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(Circulation Research. 2008;102:e1.)
© 2008 American Heart Association, Inc.

UltraRapid Communications

AKAP150 Is Required for Stuttering Persistent Ca²⁺ Sparklets and Angiotensin II–Induced Hypertension

Manuel F. Navedo, Madeline Nieves-Cintrón, Gregory C. Amberg, Can Yuan, V. Scott Votaw, W. Jonathan Lederer, G. Stanley McKnight, Luis F. Santana

From the Departments of Physiology & Biophysics (M.F.N., M.N.-C., G.C.A., C.Y., V.S.V., L.F.S.) and Pharmacology (G.S.M.), University of Washington, Seattle; and Medical Biotechnology Center (W.J.L.), University of Maryland Biotechnology Institute, Baltimore. G.C. Amberg is currently with the Department of Biomedical Sciences, Colorado State University, Fort Collins.

Correspondence to Luis F. Santana, Department of Physiology & Biophysics, Box 357290, University of Washington, Seattle, WA 98195. E-mail santana{at}u.washington.edu

	Abstract

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Hypertension is a perplexing multiorgan disease involving renal primary pathology and enhanced angiotensin II vascular reactivity. Here, we report that a novel form of a local Ca²⁺ signaling in arterial smooth muscle is linked to the development of angiotensin II–induced hypertension. Long openings and reopenings of L-type Ca²⁺ channels in arterial myocytes produce stuttering persistent Ca²⁺ sparklets that increase Ca²⁺ influx and vascular tone. These stuttering persistent Ca²⁺ sparklets arise from the molecular interactions between the L-type Ca²⁺ channel and protein kinase C at only a few subsarcolemmal regions in resistance arteries. We have identified AKAP150 as the key protein, which targets protein kinase C to the L-type Ca²⁺ channels and thereby enables its regulatory function. Accordingly, AKAP150 knockout mice (AKAP150^–/–) were found to lack persistent Ca²⁺ sparklets and have lower arterial wall intracellular calcium ([Ca²⁺]_i) and decreased myogenic tone. Furthermore, AKAP150^–/– mice were hypotensive and did not develop angiotensin II–induced hypertension. We conclude that local control of L-type Ca²⁺ channel function is regulated by AKAP150-targeted protein kinase C signaling, which controls stuttering persistent Ca²⁺ influx, vascular tone, and blood pressure under physiological conditions and underlies angiotensin II–dependent hypertension.

Key Words: L-type Ca²⁺ channels • protein kinase C • myogenic tone • total internal reflection fluorescence microscopy

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
Hypertension is a major risk factor for the development of stroke, coronary artery disease, heart failure, and renal disease.¹ Although the principal cause of hypertension is renal, vascular dysfunction is critical,² and the increased arterial tone associated with hypertension contributes to the development of the pathology. This is highlighted by recent studies indicating that the endogenous vasoconstrictor angiotensin II is a likely contributor to vascular dysfunction in human^3,4 and model^5–13 hypertension. Accordingly, the angiotensin II signaling system is the target for two major classes of pharmacological agents (angiotensin-converting enzyme inhibitors and angiotensin II receptor antagonists) that are widely used clinically for the treatment of hypertension in humans. At present, however, the molecular mechanisms by which angiotensin II signaling causes vascular dysfunction and contributes to hypertension are unclear. Here, we address that conundrum and provide a novel and surprising answer.

Increased L-type Ca²⁺ channel activity in arterial myocytes is thought to be necessary for the development of vascular dysfunction during hypertension.^9,11,14–17 Ca²⁺ entry into arterial myocytes occurs through the opening of a single or a cluster of L-type Ca²⁺ channels. These channels produce local elevations in intracellular calcium ([Ca²⁺]_i) called "Ca²⁺ sparklets".^18–20 The activation of multiple Ca²⁺ sparklets increases global [Ca²⁺]_i, which activates myosin light chain, resulting in contraction.

Identification of sarcolemmal L-type Cav1.2 channels as the molecular entity underlying Ca²⁺ sparklets in arterial myocytes is based on multiple pharmacological, biophysical, and molecular biological studies. First, Ca²⁺ sparklets are eliminated by dihydropyridine antagonists and enhanced by dihydropyridine agonists and are insensitive to intracellular store depletion with thapsigargin.^18–20 Furthermore, unlike Ca²⁺ release events from intracellular stores, Ca²⁺ sparklets are rapidly eliminated by the removal of extracellular Ca²⁺.^18,19,21 Second, unlike Ca²⁺ release events from intracellular Ca²⁺ stores, Ca²⁺ sparklets are associated with an inward L-type Ca²⁺ current.^18,20 Consistent with this, Ca²⁺ sparklets have similar voltage dependencies of amplitude and activity as Cav1.2 channels.^18–21 Third, Ca²⁺ sparklets in tsA-201 cells expressing Cav1.2 channels reproduce all the basic features (eg, pharmacology, amplitude of quantal event, voltage dependence, activity, etc) of native Ca²⁺ sparklets.^19,21 Finally, arterial myocytes expressing dihydropyridine-insensitive Cav1.2 channels produce dihydropyridine-insensitive Ca²⁺ sparklets.²¹

A striking feature of Ca²⁺ sparklet activity in vascular smooth muscle is that it is modal.^18,19 In low activity mode, rare stochastic openings of solitary L-type Ca²⁺ channels produce limited Ca²⁺ influx. In contrast, discrete clusters of L-type Ca²⁺ channels that colocalized with protein kinase (PK) C operate in a sustained, high-activity mode, resulting in substantial Ca²⁺ influx (ie, "stuttering persistent Ca²⁺ sparklets").^18–20 Thus, in principle, vasoconstrictors such as angiotensin II, which activate PKC, could increase myogenic tone and blood pressure by specifically stimulating local persistent Ca²⁺ sparklet activity.

Although produced by <1% of the functional L-type Ca²⁺ channels located in an area representing <1% of the total surface membrane, persistent Ca²⁺ sparklets are a major contributor to Ca²⁺ influx and [Ca²⁺]_i in arterial myocytes.¹⁸ These findings are significant because they raise the intriguing possibility that the targeting of PKC to the sarcolemma of arterial myocytes is critical for local persistent Ca²⁺ sparklet activity and hence for the regulation of arterial wall [Ca²⁺]_i, myogenic tone, blood pressure, and the development of hypertension. Thus, delocalization of PKC from the sarcolemma could be a rational strategy for the selective elimination of persistent Ca²⁺ sparklets and the treatment of hypertension. However, the targeting mechanism directing PKC to the sarcolemma is unknown.

