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Physiology and Pharmacology  |   May 2014
Role of Ion Channels and Subcellular Ca2+ Signaling in Arachidonic Acid–Induced Dilation of Pressurized Retinal Arterioles
Author Notes
  • Centre for Experimental Medicine, School of Medicine, Dentistry and Biomedical Sciences, Queen's University of Belfast, Northern Ireland, United Kingdom 
  • Correspondence: Tim M. Curtis, Centre for Experimental Medicine, Queen's University of Belfast, Institute of Clinical Sciences Block A, Grosvenor Road, Royal Victoria Hospital, Belfast, BT12 6BA; t.curtis@qub.ac.uk
  • Footnotes
     Current affiliation: *Department of Neuroscience, University of Minnesota, Minneapolis, Minnesota, United States.
  • Footnotes
     Faculty of Pharmaceutical Sciences, Naresuan University, Phitsanulok, Thailand.
Investigative Ophthalmology & Visual Science May 2014, Vol.55, 2893-2902. doi:10.1167/iovs.13-13511
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      Joanna Kur, Mary K. McGahon, Jose A. Fernández, C. Norman Scholfield, J. Graham McGeown, Tim M. Curtis; Role of Ion Channels and Subcellular Ca2+ Signaling in Arachidonic Acid–Induced Dilation of Pressurized Retinal Arterioles. Invest. Ophthalmol. Vis. Sci. 2014;55(5):2893-2902. doi: 10.1167/iovs.13-13511.

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      © ARVO (1962-2015); The Authors (2016-present)

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Abstract

Purpose.: To investigate the mechanisms responsible for the dilatation of rat retinal arterioles in response to arachidonic acid (AA).

Methods.: Changes in the diameter of isolated, pressurized rat retinal arterioles were measured in the presence of AA alone and following pre-incubation with pharmacologic agents inhibiting Ca2+ sparks and oscillations and K+ channels. Subcellular Ca2+ signals were recorded in arteriolar myocytes using Fluo-4–based confocal imaging. The effects of AA on membrane currents of retinal arteriolar myocytes were studied using whole-cell perforated patch clamp recording.

Results.: Arachidonic acid dilated pressurized retinal arterioles under conditions of myogenic tone. Eicosatetraynoic acid (ETYA) exerted a similar effect, but unlike AA, its effects were rapidly reversible. Arachidonic acid–induced dilation was associated with an inhibition of subcellular Ca2+ signals. Interventions known to block Ca2+ sparks and oscillations in retinal arterioles caused dilatation and inhibited AA-induced vasodilator responses. Arachidonic acid accelerated the rate of inactivation of the A-type Kv current and the voltage dependence of inactivation was shifted to more negative membrane potentials. It also enhanced voltage-activated and spontaneous large-conductance calcium-activated K+ (BK) currents, but only at positive membrane potentials. Pharmacologic inhibition of A-type Kv and BK currents failed to block AA-induced vasodilator responses. Arachidonic acid suppressed L-type Ca2+ currents.

Conclusions.: These results suggest that AA induces retinal arteriolar vasodilation by inhibiting subcellular Ca2+-signaling activity in retinal arteriolar myocytes, most likely through a mechanism involving the inhibition of L-type Ca2+-channel activity. Arachidonic acid actions on K+ currents are inconsistent with a model in which K+ channels contribute to the vasodilator effects of AA.

