Ca²⁺ entry following P2X receptor activation induces IP₃ receptor-mediated Ca²⁺ release in myocytes from small renal arteries

Abstract

BACKGROUND AND PURPOSE

P2X receptors mediate sympathetic control and autoregulation of the renal circulation triggering contraction of renal vascular smooth muscle cells (RVSMCs) via an elevation of intracellular Ca²⁺ concentration ([Ca²⁺]_i). Although it is well-appreciated that the myocyte Ca²⁺ signalling system is composed of microdomains, little is known about the structure of the [Ca²⁺]_i responses induced by P2X receptor stimulation in vascular myocytes.

EXPERIMENTAL APPROACHES

Using confocal microscopy, perforated-patch electrical recordings, immuno-/organelle-specific staining, flash photolysis and RT-PCR analysis we explored, at the subcellular level, the Ca²⁺ signalling system engaged in RVSMCs on stimulation of P2X receptors with the selective agonist αβ-methylene ATP (αβ-meATP).

KEY RESULTS

RT-PCR analysis of single RVSMCs showed the presence of genes encoding inositol 1,4,5-trisphosphate receptor type 1(IP₃R1) and ryanodine receptor type 2 (RyR2). The amplitude of the [Ca²⁺]_i transients depended on αβ-meATP concentration. Depolarization induced by 10 µmol·L⁻¹αβ-meATP triggered an abrupt Ca²⁺ release from sub-plasmalemmal (‘junctional’) sarcoplasmic reticulum enriched with IP₃Rs but poor in RyRs. Depletion of calcium stores, block of voltage-gated Ca²⁺ channels (VGCCs) or IP₃Rs suppressed the sub-plasmalemmal [Ca²⁺]_i upstroke significantly more than block of RyRs. The effect of calcium store depletion or IP₃R inhibition on the sub-plasmalemmal [Ca²⁺]_i upstroke was attenuated following block of VGCCs.

CONCLUSIONS AND IMPLICATIONS

Depolarization of RVSMCs following P2X receptor activation induces IP₃R-mediated Ca²⁺ release from sub-plasmalemmal (‘junctional’) sarcoplasmic reticulum, which is activated mainly by Ca²⁺ influx through VGCCs. This mechanism provides convergence of signalling pathways engaged in electromechanical and pharmacomechanical coupling in renal vascular myocytes.

Introduction

Renal blood flow (RBF) accounts for 20–25% of cardiac output at rest (Malpas and Leonard, 2000; Cupples and Braam, 2007). The mechanisms controlling contraction/relaxation of renal vascular smooth muscle cells (RVSMCs), which regulate the diameter of renal resistance arteries and hence RBF and glomerular filtration rate, play a fundamental role in the control of systemic blood pressure (DiBona, 2004; Navar, 2005).

Vascular myocytes are stimulated to contract by elevation of intracellular Ca²⁺ concentration ([Ca²⁺]_i) resulting in activation of Ca²⁺/calmodulin-dependent myosin light chain kinase which causes force development via myosin light chain phosphorylation (Walsh, 1994). An increase in [Ca²⁺]_i in smooth muscle cell (SMC) is caused by either depolarization of the myocyte membrane (electromechanical coupling; Somlyo and Somlyo, 1968) resulting from activation of ionotropic receptors or metabotropic receptor-coupled cationic channels and leading to Ca²⁺ influx via voltage-gated Ca²⁺ channels (VGCCs), or by activation of metabotropic receptors (pharmacomechanical coupling; Somlyo and Somlyo, 1968) coupled via G_q/11-protein to phospholipase C (PLC) activation followed by inositol 1,4,5-trisphosphate (IP₃) liberation and IP₃ receptor (IP₃R)-mediated Ca²⁺ release, or by a combination of these mechanisms (Bolton et al., 1999; Davis and Hill, 1999; Wray and Burdyga, 2010). Either of these events, which cause an initial rise in [Ca²⁺]_i, may be further augmented by ryanodine receptor (RyR)-mediated Ca²⁺ release activated via a Ca²⁺-induced Ca²⁺ release (CICR) mechanism (Fabiato and Fabiato, 1975). However, the recruitment of RyR-mediated Ca²⁺ release by voltage-gated Ca²⁺ entry and/or IP₃R-mediated Ca²⁺ release in smooth muscles is still an area of extensive debate and controversy (reviewed in Laporte et al., 2004; Wray and Burdyga, 2010). Indeed, the relative contribution of RyRs and IP₃Rs to intracellular [Ca²⁺]_i mobilization varies in different types of SMCs, and often depends on the strengths and mechanism of SMC stimulation. In some phasic smooth muscles, for example, urinary bladder and vas deferens, Ca²⁺ entry through VGCCs is tightly coupled to RyR-mediated Ca²⁺ release. Electrical stimulation of these SMCs, in which close proximity of VGCCs and RyRs within the caveolar domains was confirmed with 3D-immunofluorescence and electron microscopy (Moore et al., 2004), triggers an abrupt RyR-mediated Ca²⁺ release at multiple sub-PM regions (‘hot spots’, Ohi et al., 2001; Morimura et al., 2006). The ability of Ca²⁺ entering the SMC via VGCCs to induce RyR-mediated Ca²⁺ release has also been demonstrated in voltage-clamp experiments performed on SMCs from pregnant myometrium (Shmigol et al., 1998), ileum (Kohda et al., 1997), mesenteric artery (Bolton and Gordienko, 1998), cerebral arteries (Kamishima and McCarron, 1997) and portal vein (Coussin et al., 2000). However, other research groups working on different or even the same SMC type failed to detect any evidence of CICR (Fleischmann et al., 1996; Kamishima and McCarron, 1996; Bradley et al., 2002). The ability of Ca²⁺ liberated via IP₃Rs to trigger RyR-mediated Ca²⁺ release also varies among different types of smooth muscles and depends on receptor distribution and isoform expression (reviewed in Wray and Burdyga, 2010). Emerging evidence highlights the importance of RyRs and CICR in calcium signalling and arteriolar contraction in the renal microcirculation. It was demonstrated that in these arterioles RyR/CICR contribute to [Ca²⁺]_i responses initiated by Ca²⁺ entry and by IP₃R-mediated Ca²⁺ release (reviewed in Arendshorst and Thai, 2009).

Two major sympathetic cotransmitters, noradrenaline and ATP, mediate vasoconstriction of small arteries (Wier et al., 2009) by acting on metabotropic α₁-adrenoceptors and ionotropic P2X receptors respectively. Thus, P2X receptors provide an important therapeutic target, mediating substantial vasoconstrictor drive resistant to adrenoceptor antagonists (Wier et al., 2009).

In the kidney, apart from sympathetic transmission, ATP is involved in paracrine regulation. On being released by apical macula densa cells in response to an increase in NaCl concentration in the distal tubular fluid (Bell et al., 2003), ATP triggers P2X receptor-mediated vasoconstriction of afferent arterioles (Inscho et al., 2003), thus decreasing RBF and glomerular filtration rate. This tubuloglomerular feedback mechanism (Nishiyama et al., 2000; Guan et al., 2007a; Surprenant and North, 2009) and myogenic mechanism are two important components of renal autoregulation (Cupples and Braam, 2007).

The respective role of RyRs and IP₃Rs in smooth muscle contraction is an area of active investigation. Although RyRs and IP₃Rs are both Ca²⁺-sensitive and can activate each other via CICR mechanism, leading to an all-or-none regenerative response, graded Ca²⁺ signals have been reported to be evoked by increasing stimuli targeting RyRs (Isenberg and Han, 1994) or IP₃Rs (Bootman et al., 1994). The resolution of this paradox has been suggested to lie in the local control of Ca²⁺ microdomains that function and are regulated autonomously (Berridge, 1997), the gradual recruitment of which by gradually increasing stimuli would result in graded Ca²⁺ signals. Imaging microdomain Ca²⁺ in muscle cells has reshaped our understanding of Ca²⁺ signalling and provided direct evidence to validate the local control theory in skeletal, cardiac and smooth muscles (Wang et al., 2004; McCarron et al., 2006).

In smooth muscles, IP₃R-mediated Ca²⁺ release is essential for [Ca²⁺]_i mobilization and contraction, especially (but not exclusively) induced by stimulation with neurotransmitters and hormones (Boittin et al., 1999; Bayguinov et al., 2000; McCarron et al., 2002; Zhang et al., 2003; Lamont and Wier, 2004; MacMillan et al., 2005; Gordienko et al., 2008). The role of IP₃Rs seems to be to facilitate CICR within and between RyRs domains (Boittin et al., 1998; Gordienko and Bolton, 2002; White and McGeown, 2002; Zhang et al., 2003). However, a cooperative relationship between IP₃Rs and RyRs is not always the rule. When both receptor types share the same calcium store, the interplay between them can be regulated via alterations in the sarcoplasmic reticulum (SR) calcium load. For example, in SMCs from both the gastric antrum and vas deferens block of IP₃Rs, which caused an elevation of the caffeine-releasable store content, increased spontaneous Ca²⁺ spark activity (White and McGeown, 2002, 2003).

Another important aspect of the SMC Ca²⁺ signalling system is the regulation of IP₃R-mediated Ca²⁺ release by [Ca²⁺]_i. In vascular SMCs of small arteries, Lamont and Wier (2004) concluded that if RyRs were blocked, cell-wide Ca²⁺ waves could still be evoked by strong activation of adrenoceptors leading to myocyte and vessel contraction. Thus, CICR apparently can occur among IP₃Rs alone, although normally, at low levels of adrenoceptor activation RyRs are needed to trigger IP₃R dependent Ca²⁺ waves (Wier and Morgan, 2003). Furthermore, activation of IP₃Rs by a localized increase in [Ca²⁺]_i was directly demonstrated in an elegant experiment conducted on rabbit urinary bladder myocytes; IP₃R-mediated Ca²⁺ release was found to be induced by local Ca²⁺ uncaging (Ji et al., 2006).

We have previously demonstrated that following activation of muscarinic receptors in intestinal myocytes IP₃R-mediated Ca²⁺ release is facilitated by Ca²⁺ influx via VGCCs (Gordienko et al., 2008). Although M₃ receptors are linked to a G_q/11/PLC/IP₃/Ca²⁺ signalling pathway, we hypothesized that in contrast to cardiac muscle, excitation–contraction (E–C) coupling in smooth muscle occurs by Ca²⁺ entry through VGCCs, which evokes IP₃R-mediated Ca²⁺ release via CICR mechanism. This hypothesis needs to be tested in vascular myocytes regulated by ionotropic receptors, which are not coupled to the G_q/11-GTP/PLC/IP₃ system.

