Intravascular pressure regulates local and global Ca2+ signaling in cerebral artery smooth muscle cells

Jonathan H. Jaggar

Abstract

The regulation of intracellular Ca2+ signals in smooth muscle cells and arterial diameter by intravascular pressure was investigated in rat cerebral arteries (∼150 μm) using a laser scanning confocal microscope and the fluorescent Ca2+ indicator fluo 3. Elevation of pressure from 10 to 60 mmHg increased Ca2+spark frequency 2.6-fold, Ca2+ wave frequency 1.9-fold, and global intracellular Ca2+ concentration ([Ca2+]i) 1.4-fold in smooth muscle cells, and constricted arteries. Ryanodine (10 μM), an inhibitor of ryanodine-sensitive Ca2+ release channels, or thapsigargin (100 nM), an inhibitor of the sarcoplasmic reticulum Ca2+-ATPase, abolished sparks and waves, elevated global [Ca2+]i, and constricted pressurized (60 mmHg) arteries. Diltiazem (25 μM), a voltage-dependent Ca2+ channel (VDCC) blocker, significantly reduced sparks, waves, and global [Ca2+]i, and dilated pressurized (60 mmHg) arteries. Steady membrane depolarization elevated Ca2+ signaling similar to pressure and increased transient Ca2+-sensitive K+ channel current frequencye-fold for ∼7 mV, and these effects were prevented by VDCC blockers. Data are consistent with the hypothesis that pressure induces a steady membrane depolarization that activates VDCCs, leading to an elevation of spark frequency, wave frequency, and global [Ca2+]i. In addition, pressure induces contraction via an elevation of global [Ca2+]i, whereas the net effect of sparks and waves, which do not significantly contribute to global [Ca2+]i in arteries pressurized to between 10 and 60 mmHg, is to oppose contraction.

  • ryanodine-sensitive calcium release channel
  • voltage-dependent calcium channel
  • calcium-sensitive potassium channel
  • spark
  • wave

calcium(Ca2+) is an important second messenger that regulates a variety of cellular processes, including contraction and gene expression (for review, see Ref. 11). In small (“resistance size”) arteries, an elevation of intravascular pressure leads to a graded membrane potential depolarization (16,24) that activates voltage-dependent Ca2+ channels on smooth muscle cells of the arterial wall (24). This leads to a graded elevation in the intracellular Ca2+concentration ([Ca2+]i) of the arterial wall and a maintained constriction termed “myogenic tone” (1, 24, for review see Ref. 13).

Previous studies that investigated the regulation of arterial smooth muscle [Ca2+]i by pressure employed techniques that measured the average [Ca2+]iof many cells in the arterial wall (e.g., see Refs. 24 and29). However, recent studies have shown that the [Ca2+]i of smooth muscle cells is not homogeneously distributed, and elevations of [Ca2+]i occur that differ in respect to spatial localization, temporal kinetics, and physiological function (19, 21, 30, 32, 39). The effect of pressure changes on the properties of these intracellular Ca2+ signals is unknown. Understanding the regulation of intracellular Ca2+signaling events by pressure may provide important insights of signal transduction pathways that modulate arterial diameter.

Three cytosolic Ca2+ signaling modalities have been described in arterial smooth muscle cells: Ca2+ sparks, Ca2+ waves, and global [Ca2+]i. Ca2+ sparks are highly localized elevations of cytosolic Ca2+ that occur due to the activation of a number of ryanodine-sensitive Ca2+ release (RyR) channels on the sarcoplasmic reticulum (SR) (for review, see Ref. 21). In arterial smooth muscle, most Ca2+ sparks occur in close proximity to the plasma membrane and activate a number of Ca2+-sensitive K+ (KCa) channels to induce a transient outward K+ current [previously referred to as a “spontaneous transient outward current” (STOC); Ref.2] (21, 32, 34). At physiological potentials, Ca2+ sparks signal KCa channels in arterial smooth muscle primarily via the associated β1-subunit (6, 36) and are estimated to increase KCa channel activity 104–106-fold at −40 mV (34). Inhibition of Ca2+ sparks leads to a membrane potential depolarization, activation of voltage-dependent Ca2+channels, an elevation of arterial wall [Ca2+]i, and constriction (25,32). Although the local change in [Ca2+]i sensed by KCa channels is estimated to be high (10–100 μM) (34), the effect of Ca2+ sparks on global cytosolic Ca2+ is low (<2 nM), since Ca2+ sparks occur at a low frequency (∼1 Hz) and spread into only ∼1% of the cellular volume (21,32).

In contrast, Ca2+ waves are propagating elevations of cytosolic Ca2+ that have been proposed to mediate cellular contraction because Ca2+ wave frequency is elevated by vasoconstrictors that elevate inositol trisphosphate (IP3) (19, 20, 30, 39) or by high concentrations of caffeine, an activator of RyR channels, that induce contraction (39). UTP, a vasoconstrictor that activates phospholipase C, inhibits Ca2+ sparks and activates Ca2+ waves in arterial smooth muscle cells, suggesting that vasoconstrictors shift Ca2+ signaling from dilatory to contractile modalities (20). Global [Ca2+]i is the cytosolic [Ca2+] of arterial smooth muscle cells that may result from Ca2+entry through voltage-dependent Ca2+ channels and Ca2+ release from intracellular stores. Blockers of voltage-dependent Ca2+ channels decrease arterial wall [Ca2+]i and dilate arteries (5,24), suggesting the global [Ca2+]i of arterial smooth muscle cells is an important regulator of cellular contraction and thus arterial diameter.