Introduction
Arachidonic acid (AA) is an omega-6 long-chain polyunsaturated fatty acid. It is a major constituent of neuronal tissues of the retina and brain and represents approximately 10% of total fatty acids in intact vessels of the bovine retina and confluent endothelial cell/pericyte monolayers. 1  
In resting cells, AA is stored within the cell membrane, esterified to glycerol phospholipids. The highest concentrations of AA in the retina are found in phosphatidylcholine and phosphatidylethanolamine. 2 Arachidonic acid can be liberated via receptor- and Ca2+-dependent and/or Ca2+-independent pathways. 35 Arachidonic acid can be rapidly freed from the sn-2 position of membrane phospholipids by the enzyme phospholipase A2. 6 It can also be generated indirectly by other lipases from lipid products containing arachidonate, such as diacylglycerol, anandamide, or 2-arachidonylglycerol. 6 In many vascular beds, AA is known to be released from the endothelium to affect the overlying smooth muscle cells. 6,7 In the brain and retina, however, it can also be released from neuronal and glial cells that surround the blood vessels. 810  
Arachidonic acid and its metabolites are believed to be involved in the regulation of blood flow under basal conditions as well as during periods of increased metabolic demand. Vasodilator reactions induced by the electrical stimulation of cremaster muscle 11 or cortical perivascular nerve fibers, 12 for example, have been shown to be dependent upon AA signaling. In isolated perfused mesenteric arteries, the vascular response to AA is complex, with time-dependent constrictions and dilations being observed. 13 In addition, when injected intravenously, AA has been shown to lower systemic arterial blood pressure by decreasing both cardiac output and systemic vascular resistance. 14  
There is large body of evidence demonstrating that the effects of AA are mediated by modulation of ion channel function and Ca2+-signaling mechanisms. 15 In many vascular beds, AA-induced vasomotor effects have been linked to the modulation of K+-channel activity, including KCa, KATP, and Kv channels. 13,16 Arachidonic acid has also been implicated in the regulation of intracellular Ca2+ dynamics in various tissues, including cardiac myocytes and vascular smooth muscle cells, 1720 and has been reported to inhibit several different classes of voltage-dependent Ca2+ channels. 21  
Presently, the role of AA in the control of vascular tone in the retina is not well understood. Here, we report that AA induces dilation of isolated, myogenically active rat retinal arterioles. Experiments were also conducted to examine the mechanisms responsible for AA-induced vasodilation by investigating the effects of AA on cell membrane ion channels and subcellular Ca2+ signaling in retinal arteriolar myocytes. 
Methods
Animal use conformed to the standards in the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research and the UK Animals (Scientific Procedures) Act of 1986. Male Sprague-Dawley rats (8- to 12-weeks old; Harlan, Bicester, UK) were killed by CO2 asphyxiation (electrophysiology data) or injection with a lethal dose of sodium pentobarbital (300 mg/kg of body weight, given intraperitoneally; all other data sets). 
Arteriole Cannulation and Pressurization
First-order retinal arterioles were isolated from dissected retinas and cannulated as described previously. 22 Arterioles were inflated and maintained at 70 mm Hg for at least 15 minutes in normal Hanks' superfusate to allow for development of a stable level of myogenic tone. Intraluminal pressure was regulated by changing the height of a fluid reservoir connected to the inflow cannula and monitored using a pressure transducer. Preparations were leak-tested by observing an air bubble introduced into the tubing connecting the cannula to the transducer. If, at fixed high pressure, the bubble was moving this suggested that the preparation was leaky and the experiment was terminated. 
Arteriolar Diameter Measurements
Changes in arteriolar diameter were recorded using videomicroscopy and analyzed as described previously. 22 Briefly, a section of the arteriole, at least 40-μm away from the tip of the cannula was viewed under a ×40, NA 0.6 objective and imaged using a MCN-B013-U camera (Mightex, Pleasanton, CA, USA). The average internal diameter for a specified region of interest was measured using ImageJ software (http://imagej.nih.gov/ij/; provided in the public domain by the National Institutes of Health, Bethesda, MD, USA). The output was calibrated in micrometers using a graticule image. In the present study, it was not possible to use either the maximum diameter of the vessels at 70 mm Hg in Ca2+-free solution or the initial diameter following the pressure step to 70 mm Hg as a measure of the maximum passive diameter of the vessels for data normalization. Specifically, in some arterioles the dilation in Ca2+-free solution did not reach the initial diameter following the pressure step to 70 mm Hg (strongly suggesting that they had not reached their maximum passive diameter), whereas in other vessels the dilation went beyond that initial value (preventing us from using the diameter following the initial pressure step for normalization). Data has therefore been expressed as a percentage change in diameter from steady-state baseline values in the text and in absolute units in relevant tables. 
Subcellular Ca2+ Imaging
Smooth muscle Ca2+ signals in pressurized retinal arterioles pre-incubated with Fluo-4AM (10 μM for 2 hours; Molecular Probes, Inc., Eugene, OR, USA) were imaged using a confocal scanning laser microscope (MR-A1; Bio-Rad, Richmond, CA, USA). 22 The Fluo-4 was excited by an argon laser beam (488 nm) and emitted light was filtered through a 530- to 560-nm bandpass filter and detected using a photomultiplier tube (PMT). Calcium sparks and oscillations were analyzed with custom-made software and their spatiotemporal characteristics measured as described previously. 22 Changes in arteriolar diameter could not be simultaneously imaged with the Fluo-4–emitted fluorescence. Therefore, we collected brightfield and confocal recordings sequentially. Two different regions of the same vessel were imaged under control conditions and 5 minutes after AA treatment. 
Electrophysiological Recordings
Whole-cell membrane currents were recorded from individual smooth muscle cells still embedded within their parental arterioles using the perforated-patch clamp technique. 23 Supplementary Table S1 provides details of protocols used to isolate individual current components. Pipette resistances were 1 to 2 MΩ. Membrane currents were recorded using an Axopatch-1D (Axon Instruments, Union City, CA, USA) amplifier, low-pass filtered at 0.5 kHz and sampled at 2 kHz by National Instruments PC1200 interface (National Instruments, Newbury, UK) using WinWCP (v 3.3.3; University of Strathclyde, Glasgow, UK) software. The same software was also used for data analysis. Leak currents were subtracted off-line from the active currents using the standard leak subtraction protocol embedded within the WinWCP software (University of Strathclyde). Series resistance (15–30 MΩ) was routinely compensated by greater than 70%. Currents were normalized to cell capacitance and expressed as current densities (pA/pF). Cell membrane capacitance was determined from the time constant of a capacitance transient elicited by a hyperpolarizing step from −60 mV to −80 mV with a sampling frequency of 20 kHz. 
The voltage dependence of Kv channel activation was calculated by converting peak currents to a conductance (G) using the following equation: G = I/(V mE k), where: I is the current amplitude, V m is the command potential and E k is the equilibrium potential for K+. Based on the Nernst equation, E K was estimated to be −80 mV under recording conditions ([K+]o = 6 mM and [K+]i = 138 mM). Values were then normalized to the maximum conductance (G/G max). The voltage dependence of Kv channel inactivation was investigated by holding the retinal arteriolar myocytes at different membrane potentials during a 3-second conditioning prepulse and then applying a common test pulse (+60 mV; 1 second). The peak current following each test pulse was expressed relative to the maximum current recorded following a conditioning prepulse at −100 mV (I/I max). The resulting G/G max and I/I max values were fitted with a Boltzmann equation. 24  
Drugs and Solutions
The composition of the solutions used was as follows: (1) Hanks' solution: 140 mM NaCl; 6 mM KCl; 5 mM D-glucose; 2 mM CaCl2; 1.3 mM MgCl2; 10 mM HEPES (Sigma, Poole, UK); pH set to 7.4 with NaOH; (2) Divalent-free solution: 140 mM NaCl; 6 mM KCl; 5 mM D-glucose; 10 mM HEPES; 0.5 mM EGTA; (3) K+-based pipette solution: 138 mM KCl; 1 mM MgCl2; 0.5 mM EGTA; 0.2 mM CaCl2; 10 mM HEPES; pH set to 7.2 with KOH; (4) Cs+-based pipette solution: 138 mM CsCl; 1 mM MgCl2; 0.2 mM CaCl2; 10 mM HEPES; 0.5 mM EGTA; 2 mM ATP-2Na; 0.1 mM GTP-2Na; pH set to 7.2 with CsOH; and (5) Nominal Ca2+-free solution was of Hanks' composition, only Ca2+ was omitted. Amphotericin B (600 mg/mL) was dissolved in the pipette solutions as the pore-forming agent. 
Unless otherwise stated, stock solutions of drugs were initially prepared in dimethyl sulfoxide (DMSO) and then diluted to the final concentration. The final bath concentration of DMSO was less than or equal to 0.01%. 4-aminopyridine (4-AP), amphotericin B, AA, DIDS, EGTA, eicosatetraynoic acid (ETYA), and penitrem A were purchased from Sigma. Cyclopiazonic acid and nimodipine were from Alexis Biochemicals (Exeter, UK). Ryanodine was purchased from Ascent Scientific (Bristol, UK) and dissolved in absolute ethanol (bath vehicle concentration of 0.1%). All drugs were introduced abluminally via superfusion at 2 to 3 mL/min. Bath solutions were maintained at 37°C by passing the solution through a heat exchanger. 25  
In vehicle control experiments, application of either DMSO or ethanol, at the maximal concentrations used in these studies, had no effect on arteriolar diameter (0.0 ± 1.1% for DMSO at 0.01% vol/vol, n = 5 and 0.6 ± 0.3% for ethanol at 0.1% vol/vol, n = 6; P = 0.94 and P = 0.06, respectively, paired t-test; Table 1). A difference in the properties of Ca2+ sparks was observed in vehicle control experiments with DMSO (0.01% vol/vol). Specifically, it caused a small yet significant increase in the frequency of these events (from a control value of 0.040 ± 0.020 cell−1s−1 to 0.058 ± 0.011 cell−1s−1 after DMSO exposure, n = 78, P < 0.01, Mann-Whitney U test). Note, however, that the action of DMSO on Ca2+ spark frequency was opposite to the effect of AA (see Results). 
Table 1
 
Mean (±SEM) Changes in Internal Diameter of Pressurized Retinal Arterioles in Response to AA, ETYA, and Vehicle Controls
Table 1
 
