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Effects of hypoxia, anoxia, and metabolic inhibitors on K_ATP channels in rat femoral artery myocytes

Department of Human Anatomy and Cell Biology, School of Biomedical Sciences, Liverpool University, Liverpool, United Kingdom

Submitted 19 October 2005 ; accepted in final form 12 February 2006

	ABSTRACT

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Vascular ATP-sensitive potassium (K_ATP) channels have an important role in hypoxic vasodilation. Because K_ATP channel activity depends on intracellular nucleotide concentration, one hypothesis is that hypoxia activates channels by reducing cellular ATP production. However, this has not been rigorously tested. In this study we measured K_ATP current in response to hypoxia and modulators of cellular metabolism in single smooth muscle cells from the rat femoral artery by using the whole cell patch-clamp technique. K_ATP current was not activated by exposure of cells to hypoxic solutions (PO₂ 35 mmHg). In contrast, voltage-dependent calcium current and the depolarization-induced rise in intracellular calcium concentration ([Ca²⁺]_i) was inhibited by hypoxia. Blocking mitochondrial ATP production by using the ATP synthase inhibitor oligomycin B (3 µM) did not activate current. Blocking glycolytic ATP production by using 2-deoxy-D-glucose (5 mM) also did not activate current. The protonophore carbonyl cyanide m-chlorophenylhydrazone (1 µM) depolarized the mitochondrial membrane potential and activated K_ATP current. This activation was reversed by oligomycin B, suggesting it occurred as a consequence of mitochondrial ATP consumption by ATP synthase working in reverse mode. Finally, anoxia induced by dithionite (0.5 mM) also depolarized the mitochondrial membrane potential and activated K_ATP current. Our data show that: 1) anoxia but not hypoxia activates K_ATP current in femoral artery myocytes; and 2) inhibition of cellular energy production is insufficient to activate K_ATP current and that energy consumption is required for current activation. These results suggest that vascular K_ATP channels are not activated during hypoxia via changes in cell metabolism. Furthermore, part of the relaxant effect of hypoxia on rat femoral artery may be mediated by changes in [Ca²⁺]_i through modulation of calcium channel activity.

smooth muscle; ion channels; potassium channels

In the skeletal muscle circulation, hypoxic vasodilation in vivo is blocked by the K_ATP channel inhibitor glibenclamide (e.g., 19, 30). Vasodilation is endothelium dependent in small arteries and arterioles (e.g., 15, 31). Smooth muscle K_ATP channels may therefore be activated indirectly through the release of endothelium-dependent factors such as prostacylins and nitric oxide (15, 16, 31). However, endothelial removal only blunts hypoxic relaxation, and the smooth muscle cells must therefore possess intrinsic oxygen sensitivity (16). Indeed, hypoxic relaxation of larger arteries such as the main femoral artery is endothelium independent (6, 29). The role of K_ATP channels in hypoxic relaxation of these larger arteries is unknown.

Although the role of K_ATP channels in hypoxic vasodilation is well established at the tissue and organ level, only two studies have addressed how this occurs at a cellular level. Dart and Standen (9) have shown activation of K_ATP current by hypoxia in single smooth muscle cells from pig coronary arteries. This provides support for the proposal that hypoxic vasodilation in the heart is caused by direct activation of smooth muscle K_ATP channels, possibly through changes in cell metabolism (44). In contrast, Jackson (20) reported a lack of effect of hypoxia on K_ATP current in cells from arteries supplying rat cremaster muscle. Neither of these studies probed metabolic regulation of K_ATP channels. There is, therefore, a lack of clear information about how K_ATP channels are regulated by cellular metabolism or by changes in PO₂ in arterial smooth muscle cells. In particular, whether exposing single smooth muscle cells to a hypoxic environment changes nucleotide levels sufficiently to activate K_ATP channels has not been rigorously tested. In this study, we have addressed this issue by recording K_ATP currents on exposure of cells to solutions of differing PO₂ or containing the metabolic inhibitors oligomycin B, carbonyl cyanide m-chlorophenylhydrazone (CCCP), 2-deoxy-D-glucose (2-DG), or combinations of these agents.

