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This Article Abstract Full Text (PDF) Supplemental data All Versions of this Article: 584/1/205    most recent jphysiol.2007.138982v2 jphysiol.2007.138982v1 Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Essin, K. Articles by Moosmang, S. Search for Related Content PubMed PubMed Citation Articles by Essin, K. Articles by Moosmang, S. Related Collections Cardiovascular Related Article

¹ Department of Nephrology and Medical Intensive Care, Charité Campus Virchow-Klinikum, Berlin, Germany
² Franz Volhard Clinic, HELIOS Klinikum Berlin, Charité Campus Berlin-Buch, and Max Delbrück Center for Molecular Medicine, Berlin, Germany
³ Department of Pharmacology and Toxicology, Technical University München, Munich, Germany

	Abstract

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In arterial vascular smooth muscle cells (VSMCs), Ca²⁺ sparks stimulate nearby Ca²⁺-activated K⁺ (BK) channels that hyperpolarize the membrane and close L-type Ca²⁺ channels. We tested the contribution of L-type Ca_v1.2 channels to Ca²⁺ spark regulation in tibial and cerebral artery VSMCs using VSMC-specific Ca_v1.2 channel gene disruption in (SMAKO) mice and an approach based on Poisson statistical analysis of activation frequency and first latency of elementary events. Ca_v1.2 channel gene inactivation reduced Ca²⁺ spark frequency and amplitude by 50% and 80%, respectively. These effects were associated with lower global cytosolic Ca²⁺ levels and reduced sarcoplasmic reticulum (SR) Ca²⁺ load. Elevating cytosolic Ca²⁺ levels reversed the effects completely. The activation frequency and first latency of elementary events in both wild-type and SMAKO VSMCs weakly reflected the voltage dependency of L-type channels. This study provides evidence that local and tight coupling between the Ca_v1.2 channels and ryanodine receptors (RyRs) is not required to initiate Ca²⁺ sparks. Instead, Ca_v1.2 channels contribute to global cytosolic [Ca²⁺], which in turn influences luminal SR calcium and thus Ca²⁺ sparks.

(Received 19 June 2007; accepted after revision 27 July 2007; first published online 2 August 2007)
Corresponding author M. Gollasch: Charité Campus Virchow Klinikum, Humboldt University, Augustenburger Platz 1, 13353 Berlin, Germany. Email: maik.gollasch{at}charite.de

	Introduction

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Ca²⁺ sparks are optical images of elementary Ca²⁺ release from a single Ca²⁺ release unit (CRU) composed of a group of ryanodine receptors (RyRs) in the sarcoplasmic reticulum (SR) (Wang et al. 2004). In arterial vascular smooth muscle cells (VSMCs), membrane depolarizations produce moderate (100–300 nM) increases in the global [Ca²⁺]_i that produce contraction (Nelson et al. 1990). These contractions depend on Ca²⁺ ion influx through Ca_v1.2 Ca²⁺ channels (Moosmang et al. 2003). Ca²⁺ spark inhibition paradoxically produces vasoconstriction for two reasons (Nelson et al. 1995; Knot et al. 1998). First, a single spark is capable of producing a very high (10–100 µM) local (1% of the cell volume) increase in [Ca²⁺]_i (Perez et al. 1999, 2001) while increasing the global [Ca²⁺]_i by only < 2 nM (Nelson et al. 1995; Jaggar et al. 2000). Second, Ca²⁺ sparks occur in close proximity to the cell membrane, where every Ca²⁺ spark activates numerous BK channels (Perez et al. 1999; Brenner et al. 2000; Pluger et al. 2000; Sausbier et al. 2005). The resultant ‘spontaneous transient outward currents’ (STOCs) cause hyperpolarization of the cell membrane, thereby shutting off tonic Ca²⁺ entry through Ca_v1.2 channels. Therefore, the net result of the spark–STOC coupling is decreased global [Ca²⁺]_i and vasorelaxation (Nelson et al. 1995; Gollasch et al. 1998).

Arterial VSMCs do not exhibit action potentials in vivo to elevate global [Ca²⁺]_i. Instead, steady-state membrane VSMC depolarization increases voltage-dependent open probability of L-type channels, global [Ca²⁺]_i, and presumably SR [Ca²⁺] (Jaggar et al. 2000). Ca²⁺ spark frequency at steady-state potentials is modulated by Ca²⁺ influx through L-type Ca²⁺ channels (Nelson et al. 1995; Jaggar et al. 1998; Remillard et al. 2002; Cheng & Jaggar, 2006). However, whether or not local Ca²⁺ influx regulates the probability of these rather ‘spontaneous’ sparks in arterial VSMCs is unresolved. An elevation in local Ca²⁺ from opening of L-type Ca²⁺ channels, elevated global [Ca²⁺]_i, or increased luminal SR [Ca²⁺] could each contribute to the observed steady-state depolarization-induced increase in Ca²⁺ spark frequency and amplitude.