Mean (±SEM) Changes in Internal Diameter of Pressurized Retinal Arterioles in Response to AA, ETYA, and Vehicle Controls
Baseline Diameter, μm Maximum Diameter in Presence of Drug, μm Diameter Following 5 Min of Wash Out
28.3 ± 1.5 AA (10 μM; n = 11) 30.8 ± 1.5† 30.3 ± 1.6†
25.5 ± 2.5 ETYA (10 μM; n = 8) 28.5 ± 2.4* 26.0 ± 2.2 NS
27.7 ± 3.0 DMSO (0.01% vol/vol; n = 5) 27.7 ± 3.0 NS NA
27.4 ± 1.4 Ethanol (0.1% vol/vol; n = 6) 27.6 ± 1.5 NS NA
Statistical Analysis
Data are presented as the mean ± SEM. The statistical significance of differences in arteriole diameter under control and experimental conditions were determined using a paired t-test or repeated measures ANOVA followed by Bonferroni multiple comparison tests as appropriate. Paired data for the percentage of smooth muscle cells displaying Ca2+ sparks and/or Ca2+ oscillations before and during AA treatment were tested using the Wilcoxon rank sum test. Spark or oscillation population data, consisting of all spatiotemporal measurements for individual Ca2+ events, were compared using the Mann-Whitney U test. Summary frequency data were generated by averaging the frequency seen in each cell, with inactive (quiescent) myocytes counted as zero. Differences in whole-membrane current densities and time constants of inactivation were tested with two-way ANOVA or paired t-test. In all comparisons of mean data, the 95% level was accepted as statistically significant (in all graphical representations of the data, statistical significance is indicated as follows: NS, P > 0.05; *P < 0.05; **P < 0.01; ***P < 0.001). 
Results
AA-Induced Vasodilation and Inhibition of Ca2+ Sparks and Oscillations
Because physiological concentrations of AA are frequently in the low to mid micromolar range (1–150 μM), 26 we chose to test AA at a concentration of 10 μM. Application of 10 μM AA (5 minutes) to isolated, pressurized rat retinal arterioles resulted in an increase in vessel diameter that averaged 9.2 ± 1.3% of the steady-state diameter under conditions of myogenic tone (P < 0.001, repeated measures ANOVA; Figs. 1Ai–Ci and Table 1). This effect was irreversible within the time limit of the wash out period (5 minutes; 7.2 ± 1.9%, P < 0.001 versus baseline). The nonmetabolizable analogue of AA, ETYA (10 μM), also elicited dilation of myogenically active, pressurized rat retinal arterioles (diameter increase of 13.7 ± 4.2%; P < 0.01; Figs. 1Aii–Cii and Table 1), but unlike AA, its effects were rapidly reversible (5 minutes; 3.8 ± 4.9% of the initial steady-state diameter prior to ETYA addition; P > 0.05 versus baseline). 
Figure 1
 
Dilation of pressurized rat retinal arterioles in response to AA and its nonmetabolizable analogue, ETYA. (A) Time-course records showing changes in internal diameter of pressurized rat retinal arterioles during application of 10 μM AA (Ai) and 10 μM ETYA (Aii). Bars represent drug application intervals. (B) Photomicrographs of the same arterioles at the time points indicated by arrows in (Ai) and (Aii). (C) Mean (±SEM) percentage changes in the diameter of arterioles superfused for 5 minutes with either 10 μM AA (Ci) or ETYA (Cii) and after 5 minutes of wash out. (Data was collected from a minimum of eight vessels from six rats; ns = P > 0.05, **P < 0.01, ***P < 0.001 relative to baseline; repeated measures ANOVA.)
Figure 1
 
Dilation of pressurized rat retinal arterioles in response to AA and its nonmetabolizable analogue, ETYA. (A) Time-course records showing changes in internal diameter of pressurized rat retinal arterioles during application of 10 μM AA (Ai) and 10 μM ETYA (Aii). Bars represent drug application intervals. (B) Photomicrographs of the same arterioles at the time points indicated by arrows in (Ai) and (Aii). (C) Mean (±SEM) percentage changes in the diameter of arterioles superfused for 5 minutes with either 10 μM AA (Ci) or ETYA (Cii) and after 5 minutes of wash out. (Data was collected from a minimum of eight vessels from six rats; ns = P > 0.05, **P < 0.01, ***P < 0.001 relative to baseline; repeated measures ANOVA.)
Calcium sparks are brief, highly localized, subcellular Ca2+ release events. In retinal arteriolar myocytes, Ca2+ sparks can summate to generate more prolonged global Ca2+ oscillations, which in turn can lead to myocyte contraction and the generation of myogenic tone. 22,27 We therefore tested whether AA modulates subcellular Ca2+-signaling events in these vessels. Brightfield (vessel diameter) and confocal (Ca2+-signaling) images were recorded sequentially from myogenically active arterioles maintained at 70 mm Hg and loaded with the fluorescent Ca2+ indicator dye, Fluo-4. The average AA-induced vasodilation in these vessels was 6.6 ± 0.9% (P < 0.001, paired t-test). This did not differ significantly from the vasodilatory response observed in unloaded vessels (P = 0.17; Mann-Whitney U test). 
Under control conditions, myogenically active arterioles exhibited spontaneous subcellular Ca2+-signaling activity, as previously described. 22 Arachidonic acid–induced vasodilatory effects were associated with a pronounced inhibition of subcellular Ca2+ signals (Fig. 2). The percentage of smooth muscle cells displaying Ca2+ sparks decreased from 52.3 ± 5.1% under control conditions to 23.8 ± 8.9% following AA exposure (P < 0.05, Wilcoxon rank sum test). With the exception of frequency, which decreased from 0.066 ± 0.013 cell−1s−1 under control conditions to 0.019 ± 0.005 cell−1s−1 after AA addition (P < 0.001, Mann-Whitney U test), other spatiotemporal features of Ca2+ sparks were unaffected (Table 2). 
Figure 2
 
Arachidonic acid inhibits subcellular Ca2+ signals in pressurized rat retinal arterioles. Low (A) and high (B) temporal resolution confocal line scan images and time series plots showing changes in normalized fluorescence (ΔF/F0) for representative adjacent arteriolar myocytes (labeled from a–h). Paired images in (A, B) were recorded from two different regions of the same vessel under control conditions and 5 minutes after application of 10 μM AA. The regions marked with the dashed boxes in (A) correspond to the panels shown below in (B). Two types of subcellular Ca2+-signaling events can be observed: prolonged Ca2+ oscillations, which are identifiable in both low- and high-temporal resolution images and brief Ca2+ sparks (indicated by asterisks in [B]), which can only be resolved in high-temporal resolution images. Note a decrease in the frequency of both Ca2+ sparks and oscillations following AA treatment.
Figure 2
 