Here, we examined the role of the scaffolding protein AKAP150 in the regulation of Ca²⁺ sparklet activity, arterial wall [Ca²⁺]_i, myogenic tone, blood pressure, and angiotensin II–dependent hypertension. We focused on AKAP150 because it binds to PKC,^22–24 PKA, calcineurin, and L-type Ca²⁺ channels,²⁵ making this scaffolding protein a potential key regulator of persistent Ca²⁺ sparklet activity and thus vascular function and blood pressure. Our data indicate that AKAP150 is required for local targeting of PKC to the sarcolemma of arterial myocytes. Furthermore, we found that AKAP150 was necessary for PKC-dependent persistent Ca²⁺ sparklet activity, but not for rare, stochastic L-type Ca²⁺ channel activity. This suggests that AKAP150/PKC signaling does not increase L-type Ca²⁺ channel activity globally. Rather, this signaling complex increases Ca²⁺ influx locally by selectively coercing small clusters of L-type Ca²⁺ channels to operate in a persistent gating mode that create sites of sustained Ca²⁺ influx. We also found that AKAP150^–/– arteries had lower [Ca²⁺]_i and developed less myogenic tone than wild-type (WT) arteries. Consistent with this, AKAP150^–/– mice were hypotensive under control conditions and, unlike WT, did not develop hypertension during chronic angiotensin II infusion. We propose that stuttering persistent Ca²⁺ sparklet activity, myogenic tone, and blood pressure are regulated locally by the subcellular targeting of PKC by AKAP150.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Isolation of Arterial Myocytes
WT (C57BL/6J), AKAP150 knockout²⁵ (AKAP150^–/–, genetic background C57BL/6J), and mice expressing an AKAP150 lacking its PKA binding site (the carboxyl-terminal 36 amino acids [AKAP15036]) were euthanized with a lethal dose of sodium pentobarbital (250 mg/kg IP), as approved by the University of Washington Institutional Animal Care and Use Committee. Myocytes were dissociated from mesenteric and cerebral arteries using standard enzymatic techniques described elsewhere.²⁶ After dissociation, cells were maintained in a nominally Ca²⁺-free Ringer’s solution until used. Thapsigargin (1 µmol/L) was included in all solutions used to record Ca²⁺ sparklets to eliminate Ca²⁺ release from intracellular stores during experimentation.

RT-PCR and Western Blots
Total RNA was isolated from WT or AKAP150^–/– of 60 arterial myocytes (Figure 1A) or segments (Figures 2B and 3C) using the RNAeasy Micro kit (Qiagen, Valencia, Calif) as instructed by the manufacturer. We designed primers specific for PKC (GenBank accession No. NM_011101; sense nucleotides 1120 to 1140 and antisense nucleotides 1312 to 1331; amplicon=211 bp) and AKAP150 (GenBank accession XM_915706; sense nucleotides 58 to 80 and antisense nucleotides 444 to 464, amplicon=406 bp). The L-type Cav1.2 subunit (GenBank accession No. NM_012517) was detected with a set of commercial primers (SuperArray) that amplify a sequence between nucleotides 6548 to 6567 (amplicon=164 bp). We used β-actin (GenBank accession V01217; sense nucleotides 2384 to 2404 and antisense nucleotides 3071 to 3091; amplicon=496 bp) transcript levels as an internal control for these experiments. β-Actin primers amplify a region between exons 4 and 6 such that genomic contamination within the RNA preparation is identified by the presence of a 708-bp band in addition to the 496-bp band. Reverse transcription and amplification was performed using the OneStep RT-PCR kit from Qiagen following the instructions of the manufacturer. To do this, we used an Eppendorf Thermal Cycler running the following program: 35 cycles at 94°C for 1 minute; 56°C for 1 minute; 72°C for 1 minute each cycle, with a final extension step of 72°C for 10 minutes. The amplicons were visualized using 2% agarose gel electrophoresis. Relative abundance of the transcripts in AKAP150^–/– mice was normalized to the expression of the WT transcripts. An optical density analysis was used to quantify amplicons.

View larger version (28K): [in this window] [in a new window]   Figure 1. AKAP150 is expressed in arterial myocytes and is located in foci that colocalize with PKC near the sarcolemma of arterial myocytes. A, Image of an agarose gel showing AKAP150, PKC, and β-actin mRNA amplicons. mRNA was obtained from WT arterial myocytes. B, Representative confocal images of WT arterial myocytes labeled with Alexa 594–conjugated WGA (Bi) to identify the sarcolemma and a PKC-specific antibody (Bii) or WGA (Bv) and AKAP150-specific antibody (Bvi). Bii shows a surface plot of PKC-associated fluorescence in a typical WT cell. The images in Biii and Bvii were generated by merging the images Bi and Bii or Bv and Bvi, respectively. Biv and Bviii are surface plots of PKC-associated fluorescence (green wire frame) and WGA-associated fluorescence (solid, red surface plot) in the region within the white box in panels Biii and Bvii. C, Confocal images of a representative WT arterial myocyte labeled with AKAP150-specific (left) and PKC-specific (center) antibodies. The image to the right was produced by merging the AKAP150 and PKC images. The graph plots a frequency histogram of the PKC and AKAP150 fluorescence intensity pixel pairing in the membrane region in the images described above. The pixel pairing used for this plot had Pearson’s and Mander’s coefficients of +0.84 and +0.92, respectively.

View larger version (27K): [in this window] [in a new window]   Figure 2. AKAP150 is required for targeting of PKC to the sarcolemma of arterial myocytes. A, Confocal images of WGA- and PKC-associated fluorescence in representative WT (top row) and AKAP150–/– (bottom row) myocytes. Ai (WT) and Aiv (AKAP150–/–) are 2D images of WGA. Aii (WT) and Av (AKAP150–/–) are surface plots of PKC fluorescence. Shown in Aiii (WT) and Avi (AKAP150–/–) are 2D images produced by merging the WGA and PKC images. The graph plots the mean±SEM of the PKC surface/cytosol fluorescence ratio in WT (n=10) and AKAP150–/– (n=10) myocytes. B, Image of a gel showing PKC, AKAP150, and β-actin (control) mRNA amplification products from WT and AKAP150–/– arterial myocytes. The graph plots the mean±SEM of the relative (AKAP150–/–/WT) band intensity. C, Western blots of PKC, AKAP150, and β-actin (control) protein in WT and AKAP150–/– arteries. The bar plot shows the amount of PKC (relative to β-actin) in WT and AKAP150–/– arteries. *P<0.05 (2-tailed Student’s t test).

View larger version (15K): [in this window] [in a new window]   Figure 3. AKAP150 is required for basal, PKC-dependent regulation of ICa. A, Representative ICa recordings from WT and AKAP150–/– myocytes under control conditions and after the application of the PKC inhibitor Gö6976 (200 nmol/L) or the PKC agonist PDBu (200 nmol/L). The graph plots the mean±SEM of the amplitude of ICa (at +30 mV) in WT (n=9), WT+Gö6976 (n=4), WT+PDBu (n=4), AKAP150–/– (n=14), AKAP150–/–+Gö6976 (n=5), and AKAP150–/–+PDBu (n=9) myocytes. B, Agarose gel showing Cav1.2 and β-actin (control) mRNA amplicons from WT and AKAP150–/– arterial myocytes. The bar plot shows the mean± SEM of the normalized (AKAP150–/–/WT) band intensity. *P<0.05 (2-tailed Student’s t test).

