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Ca_v1.3 channels produce persistent calcium sparklets, but Ca_v1.2 channels are responsible for sparklets in mouse arterial smooth muscle

¹Departments of Physiology and Biophysics and ²Pharmacology, University of Washington, Seattle, Washington; and ³Department of Pharmacology and Toxicology, Institute of Pharmacy, University of Innsbruck, Innsbruck, Austria

Submitted 12 April 2007 ; accepted in final form 21 May 2007

	ABSTRACT

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Ca²⁺ sparklets are local elevations in intracellular Ca²⁺ produced by the opening of a single or a cluster of L-type Ca²⁺ channels. In arterial myocytes, Ca²⁺ sparklets regulate local and global intracellular Ca²⁺. At present, the molecular identity of the L-type Ca²⁺ channels underlying Ca²⁺ sparklets in these cells is undetermined. Here, we tested the hypotheses that voltage-gated calcium channel- 1.3 subunit (Ca_v1.3) can produce Ca²⁺ sparklets and that Ca_v1.2 and/or Ca_v1.3 channels are responsible for Ca²⁺ sparklets in mouse arterial myocytes. First, we investigated the functional properties of single Ca_v1.3 channels in tsA201 cells. With 110 mM Ba²⁺ as the charge carrier, Ca_v1.3 channels had a conductance of 20 pS. This value is similar to that of Ca_v1.2 and native L-type Ca²⁺ channels. As previously shown for Ca_v1.2 channels, Ca_v1.3 channels can operate in two gating modes characterized by short and long open times. Expressed Ca_v1.3 channels also produced Ca²⁺ sparklets. Ca_v1.3 sparklets had properties similar to those produced by Ca_v1.2 and native L-type channels, including quantal amplitude, dihydropyridine sensitivity, bimodal gating, and dual-event duration times. However, the voltage dependencies of conductance and steady-state inactivation of the Ca²⁺ current (I_Ca) in arterial myocytes were similar to those recorded from cells expressing Ca_v1.2 but not Ca_v1.3 channels. Furthermore, nifedipine (10 µM) eliminated Ca²⁺ sparklets in wild-type myocytes but not in myocytes expressing dihydropyridine-insensitive Ca_v1.2 channels. Accordingly, Ca_v1.3 transcript and protein were not detected in isolated arterial myocytes. We conclude that although Ca_v1.3 channels can produce Ca²⁺ sparklets, Ca_v1.2 channels underlie I_Ca, Ca²⁺ sparklets, and hence dihydropyridine-sensitive Ca²⁺ influx in mouse arterial myocytes.

voltage-gated calcium channel-1C subunit; voltage-gated channel-1D subunit; total internal reflection fluorescence microscopy; L-type calcium channels

The channels underlying Ca²⁺ sparklets in arterial smooth muscle meet the general criteria used to identify L-type Ca²⁺ channels (22): Ca²⁺ sparklets are activated by the dihydropyridine agonist BAY K 8644, are eliminated by the dihydropyridine antagonist nifedipine, and have similar voltage dependencies (activity and amplitude) to L-type Ca²⁺ channels. Consistent with this, voltage-dependent Ca_v1.2 L-type Ca²⁺ channels are expressed in arterial smooth muscle (26). We recently suggested that Ca²⁺ sparklets produced by heterologously expressed Ca_v1.2 channels resembled native Ca²⁺ sparklets in arterial myocytes (35). Thus, according to an indirect comparative approach, Ca_v1.2 channels could, at least in principle, underlie Ca²⁺ sparklets in arterial myocytes. At present, however, whether or not Ca_v1.2 channels underlie Ca²⁺ sparklets in arterial myocytes remains to be tested directly.

A striking feature of persistent Ca²⁺ sparklets in arterial myocytes is that they are observed at hyperpolarized membrane potentials (e.g., –70 mV), a voltage where the open probability (P_o) of L-type Ca²⁺ channels is very low (10^–6–10^–8) (41, 43). This raises the intriguing possibility that another L-type Ca²⁺ channel with a voltage dependence of activation more negative than Ca_v1.2 channels underlies Ca²⁺ sparklets in arterial myocytes. Ca_v1.3 L-type Ca²⁺ channels meet this condition, because their threshold for activation is 10–20 mV more negative than that of Ca_v1.2 channels (27, 52). Furthermore, Ca_v1.3 transcript and protein have been detected in canine basilar artery (37). These observations raise four important questions: 1) What are the biophysical properties of single Ca_v1.3 channels? 2) Are Ca_v1.3 channels capable of producing Ca²⁺ sparklet activity? 3) If so, are Ca_v1.3 sparklets similar to Ca²⁺ sparklets in mouse cerebral arterial myocytes? 4) Are Ca_v1.3 channels expressed in mouse cerebral arterial myocytes, and do they contribute to Ca²⁺ influx in these cells?

The goal of this study was to address these four fundamental questions. Beyond the obvious relevance of determining whether Ca_v1.3 channels contribute to Ca²⁺ influx in mouse arterial myocytes (question 4), by addressing questions 1, 2, and 3 we provide what to our knowledge is the first single-channel analysis of heterologously expressed Ca_v1.3 channels and their ability to transport Ca²⁺. Given the importance of Ca_v1.3 channels in nerve (5) and muscle (37, 55), addressing these issues would impact multiple fields.

	METHODS

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Isolation of arterial myocytes. We used wild-type mice (C57BL/6J) and mice genetically engineered to express Ca_v1.2 channels that are insensitive to dihydropyridines (DHP^–/–; C57BL/6J background) (44). Mice were euthanized with a lethal dose of sodium pentobarbital (250 mg/kg ip) as approved by the University of Washington Institutional Animal Care and Use Committee. Myocytes were dissociated from cerebral arteries by using standard enzymatic techniques described elsewhere (4). After dissociation, cells were maintained in a nominally Ca²⁺-free Ringer solution until they were used. Thapsigargin (1 µM) was included in all solutions used to record Ca²⁺ sparklets to eliminate Ca²⁺ release from intracellular stores during experimentation.