Arachidonic acid inhibits subcellular Ca2+ signals in pressurized rat retinal arterioles. Low (A) and high (B) temporal resolution confocal line scan images and time series plots showing changes in normalized fluorescence (ΔF/F0) for representative adjacent arteriolar myocytes (labeled from a–h). Paired images in (A, B) were recorded from two different regions of the same vessel under control conditions and 5 minutes after application of 10 μM AA. The regions marked with the dashed boxes in (A) correspond to the panels shown below in (B). Two types of subcellular Ca2+-signaling events can be observed: prolonged Ca2+ oscillations, which are identifiable in both low- and high-temporal resolution images and brief Ca2+ sparks (indicated by asterisks in [B]), which can only be resolved in high-temporal resolution images. Note a decrease in the frequency of both Ca2+ sparks and oscillations following AA treatment.
Table 2
 
The Percentage of Myocytes Exhibiting Ca2+ Sparks and Oscillations and the Spatiotemporal Features of These Events in Pressurized Rat Retinal Arterioles Under Control Conditions and Following AA Treatment
Table 2
 
The Percentage of Myocytes Exhibiting Ca2+ Sparks and Oscillations and the Spatiotemporal Features of These Events in Pressurized Rat Retinal Arterioles Under Control Conditions and Following AA Treatment
Ca2+ Sparks Ca2+ Oscillations
Control AA Control AA
Active cells, % 52.3 ± 5.1 (105) 23.8 ± 8.9* (85) 70.0 ± 2.4 (105) 43.9 ± 9.8* (85)
Amplitude, F/F0 1.53 ± 0.06 (414) 1.55 ± 0.11 (99) 2.17 ± 0.07 (493) 1.70 ± 0.07† (156)
FDHM, ms 23.99 ± 0.88 (190) 23.07 ± 1.92 (41) 866.6 ± 0.03 (493) 770.6 ± 0.05‡ (493)
FWHM, μm 0.46 ± 0.01 (414) 0.41 ± 0.01 (99) NA NA
Frequency (cell−1 s−1) 0.066 ± 0.013 (105) 0.019 ± 0.005§ (85) 0.078 ± 0.008 (105) 0.031 ± 0.005§ (85)
Arachidonic acid also altered the percentage of smooth muscle cells displaying Ca2+ oscillations, which decreased from 70.0 ± 2.4% under control conditions to 43.9 ± 9.8% after application of AA (P < 0.05, Wilcoxon rank sum test). The average frequency of Ca2+ oscillations was also decreased, from a control value of 0.078 ± 0.008 cell−1s−1 to 0.031 ± 0.005 cell−1s−1 during AA exposure (Table 2; P < 0.001 Mann-Whitney U test). Arachidonic acid also reduced the amplitude and duration of these Ca2+ events. When averaged, the peak amplitude (F/F0) was 2.17 ± 0.07 in control and 1.70 ± 0.07 following AA application (P < 0.05, Mann-Whitney U test), while the full duration at half maximum (FDHM) decreased from 866.6 ± 0.03 ms under control conditions to 770.6 ± 0.05 ms after AA addition (P < 0.01, Mann-Whitney U test). 
Since AA blocked subcellular Ca2+ signals in pressurized rat retinal arterioles, we wanted to explore in more detail whether these effects were causally related to the dilatory response. As shown in Table 3, pharmacologic interventions that are known to block Ca2+ sparks and Ca2+ oscillations in retinal arterioles 27 inhibited the AA-induced vasodilatory response. In general, these results could not be attributed to the vessels reaching their maximal vasodilatory capacity prior to AA exposure since in the majority of vessels tested (>70%) a greater dilation could be evoked by applying nominal Ca2+-free solution than in the presence of the various Ca2+-signaling inhibitors (Supplementary Fig. S1). 
Table 3
 
Mean (±SEM) Changes in Internal Diameter of Pressurized Retinal Arterioles in Response to AA Following Pretreatment With Pharmacologic Inhibitors
Table 3
 
Mean (±SEM) Changes in Internal Diameter of Pressurized Retinal Arterioles in Response to AA Following Pretreatment With Pharmacologic Inhibitors
Baseline Diameter, μm Maximum Diameter in Presence of Inhibitor, μm Maximum Diameter in Presence of Inhibitor and AA (10 μM), μm
28.5 ± 1.8 Ryanodine (100 μM; n = 9) 31.5 ± 1.1* Ryanodine + AA 32.4 ± 1.1†
29.3 ± 1.0 CPA (20 μM; n = 5) 31.6 ± 1.0‡ CPA + AA 31.6 ± 0.9‡
30.3 ± 1.5 Nimodipine (1 μM; n = 6) 32.5 ± 1.4† Nimodipine + AA 33.3 ± 1.6‡
28.4 ± 3.8 4-AP (10 mM), Penitrem A (100 nM; n = 5) 26.6 ± 3.7† 4-AP + Penitrem A + AA 28.5 ± 3.7
AA-Modulation of Ion Channels in Arteriolar Smooth Muscle Cells
Studies in other vascular beds have demonstrated that AA-induced vasodilation depends on modulation of K+ channel function. 16,28,29 We therefore decided to examine the effects of AA on the major K+ currents (A-type Kv and large-conductance calcium-activated K+ [BK] currents) previously characterized in retinal arteriolar smooth muscle cells. 23,30,31  
Following AA exposure (10 μM), the sustained component of the A-type Kv current was inhibited at potentials between +20 mV and +100 mV, whereas the maximal (peak) current was unchanged (Figs. 3A, 3B). Inspection of the kinetics of the A-type Kv current profile revealed a dramatic increase in the inactivation rate following treatment with AA. In the presence of AA, the mean time constant of fast inactivation was reduced from 37 ± 3 ms to 16 ± 7 ms (P < 0.05, paired t-test), while the mean time constant of slow inactivation decreased from 370 ± 64 ms to 77 ± 19 ms (P < 0.001, paired t-test; Fig. 3C). To establish the range of membrane potentials over which KV channels are capable of mediating a sustained outward “window current,” we examined the voltage-dependence of activation and inactivation before and after AA exposure (Fig. 3D). We found that the voltage-dependence of activation was not altered in the presence of AA (P = 0.52, repeated measures two-way ANOVA). In contrast, the voltage dependence of inactivation was shifted negatively by AA. Under control conditions the A-type Kv current was fully inactivated at −22.7 ± 2.4 mV, whereas following AA treatment full inactivation was observed at −35.7 ± 1.9 mV (P < 0.01, paired t-test). 
Figure 3
 
