Ca_V1.2/Ca_V3.x channels mediate divergent vasomotor responses in human cerebral arteries

The spatial and temporal distribution of brain blood flow is set by networks of resistance arteries. Cerebral arteries are responsive to a range of humoral and mechanical stimuli (Furchgott and Zawadzki, 1980; Segal, 2000; Filosa et al., 2006), one of the key being arterial pressure (Knot and Nelson, 1998). Under dynamic conditions, elevated intraluminal pressure initiates arterial constriction by modulating ionic conductances that depolarize arterial smooth muscle membrane potential (V_M) (Knot and Nelson, 1998; Welsh et al., 2000, 2002). This electrical event in turn grades the opening of voltage-gated Ca²⁺ (Ca_V) channels, and the subsequent rise in cytosolic [Ca²⁺]_i triggers the activation of myosin light chain kinase and actin–myosin cross-bridge cycling (Gallagher et al., 1997; Cole and Welsh, 2011). As Ca_V channels play a key role in setting arterial tone, these channels have long been a target of therapeutic interest, with blockers commonly used to moderate systemic hypertension and cerebral vasospasm (Godfraind, 2014).

The Ca_V channel family comprises 10 α₁ pore–forming subtypes, among which the Ca_V1.2 channel predominates in the transduction of vascular smooth muscle contractility (Catterall, 2011). Given the robust vasodilatory response to L-type blockers (e.g., dihydropyridines), it is often assumed that this channel is the sole subtype expressed in rodent and human arteries (Knot and Nelson, 1998; Moosmang et al., 2003). Although a principal conductance, emerging animal studies indicate the presence of additional subtypes (Jensen et al., 2004; Kuo et al., 2010; Abd El-Rahman et al., 2013). Of particular note are T-type Ca²⁺ channels whose voltage profiles are leftward shifted, compared with L-type, positioning them to be more active at hyperpolarized V_M. Initial data argued that T-type channels were functionally similar to L-type in elevating cytosolic [Ca²⁺]_i, albeit at lower intraluminal pressures where arteries are hyperpolarized (Abd El-Rahman et al., 2013; Björling et al., 2013). Recent reports, however, indicate that T-type regulation of arterial tone is more sophisticated, with one subtype (Ca_V3.1) driving constrictor responses and the other (Ca_V3.2) mediating feedback dilation (Chen et al., 2003; Harraz et al., 2014; Thuesen et al., 2014). These animal observations are intriguing, but their translational significance has remained elusive as mechanistic studies have not extended to the human vasculature.

The goal of this study was to delineate Ca²⁺ channels in native human cerebral arterial smooth muscle and to determine how each conductance modulates arterial tone development. We hypothesized that both L- and T-type Ca²⁺ channels are present and that each conductance mediates distinctive vasomotor responses. Our investigation progressed from cells to tissues and involved the integrative use of pressurized myography, Western blot analysis, PCR, electrophysiology, and computational modeling. In human cerebral arterial smooth muscle cells, we documented for the first time the expression of L- (Ca_V1.2) and T-type (Ca_V3.2/Ca_V3.3) channels and characterized their unique physiological and pharmacological profiles. Subsequent observations showed that Ca_V1.2 and Ca_V3.3 augmented myogenic arterial tone in a manner consistent with their voltage dependence. In stark contrast, human Ca_V3.2 antagonized tone development and enabled indirect arterial vasodilation through its influence on large-conductance Ca²⁺-activated K⁺ (BK_Ca) channels. The divergent roles of Ca²⁺ channels are therapeutically important, as subtype-specific targeting could either suppress or enhance arterial tone. In summary, this human-based study is the first to illustrate the expression of three different Ca_V channels and to encode their distinctive influences on human cerebral arterial tone development.

Excised human brain samples were obtained after institutional review board approval, written informed consent, and in accordance with the guidelines of the Declaration of Helsinki. Brain tissues were collected and placed in cold PBS, pH 7.4, containing (mM) 138 NaCl, 3 KCl, 10 Na₂HPO₄, 2 NaH₂PO₄, 5 glucose, 0.1 CaCl₂, and 0.1 MgSO₄, and transferred to the laboratory. Small superficial cerebral arteries (∼150–250-µm diameter) were carefully dissected out of surrounding tissue and cut into segments. Animal procedures were approved by the Animal Care and Use Committee at the University of Calgary. In brief, female Sprague–Dawley rats (2–4-mo old) or C57BL/6 mice (2–4-mo old) were euthanized via CO₂ asphyxiation. The brain was carefully removed and placed in cold PBS. Middle and posterior cerebral arteries were carefully dissected out of surrounding tissue and cut into ∼2-mm segments for enzymatic digestion.