Taking into account the importance of ionotropic P2X receptors (Burnstock, 2007; Surprenant and North, 2009) in the regulation of renal circulation (Malpas and Leonard, 2000; Inscho et al., 2003; Cupples and Braam, 2007; Guan et al., 2007a), we tested our hypothesis on RVSMCs, which, as we have recently demonstrated, express functional monomeric P2X1 and heteromeric P2X1/4 receptors (Harhun et al., 2010). We found that depolarization of RVSMCs following P2X receptor activation induces IP₃R-mediated Ca²⁺ release from sub-plasmalemmal (‘junctional’) sarcoplasmic reticulum (jSR), which is activated mainly by Ca²⁺ influx through VGCCs. Our results support the notion that, in contrast to cardiac muscle, E–C coupling in vascular smooth muscle may occur by Ca²⁺ entry through VGCCs, which evokes an initial IP₃R-mediated Ca²⁺ release activated via a CICR mechanism. A preliminary account of this study has been published in abstract form (Povstyan et al., 2009).

Methods

Isolation of renal vascular smooth muscle cells (RVSMCs)

Male Wistar Kyoto rats (180–250 g, 65 animals obtained from Charles Rivers laboratories) were humanely killed by decapitation after cervical dislocation as approved under Schedule 1 of the UK Animals (Scientific Procedures) Act 1986. Interlobar and arcuate arteries were dissected from the kidney as described previously (Gordienko et al., 1994). Following the dissection, the blood vessels with outer diameter of 50–150 µm were cut into 1–2 mm pieces and collected in ice-cold physiological salt solution (PSS) containing (in mmol·L⁻¹): NaCl 120, KCl 6, CaCl₂ 2.5, MgCl₂ 1.2, glucose 12, HEPES 10; pH adjusted to 7.4 with NaOH. After a 10 min rinse in Ca²⁺-free PSS, the tissue samples were transferred into the same solution supplemented with (mg·mL⁻¹): protease (Type X) 0.75, collagenase (Type 1A) 1.5, soybean trypsin inhibitor 1 and bovine serum albumin 1, and incubated for 20 min at 37 °C. The tissue samples were then rinsed for 10 min in an enzyme-free Ca²⁺-free solution and triturated using a glass Pasteur pipette. Isolated RVSMCs were obtained by several cycles of trituration, each followed by transfer to fresh solution with gradually increasing [Ca²⁺] (from 0.125 to 1.25 mmol·L⁻¹). Transfer of the tissue samples to fresh solution before trituration facilitated removal of debris and damaged cells from the suspension, while gradually increasing [Ca²⁺] in the medium prevented the myocytes from ‘calcium shock’ and hypercontraction (Gordienko and Zholos, 2004). Small aliquots of the cell suspension were transferred to the experimental chambers and diluted with PSS. RVSMCs were then either collected for reverse transcription polymerase chain reaction (RT-PCR) experiments or used for electrical recording and confocal imaging within 8 h of isolation.

RNA isolation and RT-PCR analysis

Reverse transcription polymerase chain reaction analysis was performed using single isolated RVSMCs. The myocytes (pool of ∼500 cells per sample) were collected under a microscope using a wide-bore glass micropipette and frozen immediately after collection. Total RNA was extracted using the Qiagen RNeasy extraction kit (Qiagen, Crawley, UK). cDNA was obtained using Superscript II Reverse Transcriptase (Invitrogen, Paisley, UK) and used in RT-PCR. cDNA was used as a template in a 50 µL RT-PCR reaction containing 1.5 mmol·L⁻¹ MgCl₂, 0.2 mmol·L⁻¹ deoxynucleoside triphosphates, 0.2 µmol·L⁻¹ forward and reverse primers (Invitrogen), and 2.5 units of platinum Taq DNA polymerase (Invitrogen). Amplification was performed using a Touchgene Thermocycler (Bibby Scientific Ltd., Staffordshire, UK) according to the following schedule: 94°C for 2 min; 40 cycles (for brain and liver tissue preparations) or 45 cycles (for RVSMC samples) of 94°C for 30 s; 57°C for 60 s; and 72°C for 3 min, followed by a final elongation period of 10 min at 72°C. No-template control PCR was also performed simultaneously with every reaction. To avoid detecting genomic DNA contamination, primers were designed to span at least one intron of the genomic sequence for genes encoding proteins of interest. The experiments were repeated with at least four samples of RVSMCs obtained from different animals. The purity of the RVSMC sample was confirmed by the expression of the gene encoding the SMC marker, smooth muscle myosin heavy chain, but not the markers for other types of cell present in the vascular wall such as: fibroblasts and endothelial cells (CD34), neurones (PGP9.5) and pericytes (NG2), as previously described (Harhun et al., 2009; 2010;). PCR products were separated and visualized in ethidium bromide-stained 2% agarose gel by electrophoresis.

The following primers (the data in parentheses show: Genebank accession number, the sense bordering nucleotide position, the anti-sense bordering nucleotide position) were used in this study: β-actin (NM_031144, 306–325, 1007–1026), smooth muscle myosin heavy chain for SMCs (X16262, 447–466, 1182–1191), CD34 for endothelial cells and fibroblasts (NM_001107202, 50–69, 882–901), PGP9.5 for neurones (D10699, 54–73, 544–563), NG2 proteoglycan for pericytes (NM_031022, 1815–1834, 2791–2810), inositol 1,4,5-trisphosphate (IP₃) receptor type 1 (NM_001007235, 6244–6263, 7030–7049), IP₃ receptor type 3 (NM_013138, 6142–6161, 7088–7107), ryanodine receptor type 1 (XM_001078539, 8038–8057, 8811–8830), ryanodine receptor type 2 (NM_032078, 8513–8532, 9317–9336), ryanodine receptor type 3 (XM_342491, 8035–8054, 9101–9120).

Drug application

Renal vascular smooth muscle cells were stimulated with αβ-methylene ATP (αβ-meATP) and caffeine which were applied as a 2 s pulse through a glass micropipette attached to the outlet of a pressure ejector Picospritzer III (Intracel Ltd, Royston, Hertfordshire, UK). Similar application of the control solution (without agonist) had no effect on [Ca²⁺]_i. In the experiments where the same RVSMC was stimulated with αβ-meATP and caffeine or different concentrations of αβ-meATP, the agonist-containing micropipette was replaced in between successive agonist applications. Antagonists were superfused through the experimental bath.

Visualization of [Ca²⁺]_i changes

Changes of [Ca²⁺]_i in isolated RVSMCs were imaged using the high-affinity fluorescent Ca²⁺ indicators Fluo-4 or Fluo-3. Fluo-4 was loaded by incubating the RVSMCs for 20 min with 5 µmol·L⁻¹ fluo-4 acetoxymethyl ester (Fluo-4 AM; diluted from a stock containing 2 mmol·L⁻¹ Fluo-4 AM and 0.025% (w·v⁻¹) pluronic F-127 in dimethyl sulphoxide) followed by a 40 min wash in PSS to allow time for de-esterification. Fluo-3 was loaded through the patch pipette by dialysis of the myocyte with a solution containing 0.1 mmol·L⁻¹ of fluo-3 pentapotassium salt. The intensity of Fluo-4 or Fluo-3 fluorescence was normalized to the average fluorescence intensity in the images acquired before agonist application and colour coded. The temporal profiles of the agonist-induced [Ca²⁺]_i transients are illustrated by the plots showing: (i) the time course of the normalized Fluo-4 or Fluo-3 fluorescence intensity (F/F₀) averaged within multiple sub-plasmalemmal regions where F/F₀ changes were initiated and rose above 1.5 or (ii) the time course of F/F₀ averaged within entire confocal optical slice of RVSMC (Gordienko et al., 2008).

Visualization of intracellular calcium stores and the SR

The intracellular calcium stores in RVSMCs were visualized using the low-affinity (K_d(Ca)= 42 µmol·L⁻¹) fluorescent Ca²⁺ indicator fluo-3FF, which was loaded by incubating the myocytes with 5 µmol·L⁻¹ fluo-3FF AM at room temperature for 90 min followed by a 60 min wash in PSS (Gordienko and Zholos, 2004; Gordienko et al., 2008). The SR was visualized using: (i) ER-Tracker™ Blue-White DPX, which was loaded by incubating the myocytes with 5 µmol·L⁻¹ of the dye for 40 min; and (ii) Brefeldin A BODIPY 558/568 (Abeele et al., 2006), which was loaded by incubating the myocytes with 2 µmol·L⁻¹ of the dye for 20 min followed by 1.5 h wash.

Immunostaining of the SR Ca²⁺ release channels

Prior to immunostaining, the SR in RVSMCs was stained with Brefeldin A BODIPY 558/568, as described above. The myocytes were then fixed by incubating them in 4% (w·v⁻¹) paraformaldehyde for 15 min. Non-specific binding was blocked by incubating the myocytes with 1% (w·v⁻¹) bovine serum albumin (BSA) and 0.3% (w·v⁻¹) Triton X-100 (a cell permeabilizing agent) for 1 h at room temperature. Primary and secondary antibodies were diluted in PSS supplemented with 0.3 (w·v⁻¹) Triton X-100. To visualize the distribution of IP₃ receptors (IP₃Rs), we used an IP₃R type 1 (IP₃R1)-specific antibody, because IP₃R1 has been shown to be ubiquitous in various tissues (Mackrill et al., 1997; Gordienko and Zholos, 2004; Gordienko et al., 2008) and RT-PCR analysis confirmed expression of gene encoding IP₃R1 in RVSMCs. This antibody was developed (Santa Cruz Biotechnology Inc., Santa Cruz, CA, USA) in goat and has been shown to selectively recognize IP₃R1 in other types of SMCs (Taylor and Traynor, 1995; Gordienko and Zholos, 2004; Gordienko et al., 2008). RyRs were detected with a monoclonal anti-RyR antibody derived (Sigma-Aldrich Co., RBI, Natick, MA, USA) from the 34C hybridoma (produced by the fusion of P3X 63 Ag8.653 myeloma cells and spleen cells from Balb/c mice). This antibody reacts strongly with RyR1, 2 and 3 and was previously used for immunodetection of RyRs in single SMCs (Gordienko and Zholos, 2004; Gordienko et al., 2008). RVSMCs were incubated with primary anti-IP₃R1 or anti-RyR antibodies (both at 1:200 dilution) overnight at 4°C. Following a 10 min rinse (4 times) in PSS, primary antibody-specific binding was detected by incubating the cells for 3 h at room temperature with either donkey anti-goat IgG or goat anti-mouse IgG, both conjugated to MFP 488 (both at 1:400 dilution; MoBiTec, Göttingen, Germany). Prior to confocal imaging the extracellular solution was replaced with Vectashield mounting medium (Vector Laboratories, Inc. Burlingame, CA, USA) containing DAPI for visualization of the cell nucleus. The spatial pattern of the Ca²⁺ release channel distribution was related to the spatial organisation of the SR and nucleus. In controls, incubation with primary antibody was omitted from the experimental protocol.