	MATERIALS AND METHODS

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Cell isolation. Male Sprague-Dawley rats (200–300 g) were made unconscious by exposure to a rising concentration of CO₂ and killed by exsanguination in accordance with Schedule 1 of the Animals (Scientific Procedure) Act, 1986, under license from the Home Office. Single smooth muscle cells were dissociated from branches of the femoral artery, as previously described (36).

Electrophysiology. Membrane current was recorded in conventional or perforated patch whole cell patch clamp. For conventional whole cell recording, the pipette (intracellular) solution contained (mM) 107 KCl, 33 KOH, 10 HEPES, 10 EGTA, and 1 MgCl₂ (pH 7.2). Nucleotides were added to this solution on the day of the experiment, and pH was readjusted to 7.2 with 1 M NaOH. For perforated patch recordings, electrical access was obtained by including 0.2 mg/ml amphotericin B in the pipette solution, which contained (mM) 140 KCl, 10 NaCl, 1 MgCl₂, 0.1 EGTA, and 10 HEPES (pH adjusted to 7.2 with 1 M NaOH). In some experiments, 110 mM KCl was substituted with 110 mM potassium aspartate or potassium methanesulfonate. All seals were formed in an extracellular solution containing (mM) 6 KCl, 134 NaCl, 1 MgCl₂, 0.1 CaCl₂, 10 HEPES, and 10 glucose (pH adjusted to 7.4 with 1 M NaOH). The effects of hypoxia and metabolic inhibitors were tested under conditions of symmetrical 140 mM K⁺, where the extracellular solution contained (mM) 140 KCl, 1 MgCl₂, 0.1 CaCl₂, and 10 HEPES (pH adjusted to 7.4 with 1 M NaOH). The extracellular solution also contained 25 µM Ba²⁺, and, for the CCCP experiments, contained 2 mM tetraethylammonium (TEA⁺) chloride, to inhibit inward rectifier potassium channels and calcium-activated potassium channels, respectively. TEA⁺ and Ba²⁺ have modest effects on the K_ATP current at these concentrations (the half inhibition constant at –60 mV is 100 µM for Ba²⁺ and 7 mM for TEA⁺, 37). The 140 mM K⁺ extracellular solution was glucose free. In preliminary experiments, removing glucose had no effect on membrane current over the time course of our experiments (mean current at –60 mV in 140 mM extracellular [K⁺] solution was –33.5 ± 3.5 pA in 10 mM glucose and –35.5 ± 4.0 pA after 10 min in glucose-free solution, n = 6 cells). The experimental chamber (volume, 0.5 ml) was continuously perfused with extracellular solution at a rate of 2 ml/min. All experiments were conducted at room temperature.

Current amplitude was measured at a membrane potential of –60 mV in the different solutions. Current was measured by averaging >10 s of recording once steady state had been achieved. All results are given as means ± SE. Statistical significance was assessed by using ANOVA with Tukey's test for significance used as a post hoc analysis or by using a Student's t-test.

For calcium current recordings, the intracellular solution contained (mM) 130 CsCl, 10 CsF, 10 EGTA, 10 HEPES, 4 MgCl₂, and 4 Na₂ATP; pH readjusted to 7.2 with 1 M NaOH (14). The extracellular solution contained (mM) 130 NaCl, 10 TEACl, 10 HEPES, 1 MgCl₂, and 10 BaCl₂; pH readjusted to 7.4 with 1 M NaOH.

Hypoxia and anoxia experiments. The extracellular solution was made hypoxic by bubbling with 100% N₂ gas in a separate glass perfusion reservoir for at least 30 min before the experiment was started. The reservoir was connected to the experimental chamber by using oxygen-impermeable tubing (Norton Pharmed, 1/16 in. inner diameter, 1/16 in. wall thickness). Hypoxic solutions were perfused by gravity at a rate of 5 ml/min. PO₂ was measured in the experimental chamber by using a Strathkelvin Instruments 781 meter connected to a needle oxygen probe (Diamond General model 786-20R). Measured PO₂ in the chamber was in the range of 30–40 mmHg.