Recently, Santana et al. obtained images of brief Ca²⁺ fluxing through single L-type Ca²⁺ channels in arterial VSMCs (Navedo et al. 2005). Analogous to cardiomyocytes (Wang et al. 2001), these events were named ‘Ca²⁺ sparklets’. The authors also detected single or small clusters of L-type channels in VSMCs that operate in a high activity mode, creating sites of nearly continual Ca²⁺ influx (‘persistent Ca²⁺ sparklet sites’) even at hyperpolarized membrane potentials of 70 mV (Navedo et al. 2005, 2006). Sparklets could directly activate RyR to generate Ca²⁺ sparks in VSMCs (Navedo et al. 2005). However, a rather loose coupling between Ca²⁺ entry and spark frequency has been deduced from urinary bladder experiments (Collier et al. 2000; Kotlikoff, 2003). In contrast to arterial VSMCs, bladder cells typically produce contraction by generating action potentials to induce calcium-induced calcium release through RyRs (Imaizumi et al. 1998). However, since arterial smooth muscle produces only small, steady-state membrane depolarizations of a few millivolts to produce sustained calcium influx to elevate global [Ca²⁺]_i, the coupling mechanism between Ca_v1.2 and RyRs in arterial smooth muscle is not clear.

Ca²⁺ can activate RyRs on the SR luminal side of the receptor (Ching et al. 2000). Recent evidence indicates that the frequency and amplitude of Ca²⁺ sparks depends steeply on the SR Ca²⁺ load in stomach muscle (ZhuGe et al. 1999). Experiments using phospholamban-deficient cerebral artery VSMCs supported this notion (Wellman et al. 2001). Spark-like Ca²⁺ release has been observed in aortic and cerebral VSMCs after chemical permeabilization of the cell membrane (Rueda & Valdivia, 2006) supporting the idea that spark initiation does not depend on a close L-type Ca²⁺ channel and RyR proximity. We tested the hypothesis that Ca_v1.2 channels contribute to global cytosolic calcium, which in turn influences luminal SR calcium and thus Ca²⁺ sparks in arterial VSMCs. We used VSMC-specific Ca_v1.2 channel gene inactivation in mice (SMAKO) (Moosmang et al. 2003). We provide evidence that rapid, local and tight coupling between the Ca_v1.2 channels and RyRs is not required to initiate Ca²⁺ sparks. We found that cytosolic [Ca²⁺]_i itself contributes minimally to the acute triggering of physiologically relevant proportion of Ca²⁺ sparks. Instead the most eficacious Ca²⁺ spark trigger appears to be the luminal SR Ca²⁺, which is slowly loaded via Ca²⁺ influx through Ca_v1.2 channels.

	Methods

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All animal experimental protocols were approved by the local animal care committees (Regierung von Oberbayern, Munich, Germany and LaGetSi, Berlin, Germany). The generation of mice deficient in the smooth muscle Ca_v1.2 Ca²⁺ channel (SMAKO, smooth muscle 1c-subunit Ca²⁺ channel knockout) has been described (Moosmang et al. 2003). Briefly, a conditional loxP-flanked allele (L2) of the Ca_v1.2 gene (i.e. exons 14 and 15) was generated by homologous recombination in R1 embryonic stem cells. In addition, mice carried a knock-in allele (SM-CreER T2 (ki)) (Kuhbandner et al. 2000), which expresses the tamoxifen-dependent Cre recombinase, CreER T2, from the endogenous SM22 gene locus, which is selectively expressed in smooth muscle of adult mice. Thus, tamoxifen treatment of mice results in conversion of the loxP-flanked Ca_v1.2 allele (L2) into a Cav1.2 knockout allele (L1) specifically in SMC. Animals were kept under standard conditions with water and food ad libitum. At an age of 2–3 months, SMAKO mice (Ca_v1.2 l1/L2, SM-CreER T2 (ki)+/.) and corresponding control (CTR) mice (Ca_v1.2+/L2, SM-CreER T2 (ki)+/.) were I.P. injected with tamoxifen (2 mg day^–1) for five consecutive days. After 3–4 weeks, mice were killed by cervical dislocation and the brain and tibial arteries were removed. Experiments were performed on the same day with arteries from litter-matched control and SMAKO mice.

Isolation of arterial VSMCs

SMCs from tibial and basilar arteries were isolated as described (Gollasch et al. 1998; Pluger et al. 2000). Briefly, the brain and tibial arteries were removed and quickly transferred to cold (4°C) oxygenated (95% O₂–5% CO₂) physiological salt solution (PSS) of the following composition (mM): 119 NaCl, 4.7 KCl, 25.0 NaHCO₃, 1.2 KH₂PO₄, 1.8 CaCl₂, 1.2 MgSO₄, 0.026 EDTA and 11.1 glucose. The arteries were cleaned, cut into pieces and placed in a Ca²⁺-free Hanks' solution (mM): 55 NaCl, 80 sodium glutamate, 5.6 KCl, 2 MgCl₂, 1 mg ml^–1 bovine serum albumin (BSA, Sigma), 10 glucose and 10 Hepes (pH 7.4 with NaOH) containing 1.0 mg ml^–1 papain (Sigma) and 1 mg ml^–1 DTT for 15 (cerebral arteries) to 45 min (tibial arteries) at 36°C. The segments were then placed in Hanks' solution containing 1 mg ml^–1 collagenase (Sigma, type F and H; ratio 30% and 70%) and 0.1 mM CaCl₂ for 6 min (cerebral arteries) to 10 min (tibial arteries) at 36°C. Following several washes in Ca²⁺-free Hanks' solution (containing 1 mg ml^–1 BSA), single cells were dispersed from artery segments by gentle trituration. Cells were then stored in the same solution at 4°C.

