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1 Department of Physiology and Pharmacology, Wake Forest University School of Medicine, Winston-Salem, North Carolina 27157; and 2 Department of Pharmacology and Toxicology, Medical College of Georgia, Augusta, Georgia 30912
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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We assessed the effect of epoxyeicosatrienoic acids
(EETs) in intact mesenteric arteries and Ca2+-activated
K+ (BKCa) channels of isolated vascular
smooth muscle cells from control and insulin-resistant (IR) rats. The
response to 11,12-EET and 14,15-EET was assessed in small mesenteric
arteries from control and IR rats in vitro. Mechanistic studies were
performed in endothelium intact or denuded arteries and in the presence
of pharmacological inhibitors. Moreover, EET-induced activation of the
BKCa channel was assessed in myocytes in both the
cell-attached and the inside-out (I/O) patch-clamp configurations. In
control arteries, both EET isomers induced relaxation. Relaxation was
impaired by endothelium denudation,
N
-nitro-L-arginine, or
iberiotoxin (IBTX), whereas it was abolished by IBTX + apamin or
charybdotoxin + apamin. In contrast, the EETs did not
relax IR arteries. In control myocytes, the EETs increased BKCa activity in both configurations. Conversely, in
the cell-attached mode, EETs had no effect on BKCa
channel activity in IR myocytes, whereas in the I/O configuration,
BKCa channel activity was enhanced. EETs induce
relaxation in small mesenteric arteries from control rats through
KCa channels. In contrast, arteries from IR rats do
not relax to the EETs. Patch-clamp studies suggest impaired relaxation
is due to altered regulatory mechanisms of the BKCa channel.
calcium-dependent K+ channels; vascular smooth muscle; endothelium
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INTRODUCTION |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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INSULIN RESISTANCE AND HYPERINSULINEMIA are common in patients with essential hypertension and are implicated in the pathogenesis of this disease as well as in its complications (8, 13, 20). Although the mechanisms that link insulin resistance and vascular dysfunction remain unclear, impairment of endothelial function has been proposed as one potential mechanism. This hypothesis is supported by studies in both insulin-resistant human subjects (27) as well as animals (16, 21, 28) that demonstrated impaired endothelium-dependent relaxation.
Previous studies (21, 22, 28) using the fructose-fed rat model of insulin resistance have documented an impaired endothelium-dependent relaxation, as defined by a decreased response to acetylcholine and/or bradykinin, in small mesenteric and coronary arteries. Furthermore, this impaired endothelium-dependent relaxation is related to a defect in a nitric oxide/prostacyclin-independent relaxing factor that induces vasodilation through activation of Ca2+-dependent K+ channels (KCa) (17). On the basis of the current literature (23, 25), this relaxation is likely due to endothelium-derived hyperpolarizing factor (EDHF).
To date, the identity of EDHF is unclear. On the basis of the current literature, it is likely that there is more than one definitive EDHF depending on the animal species and vascular bed studied. However, consistent throughout the majority of data is the suggestion that EDHF is a metabolite of arachidonic acid (3, 9, 12). Although this area remains controversial, considerable evidence (19, 24, 26, 33) has clearly shown that arachidonic acid metabolites of cytochrome P-450 monooxygenase enzyme system, such as epoxyeicosatrienoic acids (EET) and their dihydroxyeicosatrienoic acid metabolites, exhibit EDHF-like activity in coronary, cerebral, renal, and mesenteric arteries of various species.
A recent study (18) by our laboratory has demonstrated that endothelium-dependent and EDHF-mediated relaxation can be restored in small mesenteric arteries by induction of the cytochrome P-450 monooxygenase enzyme system. These data suggest that decreased EDHF production is the mechanism for impaired endothelium-mediated relaxation; however, other issues could be involved, such as an enhanced breakdown of EDHF or impaired K+ channel function on vascular smooth muscle (VSM) or the endothelium. The current study was designed to assess the ability of the EETs, a putative EDHF, to induced relaxation in small mesenteric arteries of control and insulin-resistant rats.