Western blot analysis was performed using a previously described method⁹ with minor modifications. Briefly, WT or AKAP150^–/– mesentery and cerebral arteries were harvested and homogenized in ice-cold NP-40 buffer (1% Triton X-100, 0.15 mol/L NaCl, 0.01 mol/L sodium phosphate). These homogenates were then centrifuged at 10 000g for 10 minutes (4°C), and the supernatant was recovered. Protein concentration was determined using a Bio-Rad BCA kit. Total protein extracts were separated on 4% to 15% SDS–polyacrylamide gels (Bio-Rad) and transferred to nitrocellulose membranes. The membranes were blocked in Tris-buffered saline with Tween (TBST) (25 mmol/L Tris-HCL, pH7.3, 0.15 mol/L NaCl, 0.1% Tween-20) containing 5% milk and probed with primary antibody (mouse monoclonal anti-PKC antibody from Abcam; goat polyclonal anti-AKAP150 antibody from Santa Cruz Biotechnology, Santa Cruz, Calif; mouse monoclonal anti-Actin antibody from Chemicon) diluted in blocking buffer (1:250) for 1 hour. After extensive wash in TBST, membranes were probed with donkey anti-mouse or anti-goat IgG (Santa Cruz Biotechnology) and then washed in TBST. Horseradish peroxidase bound to immunoblot was visualized with enhanced chemiluminescence (ECL) (Amersham Biosciences) and Amersham ECL film. Protein was quantified using densitometry.

Immunofluorescence
To label the sarcolemma of arterial myocytes, we used an Alexa 594–conjugated wheat germ agglutinin (WGA), which binds to sialic acid residues of sarcolemmal glycoproteins. Briefly, isolated arterial myocytes were incubated in 10 µg/mL WGA for 1 hour before being plated on laminin-coated coverslips and allowed to settle for 2 hours. After plating, but before fixation and permeabilization (see below), cells were washed in PBS to remove any excess of WGA.

Immunofluorescence labeling of dispersed myocytes was performed as described previously²⁰ using a monoclonal antibody specific for PKC (Abcam, Cambridge, Mass), a polyclonal antibody specific for AKAP150 (Santa Cruz Biotechnology), and/or an Alexa 594–conjugated WGA to label the surface membrane (Molecular Probes). The secondary antibodies were an Alexa Fluor 488–conjugated rabbit anti-mouse (5 mg/mL), Alexa Fluor 488–conjugated donkey anti-goat, or an Alexa Fluor 568–conjugated donkey anti-goat (5 mg/mL) from Molecular Probes. Cells were imaged (512x512 pixel images) using our confocal system coupled with a Nikon x60 oil immersion lens (NA=1.4) and a zoom of 3.5 (pixel size=0.1 µm). The point spread function of our confocal system showed that it has a lateral and axial resolution of 0.30 µm and 0.80 µm, respectively. Images were collected at multiple optical planes (z-axis step size=1 µm).

We tested the specificity of our labeling by performing a negative control experiment in which the primary antibodies were substituted with PBS. PKC- or AKAP150-associated fluorescence was undetectable under these conditions (data not show). Double-labeling experiments with anti-PKC or anti-AKAP150 were used for colocalization analysis on paired of images (at multiple z-axis planes) using NIH ImageJ software with the colocalization analysis plug-in. Pearson’s correlation and Mander’s overlap coefficients²⁷ were calculated from those images. These analyses quantify the correlation or overlap between PKC and AKAP150 signals, ranging from 0 for no colocalization to 1 for complete colocalization. Fluorescence was quantified by measuring the intensity of pixels above a set threshold defined as the mean nonspecific fluorescence intensity within cells (ie, cell background, secondary antibody only) plus 3 times its SD in WT and AKAP150^–/– arterial myocytes. The location of PKC and AKAP150 clusters with respect to the membrane was determined by measuring the shortest distance between the epicenter of these clusters and the center of the WGA fluorescence at multiple focal planes.

We obtained the ratio of near-membrane to cytosol PKC using an approach similar to the one used by Khalil et al.²⁸ Briefly, for each cell, pixel intensities within a 1-µm section from the edge of the cell (which includes the cell membrane) were determined. Next, a spatially equivalent section of cytosol was selected and the intensity of each pixel averaged. From these measurements, we determined the ratio of PKC fluorescence at the different locations and used this as an indicator of PKC translocation and activity.²⁸ The delimitation of the cell was obtained using two independent approaches. (1) In some experiments we used a threshold analysis in which the boundary of the cell (ie, surface membrane) was detected as having background fluorescence intensity that was larger than the mean plus 3 SDs of the image intensity outside the cell. (2) We also used WGA-associated fluorescence to determine the location of the surface membrane. Note that WGA-associated fluorescence overlapped with the edge of the cell, as determined using the threshold analysis described above, indicating that either of these 2 approaches can be used to estimate the location of the sarcolemma of arterial myocytes.

Patch-Clamp Electrophysiology
We used the conventional whole-cell patch-clamp technique to control membrane voltage and record L-type Ca²⁺ currents using an Axopatch 200B amplifier. During experiments, cells were continuously superfused with a solution containing (mmol/L): 120 N'-methyl-D-glucamine, 5 CsCl, 1 MgCl₂, 10 glucose, 10 HEPES, and 20 CaCl₂ adjusted to pH 7.4. Pipettes were filled with a solution composed of (mmol/L) 87 Cs-aspartate, 20 CsCl, 1 MgCl₂, 5 MgATP, 10 HEPES, 10 EGTA, and adjusted to pH 7.2 with CsOH. For Ca²⁺ sparklets experiments, 0.2 mmol/L fluo-5F was added to the pipette solution. A voltage error of 10 mV resulting from liquid junction potential of these solutions was corrected for. Whole-cell L-type Ca²⁺ currents (I_Ca) were recorded using a step depolarization of 200 ms from the holding potential of –70 mV to +30 mV. I_Ca was measured as the difference between the peak and the sustained current at the end of the 200-ms pulse. Because murine cerebral arterial smooth muscle cells do not express functional T-type Ca²⁺ channels and are insensitive to dihydropyridine agonists and antagonists,^29,30 the voltage-gated, time-dependent currents evoked under these experimental conditions (ie, Na⁺ and K⁺ free external solution) have been shown to be produced by L-type Ca²⁺ channels. Currents were sampled at 20 kHz and low pass-filtered at 2 kHz.