Ca²⁺ sparks

VSMCs were seeded onto glass coverslips and incubated with the Ca²⁺ indicator fluo-3-AM (5 µM) and pluronic acid (0.005%; w/v) for 30 min at room temperature in Ca²⁺-free Hanks' solution (Lohn et al. 2000; Pluger et al. 2000; Lohn et al. 2001a). After loading of the cells with fluo-3, the cells were washed with a Hepes-buffered physiological saline solution (Hepes-PSS) for 30–40 min at room temperature. The Hepes-PSS had the following composition (mM): 135 NaCl, 5.4 KCl, 1.8 CaCl₂, 1 MgCl₂, 10 Hepes and 10 glucose (pH 7.4 with NaOH). Single SMCs were imaged using a Bio-Rad (Munich, Germany) laser scanning confocal microscope attached to a Nikon Diaphot microscope (Furstenau et al. 2000; Lohn et al. 2001a). Images were obtained by illumination with a krypton–argon laser at 488 nm, and recording all emitted light above 500 nm. Ca²⁺ sparks were measured in Hepes-PSS. Cells were scanned in the ‘line scan’ mode for 10 s. Ca²⁺ spark analysis was performed off-line using custom software written in C++ by K. Essin. Ca²⁺ sparks were defined as local fractional fluorescence increases greater than 1.2. The site of a Ca²⁺ spark was determined as the centre of the spark at the time of its initiation. Ca²⁺ spark width was determined at 50% maximal amplitude; decay was measured from peak to half-maximal amplitude. The frequency was estimated as the number of detected sparks divided by the total scan time. The amplitudes were expressed as fractional fluorescence increase (F/F₀) or in absolute values relative to the global resting cytosolic [Ca²⁺] using the following equation: (Cheng et al. 1993; Jaggar et al. 1998; Herrera et al. 2001)

(1)

where R is the fractional fluorescence increase (F/F₀), [Ca²⁺]_r is the free resting cytosolic Ca²⁺ concentration, and K is the apparent affinity of fluo-3 for Ca²⁺ (400 nM) (Cheng et al. 1993). [Ca²⁺]_r was measured directly using Fura-2 in separate experiments (see below).

In the experiments to determine the first latency to occurrence of a Ca²⁺ spark after membrane depolarization, fast two-dimensional confocal microscopy was used in VSMCs clamped by the perforated whole-cell patch technique (see below). Cells were loaded with fluo-4-AM (5 µM) and pluronic acid (0.005%; w/v) for 30 min at room temperature in Ca²⁺-free solution (mM: 10 Hepes, 55 NaCl, 5.6 KCl, 80 sodium glutamate, 2 MgCl₂, and 10 D-glucose; pH to 7.4 with NaOH) and washed for 30 min with Ca²⁺containing solution (for external solution for STOC recording, see below). Cells were clamped at –40 mV and depolarized according to the protocol presented in the online Supplemental Fig. 2, similar to that in Lopez-Lopez et al. (1995). Synchronicity of the command voltage onset and fluorescence measurment was achieved by means of light-emitting diode placed near the recording chamber and switched on and off from a D/A output of CED 1401 interface before and after the acqusition protocol. Images were taken at 25–50 frames s^–1 on a PerkinElmer, Nipkow disc-based UltraView LCI confocal scanner linked to a fast digital camera. The confocal system was mounted in an inverted Diaphot microscope with a x40 oil-immersion objective (NA 1.3; Nikon). Fluo-4 was excited by the 488 nm line of an argon ion laser and emitted fluorescence was collected at wavelengths > 515 nm. Image analysis was done using PerkinElmer's ImagingSuite 5.2 software. Average fluorescence intensity outside of the cell was subtracted from the mean fluorescence intensities to correct for background fluorescence. After background correction, fluorescence versus time traces were further analysed in Origin 6.1 (OriginLab Corp., Northampton, MA, USA) and represent the averaged fluorescence from a region of interest (ROI) centred on the spark generating area. This ROI size was determined empirically to be the best compromise between temporal and spatial precision of Ca²⁺ sparks and the signal to noise ratio. Ca²⁺ spark amplitude was expressed as relative fluorescence increase F/F₀, where F is the peak fluorescence and F₀ is the baseline fluorescence in the ROI before Ca²⁺ spark appearance. The latency of occurrence of Ca²⁺ sparks was measured as the time elapsed from the onset of the pulse depolarization to the moment when Ca²⁺ spark amplitude reached 5–10% of its maximum.

