION |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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The conduction of vasomotor responses reflects the electrotonic spread
of current along the arteriolar wall. This study has distinguished the
cellular pathways by which electrical responses are conducted in vivo.
Using the hamster cheek pouch preparation, we show that
-adrenoceptor agonists initiated a depolarization on and conducted
between smooth muscle cells, independent of endothelial cells. In
contrast, ACh elicited hyperpolarization that spread along both the
endothelial cell layer and the smooth muscle cell layer. However, the
shape and the duration of hyperpolarization varied substantively and
characteristically between the respective cell types. Our findings
indicate that current can readily pass between adjacent endothelial
cells or adjacent smooth muscle cells; thus homologous electrical
coupling is present in vivo. In contrast, electrical coupling between
endothelial and smooth muscle cells was not prevalent during blood flow
control. In light of these findings, we propose that the endothelial
cell layer and the smooth muscle cell layer can each function as a
distinct electrical pathway. In vivo, the initial stimulus may thereby
determine which electrical pathway(s) may be used in signal
transmission along the arteriolar wall.
Cell labeling. The lack of Lucifer yellow dye coupling between smooth muscle and endothelial cells in cheek pouch arterioles (Fig. 1) confirms earlier reports (19, 26). In the present study, this property has enabled us to rigorously define the cell type from which electrical recordings were obtained. Whereas the spread of dye between endothelial cells indicates gap junctional coupling (26), the absence of dye coupling in smooth muscle cells (Fig. 1) does not reflect the electrical coupling clearly present within this cell layer (Figs. 2 and 3). Such discrepancy between electrical coupling and dye coupling (i.e., conductance vs. permeability of gap junctions) can be explained by the permselectivity of respective gap junctions to ions versus dye molecules (6, 19, 32). Lucifer yellow has been shown to be a nontoxic, intensely fluorescent dye that is readily visualized in living cells (31). Although this agent has recently been reported (19) to impair dye coupling in arteriolar smooth muscle cells, it has no effect on electrophysiological responses or on cell-to-cell electrical coupling (31). Moreover, this dye remains widely used to identify the electrophysiological properties of specifically labeled cells. In the present experiments, if Lucifer yellow dye could effectively block electrical coupling, then a labeled smooth muscle cell should have been electrically isolated from other smooth muscle cells. It is evident from findings in Figs. 2, 3, and 4 that electrical responses were readily and consistently conducted into and beyond smooth muscle cells that were labeled with this dye.
Em at rest.
At rest, the Em
of endothelial and smooth muscle cells averaged 36 and 35
mV, respectively. These resting values are near the equilibrium
potential for chloride [approximately 20 to 30 mV
(12, 24)], suggesting that endothelial and smooth muscle cells
may have considerable chloride permeability under physiological conditions. However, if endothelial and smooth muscle cells are "clamped" at or near chloride's equilibrium potential, then
fluctuations in K+ and
Na+ permeability may have little
effect on Em, and
one would predict stable recordings at rest. In light of the rapidity
of electrical responses when stimulated (see
RESULTS) and the fluctuations in Em observed at
rest, we suggest that
Em largely
reflects (changes in) the relative permeabilities to cations (7, 9, 28, 33).
50 mV) compared with arterioles that had not
been damaged (unpublished observations). Thus loss of pressure and
physical trauma may increase K+
conductance in both endothelial and smooth muscle cells, producing resting potentials that are closer to the equilibrium potential for
K+ than exist in arterioles
actively controlling blood flow. With hyperpolarization and loss of
vasomotor tone, the ability to study the conduction of vasodilation is
precluded. Hence, electrophysiological studies of conduction in
isolated arterioles have focused on depolarization and vasoconstriction
(29, 37, 38).
Local and conducted vasoconstriction.
Focal activation of adrenoceptors initiated a characteristic
depolarization of smooth muscle cells that, within ~2 s, gave rise to
vasoconstriction (Figs. 2 and 3). At both local and conducted sites,
these electrical events were rapid in onset and were followed by a
slower phase of repolarization that coincided with the return to
resting diameter. The similarity between the effects of NE and PE
observed (Figs. 2 and 3), together with the selectivity of PE for the
1-adrenoceptors, indicates that
NE acted primarily through the
1-subtype. These depolarizing
responses were observed through a corresponding range of
vasoconstriction (Fig. 6) and may be explained by the activation of a
cation channel permeable to both
Na+ and
Ca2+ (33), along with a reduction
in K+ conductance (22).