To generate anoxic conditions, the extracellular solution was first made hypoxic by bubbling with 100% N₂ gas, as before. Sodium dithionite (0.5 mM) was then added to this hypoxic solution, and the chamber was perfused. Reduction of O₂ by dithionite not only lowers PO₂ but also causes generation of free radicals (SO^2–) and hydrogen peroxide (2). This was prevented in our experiments by inclusion of bovine superoxide dismutase (100 U/ml) and catalase (2,000 U/ml) in the dithionite-containing solution (2).

Microfluorimetry. Intracellular calcium concentration ([Ca²⁺]_i) was reported by fura-2 in voltage-clamped single cells using a deltaRAM system (Photon Technology International), as previously described (22). The pipette solution contained (mM) 145 KCl, 10 HEPES, 3 MgCl₂, 3 Na₂ATP, and 0.05 fura-2 (pH adjusted to 7.2 with 1 M NaOH). The extracellular solution contained (mM) 134 NaCl, 6 KCl, 10 HEPES, 1 MgCl₂, and 3 CaCl₂ (pH readjusted to 7.4 with 1 M NaOH). Fluorescence emission was measured at a wavelength of 510 nm (40 nm band pass) at a frequency of 10 Hz in response to alternate excitation of the fluorophore with light at a wavelength of 340 nm (8 nm band pass) and 380 nm (8 nm band pass). Emission ratio was converted to free calcium concentration using an in vitro calibration, as previously described (22).

Mitochondrial membrane potential was measured in single cells that were not voltage clamped by using rhodamine 123. The extracellular solution contained (mM) 6 KCl, 134 NaCl, 1 MgCl₂, 0.1 CaCl₂, and 10 HEPES (pH adjusted to 7.4 with 1 M NaOH). Dye was loaded by incubating cells in a 10 µg/ml solution at room temperature for 10–15 min (22). The dye was excited at 500 nm (25 nm bandpass), and emission was measured at 545 nm (35 nm bandpass) at 10 Hz. Rhodamine 123 is a potentiometric dye that accumulates in mitochondria due to their negative membrane potential (33). Dye accumulation results in aggregation of dye molecules and quenching of the emission signal. Mitochondrial depolarization causes release of dye, with consequent unquenching and an increase in the fluorescence signal.

Myography. Isometric tension generation was measured in a small artery myograph (model 500A, Danish Myotechnology, Aarhus, Denmark). Femoral arteries were dissected in cold 5.4 mM K⁺ saline solution containing (mM) 137 NaCl, 5.4 KCl, 0.44 NaH₂PO₄, 0.42 Na₂HPO₄, 1 MgCl₂, 2 CaCl₂, 10 HEPES, and 10 glucose (pH adjusted to 7.4 with NaOH). Ring segments of artery were then mounted by threading two strands of tungsten wire (diameter 40 µm) through the vessel lumen. After vessel mounting and equilibration at 37°C were completed, passive tension was adjusted to allow measurement of active force production (32). Arteries were precontracted by adding the ₁-adrenoreceptor agonist phenylephrine (3 or 10 µM) to the 5.4 mM K⁺ saline. In some experiments, arteries were activated by an 80 mM K⁺ containing saline, which had the composition of the 5.4 mM K⁺ saline except 74.6 mM NaCl was substituted with KCl. The endothelium was removed from some arteries by rubbing a human hair through the lumen. Removal of a functional endothelium was confirmed by absence of relaxation to 10 µM acetylcholine.

Drugs. All salts were obtained from BDH or Sigma. Fura-2 and rhodamine 123 were obtained from Molecular Probes. Oligomycin B and CCCP were dissolved in DMSO as 3 and 10 mM stock solutions, which were added to the extracellular solution to give the appropriate final concentration.