K⁺ current recordings

K⁺ currents were measured by the conventional whole-cell or perforated whole-cell patch technique (Gollasch et al. 1996; Lohn et al. 2001a). In perforated patch recordings, whole cell access was achieved by amphotericin B within 10 min of seal formation at room temperature (20–24°C). Amphotericin B (Sigma) was dissolved in dimethyl sulfoxide (DMSO) and diluted into the pipette solution to 200 µg ml^–1. Patch pipettes (resistance, 3–5 M) were filled with a solution containing (mM): 110 potassium aspartate, 30 KCl, 10 NaCl, 1 MgCl₂, 10 Hepes and 0.05 EGTA (pH 7.2). The external solution contained (mM): 134 NaCl, 6 KCl, 1 MgCl₂, 2 CaCl₂, 10 glucose and 10 Hepes (pH 7.4).

To clamp the global [Ca²⁺]_i at different levels, STOCs were recorded in the conventional whole-cell mode using the following pipette solutions (mM): 80 potassium aspartate, 50 KCl, 10 NaCl, 1 MgCl₂, 3 MgATP, 10 Hepes, 10 EGTA and different [CaCl₂]_i (pH 7.2). [CaCl₂]_i were 0.01, 4 and 8 mM and equalled 0.18, 120 and 1000 nM free cytosolic [Ca²⁺]. The free cytosolic [Ca²⁺] was calculated by the ‘Cabuf’ program written by Prof G. Droogmans (available at ftp.cc.kuleuven.ac.be/pub/droogmans/cabuf.zip) and based on the stability constants given by Fabiato & Fabiato (1979). To set the free [Ca²⁺]_i at 100 nM and the free [EGTA]_i at different levels, STOCs were recorded in the conventional whole-cell mode using the following pipette solutions (mM): (a) 80 potassium aspartate, 45 KCl, 10 NaCl, 1 MgCl₂, 3 MgATP, 10 Hepes, 17 EGTA and 7 CaCl₂ (pH 7.2 with KOH) – (10 mM free [EGTA]_i, 100 nM free [Ca²⁺]_i); or (b) 80 potassium aspartate, 45 KCl, 10 NaCl, 1 MgCl₂, 3 MgATP, 30 glucose, 10 Hepes, 1.7 EGTA and 0.7 CaCl₂ (pH 7.2 with KOH) – (1 mM free [EGTA]_i, 100 nM free [Ca²⁺]_i); or (c) 80 potassium aspartate, 45 KCl, 10 NaCl, 1 MgCl₂, 3 MgATP, 30 glucose, 10 Hepes, 0.17 EGTA and 0.07 CaCl₂ (pH 7.2 with KOH) – (0.1 mM free [EGTA]_i, 100 nM free [Ca²⁺]_i). When indicated, 20 µM Fluo4-AM was added into the pipette solutions to monitor caffeine-induced Ca²⁺ release. Whole cell currents were recorded 5–7 min after disruption of the membrane.

Whole cell currents were recorded using an EPC 7 amplifier (List, Darmstadt, Germany), digitized at 5 kHz, using a CED 1401 series interface (Cambridge Electronic Design Ltd, Cambridge, UK), and CED patch and voltage clamp software version 6.08 or an EPC9 amplifier under contol of Pulse software (HEKA Electronik, Lambrecht, Germany). STOC latency to occurrence upon membrane depolarization was measured using the pulse protocol shown in the online Supplemental Fig. 2, similar to that in Lopez-Lopez et al. (1995). The latency of occurrence (or waiting time) of STOCs was measured as the time elapsed from the onset of the pulse depolarization to the peak of STOC minus 15 ms (average time to peak). STOC analysis was performed off-line using custom software written in C++ by K. Essin or Origin 6.1.

Recording of global intracellular [Ca²⁺]

Intracellular [Ca²⁺]_i was monitored at 35°C as described (Gollasch et al. 1991). Briefly, isolated tibial myocytes were loaded with 3 µM Fura-2-AM for 30 min in buffer solution (mM: 137 NaCl, 5.4 KCl, 1.8 CaCl₂, 1 MgCl₂, 10 Hepes and 5.6 glucose). In some experiments, cells were incubated with EGTA-AM at different concentrations (10 min) after the Fura-2-AM loading. [Ca²⁺]_i was continuously recorded as fluorescence intensity (at 510 nm) at alternating 350 (F₃₅₀) and 380 nm (F₃₈₀) excitation wavelengths and their respective ratio (F₃₅₀/F₃₈₀) by using TILL vision devices (http://www.till-photonics.de). [Ca²⁺] was calculated using the following equation:

(2)

with R_min and R_max measured from ionomycin treated cells, and K_d determined (Moosmang et al. 2003). The pooled mean values for R_max, R_min and K_d were 8.97, 1.68 and 307 nM, respectively. Stimulation was performed by local application of caffeine (10 mM) via a syringe device. The resting global cytosolic [Ca²⁺]_r was used for estimation of Ca²⁺ spark amplitudes.

Materials

Fluo-3-AM, Fluo-4-AM, EGTA-AM and Fura-2-AM were purchased from Molecular Probes (Eugene, OR, USA). Stock solutions (0.25 mM) of fluo-3-AM were made using DMSO as the solvent. Ryanodine was obtained from Calbiochem (Bad Soden, Germany). All salts and other drugs were obtained from Sigma-Aldrich (Deisenhofen, Germany) or Merck (Darmstadt, Germany). High external potassium solutions were made by iso-osmotic substitution of NaCl with KCl in the PSS.