Nevertheless, the ionic bases of these responses remain to be defined.
1-adrenoceptors (or their
signaling pathway for depolarization) were not present on endothelial
cells and that smooth muscle cells were not electrically coupled to
endothelial cells. It is important to recognize that these findings
cannot be explained by the inability of endothelial cells to depolarize
in vivo, because superfusing the cheek pouch with 55 mM KCl depolarized
these same cells by an average of 23 mV.
The inability of PE to influence the
Em of endothelial
cells is in contrast to a recent in vitro study (38) in which this agonist was found to elicit depolarization in endothelial as well as
smooth muscle cells. On the basis of the similarity of electrical responses between cell types and the assumption that PE activated 1-adrenoceptors only on smooth
muscle cells, it was concluded that myoendothelial gap junctions (i.e.,
heterologous cell couplings) were present in cheek pouch arterioles and
that this enabled the rapid spread of current from smooth muscle to
endothelium (38). As an explanation of such a discrepancy with the
present findings, it is possible that myoendothelial junctions are
indeed present but are effectively closed under physiological
conditions. As a result, current would not pass between the two cell
types, as observed in the present study (Figs. 2 and 3). We hypothesize that myoendothelial gap junctional conductance could be reduced by
cyclic nucleotides, Ca2+, or
nitric oxide (20, 30); the production of these second messengers is
sensitive to both pressure and flow. Alternatively, the input
resistance of endothelial cells in vivo may be considerably less than
that of smooth muscle cells; this could selectively limit the effect of
current spread from smooth muscle cells on endothelial cell
Em (6). Finally,
high local concentrations of PE may well activate an endothelial cell
adrenoceptor (11). If such receptors were coupled to chloride channels,
then cells with a resting
Em near 65
mV will rapidly depolarize, whereas those in the present study, which
rest closer to chloride's equilibrium potential, may experience little
or no change in
Em. Because this interpretation remains speculative, it is clear that additional work is
necessary to resolve the differences between in vivo and in vitro
findings.
The release of KCl from a micropipette initiates the conduction of
vasoconstriction and depolarization in arterioles (28, 29, 37, 38).
Figure 4 uniquely illustrates that in vivo electrical responses to KCl
were quite similar between smooth muscle and endothelial cells.
Moreover, the onset of the electrical response preceded the vasomotor
response by ~2 s, as seen with the adrenoceptor agonists. Unlike NE
and PE, which must first interact with receptors to elicit a cellular
response, the elevation of external
K+ produces depolarization through
a Nernst effect via changes in the
K+ equilibrium potential. Although
KCl has been useful in defining the contractile behavior of vascular
smooth muscle (23) and the cable properties of arterioles (29), its
lack of specificity on the
Em of excitable
cells disqualifies it as a probe for myoendothelial coupling.
Nevertheless, the consistency of the delay between the onset of
depolarization and vasoconstriction across these stimuli indicates that
a common final pathway is used in the process of electromechanical
coupling (37).
Local and conducted vasodilation. Whereas ACh triggered hyperpolarization in both endothelial and smooth muscle cells, there was a concomitant and characteristic difference between respective cell types (Fig. 5). The endothelial response was rapid in both onset and recovery; in contrast, the smooth muscle response was prolonged and was followed by an afterdepolarization. These electrophysiological "fingerprints" were maintained with high fidelity as hyperpolarization spread along the arteriolar wall (Fig. 5). Moreover, as discussed earlier with depolarization and constriction, the onset of the electrical response to ACh preceded that of the vasomotor response by ~2 s. Thus the opening and closing of voltage-sensitive Ca2+ channels (on depolarization and hyperpolarization, respectively) in smooth muscle cells appears coupled, with similar time course, to the activation and inhibition of smooth muscle cell contraction. In turn, this suggests that, under physiological conditions, electromechanical coupling effectively contributes to the control of both increases and decreases in tissue blood flow.