The goal of the present study was to investigate the regulation of Ca2+ sparks, Ca2+ waves, and global [Ca2+]i in arterial smooth muscle cells by intravascular pressure and to examine the regulation of arterial diameter by these intracellular Ca2+ signaling modalities. Cytosolic Ca2+ signals were measured in arterial smooth muscle cells of small (∼150 μm) cerebral arteries at 37°C using a laser scanning confocal microscope and the fluorescent Ca2+indicator fluo 3. Data were consistent with the hypothesis that elevating pressure from 10 to 60 mmHg induced a steady membrane depolarization that activated voltage-dependent Ca2+channels, leading to an elevation of Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in arterial smooth muscle cells and vasoconstriction. Data suggest that pressure constricts via an elevation in global [Ca2+]i, which occurs primarily due to Ca2+ entry via voltage-dependent Ca2+ channels. Sparks and waves occur due to Ca2+ release through RyR channels on the SR and do not contribute significantly to global [Ca2+]i, and the net effect of sparks and waves is to oppose contraction.

METHODS

Tissue preparation.

Sprague-Dawley rats (∼250 g) of either sex were euthanized by peritoneal injection of pentobarbital sodium solution (150 mg/kg). The brain was removed and placed into ice-cold (4°C), oxygenated (95% O2-5% CO2), bicarbonate solution containing (in mM) 119 NaCl, 4.7 KCl, 24 NaHCO3, 1.2 KH2PO4, 1.6 CaCl2, 1.2 MgSO4, 0.023 EDTA, and 11 glucose (pH 7.4). Posterior cerebral and cerebellar arteries (100–200 μm) were removed, cleaned of basolateral connective tissue, and maintained in ice-cold bicarbonate solution until used.

Pressurization of cerebral artery segments and confocal Ca2+ measurements.

Arterial segments (1–2 mm in length) were placed into a HEPES-buffered saline solution of the following composition (in mM): 136 NaCl, 6 KCl, 10 HEPES, 2 CaCl2, 1 MgCl2, and 10 glucose (pH 7.4 with NaOH), containing 15 μM fluo 3-AM (Molecular Probes) and 0.05% Pluronic acid (Molecular Probes) for 60 min at 22°C. To allow indicator deesterification, arteries were subsequently placed into HEPES-buffered saline solution (in the absence of fluo 3) for 30 min at 22°C.

All the following procedures were performed in a HEPES/bicarbonate solution of composition (in mM) 112 NaCl, 4.8 KCl, 10 HEPES, 26 NaHCO3, 1 CaCl2, 1.2 MgSO4, 1.2 KH2PO4, and 5 glucose, gassed with 74% N2-21% O2-5% CO2. Arterial segments were cannulated at each end in a vessel chamber (Living Systems Instrumentation, Burlington, VT). The vessel chamber was placed on the stage of a Nikon TE300 microscope and superfused (3–6 ml/min) with warmed (37°C), gassed, HEPES/bicarbonate solution. At 37°C the pH, Po 2, and Pco 2 of the HEPES/bicarbonate solution in the experimental chamber were 7.4, 150–160 mmHg, and 35–40 mmHg, respectively, as measured using a blood gas analyzer (Instrumentation Laboratory, Lexington, MA). After a 10-min equilibration period, one cannula was stoppered and the other was attached to a reservoir that could be elevated or lowered to attain steady intravascular transmural pressures of either 10 or 60 mmHg (measured with a perfusion pressure monitor; Living Systems Instrumentation). Smooth muscle cells within the artery wall were imaged using a Noran Oz laser scanning confocal microscope and a ×60 water-immersion lens (numerical aperture = 1.2) by illuminating with a krypton/argon laser at 488 nm. Emitted light >500 nm was captured. To ensure efficient quantum capture, the artery was placed near the bottom of the chamber. Images of the arterial wall (56.3 × 52.8 μm or 256 × 240 pixels) were recorded every 16.7 ms (i.e., 60 images/s). Under each condition, at least two different representative areas of the same arterial segment were each scanned for 10 s. The same area of artery was only scanned once to avoid any laser-induced changes in Ca2+signaling, and the effects of pressure or drugs were measured in paired experiments. In some experiments, an elevated K+ (30 mM K+) HEPES/bicarbonate solution of composition (in mM) 88 NaCl, 28.8 KCl, 10 HEPES, 26 NaHCO3, 1 CaCl2, 1.2 MgSO4, 1.2 KH2PO4, and 5 glucose (gassed with 74% N2-21% O2-5% CO2, pH 7.4 at 37°C) was used to depolarize arteries pressurized to 10 mmHg to a membrane potential of approximately −40 mV (3, 15, 17), which is similar to the membrane potential of arteries pressurized to 60 mmHg (16, 24). Luminal diameter was measured by acquiring images of pressurized artery segments under various conditions using a ×10 objective (numerical aperture = 0.25) and the transmitted light detector of the confocal microscope. The passive diameter of arteries was determined with diltiazem (25 μM, Fig.1 A), which dilates rat cerebral arteries similarly to Ca2+-free bath conditions (24).

Fig. 1.

Intravascular pressure elevates Ca2+ spark frequency, Ca2+ wave frequency, and global intracellular Ca2+ concentration ([Ca2+]i) in smooth muscle cells and constricts cerebral arteries. A: transmitted light images of the same rat cerebral artery pressurized to 10 mmHg, to 60 mmHg, and to 60 mmHg after application of diltiazem (dilt; 25 μM). Mean diameters of 6 cerebral arteries were: 10 mmHg, 128 ± 4 μm; 10 mmHg + 25 μM diltiazem, 136 ± 8 μm; 60 mmHg, 123 ± 12 μm, 60 mmHg + 25 μM diltiazem, 174 ± 15 μm. B: average fluorescence (100 of 600 images) over 10 s of 2 different 56.2 μm × 52.8 μm areas of the same cerebral artery illustrated in A pressurized to 10 and 60 mmHg. The locations of Ca2+ sparks that occurred during 10 s are indicated by white boxes (1.54 μm × 1.54 μm). Black boxes (2.2 μm × 2.2 μm) are located in cells in which propagating Ca2+ waves occurred and in a cell in which a Ca2+ wave did not occur (box 2). Elevating pressure from 10 to 60 mmHg increased average global [Ca2+]i fluorescence 1.34-fold in smooth muscle cells of this artery. C: changes in Ca2+fluorescence for corresponding boxes represented in B. At 10 mmHg, 16 Ca2+ sparks and 1 Ca2+ wave (box 1) occurred over 10 s in this acquisition area. Representative Ca2+ sparks (boxes a–d), the Ca2+ wave (box 1), and a smooth muscle cell in which no Ca2+ wave occurred (box 2) are illustrated. At 60 mmHg, 34 Ca2+ sparks and 5 Ca2+ waves (boxes 3–6) occurred over 10 s. Representative Ca2+ sparks in smooth muscle cells at 60 mmHg are illustrated (boxes e–h).D: time course of the Ca2+ spark located atbox e in B, i.e., at 60 mmHg. The colored bar illustrates the F/F0 scale of the images.