[Ca²⁺]_i and Myogenic Tone in Pressurized Arteries
Arterial wall [Ca²⁺]_i was imaged using a Radiance 2100 confocal system (Cambridge, Mass) coupled to a Nikon TE2000S inverted microscope equipped with a Nikon 20X (NA=0.75) lens. Small mesenteric arteries (types II and III) from WT and AKAP150^–/– mice were removed and cleaned in normal Ringer’s solution. Isolated arteries were cannulated and mounted in a close-working-distance arteriograph. The endothelium of the artery was disrupted by passing air bubbles through their lumen. Endothelial damage was confirmed by the absence of a dilator response to acetylcholine (5 µmol/L) after the development of tone (data not shown). The arteriograph was placed on the stage of the inverted microscope and the artery was extraluminally perfused (3 to 6 mL/min) at 37°C with a bicarbonate-based physiological saline solution composed of (mmol/L): 119 NaCl, 4.7 KCl, 24 NaHCO₃, 1.2 KH₂PO₄, 1.6 CaCl₂, 1.2 MgSO₄, and 11 glucose with the pH set to 7.4 by bubbling with a gas mixture of O₂ (95%) and CO₂ (5%). After equilibration (20 minutes), intravascular pressure was increased to 80 mm Hg for 20 minutes. Arteries were loaded with the Ca²⁺ indicator fluo-4 as described elsewhere.²⁰ All fluorescence recordings were corrected for background (eg, autofluorescence, etc) before analysis. Arterial viability was tested by raising external K⁺ to 60 mmol/L. Arteries that failed to contract robustly in response to high [K⁺] were discarded.

Pressurized artery diameter measurements from WT and AKAP150^–/– mice were obtained from small mesenteric arterial segments 1 to 2 mm in length. These arterial segments were cannulated and mounted in a close-working-distance arteriograph. The arteriograph was placed on the stage of the inverted microscope (Nikon TE 2000S), and the artery segment was extraluminally perfused (3 to 6 mL/min) at 37°C with the same bicarbonate solution described above. After equilibration at 0 mm Hg for 20 minutes, steady-state changes in intravascular pressure were achieved by elevating or lowering an attached reservoir and monitored using a pressure transducer. Internal diameters were measured from live video images with the length-calibrated edge-detection function of an IonOptix imaging software at a sampling rate of 1 Hz. Myogenic tone was assessed by subjecting the vessels to a series of pressure steps between 40 and 100 mm Hg. Spontaneous myogenic tone was allowed to develop at each step. Experiments were terminated by the addition of 10 µmol/L diltiazem in nominally Ca²⁺-free bicarbonate-base solution, after which the pressure response curve was repeated to obtain the passive diameter of the artery. Myogenic tone was calculated as the percentage difference in active diameter versus passive diameter at each pressure. To determine angiotensin II–induced constriction, arteries were pressurized to 80 mm Hg and superfused with a solution containing 100 nmol/L angiotensin II. Induced constriction was defined as the percentage decrease in internal diameter after the application of the contractile stimuli versus passive diameter. Again, arterial viability was tested by raising external K⁺ to 60 mmol/L. Arteries that failed to contract robustly in response to high [K⁺] were discarded.

Analysis of fluorescence signals proceeded as follows. Background fluorescence (ie, arterial and nonarterial autofluorescence) was subtracted from the total fluorescence signal of fluo-4/5F–loaded arteries and cells. Fluo-4 and fluo-5F fluorescence signals were then converted to Ca²⁺ concentration units using the "F_max" equation³¹: equation

as described in detail previously.^19,20 Briefly, F is fluorescence, F_max is the fluorescence intensity of fluo-4/5F or rhod-2 in the presence of saturating free Ca²⁺, K_d is the dissociation constant of (fluo-5F=1280 nmol/L; fluo-4=800 nmol/L), and R_f (fluo-5F=286; fluo-4=150) is the F_max/F_min ratio of the indicator. F_min is the fluorescence intensity of the indicator in a solution in which the Ca²⁺ concentration is 0. K_d and R_f values were determined in vitro using standard methods and are similar to those reported by others.³² F_max was determined at the end of each experiment by exposing cells or arteries to the Ca²⁺ ionophore ionomycin (10 µmol/L) and 20 mmol/L external Ca²⁺.

We used the F_max method developed by Maravall et al³¹ to convert fluorescence signals from a nonratiometric Ca²⁺ indicator (fluo-5F or fluo-4) to calcium concentration units for multiple reasons. First, the F_max method has been thoroughly tested and, with indicators like fluo-4/5F, found to give [Ca²⁺]_i estimates that are similar to those obtained with ratiometric indicators. Second, we met all the conditions required to apply this method (ie, negligible rates of photobleaching; we used an indicator with a known dissociation constant that has a high affinity for Ca²⁺, as well as a high dynamic range, and we determined F_max for each cell). Third, unlike the pseudoratio method,³³ the F_max method does not require one to assume resting levels of [Ca²⁺]_i. This last point represents a major advantage of the F_max method.

Total Internal Reflection Fluorescence Microscopy
Ca²⁺ sparklets were recorded in patch-clamped (whole-cell configuration) arterial myocytes (from cerebral and mesenteric arteries) as previously described.^18–21 Cells were held at the hyperpolarized potential of –70 mV to increase the driving for Ca²⁺ entry (ie, relatively large Ca²⁺ sparklet amplitude) and thus allow the recording of quantal Ca²⁺ sparklets. Briefly, we used a through-the-lens total internal reflection fluorescence (TIRF) microscope built around an inverted Olympus IX-70 microscope equipped with an Olympus PlanApo (x60; NA=1.45) oil immersion lens and an Andor iXON CCD camera (South Windsor, Conn). To monitor [Ca²⁺]_i, cells were loaded with the calcium indicator fluo-5F as described above. As in previous studies,^18–21 we used the relatively slow Ca²⁺ buffer EGTA (on-rate 100-fold slower than fluo-5F) to enhance the "TIRF effect" and further restrict the fluorescence of fluo-5 to near the site of calcium entry (1 µm).³⁴ Note that a similar strategy has been used to record Ca²⁺ release sites in ventricular myocytes using confocal microscopy.^35,36 In this scenario, calcium entering the cell preferentially interacts with the faster fluo-5F, producing fluorescence, but then quickly (on average 2 ms)³⁴ binds to the more abundant and nonfluorescent EGTA. Thus, in our TIRF experiments in which fluo-5F and EGTA are used, intracellular calcium signals are limited to the submembrane space near the mouth of L-type Ca²⁺ channels. For a more detailed discussion of these issues, see Amberg et al.²⁰

Excitation of fluo-5F was achieved with the 488-nm line of an argon laser (Dynamic Lasers, Salt Lake City, Utah). Images were acquired at 100 to 300 Hz. As before,^18,19 we determined the activity of Ca²⁺ sparklets by calculating the nP_s of each Ca²⁺ sparklet site, where n is the number of quantal levels and P_s is the probability that a quantal Ca²⁺ sparklet event is active. As previously reported, all data sets were submitted to a normality test to determine whether they had a Gaussian distribution. Because Ca²⁺ sparklet activity has a bimodal distribution, Ca²⁺ sparklet sites were grouped into 3 categories: silent (by default has an nP_s of 0), low (nP_s between 0 and 0.2), and high (nP_s higher than 0.2). Comparison of the entire nP_s data set was performed using nonparametric data analyses (Mann–Whitney test). Comparison between low and high nP_s groups was performed using parametric data analyses (2-tailed Student’s t test). A detailed description of this analysis can be found in Navedo et al.¹⁹

Telemetry Blood Pressure Recordings
Blood pressure was monitored in conscious adult WT and AKAP150^–/– mice before and after angiotensin II treatment using a telemetry system (Data Science International). Briefly, mice were anesthetized with isoflurane. A ventral midline incision from the lower mandible to the sternum was made to isolate the left common carotid. Two lengths of 7-0 silk sutures were threaded beneath the vessel for retraction and ligation. The artery was permanently ligated at the level of the bifurcation between the interior and exterior carotid. The second suture was used to occlude blood flow temporally and allow insertion of the catheter. A 25-gauge syringe needle was used to make an incision in the artery through which the tip of the catheter was inserted. The catheter was advanced to the thoracic aorta and tied in place with the suture. A subcutaneous pocket was then made to place the transmitter body. After placement of the transmitter body the incision was closed with 5-0 sutures.