Heterologous expression of Ca_v1.3 and PKC in tsA201 cells. Cultures of tsA201 cells were maintained in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum, 2 mM L-glutamine, and a 1% streptomycin and penicillin solution. Unitary Ca_v1.3 channel activity (Fig. 1) was recorded from tsA201 cells transiently transfected with the pcDNA clones of Ca_v1.3 (GenBank accession no. AF37009), Ca_v3 (GenBank accession no. M88751), Ca_v21 (GenBank accession no. AF286488; a generous gift from Dr. Dianne Lipscombe), and the enhanced green fluorescent protein (EGFP; kindly provided by Dr. John Exton) by using Lipofectamine 2000. Ca_v1.3 sparklets and whole cell, macroscopic Ca_v1.2 and Ca_v1.3 currents were recorded from tsA201 cells expressing Ca_v1.2 (GenBank accession no. AY728090) or Ca_v1.3 and Ca_v3, Ca_v21, and PKC tagged with EGFP. Successfully transfected cells were identified on the basis of EGFP fluorescence.

View larger version (37K): [in this window] [in a new window]   Fig. 1. Unitary Cav1.3 channel currents. A: representative single Cav1.3 channel records obtained during step depolarization (1 s) to voltages ranging from –40 to +20 mV with 110 mM Ba2+ as the charge carrier. Arrow marks 0-current level (i.e., closed channel). B: Cav1.3 channel open probability (Po)-voltage relationship (n = 6 cells). Solid line represents best-fit curve to data determined by a least-squares method using a Boltzmann equation: y = [(A1- A2)/ 1+e(V–V1/2/k)], where A1 (0.008), A2 (0.686), V1/2 (–2.4 mV), and k (6.4) are the initial value, final value, the voltage at which 50% of the current or conductance was observed, and the slope factor. C: current-voltage relationship of unitary Cav1.3 currents (n = 6 cells). Solid line represents best-fit curve to data using a linear equation. Mean unitary Cav1.3 currents for different membrane potentials studied are –1.65 pA (–40 mV), –1.47 pA (–30 mV), –1.27 pA (–20 mV), –1.07 pA (–10 mV), –0.85 pA (0 mV), –0.63 pA (+10 mV), and –0.43 pA (+20 mV). D: open-time histogram of single Cav1.3 channel at –10 mV (n = 2,806 events). Solid line represents best-fit curve to data using a log-normal function: y=A·e(–{[ln(x) –ln(s)]2})/2s2+ B·e(–{[ln(x) –ln(l)]2})/2l2, where A and B are constants, s (1.6 ms; 67%) is the short time constant, l (9.5 ms; 33%) is the long time constant, and s (0.7 ms) and l (0.6 ms) are standard deviations of s and l, respectively. Dashed lines are individual fits for short and long opening. Sqrt, square root.

To obtain the current-voltage relationship of Ca²⁺ currents, cells were depolarized for 200 ms from the holding potential of –70 mV to voltages ranging from –70 to +70 mV. For the voltage dependence of activation, peak I_Ca currents at test potentials between –40 and +50 mV were converted into conductances (G = I_Ca/[test pulse potential – reversal potential of I_Ca]), normalized (G/G_Max), and plotted as a function of test potential. We used a standard two-test pulse to obtain the voltage dependence of steady-state inactivation of Ca²⁺ currents. Briefly, cells were submitted to prepulses (E_pre) from –40 to +30 mV for 2 s, after which they were depolarized to a test potential (E_test) of +20 mV for 200 ms.

Single-channel recordings of Ca_v1.3 currents were obtained in the cell-attached configuration of the patch-clamp technique. The bath solution contained (in mM) 145 KCl, 2 MgCl₂, 0.1 CaCl₂, 10 HEPES, and 10 glucose adjusted to pH 7.3 with KOH to set the membrane resting potential to 0 mV. The pipette solution contained (in mM) 110 BaCl₂ and 10 HEPES adjusted to pH 7.2 with CsOH. Pipette electrodes were pulled from thin-wall borosilicate glass capillaries (Sutter Instruments, Novato, CA) and were fire polished to resistances between 10 and 15 M when filled with the pipette solution. Single-channel currents were amplified, low-pass filtered at 5 kHz, and sampled at 10 kHz with the Axopatch 200B amplifier using a DigiData 1322A acquisition board and pClamp 9.0 software and were stored directly on a PC hard drive. The data were filtered with a Gaussian filter (500 Hz) during analysis. Currents were elicited by stepping pulses from a holding potential of –80 mV to indicated test potentials for 1 s. Capacitative currents were compensated by subtraction of blank sweeps. All experiments were performed at room temperature (22–25°C).

Single-channel events were detected with the threshold detection algorithm of pClamp 9. To determine unitary conductance of Ca_v1.3 channels, single-channel amplitude was measured at voltages ranging from –40 to +20 mV. Single-channel conductance was calculated as the slope of the current-voltage relationship. The open-time histogram was constructed by using pClamp 9. The voltage of –10 mV was selected for open-time analysis on the basis of sufficient channel activity and ability to clearly resolve channel openings and closings.

Arterial wall [Ca²⁺]_i was imaged by using a Radiance 2100 confocal system (Cambridge, MA) coupled to a Nikon TE300 inverted microscope equipped with a Nikon x20 (numerical aperture = 0.75) lens. Small mesenteric arteries [types II and III (46)] from wild-type and DHP^–/– mice were removed and cleaned in normal Ringer solution. Isolated arteries were cannulated and mounted in a close-working-distance arteriograph, and the endothelium was disrupted by passing air bubbles through the artery. The arteriograph was placed on the stage of the inverted microscope and was extraluminally perfused (3–6 ml/min) at 37°C with a bicarbonate-based physiological saline solution composed of (in mM) 119 NaCl, 4.7 KCl, 24 NaHCO₃, 1.2 KH₂PO₄, 1.6 CaCl₂, 1.2 MgSO₄, and 11 glucose with the pH set to 7.4 by bubbling with a gas mixture of O₂ (95%) and CO₂ (5%). After equilibration (20 min), intravascular pressure was increased to 80 mmHg for 20 min. Arteries were loaded with the Ca²⁺ indicator fluo-4 as described elsewhere (2). Arterial viability was tested by raising external K⁺ to 60 mM; arteries that failed to contract robustly in response to high K⁺ concentrations were discarded.