Human arterial segments were mounted in a customized arteriograph and superfused with warm (37°C) physiological saline solution (PSS) containing: 119 mM NaCl, 4.7 mM KCl, 20 mM NaHCO₃, 1.1 mM KH₂PO₄, 1.2 mM MgSO₄, 1.6 mM CaCl₂, and 10 mM glucose, pH 7.4 (21% O₂, 5% CO₂, balance N₂). To limit the endothelial feedback influence on myogenic tone development, air bubbles were passed through the vessel lumen (1–2 min). Arteries were equilibrated at 15 mmHg, and the contractile responsiveness was assessed by brief application of 60 mM KCl. After equilibration, intraluminal pressure was incrementally elevated from 20 to 100 mmHg and arterial external diameter was monitored under control conditions and in the presence of 200 nM nifedipine (Ca_V1.2 inhibitor), 1 µM NNC 55–0396 (T-type blocker), and/or 50 µM Ni²⁺ (Ca_V3.2 blocker). Maximal arterial diameter was subsequently assessed in Ca²⁺-free PSS (zero externally added Ca²⁺ plus 2 mM EGTA). Percentage of myogenic tone was used as a measure of myogenic responsiveness and was calculated at each intraluminal pressure as follows: % myogenic tone = 100*(D₀ − D)/D₀, where D is external diameter under control conditions (Ca²⁺ PSS) or treated conditions (nifedipine, NNC 55–0396 or Ni²⁺), and D₀ is external diameter in Ca²⁺-free PSS. Percentage of maximal tone was used to ascertain the relative contribution of L- and T-type channels and was calculated at each intraluminal pressure as: % maximal tone = 100*ΔD_X/D₀ − D, where ΔD_X is change in arterial diameter induced by nifedipine or NNC 55–0396, D₀ is external diameter in Ca²⁺-free PSS, and D is arterial diameter under control conditions (Ca²⁺ PSS).

Smooth muscle cells from human, rat, or mouse cerebral arteries were enzymatically isolated as described previously (Harraz and Welsh, 2013). In brief, arterial segments (1–2-mm long) were placed in an isolation medium (at 37°C for 10 min) containing (mM): 60 NaCl, 80 Na-glutamate, 5 KCl, 2 MgCl₂, 10 glucose, and 10 HEPES with 1 mg/ml BSA, pH 7.4. Vessels were then exposed to a two-step digestion process that involved: (1) 10–15-min incubation in isolation medium (37°C) containing 0.5 mg/ml papain and 1.5 mg/ml dithioerythritol; and (2) a 15-min incubation in isolation medium containing 100 µM Ca²⁺, 0.7 mg/ml of type F collagenase, and 0.3 mg/ml of type H collagenase. After incubation, tissues were washed repeatedly with ice-cold isolation medium and triturated with a fire-polished pipette. Isolated cells were kept in ice-cold isolation medium for use within ∼6 h.

Total RNA was isolated from human cerebral arteries or isolated smooth muscle cells using the RNeasy micro kit (QIAGEN) according to the manufacturer’s recommendations. Control total RNA preparations from human whole brain or skeletal muscle were obtained from Takara Bio Inc., whereas human retina total RNA was provided by P. Schnetkamp (University of Calgary, Calgary, Canada). Reverse transcription was performed using the Quantitect reverse transcription kit (QIAGEN). For the negative control groups, all components except the reverse transcriptase were included in the reaction mixtures. Real-time PCR using intron-spanning primer sequences obtained from the qPrimer depot (Table S2; Cui et al., 2007) was performed using the Kapa SYBR Fast Universal qPCR kit (Kapa Biosystems). Human glyceraldehyde 3-phosphate dehydrogenase (GAPDH) gene was used as the reference gene. Control reactions and those containing cDNA from cerebral arteries were performed with 1 ng of template per reaction. Because of the limited quantities of RNA obtained from isolated smooth muscle cells (∼200 cells), the entire cDNA yield from each preparation was used to assay the full set of test and housekeeping genes. The running protocol extended to 45 cycles consisting of 95°C for 5 s, 55°C for 10 s, and 72°C for 8 s using an Eppendorf Realplex 4 Mastercycler. PCR specificity was checked by dissociation curve analysis. Assay validation was confirmed by testing serial dilutions of pooled template cDNAs, suggesting a linear dynamic range of 2.8–0.0028 ng of template and yielding percent efficiencies ranging from 80.4 to 108% (Table S3). No-template controls yielded no detectable fluorescence. Expression of the various Ca_V subtypes in human cerebral arteries or smooth muscle cells relative to whole brain (Ca_V1.2, Ca_V1.3, Ca_V2.1, Ca_V2.2, Ca_V2.3, Ca_V3.1, Ca_V3.2, and Ca_V3.3), skeletal muscle (Ca_V1.1), or retina (Ca_V1.4) was determined using the relative expression software tool (REST) version 2.0.13.