Arachidonic acid–induced modulation of A-type KV current in retinal arteriolar myocytes. (A) A-type Kv currents evoked in response to depolarizing voltage steps (protocol shown in inset) under control conditions (Ai) and in the presence (Aii) of 10 μM AA (7 minutes). (B) Mean (±SEM) peak (Bi) and steady-state (Bii) current densities recorded under control conditions (closed circles) and in the presence of 10 μM AA (open circles; n = 6). (Ci) A-type Kv currents evoked in response to a single voltage step from −80 mV to +40 mV in the absence and presence of AA (extracted from [A]). Currents were fitted with double exponential curves (solid lines superimposed on current traces) and inactivation time constants calculated. (Cii) Mean (±SEM) fast and slow time constants before and following AA exposure (n = 8). Note a decrease in the time constants in the presence of AA indicating acceleration of the rate of Kv current inactivation. (Di) Representative current traces resulting from a double-pulse protocol (shown in inset) applied to obtain the steady-state, voltage-dependent inactivation of the A-type Kv current in the presence and absence of AA. (Dii) The voltage dependence of activation (open circles, n = 6) and inactivation (closed circles, n = 7) were calculated as described in the Methods and plotted against membrane potential. The lines are the fit of Boltzmann relationships to the data for control (solid line) and AA (dashed line) conditions. Under control conditions the activation and inactivation curves overlap substantially revealing a “window current” (area under the two intersecting curves). Note that AA exposure resulted in a negative shift in the inactivation curve causing a reduction in the window current. Symbols *, **, *** denote P < 0.05, P < 0.01, and P < 0.001 significance level, respectively.
Figure 3
 
Arachidonic acid–induced modulation of A-type KV current in retinal arteriolar myocytes. (A) A-type Kv currents evoked in response to depolarizing voltage steps (protocol shown in inset) under control conditions (Ai) and in the presence (Aii) of 10 μM AA (7 minutes). (B) Mean (±SEM) peak (Bi) and steady-state (Bii) current densities recorded under control conditions (closed circles) and in the presence of 10 μM AA (open circles; n = 6). (Ci) A-type Kv currents evoked in response to a single voltage step from −80 mV to +40 mV in the absence and presence of AA (extracted from [A]). Currents were fitted with double exponential curves (solid lines superimposed on current traces) and inactivation time constants calculated. (Cii) Mean (±SEM) fast and slow time constants before and following AA exposure (n = 8). Note a decrease in the time constants in the presence of AA indicating acceleration of the rate of Kv current inactivation. (Di) Representative current traces resulting from a double-pulse protocol (shown in inset) applied to obtain the steady-state, voltage-dependent inactivation of the A-type Kv current in the presence and absence of AA. (Dii) The voltage dependence of activation (open circles, n = 6) and inactivation (closed circles, n = 7) were calculated as described in the Methods and plotted against membrane potential. The lines are the fit of Boltzmann relationships to the data for control (solid line) and AA (dashed line) conditions. Under control conditions the activation and inactivation curves overlap substantially revealing a “window current” (area under the two intersecting curves). Note that AA exposure resulted in a negative shift in the inactivation curve causing a reduction in the window current. Symbols *, **, *** denote P < 0.05, P < 0.01, and P < 0.001 significance level, respectively.
Voltage-activated BK currents were investigated in retinal arteriolar myocytes by stepping the membrane potential from −80 mV to +70 mV in 10 mV increments. Using this protocol, changes in BK currents following the addition of 10 μM AA are illustrated in Figure 4Ai. It is apparent that AA treatment substantially enhanced these currents, but only at membrane potentials positive to +30 mV (Fig. 4Aii). We also examined spontaneous BK currents at steady-state holding potentials between −60 mV and +60 mV. 31 Similar to our findings for voltage-activated BK currents, spontaneous BK currents were significantly increased by 10 μM AA, but only at holding potentials positive to +20 mV (Figs. 4Bi, 4Bii). 
Figure 4
 
Arachidonic acid–induced activation of the BK current in retinal arteriolar myocytes. (Ai) BK currents evoked in response to depolarizing voltage steps (protocol shown in inset) under control conditions (upper panel) and following 5-minute application of 10 μM AA (lower panel). (Aii) Mean (±SEM) BK current densities (averaged across each depolarizing voltage step) under control conditions (closed circles) and in the presence of 10 μM AA (open circles). Note an enhancement of BK current at membrane potentials positive to +30 mV (n = 5; *P < 0.05, ***P < 0.001, two-way ANOVA). (Bi) Spontaneous BK currents recorded at various holding potentials under control conditions (left panel) and following 5 minute application of 10 μM AA (right panel). (Bii) Mean (±SEM) integrated spontaneous BK current densities under control conditions (closed circles) and in the presence of 10 μM AA (open circles). Enhancement of spontaneous BK current activity was observed at holding potentials positive to +20 mV (n = 4; ***P < 0.001, two-way ANOVA).
Figure 4
 
Arachidonic acid–induced activation of the BK current in retinal arteriolar myocytes. (Ai) BK currents evoked in response to depolarizing voltage steps (protocol shown in inset) under control conditions (upper panel) and following 5-minute application of 10 μM AA (lower panel). (Aii) Mean (±SEM) BK current densities (averaged across each depolarizing voltage step) under control conditions (closed circles) and in the presence of 10 μM AA (open circles). Note an enhancement of BK current at membrane potentials positive to +30 mV (n = 5; *P < 0.05, ***P < 0.001, two-way ANOVA). (Bi) Spontaneous BK currents recorded at various holding potentials under control conditions (left panel) and following 5 minute application of 10 μM AA (right panel). (Bii) Mean (±SEM) integrated spontaneous BK current densities under control conditions (closed circles) and in the presence of 10 μM AA (open circles). Enhancement of spontaneous BK current activity was observed at holding potentials positive to +20 mV (n = 4; ***P < 0.001, two-way ANOVA).
To further examine the possible contribution of A-type Kv and BK currents in the AA-induced dilation of pressurized retinal arterioles, a set of experiments was carried out in the presence of the A-type Kv and BK channel blockers, 4-AP (10 mM) and Penitrem A (100 nM), respectively. Addition of the K+-channel blockers to myogenically active retinal arterioles resulted in significant vasoconstriction (6.4 ± 1.3% decrease in vessel diameter; P < 0.01, repeated measures ANOVA; Table 3). Subsequent addition of AA in the continued presence of the K+-channel blockers evoked a robust vasodilatory response (8.0 ± 1.8% increase in vessel diameter; P < 0.01, repeated measures ANOVA; Table 3) that did not differ significantly from that previously observed with AA alone (P = 0.82; Mann-Whitney U test). 
Data from our own laboratory have shown that Ca2+ influx through L-type Ca2+ channels contributes to myogenic tone development and the generation of subcellular Ca2+-signaling activity in retinal arterioles. 22,27 It was therefore of interest to examine the effects of AA on these channels. As shown in Figure 5, L-type Ca2+ currents were AA sensitive. On average, the peak inward current during voltage steps from −80 to 0 mV decreased from −11.7 ± 3.5 pA/pF under control conditions to −2.4 ± 2.7 pA/pF after AA treatment (P < 0.05, paired t-test). 
Figure 5
 
Arachidonic acid–induced inhibition of the L-type Ca2+ current in retinal arteriolar myocytes. (A) L-type Ca2+ currents under control conditions and following approximately 1-minute exposure to 10 μM AA evoked by depolarizing steps from −80 mV to 0 mV (inset). (B) Mean (±SEM) peak L-type Ca2+ current densities for the experimental conditions shown in (A) (n = 6; *P < 0.05 paired t-test).
Figure 5
 