All values are given as means ± S.E.M. Data were compared with Student's t test (P < 0.05). The term ‘n’ represents the cell number tested.

	Results

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Elementary SR release in SMAKO cells

Figure 1A shows confocal line-scan images of representative tibial artery VSMC isolated from control and SMAKO mice. Ca²⁺ sparks were observed only in close proximity to the cell surface in both control and SMAKO cells. Ca²⁺ sparks in control cells had an estimated mean amplitude of 418 ± 26 nM, a mean rise time of 22.4 ± 1.3 ms, a decay half-life of 51.6 ± 4.4 ms, and a width at half-maximal amplitude of 3.3 ± 0.1 µm. The frequency of Ca²⁺ sparks was 1.1 ± 0.1 Hz (n = 162; Fig. 1B). In contrast, Ca²⁺ spark frequency and amplitude were reduced by 50% and 80% in SMAKO cells, compared to control cells. The Ca²⁺ spark frequency and estimated spark amplitude were 0.6 ± 0.1 Hz and 55 ± 7 nM in SMAKO cells (n = 79 cells) (Fig. 1B). However, the rise time, decay and width of Ca²⁺ sparks were not different in SMAKO cells. The mean rise time, decay and width were 20.8 ± 1.8 ms, 48.7 ± 5.9 ms and 3.2 ± 0.2 µm, respectively (n = 40 cells). Similar effects were observed in cerebral artery VSMCs (online Supplemental Fig. 1). Since SMAKO cells lack functional L-type Ca_v1.2 Ca²⁺ channels (Moosmang et al. 2003), the data indicate that Ca_v1.2 channels play an important role in the generation of VSMC Ca²⁺ sparks.

View larger version (22K): [in this window] [in a new window]   Figure 1.  Ca2+ sparks in control and SMAKO tibial VSMC A, upper panel, confocal line-scan image of fluo-3-loaded wild-type (WT) cell with the time course of Ca2+ sparks indicated below. The fluorescence time course of the Ca2+ sparks was determined over the line indicated by the two arrows. Each line-scan image is a plot of fluorescence along a scanned line (ordinate) versus time (abscissa). The line-scan image duration was 5 s, and each line was 4 ms. Lower panel, confocal line-scan image of a fluo-3-loaded SMAKO cell. Amplitudes of Ca2+ sparks are expressed as absolute values ([Ca2+]i) relative to the global resting cytosolic [Ca2+]r at F0 using eqn (1). B, comparison of spatial–temporal characteristics of Ca2+ sparks in WT and SMAKO cells.

View larger version (22K): [in this window] [in a new window]   Figure 2.  STOCs caused by Ca2+ sparks A, STOCs in representative cells isolated from tibial arteries from control (upper panels, WT) and SMAKO mice (lower panels). The holding potential was stepwise increased in 20 mV increments from –40 mV to 0 mV. B, comparison of STOCs characteristics. The holding potential was –40 mV, –20 mV or 0 mV, *P < 0.05; n.s., not significant.

Dihydropyridines modulate the open probability of L-type Ca²⁺ channels (Catterall & Striessnig, 1992; Hofmann et al. 1999) and therefore should affect spark frequency in wild-type VSMCs. The effects of the dihydropyridines on Ca²⁺ sparks were studied between 1 and 15 min, between 15 and 30 min, and between 30 and 90 min after application of the drugs (Fig. 3A and C). Nimodipine (1 µM) did not affect Ca²⁺ sparks (n = 145), compared to control cells (n = 115) in the absence of the drug, when sparks were determined 1–15 min after application of nimodipine. However, nimodipine reduced the Ca²⁺ sparks frequency 15 and 90 min after application (Fig. 3C). As expected, the Ca_v1.2 channel activator BayK 8644 (1 µM) increased the frequency of Ca²⁺ sparks 60–90 min after application. BayK 8644 and nimodipine did not affect the decay and width of Ca²⁺ sparks (not shown).

View larger version (16K): [in this window] [in a new window]   Figure 3.  Effects of L-type Ca2+ channel modulators on Ca2+ spark frequency in control (WT, A) and SMAKO (B) cells isolated from tibial arteries Ca2+ sparks were recorded in the absence (Control) and presence of 1 µM BayK 8644 (BayK), 1 µM nimodipine (Nim), 30 mM KCl (KCl), and 30 mM KCl plus 1 µM nimodipine (KCl Nim); n 40 cells for each group. Cells were preincubated with the compounds for 30–90 min. C, summary of Ca2+ spark frequency. WT cells from tibial arteries were preincubated with 1 µM nimodipine for 1–15 min, 15–30 min and 30–90 min before the recording. n 40 for each group, *P < 0.05.

We next measured STOCs in the absence and presence of dihydropyridines and Cd²⁺. The effects were analysed between 1 and 15 min after drug application. Figure 4 shows that BayK 8644 (1 µM), nimodipine (1 µM) or nimodipine (1 µM) plus the inorganic L-type Ca_v1.2 channel blocker Cd²⁺ (100 µM) did not affect STOCs in both control cells and SMAKO tibial artery VSMCs. Similar results were observed in cerebral VSMCs (not shown). However, STOCs were inhibited in control cells by 50% after 30–90 min application of 1 µM nimodipine (not shown).