Fluo-4, fluo-5F, and rhod-2 fluorescence signals were converted to Ca²⁺ concentration units by using the "F_max" equation (32)

as described in detail previously (13, 35). Briefly, F is fluorescence, F_max is the fluorescence intensity of fluo-4/5F or rhod-2 in the presence of saturating free Ca²⁺, K_d is the dissociation constant (fluo-5F = 1,280 nM; fluo-4 = 800 nM; rhod-2 = 700 nM), and R_f is the indicator's F_max/F_min ratio (fluo-5F = 286; fluo-4 = 150; rhod-2 = 125). F_min is the fluorescence intensity of the indicator in a solution where the Ca²⁺ concentration is 0. K_d and R_f values were determined in vitro by using standard methods and are similar to those reported by others (49). F_max was determined at the end of each experiment by exposing cells or arteries to the Ca²⁺ ionophore ionomycin (10 µM) and 20 mM external Ca²⁺.

Total internal reflection fluorescence microscopy. Ca²⁺ sparklets were recorded at –70 mV as previously described (34, 35). Briefly, we used a through-the-lens total internal reflection fluorescence (TIRF) microscope built around an inverted Olympus IX-70 microscope equipped with an Olympus PlanApo (x60, numerical apperture = 1.45) oil-immersion lens and an Andor iXON charge-coupled device camera (South Windsor, CT). To monitor [Ca²⁺]_i, cells were loaded with the calcium indicator fluo-5F or rhod-2. Rhod-2 was used in all experiments in which the EGFP was expressed. Excitation of fluo-5F or rhod-2 was achieved with the 488 or 568 nm line of an argon or krypton laser, respectively (Dynamic Lasers). Images were acquired at 100–300 Hz. As before (34, 35), we determined the activity of Ca²⁺ sparklets by calculating the nP_s of each Ca²⁺-sparklet site, where n is the number of quantal levels and P_s is the probability that a quantal Ca²⁺-sparklet event is active. Using this analysis, we have grouped Ca²⁺-sparklet sites into three categories: silent (by default has an nP_s of 0), low (nP_s between 0 and 0.2), and high (nP_s > 0.2). A detailed description of this analysis can be found in Navedo et al. (35).

Ca²⁺-sparklet signal mass and duration analysis. We used the "signal mass" approach developed by Zou et al. (56, 57) to determine the amount of Ca²⁺ flux (Q_Ca, measured in coulombs) associated with Ca²⁺ sparklets at –70 mV as described in Amberg et al. (2). Briefly, for this analysis the total fluorescence intensity (F_total) associated with a Ca²⁺ sparklet is determined from raw images by summing the fluorescence from all the pixels within an area of the image larger than the entire fluorescence signal produced by a Ca²⁺ sparklet. The change in F_total (F_total) is then determined by subtracting the total fluorescence before the channel(s) open from the total fluorescence at each time point during the opening. Signal mass was obtained by determining the peak of the integral of F_total trace over time (F_total dt) for each Ca²⁺ sparklet. The relationship between sparklet signal mass and Q_Ca is linear (2, 47, 56, 57). Thus the slope (555 peak F_total dt units·fC^–1, where fC^–1 is inverse fentocoulombs) of a previously obtained signal mass-Q_Ca relationship for an L-type Ca²⁺ channel (2) was used to calculate the Q_Ca of Ca_v1.3 sparklets from F_total dt values.

Ca²⁺-sparklet event-duration times at the membrane potential of –70 mV were obtained from fits to Ca²⁺-sparklet records by using pClamp 9 "threshold detection analysis" as described in detail elsewhere (2). This analysis is similar to the one implemented by the Parker group for the study of Ca²⁺ signals via single N-type Ca²⁺ channels (11) and acetylcholine receptors (12).

RNA isolation and RT-PCR. Total RNA was isolated from 60 arterial myocytes by using the RNeasy Micro kit (Qiagen, Valencia, CA) as instructed by the manufacturer. We designed primers specific for the Ca_v1.2 subunit (GenBank accession no. NM_009781; sense nt 5292–5316 and antisense nt 5884–5910; amplicon = 618 bp) and Ca_v1.3 subunit (GenBank accession no. NM_028981; sense nt 4589–4613 and antisense nt 5065–5090; amplicon = 501 bp) of this channel. We used -actin (GenBank accession no. V01217; sense nt 2384–2404 and antisense nt 3071–3091; amplicon = 496 bp) transcript levels as an internal control for these experiments. -Actin primers amplify a region between exons 4 and 6 such that genomic contamination within the RNA preparation is identified by the presence of a 708-bp band in addition to the 496-bp band. Reverse transcription and amplification was performed by using the OneStep RT-PCR kit from Qiagen following the manufacturer's instructions. To do this, we used a Eppendorf thermal cycler running the following program: 35 cycles at 94°C for 1 min, 56°C for 1 min, and 72°C for 1 min/cycle, with a final extension step of 72°C for 10 min. Amplicons were separated by using 2% agarose gel electrophoresis.

Immunofluorescence. Isolated arterial myocytes were plated on laminin-coated coverslips and were allowed to settle for 2 h. Cells were then fixed with 4% paraformaldehyde for 45 min, rinsed in 0.1 M phosphate buffer, rinsed in 0.1 M Tris buffer (TB) for 15 min, rinsed in 0.1 M Tris-buffered saline (TBS) for 15 min, blocked in 2% avidin in TBS for 30 min, rinsed in TBS for 30 min, blocked in 2% biotin for 30 min, and then rinsed in TBS for 30 min. Cells were then incubated in the anti-Ca_v1.2 (chicken, diluted 1:50) or anti-Ca_v1.3 (rabbit, diluted 1:50) antibodies overnight at 4°C (48). Cells were rinsed and incubated in biotinylated goat anti-chicken IgG (diluted 1:300; Vector) or biotinylated goat anti-rabbit IgG for 2 h at room temperature, rinsed, and then incubated in avidin D fluorescein (diluted 1:300; Vector) for 2 h at room temperature, rinsed in TBS, rinsed in TB, and then coverslipped. As a control, the primary antibody was omitted in some experiments.