Arachidonic acid–induced inhibition of the L-type Ca2+ current in retinal arteriolar myocytes. (A) L-type Ca2+ currents under control conditions and following approximately 1-minute exposure to 10 μM AA evoked by depolarizing steps from −80 mV to 0 mV (inset). (B) Mean (±SEM) peak L-type Ca2+ current densities for the experimental conditions shown in (A) (n = 6; *P < 0.05 paired t-test).
Discussion
AA-Induced Modulation of Myogenic Tone and Subcellular Ca2+ Signaling
The results presented here provide the first direct evidence for a role of AA in modulating retinal vascular tone. Our findings are consistent with previous reports showing that AA is capable of mediating vasodilation in coronary, 32 mesenteric, 3,33 femoral, 34 intrapulmonary, 35 and placental artery. 36 They also demonstrate that AA is not only an important mediator of vasodilation in large arterial vessels, but also acts at the level of the microcirculation. 
Confocal Ca2+ imaging in pressurized retinal arterioles demonstrated that subcellular Ca2+ signals in retinal arteriolar myocytes are inhibited by AA. To the best of our knowledge, this is the first direct evidence demonstrating that AA is capable of modulating localized Ca2+ signals in any tissue. Our data also strongly supports the idea that inhibition of Ca2+-spark and Ca2+-oscillation activity is the primary mechanism underlying the effects of AA on retinal arteriolar tone, since interventions that blocked Ca2+ sparks and Ca2+ oscillations prevented the vasodilatory effects of AA (although we cannot fully rule out the possibility that a few of the vessels included in this dataset had already reached their maximal vasodilatory capacity prior to AA addition). While this study is the first to describe the effects of AA on subcellular Ca2+ signaling, previous studies have reported changes in Ca2+ sparks and global Ca2+-signaling activity in response to other polyunsaturated fatty acids (PUFAs). For instance, in cardiac myocytes, the omega-3 PUFA, eicosapentaenoic acid (EPA), has been found to reduce both the width and duration of Ca2+ sparks 37 and the amplitude and rate of propagation of spontaneous Ca2+ waves. 38 Taken together, these results and those of the present study suggest that the suppression of subcellular Ca2+-signaling activity may represent a common mechanism underlying the physiological effects of PUFAs in both cardiac myocytes and vascular smooth muscle cells. 
AA-Induced Modulation of K+ Currents
From a physiological perspective, it is well established that the activation of K+ channels counteracts the development of myogenic tone by providing a hyperpolarizing influence, which reduces Ca2+ influx through voltage-dependent Ca2+ channels, and thereby limits the degree of vasoconstriction. 39 K+-channel activation has been implicated in the dilator effects of AA in a number of vascular beds. 13,16 However, as described below, our electrophysiological data are inconsistent with the idea that K+ channels contribute to the vasodilator effects of AA in retinal arterioles, and notably, we observed no significant difference in the degree of AA-induced vasodilation in the absence and presence of K+ channel inhibitors. 
We found that AA caused acceleration in the time course of inactivation and a negative shift in the voltage-dependence of inactivation of the A-type KV current in retinal arteriolar myocytes, reducing the steady state KV window current. A reduction in KV window current would be expected to cause cell membrane potential depolarization, increased Ca2+ influx through voltage-dependent Ca2+ channels, and thus vessel constriction rather than dilation. Our observation of accelerated inactivation concurs with previous studies examining the effects AA on fast inactivating Kv currents in rat pituitary melanotrophs 40 and rat hippocampal and pyramidal neurons. 41,42 Studies using heterologous expression systems suggest that, at a molecular level, AA increases the rate of inactivation of KV channels by inducing conformational alterations in the selectivity filter. 43  
We found that AA increased the amplitude of voltage-activated and spontaneous BK currents in retinal arteriolar myocytes. These findings are consistent with previous observations in gastric myocytes of guinea pig and rabbit pulmonary artery smooth muscle cells. 44,45 Recently, it has been reported that AA may act to potentiate BK currents through an indirect mechanism involving metabolites of the cyclooxygenase (COX) pathway. 28 Although we observed similar effects of AA on BK currents as those reported in other types of smooth muscle, it is important to note that the resting membrane potential of retinal arteriolar myocytes is far more negative (−40 mV) 23 than the threshold for AA-induced activation of BK currents in these cells (+40 mV; Fig. 4). Thus, it would appear that this mechanism is not likely to be relevant at physiological potentials and this most likely explains why inhibition of these channels failed to attenuate the vasodilatory actions of AA on myogenically active retinal arterioles. 
AA Effects on L-Type Ca2+ Currents
The results of present study show that AA inhibits L-type channel currents. Similar findings have been reported for smooth muscle cells from the vas deferens of the guinea pig 46 and rabbit intestine, 47 and in cardiac myocytes from frog ventricle. 48 There is no consensus on the mechanism responsible. Khurana and Bennett 49 suggested that in ciliary ganglion cells leukotrienes synthesized from AA by lipoxygenase act as second messengers responsible for AA-induced block of voltage-dependent Ca2+ channels. Other studies, meanwhile, have reported that the inhibition of voltage-gated Ca2+ channels by AA may, at least in part, be due to superoxide radicals derived from AA oxidation. 46 Regardless of the mode of action at the ion channel level, our results are consistent with the idea that the effects of AA on subcellular Ca2+ signaling, and hence vascular tone in retinal arterioles are most likely due to the inhibition of voltage-dependent Ca2+ entry. Indeed in previous work, we have shown that blockade of L-type Ca2+ channels suppresses Ca2+ sparks and oscillations in retinal arterioles, and causes vasodilation through a pathway associated with a reduction in the sarcoplasmic reticulum (SR) Ca2+ content. 27 It is worth stressing, however, that at this stage other possible contributory mechanisms cannot be fully discounted. For example, possible effects of AA on sarcoplasmic/endoplasmic reticulum Ca2+-ATPase (SERCA) activity and store-operated Ca2+ entry could also contribute to the inhibition of subcellular Ca2+-signaling activity by modifying SR Ca2+ content. It is also possible that AA may act to block the function of ryanodine receptors on the SR, which are known to underlie Ca2+ spark and oscillation activity in retinal arteriolar myocytes. 27 Previous biophysical studies using lipid-bilayer systems, however, have suggested that AA is not a direct inhibitor of these channels. 20  
Conclusions
In conclusion, our experiments provide evidence that AA contributes to the modulation of the contractile, Ca2+ signaling and cell membrane ion-channel activity of retinal arteriolar myocytes. Our patch-clamp data and pressure myography experiments suggest that AA actions on individual K+ conductances cannot explain the vasodilatory effects of this lipid-signaling molecule. The most likely mechanism underlying the actions of AA on retinal arteriolar tone is through a pathway involving the inhibition of Ca2+ sparks and Ca2+ oscillations secondary to blockade of L-type Ca2+ channels. A major question that remains is whether these effects are exerted directly through AA or result from the generation of AA metabolites. Our studies using the nonmetabolizable analogue of AA, ETYA, suggest that AA may be capable of directly causing retinal arteriolar vasodilation, although differences in the reversibility of these compounds indicates that a contribution by AA metabolites cannot be fully excluded. Elucidating a possible role for AA metabolites in the vasodilator effects of AA on retinal arterioles provides a clear direction for future research in this area. 
Supplementary Materials
Acknowledgments
Supported by grants from Fight for Sight, UK, and BBSRC (BB/I026359/1). 
Disclosure: J. Kur, None; M.K. McGahon, None; J.A. Fernández, None; C.N. Scholfield, None; J.G. McGeown, None; T.M. Curtis, None 
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Footnotes
 JGM and TMC contributed equally to the work presented here and should therefore be regarded as equivalent authors.
Figure 1
 