ACh acts on muscarinic receptors of endothelial cells to elicit hyperpolarization (9, 10, 25). In contrast, the corresponding response of smooth muscle cells is indirect and depends on the release of endothelial-derived hyperpolarizing factors (EDHF) (5, 10) or the spread of current between endothelial cells and smooth muscle cells via myoendothelial gap junctions (19, 38). If heterologous cell coupling is present in cheek pouch arterioles, then ACh should have elicited parallel electrical responses in both cell types. Figure 5 illustrates that hyperpolarization was rapid in endothelial cells and in smooth muscle cells; however, the shape and duration of these respective responses did not parallel each another. Furthermore, when expressed relative to the vasomotor response, the duration of the electrical response in smooth muscle cells was nearly twice as long as in endothelial cells. In light of these observations, we suggest that smooth muscle hyperpolarization at the site of ACh release does not depend on the transfer of current between endothelium and smooth muscle but instead reflects the rapid release of an EDHF (3, 8, 34). In turn, the basal release of an autacoid (in contrast to heterologous electrical coupling) may explain the similarity of Em between cell types at rest. The spread of hyperpolarization was clear in both cell layers, with each cell type maintaining a distinctive electrical response (Fig. 5). This distinction of electrical responses between cell layers reinforces our view that endothelial cells are not electrically coupled to smooth muscle during blood flow control. Our findings with ACh raise the question of whether endothelial cell hyperpolarization per se gives rise to smooth muscle cell relaxation; i.e., does the spread of hyperpolarization along the endothelium give rise to the release of EDHF along the arteriolar wall? Alternatively, the release of an EDHF at the site of ACh application could generate an electrical signal that then spreads along the smooth muscle cell layer in parallel with the signal spreading along the endothelium (see Figs. 4 and 5). Assuming that the half-life of the EDHF is shorter than that for ACh released from a pipette, we speculate that the afterdepolarization of smooth muscle cells may provide insight into this dichotomy. Thus, once the action of an EDHF wanes at the site of ACh release, the afterdepolarization may reflect the direct actions of any residual ACh on smooth muscle cells (4). Because the direct effect of ACh is present only where the agonist is released (15, 27, 28), the presence of afterdepolarization along the vessel implies that that electrical events in smooth muscle originated locally and were conducted along the smooth muscle layer. In contrast, if EDHF was released along the vessel as hyperpolarization spread along the endothelium, the afterdepolarization should not have been present at conducted sites. Although six of eight experiments were consistent with this explanation, additional studies are required to resolve this matter. In summary, the present data represent the first cell-specific electrophysiological records of conduction obtained in arterioles controlling blood flow.-Adrenoceptor and muscarinic agonists elicited local changes in
Em that were
readily conducted along the vessel wall. Specifically, NE and PE
initiated a depolarization that spread along smooth muscle cells
independent of endothelial cells. In contrast, ACh elicited
hyperpolarizing responses that were distinct between cell types and
were conducted along the respective cell layers. Our findings indicate
that current flow along the wall of arterioles occurs through
homologous coupling between endothelial cells and between smooth muscle
cells; heterologous coupling between endothelial and smooth muscle
cells was not prevalent. We propose that the endothelium and the smooth
muscle monolayers form parallel electrical pathways along the arteriole
wall. Whereas hyperpolarization and depolarization can rapidly spread
along both cellular pathways, it appears that the particular stimulus will determine which cell layer is used for signaling. Differences between the behavior of similar vessels studied in vitro may be explained by the loss of physiological regulation of gap junctional coupling.
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ACKNOWLEDGEMENTS |
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We are grateful to Dr. Timothy O. Neild for contributions.
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FOOTNOTES |
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This study was supported by National Heart, Lung, and Blood Institute Grants RO1-HL-41026 and RO1-HL-56786. This work was completed during the tenure of an Established Investigatorship Award from the American Heart Association and Genentech, Inc.
Address for reprint requests: S. S. Segal, The John B. Pierce Laboratory, Yale Univ. School of Medicine, 290 Congress Ave., New Haven, CT 06519 (E-mail: sssegal{at}jbpierce.org).
Received 24 June 1997; accepted in final form 8 September 1997.
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REFERENCES |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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, local endothelial
responses; 
, conducted endothelial responses; 
, local smooth
muscle responses; 
, conducted smooth muscle responses. Data were
obtained from a total of 40 endothelial cells and 23 smooth muscle
cells.