Chemicals and statistics. Gö-6976 and ionomycin were from Calbiochem (San Diego, CA); Dulbecco's modified Eagle's medium was from GIBCO. Lipofectamine 2000 was from Invitrogen. All other chemicals were from Sigma-Aldrich (St. Louis, MO) unless otherwise stated. Normally distributed data are presented as means ± SE. Two-sample comparisons were made by using a Student's t-test; multigroup comparisons were made by ANOVA, which, if necessary, was followed by Tukey's multicomparison test. Nonparametric statistical analyses (Mann-Whitney test) were used for nonnormally distributed data. P values <0.05 were considered significant. Asterisks used in the figures indicate a significant difference between groups.

	RESULTS

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Properties of single Ca_v1.3 channels. We recorded single Ca_v1.3-channel activity in tsA201 cells (Fig. 1). For these experiments, the pore-forming Ca_v1.3 subunit was coexpressed with accessory Ca_v3 and Ca_v21 subunits and EGFP. Transfected cells were identified on the basis of EGFP fluorescence. Ca_v1.3-channel currents were recorded in cell-attached patches, with 110 mM Ba²⁺ in the pipette solution used as the charge carrier.

Figure 1A shows a representative family of single Ca_v1.3-channel sweeps during step depolarization (1 s) from a holding potential of –80 mV to voltages ranging from –40 to +20 mV. Note that membrane depolarization increased the P_o of Ca_v1.3 channels and that the threshold for the activation of these channels was –40 mV. Similar findings were obtained in four additional independent experiments. Indeed, the P_o of Ca_v1.3 channels was 0.003 ± 0.001 and 0.686 ± 0.006 at –40 and +20 mV, respectively (n = 5; Fig. 1B). The voltage dependence of Ca_v1.3 channels was fitted (R² = 0.99) with a Boltzmann function with a maximum P_o of 0.69 ± 0.006 at +20 mV, the voltage at which 50% of the maximum P_o was observed (V_1/2) of –2.4 ± 0.2 mV, and a slope factor of 6.4 ± 0.2.

The voltage dependence of unitary Ca_v1.3-current amplitudes is shown in Fig. 1C. At –40, –10, and +20 mV, the amplitude of single Ca_v1.3-channel currents was 1.65 ± 0.04, 1.07 ± 0.02, and 0.63 ± 0.04 pA, respectively (n = 6). These data were fit with a linear function that revealed a slope conductance of 20 ± 1 pS (n = 6; R² = 0.99).

We also analyzed the open times of single Ca_v1.3-channel openings at –10 mV (Fig. 1D). The Ca_v1.3-channel open-times histogram at this voltage was fit with the sum of two log-normal probability density functions (PDFs) with a short (_s) and long (_l) time constant of 1.6 (standard deviation, _s = 0.7 ms; 67%) and 9.5 ms (standard deviation, _l = 0.6 ms; 33%), respectively (²= 1.7; n = 2,806 events). This analysis was validated by using Akaike's Information Criterion, which determines the probability that a data set could be described by a particular set of competing models (1). Indeed, this test revealed that the probability that the Ca_v1.3 open-time data described above could be fit by the sum of two log-normal PDFs was >99.99% but was <0.01% with a single log-normal PDF. Together, these data suggest that Ca_v1.3 channels, like Ca_v1.2 channels (21), could operate in two gating modes with short and long open times.

Ca_v1.3 channels produce Ca²⁺ sparklets. Having examined the function of single Ca_v1.3 channels, we investigated the mechanisms of Ca²⁺ influx through Ca_v1.3 channels by imaging near-membrane [Ca²⁺]_i in tsA201 cells expressing Ca_v1.3 channels with the use of TIRF microscopy. We and others (11, 12, 34, 35) have used this approach to image Ca²⁺ influx via single Ca²⁺ channels with high temporal and spatial resolution. Because PKC was necessary for high-activity persistent Ca²⁺ sparklets with Ca_v1.2 channels (35), we coexpressed PKC-EGFP (PKC) with Ca_v1.3 for these experiments. Expression of functional Ca_v1.3 channels in tsA201 cells was verified by the application of a brief (200 ms) voltage step from the holding potential of –70 mV to +10 mV. As illustrated in Fig. 2A, application of this voltage protocol evoked robust whole cell I_Ca. I_Ca was never observed in tsA201 cells expressing PKC-EGFP only.

View larger version (23K): [in this window] [in a new window]   Fig. 2. Cav1.3 sparklets in the heterologous expression system. A: representative Ca2+ current (ICa) records from tsA201 cells transfected with protein kinase C (PKC) or Cav1.3 (plus Cav3 and Cav21) and PKC. ICa was evoked by a test pulse from –70 to +10 mV, with 20 mM Ca2+ as charge carrier. B: total internal reflection fluorescence (TIRF) image from a tsA201 cell expressing Cav1.3 and PKC {extracellular Ca2+concentration ([Ca2+]o) = 20 mM and –70 mV}. Traces at right and bottom show time courses of intracellular Ca2+ concentration ([Ca2+]i) in region within green circle before (control) and after application of a nominal Ca2+-free solution, respectively. C: time courses of [Ca2+]i from a Cav1.3-sparklet site under control conditions and after application of 10 µM nifedipine. D: mean percent decrease in nPs (where n is number of quantal levels and Ps is probability that a quantal Ca2+-sparklet event is active) of Cav1.3 sparklets after application of 10 µM nifedipine or 0 mM [Ca2+]o. Application of nifedipine decreased Cav1.3-sparklet activity (i.e., nPs) by 34 ± 9% (n = 8 sparklet sites from 4 cells; P < 0.05). E: amplitude histogram of Cav1.3 sparklets ([Ca2+]o = 20 mM; n = 339 events). Black line is best fit to Cav1.3 sparklets with a multicomponent Gaussian function where a and b are constants and q (34 nM) is quantal unit of Ca2+ influx. F: voltage dependence of quantal Cav1.3-sparklet amplitude (n = 25 events/point). Solid red line is a linear fit to data. Inset: representative quantal Cav1.3 sparklets at –90 and –50 mV. Dotted lines mark quantal level for Cav1.3 sparklets at –90 and –50 mV.