Confocal microscopy

Experimental chambers with RVSMCs were placed on the stage of either Axiovert 100 M or Axiovert 200 M inverted microscopes attached to a LSM 510 and LSM 510 META laser-scanning units respectively (Zeiss, Oberkochen, Germany). The SCSi interface of the confocal microscopes was hosted by a Pentium PC (32-bit Windows NT 4.0 operating system) running LSM 510 software (Zeiss, Oberkochen, Germany).

During time series protocol, the x-y confocal images of Fluo-4 or Fluo-3 fluorescence were acquired at 20–40 Hz using a Zeiss plan-Apochromat 40× 1.3 N.A. oil-immersion objective. Fluo-4 or Fluo-3 fluorescence was excited by the 488 nm line of a 30 mW argon ion laser (Laser-Fertigung, Hamburg, Germany) and was captured at wavelengths above 505 nm. The illumination intensity was attenuated to 0.6–0.7% with an acousto-optical tuneable filter (Zeiss, Oberkochen, Germany). To optimize signal quality, the pinhole was set to provide a confocal optical section below 1.2 µm (measured with 0.2 µm fluorescent beads). The focus was adjusted to acquire the images from the middle of the myocyte depth. In the images acquired in this way, the events initiated within 1 µm of the cell surface (edge) were considered to have sub-plasmalemmal origin and their temporal profiles were analysed (see above).

The imaging of the myocyte structures and spatial distribution of the SR Ca²⁺ release channels was performed using a Zeiss plan-Apochromat 63× 1.4 N.A. oil-immersion objective. The pinhole was set to provide an x-y resolution of 0.4 µm and a confocal optical section (z resolution) of 0.8–1 µm (measured with 0.2 µm fluorescent beads). Averaging of four frames was used to reduce a shot noise. To avoid any bleed-through of the fluorescence signal in multistaining experiments, fluorochromes with well-separated excitation and emission spectra were used and imaging was performed using the frame-by-frame multitrack mode of the confocal scanner: sequential acquisition via well-separated optical channels of the x-y images produced by fluorescence of different fluorochromes. In these experiments, fluo-4, fluo-3FF and MFP 488 fluorescence were excited by the 488 nm line of a 30 mW argon ion laser and detected at wavelengths of 505–550 nm; ER-Tracker™ Blue-White DPX and DAPI fluorescence were excited by the 405 nm line of a 25 mW laser diode and detected at wavelengths of 420–480 nm; Brefeldin A BODIPY 558/568 was excited by the 543 nm line of a 1 mW helium/neon ion laser and detected at wavelengths above 560 nm. The photomultiplier gain and offset in each optical channel were set individually to achieve similar signal intensity at each channel and remove subsignal noise from the images. The adequacy of the imaging protocol applied to the multilabelled myocytes was confirmed by control experiments on the monolabelled cells.

Electrophysiological recordings

Electrical recordings from single RVSMCs preloaded with fluo-4 AM (see above) were performed by using perforated-patch (200 µg·mL⁻¹ amphotericin B) technique. This allowed low-resistance access to the cell while minimally interfering with [Ca²⁺]_i (because pores formed by amphotericin B are Ca²⁺-impermeable). The myocytes were bathed in PSS, dialysed with solution composed of (in mmol·L⁻¹): KCl 115; NaCl 6, HEPES 10 (pH adjusted to 7.4 with KOH), and stimulated with 2 s pulses of 10 µmol·L⁻¹αβ-meATP applied at 20 min intervals (see above). Fire-polished borosilicate patch pipettes filled with pipette solution were connected to the head stage of an Axopatch 200A (Molecular Devices Co., Sunnyvale, CA, USA) input amplifier and had a free-tip resistance of 2–5 MΩ. Electrical recordings were synchronized with confocal [Ca²⁺]_i imaging using a TTL synchronizing pulse generated by the confocal scanner at the beginning of the time series protocol. The electrical signals were filtered at 1 kHz (–3 dB frequency) by four-pole low-pass Bessel filter and digitized at 5 kHz using a DigiData 1200 hosted by a Pentium PC running pCLAMP 6.0 software (Molecular Devices Co., CA, USA).

Flash photolysis

Inositol 1,4,5-trisphosphate was rapidly and uniformly released within RVSMC from d-myo-inositol 1,4,5-trisphosphate, P⁴⁽⁵⁾-1-(2-nitrophenyl)ethyl ester trisodium salt (‘caged’ IP₃) by 1 ms flash of light (300–380 nm) from a xenon arc lamp, as described previously (Gordienko and Zholos, 2004). In these experiments, the myocytes were bathed in PSS and dialysed with solution composed of (in mmol·L⁻¹): KCl 115, NaCl 6, MgATP 5, Li₃GTP 1, HEPES 10 (pH adjusted to 7.4 with KOH) and supplemented with 30 µmol·L⁻¹‘caged’ IP₃ and 100 µmol·L⁻¹ fluo-3 pentapotassium salt. To ensure equilibration between the pipette solution and cytosol, confocal [Ca²⁺]_i imaging was commenced at least 5 min after establishment of the whole-cell configuration.

Data analysis and statistical procedures

Image processing was carried out using an Indy workstation (Silicon Graphic, Inc., Mountain View, CA, USA) with custom routines written in IDL (Research Systems, Inc., Boulder, CO, USA). The final figures were produced using MicroCal Origin (MicroCal Software Inc., Northampton, MA, USA) and CorelDraw 7.0 (Corel Corporation, Ottawa, Ontario, Canada). The data are expressed as mean values ± SEM for the number of cells (n) analysed. Comparative analysis of the data groups was performed using Student's t-test with the threshold for statistical significance set at the 0.05 level.

Drug/molecular target nomenclature

Nomenclature of drugs/molecular targets used in this work conforms to BJP's Guide to Receptors and Channels, 4th edition (Alexander et al., 2009).

Drugs and chemicals

Fluo-4 AM, fluo-3 pentapotassium salt, ER-Tracker™ Blue-White DPX, Brefeldin A BODIPY 558/568 and pluronic F-127 were obtained from Invitrogen Ltd. (Paisley, UK). Fluo-3FF acetoxymethyl ester was from TefLabs (Austin, TA, USA). Donkey anti-goat IgG and goat anti-mouse IgG, both conjugated to MFP 488 were from MoBiTec (Göttingen, Germany). d-myo-inositol 1,4,5-trisphosphate, P⁴⁽⁵⁾-1-(2-nitrophenyl)ethyl ester, trisodium salt (‘caged’ IP₃) was from Calbiochem-Novabiochem (UK) Ltd. (Beeston, Nottingham, UK). αβ-meATP trisodium salt, NF279, MRS2578 and (7R)-4-Hydroxy-7-methoxy-N,N,N-trimethyl-3,5,9-trioxa-4-phosphaheptacosan-1-aminium-4-oxide (edelfosine) were from Tocris (Bristol, UK). The RNA extraction kit was purchased from Qiagen (Crawley, UK); the primers and other reagents for RT-PCR experiments were purchased from Invitrogen (Paisley, UK). 1-[6-([(17β)-3-methoxyestra-1,3,5(10)-trien-17-yl]amino)hexyl]-1H-pyrrole-2,5-dione (U-73122), 1-[6-([(17β)-3-methoxyestra-1,3,5(10)-trien-17-yl]amino)hexyl]-2,5-pyrrolidinedione (U-73343), protease (Type X), collagenase (type 1A), soybean trypsin inhibitor (Type II-S), BSA, ATP (disodium salt), GTP (dilithium salt), creatine, HEPES, 1,3,7-trimethylxanthine (caffeine), dimethyl sulphoxide, paraformaldehyde and Triton X-100 were obtained from Sigma-Aldrich (Poole, UK). All other chemicals were from BDH Laboratory Supplies, AnalaR grade (Pool, UK).

Results

Dependence of [Ca²⁺]_i transients on P2X receptor agonist concentration

Fluo-4-loaded RVSMCs were stimulated with gradually increasing concentrations of the P2X receptor agonist αβ-meATP applied through a glass micropipette as a 2 s pulse (Figure 1A). αβ-meATP was applied at 20 min intervals, during which the application pipette was replaced with a new one containing a different concentration of the agonist. The same RVSMC was stimulated with at least three different agonist concentrations followed by application of 100 µmol·L⁻¹αβ-meATP. The intensity of fluo-4 fluorescence was normalized to that before agonist application and colour-coded. Traces of self-normalized fluo-4 fluorescence (F/F₀) reflect the dynamics of [Ca²⁺]_i changes induced by different αβ-meATP concentrations (Figure 1A; top left). Each of the four panels in Figure 1 (bottom) show 30 sequential images captured from the same RVSMC following application of 0.1, 0.25, 0.5 and 100 µmol·L⁻¹αβ-meATP (i–iv, respectively) during the periods highlighted in the traces. To quantify the relationship between the response amplitude and αβ-meATP concentration, averaged peaks of ΔF/F₀ transients were normalized to that induced by 100 µmol·L⁻¹αβ-meATP in the same cell and plotted versus αβ-meATP concentrations (Figure 1A; top right). A dose–response curve fitted to the data using a non-linear least-squares minimization algorithm revealed EC₅₀= 0.18 ± 0.01 µmol·L⁻¹ (n= 5–10). On the one hand, this value is more than six times lower than that which we have recently reported for P2X receptor-mediated current in RVSMCs (Harhun et al., 2010), suggesting that some amplification mechanism contributes to Ca²⁺ mobilization. On the other hand, a gradual increase in the amplitude and rate of rise of the [Ca²⁺]_i response with increasing αβ-meATP concentration suggests that either no regenerative mechanisms are involved or that these mechanisms are restricted to Ca²⁺ microdomains that function autonomously and are gradually recruited with increasing stimuli (Berridge, 1997). In support of the latter, localized Ca²⁺ events were observed following stimulation with low (≤0.5 µmol·L⁻¹) concentrations of αβ-meATP (Figure 1A; panels i–iii). In response to high (≥1 µmol·L⁻¹) concentrations of αβ-meATP, Ca²⁺ mobilisation was initiated by a sub-plasmalemmal [Ca²⁺]_i upstroke (SPCU) (Gordienko et al., 2008), an abrupt increase in [Ca²⁺]_i at multiple sub-plasmalemmal regions, which propagates through the entire cell volume within 300 ms (Figure 1A, panel iv and B, panels i–iii).