Ca2+ spark, Ca2+ wave, and global change in fluorescence analysis.

Ca2+ sparks were detected in smooth muscle cells using custom analysis software written using IDL 5.0.2 (Research Systems, Boulder, CO), kindly provided by Drs. M. T. Nelson and A. D. Bonev (University of Vermont). Detection of Ca2+ sparks was performed by dividing an area 1.54 μm (7 pixels) × 1.54 μm (7 pixels) (i.e., 2.37 μm2) in each image (F) by a baseline (F0), which was determined by averaging six images without Ca2+ spark activity. Ca2+ spark amplitude was calculated as F/F0. Ca2+ waves were detected by placing 2.2 × 2.2-μm (10 × 10 pixels) boxes in individual smooth muscle cells and refer to a change in F/F0 of >1.2 that propagated for at least 20 μm. In each artery, Ca2+ spark and Ca2+ wave frequency (in Hz) was calculated from at least two different 56.3 × 52.8-μm areas of the arterial wall, each scanned for 10 s. Ca2+ sparks and Ca2+ waves were measured in the same cells, and mean data for each condition was the mean ± SE of the mean of frequency and amplitude values. Global Ca2+fluorescence was calculated from the same images used for Ca2+ spark and Ca2+ wave analysis and was the mean pixel value of 100 different images acquired over 10 s. Changes in global [Ca2+]i were calculated using methods described previously (7, 10, 22).

Isolation of arterial smooth muscle cells and patch-clamp electrophysiology.

Individual smooth muscle cells were enzymatically dissociated from cerebral arteries using an isolation solution of the following composition (in mM): 55 NaCl, 80 sodium glutamate, 5.6 KCl, 2 MgCl2, 10 HEPES, and 10 glucose (pH 7.3 with NaOH). Briefly, cerebral arteries were placed into isolation solution containing 0.3 mg/ml papain, 1 mg/ml dithioerythreitol, and 1 mg/ml bovine serum albumin (BSA) for 18 min (at 37°C) and were immediately transferred to isolation solution containing 1 mg/ml collagenase, 100 μM CaCl2 and 1 mg/ml BSA for 9 min (at 37°C). Arteries were subsequently washed in ice-cold isolation solution for 10 min and triturated using a polished glass pasteur pipette to yield single smooth muscle cells. Cells were allowed to adhere to a glass coverslip at the bottom of a chamber for 10 min before experimentation.

K+ currents were measured using the whole cell, perforated-patch configuration (18) of the patch-clamp technique (14), using an Axopatch 200B amplifier and pCLAMP 8 (Axon Instruments, Foster City CA). Bath solution contained (in mM) 134 NaCl, 6 KCl, 2 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose (pH 7.4 with NaOH). The pipette solution contained (in mM) 110 potassium aspartate, 30 KCl, 10 NaCl, 1 MgCl2, 10 HEPES, and 0.05 EGTA (pH 7.2 with KOH). Membrane currents were recorded (sample rate 2 kHz; filtered at 500 Hz) at steady holding voltages between −50 and −10 mV in 10-mV increments. STOC analysis was performed off-line using a custom analysis program provided by Drs. M. T. Nelson and A. D. Bonev (University of Vermont). The threshold of detection for STOCs was set at 2.5 times the single KCa channel amplitude. In the presence of ryanodine or thapsigargin, the simultaneous opening of three single KCa channels was not observed at −40 mV (4, 32, 37). Frequency and amplitude analysis of STOCs was not performed until 60 s after each voltage was applied to avoid transient effects of voltage on STOC activity, and in each cell at least 120 s of data were analyzed at each voltage. Data are from cells that were clamped at all voltages between −50 and −10 mV.

Statistical analysis.

Values are expressed as means ± SE. Paired and unpaired Student's t-tests were performed where appropriate, withP < 0.05 considered significant.

Chemicals.

Unless stated otherwise, all chemicals used in this study were obtained from Sigma Chemical (St. Louis, MO). Papain was purchased from Worthington Biochemical (Lakewood, NJ), ryanodine from LC Laboratories (Woburn, MA), and fluo 3-AM from Molecular Probes (Eugene, OR).

RESULTS

Elevation of intravascular pressure increases Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in cerebral artery smooth muscle cells.

In small cerebral arteries, an elevation of intravascular pressure from 10 to 60 mmHg induces a membrane potential depolarization from approximately −60 to −40 mV, an increase in arterial wall [Ca2+]i from ∼100 to 200 nM, and a maintained constriction (1, 5, 16, 24, 28). To investigate the effect of elevating intravascular pressure on intracellular Ca2+ signals, Ca2+ sparks, Ca2+waves, and global [Ca2+]i were measured at 37°C in the smooth muscle cells of the same cerebral arteries pressurized to 10 and 60 mmHg, with the use of a laser scanning confocal microscope and the fluorescent Ca2+ indicator fluo 3.