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METHODS |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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The animal care committees at Wake Forest University School of Medicine and the Medical College of Georgia approved the current protocol. Male Sprague-Dawley rats were obtained at 6 wk of age and randomized into one of the following two groups: 1) insulin-resistant (n = 24) and 2) control (n = 40) rats. Animals in the insulin-resistant group were fed a fructose-rich diet containing 66% fructose, 22% casein, and 12% lard, plus essential vitamins and minerals (Teklad Labs; Madison, WI), whereas control animals received standard rat chow.
After a 4-wk diet treatment, the rats (in a fasting state) were
anesthetized with pentobarbital sodium (50 mg/kg ip) and anticoagulated with heparin sodium (500 units ip). A midline incision was made and the abdominal and chest cavities were opened. Approximately 1 ml of
blood was removed for evaluation of insulin and glucose concentrations.
Hyperinsulinemia was used as a marker of insulin resistance in this
model (31). Subsequently, a section of the small intestine
was clamped, removed, and placed in a chilled oxygenated modified
Krebs-Ringer bicarbonate solution concentration composed of (in mM)
118.3 NaCl, 4.7 KCl, 2.5 CaCl2, 1.2 MgSO4, 1.2 KH2PO4, 25 NaHCO3, and 11.1 dextrose. Fourth-order branches of the superior mesenteric artery
(
210 µm in diameter) were isolated from surrounding perivascular
tissue and removed from the mesenteric vascular bed for either
functional studies or VSM patch-clamp experiments.
Determination of vascular reactivity. Small mesenteric arteries (~2 mm in length) isolated from the mesenteric vascular bed were transferred to a vessel chamber and mounted and secured between two glass micropipettes with a 10-0 ophthalmic suture. The vessel chamber was transferred to an inverted light microscope stage coupled to a video dimension analyzer (Living Systems Instrumentation; Burlington, VT). The video dimension analyzer was connected to both a video monitor (for visualization of the vessel) and to a strip-chart recorder (Kipp and Zonen) for constant recording of the intraluminal diameter of the vessel. Oxygenated (20% O2-5% CO2) Krebs solution, maintained at 37°C, was continuously circulated through the vessel bath. In addition, the lumen of the vessel was filled with Krebs solution through the micropipettes and maintained at a constant pressure of 40 mmHg. Only one concentration-response experiment was performed per artery; however, several arteries were taken from each rat.
Mesenteric arteries were allowed to equilibrate for 30 min and subsequently preconstricted to ~40% of their resting diameter with phenylephrine, an
1-receptor agonist.
Concentration-response experiments were performed with two of the EET
regioisomers: 11,12-or 14,15-EET (1 × 10
10 to
3 × 106 M) or vehicle (ethanol) in arteries from
both control and insulin-resistant rats. These two regioisomers were
chosen based on preliminary experiments where their effects were not
altered by inhibition of cyclooxygenase products with indomethacin
(data not shown). To determine the mechanism of EET-induced relaxation
in this vascular bed, additional studies were performed with 14,15-EET.
Just one regioisomer was chosen as a prototype EET because both EETs
studied induced similar responses in control arteries and the
substantial costs of performing all of the mechanistic studies with
both agents. To determine the role of the endothelium in EET-induced
relaxation, arteries were denuded of endothelium before the
concentration response experiment with 14,15-EET. Endothelial
denudation was performed by perfusing air through the lumen of the
artery. Endothelial disruption was verified by the absence of a dilator
response to acetylcholine and viability was tested by vasodilator
response to nitroprusside. In addition, these pharmacological tests
were verified by electron microscopy (Fig.
1). To determine the role of
Ca2+-dependent potassium channels (KCa)
arteries were pretreated with iberiotoxin (IBTX) (0.1 µM), IBTX (0.1 µM) + apamin (0.5 µM) or charybdotoxin (CTX) (0.1 µM) + apamin (0.5 µM). IBTX was used to specifically inhibit large
conductance KCa, whereas the combination of
IBTX + apamin was used to assess the role of large and small conductance KCa. CTX is a nonspecific antagonist of
KCa, whereas apamin is an antagonist of the
small-conductance KCa. The combination of CTX + apamin was used because it has been shown to inhibit EDHF, likely via
its effects on the intermediate and small-conductance KCa (1, 5). Finally, to determine
whether EETs induce relaxation via the release of nitric oxide,
arteries were pretreated with N-nitro-L-arginine
(L-NNA, 100 µM).