Blood pressure was recorded continuously and stored in the hard drive of a personal computer running Dataquest software (Data Science International). Control blood pressure measurements began 7 days after surgery to allow animals to recover. After this recovery period, blood pressures were recorded for 3 days before the saline or angiotensin II osmotic minipump was implanted. Angiotensin II was delivered at a rate of 800 ng/kg per minute.^9,11,37,38 Preparation and implantation of these pumps was performed as described elsewhere.^9,11 Blood pressure recordings continued for up to 7 days after pump implantation.

Chemicals and Statistics
All chemicals were from Sigma-Aldrich (St Louis, Mo) unless stated otherwise. Gö6976 was from Calbiochem. Normally distributed data are presented as means±SEM. Two-sample comparisons were made using a Student’s t test. Nonparametric statistical analyses (Mann–Whitney test) were used for nonnormally distributed data. Probability values less than 0.05 were considered significant. Asterisks used in the figures indicate a significant difference between groups.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
Investigation of the links between vascular Ca²⁺ signaling and blood pressure regulation under control conditions and following angiotensin II elevation requires the examination of L-type Ca²⁺ channels and related proteins in vascular smooth muscle. To investigate the molecular links between the L-type Ca²⁺ channel and PKC, we examined a candidate protein: AKAP150. This is a member of a family of proteins³⁹ known to be scaffolding molecules that facilitate local signaling between and among an assembly of proteins, often enabling the organization of a macromolecular signaling complex.

AKAP150 Targets PKC to the Sarcolemma of Arterial Myocytes
RT-PCR was used to determine what, if any, AKAP150 may be present in arterial smooth muscle myocytes. We found that AKAP150 transcript was expressed in arterial myocytes (Figure 1A). As a control, we also detected PKC and β-actin transcripts in these cells. This identification, however, raised the question of whether AKAP150 may be interacting with PKC as a functional entity, as hypothesized. An immunofluorescence approach was used to determine the spatial distribution of AKAP150 and PKC protein in WT arterial myocytes.

As shown in the representative cell in Figure 1B, we observed clear AKAP150- and PKC-associated fluorescence in WT arterial myocytes. For these experiments, an Alexa 594-conjugated WGA (see Figure 1Bi and 1Bv), which binds sialic acid residues in glycoproteins, was used to label the sarcolemma of arterial myocytes. PKC fluorescence was observed throughout the cell and in discrete foci near (<1 µm) WGA fluorescence in arterial myocytes (Figure 1Bii through 1Biv). Interestingly, AKAP150 fluorescence was confined to specific regions of the cell near the WGA fluorescence (Figure 1Biv through 1Bviii). A relatively small number of AKAP150 and PKC clusters were observed per cell (range=1 to 7 per cell, n=15), each with an area of 0.54±0.09 µm². AKAP150 fluorescence was never observed in AKAP150^–/– myocytes (data not shown). These data suggest that AKAP150 and PKC form clusters near the sarcolemma of arterial myocytes.

An immunofluorescence approach was also used to determine whether AKAP150 and PKC colocalize in WT arterial myocytes. As illustrated in Figure 1C, we observed colocalization of AKAP150 (red) and PKC (green) clusters near the boundaries of the cell (see Materials and Methods above for details about the spatial resolution of our microscope). Quantitative colocalization is shown in the plot of PKC fluorescence versus that of AKAP150 on a pixel-by-pixel basis. The statistical significance of the colocalization is supported by Mander’s overlap and Pearson correlation coefficients of +0.92 and +0.84, respectively. Together with the transcript and immunofluorescence data above, these data demonstrate that AKAP150 transcript and protein are expressed in arterial myocytes and that this scaffolding protein colocalizes with PKC in small subsarcolemmal clusters in these cells.

The colocalization of AKAP150 and PKC raised the question of a linkage between the 2 proteins and whether or not AKAP150 is required for PKC targeting to specific regions of the sarcolemma. If so, then PKC should not be found in puncta along the sarcolemma in the absence of AKAP150. Indeed, and unlike WT cells (Figure 2Ai through 2Aiii), clusters of PKC were never observed in AKAP150^–/– myocytes (Figure 2Aiv through 2Avi). Importantly, however, PKC was still present in the knockout cells, but was diffusely distributed throughout the cytoplasm (Figure 2Aiv through 2Avi). Consistent with this observation, we found that the overall abundance of PKC transcript (Figure 2B) and protein (Figure 2C) was similar in WT and AKAP150^–/– arteries. These data support the hypothesis that AKAP150 is required for local PKC targeting into the sarcolemma of arterial myocytes.

AKAP150 and PKC Increase Basal L-Type Ca²⁺ Channel Currents in Arterial Myocytes
Next, we examined I_Ca in WT and AKAP150^–/– myocytes (Figure 3). As demonstrated by others (eg, Cobine et al⁴⁰ and Amberg et al²⁰), inhibition of PKC with Gö69766 (200 nmol/L) decreased I_Ca in WT arterial myocytes, suggesting that basal PKC activity increases L-type Ca²⁺ channel function in these cells (Figure 3A). Conversely, application of the PKC activator phorbol 12,13-dibutyrate (PDBu) (200 nmol/L) increased I_Ca nearly 2.3-fold in WT cells (n=4; P<0.05).

A testable prediction from the immunofluorescence experiments described above is that I_Ca should be smaller in AKAP150^–/– than in WT myocytes because of loss of sarcolemmal localized PKC. Consistent with this hypothesis, we found that I_Ca was 63% smaller in AKAP150^–/– (n=14) than in WT myocytes (Figure 3A; n=9, P<0.05). In contrast to WT, application of Gö69766 or PDBu did not change I_Ca in AKAP150^–/– myocytes. Note also that the basal I_Ca in AKAP150^–/– cells was reduced to a level comparable to WT cells treated with a PKC inhibitor (n=4, P>0.05). Consistent with this, Cav1.2 (and PKC; see above), the predominant L-type Ca²⁺ channel in arterial myocytes,^21,30,41 transcript expression was similar (n=4; P>0.05) in WT and AKAP150^–/– arterial smooth muscle (Figure 3B). This suggests that differences in the expression of this pore-forming subunit do not account for differential I_Ca in WT and AKAP150^–/– myocytes. Together with the electrophysiological, immunofluorescence, and molecular biological (eg, PKC expression) data described above, these data support the hypothesis that delocalization of PKC is responsible for the decrease in I_Ca in AKAP150^–/– myocytes.