Dilation of pressurized rat retinal arterioles in response to AA and its nonmetabolizable analogue, ETYA. (A) Time-course records showing changes in internal diameter of pressurized rat retinal arterioles during application of 10 μM AA (Ai) and 10 μM ETYA (Aii). Bars represent drug application intervals. (B) Photomicrographs of the same arterioles at the time points indicated by arrows in (Ai) and (Aii). (C) Mean (±SEM) percentage changes in the diameter of arterioles superfused for 5 minutes with either 10 μM AA (Ci) or ETYA (Cii) and after 5 minutes of wash out. (Data was collected from a minimum of eight vessels from six rats; ns = P > 0.05, **P < 0.01, ***P < 0.001 relative to baseline; repeated measures ANOVA.)
Figure 1
 
Dilation of pressurized rat retinal arterioles in response to AA and its nonmetabolizable analogue, ETYA. (A) Time-course records showing changes in internal diameter of pressurized rat retinal arterioles during application of 10 μM AA (Ai) and 10 μM ETYA (Aii). Bars represent drug application intervals. (B) Photomicrographs of the same arterioles at the time points indicated by arrows in (Ai) and (Aii). (C) Mean (±SEM) percentage changes in the diameter of arterioles superfused for 5 minutes with either 10 μM AA (Ci) or ETYA (Cii) and after 5 minutes of wash out. (Data was collected from a minimum of eight vessels from six rats; ns = P > 0.05, **P < 0.01, ***P < 0.001 relative to baseline; repeated measures ANOVA.)
Figure 2
 
Arachidonic acid inhibits subcellular Ca2+ signals in pressurized rat retinal arterioles. Low (A) and high (B) temporal resolution confocal line scan images and time series plots showing changes in normalized fluorescence (ΔF/F0) for representative adjacent arteriolar myocytes (labeled from a–h). Paired images in (A, B) were recorded from two different regions of the same vessel under control conditions and 5 minutes after application of 10 μM AA. The regions marked with the dashed boxes in (A) correspond to the panels shown below in (B). Two types of subcellular Ca2+-signaling events can be observed: prolonged Ca2+ oscillations, which are identifiable in both low- and high-temporal resolution images and brief Ca2+ sparks (indicated by asterisks in [B]), which can only be resolved in high-temporal resolution images. Note a decrease in the frequency of both Ca2+ sparks and oscillations following AA treatment.
Figure 2
 
Arachidonic acid inhibits subcellular Ca2+ signals in pressurized rat retinal arterioles. Low (A) and high (B) temporal resolution confocal line scan images and time series plots showing changes in normalized fluorescence (ΔF/F0) for representative adjacent arteriolar myocytes (labeled from a–h). Paired images in (A, B) were recorded from two different regions of the same vessel under control conditions and 5 minutes after application of 10 μM AA. The regions marked with the dashed boxes in (A) correspond to the panels shown below in (B). Two types of subcellular Ca2+-signaling events can be observed: prolonged Ca2+ oscillations, which are identifiable in both low- and high-temporal resolution images and brief Ca2+ sparks (indicated by asterisks in [B]), which can only be resolved in high-temporal resolution images. Note a decrease in the frequency of both Ca2+ sparks and oscillations following AA treatment.
Figure 3
 
Arachidonic acid–induced modulation of A-type KV current in retinal arteriolar myocytes. (A) A-type Kv currents evoked in response to depolarizing voltage steps (protocol shown in inset) under control conditions (Ai) and in the presence (Aii) of 10 μM AA (7 minutes). (B) Mean (±SEM) peak (Bi) and steady-state (Bii) current densities recorded under control conditions (closed circles) and in the presence of 10 μM AA (open circles; n = 6). (Ci) A-type Kv currents evoked in response to a single voltage step from −80 mV to +40 mV in the absence and presence of AA (extracted from [A]). Currents were fitted with double exponential curves (solid lines superimposed on current traces) and inactivation time constants calculated. (Cii) Mean (±SEM) fast and slow time constants before and following AA exposure (n = 8). Note a decrease in the time constants in the presence of AA indicating acceleration of the rate of Kv current inactivation. (Di) Representative current traces resulting from a double-pulse protocol (shown in inset) applied to obtain the steady-state, voltage-dependent inactivation of the A-type Kv current in the presence and absence of AA. (Dii) The voltage dependence of activation (open circles, n = 6) and inactivation (closed circles, n = 7) were calculated as described in the Methods and plotted against membrane potential. The lines are the fit of Boltzmann relationships to the data for control (solid line) and AA (dashed line) conditions. Under control conditions the activation and inactivation curves overlap substantially revealing a “window current” (area under the two intersecting curves). Note that AA exposure resulted in a negative shift in the inactivation curve causing a reduction in the window current. Symbols *, **, *** denote P < 0.05, P < 0.01, and P < 0.001 significance level, respectively.
Figure 3
 
Arachidonic acid–induced modulation of A-type KV current in retinal arteriolar myocytes. (A) A-type Kv currents evoked in response to depolarizing voltage steps (protocol shown in inset) under control conditions (Ai) and in the presence (Aii) of 10 μM AA (7 minutes). (B) Mean (±SEM) peak (Bi) and steady-state (Bii) current densities recorded under control conditions (closed circles) and in the presence of 10 μM AA (open circles; n = 6). (Ci) A-type Kv currents evoked in response to a single voltage step from −80 mV to +40 mV in the absence and presence of AA (extracted from [A]). Currents were fitted with double exponential curves (solid lines superimposed on current traces) and inactivation time constants calculated. (Cii) Mean (±SEM) fast and slow time constants before and following AA exposure (n = 8). Note a decrease in the time constants in the presence of AA indicating acceleration of the rate of Kv current inactivation. (Di) Representative current traces resulting from a double-pulse protocol (shown in inset) applied to obtain the steady-state, voltage-dependent inactivation of the A-type Kv current in the presence and absence of AA. (Dii) The voltage dependence of activation (open circles, n = 6) and inactivation (closed circles, n = 7) were calculated as described in the Methods and plotted against membrane potential. The lines are the fit of Boltzmann relationships to the data for control (solid line) and AA (dashed line) conditions. Under control conditions the activation and inactivation curves overlap substantially revealing a “window current” (area under the two intersecting curves). Note that AA exposure resulted in a negative shift in the inactivation curve causing a reduction in the window current. Symbols *, **, *** denote P < 0.05, P < 0.01, and P < 0.001 significance level, respectively.
Figure 4
 