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VSM patch-clamp experiments. Myocytes were isolated from identical vessels used in the functional studies as previously described (4). Briefly, the endothelium was removed and the adventitia of the arteries was carefully teased away under a microscope. The remaining smooth muscle-rich media layer was enzymatic digested. The muscles were incubated at 37°C in a solution consisting of (in mg) 6 papain, 4 dithiothreitol, 2 collagenase, and 0.02% bovine serum albumin. After 30 min of gentle shaking, the muscle strips were lightly triturated, and the enzyme solution was diluted by the addition of excess enzyme-free solutions. The solution was removed and centrifuged at 500 rpm for 15 min. The pellet was resuspended in fresh medium composed of (in mM) 110 NaCl, 5 KCl, 2 CaCl2, 2 MgCl2, 10 HEPES, 10 NAHCO3, 0.5 KH2PO4, 10 glucose, 0.49 EDTA, and 10 taurine. The pellet was kept at 4°C. The recordings were performed within 6-8 h after cell dissociation.
Small aliquots of cell suspension were placed in a recording chamber (Warner Instruments). The external recording bath solution was composed of the following (in mM): 140 KCl, 10 MgCl2, 0.1 CaCl2, 10 HEPES, and 30 glucose (pH 7.2, 25°C). A gigaohm seal between the cell and the pipette was formed and a pCLAMP 7 (Axopatch 200B Amp, Axon Instruments) amplifier was used to record the current. Current and voltage traces were digitized with Digidata 1200 series (Axon Instruments) and stored for analysis. Capacitative and leakage currents were subtracted digitally. Single-K+ channels were measured in cell-attached or inside-out patches. In cell-attached configuration, the patch pipette (2-5 M
) was filled with a Ringer solution containing (in mM) 140 NaCl, 5 KCl, 2 MgCl2, 2 CaCl2, 20 HEPES, and 20 glucose (pH
7.2, 25°C), and a gigaohm seal was made on an intact cell to measure
channel activity at a voltage of +50 mV. The effect of 11,12-EET and
14,15-EET was determined at a concentration of 1 µM.
In the experiments measuring K+ channel-activity in
cell-free inside-out patches, the bathing solution exposed to the
cytoplasmic surface of the membrane composed of (in mM) 60 K2SO4, 30 KCl, 2 MgCl2, 0.16 CaCl2, 1 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid
(pCa 7), 10 HEPES, 5 ATP, and 10 glucose (pH 7.4, 22-25°C). The
solution in contact with the external surface was the Ringer solution
described above. Average channel activity in patches with multiple
Ca2+-activated K+ (BKCa)
channels was determined as described previously (29). For
the inside-out experiments, the effect of 11,12-EET and 14,15-EET was
determined at a concentration of 1 µM.
Biochemical measurements. Plasma insulin was assayed by using a dextran-coated charcoal immunoassay with rat antibody. Glucose concentrations were measured using a Glucose Trinder Kit (Sigma; St. Louis, MO).
Chemicals. The EETs were obtained from Cayman Chemicals. For all experiments, the EETs were kept in the dark and on ice to minimize metabolism. All other chemicals were obtained from Sigma. All agents were dissolved in deionized water and diluted with Krebs buffer. L-NNA was dissolved in water and titrated to a pH of ~2 with hydrochloric acid for dissolution. The pH was then titrated to physiological level (7.4) with sodium hydroxide.
Data analysis. Data from vascular reactivity studies are expressed as a percentage of relaxation after preconstriction. All data are expressed as means ± SE. All concentration response curves were evaluated for changes in maximal response and differences at each concentration using analysis of variance with repeated measures, followed by Fisher's pairwise least-significant difference test for multiple comparisons. Statistical comparison between groups for patch-clamp experiments was evaluated by Student's t-test. The criteria for significance were P < 0.05.