At 10 mmHg, mean Ca2+ spark frequency in the smooth muscle cells of six cerebral arteries was 1.28 ± 0.11 Hz, and average Ca2+ wave frequency was 0.15 ± 0.03 Hz (Figs. 1 and2). Elevating intravascular pressure from 10 to 60 mmHg caused the same six cerebral arteries to constrict on average to ∼72% of passive diameter (Fig. 1 A). Elevating pressure to 60 mmHg also significantly increased mean Ca2+spark frequency in arterial smooth muscle cells 2.6-fold and increased mean Ca2+ wave frequency 1.9-fold (Figs. 1 and 2). The mean F/F0 and the mean half time of decay of Ca2+sparks were not significantly different at 10 and 60 mmHg. Similarly, mean Ca2+ wave velocity was not significantly different at 10 and 60 mmHg. Ca2+ waves did not spread into adjacent cells and did not appear to be synchronized at either 10 or 60 mmHg.

Fig. 2.

Average effects of elevating intravascular pressure from 10 to 60 mmHg on Ca2+ spark frequency and Ca2+wave frequency in cerebral artery smooth muscle cells. Elevating intravascular pressure from 10 to 60 mmHg increased the mean frequency of Ca2+ sparks in smooth muscle cells from 1.28 ± 0.11 to 3.32 ± 0.29 Hz, or 2.6-fold (n = 6 arteries). In the same smooth muscle cells, elevating pressure to 60 mmHg increased mean Ca2+ wave frequency from 0.15 ± 0.03 to 0.29 ± 0.02 Hz, or 1.9-fold. The F/F0 and the mean half-time of decay (t ½) of Ca2+ sparks were not significantly different at 10 mmHg (1.53 ± 0.03 and 47.6 ± 4.5 ms, n = 148 sparks) compared with 60 mmHg (1. 48 ± 0.02 and 58.5 ± 6.4 ms, n = 394 sparks, P > 0.05). Mean Ca2+ wave velocity was not significantly different at 10 mmHg (33 ± 4 μm/s, range = 11–60 μm/s,n = 11) and 60 mmHg (47 ± 7 μm/s, range = 7–121 μm/s, n = 19, P > 0.05).

Elevating pressure to 60 mmHg also significantly increased the mean global Ca2+ fluorescence of the same smooth muscle cells 1.42 ± 0.13-fold (P < 0.05). Assuming a [Ca2+]i of 119 nM at 10 mmHg (24), elevating pressure increased the global [Ca2+]i of smooth muscle cells to ∼193 nM, which is similar to the mean arterial wall [Ca2+]i of cerebral arteries pressurized to 60 mmHg, when measured using fura 2 (24). These results demonstrate that elevating intravascular pressure from 10 to 60 mmHg increases Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in cerebral artery smooth muscle cells.

Ryanodine and thapsigargin inhibit Ca2+ sparks and Ca2+ waves, elevate global [Ca2+]i, and decrease diameter in pressurized myogenic cerebral arteries.

In arterial smooth muscle cells, Ca2+ sparks and Ca2+ waves are abolished by ryanodine (≥10 μM), a blocker of RyR channels, and by thapsigargin or cyclopiazonic acid, blockers of the SR Ca2+-ATPase that deplete the SR of Ca2+ (19-21, 30). Ryanodine and thapsigargin also depolarize the membrane potential of pressurized (60 mmHg) cerebral arteries by ∼10 mV, elevate arterial wall [Ca2+]i by ∼50 nM, and constrict arteries by ∼25–30% (25, 32). To investigate whether Ca2+ sparks and Ca2+ waves in smooth muscle cells of pressurized arteries occur due to Ca2+ release through RyR channels on the SR, and to explore the regulation of arterial diameter by intracellular Ca2+ signaling events, ryanodine (10 μM) or thapsigargin (100 nM) was applied to pressurized (60 mmHg) cerebral arteries.

Ryanodine (10 μM, 15-min exposure) abolished Ca2+ sparks and Ca2+ waves in smooth muscle cells of cerebral arteries pressurized to 60 mmHg. Before exposure to ryanodine, mean Ca2+ spark frequency was 3.67 ± 0.57 Hz and mean Ca2+ wave frequency was 0.26 ± 0.03 Hz (n = 4 arteries). In the same cells, ryanodine increased mean global Ca2+ fluorescence 1.20 ± 0.03-fold (i.e., from ∼193 to ∼256 nM) and decreased mean arterial diameter from 122 ± 6 to 98 ± 12 μm or from 75% to 61% of passive diameter (161 ± 15 μm, determined with diltiazem, 25 μM).

Thapsigargin (100 nM) also abolished Ca2+ sparks and Ca2+ waves in the smooth muscle cells of pressurized (60 mmHg) arteries. Before thapsigargin exposure, mean Ca2+spark frequency was 3.3 ± 0.5 Hz and mean Ca2+ wave frequency was 0.27 ± 0.04 Hz (n = 4 arteries). Similar to the effects of ryanodine, thapsigargin increased mean global Ca2+ fluorescence 1.26 ± 0.08-fold (i.e., from ∼193 to ∼278 nM) and decreased mean arterial diameter from 72% to 62% of passive diameter. These results suggest that, in the smooth muscle cells of pressurized cerebral arteries, Ca2+ sparks and Ca2+ waves occur due to Ca2+ release through RyR channels on the SR and indicate that inhibition of Ca2+sparks and Ca2+ waves elevates global [Ca2+]i and induces vasoconstriction.

Diltiazem decreases Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in smooth muscle cells of pressurized cerebral arteries.