Arachidonic acid–induced activation of the BK current in retinal arteriolar myocytes. (Ai) BK currents evoked in response to depolarizing voltage steps (protocol shown in inset) under control conditions (upper panel) and following 5-minute application of 10 μM AA (lower panel). (Aii) Mean (±SEM) BK current densities (averaged across each depolarizing voltage step) under control conditions (closed circles) and in the presence of 10 μM AA (open circles). Note an enhancement of BK current at membrane potentials positive to +30 mV (n = 5; *P < 0.05, ***P < 0.001, two-way ANOVA). (Bi) Spontaneous BK currents recorded at various holding potentials under control conditions (left panel) and following 5 minute application of 10 μM AA (right panel). (Bii) Mean (±SEM) integrated spontaneous BK current densities under control conditions (closed circles) and in the presence of 10 μM AA (open circles). Enhancement of spontaneous BK current activity was observed at holding potentials positive to +20 mV (n = 4; ***P < 0.001, two-way ANOVA).
Figure 4
 
Arachidonic acid–induced activation of the BK current in retinal arteriolar myocytes. (Ai) BK currents evoked in response to depolarizing voltage steps (protocol shown in inset) under control conditions (upper panel) and following 5-minute application of 10 μM AA (lower panel). (Aii) Mean (±SEM) BK current densities (averaged across each depolarizing voltage step) under control conditions (closed circles) and in the presence of 10 μM AA (open circles). Note an enhancement of BK current at membrane potentials positive to +30 mV (n = 5; *P < 0.05, ***P < 0.001, two-way ANOVA). (Bi) Spontaneous BK currents recorded at various holding potentials under control conditions (left panel) and following 5 minute application of 10 μM AA (right panel). (Bii) Mean (±SEM) integrated spontaneous BK current densities under control conditions (closed circles) and in the presence of 10 μM AA (open circles). Enhancement of spontaneous BK current activity was observed at holding potentials positive to +20 mV (n = 4; ***P < 0.001, two-way ANOVA).
Figure 5
 
Arachidonic acid–induced inhibition of the L-type Ca2+ current in retinal arteriolar myocytes. (A) L-type Ca2+ currents under control conditions and following approximately 1-minute exposure to 10 μM AA evoked by depolarizing steps from −80 mV to 0 mV (inset). (B) Mean (±SEM) peak L-type Ca2+ current densities for the experimental conditions shown in (A) (n = 6; *P < 0.05 paired t-test).
Figure 5
 
Arachidonic acid–induced inhibition of the L-type Ca2+ current in retinal arteriolar myocytes. (A) L-type Ca2+ currents under control conditions and following approximately 1-minute exposure to 10 μM AA evoked by depolarizing steps from −80 mV to 0 mV (inset). (B) Mean (±SEM) peak L-type Ca2+ current densities for the experimental conditions shown in (A) (n = 6; *P < 0.05 paired t-test).
Table 1
 
Mean (±SEM) Changes in Internal Diameter of Pressurized Retinal Arterioles in Response to AA, ETYA, and Vehicle Controls
Table 1
 
Mean (±SEM) Changes in Internal Diameter of Pressurized Retinal Arterioles in Response to AA, ETYA, and Vehicle Controls
Baseline Diameter, μm Maximum Diameter in Presence of Drug, μm Diameter Following 5 Min of Wash Out
28.3 ± 1.5 AA (10 μM; n = 11) 30.8 ± 1.5† 30.3 ± 1.6†
25.5 ± 2.5 ETYA (10 μM; n = 8) 28.5 ± 2.4* 26.0 ± 2.2 NS
27.7 ± 3.0 DMSO (0.01% vol/vol; n = 5) 27.7 ± 3.0 NS NA
27.4 ± 1.4 Ethanol (0.1% vol/vol; n = 6) 27.6 ± 1.5 NS NA
Table 2
 
The Percentage of Myocytes Exhibiting Ca2+ Sparks and Oscillations and the Spatiotemporal Features of These Events in Pressurized Rat Retinal Arterioles Under Control Conditions and Following AA Treatment
Table 2
 
The Percentage of Myocytes Exhibiting Ca2+ Sparks and Oscillations and the Spatiotemporal Features of These Events in Pressurized Rat Retinal Arterioles Under Control Conditions and Following AA Treatment
Ca2+ Sparks Ca2+ Oscillations
Control AA Control AA
Active cells, % 52.3 ± 5.1 (105) 23.8 ± 8.9* (85) 70.0 ± 2.4 (105) 43.9 ± 9.8* (85)
Amplitude, F/F0 1.53 ± 0.06 (414) 1.55 ± 0.11 (99) 2.17 ± 0.07 (493) 1.70 ± 0.07† (156)
FDHM, ms 23.99 ± 0.88 (190) 23.07 ± 1.92 (41) 866.6 ± 0.03 (493) 770.6 ± 0.05‡ (493)
FWHM, μm 0.46 ± 0.01 (414) 0.41 ± 0.01 (99) NA NA
Frequency (cell−1 s−1) 0.066 ± 0.013 (105) 0.019 ± 0.005§ (85) 0.078 ± 0.008 (105) 0.031 ± 0.005§ (85)
Table 3
 
Mean (±SEM) Changes in Internal Diameter of Pressurized Retinal Arterioles in Response to AA Following Pretreatment With Pharmacologic Inhibitors
Table 3
 
Mean (±SEM) Changes in Internal Diameter of Pressurized Retinal Arterioles in Response to AA Following Pretreatment With Pharmacologic Inhibitors
Baseline Diameter, μm Maximum Diameter in Presence of Inhibitor, μm Maximum Diameter in Presence of Inhibitor and AA (10 μM), μm
28.5 ± 1.8 Ryanodine (100 μM; n = 9) 31.5 ± 1.1* Ryanodine + AA 32.4 ± 1.1†
29.3 ± 1.0 CPA (20 μM; n = 5) 31.6 ± 1.0‡ CPA + AA 31.6 ± 0.9‡
30.3 ± 1.5 Nimodipine (1 μM; n = 6) 32.5 ± 1.4† Nimodipine + AA 33.3 ± 1.6‡
28.4 ± 3.8 4-AP (10 mM), Penitrem A (100 nM; n = 5) 26.6 ± 3.7† 4-AP + Penitrem A + AA 28.5 ± 3.7
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