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RESULTS |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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Vascular reactivity experiments. Resting intraluminal diameter of small mesenteric arteries did not differ between groups (207 ± 6 µm for control and 212 ± 4 µm for insulin-resistant arteries). Moreover, the percentage of arterial constriction after phenylephrine was similar between groups with 42 ± 2% for control and 41 ± 2% for the insulin-resistant group. Neither endothelial denudation nor pharmacological inhibition significantly altered the resting diameter compared with the arteries in the control group. The percentage of constriction in experiments with endothelial denudation or pharmacological inhibition also did not differ compared with normal control arteries; however, the concentration of phenylephrine was reduced by one-half (from 200 to 100 µM) to produce the same degree of vasoconstriction.
In arteries from control animals, 11,12-EET and 14,15-EET induced a concentration-dependent relaxation (Fig. 2). By contrast, neither of the EET regioisomers induced relaxation in arteries from the insulin-resistant rats. In fact, a small but significant vasoconstriction was induced with both EET regioisomers (Fig. 3). Because the EETs did not induce relaxation in arteries from the insulin-resistant rats, no further functional experiments were performed with these arteries. It should be noted that the vehicle (ethanol) induced a small vascular relaxation in both control and insulin-resistant arteries (Figs. 2 and 3) that was significant versus time control (data not shown).
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Patch-clamp experiments.
In cell-attached patches of smooth muscle cells from control and
insulin-resistant mesenteric microvessels, the BKCa
channel exhibited a single-channel conductance of 142 ± 6 and
134 ± 3 pS, respectively. These values did not differ from one
another and were similar to what has previously been described
(4). The effects 11,12- and 14,15-EET on
BKCa channel opening probability were measured in
myocytes from control and insulin-resistant rats. In control cells,
both of the EET compounds significantly increased the
BKCa channel opening probability (Fig.
6). The addition of 11,12-EET
(n = 3) and 14,15-EET (n = 3) caused an
increase in BKCa channel opening probability by 60- and 79-fold, respectively. In contrast, these compounds had no effect
on channel opening probability in myocytes from insulin-resistant rats
recorded in the cell-attached configuration (Fig. 6).
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Biochemical measurements. Mean body wt (303 ± 8 g for control and 310 ± 6 g for insulin resistant) and fasting glucose (149 ± 11 mg/dl for control and 142 ± 8 mg/dl for insulin resistant) were similar among control and insulin-resistant rats. In contrast, fasting plasma insulin (97 ± 27 pmol/l for control and 234 ± 37 pmol/l for insulin resistant, P < 0.05) was significantly elevated in insulin-resistant rats compared with control.
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DISCUSSION |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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The current study assessed the effect of EETs on vascular relaxation and VSM BKCa activation in mesenteric arteries from control and insulin-resistant rats. There are several important findings herein. First, in control mesenteric arteries both of the EET regioisomers tested induced a similar concentration-dependent relaxation. Second, in myocytes from control mesenteric arteries the EETs enhanced the open probability of the BKCa channel both in the cell-attached and inside-out patch-clamp configurations. Third, although EET-induced relaxation in control arteries appears to be mostly due to their effect on VSM, the endothelium is also involved because endothelial denudation and inhibition of nitric oxide synthase reduced relaxation to 14,15-EET. Fourth, the BKCa channel is not the only KCa channel activated by the EETs because IBTX, a specific inhibitor of BKCa channels, reduced, but did not abolish, relaxation to 14,15-EET. In contrast, the combination of IBTX + apamin or CTX + apamin eliminated 14,15-EET-induced relaxation. Fifth, neither of the EET regioisomers induced relaxation in arteries from insulin-resistant rats. Likewise, in the cell-attached mode, EETs did not affect the open probability of BKCa channels in myocytes from insulin-resistant rats. Finally, both EETs increased BKCa channel open probability in inside-out patches of myocytes from insulin-resistant animals similar to that observed in control myocytes.