Figure 1.

(A) Dependence of [Ca²⁺]_i transients induced by P2X receptor stimulation in renal vascular smooth muscle cells (RVSMCs) on αβ-methylene ATP (αβ-meATP) concentration. Traces of self-normalized Fluo-4 fluorescence (F/F₀) averaged within confocal (<1.2 µm) slice of the cell reflect [Ca²⁺]_i transients induced by different αβ-meATP concentrations, as indicated (the drug applications are denoted by magenta bars above the traces, top left). Please note that the traces shown were obtained from two RVSMCs: cell 1 – the responses to 0.1, 0.25, 0.5 and 100 µmol·L⁻¹αβ-meATP and cell 2 – the responses to 0.01, 1.0 and 10 µmol·L⁻¹αβ-meATP. Averaged peaks of ΔF/F₀ transients normalized to that induced by 100 µmol·L⁻¹αβ-meATP in the same cell are plotted versus corresponding αβ-meATP concentrations (top right). Dose–response curve fitted to the data revealed EC₅₀= 0.18 ± 0.01 µmol·L⁻¹ (n = 5–10). The four panels (bottom) of 30 colour-coded confocal images were sequentially captured during the periods highlighted in the traces (top left), i–iv respectively. (B) Traces of self-normalized Fluo-4 fluorescence averaged within sub-plasmalemmal regions of initiations (insets) reflect high reproducibility of [Ca²⁺]_i transients induced by repetitive (with 20 min interval) applications of 10 µmol·L⁻¹αβ-meATP (top). The three panels (bottom) of 30 images were captured during the highlighted periods (top), i–iii respectively.

To compensate for cell-to-cell variations in Fluo-4 load and/or resting [Ca²⁺]_i and, thereby, to permit the comparison of findings between different RVSMCs, we evaluated the effect of different drugs on SPCU by relating the fluorescent response (ΔF/F₀) induced by 10 µmol·L⁻¹αβ-meATP in the presence of a drug to that obtained in control in the same myocyte. In these experiments, the same Fluo-4-loaded RVSMCs were stimulated with 2 s pulses of αβ-meATP applied with a 20 min interval. The response to the succeeding stimulation (Test) was related to the response to the first stimulation (Control). The test response was obtained either in control conditions (to evaluate the response reproducibility) or following 20 min incubation with a drug or drug combination. This experimental strategy was validated by high reproducibility of [Ca²⁺]_i transients induced in control conditions by 10 µmol·L⁻¹αβ-meATP applied at 20 min intervals (Figure 1B, top). On average, the reproducibility of the response amplitude was 99 ± 1% (n= 20) (Figure 2E, ‘Untreated’ and 7A). Comparing images in the panels also revealed a remarkable constancy of the positions of the sites of initiation of Ca²⁺ mobilization associated with each αβ-meATP application (Figure 1B, panels i–iii). This allowed us to analyse SPCU in all subsequent experiments.

Figure 2.

Stimulation of renal vascular smooth muscle cells (RVSMCs) with 10 µmol·L⁻¹αβ-methylene ATP (αβ-meATP) selectively activates P2X receptors. Traces (A–D) of relative changes in Fluo-4 fluorescence (ΔF/F₀) illustrate the experimental protocol. RVSMCs were stimulated with 2 s pulse of αβ-meATP applied twice at a 20 min interval. The peak amplitude of the response (ΔF/F₀) to the second stimulation (Test) was normalized to that of the response to the first stimulation (Control). The Test response was obtained either in the absence an antagonist [A, ‘Untreated’ in (E)] or following incubation with 2 µmol·L⁻¹ (B) or with 2 mmol·L⁻¹ (C) of the P2X receptor antagonist NF 279, or with 10 µmol·L⁻¹ of the P2Y receptor antagonist MRS 2578 (D). Statistical analysis (E) revealed that the αβ-meATP-induced responses were significantly reduced by NF 279 but were insensitive to MRS 2578. ***P < 0.001.

The ability of 10 µmol·L⁻¹αβ-meATP to selectively activate P2X receptors was confirmed in separate experiments (Figure 2), where the effects of the P2X receptor antagonist NF 279 and P2Y6 receptor antagonist MRS 2578 on αβ-meATP-induced [Ca²⁺]_i transients were tested. Of the eight recombinant P2Y receptor subtypes (P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13, and P2Y14), only four subtypes were found to be expressed in the kidney: P2Y1, P2Y2, P2Y4 and P2Y6 (reviewed in Guan et al., 2007b). Our RT-PCR analysis of the expression of the genes encoding P2Y receptors performed using single isolated RVSMCs (see Methods) revealed the expression of genes encoding only two P2Y receptor subtypes: P2Y2 and P2Y6 (data not shown). Although both of these P2Y receptor subtypes are uridine nucleotide-sensitive rather than adenine nucleotide-sensitive and should not be affected by 10 µmol·L⁻¹αβ-meATP used in this study, we still tested the effect of the P2Y6 receptor antagonist MRS 2578 (as there is no commercially available selective P2Y2 receptor antagonist) on αβ-meATP-induced [Ca²⁺]_i transients. We found that the peak amplitude of the response (ΔF/F₀) of Fluo-4-loaded RVSMCs to 10 µmol·L⁻¹αβ-meATP was reduced by 61 ± 3% (n= 12) following a 20 min incubation with 2 µmol·L⁻¹ NF 279 and by 98 ± 1% (n= 6) following incubation with 2 mmol·L⁻¹ NF 279, but was unaffected by 10 µmol·L⁻¹ MRS 2578 (n= 6; P= 0.29).

SPCU and plasmalemmal electrical events

Confocal imaging of [Ca²⁺]_i transients induced in RVSMCs by 10 µmol·L⁻¹αβ-meATP was combined with electrical recordings performed using the perforated patch technique (Gordienko et al., 2008). Under current clamp (Figure 3A), SPCU was associated with a spike-like depolarization, which reached a peak of −2.8 ± 1.0 mV within 90 ± 15 ms (n= 14). Only three out of 14 RVSMCs revealed an overshoot (3.2 ± 1.1 mV). This suggests that the αβ-meATP-induced depolarization is mainly caused by the P2X receptor-mediated rather than VGCC-mediated current. Concurrent elevation of [Ca²⁺]_i peaked within 792 ± 35 ms (n= 14) and persisted after recovery of the membrane potential to the resting level, thus, implying that mechanisms other than P2X-mediated Ca²⁺ entry contribute to the intracellular Ca²⁺ mobilization.

Figure 3.

Relationship between the sub-plasmalemmal [Ca²⁺]_i upstroke (SPCU) induced in renal vascular smooth muscle cells (RVSMCs) by 10 µmol·L⁻¹αβ-methylene ATP (αβ-meATP) and the plasmalemmal electrical events. Changes in the cell membrane potential (A) and transmembrane current at V_h=−60 mV (B) were recorded in Fluo-4-loaded RVSMC using the perforated-patch technique (A, top inset) simultaneously with confocal imaging of [Ca²⁺]_i changes. In (A) and (B): electrical recordings (top), traces of self-normalized Fluo-4 fluorescence averaged within sub-plasmalemmal regions of initiations (middle), gallery (bottom) of colour-coded (F/F₀) images sequentially captured during the highlighted (middle) periods.

When the same RVSMC was voltage-clamped (at −60 mV), P2X receptor activation did not induce depolarization of the cell membrane, thus preventing a contribution of VGCCs to the αβ-meATP-induced response. Under these conditions, the peak of the Fluo-4 response (ΔF/F₀) was reduced by 43% (Figure 3B). On average, the peak of the αβ-meATP-induced response (ΔF/F₀) detected under voltage-clamp conditions was reduced by 37 ± 4% (n= 14) relative to that detected under current-clamp conditions. This statistically significant reduction (P < 0.00008; paired t-test) suggests that Ca²⁺ entry through VGCCs contributes to the αβ-meATP-induced SPCU. Under voltage-clamp conditions P2X-mediated inward current reached a peak within 106 ± 18 ms, while concurrent SPCU peaked within 1508 ± 40 ms (n= 14). This discrepancy suggests that Ca²⁺ release from the SR is recruited following P2X receptor activation. Indeed, even though VGCCs were not activated under these conditions, the elevation of [Ca²⁺]_i continued to advance when the P2X-mediated current started to decline. Deceleration of the SPCU resulting from VGCC deactivation implies that the SR Ca²⁺ release can be speeded up by Ca²⁺ entry through VGCCs. Thus, under voltage-clamp conditions, when depolarization of the cell membrane by P2X receptor-mediated current does not occur, the link between P2X receptor activation and Ca²⁺ entry via VGCCs is lost and [Ca²⁺]_i mobilization evoked by P2X receptor activation is substantially distorted. Therefore, all subsequent experiments were conducted on non-clamped myocytes.

Contribution of voltage-gated Ca²⁺ entry and the SR Ca²⁺ release to SPCU

In these experiments, nicardipine was applied 30 s before αβ-meATP to minimize the possible effect of VGCC block on the SR calcium load. The ability of the external application of 5 µmol·L⁻¹ nicardipine to completely block VGCCs within 16 s was confirmed in separate voltage-clamp experiments (n= 5; data not shown). The effects of blocking VGCCs following calcium store depletion (Figure 4A) and calcium store depletion following block of VGCCs (Figure 4B) on SPCU induced by 10 µmol·L⁻¹αβ-meATP were tested. Incubation of RVSMCs for 20 min with 50 µmol·L⁻¹ cyclopiazonic acid (CPA), an inhibitor of the sarco-/endoplasmic reticulum Ca²⁺-ATPase (SERCA), which has no direct effect on VGCCs, Ca²⁺-dependent K⁺ channels or voltage-dependent K⁺ channels (Suzuki et al., 1992), completely depleted intracellular calcium stores in RVSMCs, as assessed with 5 mmol·L⁻¹ caffeine (n= 9; data not shown). The fraction of the calcium store depletion induced by a single application of 5 mmol·L⁻¹ caffeine was estimated in RVSMCs pre-incubated for 5 min with 10 µmol·L⁻¹ CPA (to inhibit SERCA without the calcium store depletion) and was found to be 94 ± 2% (n= 12; data not shown), thus confirming that 5 mmol·L⁻¹ is a sufficient concentration of caffeine to test the SR Ca²⁺ content in these cells. Depletion of calcium stores by a 20 min incubation with 50 µmol·L⁻¹ CPA reduced the peak of the αβ-meATP-induced response (ΔF/F₀) by 53%. Subsequent block of VGCCs with 5 µmol·L⁻¹ nicardipine (while keeping calcium stores depleted) resulted in an additional reduction of the peak response by 20% (Figure 4A). The peak response was attenuated by 44% following block of VGCCs alone, and was reduced further by 10% following subsequent calcium store depletion (Figure 4B). These observations suggest that both Ca²⁺ entry through VGCCs and the SR Ca²⁺ release contribute to the SPCU. The response persisting in the presence of both nicardipine and CPA arose from Ca²⁺ entry though P2X receptors and had a peak amplitude 29 ± 3% of that in control (n= 20).