AKAP150 Is Required for Persistent Ca²⁺ Sparklet Activity in Arterial Myocytes
We imaged the elementary Ca²⁺ influx events underlying I_Ca in arterial myocytes: Ca²⁺ sparklets. This is critical because Ca²⁺ sparklet activity varies within the sarcolemma of arterial myocytes,¹⁹ which cannot be detected with the conventional whole-cell patch-clamp approaches used above. As described in the introduction, the majority of L-type Ca²⁺ channels in these cells operate in low open probability mode that causes small, sporadic elevations in [Ca²⁺]_i (ie, low-activity Ca²⁺ sparklets). In contrast, discrete clusters of L-type Ca²⁺ channels associated with PKC operate in a sustained, high-activity mode that result in substantial Ca²⁺ influx (ie, stuttering persistent Ca²⁺ sparklets).^18–20 Regional variations in PKC activity have been suggested to underlie this novel Ca²⁺ signaling modality. However, the mechanisms underlying heterogeneous PKC distribution and Ca²⁺ sparklet activity in these cells are unclear. One potential mechanism suggested by the data described above is that AKAP150 targets PKC to subsarcolemmal microdomains, where it can modulate the function of nearby L-type Ca²⁺ channels and induce stuttering persistent Ca²⁺ sparklet activity.

One testable prediction of this hypothesis is that persistent Ca²⁺ sparklet activity should either be lower or absent in AKAP150^–/– compared with WT arterial myocytes. Consistent with this, Ca²⁺ sparklet activity (nP_s; –70 mV) was lower in AKAP150^–/– than in WT cells (Figure 4). Indeed, Ca²⁺ sparklets were rarely observed in AKAP150^–/– myocytes (Figure 4A and 4C). The lower level of Ca²⁺ sparklet activity in AKAP150^–/– myocytes was attributable to an absence of high activity, persistent Ca²⁺ sparklet sites (nP_s>0.2) (Figure 4B and 4D) as the mean nP_s value of low activity Ca²⁺ sparklets was similar in WT (0.11±0.05; n=55) and AKAP150^–/– myocytes (0.10±0.01; n=8; P=0.38).

View larger version (23K): [in this window] [in a new window]   Figure 4. AKAP150 is required for persistent Ca2+ sparklet activity in arterial myocytes. A, TIRF images of representative WT and AKAP150–/– arterial myocytes. Traces below each image show the time course of [Ca2+]i in the green circle before (top) and after PDBu (200 nmol/L) (bottom) treatment. B, Scatter plot of nPs values for individual Ca2+ sparklet sites in WT (n=7) and AKAP150–/– (n=5) under control conditions and after application of PDBu. *P<0.05 (Mann–Whitney test). C, Time courses of [Ca2+]i in Ca2+ sparklet sites from representative WT and AKAP150–/– cells before and after application of 100 nmol/L angiotensin II. D, Scatter plot of nPs values for individual Ca2+ sparklet sites in WT (n=7) and AKAP150–/– (n=5) under control conditions and after application of angiotensin II. The dashed lines in B and D represent the threshold for high nPs sites. *P<0.05 (Mann–Whitney test).

Activation of PKC increases persistent Ca²⁺ sparklet activity in arterial myocytes.¹⁹ Thus, we examined the effects of direct and receptor-mediated PKC activation with PDBu (200 nmol/L) and angiotensin II (100 nmol/L) on Ca²⁺ sparklet activity in WT and AKAP150^–/– myocytes. We found that application of these vasoconstrictors evoked persistent Ca²⁺ sparklet activity in WT but not in AKAP150^–/– myocytes (Figure 4A through 4D; see also Movie 1 in the online data supplement).

In addition to PKC, AKAP150 also binds calcineurin and PKA.²³ In WT myocytes, inhibition of calcineurin with cyclosporin A increased Ca²⁺ sparklet activity by 3.1-fold¹⁹ but was without effect in AKAP150^–/– arterial myocytes (n=5; data not shown). We also investigated the possibility that PKA binding to AKAP150 was participating in the regulation of PKC-dependent persistent Ca²⁺ sparklet activity. To do this, we examined Ca²⁺ sparklet activity in WT cells and in arterial myocytes from a mouse expressing an AKAP150 lacking its PKA binding site (the carboxyl-terminal 36 amino acids; referred to as AKAP15036) before and after application of PDBu (see Figure 5). Unlike AKAP150^–/– cells, WT levels of low and high nP_s Ca²⁺ sparklets were observed in AKAP15036 myocytes under control conditions. Furthermore, application of PDBu (200 nmol/L) or cyclosporin A (500 nmol/L; Figure 5) increased persistent Ca²⁺ sparklet activity to the same extent in AKAP15036 (n=5) and WT myocytes (n=5, P>0.05) but not in AKAP150^–/– myocytes. Together with the AKAP150^–/– data above, these findings demonstrate that association of PKC, but not PKA, with AKAP150 is required for localized persistent Ca²⁺ sparklet activity in arterial myocytes.

View larger version (13K): [in this window] [in a new window]   Figure 5. PKA binding to AKAP150 is not required for PKC-dependent or calcineurin regulation of persistent Ca2+ sparklet activity in arterial myocytes. A, Time courses of [Ca2+]i in Ca2+ sparklet sites from representative WT and AKAP15036 cells before and after application 200 nmol/L PDBu. B, Scatter plot of nPs values for individual Ca2+ sparklet sites in WT (n=7) and AKAP15036 (n=6) under control conditions and after application of PBDu. *P<0.05 (Mann–Whitney test). C, Time courses of [Ca2+]i in Ca2+ sparklet sites from a representative AKAP15036 cell before and after application of 500 nmol/L cyclosporine A (CSA). D, Scatter plot of nPs values for individual Ca2+ sparklet sites in AKAP15036 under control conditions and after application of cyclosporine A (n=6). *P<0.05 (Mann–Whitney test). The dashed lines in B and D represent the threshold for high nPs sites.

Loss of AKAP150 Decreases Arterial Wall [Ca²⁺]_i and Tone
Next, we compared arterial wall [Ca²⁺]_i in pressurized (80 mm Hg) arterial segments from WT and AKAP150^–/– arterial segments (Figure 6A). We found that under control conditions, [Ca²⁺]_i was lower in AKAP150^–/– (n=5) than in WT (n=5, P<0.05) arteries. Transient application of angiotensin II (100 nmol/L) increased [Ca²⁺]_i in AKAP150^–/– arteries slightly, but to a significantly smaller extent, than in WT arteries (Figure 6B; P<0.05). Note that in the presence of the L-type Ca²⁺ channel blocker nifedipine (1 µmol/L) arterial wall [Ca²⁺]_i was similar in WT and AKAP150^–/– arteries (100 nmol/L; P>0.05), as expected if the L-type Ca²⁺ channel were ultimately responsible for the effect.