Accordingly, we performed analyses on Ca_v1.3 sparklets developed for detecting and quantifying the activity, amplitude, and duration of Ca²⁺ sparklets in arterial myocytes. Detailed descriptions of these analyses can be found in METHODS and elsewhere (2, 34, 35). Ca²⁺-sparklet activity was quantified by determining the nP_s of Ca_v1.3-sparklet sites under control conditions and after the application of a Ca²⁺-free solution or a solution containing 20 mM Ca²⁺ and 10 µM nifedipine (Fig. 2, B–D). As expected, Ca_v1.3 sparklets were completely eliminated by the application of a Ca²⁺-free external solution, confirming that they resulted from Ca²⁺ entry, presumably via Ca_v1.3 channels (Fig. 2, B and D).

Next, we recorded Ca_v1.3 sparklets (at –70 mV) before and after application of 10 µM nifedipine. Previous studies suggest that Ca_v1.3 channels are less sensitive (9-fold) to dihydropyridine blockers than Ca_v1.2 channels (20, 27, 33, 52). Accordingly, we found that 10 µM nifedipine decreased Ca_v1.3-sparklet activity (i.e., nP_s) by 34 ± 9% (n = 8 sparklet sites from 4 cells; P < 0.05; Fig. 2, C and D). Although 10 µM nifedipine partially inhibited Ca_v1.3-sparklet activity at –70 mV, this concentration of nifedipine completely eliminated Ca_v1.2 sparklets in tsA201 cells at the same membrane potential (35). It is important to note that the 34 ± 9% decrease in Ca_v1.3-sparklet activity by 10 µM nifedipine observed here is consistent with a recent study (33) suggesting that a similar concentration of this dihydropyridine decreased Ca_v1.3 currents in outer hair cells by 36% (holding potential = –90 mV).

Figure 2E shows an amplitude histogram of Ca_v1.3 sparklets at –70 mV with 20 mM external Ca²⁺ concentration ([Ca²⁺]_o). As for Ca_v1.2 channels and native arterial myocyte sparklets under similar experimental conditions (34, 35), this histogram was fit with a multi-Gaussian function with a quantal unit of Ca²⁺ influx of 34 nM (n = 339 events; ² = 1.5). This value is similar to that of Ca_v1.2 (36 nM) and arterial myocyte Ca²⁺ sparklets (38 nM) under similar experimental conditions (34, 35). Thus, as in arterial myocytes and tsA201 cells expressing Ca_v1.2 channels, our analysis indicates that Ca²⁺ entry via Ca_v1.3 channels is quantal in nature and that multi-amplitude Ca_v1.3 sparklets result from coincidental activation of multiple quantal units.

We also determined the voltage dependence of the amplitude of quantal Ca_v1.3 sparklets (Fig. 2F). As expected from the single Ca_v1.3-channel data described above, the amplitude of Ca_v1.3 sparklets decreased with membrane depolarization as the driving force for Ca²⁺ influx decreased. Indeed, the Ca_v1.3 sparklet amplitude-voltage relationship was fit with a linear function with a slope of 0.44 nM/mV (R² = 0.99).

As with Ca_v1.2 and arterial smooth muscle Ca²⁺ sparklets under similar conditions, Ca_v1.3-sparklet activity was bimodal (Fig. 3, A and B). Ca_v1.3 sparklets could be divided into low and high nP_s sites. On average, the nP_s for low and high Ca_v1.3-sparklet sites were 0.07 ± 0.01 (n = 37) and 0.46 ± 0.1 (n = 20), respectively (Fig. 3B). Interestingly, these nP_s values are similar to those for Ca_v1.2 and arterial myocyte Ca²⁺ sparklets (34, 35). Application of the dihydropyridine agonist BAY K 8644 (500 nM) evoked Ca²⁺-sparklet activity in previously silent sites and increased the activity of low nP_s sites from 0.06 ± 0.01 (n = 45) to 0.31 ± 0.1 (n = 35), a value similar to that of high nP_s sites. BAY K 8644 did not increase the activity of high nP_s Ca_v1.3-sparklet sites, indicating that these channels were maximally activated (Fig. 3, A and B).

View larger version (16K): [in this window] [in a new window]   Fig. 3. BAY K 8644 increases Cav1.3-sparklet activity. A: representative silent, low nPs, and high nPs Cav1.3 sparklet sites before and after application of BAY K 8644 ([Ca2+]o = 20 mM). B: means ± SE of silent, low nPs, and high nPs sites before and after 500 nM BAY K 8644 (n = 6 cells).

View larger version (15K): [in this window] [in a new window]   Fig. 4. Analysis of duration and signal mass of Cav1.3 sparklets. A: scatter plot of signal mass of Cav1.3-sparklet events from low (n = 22 events) and high nPs (n = 35 events) sparklets. Red lines represent median values. Histograms of duration of low (B;n = 86 events) and high (C; n = 100 events) nPs Cav1.3-sparklet events. Solid black line in low nPs histograms is a fit to data with a single exponential function: y = A·e(–x/t) + yi, where A is amplitude (0.51), is the time constant (31 ms), and yi is the y intercept (0.008). Solid black line in high nPs histograms is a fit to data for a double-exponential function: y = A1·e(–x/t1)+ A2·e(–x/t2)+ yi, where A1 is amplitude of the first component (0.35), A2 is amplitude of the second component (0.04), 1 is the fast time constant (37 ms), 2 is the slow time constant (126 ms), and yi is the y intercept (0.002). Insets in B and C show representative Cav1.3 sparklets from low and high nPs sites.