The present study provides direct evidence in intact microvessels and single myocytes from mesenteric arteries of control rats that EETs induce relaxation through KCa channels. This relaxation is primarily mediated through BKCa channels because a significant portion of EET-induced relaxation was inhibited by IBTX, a specific inhibitor of the BKCa channel. Moreover, we noted a marked increase in the open probability of the BKCa channel in the presence of the EETs. The relaxation that is resistant to IBTX appears to be due to activation of small conductance KCa channels because it was completely abolished by the combination of IBTX + apamin or CTX + apamin. The majority of studies (10, 12, 24) support these findings where EET-induced relaxation was inhibited by nonspecific antagonists of KCa (tetraethylammonium, tetrabutylammonium, and CTX) in porcine coronary, canine coronary, and cat cerebral arteries.
Regarding the effect of EETs on BKCa channels of control myocytes, we have shown that the EETs increase the open probability of the BKCa channel using both the cell-attached and inside-out VSM patch-clamp configurations. These data suggest that the EETs directly activate the channel without the need of G proteins or other second-messenger systems. Previous studies (14, 19) using the patch-clamp technique have also shown that the EETs activate VSM BKCa channels in control myocytes. In addition, they have been shown to activate BKCa in other tissues including porcine coronary endothelial cells (2) and pituitary GH3 cells (30). There have been two previous studies that have described the effect of EETs on VSM BKCa in the cell attached and inside-out patch-clamp configurations; however, their results differ from our own. Hu and Kim (14) assessed the effect of EETs on VSM cells from rabbit portal vein, rat caudal artery, guinea pig aorta, and porcine coronary artery. These investigators found that all EET isomers potentiated BKCa channel activity in the cell-attached mode. In contrast, these investigators found that the EETs had no effect in the inside-out configuration. Likewise, Li and Campbell (19) assessed EET-induced BKCa channel activity in bovine coronary VSM cells and found that BKCa channel activity was increased in the cell attached mode, but not in the inside-out configuration in the absence of guanosine 5'-triphosphate (GTP). However, in the presence of GTP, BKCa activity was enhanced in the inside-out configuration, leading these authors to conclude that EET-induced activation of BKCa involves GTP binding proteins. In the current study, we demonstrated that EETs could activate BKCa in the inside-out configuration in the absence of GTP. We are not sure why our findings differ from those of Li and Campbell; however, it may be explained by differences in animal model, vascular bed, artery size, or free Ca2+ content. In contrast to these studies, it has been shown in porcine coronary artery endothelial cells that the EETs enhance BKCa activation in the inside-out configuration without the addition of GTP (2). Similar findings are also reported in pituitary GH3 cells (30).
The current study also assessed the role of the endothelium in EET-induced relaxation of control arteries. We demonstrate that removal of the endothelium or pretreatment with L-NNA in control arteries diminishes EET-induced relaxation to a similar degree, suggesting that these two interventions eliminate the same mechanism of vasodilation. In addition, it is likely that endothelium-dependent EET-induced relaxation is mediated through activation of KCa channels because pretreatment with IBTX + apamin or CTX + apamin completely abolished vasodilation. In other words, it appears from the current data that EET stimulation results in the activation of endothelial cell KCa channels and the production of nitric oxide. These data are supported by a previous study (11) in cultured endothelial cells from bovine coronary arteries and human umbilical cord, where 5,6-EET increased intracellular Ca2+ to a similar degree as what is observed with bradykinin. In addition, this study showed that inhibiting the production of the EETs resulted in a decreased formation of nitric oxide (11). In contrast to these data and to our own, endothelial denudation of porcine and canine coronary arteries does not alter the vasodilatory response to the EETs (12, 24).
In contrast to experiments with control arteries, arteries from insulin-resistant rats did not relax to any of the EET regioisomers. In fact, a small but significant concentration-dependent vasoconstriction was induced with each of the three EET regioisomers. Recent data (7) suggest that this vasoconstriction may be due to the EETs ability to increase VSM intracellular Ca2+ by enhanced influx of extracellular Ca2+. We did not address this mechanism in the current study. It should be noted that we have previously shown that these arteries will respond normally to sodium nitroprusside (21), thus they do have the capability to vasodilate.