Figure 4.

Ca²⁺ entry via voltage-gated Ca²⁺ channels (VGCCs) following P2X receptor activation in renal vascular smooth muscle cells (RVSMCs) induces Ca²⁺ release from intracellular calcium stores. Effect of (A) block of VGCCs (with 5 µmol·L⁻¹ nicardipine) following calcium store depletion (by 20 min incubation with 50 µmol·L⁻¹ cyclopiazonic acid, CPA) and (B) calcium store depletion following block of VGCCs on SPCU triggered by repetitive (with 20-min interval) applications of 10 µmol·L⁻¹αβ-methylene ATP (αβ-meATP). The panels below the traces of self-normalized Fluo-4 fluorescence show colour-coded (F/F₀) confocal images sequentially captured during the periods highlighted in the traces, i–iii respectively. (C) The amplitude of ΔF/F₀ transients detected following calcium store depletion (Test) was normalized to that detected before incubation with CPA (Control) before and following block of VGCCs (left) and compared (right). ***P < 0.001.

To assess the role of Ca²⁺ entry through VGCCs in the activation of the SR Ca²⁺ release, we compared the effect of calcium store depletion following block of VGCCs with that observed in the absence of nicardipine (Figure 4C). This experimental strategy (illustrated by traces obtained from two other RVSMCs) revealed that in control conditions calcium store depletion reduced the response (ΔF/F₀) to αβ-meATP by 53 ± 3% (n= 22), while when VGCCs were blocked, the calcium store depletion decreased the response by only 23 ± 5% (n= 8). Significant attenuation of the effect of the calcium store depletion on the SPCU following block of VGCCs implies that Ca²⁺ release from the SR following P2X receptor activation is induced mainly by Ca²⁺ entry through VGCCs.

Contribution of RyRs and IP₃Rs to the SR Ca²⁺ release following P2X receptor activation

Ryanodine receptors activity was assessed with 100 µmol·L⁻¹ ryanodine, a highly selective agent, which at micromolar concentrations blocks RyR-mediated Ca²⁺ release without calcium store depletion (Sutko et al., 1997; Janiak et al., 2001; Gordienko and Bolton, 2002) and does not affect IP₃Rs (Janiak et al., 2001) or VGCCs (Balke and Wier, 1991). IP₃R activity was assessed with 30 µmol·L⁻¹ 2-aminoethoxydiphenyl borate (2-APB). Because it was previously suggested that 2-APB may affect mechanisms of intracellular Ca²⁺ homeostasis other than those mediated by IP₃R, including inhibition of store-operated Ca²⁺ entry and SERCA (reviewed by Bootman et al., 2002), we tested the effect of 30 µmol·L⁻¹ 2-APB on the SR calcium load and Ca²⁺ entry mechanisms evoked by P2X receptor activation in RVSMCs (Figure 5).

Figure 5.

2-aminoethoxydiphenyl borate (2-APB) (30 µmol·L⁻¹) exerts no effect on (A) the P2X receptor-mediated cationic current (B) the Ca²⁺ entry mechanisms induced by P2X receptor activation and (C) the sarcoplasmic reticulum (SR) calcium load. Left panels (in A–C) illustrate experimental protocols. Renal vascular smooth muscle cells were stimulated with 2 s pulses of 10 µmol·L⁻¹αβ-methylene ATP (αβ-meATP) (A, B) or 5 mmol·L⁻¹ caffeine (C) applied twice from a glass micropipette at 10-min (A) or 20-min (B, C) intervals and the peak amplitude of the responses was measured: (A) current density and (B, C) relative changes in Fluo-4 fluorescence (ΔF/F₀). Depleting the calcium store by at least 20 min incubation with 50 µmol·L⁻¹ CPA unmasked Ca²⁺ entry induced by P2X receptor stimulation (B). The peak amplitude of the response (ΔF/F₀) to caffeine reflects the SR calcium load (C). The peak amplitude of the response to the second application of the agonist (Test) was normalized to that of the response to the first application of the agonist (Control) in each case (A–C). The Test response was obtained either in the absence of 2-APB (to asses reproducibility of the response to the agonist) or following incubation with 30 µmol·L⁻¹ 2-APB (to assess the effect of the drug). Statistical analysis (right panels in A–C), revealed no significant effect of 30 µmol·L⁻¹ 2-APB: P= 0.36 (A), P= 0.28 (B) and P= 0.45 (C).

P2X receptor-mediated current (Figure 5A) was unaffected by a 10 min incubation with 30 µmol·L⁻¹ 2-APB (n= 5). To unmask Ca²⁺ entry induced by P2X receptor stimulation in RVSMCs, the intracellular calcium stores were depleted by at least 20 min incubation with 50 µmol·L⁻¹ CPA prior to stimulation with 10 µmol·L⁻¹αβ-meATP (Figure 5B). Under these experimental conditions, αβ-meATP-induced [Ca²⁺]_i transients were insensitive to 30 µmol·L⁻¹ 2-APB (n= 9). The SR calcium load, assessed with 5 mmol·L⁻¹ caffeine (Figure 5C), was also unaffected by the 20 min incubation with 30 µmol·L⁻¹ 2-APB (n= 8). These observations validate 30 µmol·L⁻¹ 2-APB as a selective probe for IP₃R-mediated Ca²⁺ release in RVSMCs.

To analyse the contribution of RyRs and IP₃Rs to the SR Ca²⁺ release evoked by P2X receptor activation, the effects of successive cumulative inhibition of IP₃Rs after RyRs (Figure 6A) and RyRs after IP₃Rs (Figure 6B) on SPCU induced by 10 µmol·L⁻¹αβ-meATP were tested. Block of RyRs reduced the peak of the response (ΔF/F₀) to αβ-meATP by only 18%. Subsequent inhibition of IP₃Rs resulted in an additional reduction of the response by 57% (Figure 6A). The response was attenuated by 60% following inhibition of IP₃Rs alone, and reduced further by 8% following subsequent block of RyRs (Figure 6B).

Figure 6.

Relative contribution of ryanodine receptor (RyR)- and inositol 1,4,5-trisphosphate receptor (IP₃R)-mediated Ca²⁺ release to sub-plasmalemmal [Ca²⁺]_i upstroke (SPCU) induced by P2X receptor activation in renal vascular smooth muscle cells. Effect of successive cumulative inhibition of (A) IP₃Rs [with 30 µmol·L⁻¹ 2-aminoethoxydiphenyl borate (2-APB) ] after RyRs (with 100 µmol·L⁻¹ ryanodine) and (B) RyRs after IP₃Rs on SPCU triggered by repetitive applications of 10 µmol·L⁻¹αβ-methylene ATP (αβ-meATP). The cells were incubated with the Ca²⁺ release channel inhibitors for 20 min before the application of αβ-meATP. The panels below the traces of self-normalized Fluo-4 fluorescence show colour-coded confocal images sequentially captured during the periods highlighted in the traces, i–iii respectively. (C) The peak amplitude of the response to the second application of the agonist (Test) was normalized to that of the response to the first application of the agonist (Control). The Test response was obtained either in the presence of ryanodine alone (to assess possible effect of ryanodine on the calcium store load; left: top panel) or in the presence of both 2-APB and ryanodine (to asses the cumulative effect of the drugs; left: middle and bottom panels). The amplitudes of ΔF/F₀ transients detected in the presence of the two inhibitors were normalized to that detected in the presence of one of them and in both cases were compared (right). ***P < 0.001.

An important condition of the above strategy is the ability of 100 µmol·L⁻¹ ryanodine to block RyRs without calcium store depletion. This was tested in separate experiments. In these experiments (Figure 6C, left: top panel), after 20 min incubation with 100 µmol·L⁻¹ ryanodine the RVSMCs were stimulated with 10 µmol·L⁻¹αβ-meATP twice. The αβ-meATP pulses were applied at 20 min intervals. The amplitude of the [Ca²⁺]_i transient induced by the second αβ-meATP application (Test) was related to that induced by the first one (Control). On average, the reproducibility of the amplitude of the response to 10 µmol·L⁻¹αβ-meATP in the presence of 100 µmol·L⁻¹ ryanodine was found to be 98 ± 5% (n= 6) (Figure 6, right); this was not statistically different (P= 0.778) from that observed in RVSMCs not treated with ryanodine, 99 ± 1% (n= 20) (Figure 2E). This observation indicates that, with the experimental protocol used in this study, 100 µmol·L⁻¹ ryanodine blocks RyR-mediated Ca²⁺ release without depleting the calcium store.

Ryanodine receptors and IP₃Rs are both Ca²⁺-sensitive and may activate each other via CICR mechanism (Fill and Copello, 2002; Foskett et al., 2007). Thus, by blocking one receptor type the effect of CICR on the other receptor type will also be lost. To eliminate the effect of possible cross-activation of these receptors from the analysis, the responses (ΔF/F₀) to αβ-meATP in the presence of both blockers were normalized to the responses in the presence of each blocker alone and compared (Figure 6C). This revealed that when RyRs were initially blocked, inhibition of IP₃Rs reduced the response by 61 ± 4% (n= 7). In contrast, when IP₃Rs were initially blocked, inhibition of RyRs reduced the response by only 20 ± 3% (n= 5). This suggests that Ca²⁺ release from the SR following P2X receptor activation is mediated mainly by IP₃Rs.