Ca_v1.2 channels underlie I_Ca in mouse arterial myocytes. After demonstrating that Ca_v1.3 channels can produce Ca²⁺ sparklets similar in amplitude, activity, and duration to those observed for Ca_v1.2 and arterial myocyte Ca²⁺ sparklets, we investigated whether these channels contribute to I_Ca in cerebral arterial myocytes. We recorded Ca²⁺ currents with 20 mM external Ca²⁺ from cerebral arterial myocytes and tsA201 cells expressing Ca_v1.2 or Ca_v1.3 channels (Fig. 5). Ca²⁺ currents were evoked by 200-ms voltage pulses from –70 mV to potentials ranging from –70 to +70 mV. As reported by others (27, 52), Ca_v1.3 currents activated at more negative potentials than Ca_v1.2 currents (Fig. 5, A and B). Similar to that of Ca_v1.2 currents, the current-voltage relationship of I_Ca in cerebral arterial myocytes was shifted toward more depolarized potentials. The voltage at which 50% of I_Ca was observed (V_1/2) in cerebral arterial myocytes and tsA201 cells expressing Ca_v1.2 was +33.8 ± 1.2 mV (n = 7) and +31.9 ± 1.2 mV (n = 7), respectively, whereas the V_1/2 for tsA201 cells expressing Ca_v1.3 was +9.7 ± 2.9 mV (n = 8; P < 0.05).

View larger version (38K): [in this window] [in a new window]   Fig. 5. Cav1.2 channels underlie ICa in arterial myocytes. A: ICa records (at –20, 0, +20, +40, and +60 mV) from arterial myocytes and tsA201 cells expressing Cav1.2 or Cav1.3 channels. ICa were evoked by 200-ms depolarization steps from holding potential of –70mV to voltages ranging from –70 to +70 mV. B: current-voltage relationships of ICa from arterial myocytes (n = 7 cells) and tsA201 cells expressing Cav1.2 (n = 7 cells) or Cav1.3 channels (n = 8 cells). C: ICa steady-state inactivation records (at –30, 0, and +30 mV) from arterial myocytes (n = 8 cells) and tsA201 cells expressing Cav1.2 (n = 6 cells) or Cav1.3 channels (n = 6 cells). Cells were submitted to prepulses (Epre) from –40 to +30 mV for 2 s, after which cells were depolarized to a test potential (Etest) of +20 mV for 200 ms. Inset: diagrammatic representation of voltage protocol used in these experiments. D: ICa steady-state inactivation is shown as a plot of normalized peak current (I/IMax) as a function of conditioning potential. For voltage dependence of activation, peak ICa currents at test potentials between –40 and +50 mV were converted into conductance (G = ICa/[test pulse potential – reversal potential of ICa]), normalized (G/GMax), and plotted as a function of test potential. Smooth lines represent best-fit curves to data determined by a least-squares method using a Boltzmann equation similar to one used in Fig. 1. E: summary of parameters used to fit conductance and steady-state inactivation relationships of ICa in arterial myocytes and tsA201 cells expressing Cav1.3 and Cav1.2 channels. SMC, smooth muscle cell. *P < 0.05.

Ca_v1.2 channels, not Ca_v1.3 channels, underlie Ca²⁺ sparklets and control dihydropyridine-sensitive [Ca²⁺]_i in mouse arterial myocytes. With the data presented above, we cannot rule out that low-level expression of Ca_v1.3 channels in arterial myocytes is sufficient to contribute to Ca²⁺-sparklet activity without significantly influencing I_Ca. To address this issue, we recorded Ca²⁺ sparklets in arterial myocytes from wild-type and DHP^–/– mice. These DHP^–/– Ca_v1.2 channels have a single-point mutation (Thr¹⁰⁶⁶Tyr) that renders them insensitive to inhibition by dihydropyridines without otherwise altering their functional properties (27, 44). DHP^–/– mice are useful because dihydropyridine L-type Ca²⁺ channel antagonists can be used to selectively block Ca_v1.3 (relative to Ca_v1.2) channels and to determine their contribution to Ca²⁺-sparklet activity. If, as our data suggest, Ca_v1.2 and not Ca_v1.3 channels underlie Ca²⁺ sparklets in arterial myocytes, then we expect that nifedipine would eliminate Ca²⁺ sparklets in wild-type but not in DHP^–/– cells.

Figure 6A shows Ca²⁺-sparklet records (at –70 mV) from wild-type and DHP^–/– arterial myocytes before and after the application of 10 µM nifedipine. Note that nifedipine eliminated Ca²⁺ sparklets in wild-type but not in DHP^–/– cells (Fig. 6, A–C). This is consistent with the hypothesis that Ca_v1.2 and not Ca_v1.3 channels underlie Ca²⁺ sparklets in mouse arterial myocytes. To provide further support to this hypothesis, we examined arterial wall [Ca²⁺]_i in pressurized (80 mmHg) intact mesenteric arterial segments from wild-type and DHP^–/– mice. As expected (25), application of 10 µM nifedipine decreased wild-type arterial wall [Ca²⁺]_i from 210 to 112 nM [Ca²⁺]_i. Application of a Ca²⁺-free solution did not produce any further decrease in [Ca²⁺]_i. Consistent with our Ca²⁺-sparklet data, and unlike in wild-type arteries, we found that 10 µM nifedipine did not decrease [Ca²⁺]_i in pressurized DHP^–/– arteries. Application of a Ca²⁺-free solution, however, decreased arterial wall [Ca²⁺]_i in DHP^–/– arteries to a similar extent (100 ± 5 nM; Fig. 6D) to the decrease caused by nifedipine in wild-type arteries (210 to 107 nM; Fig. 6, D and E).