View larger version (11K): [in this window] [in a new window]   Figure 4.  Effects of L-type Ca2+ channel modulators on STOC frequencies in wild type (A) and SMAKO (B) SMCs Cells were preincubated for 1–15 min with the indicated compounds. STOCs were recorded at a holding potential of –20 mV. STOCs were recorded in the absence (Control) and presence of 1 µM BayK 8644 (BayK), 1 µM nimodipine (Nim), and 1 µM nimodipine plus 300 µM Cd2+ (Nim Cd). n 7 cells for each group.

View larger version (15K): [in this window] [in a new window]   Figure 5.  Effects of external Ca2+ and 2-aminoethoxydiphenyl borate (2-APB) on Ca2+ spark and STOC frequency of control (WT) cells isolated from tibial arteries A, Ca2+ sparks were recorded in the presence of 1.8 mM external Ca2+ (Control) and in nominally Ca2+-free solution (0 [Ca2+]o), 0 mM external Ca2+ plus 10 µM 2-APB, 0 mM external Ca2+ plus 50 µM 2-APB, and 50 µM 2-APB; n 30 cells per group. B, STOCs were recorded the absence (Control) and presence of 100 µM 2-APB at a holding potential of –20 mV (n = 5, each). *P < 0.05.

Resting [Ca²⁺]_i was lower in SMAKO cells compared to control cells (30 nM versus 130 nM in SMAKO versus control cells) (Fig. 6B), suggesting that steady-state [Ca²⁺]_i is tightly controlled by Ca²⁺ influx through Ca_v1.2 channels. SR Ca²⁺ load can be analysed by the use of caffeine, which activates each of the RyRs. Caffeine (10 mM) evoked smaller Ca²⁺ transients in SMAKO cells, as compared to control cells (Fig. 6A and C), indicating that ryanodine-sensitive stores are depleted in SMAKO cells. We used a multipulse application protocol to further characterize the disrupted Ca²⁺ uptake into ryanodine-sensitive stores in SMAKO mice. Eight minutes after a conditioning caffeine pulse, subsequent applications of caffeine (10 mM) induced [Ca²⁺]_i transients only in control cells, but not in SMAKO VSMCs. Caffeine did not induce [Ca²⁺]_i elevations at earlier time points in both cell types. The poor recovery of the Ca²⁺ transient in SMAKO VSMCs again suggested that refilling of ryanodine-sensitive Ca²⁺ stores mainly depends on Ca²⁺ influx through Ca_v1.2 channels. The results support the interpretation that RyR stores are exposed to lower global cytosolic [Ca²⁺]_i in SMAKO VSMCs resulting in a reduced driving force for SR Ca²⁺ loading and a reduced SR Ca²⁺ content. Furthermore, the data indicate that Ca_v1.2 channels significantly contribute to Ca²⁺ store refilling after Ca²⁺ store depletion in VSMC.

View larger version (18K): [in this window] [in a new window]   Figure 6.  Effects of caffeine on [Ca2+]i in control (WT) and SMAKO cells isolated from tibial arteries A, time course of caffeine's effect on [Ca2+]i. Horizontal lines indicate the presence of caffeine (10 mM) in the bath solution. B, comparison of resting [Ca2+]i in WT and SMAKO cells (n = 70, each). C, comparison of 10 mM caffeine-induced Ca2+ release in WT and SMAKO cells (n = 74 cells for each group). [Ca2+]i increases induced by the first application of caffeine (0 min) and after a time interval of 8 min are compared. The Ca2+ responses are normalized to the effects (100% response) of 10 mM caffeine in WT cells at 0 min. AUC, areas under the curve.

We next investigated whether or not RyRs are able to sense local Ca²⁺ entry by performing experiments in which we clamped the global [Ca²⁺]_i at 0.18 nM, 120 nM and 1000 nM while maintaining high mobile intracellular Ca²⁺ buffer (10 mM EGTA). Figure 7A shows that STOCs were strongly controlled by the global cytosolic [Ca²⁺]_i with significantly higher frequencies of STOCs at increasing global [Ca²⁺]_i. Moreover, STOC frequencies were similar in SMAKO and control cells at identical [Ca²⁺]_i suggesting that the source leading to increased [Ca²⁺]_i is irrelevant for the induction of Ca²⁺ sparks.

View larger version (15K): [in this window] [in a new window]   Figure 7.  Effects of clamped [Ca2+]i and EGTA-AM on Ca2+ spark, STOC frequency and caffeine-induced Ca2+ release in tibial VSMCs A, STOCs were recorded in the whole-cell mode at –40 mV for 2 min. [Ca2+]i was clamped to 0.18, 120 or 1000 nM in control (WT) and SMAKO cells (n 8, each). B, effects of different concentrations of EGTA-AM on Ca2+ spark frequency in WT cells (n 30, each). C, effects of different concentrations of EGTA-AM on caffeine-induced Ca2+ release in WT cells (n 10, each). The Ca2+ responses were normalized to the effect of 10 mM caffeine (100% response) in control cells. AUC, area under the curve.