	RESULTS

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K_ATP current in rat femoral artery smooth muscle cells. In initial experiments, the nucleotide sensitivity of the K_ATP current in these cells was investigated by dialyzing the cell with pipette solutions containing differing levels of ATP and ADP using conventional whole cell patch clamp (Fig. 1A). Whole cell recording was established in an extracellular solution containing 6 mM K⁺ and at a holding potential of –60 mV. The extracellular solution was then exchanged for one containing 140 mM K⁺, changing the driving force on K⁺ movement from an outward to an inward direction and resulting in development of an inward holding current (Fig. 1A). This reflects the basal K⁺ current in these cells. K_ATP current was then induced by the potassium channel opener pinacidil (10 µM). After pinacidil-induced current had developed, the K_ATP channel inhibitor glibenclamide (10 µM) was added to the extracellular solution. The magnitude of the pinacidil-induced current increased when ATP was lowered from 3.0 to 0.2 mM in the presence of 0.2 mM ADP or when ADP was raised from 0.2 to 1 mM in the presence 3 mM ATP (Fig. 1B). Rat femoral artery smooth muscle cells therefore express nucleotide-sensitive K_ATP channels similar to those identified in other vascular cells (37).

View larger version (14K): [in this window] [in a new window]   Fig. 1. Nucleotide modulation of ATP-sensitive K (KATP) current in rat femoral artery myocytes. A: conventional whole cell recordings of membrane current from a rat femoral artery myocyte. Cells were dialyzed with pipette solutions containing ATP and ADP at the concentrations (in mM) indicated beside the traces. Arrow, after cell recording was started, extracellular K+ was changed from 6 to 140 mM. Pinacidil (10 µM, Pin) and glibenclamide (10 µM, Glib) were added to the 140 mM K+ extracellular solution as indicated. Intracellular solution contained 140 mM K+. Holding potential was –60 mV. Calibration bar applies to both recordings. B: mean (±SE) current measured at different intracellular nucleotide concentrations in an extracellular solution containing: 6 mM K+, 140 mM K+, 140 mM K+ + 10 µM Pin, 140 mM K+ + 10 µM Pin + 10 µM Glib (n = 6–7 cells).

View larger version (13K): [in this window] [in a new window]   Fig. 2. Effect of hypoxia (Hypox) on KATP current in femoral artery myocytes. A: perforated patch recording of whole cell membrane current. Arrow, after cell recording was started, extracellular K+ was changed from 6 to 140 mM. After the current had reached a steady-state level, the cell was perfused with a hypoxic extracellular solution. Pin (10 µM) and Glib (10 µM) were added to the 140 mM K+ extracellular solution as indicated. Intracellular solution contained 140 mM K+. Dotted line, zero current. Holding potential was –60 mV. Hypoxia had little effect on current, even though there are functional KATP channels, as shown by response to Pin and Glib. B: mean (±SE) current from 7 experiments measured in an extracellular solution containing 6 mM K+, 140 mM K+, 140 mM K+ hypoxic solution, 140 mM K+ + 10 µM Pin, and 140 mM K+ + 10 µM Pin + 10 µM Glib.

View larger version (16K): [in this window] [in a new window]   Fig. 3. Inhibition of Ca2+ current and intracellular calcium ([Ca2+]i) by hypoxia. A: calcium current recorded in response to a depolarizing voltage step from –70 to 0 mV. Three superimposed traces are shown in normoxia and recovery (normox) and during exposure of the cell to a hypoxic solution (hypox). Ba2+ (10 mM) was present in the extracellular solution as charge carrier, and pipette contained 140 mM Cs+. B: time course of inhibition of peak Ca2+ current measured at 0 mV. C: hypoxic inhibition of [Ca2+]i. [Ca2+]i was measured by microspectrofluorimetry using the Ca2+-sensitive dye fura-2. Cell was gradually depolarized to activate calcium current, causing a sustained rise in [Ca2+]i. It was then superfused with a solution made hypoxic by bubbling with 100% N2. Superfusion with hypoxic solution caused [Ca2+]i to fall, an effect that was partially reversed on returning to normoxic solution.