Figure 7 summarizes the effects of VGCC/SERCA/RyR/IP₃R/PLC inhibitors on the SPCU induced in RVSMCs by 10 µmol·L⁻¹αβ-meATP. The analysis strategy was the same as described above. The response to the succeeding stimulation (Test) was related to the response to the first stimulation (Control). The test response was obtained either in control conditions (to evaluate the response reproducibility) or following 20 min incubation with a drug or drug combination. Figure 7A shows sample traces illustrating this experimental protocol. Relative changes in the response peak amplitude (Figure 7B) and time-to-peak (Figure 7C) were analysed. In control conditions (‘Untreated’), the amplitude and time-to-peak of the test response constituted 99 ± 1% and 109 ± 5% of the control response respectively (n= 20). Block of RyRs (100 µmol·L⁻¹ ryanodine) reduced the amplitude by only 18 ± 4% (n= 7) and had no significant effect on time-to-peak (P= 0.08). The effect of IP₃R inhibition (30 µmol·L⁻¹ 2-APB) was significantly stronger: reduction of the amplitude by 48 ± 6% and increase of the time-to-peak by 98 ± 15% (n= 10). The effect of calcium store depletion (20 min incubation with 50 µmol·L⁻¹ CPA) was also significantly stronger than that of RyR inhibition: reduction of the amplitude by 53 ± 3% and increase of the time-to-peak by 51 ± 10% (n= 22). The effect of simultaneous block of RyRs and IP₃Rs was similar to the effect of calcium store depletion: reduction of the amplitude by 60 ± 4% and increase of the time-to-peak by 48 ± 11% (n= 12). Block of VGCCs (5 µmol·L⁻¹ nicardipne) reduced the amplitude by 39 ± 2% and increased time-to-peak by 81 ± 11% (n= 18). Block of VGCCs following calcium store depletion reduced the amplitude by 70 ± 3% and decreased time-to-peak by 21 ± 8% (n= 20). It should be noted that calcium store depletion (with CPA) reduced the αβ-meATP-induced response significantly (P < 0.0006) more than block of VGCCs, suggesting that Ca²⁺ release from the SR is partially induced by Ca²⁺ entering the cell via P2X receptors. Calcium store depletion with simultaneous block of VGCCs reduced the response significantly (P < 0.0002) more than calcium store depletion alone, suggesting that Ca²⁺ entry via VGCCs makes a significant contribution to the αβ-meATP-induced Ca²⁺ mobilization. Comparison of mean amplitudes of the responses (ΔF/F₀) to αβ-meATP in RVSMCS with depleted calcium stores obtained in the absence and in the presence of nicardipine suggests that the Ca²⁺ influx via P2X receptors is ∼1.8 times larger than that via VGCCs. Thus, Ca²⁺ influx following P2X receptor activation induces Ca²⁺ release from the SR, which is mediated mainly (but not solely) via IP₃Rs and accelerates intracellular Ca²⁺ mobilization.

Figure 7.

Summary of the effects of voltage-gated Ca²⁺ channel/sarco-/endoplasmic reticulum Ca²⁺-ATPase/ryanodine receptor/inositol 1,4,5-trisphosphate receptor/phospholipase C inhibitors on sub-plasmalemmal [Ca²⁺]_i upstroke in renal vascular smooth muscle cells (RVSMCs). Experimental protocol is illustrated in (A). Fluo-4 loaded RVSMCs were stimulated with 2 s pulses of 10 µmol·L⁻¹αβ-methylene ATP (αβ-meATP) applied with a 20 min interval. The response (ΔF/F₀ averaged at sub-plasmalemmal regions of initiation) to the second stimulation (Test) was related to the response to the first stimulation (Control). The Test response was obtained either in control conditions (‘Untreated’) or following incubation with a drug or drug combination. Two parameters were examined and summarized: (B) relative change in peak amplitude and (C) relative change in time-to-peak. *P < 0.05, **P < 0.01 and ***P < 0.001 versus control, unless denoted otherwise. 2-APB, 2-aminoethoxydiphenyl borate; CPA, cyclopiazonic acid.

While P2X receptors do not signal via phospholipase C-β (PLC-β), basal levels of [IP₃]_i produced by spontaneous activity of PLC (Prestwich and Bolton, 1991; Gordienko and Bolton, 2002; Horowitz et al., 2005; Peng et al., 2007) may play a permissive role in activation of IP₃R-mediated Ca²⁺ release by voltage-gated Ca²⁺ entry. Although the rates of hydrolysis by G_q-activated PLC-β are accelerated by elevation of [Ca²⁺]_i, the results of experiments on permeabilized cells have suggested that [Ca²⁺]_i levels achieved during cell signalling do not activate PLC-β by themselves but are required for PLC-β activation by G_q (Fisher et al., 1989; Horowitz et al., 2005). Thus, it is unlikely thatαβ-meATP-induced [Ca²⁺]_i mobilization could per se activate PLC-β. Activation of any G_q/11-coupled receptors in RVSMCs by 10 µmol·L⁻¹αβ-meATP is also unlikely, because the αβ-meATP-induced [Ca²⁺]_i transients were completely abolished by selective P2X antagonist NF 279, but were unaffected by P2Y antagonist MRS 2578 (see Figure 2). Nevertheless, we tested the effect of commercially available PLC inhibitors on the αβ-meATP-induced [Ca²⁺]_i transients. The aminosteroid compound U-73122 and its relatively inactive analogue U-73343 (Smith et al., 1990) both significantly reduced the amplitude and increased the time-to-peak of the response (Figure 7B and C). However, the effect of 2.5 µmol·L⁻¹ U-73122 (the reduction of the amplitude by 44 ± 7% and increase of the time-to-peak by 99 ± 22%; n= 6) was significantly stronger than that of 2.5 µmol·L⁻¹ U-73343 (the reduction of the amplitude by 22 ± 3% and increase of the time-to-peak by 32 ± 15%; n= 5). Although these compounds are known to have many side effects attributable to alkylation of various proteins (Horowitz et al., 2005), the difference between the effect of U-73122 and that of U-73343 suggests a permissive role of IP₃ in activation of IP₃Rs by Ca²⁺ in RVSMCs. To test this further, we examined the effect of another PLC inhibitor, the ether lipid analogue edelfosine (Powis et al., 1992; Horowitz et al., 2005). Pre-incubation with 10 µmol·L⁻¹ edelfosine (Figure 7) reduced the amplitude of the αβ-meATP-induced [Ca²⁺]_i transients by 40 ± 4% and increased time-to-peak by 82 ± 11% (n= 6). We therefore concluded that the ability of Ca²⁺ entering the RVSMCs following P2X receptor stimulation to induce IP₃R-ediated Ca²⁺ release depends on [IP₃]_i which is determined by spontaneous basal activity of PLC.

To assess the role of Ca²⁺ entry through VGCCs in IP₃R activation, we compared the effect of IP₃R inhibition following block of VGCCs with that observed in the absence of nicardipine (Figure 8). This experimental strategy was validated by high reproducibility of the responses to 10 µmol·L⁻¹αβ-meATP in the presence of 30 µmol·L⁻¹ 2-APB (Figure 8A). The effect of IP₃R inhibition on the SPCU was attenuated by block of VGCCs (Figure 8B). To quantify this, the responses (ΔF/F₀) in the presence of 2-APB were normalized to the responses before IP₃R inhibition, both in control and in the presence of nicardipine, and compared (Figure 8C). This revealed that, when VGCCs were initially blocked, IP₃R inhibition decreases the response by only 16 ± 4% (n= 12), while in control conditions (without pretreatment with nicardipine) IP₃R inhibition reduced the response (ΔF/F₀) to αβ-meATP by 48 ± 6% (n= 10). Significant attenuation of the effect of IP₃R inhibition on SPCU by block of VGCCs implies that IP₃R-mediated Ca²⁺ release is induced mainly (but not solely; see above) by Ca²⁺ influx via VGCCs.

Figure 8.

Relative contribution of P2X receptor- and voltage-gated Ca²⁺ channel (VGCC)-mediated Ca²⁺ entry to activation of inositol 1,4,5-trisphosphate receptor (IP₃R)-mediated Ca²⁺ release in renal vascular smooth muscle cells. Effect of IP₃R inhibition (A) and IP₃R inhibition after block of VGCCs (B) on sub-plasmalemmal [Ca²⁺]_i upstroke triggered by repeated applications of 10 µmol·L⁻¹αβ-methylene ATP (αβ-meATP). The panels below the traces of self-normalized Fluo-4 fluorescence show colour-coded (F/F₀) confocal images sequentially captured during the periods highlighted in the traces, i–iii respectively. (C) The amplitude of ΔF/F₀ transients detected following IP₃R inhibition (Test) was normalized to that detected before incubation with 2-aminoethoxydiphenyl borate (2-APB) (Control), both without (left, top panel) and in the presence of nicardipine (left, bottom panel), and compared (right). **P < 0.01.

Sub-plasmalemmal SR is enriched with IP₃Rs

Tight functional coupling between VGCCs and IP₃Rs suggests the expression of IP₃Rs in jSR. Intracellular calcium stores, visualized in RVSMCs with the low-affinity Ca²⁺ indicator Fluo-3FF, coincided with SR elements, visualized with ER-Tracker Blue-White or Brefeldin A BODIPY (n= 27), and consisted of some central elements and a sub-plasmalemmal SR network (Figure 9A). Similar organization of the SR has been reported in other SMC types (Gordienko et al., 2001; 2008; Gordienko and Zholos, 2004). RT-PCR analysis of single RVSMCs revealed the expression of genes encoding IP₃R type 1 and RyR type 2 (Figure 9B). Immunodetection of IP₃Rs and RyRs in RVSMCs stained with Brefeldin A BODIPY and DAPI (Figure 9C) revealed that type 1 IP₃Rs are primarily expressed in sub-plasmalemmal SR elements (n= 21), while RyRs predominate in deeper SR, particularly in the perinuclear region of the myocyte (n= 16). Based on this distribution of IP₃Rs and RyRs, Ca²⁺ responses to stimuli selectively targeting IP₃Rs and RyRs are expected to initiate from different positions within the cell. In the same RVSMC loaded with Fluo-4 and stained with ER-Tracker (Figure 9D), the sites of initiation of αβ-meATP-induced response coincide with positions of subplasmalemmal SR elements, while the site of initiation of the response to caffeine (which selectively activates RyR-mediated Ca²⁺ release) coincides with the position of deeper central SR elements (n= 8), as evident from a comparison of the images number 2 in the panels (Figure 9D, bottom) showing sequential images captured during the initial phase of the response to αβ-meATP (left) and caffeine (middle). Furthermore, flash release of IP₃ (which selectively activates IP₃Rs) within the entire cell volume (Figure 9D) initiated Ca²⁺ release at the cell periphery (n= 5), similar to P2X receptor stimulation.

Figure 9.