VSM patch-clamp studies of myocytes from insulin-resistant arteries reveal that the EETs do not affect the open probability of BKCa in the cell-attached mode. Conversely, the open probability of the BKCa channel was enhanced by the EETs in the inside-out mode similar to what was attained in control myocytes in this configuration. These findings suggest that BKCa channel itself is not altered in insulin-resistant arteries because the EET compounds are able activate the channel in the inside-out configuration. However, in myocytes from insulin-resistant arteries, there is an alteration in the regulatory mechanisms of the channel such that it cannot be activated in the cell-attached mode, nor can EETs induce relaxation in these arteries. We do not know the mechanism of this defect. However, we propose that it may be due to one of two possibilities. First, there may be a decreased availability of Ca2+ for channel activation. In the inside-out patch-clamp configuration, free Ca2+ is controlled by the experimental conditions; thus Ca2+ would always be available for channel activation. However, in the cell-attached configuration, free Ca2+ availability is dependent on the intracellular stores of the cell. Thus, whether the mechanism by which Ca2+ is released were dysfunctional or its access to the KCa channel impeded, then the channel would not be activated regardless of the stimulus. The second possibility is that mesenteric arteries from insulin-resistant rats produce a substance, such as 20-hydroxyeicosatetraenoic acid, that inactivates the KCa channels. This has previously been demonstrated in the renal arterioles of the rat (32). This product could impair channel activation in the cell-attached mode (and impair relaxation). However, when the cell membrane is ruptured to perform the inside-out patch-clamp configuration, the 20-hydroxyeicosatetraenoic acid inhibition is eliminated, and thus the channel can be activated. These hypotheses are only speculative at this time and will be addressed in future studies.
Thus it appears from the current findings that impaired relaxation in mesenteric arteries from insulin-resistant rats as previously described (17, 21) is not due primarily to endothelial dysfunction but rather due to an inability of the VSM K+ channels to respond to endothelium-derived relaxing factors. However, if KCa channels are involved in the production of EDHF and other endothelium-derived relaxing factors, as has been suggested (5, 6), then it may be that insulin resistance affects both the production and activity of endothelium-derived relaxing factors. Regardless, the KCa channels are important mediators of VSM vasodilation, particularly in smaller arteries (4, 15). The current findings may explain the mechanisms of how insulin resistance promotes hypertension and vascular dysfunction.
In summary, the EETs induce a concentration-dependent relaxation of small mesenteric arteries from control rats. In addition, the EETs enhance the open probability of the VSM BKCa channel in both the cell attached and inside-out configurations in control myocytes from rat mesenteric arteries. In contrast, the EETs induce a small vasoconstriction in small mesenteric arteries from insulin-resistant rats and they do not affect the VSM BKCa channel as assessed in the cell-attached patch-clamp configuration. Interestingly, the activation of the BKCa channel in the inside-out patch-clamp configuration is not different from control. Thus the impaired response to EETs in insulin-resistant arteries is not due to a direct BKCa channel dysfunction, but is more likely due to an alteration of the signal transduction pathways regulating the BKCa channel opening.
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ACKNOWLEDGEMENTS |
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This work was supported by an American Heart Association grant (to A. W. Miller, R. E. White, and G. O. Carrier) and by National Heart, Lung, and Blood Institute Grants HL-64779 (to G. O. Carrier and R. E. White), HL-54844 (to R. E. White), HL-30260, HL-46558, and HL-50587 (all to D. W. Busija). A. W. Miller is also supported by the American Foundation for Pharmaceutical Education.
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FOOTNOTES |
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Address for reprint requests and other correspondence: A. W. Miller, Dept. of Physiology/Pharmacology, Medical Center Blvd., Hanes 1002, Wake Forest Univ. School of Medicine, Winston-Salem, NC 27157.
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 14 February 2001; accepted in final form 29 June 2001.
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TOP
ABSTRACT
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METHODS
RESULTS
DISCUSSION
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