View larger version (13K): [in this window] [in a new window]   Fig. 4. Effect of 2-deoxyglucose (2-DG) on KATP current (I). A: perforated patch recording of whole cell membrane current. Extracellular K+ was changed from 6 to 140 mM, as indicated. 2-DG (5 mM), Pin (10 µM), and Glib (10 µM) were added to the extracellular solution as indicated. Intracellular solution contained 140 mM K+. Dotted line, zero current. Holding potential was –60 mV. B: mean (±SE) data from 6 experiments measured in an extracellular solution containing 6 mM K+, 140 mM K+, 140 mM K+ + 5 mM 2-DG, 140 mM K+ + 10 µM Pin, and 140 mM K+ + 10 µM Pin + 10 µM Glib.

View larger version (14K): [in this window] [in a new window]   Fig. 5. Effect of oligomycin (Oligo B) on KATP current. A: perforated patch recording of whole cell membrane current. Extracellular K+ was changed from 6 to 140 mM, as indicated. Oligo B (3 µM), Pin (10 µM), and Glib (10 µM) were added to extracellular solution as indicated. Intracellular solution contained 140 mM K+. Dotted line, zero current. Holding potential was –60 mV. B: mean (±SE) data from 5 experiments measured in an extracellular solution containing 6 mM K+, 140 mM K+, 140 mM K+ + 3 µM Oligo B; 140 mM K+ + 10 µM Pin, and 140 mM K+ + 10 µM Pin + 10 µM Glib.

View larger version (17K): [in this window] [in a new window]   Fig. 6. Effect of carbonyl cyanide m-chlorphenyhdrazone (CCCP) on KATP current. A and B: perforated patch recording of whole cell membrane current. Extracellular K+ was changed from 6 to 140 mM, as indicated. CCCP (1 µM), Pin (10 µM), and Glib (10 µM) were added to the extracellular solution as indicated. Intracellular solution contained 140 mM K+. Dotted line, zero current. Holding potential was –60 mV. C: mean (±SE) data from 8 experiments measured in an extracellular solution containing 6 mM K+, 140 mM K+, 140 mM K+ + 1 µM CCCP, 140 mM K+ + 10 µM Pin, and 140 mM K+ + 10 µM Pin + 10 µM Glib. D: rhodamine 123 fluorescence signal recorded from a myocyte on exposure to CCCP (1 µM). Upward deflection reflects unquenching of the dye due to mitochondrial depolarization.

View larger version (14K): [in this window] [in a new window]   Fig. 7. Effect of Oligo B on CCCP-induced current. A: perforated patch recordings of whole cell membrane current. Extracellular K+ was changed from 6 to 140 mM, as indicated. Drugs were added to the extracellular solution at the concentrations indicated. Intracellular solution contained 140 mM K+. Dotted line, zero current. Holding potential was –60 mV. B: mean (±SE) data from 6 experiments measured in an extracellular solution containing 6 mM K+, 140 mM K+, 140 mM K+ + 1 µM CCCP + 300 nM Pin, 140 mM K+ + 1 µM CCCP + 300 nM Pin + 3 µM Oligo B, and 140 mM K+ + 1 µM CCCP + 300 nM Pin + 3 µM Oligo B + 10 µM Glib.

View larger version (15K): [in this window] [in a new window]   Fig. 8. Effect of combined application of CCCP and 2-DG on current. A: perforated patch recording of whole cell membrane current. Extracellular K+ was changed from 6 to 140 mM, as indicated. CCCP (1 µM) and 2-DG (5 mM) were added to the extracellular solution where indicated. Intracellular solution contained 140 mM K+. Dotted line, zero current. Holding potential was –60 mV. B: mean (±SE) data from 5 experiments measured in an extracellular solution containing: 6 mM K+, 140 mM K+, 140 mM K+ + 1 µM CCCP + 5 mM 2-DG, 140 mM K+ + 1 µM CCCP + 5 mM 2-DG + 10 µM Glib, and 140 mM K+ + 10 µM Pin.