Spatial distribution of the sarcoplasmic reticulum (SR) Ca²⁺ release channels in renal vascular smooth muscle cells (RVSMCs). (A) Confocal images of the ER-tracker Blue-White, Fluo-3FF and Brefeldin A BODIPY fluorescence, and their overlay illustrate spatial organization of the SR in RVSMC. (B) RT-PCR analysis of inositol 1,4,5-trisphosphate receptor (IP₃R) and ryanodine receptor (RyR) expression in RVSMCs. Primers designed to amplify genes encoding IP₃R and RyR subtypes: IP₃R1 (806 bp), IP₃R3 (966 bp), RyR1 (793 bp), RyR2 (824 bp), RyR3 (1086 bp) were tested for their specificity using cDNA preparations of rat brain and liver tissue (top) before using them on RVSMC preparations (bottom). RVSMCs were found to express genes encoding IP₃R1 and RyR2. (C) Immunodetection of IP₃R1 (top) and RyR (middle) in RVSMC stained with Brefeldin A BODIPY and DAPI. Primary antibody binding was detected with MFP488-conjugated secondary antibody, which shows no non-specific binding in the absence of primary antibody (bottom). Confocal images highlight localization of IP₃R1 in sub-plasmalemmal SR and RyR in deeper SR. (D) RVSMCs were either pre-incubated with Fluo-4AM and ER-tracker Blue-White (pseudocolour confocal images) and stimulated sequentially with 10 µmol·L⁻¹αβ-methylene ATP (αβ-meATP) (left) or 5 mmol·L⁻¹ caffeine (middle), or loaded with Fluo-3 and ‘caged’ IP₃ via patch pipette (transmitted light and confocal images) and stimulated by whole-cell UV flash (right). Galleries of five sequential images (labelled 1–5, bottom) highlight spatial patterns of the initial phase of the responses.

Discussion

The nature of mechanisms coupling excitation to [Ca²⁺]_i elevation, as well as the spatial organization and molecular composition of intracellular Ca²⁺ release units are important determinants of the contractile response in all types of muscles (Berridge, 1997; Wier and Balke, 1999; Lamb, 2002; Blatter et al., 2003; Wang et al., 2004; McCarron et al., 2006). In skeletal muscles, excitation triggers Ca²⁺ release from the SR via protein–protein interactions between VGCCs and RyRs (Lamb, 2002). In cardiac muscles, Ca²⁺ entry through VGCCs triggers RyR-mediated Ca²⁺ release via CICR mechanism (Wier and Balke, 1999; Blatter et al., 2003; Wang et al., 2004). In both cases, the structural basis for E–C coupling is close juxtaposition of the SR RyRs and VGCCs. In smooth muscles, elevation of [Ca²⁺]_i is caused either by cell membrane depolarization (electromechanical coupling; Somlyo and Somlyo, 1968) leading to opening of VGCCs, which are manifested by the generation of action potentials or allow Ca²⁺ to ‘leak’ into the cell giving rise to ‘sparklets’ (Amberg et al., 2007), or by activation of metabotropic receptors (pharmacomechanical coupling; Somlyo and Somlyo, 1968) usually linked to G_q/11-GTP/PLC/IP₃/IP₃R-mediated Ca²⁺ release, or by a combination of these mechanisms (Bolton et al., 1999; Davis and Hill, 1999). The initial Ca²⁺ mobilization may be augmented by RyR-mediated Ca²⁺ release recruited via a CICR mechanism (Bolton and Gordienko, 1998; Bayguinov et al., 2000; White and McGeown, 2002; Kotlikoff, 2003; Zhang et al., 2003; Hotta et al., 2007; Gordienko et al., 2008). The contribution of these mechanisms to intracellular Ca²⁺ mobilization may vary in different SMC types, depending on the type and strength of stimuli, SMC excitability and spatial organisation of intracellular Ca²⁺ release units (Walsh, 1994; Bolton and Gordienko, 1998; Bolton et al., 1999; Davis and Hill, 1999; Bayguinov et al., 2000; Gordienko et al., 2001, 2008; Janiak et al., 2001; Gordienko and Bolton, 2002; White and McGeown, 2002; Kotlikoff, 2003; Wier and Morgan, 2003; Zhang et al., 2003; Lamont and Wier, 2004; McCarron et al., 2006; Amberg et al., 2007; Hotta et al., 2007; Wier et al., 2009).

We hypothesized that in vascular myocytes Ca²⁺ entry following activation of ionotropic P2X receptors may induce IP₃R-mediated Ca²⁺ release. Our hypothesis was based on the following findings reported in visceral and vascular myocytes: (i) a localized increase in [Ca²⁺]_i induced by local flash release of Ca²⁺ from a ‘caged’ precursor may trigger IP₃R-mediated Ca²⁺ release (Ji et al., 2006); (ii) Ca²⁺ entry via VGCCs facilitates IP₃R-mediated Ca²⁺ release following stimulation of metabotropic receptors (Gordienko et al., 2008); (iii) the stoichiometric ratio of RyRs to IP₃Rs in SMCs is 1:9–10, suggesting the existence of a Ca²⁺-storage compartment devoid of RyRs but equipped with IP₃Rs (Wibo and Godfraind, 1994); (iv) caffeine/ryanodine-releasable and IP₃-releasable calcium stores in the SR of vascular myocytes are at least partially distinct with about 56% of the SR being solely IP₃-sensitive (Blaustein et al., 2002); and (v) visualization of Ca²⁺ release sites recruited by selective IP₃R stimulation and IP₃R immunodetection in single SMCs suggests that they are predominantly expressed in sub-plasmalemmal SR (Bayguinov et al., 2000; Zhang et al., 2003; Gordienko and Zholos, 2004; Gordienko et al., 2008).

It should be noted, however, that there is no general agreement about the calcium store arrangement in SMCs. Indeed, the results of early experiments conducted on permeabilized strips of portal vein, pulmonary artery and taenia caeci suggested the existence of two separate calcium stores: S_α expressing both RyRs and IP₃Rs and S_β expressing only IP₃Rs. The contribution of the S_β store to IP₃-induced Ca²⁺ release was estimated to be 60% in taenia caecum, 40% in pulmonary artery and only 6% in portal vein (reviewed in Wray and Burdyga, 2010). In contrast, the results of more recent experiments conducted on isolated pulmonary artery myocytes suggested that IP₃-releasable and caffeine-releasable calcium stores are virtually completely separate (Janiak et al., 2001). A similar conclusion was made from the experiments performed on in intact precapillary vessels, where agonist-induced [Ca²⁺]_i oscillations were insensitive to the combined action of caffeine and ryanodine (Borisova et al., 2009). An elegant study recently conducted by Rainbow et al. (2009) on guinea-pig hepatic portal vein myocytes provided strong evidence for the existence of a single luminally continuous SR where the opening of IP₃Rs and RyRs is regulated by luminal [Ca²⁺], so that when the luminal [Ca²⁺] declines, the receptor opening also declines and stops. This model therefore suggests that the appearance of multiple stores in some functional studies may arise from the regulation of luminal [Ca²⁺] by RyRs and IP₃Rs expressed in a single luminally continuous SR.

We tested our hypothesis that Ca²⁺ entry following P2X activation may induce IP₃R-mediated Ca²⁺ release on RVSMCs, because: (i) P2X receptors mediate sympathetic control and autoregulation of the renal circulation (Malpas and Leonard, 2000; Nishiyama et al., 2000; Bell et al., 2003; Inscho et al., 2003; Cupples and Braam, 2007; Guan et al., 2007a; Surprenant and North, 2009); and (ii) RVSMCs express functional monomeric P2X1 and heteromeric P2X1/4 receptors (Harhun et al., 2010).

We found that depolarization of RVSMC induced by selective stimulation of P2X receptors with αβ-meATP triggers Ca²⁺ release from sub-plasmalemmal SR enriched with IP₃Rs but poor in RyRs, similar to direct stimulation of IP₃Rs by uniform cell-wide release of IP₃ from its ‘caged’ precursor. The αβ-meATP-induced SPCU was significantly reduced by block of VGCCs or depletion of the intracellular calcium stores, indicating that activation of P2X receptors in RVSMCs recruits both voltage-gated Ca²⁺ entry and the SR Ca²⁺ release to intracellular Ca²⁺ mobilization. Activation of voltage-gated Ca²⁺ influx through L-type Ca²⁺ channels following P2X receptor stimulation was previously demonstrated in preglomerular microvascular SMCs of rat kidney (White et al., 2001). However, the involvement of the SR Ca²⁺ release mechanisms was not addressed in that study. Also, [Ca²⁺]_i transients induced by 10 µmol·L⁻¹αβ-meATP in preglomerular microvascular SMCs were completely abolished by 20 µmol·L⁻¹ NF 279, while in RVSMCs from interlobar and arcuate arteries 2 mmol·L⁻¹ NF 279 reduced the [Ca²⁺]_i transients by 98%. This discrepancy may arise from variations in the distribution of P2X receptor subtypes in SMCs from different regions of renal vascular bed and may reflect the contribution of heteromeric P2X1/4 and monomeric P2X4 receptors to intracellular Ca²⁺ mobilization in RVSMCs from interlobar and arcuate arteries (Harhun et al., 2010). Here we not only demonstrated that the SR Ca²⁺ release is recruited following P2X receptor activation, but also provided strong evidence suggesting that this release is mediated mainly by IP₃Rs. Indeed, depletion of calcium stores, block of VGCCs or IP₃Rs reduced the amplitude and rate of rise of αβ-meATP-induced SPCU significantly more than block of RyRs. This implies that Ca²⁺ entry following P2X receptor activation induces IP₃R-mediated Ca²⁺ release which serves to accelerate [Ca²⁺]_i elevation and is initiated at sub-plasmalemmal SR enriched with IP₃Rs but poor in RyRs.

It is unlikely that stimulation of RVSMCs with 10 µmol·L⁻¹αβ-meATP recruited any metabotropic receptors coupled to a G_q/11/PLC/IP₃ signalling pathway, because the αβ-meATP-induced [Ca²⁺]_i transients were completely abolished by the selective P2X antagonist NF 279, but were unaffected by the P2Y antagonist MRS 2578. Even though activation of PLC-β by G_q is accelerated at [Ca²⁺]_i levels achieved during cell signalling, direct activation of PLC-β by Ca²⁺ is also unlikely (Fisher et al., 1989; Horowitz et al., 2005). Nevertheless, the effect of PLC-β inhibition with either U-73122 or edelfosine on αβ-meATP-induced [Ca²⁺]_i transients in RVSMCs was not statistically different from that of IP₃R inhibition with 2-APB. This suggests that basal levels of [IP₃]_i produced by spontaneous activity of PLC (Prestwich and Bolton, 1991; Gordienko and Bolton, 2002; Horowitz et al., 2005; Peng et al., 2007) may play a permissive role in the activation of IP₃R-mediated Ca²⁺ release by elevated [Ca²⁺]_i achieved following P2X receptor activation. It should be noted, however, that U-73122 is known for its side effects attributable to alkylation of various proteins (Horowitz et al., 2005), while edelfosine appears to be cytotoxic causing cell swelling and eventual lysis, and therefore may well have its own side effects (Horowitz et al., 2005). Indeed, out of the16 RVSMCs tested, edelfosine caused lysis of four myocytes and induced substantial elevation of [Ca²⁺]_i in five cells.