View larger version (20K): [in this window] [in a new window]   Figure 8.  Effects of different [EGTA]i on STOC frequency and caffeine-induced Ca2+ release in tibial VSMCs The free intracellular Ca2+ concentration [Ca] was clamped at 100 nM. A, STOC frequency and amplitude recorded in the whole-cell mode at –40, –20 and 0 mV for 2 min. Free [EGTA]i was set to 0.1, 1 or 10 mM (n cells 10, each concentration); free [Ca] was 100 nM. B, left, caffeine-induced Ca2+ release in cells loaded with free [Ca2+]i at 100 nM and free EGTA at 0.1 mM (upper panel) and with free [Ca2+]i at 100 nM and free EGTA at 10 mM (lower panel). The holding potential was –40 mV. Right, summary of the results. The Ca2+ responses were normalized to the effect of 10 mM caffeine (100% response). AUC, areas under the curve. (n 6 cells, each.)

View larger version (30K): [in this window] [in a new window]   Figure 9.  Analysis of first-latency events (Ca2+ sparks) A, histograms of first-latency events from the start of the depolarization to the first identified Ca2+ spark. Plots show histograms of latencies to the onset of Ca2+ sparks elicited by 500 ms depolarizing pulses to the indicated membrane potentials (–30 mV, –20 mV, –10 mV, 0 mV, +10 mV, +20 mV, +30 mV, +40 mV, +50 mV, +60 mV, +70 mV, +80 mV, +90 mV and +100 mV) for wild-type tibial VSMCs (300 nM nimodipine; n = 15 cells of 5 mice). B, amplitude of first-latency Ca2+ sparks was plotted as a function of the depolarizing voltage. C, average latency of first-latency Ca2+ sparks was plotted as a function of the depolarizing voltage. D, first-latency histogram peak NSparks/s was plotted as a function of depolarizing voltage. The continuous lines represent polynomial second order fits to the data. Curve fitting revealed maximal values of peak NSparks/s at 18.8 mV.

	Discussion

Top Abstract Introduction Methods Results Discussion References

Rapid, local and tight coupling between the Ca_v1.2 channels and RyR is not required

Our data show that Ca_v1.2 channels do not directly control RyRs in the CRU via elevation of Ca²⁺ locally restricted at the cytosolic site of the Ca_v1.2 channels in VSMCs. We reach this conclusion for several reasons. First, the effects of nimodipine resembled the results obtained with the SMAKO VSMCs but developed extremely slowly over many minutes. This observation is in contrast to the rapid inhibitory effects of this drug on L-type channels, which can be observed within seconds (Ruth et al. 1985; McCarthy & Cohen, 1989; Lohn et al. 2002). The failure of nimodipine to induce rapid (within seconds) inhibition of Ca²⁺ sparks indicates that many of the RyRs in the CRU are not directly controlled by high local [Ca²⁺]_i caused by the opening of adjacent Ca_v1.2 channels. Second, the robustness and latency analysis showed that the Ca²⁺ release events did not reflect the voltage dependency and kinetic properties of single L-type channels. In these experiments, we first analysed the histograms of these events using VSMC cells treated with an L-type channel blocker to satisfy Poisson distribution of Ca_v1.2 channel openings (Lopez-Lopez et al. 1995; Collier et al. 1999). Unlike in cardiac muscle, the histograms of these events did not show steep bell-shaped curves characteristic for Ca²⁺ sparks triggered by an elevation in local cytosolic Ca²⁺ from single L-type Ca²⁺ channels (Lopez-Lopez et al. 1995; Cleemann et al. 1998; Collier et al. 1999). In addition, we also found that important features of the nimodipine-treated VSMC histograms, namely those cells with low Ca_v1.2 channel activity, did not differ from histograms of control cells (with high Ca_v1.2 channel activity) and from SMAKO cells (without Ca_v1.2 channels). Third, Ca²⁺ sparks were completely suppressed by excessive concentrations of a slow, high affinity cytosolic Ca²⁺ buffer (millimolar concentrations of EGTA), which suppresses global cytosolic [Ca²⁺] and causes a slow depletion of ryanodine-sensitive SR Ca²⁺ content in VSMCs, but which should not affect Ca²⁺ communication between Ca_v1.2 channels and RyR occurring on the nanometer scale (Stern, 1992; Collier et al. 2000). Finally, the substantially reduced frequency and amplitude of Ca²⁺ sparks in SMAKO VSMCs were completely reversed by elevating cytosolic Ca²⁺ levels demonstrating that the source leading to increased global [Ca²⁺]_i is irrelevant for the induction of Ca²⁺ sparks. Taken together, these results strongly suggest that the intimate association between the trigger Ca_v1.2 L-type channel and target RyRs is not crucial for RyRs to generate Ca²⁺ sparks in VSMCs. Therefore, we suggest that the intermolecular mechanisms leading to elementary SR Ca²⁺ release differ substantially between arterial, cardiac and skeletal muscle (Lopez-Lopez et al. 1995; Klein et al. 1997; Grabner et al. 1999; Collier et al. 2000; Papadopoulos et al. 2004).