Pressure could conceivably elevate Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in smooth muscle cells by inducing a membrane depolarization that would activate voltage-dependent Ca2+ channels (see e.g., Ref. 22). However, although intravascular pressure induces membrane depolarization, pressure may also activate other signal transduction pathways that could positively or negatively regulate Ca2+ signaling in smooth muscle cells (for reviews, see Refs. 13 and 21). For example, inositol phosphates increase in smooth muscle in response to pressure or stretch (31, 41), suggesting that IP3-gated Ca2+ release channels could underlie pressure-induced Ca2+ signaling.

To investigate whether pressure elevated Ca2+ signaling via activation of voltage-dependent Ca2+ channels, Ca2+ sparks, Ca2+ waves, and global [Ca2+]i were measured in the smooth muscle cells of pressurized (60 mmHg) cerebral arteries, before and after addition of diltiazem (25 μM), a selective voltage-dependent Ca2+ channel blocker.

Diltiazem increased the mean diameter of eight pressurized cerebral arteries from 123 to 171 μm (i.e., 72% tone in the absence of diltiazem, see Fig. 1 A). Diltiazem also significantly reduced mean Ca2+ spark and Ca2+ wave frequency in smooth muscle cells of the same cerebral arteries (Fig.3). Mean F/F0 of Ca2+ sparks was not significantly different in diltiazem, when compared with control. Diltiazem also decreased mean global Ca2+ fluorescence (F/F0) to 0.58 ± 0.07 of control, or from 193 to ∼93 nM (Fig. 3). These results suggest that pressure elevates Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in cerebral artery smooth muscle cells via activation of voltage-dependent Ca2+ channels.

Fig. 3.

Diltiazem reduced Ca2+ spark frequency and Ca2+ wave frequency in the smooth muscle cells of cerebral arteries pressurized to 60 mmHg. Diltiazem (25 μM) increased the mean diameter of pressurized arteries from 123 ± 9 to 171 ± 12 μm (i.e., 73% tone in the absence of diltiazem, n = 8). Diltiazem reduced the mean frequency of Ca2+ sparks in smooth muscle cells of the same pressurized (60 mmHg) cerebral arteries (n = 8) from 3.34 ± 0.35 to 0.13 ± 0.03 Hz and reduced mean Ca2+ wave frequency in the same smooth muscle cells from 0.30 ± 0.02 to 0.04 ± 0.02 Hz. Mean F/F0 of Ca2+ sparks was unaltered by diltiazem (60 mmHg, 1.48 ± 0.03, n = 384 sparks; 60 mmHg + 25 μM diltiazem, 1.53 ± 0.11, n = 15 sparks).

Diltiazem reverses depolarization-induced elevation of Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in cerebral artery smooth muscle cells.

To investigate whether pressure elevated Ca2+ sparks, Ca2+ waves, and global [Ca2+]i in smooth muscle cells via depolarization-induced activation of voltage-dependent Ca2+ channels, intracellular [Ca2+]i was measured in the smooth muscle cells of the same cerebral arteries at low pressure (10 mmHg), before and after elevation of bath K+ from 6 to 30 mM, which depolarizes arteries from approximately −60 to approximately −40 mV (3, 15, 17). This depolarization is similar to the effect of elevating intravascular pressure from 10 to 60 mmHg (16,24).

Elevation of bath K+ from 6 to 30 mM increased mean Ca2+ spark frequency in smooth muscle cells 2.9-fold (n = 6 arteries, Fig.4 A). Mean Ca2+spark amplitude (F/F0) was not significantly different in 6 mM K+, when compared with 30 mM K+. Elevating K+ also increased mean Ca2+ wave frequency in the same smooth muscle cells 2.4-fold (Fig. 4 A) and elevated global Ca2+ fluorescence 1.35 ± 0.1-fold, indicating that [Ca2+]i increased from 119 nM (24) to ∼179 nM. These results indicate that a membrane depolarization from approximately −60 to approximately −40 mV, or a pressure elevation from 10 to 60 mmHg, induces a similar increase of Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in the smooth muscle cells of cerebral arteries.

Fig. 4.

Depolarization-induced elevation of Ca2+spark frequency and Ca2+ wave frequency in cerebral artery smooth muscle cells is reversed by diltiazem. A: in arteries at low pressure (10 mmHg), an elevation of bath K+ from 6 to 30 mM significantly increased mean Ca2+ spark frequency in smooth muscle cells from 1.33 ± 0.1 to 3.8 ± 0.48 Hz, or 2.9-fold, and increased mean Ca2+ wave frequency in the same cells from 0.14 ± 0.02 to 0.33 ± 0.02 Hz, or 2.4-fold (P < 0.05 for each, n = 6 arteries). Mean Ca2+ spark amplitude (F/F0) was not significantly different in 6 mM K+ (1.49 ± 0.02,n = 128 sparks) compared with 30 mM K+(1.52 ± 0.02, n = 319 sparks; P> 0.05). B: diltiazem (25 μM) significantly reduced mean Ca2+ spark frequency in the smooth muscle cells of depolarized (30 mM K+) arteries from 3.9 ± 0.53 to 0.22 ± 0.18 Hz and decreased the mean frequency of Ca2+ waves in the same cells from 0.33 ± 0.02 to 0.03 ± 0.02 Hz (P < 0.05 for each,n = 5 arteries). Diltiazem did not significantly change mean Ca2+ spark F/F0 (30 K+, 1.52 ± 0.02, n = 314 sparks; 30 K+ + 25 μM diltiazem, 1.62 ± 0.19,n = 13 sparks, P > 0.05).

To investigate whether membrane depolarization elevated Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in smooth muscle cells via activation of voltage-dependent Ca2+ channels, intracellular Ca2+ signaling modalities were measured in depolarized (30 mM K+) arteries pressurized to 10 mmHg before and after application of diltiazem (25 μM). Diltiazem significantly reduced mean Ca2+ spark and Ca2+wave frequency in the smooth muscle cells of depolarized (30 mM K+) arteries (n = 5 arteries, Fig.4 B), although diltiazem did not significantly change mean Ca2+ spark F/F0. Diltiazem also significantly reduced mean global Ca2+ fluorescence (F/F0, 0.63 ± 0.04) from ∼179 to ∼97 nM. These results suggest that diltiazem similarly decreased depolarization- and pressure-induced elevations of Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i (Figs. 3 and4 B). Taken together, data suggest that pressure elevates Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in cerebral artery smooth muscle cells by inducing a steady membrane depolarization that activates voltage-dependent Ca2+ channels.