View larger version (29K): [in this window] [in a new window]   Fig. 6. Cav1.2 channels underlie Ca2+ sparklets in mouse arterial myocytes. A: representative time courses of [Ca2+]i from Ca2+-sparklet sites recorded from wild-type and dihydropyridine-insensitive Cav1.2 (DHP–/–) arterial myocytes before (control) and after 10 µM nifedipine ([Ca2+]o = 20 mM and membrane potential = –70 mV). B: scatter plot of nPs of Ca2+-sparklet sites (n = 5 cells/group). Gray lines represent median values. C: means ± SE of number of Ca2+-sparklet sites per cell in arterial myocytes from wild-type or DHP–/– cells before and after 10 µM nifedipine. D: arterial wall [Ca2+]i in pressurized (80 mmHg) wild-type and DHP–/– arteries during application of 1 µM nifedipine or a solution with nominal Ca2+. E: means ± SE of arterial wall [Ca2+]i under control conditions and after application of nifedipine or 0-Ca2+ solution in wild-type and DHP–/– arteries. *P < 0.05.

View larger version (38K): [in this window] [in a new window]   Fig. 7. Mouse vascular smooth muscle cells express Cav1.2 but not Cav1.3 channels. A: representative gel of Cav1.2- and Cav1.3-subunit transcripts in brain (positive control) and single vascular smooth muscle cells (SMC); -actin was used as an internal control. B and C: representative confocal images of single vascular smooth muscle cells stained with antibodies targeted against Cav1.2 and Cav1.3, respectively. D: a preparation that does not include primary antibody was used as a negative control for secondary antibody. Note that no staining was observed in this preparation, suggesting that our primary antibodies are highly specific for Cav1.2 and Cav1.3 subunits. Insets in C and D are differential interference contrast images of cells examined.

	DISCUSSION

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In this study, we report three fundamental observations. First, we provide the first examination of single Ca_v1.3-channel function. Second, we demonstrate that as with Ca_v1.2 channels, Ca_v1.3 channels can produce persistent Ca²⁺ sparklets. Third, we found that Ca_v1.2 channels, not Ca_v1.3 channels or any other Ca²⁺-permeable channel, underlie I_Ca, Ca²⁺ sparklets and hence dihydropyridine-sensitive Ca²⁺ influx in mouse arterial myocytes. These findings and their physiological implications are discussed below.

There are four known members of the Ca_v1.X family of L-type Ca²⁺ channels (1.1, 1.2, 1.3, and 1.4) (7, 29). A hallmark of Ca_v1.X channels is that they are voltage gated and are sensitive, although to differing degrees, to dihydropyridines. Ca_v1.3 channels have a single-channel conductance of 21 pS with 110 mM Ba²⁺, which is similar to that of Ca_v1.1 (16 pS) (8) and Ca_v1.2 (19 pS) (9, 15) channels but larger than Ca_v1.4 channels (4 pS) (15) under similar ionic conditions. Our data also suggest that like Ca_v1.1 and Ca_v1.2 channels, Ca_v1.3 channels could operate in two gating modes with short (1.6 ms) and long (9.5 ms) open times (8–10, 14, 21, 39). Interestingly, Ca_v1.4 channels apparently do not have this bimodal gating behavior (15). Together, these data indicate that Ca_v1.1–3 channels form part of a subgroup within the Ca_v1 family of channels with similar conductance and gating modalities.

The hyperpolarized voltage dependence of activation of Ca_v1.3 channels suggests that these channels contribute to Ca²⁺ influx at relatively negative potentials (27, 52). Our single Ca_v1.3-channel data provide important quantitative insight into mechanisms of Ca²⁺ entry via these channels. For example, recording Ca_v1.3 channel activity at –70 mV with conventional patch-clamp techniques is difficult because of the low P_o of these channels at this voltage. However, the Boltzmann function used to fit our Ca_v1.3-channel P_o-voltage relationship can be used to predict the P_o of these channels at –70 mV. Under this approach, the estimated P_o of a typical Ca_v1.3 channel is calculated to be 10^–5 at –70 mV, a value that is one to three orders of magnitude higher than the average P_o for Ca_v1.2 channels (P_o 10^–8–10^–6) under similar experimental conditions (42, 50).

Our Ca_v1.3-sparklet data suggest that these channels could contribute to Ca²⁺ influx at hyperpolarized potentials. As with Ca_v1.2 channels (35), heterologously expressed Ca_v1.3 channels are capable of producing persistent Ca²⁺ sparklets at hyperpolarized potentials. As expected from our single Ca_v1.3-channel analysis (see above), we found many similarities between Ca_v1.2 and Ca_v1.3 sparklets. Ca_v1.3 sparklets have a quantal amplitude of 34 nM, which is similar to that of Ca_v1.2 (36 nM) and native smooth muscle (38 nM) Ca²⁺ sparklets (34, 35). This is consistent with our observation that Ca_v1.2 and Ca_v1.3 channels have similar single-channel conductances. Another similarity between Ca_v1.2, Ca_v1.3, and smooth muscle cell Ca²⁺ sparklets is that their duration and activity are bimodal. This feature of Ca²⁺ sparklets likely reflects the ability of Ca_v1.2 and Ca_v1.3 channels to operate in two gating modes with either a brief (i.e., mode 1) or long (i.e., mode 2) open time. It is important to note, however, that the total nP_s of Ca_v1.3 sparklets is significantly higher (P < 0.05; see the supplemental figure for this article, available online at the American Journal of Physiology-Heart and Circulatory Physiology website) than that of Ca_v1.2 sparklets at –70 mV. This is consistent with greater Ca_v1.3 activity than Ca_v1.2 activity at this negative potential.

Although Ca_v1.2 and Ca_v1.3 channels produce Ca²⁺ sparklets with similar properties, we provide compelling evidence supporting the hypothesis that Ca_v1.2 channels, not Ca_v1.3 channels, underlie Ca²⁺ sparklets and I_Ca in mouse arterial myocytes. First, the voltage dependencies of activation and inactivation of I_Ca in arterial myocytes are similar to those of Ca_v1.2 but not of Ca_v1.3 currents. Second, nifedipine eliminated Ca²⁺ sparklets in wild-type myocytes but was without effect in Ca_v1.2 DHP^–/– arterial myocytes. This finding is of particular importance because it eliminates the possibility that Ca²⁺ sparklets are produced by a dihydropyridine-sensitive, non-Ca_v1.X channel (e.g., a member of the transient receptor potential superfamily of channels) in arterial myocytes. Third, Ca_v1.2, but not Ca_v1.3, transcript and protein were detected in isolated arterial myocytes. Thus, although Ca_v1.3 channels are capable of producing persistent Ca²⁺ sparklets, Ca_v1.2 channels underlie I_Ca, Ca²⁺ sparklets and hence dihydropyridine-sensitive Ca²⁺ influx in mouse cerebral arterial myocytes.