View larger version (21K): [in this window] [in a new window]   Figure 6. Arterial wall [Ca2+]i and tone are lower in pressurized AKAP150–/– than in WT arteries. A, Time course of arterial wall [Ca2+]i in pressurized (80 mm Hg) WT and AKAP150–/– arteries during control conditions and after the application of 100 nmol/L angiotensin II or 1 µmol/L nifedipine. B, Bar plot of the mean±SEM of the [Ca2+]i in WT (n=5) and AKAP150–/– (n=5) arteries under control conditions and after the application of 100 nmol/L angiotensin II or 1 µmol/L nifedipine. C, Time course of arterial diameter during an increase in intravascular pressure from 40 to 60 mm Hg under control conditions (ie, when the artery has tone) and in the presence of an external Ca2+-free solution containing 10 µmol/L diltiazem. The horizontal lines indicate the time the artery was maintained at the indicated pressure. D, Plot of myogenic tone as a function of intravascular pressure in WT (n=5) and AKAP150–/– (n=5) arteries. E, Bar plot of the mean±SEM of the contraction induced by angiotensin II (80 mm Hg) in WT and AKAP150–/– arteries. *P<0.05 (2-tailed Student’s t test).

To examine the physiological consequences of reduced I_Ca, Ca²⁺ sparklet activity, and [Ca²⁺]_i in AKAP150^–/– smooth muscle, we evaluated the effect of pressure on the diameter of WT and AKAP150^–/– arteries (Figure 6C and 6D). Elevation of intravascular pressure constricts small resistance arteries.⁴² Consistent with our I_Ca, Ca²⁺ sparklet, and [Ca²⁺]_i data, AKAP150^–/– arteries (n=5) were significantly less constricted than WT arteries at all intravascular pressures examined (Figure 6D; n=5; P<0.05). Furthermore, angiotensin II evoked a smaller contraction in AKAP150^–/– than in WT arteries (Figure 6E). These findings suggest that AKAP150 is required for basal and angiotensin II–induced changes in arterial [Ca²⁺]_i and is an important component of myogenic tone. Furthermore, our results are consistent with the hypothesis that physiological modulation of vascular tone by angiotensin II depends on the modulation of stuttering persistent Ca²⁺ sparklets.

In addition to increasing [Ca²⁺]_i, PKC enhances smooth muscle contraction by increasing the sensitivity of contractile filaments to [Ca²⁺]_i.^43,44 AKAP150 could thus be necessary for PKC-induced sensitization of myofilaments during angiotensin II signaling. To provide insight into this fundamental issue, we calculated the ratio of the mean arterial tone to [Ca²⁺]_i during angiotensin II treatment in WT and AKAP150^–/– arteries (see Figure 6). We found that, on average, the ratio of arterial tone to [Ca²⁺]_i was similar in AKAP150^–/– (29% tone per 216 nmol/L=0.13% tone per nmol/L) and WT (50% tone per 369 nmol/L=0.14% tone per nmol/L) arteries. This analysis suggests that the amount of myogenic tone developed per unit of [Ca²⁺]_i was similar in WT and AKAP150^–/– arteries during angiotensin II signaling. Thus, although targeting of PKC to the sarcolemma by AKAP150 is required for activation of persistent Ca²⁺ sparklet activity, it is not essential for the modulation of the Ca²⁺ sensitivity of contractile filaments during angiotensin II signaling.

AKAP150 Is Required for Angiotensin II–Induced Hypertension
Chronic activation of angiotensin II signaling contributes to the development of hypertension in mice and humans.^3,4,9,11 To examine the role of AKAP150/PKC-dependent, persistent Ca²⁺ sparklets in this process, we examined blood pressure in AKAP150^–/– and WT mice infused with saline (control) or angiotensin II (Figure 7A). Mean arterial pressure (MAP) was significantly lower in AKAP150^–/– (102±2 mm Hg, n=9) than in WT (116±3 mm Hg, n=6; P<0.05) mice infused with saline. As expected, angiotensin II infusion significantly increased MAP in WT to 149±3 mm Hg (P<0.05). Interestingly, MAP in angiotensin II–infused AKAP150^–/– mice (120±3 mm Hg) was similar (P>0.05) to that of normotensive WT mice infused with saline. Thus, although angiotensin II induced a small increase in blood pressure in AKAP150^–/– mice (MAP=14±4 mm Hg), this response was 2.4-fold lower than in WT mice (MAP=33±4 mm Hg; P<0.05) and equalized MAP in AKAP150^–/– infused with angiotensin II and normotensive, saline-infused WT mice (P>0.05). These data demonstrate that AKAP150 regulates basal blood pressure and plays a critical role in the development of angiotensin II–dependent hypertension.

View larger version (14K): [in this window] [in a new window]   Figure 7. AKAP150 is required for angiotensin II–induced hypertension. A, Representative pressure waveforms of conscious WT and AKAP150–/– mice before and after angiotensin II administration (right). B, Bar plot of MAP in WT (n=6) and AKAP150–/– (n=9) mice infused with saline (control) or angiotensin II. MAP was similar in normotensive, saline-infused WT mice and AKAP150–/– angiotensin II–infused mice (P>0.05). Angiotensin II–infused WT and AKAP150–/– mice had MAPs of 149±3 mm Hg (n=5) and 120±3 mm Hg (n=8), respectively. *P<0.05 (2-tailed Student’s t test). C, Proposed mechanism by which subcellular targeting of PKC and calcineurin by AKAP150 control local stuttering persistent Ca2+ sparklets. Under physiological conditions, low activity and stuttering persistent Ca2+ sparklets contribute to arterial [Ca2+]i and hence to the development of myogenic tone and the regulation of blood pressure. Increased angiotensin II signaling activates stuttering persistent Ca2+ sparklets signals in arterial myocytes via AKAP150/PKC signaling. This increases arterial [Ca2+]i and myogenic tone and contributes to hypertension.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
The findings reported here suggest that AKAP150 is required for stuttering persistent Ca²⁺ sparklets and for the regulation of myogenic tone, blood pressure, and the development of angiotensin II–induced hypertension. Indeed, our data support a new model of Ca²⁺ signaling in vascular smooth muscle in which AKAP150 targets PKC to relatively small regions of the sarcolemma of arterial myocytes. Local targeting of PKC by AKAP150 is responsible for persistent Ca²⁺ sparklet activity, but not for rare, stochastic L-type Ca²⁺ channel openings (ie, low-activity Ca²⁺ sparklets). Our data suggest that AKAP150 is required for PKC-dependent persistent Ca²⁺ sparklets and thus regulates arterial [Ca²⁺]_i, myogenic tone, and blood pressure. Furthermore, AKAP150 is necessary for the development of angiotensin II–induced hypertension. Thus, we propose that AKAP150 regulates myogenic tone, blood pressure, and the development of angiotensin II–induced hypertension, at least in part, by local control of PKC-dependent stuttering persistent Ca²⁺ influx. The implications of these findings are discussed below.