Steady membrane depolarization elevates STOC frequency and amplitude in cerebral artery smooth muscle cells.

Elevating intravascular pressure activates KCa channels in arterial smooth muscle cells, inducing a negative-feedback pathway that limits arterial constriction (5). However, the pressure-induced signaling pathways that activate KCachannels are unclear. The present study suggests that an elevation of intravascular pressure induces a steady membrane depolarization that increases Ca2+ spark frequency in cerebral artery smooth muscle cells. Virtually all Ca2+ sparks activate STOCs in cerebral artery smooth muscle cells (>95%, see Ref. 34), suggesting that pressure may elevate KCa channel activity by increasing Ca2+ spark and thus STOC frequency. Depolarization is known to activate STOCs in smooth muscle cells (2, 36, 40), but the steady-state voltage dependence of STOC frequency and amplitude has not been quantified. To determine the effect of steady membrane depolarization on STOCs, K+currents were recorded in isolated cerebral artery smooth muscle cells at holding potentials between −50 and −10 mV, using the patch-clamp technique. To ensure that the effects of voltage were studied under steady-state conditions, STOCs were not analyzed until 60 s after each voltage was applied to avoid transient effects of depolarization, and for each cell, at least 120 s of data were analyzed at each holding potential.

Steady membrane depolarization increased STOC frequency and amplitude (Fig. 5). The relationship between voltage and STOC frequency was fit with a Boltzmann function (Fig.5 B), indicating that STOCs were 50% activated at −35 mV (n = 8 cells). The steepness factor, which provides an indication of the sensitivity of STOCs to voltage, demonstrated that STOC frequency increased e-fold for 7.4 mV depolarization. These data demonstrate that steady membrane depolarization elevates STOC frequency and amplitude in cerebral artery smooth muscle cells and support data obtained in pressurized arteries, suggesting that intravascular pressure elevates Ca2+ spark frequency by inducing steady membrane depolarization.

Fig. 5.

Steady membrane depolarization increases spontaneous transient outward current (STOC) frequency and amplitude in isolated voltage-clamped cerebral artery smooth muscle cells. A: K+ currents measured at steady voltages of −40 and −20 mV in the same smooth muscle cell, illustrating the effect of steady depolarization on STOC frequency and amplitude. Dotted line represents the baseline current on which STOCs occurred. B: STOC frequency increased with steady membrane depolarization. Mean STOC frequencies at each voltage were (in Hz) −50 mV, 0.27 ± 0.08; −40 mV, 0.61 ± 0.12; −30 mV, 1.01 ± 0.15; −20 mV, 1.39 ± 0.21; −10 mV, 1.44 ± 0.25 (n = 8 cells). The relationship between voltage and STOC frequency was fit with a Boltzmann function, indicating a V 0.5 of −35 mV and a steepness factor of 7.4 mV. C: STOC amplitude increased with depolarization. Mean STOC amplitudes were (in pA) −50 mV, 12.8 ± 1.7; −40 mV, 22.3 ± 3.8; −30 mV, 31.5 ± 5.6; −20 mV, 45.2 ± 7.5; −10 mV, 69.2 ± 11.7 (n = 8 cells).

Cd2+, a voltage-dependent Ca2+ channel blocker, abolishes the voltage dependence of STOC frequency in cerebral artery smooth muscle cells.

In arterial smooth muscle cells, Ca2+ entry through voltage-dependent Ca2+ channels regulates Ca2+spark (21) and STOC frequency (4, 20, 40). If steady membrane depolarization elevates Ca2+ spark frequency due to the activation of voltage-dependent Ca2+channels, then inhibitors of these channels should block the voltage dependence of STOCs. To examine whether membrane depolarization increased STOC frequency and amplitude via activation of voltage-dependent Ca2+ channels, STOCs were recorded at steady membrane potentials of −40 and −20 mV, before and after application of Cd2+ (250 μM), a voltage-dependent Ca2+ channel blocker.

Steady membrane depolarization from −40 to −20 mV significantly increased mean STOC frequency in the same cerebral artery smooth muscle cells from ∼0.66 to ∼1.64 Hz and significantly elevated mean STOC amplitude from ∼24 to ∼50 pA, respectively (n = 5 cells, Fig. 6). In the same cells after application of Cd2+, membrane depolarization from −40 to −20 mV did not significantly elevate mean STOC frequency, but mean STOC amplitude increased in a manner similar to that observed before Cd2+ application (Fig. 6, A and B). These data suggest that steady membrane depolarization elevates STOC frequency in cerebral artery smooth muscle cells via activation of voltage-dependent Ca2+ channels, supporting the concept that pressure elevates Ca2+ spark frequency via steady membrane depolarization. These data also suggest that membrane depolarization elevates STOC amplitude independent of Ca2+entry through voltage-dependent Ca2+ channels.

Fig. 6.

Cd2+ (250 μM), a voltage-dependent Ca2+channel blocker, inhibits the voltage dependence of STOC frequency but does not affect the voltage dependence of STOC amplitude in cerebral artery smooth muscle cells. A: steady membrane depolarization (open bars) from −40 to −20 mV significantly elevated mean STOC frequency from 0.66 ± 0.16 to 1.64 ± 0.34 Hz, respectively (P < 0.05, n = 5 cells). In the same cells, Cd2+ (250 μM, shaded bars) blocked the depolarization-induced increase in STOC frequency (−40 mV, 0.34 ± 0.09 Hz; −20 mV, 0.49 ± 0.1 Hz, P > 0.05).B: steady depolarization (open bars) significantly elevated mean STOC amplitude from 23.7 ± 2.6 pA at −40 mV to 49.6 ± 5.7 pA at −20 mV (P < 0.05, n = 5 cells). Cd2+ (250 μM, shaded bars) did not significantly change mean STOC amplitude at −40 mV (25.4 ± 3.1 pA,P > 0.05) or −20 mV (52.6 ± 8.4 pA,P > 0.05).