View larger version (14K): [in this window] [in a new window]   Fig. 9. Effect of anoxia induced by dithionite on KATP current in femoral artery myocytes. A: perforated patch recording of whole cell membrane current. Arrow, after the recording was started, extracellular K+ was changed from 6 to 140 mM. After the current had reached a steady-state level, the cell was perfused with a hypoxic extracellular solution to which 0.5 mM sodium dithionite had been added. The solution also contained 100 U/ml bovine superoxide dismutase and 2,000 U/ml catalase. Glib (10 µM) was added to the extracellular solution as indicated. Intracellular solution contained 140 mM K+. Dotted line, zero current. Holding potential was –60 mV. B: mean (±SE) current from 6 experiments measured in an extracellular solution containing 6 mM K+, 140 mM K+, 140 mM K+ + 0.5 mM dithionite, 140 mM K+ + 0.5 mM dithionite + 10 µM Glib, and 140 mM K+ + 10 µM Pin. C: rhodamine 123 fluorescence signal recorded from a myocyte on exposure to dithionite (0.5 mM). Upward deflection reflects unquenching of dye due to mitochondrial depolarization.

View larger version (18K): [in this window] [in a new window]   Fig. 10. Effect of hypoxia on phenylephrine (PE) and 80 mM K+-induced contractions of femoral artery. A and B: recording of tension in a femoral artery mounted in a myograph exposed to hypoxic solution. Artery was contracted by PE (A) or an 80 mM K+-containing solution (B). C and D: mean maximal tension recorded from experiments such as those illustrated in A and B. rec, Recovery after hypoxia.

	DISCUSSION

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Given the known role for K_ATP channels in hypoxic vasodilation in the skeletal muscle circulation, our failure to activate the K_ATP current in response to hypoxia was initially surprising and led us to question whether the oxygen sensor in our cells was disrupted by the cell isolation procedure or if we were achieving appropriate levels of hypoxia in our experimental chamber. However, voltage-dependent calcium current and [Ca²⁺]_i were sensitive to hypoxia in our cells (Fig. 3). To understand why hypoxia did not activate K_ATP current, we further probed metabolic regulation of these channels. Surprisingly, blocking glycolytic or mitochondrial ATP production alone caused no significant measurable activation of current (Figs. 4 and 5). It appears that blocking ATP production is insufficient to activate the K_ATP current in our cells, at least over a period of 10 min. Indeed, even the mitochondrial uncoupler CCCP, which would be expected to result in rapid hydrolysis of cellular ATP (33), only caused moderate current activation compared with that induced by pinacidil (Fig. 6). The observation that K_ATP currents activated by combined application of 300 nM pinacidil and 1 µM CCCP were inhibited by 3 µM oligomycin B (Fig. 7) suggests that CCCP may be working by mitochondrial depolarization. Collapse of the mitochondrial membrane potential causes the ATP synthase to work in reverse mode, therefore, hydrolyzing cellular ATP (33). The activation of K_ATP current seen in our cells during exposure to anoxic solution is also consistent with this interpretation (Fig. 9). Anoxia, unlike hypoxia, will cause cessation of electron transport and collapse of the mitochondrial membrane potential, an effect we were able to measure (Fig. 9C). Therefore, not only will energy production cease due to lack of O₂ availability for enzymes of the mitochondrial respiratory chain, but also cellular ATP will be consumed by ATP synthase working in reverse mode (4, 24, 33, 39).

These patch-clamp experiments looking at the effects of hypoxia and metabolic inhibitors were conducted at room temperature, which may be unfavorable to seeing K_ATP channel activation. It is possible that K_ATP current activation might be seen if experiments were conducted at 37°C in mechanically loaded cells and for a longer period of exposure. A raised temperature along with mechanical loading would increase ATP consumption, which our results with CCCP and dithionite suggest is a critical determinant of K_ATP channel activation. However, arguing against this, the K_ATP channel inhibitor glibenclamide did not alter the hypoxic relaxation of intact femoral artery mounted in a myograph (Fig. 10). These results are consistent with our patch-clamp data and support the conclusion that hypoxia does not activate K_ATP channels even under conditions closer to the physiological situation.