The importance of Ca_v1.2 with respect to I_Ca and myogenic tone was recently examined in small mesenteric arteries (54). Although the authors of this study did not measure Ca²⁺ sparklets and arterial wall [Ca²⁺]_i, they found that Ca_v1.2 channels play a predominant role in I_Ca and myogenic tone in small mesenteric arteries. Together with the data presented here, these data suggest that Ca_v1.2 channels are crucial in the regulation of Ca²⁺ influx in smooth muscle from multiple arterial beds.

Although Ca_v1.3 channels do not contribute to Ca²⁺-sparklet activity and I_Ca in mouse cerebral (this study) and mesenteric (54) artery smooth muscle, we cannot rule out that these channels modulate Ca²⁺ influx in arterial smooth muscle from other species. Indeed, Ca_v1.3 transcript and protein has been detected in canine basilar artery (37). Future studies should examine Ca²⁺ sparklets in smooth muscle cells from basilar arteries from dog and other species and should assess the contribution of Ca_v1.3 channels to these events.

Ca_v1.2 and Ca_v1.3 channels, as well as PKC, are expressed in multiple excitable cells, including neuronal, cardiac, smooth muscle, endocrine, and outer hair cells (7, 30, 33, 37, 38, 40). Thus our observation that Ca_v1.2 and Ca_v1.3 channels in association with PKC could produce persistent Ca²⁺-sparklet activity has broad physiological implications. For example, the findings in this study, in combination with recent work from our laboratory (2), indicate that persistent Ca_v1.2 sparklets regulate local and global [Ca²⁺]_i in murine arterial smooth muscle. Future studies should examine whether, as in arterial myocytes, persistent Ca²⁺ sparklets modulate physiological processes in which Ca_v1.2 and/or Ca_v1.3 channels and PKC have been shown to be involved. These include synaptic plasticity (28, 51), gene expression and excitability (3, 16), hormone release (53), cardiac excitation-contraction coupling (6), and cardiac pacemaking activity (31).

The Ca_v1.3-sparklet data presented here add to a growing body of evidence suggesting that the activity of dihydropyridine-sensitive, voltage-gated Ca²⁺ channels is heterogeneously dispersed along the plasma membrane (2, 34, 35). In arterial myocytes, Ca²⁺-sparklet activity varies regionally depending on the relative activities of nearby PKC and protein phosphatases 2A and 2B (35). Although the specific mechanisms underlying regional variations in Ca²⁺-sparklet activity (or lack thereof) may vary in different cell types, the findings underscore the power of TIRF microscopy to optically record Ca²⁺ influx via single Ca²⁺ channels from a relatively large surface area (11, 12).

The current-voltage relationship in Fig. 5B shows that, under our experimental conditions (i.e., [Ca²⁺]_o = 20 mM), Ca²⁺ currents produced by Ca_v1.3, Ca_v1.2, and native smooth muscle L-type calcium channels peaked at +20 mV, +40 mV, and +40 mV, respectively. This represents a +30- to +40-mV shift in the current-voltage relationships of these currents relative to those recorded in physiological 2 mM [Ca²⁺]_o (20, 24, 43). Examination of the surface-potential model (17, 23) suggests a potential mechanism for the differences between the voltage dependencies of activation of Ca²⁺ currents at 2 and 20 mM external Ca²⁺. In this model, negatively charged particles (phospholipids and sugars) on the surface membrane of the cell produce a surface potential. This surface potential determines the intramembrane voltage, which is the voltage modulating the voltage sensor of the Ca²⁺ channels. According to the surface-potential model, increasing external Ca²⁺ from 2 to 10 mM would neutralize negatively charged particles associated with the outer leaflet of the plasma membrane, which would reduce the magnitude of the surface potential and thus increase the intramembrane potential. Consequently, a greater membrane depolarization will have to be applied for the Ca²⁺ channels to "feel" the same trans-bilayer field at 20 as at 2 mM [Ca²⁺]_o, which would result in a rightward shift in the voltage dependencies of these channels.

To conclude, we have provided the first examination of single Ca_v1.3 channels. Our data demonstrate, for the first time, that Ca_v1.3 channels have similar conductance and gating modalities to Ca_v1.2. Accordingly, Ca_v1.3 sparklets resemble most of the features of Ca²⁺ sparklets in arterial myocytes and in tsA201 cells expressing Ca_v1.2 channels. Our data indicate that although Ca_v1.3 channels can produce persistent Ca²⁺ sparklets, Ca_v1.2 channels underlie I_Ca, Ca²⁺ sparklets, and hence dihydropyridine-sensitive Ca²⁺ influx in mouse cerebral arterial myocytes.

	GRANTS

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This work was supported by National Heart, Lung, and Blood Institute Grants HL-85870, HL-77115, HL-07828, and HL-44948 and by grants from the American Heart Association (0635118N) and the Austrian Science Fund (P17159 [GenBank] ).

	ACKNOWLEDGMENTS

 
We thank V. Scott Votaw for help with image analysis, Jennifer Cabarrus for technical assistance, and Drs. Carmen A. Ufret-Vincenty and Madeline Nieves-Cintrón for reading this manuscript. Drs. Dianne Lipscombe and John Exton kindly provided Ca_v1.3 and PKC clones, respectively.

	FOOTNOTES

 

Address for reprint requests and other correspondence: L. F. Santana, Dept. of Physiology and Biophysics, Univ. of Washington, Box 357290, Seattle, WA 98195 (e-mail: santana{at}u.washington.edu)

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

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