DISCUSSION

This study demonstrates for the first time that pressure activates distinct intracellular Ca2+ signaling modalities in arterial smooth muscle cells. Data are consistent with the hypothesis that pressure induces a steady membrane potential depolarization that activates voltage-dependent Ca2+ channels, leading to an elevation of Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in cerebral artery smooth muscle cells. Data also suggest that pressure induces contraction via an elevation of global [Ca2+]i, whereas the net effect of Ca2+ sparks and Ca2+ waves, which occur due to Ca2+ release through RyR channels on the SR, is to oppose constriction.

Pressure elevates intracellular Ca2+signaling modalities in cerebral artery smooth muscle cells.

Elevation of intravascular pressure from 10 to 60 mmHg increased Ca2+ spark frequency in smooth muscle cells approximately threefold. The scanned volume corresponds to approximately three smooth muscle cells (see also Ref. 22), indicating that elevation of pressure increased Ca2+ spark frequency in a single cell from ∼0.4 to ∼1.1 Hz. Therefore, based on earlier calculations regarding the contribution of Ca2+ sparks to global [Ca2+]i (21, 32), at physiological levels of temperature and pressure Ca2+sparks do not significantly elevate global [Ca2+]i.

Elevating pressure to 60 mmHg also increased Ca2+ wave frequency approximately twofold. In agreement with others (30,39), Ca2+ waves were asynchronous and did not propagate between smooth muscle cells. The precise elevation of global [Ca2+]i that occurred due to a Ca2+ wave is unclear, because a nonratiometric imaging procedure is employed in this study, and basal [Ca2+]i in the absence of Ca2+waves is unknown. However, even in arteries pressurized to 60 mmHg, it is unlikely that smooth muscle Ca2+ waves contribute significantly to global [Ca2+]i, at least in the absence of receptor agonists that elevate IP3. This postulation is based on a number of observations. First, the frequency of Ca2+ waves is low even in arteries pressurized to 60 mmHg, and the mean velocity of Ca2+ waves (∼ 45 μm/s) indicates that the propagating wave front will be present in a cell for no more than 1.5 s. If a propagating Ca2+ wave front occupies approximately one-eighth of the volume of a cell (see also Ref. 20), is present in that cell for ∼15% of the time (1.5 s/10 s), and Ca2+ waves occur in only 30% of cells over 10 s, Ca2+ wave fronts of 100 nM [Ca2+]i would raise mean arterial wall [Ca2+]i by ∼0.5 nM. Second, pressure elevated cytosolic [Ca2+] in many cells that did not exhibit Ca2+ waves, presumably due to Ca2+influx through voltage-dependent Ca2+ channels. Thus global [Ca2+]i appears to occur primarily due to Ca2+ influx through voltage-dependent Ca2+channels. Third, ryanodine and thapsigargin blocked Ca2+sparks and Ca2+ waves and elevated global [Ca2+]i. Because Ca2+ sparks do not significantly elevate global [Ca2+]i, this suggests that the negative-feedback effect of Ca2+sparks and Ca2+ waves on Ca2+ entry (via membrane hyperpolarization) is greater than any contribution of Ca2+ waves to global [Ca2+]i. In addition, ryanodine does not alter the arterial wall [Ca2+]i of cerebral arteries pressurized to 60 mmHg when applied in the presence of iberiotoxin, a KCachannel blocker (25, 32). Thus, when Ca2+sparks and Ca2+ waves are blocked under conditions where membrane potential does not change, arterial wall [Ca2+]i is not altered. These arguments suggest that Ca2+ sparks and Ca2+ waves do not contribute to global [Ca2+]i in arteries pressurized to between 10 and 60 mmHg.

Elevating pressure to 60 mmHg increased global [Ca2+]i in smooth muscle cells to ∼193 nM, which is similar to the [Ca2+]i of these arteries at 60 mmHg when measured using fura 2 (24). Data are consistent with the idea that elevating pressure from 10 to 60 mmHg increases arterial wall [Ca2+]i via an elevation of global [Ca2+]i in cerebral artery smooth muscle cells.

Pressure elevates intracellular Ca2+signals via activation of voltage-dependent Ca2+ channels.

Pressure or depolarization induced a similar elevation of Ca2+ spark frequency, Ca2+ wave frequency, and global Ca2+, and diltiazem similarly reduced these Ca2+ signaling modalities in pressurized and depolarized arteries. In addition, STOC frequency increased e-fold for ∼7 mV depolarization, which is similar to the voltage dependence of Ca2+ currents in arterial smooth muscle cells (33,38), and Cd2+ blocked the voltage dependence of STOC frequency. These data suggest that pressure elevates Ca2+sparks, Ca2+ waves, and global [Ca2+]i in arterial smooth muscle cells via a steady membrane depolarization that activates voltage-dependent Ca2+ channels.

Global [Ca2+]i was similarly elevated by pressure and depolarization, was reduced by diltiazem, and was elevated by ryanodine and thapsigargin, agents that abolished Ca2+sparks and Ca2+ waves and depolarized pressurized cerebral arteries by ∼10 mV (25, 32). These findings suggest that pressure elevates global [Ca2+]iprimarily due to Ca2+ entry through voltage-dependent Ca2+ channels.