Although K_ATP current is activated by hypoxia in pig coronary arteries (9), it is not clear that this is a consequence of changing intracellular nucleotide levels. Cellular respiration is generally thought to be independent of oxygen tension at levels that cause vasodilation (i.e., PO₂ values between 70 and 10 mmHg), because cytochrome oxidase is saturated with oxygen at these partial pressures (e.g., 46). Also, levels of oxygen that cause vasodilation do not cause large changes in intracellular ATP or ADP concentration in vascular tissue (e.g., 6, 10, 25, 28). However, the PO₂ at the level of the enzymes of the mitochondrial respiratory chain is unknown, although it will presumably be less than that in the bulk extracellular space due to diffusion gradients. Furthermore, submembrane nucleotide concentrations are unlikely to reflect bulk cellular concentrations (34). Thus there remains the possibility that hypoxia acts via changes in nucleotide levels. Alternatively, as suggested by Dart and Standen (9), other sensors directly sensitive to PO₂, including potentially the channel itself, may control K_ATP channel activity. Our data suggests that such a sensor does not operate in femoral artery smooth muscle cells.

Relation to previous work on oxygen regulation of cardiac myocyte K_ATP current. K_ATP channel regulation by oxygen tension and by cell metabolism is more clearly understood in cardiac muscle than in smooth muscle. In cardiac cells, a consensus view has emerged that ATP must fall to low levels before current activation is seen (e.g., 26). This requires exposure of cells to complete anoxia, which, unlike hypoxia, will lead to mitochondrial uncoupling and ATP hydrolysis by reverse mode operation of the ATP synthase. Hypoxia did not cause mitochondrial depolarization in our cells, suggesting that mitochondrial uncoupling is not occurring (T. Kamishima and J. M. Quayle, unpublished observations). Even during exposure of cardiac cells to complete anoxia, activation of K_ATP current occurs only after a considerable delay [>10 min (4, 24); 5 min (39)]. The studies cited used conventional whole cell patch-clamp recording, with the cell being dialyzed with an ATP-free pipette solution at a temperature of 35–37°C. This would favor a rapid decline in intracellular ATP levels compared with the conditions used in this study or that of Dart and Standen (9) (i.e., room temperature, perforated patch recordings). That said, no simple comparison is possible between the two cell types. Cardiac myocytes are more reliant on oxidative energy conversion, whereas smooth muscle depends more on glycolytic production (34). Cardiac myocytes also have greater energy reserves in the form of creatine phosphate (11). Finally, cardiac K_ATP channels are composed of the inward rectifier potassium channel Kir6.2 and the sulfonylurea receptor SUR2A, whereas vascular channels are probably composed of Kir6.1 and SUR2B (7).

Mechanism of hypoxic vasodilation in rat femoral arteries. Although our main objective in this study was to probe metabolic and hypoxic regulation of K_ATP current in single vascular myocytes, we also investigated the role of these channels in hypoxic vasodilation in intact arteries. Glibenclamide had no effect on the hypoxic relaxation of arteries precontracted with the ₁-adrenoreceptor agonist phenylephrine (Fig. 10). Arteries precontracted with 80 mM K⁺ also relaxed to hypoxia. In this solution the membrane potential will be clamped at the potassium equilibrium potential, effectively excluding potassium channel activation as a mechanism of hypoxic relaxation. The experiments illustrated in Fig. 10 were carried out under more physiological conditions than the patch-clamp experiments and are consistent with them. Overall, the data do not support a role for direct activation of K_ATP channels by hypoxia in smooth muscle cells of larger skeletal muscle arteries.