The evidence linking PKC to persistent Ca²⁺ sparklet activity is numerous. First, pharmacological inhibition of PKC eliminates persistent Ca²⁺ sparklet activity.^18–20 Second, PKC-null (PKC^–/–) arterial myocytes, which presumably express AKAP150 and other associated proteins, have low-activity, but not high-activity, persistent Ca²⁺ sparklet activity.^19,20 Third, activation of PKC increases persistent Ca²⁺ sparklet activity. Fourth, expression of PKC and Cav1.2 channels in tsA-201 cells is sufficient to produce persistent Ca²⁺ sparklets. Ca²⁺ sparklets were never observed in control (ie, nontransfected) tsA-201 cells.^19,21 These findings provide compelling support for the hypothesis that PKC and L-type Ca²⁺ channel expression is necessary for basal and evoked stuttering persistent Ca²⁺ sparklet activity in arterial myocytes and raise an important question: What are the mechanisms leading to local PKC-dependent persistent Ca²⁺ sparklet activity?

Examination of our data provides insight into this important issue. L-type Ca²⁺ channels are expressed throughout the surface sarcolemma of smooth muscle cells.^18,45 By contrast, in these cells, PKC is found in small clusters near the sarcolemma of WT, but not in AKAP150^–/–, myocytes. Note, however, that PKC transcript and protein expression was similar in WT and AKAP150^–/– arteries. Yet AKAP150^–/– myocytes, as with PKC^–/– myocytes¹⁹ (see above), do not have persistent Ca²⁺ sparklet activity, suggesting that PKC expression alone is insufficient to induce persistent Ca²⁺ sparklet activity in arterial myocytes. Rather, these data give credence to the view that targeting of PKC by AKAP150 is crucial for the local control of stuttering persistent Ca²⁺ sparklet activity in arterial myocytes.

Although our TIRF data provide compelling support to the hypothesis that Ca²⁺ channel activity varies regionally within arterial myocytes, analysis of our immunofluorescence images provides further support for this concept. On average, the surface area of an arterial myocyte is 1000 µm².¹⁸ We found that there are 7 AKAP150/PKC clusters of 0.5 µm² per cell (see Results section above). Accordingly, these clusters would account for only 0.3% (3.5 µm²/1000 µm²x100) of the entire sarcolemma. This analysis suggests that a relatively small fraction of the sarcolemma of arterial myocytes participates in AKAP150/PKC-mediated persistent Ca²⁺ sparklet signaling, arterial function and thus blood pressure regulation.

In WT myocytes, activation of PKC increases Ca²⁺ influx by activating persistent Ca²⁺ sparklet activity.^18,19 However, AKAP150^–/– myocytes did not have persistent Ca²⁺ sparklet activity and consequently had lower arterial [Ca²⁺]_i, tone, and response during angiotensin II signaling. Importantly, AKAP150^–/– mice did not develop angiotensin II–induced hypertension even though total PKC expression is similar in WT and AKAP150^–/– arteries. On the basis of these findings, we suggest that PKC targeting to specific foci in the sarcolemma by AKAP150 is necessary for persistent Ca²⁺ sparklet activity in arterial myocytes and may contribute to the development of hypertension.

Elimination of persistent Ca²⁺ sparklet activity per se would decrease arterial [Ca²⁺]_i and hence tone and blood pressure in AKAP150^–/– mice. However, it is important to note that it is possible that other AKAP150-dependent mechanisms, in combination with the loss of persistent Ca²⁺ sparklets, also contribute to lower myogenic tone and blood pressure in these mice. For example, our data clearly indicate that angiotensin II increases Ca²⁺ sparklet activity (ie, Ca²⁺ influx) in WT arterial myocytes, presumably through the activation of PKC, with no changes in membrane potential (ie, in a voltage-clamped myocytes). However, in vivo activation of angiotensin II/PKC signaling has also been shown to inhibit voltage-gated, delayed-rectifier K⁺ channels⁴⁶ and large conductance, Ca²⁺-activated K⁺ channels,^47,48 which would lead to arterial smooth muscle depolarization. In addition, as demonstrated by Amberg et al,²⁰ membrane depolarization increases low- and high-activity persistent Ca²⁺ sparklets in arterial myocytes. Consistent with this, 2 recent studies have suggested that PKC activity plays an important role in the regulation of membrane potential, Ca²⁺ influx via L-type Ca²⁺ channels, and the development of myogenic tone in coronary⁴⁰ and cerebral arteries.⁴⁹ Thus, in vivo, increased angiotensin II/PKC signaling could increase Ca²⁺ sparklet activity by voltage-dependent and voltage-independent mechanisms.

AKAP150 binds PKC and calcineurin but also PKA.²³ Activation of PKA causes vasodilation,⁵⁰ at least in part, by decreasing arterial smooth muscle [Ca²⁺]_i. Accordingly, loss of AKAP150, which would presumably eliminate targeting of PKA to the sarcolemma, could potentially lead to increased [Ca²⁺]_i, vasoconstriction, and blood pressure. However, Ca²⁺ sparklet activity, I_Ca, [Ca²⁺]_i, and blood pressure were lower in AKAP150^–/– than in WT mice. How could these seemingly contradictory findings be reconciled? One intriguing possibility is that the primary role of AKAP150 is to target PKC, not PKA, near L-type Ca²⁺ channels in arterial myocytes. Consistent with this, Ca²⁺ sparklet activity was similar in the AKAP15036 mutant, which does not bind PKA, and WT myocytes. Furthermore, we found that activation of PKA with forskolin did not increase Ca²⁺ sparklet activity in WT arterial myocytes. Thus, an AKAP other than AKAP150 may be the predominant anchor for PKA to the sarcolemma of arterial myocytes. Alternatively, PKA activity could be relatively low under basal conditions and during angiotensin II signaling, thus providing little opposition to the actions of PKC on Ca²⁺ influx in arterial myocytes.

To conclude, our data indicate that AKAP150 plays a crucial role in the regulation of vascular function (Figure 7D). By targeting PKC, not PKA, and presumably calcineurin to specific sarcolemmal microdomains near L-type Ca²⁺ channels, AKAP150 coordinates the formation of a signaling module that controls Ca²⁺ influx, [Ca²⁺]_i, arterial tone, and hence blood pressure under physiological and pathophysiological conditions.

	Acknowledgments

 
Sources of Funding

This work was supported by NIH grants HL077115 and HL085870 (to L.F.S.) and American Heart Association Grants 0635118N (to G.C.A.) and 0735251N (to M.F.N.). L.F.S. is an Established Investigator of the American Heart Association.

Disclosures

None.

	Footnotes

 
Original received September 7, 2007; resubmission received November 13, 2007; revised resubmission received December 13, 2007; accepted December 17, 2007.

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