In nonpressurized arteries and isolated smooth muscle cells, Ca2+ sparks occur due to the activation of RyR channels (21). However, in smooth muscle, inositol phosphates increase in response to pressure or stretch (31, 41), suggesting that pressure could elevate Ca2+ sparks and Ca2+ waves via IP3-gated Ca2+release. Several lines of evidence from this study do not support a significant role for IP3-mediated Ca2+ release in the smooth muscle cells of arteries pressurized to between 10 and 60 mmHg, at least in the absence of receptor agonists that elevate IP3. First, pressure and depolarization induced a similar elevation in intracellular Ca2+ signals. Second, diltiazem similarly reduced Ca2+ spark and Ca2+ wave frequency in pressurized and depolarized arteries. Third, caffeine, an activator of RyR channels (35), elevates Ca2+wave frequency in venous smooth muscle cells (39), suggesting that RyR channels can generate Ca2+ waves in the absence of an elevation in IP3. Thus the present study suggests that Ca2+ sparks and Ca2+ waves occur in smooth muscle cells of pressurized cerebral arteries due to the activation of RyR channels. This conclusion is further supported by the finding that ryanodine abolished Ca2+ sparks and Ca2+ waves in pressurized arteries, and by evidence demonstrating that RyR channels are activated by elevations in cytosolic [Ca2+] and SR [Ca2+] (21).

Activation of voltage-dependent Ca2+ channels in smooth muscle cells could conceivably increase Ca2+ spark frequency and Ca2+ wave frequency via a number of mechanisms, including an increase in cytosolic [Ca2+]i, an elevation of SR [Ca2+], or by enhancement of “local control.” In cardiac myocytes, Ca2+ entering the cell through voltage-dependent Ca2+ channels produces a subsarcolemmal elevation of [Ca2+]i that activates a number of nearby RyR channels on the SR, leading to a Ca2+ spark (local control) (8, 27). In arterial smooth muscle, the mechanism of communication between voltage-dependent Ca2+channels and RyR channels is less clear, but local Ca2+entering through a single caveolemmal voltage-dependent Ca2+ channel has been proposed to activate several RyR channels to generate a Ca2+ spark (26). In contrast, Ca2+ entry through voltage-dependent Ca2+ channels is not tightly coupled to RyR channels in bladder smooth muscle cells (12). In skeletal muscle, physical coupling of voltage-dependent Ca2+ channels and RyR channels can activate Ca2+ release in the absence of Ca2+ entry (23). Although the mechanisms linking Ca2+ entry to Ca2+ release are not fully established in smooth muscle cells, voltage-dependent Ca2+ channel blockers inhibited the pressure- and depolarization-induced frequency elevation of Ca2+ sparks, STOCs, and waves. Thus, in arterial smooth muscle, it is unlikely that voltage-dependent Ca2+ channels activate RyR channels via physical coupling of these proteins.

Physiological relevance of pressure-induced Ca2+ signaling.

When global [Ca2+]i increased, arteries constricted, and when global [Ca2+]idecreased, arteries dilated. Thus this study suggests that pressure-induced elevations in global [Ca2+]ilead to the development of myogenic tone in cerebral arteries, presumably via the activation of Ca2+/calmodulin-dependent myosin light chain kinase. The physiological function of Ca2+ waves is less clear, primarily because the amplitude of Ca2+ waves is uncertain. Nevertheless, in arteries pressurized to between 10 and 60 mmHg, Ca2+ waves do not appear to significantly contribute to arterial wall [Ca2+]i.

Elevating intravascular pressure activates KCa channels in arterial smooth muscle cells, inducing a negative-feedback pathway that limits arterial constriction (5). Data from this study indicate that elevating pressure from 10 to 60 mmHg increases Ca2+ spark and STOC frequency 3- to 4-fold and STOC amplitude 10- to 20-fold. Thus the present study supports the concept that pressure elevates KCa current primarily by elevating STOC frequency and amplitude. This finding is supported by recent evidence suggesting that arterial smooth muscle KCachannels are relatively insensitive to global levels of cytosolic [Ca2+] (21) but are activated by the high, localized elevation of [Ca2+]i produced by a Ca2+ spark (6, 21, 32, 34). Depolarization elevated STOC frequency due to the activation of voltage-dependent Ca2+ channels, whereas STOC amplitude increased independently. STOC frequency will be dependent on Ca2+spark frequency, whereas depolarization could conceivably elevate STOC amplitude via an increase in the Ca2+ sensitivity of KCa channels (9), an increase in Ca2+ spark amplitude (22), and an increase in the driving force for K+. When averaged throughout the arterial wall, the pressure-induced increase in KCa channel activity translates to a tonic hyperpolarization of ∼10 mV at 60 mmHg, which reduces the activity of voltage-dependent Ca2+channels, decreases the [Ca2+]i by ∼50 nM, and dilates (25, 32).

In summary, data in this study are consistent with the hypothesis that pressure induces a steady membrane depolarization that activates voltage-dependent Ca2+ channels, leading to an elevation of Ca2+ spark frequency, Ca2+ wave frequency, and global [Ca2+]i in cerebral artery smooth muscle cells. Pressure elevates global [Ca2+]i primarily due to an increase in Ca2+ influx through voltage-dependent Ca2+channels, and this leads to contraction; whereas Ca2+sparks and Ca2+ waves occur due to the activation of RyR channels on the SR, these events do not significantly contribute to global [Ca2+]i, and the net effect of sparks and waves is to oppose contraction.

Acknowledgments

I thank Dr. C. W. Leffler for helpful comments on the manuscript.

Footnotes

  • This study was supported by grants from the American Heart Association National Center and Southeast Affiliate.

  • Address for reprint requests and other correspondence: J. H. Jaggar, Dept. of Physiology, College of Medicine, Univ. of Tennessee Health Science Center, 894 Union Ave., Memphis, TN 38163 (E-mail:jjaggar{at}physio1.utmem.edu).

  • The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

REFERENCES

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