Despite the above, the role of K_ATP channels in hypoxic vasodilation in the skeletal muscle circulation is well established from functional studies (e.g., 15, 16, 19, 30). In particular, there is a body of evidence that K_ATP channels are involved in hypoxic relaxation of resistance-sized arteries from the skeletal muscle circulation (e.g., 15, 16, 19, 30). However, published observations suggest that the opening of K_ATP channels in these vessels is secondary to release of endothelial dilators, rather than through direct activation of smooth muscle K_ATP channels. For example, in isolated and pressurized resistance arteries from the rat gracilis muscle, K_ATP channel blockers reduce the membrane potential hyperpolarization and vasodilation seen in response to moderate and severe, but not mild, hypoxia (16). However, relaxation of the gracilis and other small skeletal muscle arteries and arterioles is largely endothelium dependent (15, 16, 31). K_ATP channels are activated during moderate and severe hypoxia as a consequence of prostacyclin release from endothelial cells (16). In contrast to small arteries, hypoxic vasodilation in large arteries supplying skeletal muscle is endothelium independent and is not reduced by K_ATP channel inhibition (Fig. 10 and Refs. 3 and 29). This suggests a difference in the mechanism of hypoxic vasodilation in small and large arteries supplying this vascular bed.

The endothelium-independent, K_ATP channel-independent hypoxic relaxation of larger arteries supplying skeletal muscle and other tissues must reflect a direct effect of hypoxia on the arterial smooth muscle cells (e.g., Fig. 10 and Refs. 3, 6, and 29). A number of mechanisms have been implicated in this direct effect of hypoxia (41). Relaxation occurs by Ca²⁺-dependent and Ca²⁺-independent mechanisms. Ca²⁺-independent mechanisms involve a change in the Ca²⁺ sensitivity of the contractile apparatus of the cell (e.g., 1, 17, 40). Hypoxia also inhibits calcium entry (35) and lowers [Ca²⁺]_i (e.g., Fig. 3 and Refs. 1 and 14). This may occur through voltage-dependent calcium channel inhibition (Fig. 3 and Refs. 13, 14, 18, and 45). We show a mean inhibition of peak calcium current at 0 mV of 16.7% at a PO₂ of 40 mmHg, similar to an inhibition of about 20% reported previously (Fig. 6 in Ref. 13). Interestingly, hypoxia had a relatively modest effect on the calcium current at the end of the voltage pulse (Fig. 3A). This is not evident in the work of Franco-Obregón et al. (13, 14), possibly because they used much shorter voltage pulses (20 ms cf. 160 ms), terminating the calcium current recording (by repolarization) before inactivation had developed. Although this might point against a physiological role for calcium current inhibition in hypoxic vasodilation in the steady state, it is important to point out that hypoxic inhibition of current is much more prominent at the membrane potentials normally experienced by vascular smooth muscle cells (i.e., –50 to –30 mV) than at 0 mV (13, 14). Ideally, to investigate this would require measurement of the steady-state calcium current at physiological membrane potentials. However, we estimate peak steady-state current at <1 pA (see Ref. 21), and this may therefore not be feasible. For this reason, we chose instead to measure [Ca²⁺]_i, a much more sensitive indicator than membrane current. Hypoxic solutions lowered [Ca²⁺]_i at physiological membrane potentials (Fig. 3C). Our results are therefore consistent with inhibition of calcium current and a fall in [Ca²⁺]_i contributing to hypoxic vasodilation in femoral arteries.

	GRANTS

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This work was funded by Wellcome Trust Project Grant 055506/Z/98.

	ACKNOWLEDGMENTS

 
Current affiliation for M. R. Turner: School of Medicine, University of Wales Swansea, Singleton Park, Swansea, SA2 8PP, UK.

	FOOTNOTES

 

Address for reprint requests and other correspondence: J. M. Quayle, Dept. of Human Anatomy and Cell Biology, School of Biomedical Sciences, Liverpool Univ., The Sherrington Bldgs., Ashton St., Liverpool L69 3GE, UK (e-mail: Jquayle{at}liv.ac.uk)

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.

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