Ca_v1.2 channels contribute to global cytosolic [Ca²⁺], which in turn influences luminal SR calcium and thus Ca²⁺ sparks. Our study shows that the substantially reduced frequency and amplitude of Ca²⁺ sparks in SMAKO VSMCs is associated with lower global [Ca²⁺]_i levels and reduced SR Ca²⁺ load. The data indicate that the cytosolic [Ca²⁺] is mainly determined by Ca²⁺ influx through Ca_v1.2 channels. We provided direct evidence and observed that these effects are completely reversed by elevating cytosolic Ca²⁺ levels (Fig. 7A). Our results indicate that cytosolic [Ca²⁺]_i itself contributes minimally to the acute triggering of the physiologically relevant proportion of Ca²⁺ sparks. This conclusion is based on the following findings. First, RyRs in situ are relatively insensitive to global cytosolic Ca²⁺ levels (Jaggar et al. 2000). Second, the probability and latency histograms between SMAKO, nimodipine-treated and control cells were not different, despite the fact that the voltage steps used in these experiments induced relatively rapid increases in global [Ca²⁺]_i from 100 nM to 300 nM (within 500 ms to 1 s) in wild-type VSMCs, but not in VSMCs without functional L-type channels (Kamishima & McCarron, 1996; Kamishima et al. 2000; Lohn et al. 2001b). Similar data were obtained in wild-type tibial VSMCs, but not in SMAKO cells (K. Essin & M. Gollasch, unpublished observations). In agreement with these suggestions, rapid (for instance, caffeine or thapsigargin) or slow (phospholamban deficiency) modulation of SR Ca²⁺ load has profound direct effects on both amplitudes and frequency of Ca²⁺ sparks and STOCs, which follows the time course of depletion or uploading SR [Ca²⁺] (Nelson et al. 1995; Bychkov et al. 1997; Lohn et al. 2001a; Wellman et al. 2001). Furthermore, increasing levels of cytosolic [EGTA] dramatically reduced both the frequency and amplitude of STOCs in VSMCs, despite [Ca²⁺]_i being clamped at similar levels, i.e. 100 nM (Fig. 8A). These effects were associated with SR Ca²⁺ depletion (Fig. 8B) and resembled the effects observed in SMAKO cells. The data indicate that, in VSMCs, the luminal SR Ca²⁺ represents a very important physiological Ca²⁺ signal for activating elementary Ca²⁺ release. The luminal SR Ca²⁺ is controlled by relatively slow Ca²⁺ uptake from the global cytosolic [Ca²⁺]. The advantage of the rigorous SR Ca²⁺ defined regulation of Ca²⁺ sparks is uncoupling spark formation from rapid changes in global cytosolic [Ca²⁺] to enable a robust and stabile function of the Ca²⁺ spark/STOC pathway to lower global [Ca²⁺] and vascular tone.

Steady-state Ca²⁺-influx in SMAKO cells

In SMAKO cells, the global cytosolic [Ca²⁺] is only 16% of wild-type cells. In contrast, SR Ca²⁺ stores (Fig. 6) and Ca²⁺ spark frequency are reduced by only 50% in SMAKO cells, which is similar to the effects of nimodipine in wild-type cells. Importantly, the acute removal of Ca²⁺ from the extracellular solution produced effects similar to nimodipine, namely a reduction in the Ca²⁺ spark frequency by 50%. The remaining Ca²⁺ sparks were completely blocked by 2-APB (Fig. 6). The blocking effects of the IP₃ receptor antagonist 2-APB on Ca²⁺ sparks have been previously reported in portal vein VSMCs. Crosstalk between IP₃ receptors and RyRs has been proposed (Gordienko & Bolton, 2002). Taking into consideration the ability of 2-APB to non-selectively inhibit cation channels, for example TRP related channels (Albert & Large, 2006; Owsianik et al. 2006), an alternative explanation could be that ion influx though non-selective cation channels may serve as additional important regulators of Ca²⁺ sparks in arterial VSMCs. Possibly, these channels can function as caveolemmal Ca²⁺ channels that may produce a subpopulation of Ca²⁺ sparks in caveolar microdomains (Lohn et al. 2000). Future studies would clarify the possible role of TRP channels in the regulation of Ca²⁺ sparks in VSMCs.

In conclusion, Ca_v1.2 channels are important regulatory proteins in the indirect control of Ca²⁺ sparks in arterial VSMCs. We found that Ca_v1.2 channel genetic inactivation substantially lowers resting global [Ca²⁺]_i, which in turn reduces slowly luminal SR calcium and thus Ca²⁺ sparks. We found that rapid, local and tight coupling between the Ca_v1.2 channels and RyRs is not required to initiate Ca²⁺ sparks. The slow, indirect coupling between Ca_v1.2 channels and RyRs is in excellent agreement with the physiological function of Ca²⁺ sparks to serve as robust and stable negative feedback regulators for the global [Ca²⁺]_i and arterial tone.

	Footnotes

 
This paper has online supplemental material.

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