A comparison of the effect of the calcium store depletion with the cumulative effect of the calcium store depletion and block of VGCCs revealed that, following stimulation of RVSMCs with 10 µmol·L⁻¹αβ-meATP, P2X receptors contributed more than VGCCs to Ca²⁺ entry. This is consistent with the high density of P2X receptor-mediated currents in RVSMCs (Harhun et al., 2010) and high Ca²⁺ permeability of P2X1 and P2X4 receptors: a relative Ca²⁺ over Na⁺ permeability (P_Ca/P_Na) of 4.8 and 4.2 was reported for P2X1 and P2X4 receptors respectively (North, 2002; Egan et al., 2006). Nevertheless, IP₃R-mediated Ca²⁺ release was activated mainly by Ca²⁺ entering the cell via VGCCs. Indeed, the effect of IP₃R inhibition on the SPCU was reduced by threefold following block of VGCCs. This suggests the co-localization of plasmalemmal VGCCs and the jSR IP₃Rs, and ‘local control’ (Berridge, 1997; McCarron et al., 2006) of Ca²⁺ release mechanisms in RVSMCs. The latter is also supported by the gradual dependence of [Ca²⁺]_i transients on αβ-meATP concentration, despite the fact that a regenerative CICR mechanism was recruited (Arendshorst and Thai, 2009). The validation of this hypothesis, however, requires a separate electron microscopy study of ‘Ca²⁺ release units’ (Moore et al., 2004) in RVSMCs.

It is now generally accepted that jSR (which lies within 12–15 nm of plasmalemma), the overlying plasmalemma microdomains and the intervening, tiny volume of cytosol form junctional complexes that serve as the Ca²⁺‘buffer barrier’, through which Ca²⁺ can move directly between the extracellular fluid and jSR (Blaustein et al., 2002; Poburko et al., 2008). This may facilitate (i) local elevation of luminal [Ca²⁺] in jSR; and (ii) accumulation of molecules, including Ca²⁺, Na⁺ (Blaustein et al., 2002; Poburko et al., 2008), and IP₃, in the cytosolic microvolume from which the diffusion into the bulk cytoplasm is markedly restricted (Poburko et al., 2008). Taking this into account, the following factors may favour activation of IP₃Rs by Ca²⁺ influx: (i) a robust increase in [Ca²⁺]_i in the junctional cytosolic microvolume (Blaustein et al., 2002; Poburko et al., 2008); (ii) spontaneous basal activity of PLC (Prestwich and Bolton, 1991; Gordienko and Bolton, 2002; Horowitz et al., 2005; Peng et al., 2007); (iii) Ca²⁺ activation of type 1 IP₃Rs with positive co-operativity (Foskett et al., 2007); (iv) regulation of IP₃Rs by the SR luminal [Ca²⁺] (Rainbow et al., 2009); and (v) IP₃R clustering (Rahman and Taylor, 2009).

It should be noted, however, that the precise evaluation of the relative contribution of Ca²⁺ entry (P2X receptors and VGCCs) and Ca²⁺ release (IP₃Rs and RyRs) mechanisms to αβ-meATP-induced [Ca²⁺]_i transients is limited by: (i) a non-linear relationship between the intensity of the indicator fluorescence and [Ca²⁺]_I; and (ii) the possible effects of pharmacological agents targeting the SR Ca²⁺ release mechanisms (either via the SR calcium load or via inhibition of IP₃Rs/RyRs) on the cell membrane potential and, hence, the population of VGCCs available for activation by P2X receptor-mediated depolarization and/or the initial driving force for Ca²⁺ entry. The ability of calcium store modulators (CPA, ryanodine and 2-APB) to alter the cell membrane potential is usually attributed to their effect on spontaneous RyR- and/or IP₃R-mediated Ca²⁺ release events, which modulate the cell membrane potential via activation of Ca²⁺-dependent K⁺ channels and/or Ca²⁺-dependent Cl^- channels (Burdyga and Wray, 1999; Gordienko et al., 1999; Bayguinov et al., 2000; Hashitani and Brading, 2003; Hashitani et al., 2006; Zizzo et al., 2006; von der Weid et al., 2008; ZhuGe et al., 1998; 2010;). Since spontaneous Ca²⁺ release events were observed in less than 5% of RVSMCs used in this study, it is rather unlikely that inhibition of the SR Ca²⁺ release by ryanodine, 2-APB or CPA had any significant effect on resting membrane potential in these myocytes. Also, there is no evidence in the literature for a direct effect (which can be evaluated only from the single channel activity in isolated patches) of the calcium store modulators on potassium and/or chloride channels. On the contrary, it was demonstrated that neither ryanodine (Sakai et al., 1988) nor CPA (Suzuki et al., 1992) had any direct effect on Ca²⁺-dependent K⁺ channels in SMCs. In this study we have demonstrated that in RVSMCs 2-APB exerts no effect on the Ca²⁺ entry mechanisms induced by P2X receptor activation and the SR calcium load (Figure 5). Because the effect of 2-APB on αβ-meATP-induced [Ca²⁺]_i transient was significantly attenuated in the presence of nicardipine, we concluded that in RVSMCs IP₃R-mediated Ca²⁺ release following P2X receptor stimulation is activated mainly by Ca²⁺ entering the cell via VGCCs.

Although activation of the SR Ca²⁺ release by Ca²⁺ entering the cell via VGCCs has been demonstrated in voltage-clamp experiments performed on different types of visceral and vascular SMCs (Kamishima and McCarron, 1997; Kohda et al., 1997; Bolton and Gordienko, 1998; Shmigol et al., 1998; Coussin et al., 2000), there is a number of studies (e.g. Bradley et al., 2002; 2004;) demonstrating that complete depletion of the SR of Ca²⁺ (including IP₃-sensitive store) does not reduce [Ca²⁺]_i transients induced by step-like depolarization of the cell membrane. The latter results suggest that CICR is not recruited. An alternative explanation given by Bradley et al. (2004) suggests that the SR and sarcolemma may form a passive physical barrier to Ca²⁺ influx (‘a Ca²⁺ trap’), which normally limits the [Ca²⁺]_i rise occurring during depolarization. The drugs, which open the SR Ca²⁺ release channels and facilitate the SR Ca²⁺ leak, diminish the influence of ‘the Ca²⁺ trap’ and may, thereby, increase the amplitude of [Ca²⁺]_i transients resulting from Ca²⁺ entry via VGCCs even when the SR contains little or no Ca²⁺ (Bradley et al., 2004).

In cardiac myocytes, abnormal intracellular Ca²⁺ handling arising from altered expression/function of RyRs and/or IP₃Rs promotes arrhythmias, atrial fibrillation and heart failure (George and Lai, 2007; Kockskämper et al., 2008). In vascular myocytes, mechanisms coupling P2X receptor activation to IP₃R-mediated Ca²⁺ release may be altered in hypertension. Indeed, it was demonstrated that impaired Ca²⁺ signalling attenuates P2X receptor-mediated vasoconstriction of afferent arterioles in angiotensin II hypertension (Zhao et al., 2005). On the other hand, elevated PLC-β levels and activity were reported in renal arterioles of young spontaneously hypertensive rats (Peng et al., 2007) and were suggested to account for previously demonstrated exaggerated renal vasoconstriction and Ca²⁺ signalling in preglomerular arterioles induced by angiotensin II, thromboxane A₂ and vasopressin in spontaneously hypertensive versus normotensive rats (Feng and Arendshorst, 1997; Iversen and Arendshorst, 1999; Vagnes et al., 2005).

To conclude, we demonstrated that P2X receptor activation evokes IP₃R-mediated Ca²⁺ release in RVSMCs, which is induced mainly by voltage-gated Ca²⁺ entry. This mechanism provides convergence of signalling pathways engaged in electromechanical and pharmacomechanical coupling in smooth muscles of renal vasculature and should be taken into consideration when developing new therapeutic strategies. Further work, however, is needed to examine whether and how this mechanism is altered in hypertension.

Glossary

Abbreviations

2-APB

2-aminoethoxydiphenyl borate

αβ-meATP

αβ-methylene ATP

CICR

Ca²⁺-induced Ca²⁺ release

CPA

cyclopiazonic acid

IP₃

inositol 1,4,5-trisphosphate

IP₃R

inositol 1,4,5-trisphosphate receptor

jSR

sub-plasmalemmal (‘junctional’) sarcoplasmic reticulum

MRS2578

N,N''-1,4-butanediylbis[N'-(3-isothiocyanatophenyl)thio urea

NF279

8,8'-[carbonylbis(imino-4,1-phenylenecarbonylimino-4,1-phenylenecarbonyl-imino)] bis(1,3,5-naphthalene-trisulphonic acid)

PLC

phospholipase C

RBF

renal blood flow

RSNA

renal sympathetic nerve activity

RT-PCR

reverse transcription polymerase chain reaction

RVSMC

renal vascular smooth muscle cells

RyR

ryanodine receptor

SERCA

sarco-/endoplasmic reticulum Ca²⁺-ATPase

SPCU

sub-plasmalemmal [Ca²⁺]_i upstroke

SR

sarcoplasmic reticulum

U-73122

1-[6-([(17β)-3-methoxyestra-1,3,5(10)-trien-17-yl]amino)hexyl]-1H-pyrrole-2,5-dione

U-73343

1-[6-([(17β)-3-methoxyestra-1,3,5(10)-trien-17-yl]amino)hexyl]-2,5-pyrrolidinedione

edelfosine

(7R)-4-hydroxy-7-methoxy-N,N,N-trimethyl-3,5,9-trioxa-4 -phosphaheptacosan-1-aminium-4-oxide

VGCC

voltage-gated Ca²⁺ channel

Conflicts of interest

None.

Supplementary material

Supporting Information: Teaching Materials; Figs 1–9 as PowerPoint slide.