Whole human arteries were collected and frozen at −80°C in PBS, pH 7.4. The arteries were thawed on ice, centrifuged briefly to remove PBS, resuspended in low ionic strength buffer (10 mM Tris, pH 7.0, and 0.5 mM MgCl₂) containing Complete Ultra protease inhibitors (Roche), and incubated for 10 min on ice. The tissue was homogenized using a Polytron, followed by the addition of 1 volume solution A (0.5 M sucrose, 10 mM Tris, pH 7.0, 40 µM CaCl₂, 6 mM 2-mercaptoethanol, and 0.3 M KCl). After additional homogenization with the Polytron, the arteries were further subjected to 40 strokes in a Dounce homogenizer. The resulting crude lysate was centrifuged at 100,000 g for 1 h and resuspended in 50 µl of solution B (0.25 M sucrose, 10 mM Tris, pH 7.0, 20 µM CaCl₂, 3 mM 2-mercaptoethanol, and 0.15 M KCl). The protein concentration was determined with the Protein assay (Bio-Rad Laboratories) using BSA as the standard. The samples (4 µg per lane) were electrophoresed on a 5.6% polyacrylamide gel followed by transfer to PVDF membranes. The membranes were blocked for 1 h at room temperature using 0.2% Tropix I-Block (Applied Biosystems) in PBS followed by the addition of 1:200 diluted primary polyclonal antibodies raised against Ca_V1.2, Ca_V3.2, or Ca_V3.3 (Alomone Labs) overnight at 4°C. The membranes were washed three times for 10 min with PBS containing 0.1% Tween-20 followed by the addition of 1:10,000 goat anti–rabbit IRDye 800CW (Li-Cor Biosciences) for 1 h at room temperature, washed three times with PBS-Tween, and visualized using an infrared imaging system (Odyssey; Li-Cor Biosciences).

Whole-cell currents were recorded using a patch-clamp amplifier (Axopatch 200B; Molecular Devices), filtered at 1 kHz, digitized at 5 kHz, and stored on a computer for offline analysis with Clampfit 10.3 software. Whole-cell capacitance was measured with the cancellation circuitry in the voltage-clamp amplifier. A 1-M NaCl–agar salt bridge between the reference electrode and the bath solution was used to minimize offset potentials (<2 mV). All electrophysiological recordings were performed at room temperature (∼22°C).

Conventional patch-clamp electrophysiology was used to monitor whole-cell Ca_V channel currents in isolated smooth muscle cells as reported previously (Harraz and Welsh, 2013). Recording electrodes (5–8 MΩ) were pulled from borosilicate glass microcapillary tubes (Sutter Instrument) using a micropipette puller (Narishige PP-830), and backfilled with pipette solution (mM): 135 CsCl, 5 Mg-ATP, 10 HEPES, and 10 EGTA, pH 7.2. Cells were voltage clamped and equilibrated in bath solution (mM): 110 NaCl, 1 CsCl, 10 BaCl₂, 1.2 MgCl₂, 10 glucose, and 10 HEPES, pH 7.4. For the experiment depicted in Fig. 3, bath solutions consisted of (mM): 110 NaCl, 1 CsCl, 1.2 MgCl₂, 10 glucose, and 10 HEPES plus charge carrier (1.8 Ca²⁺, 1.8 or 10 Ba²⁺). To record whole-cell currents, isolated smooth muscle cells were voltage clamped at a holding potential of −60 mV and subjected to a −90-mV prepulse (200 ms) and then to 10 voltage steps (300 ms) ranging from −50 to 40 mV (10-mV interval). I-V relationships were plotted as peak current (pA), current density (pA/pF), or normalized currents (I/I_max) at different voltage steps. For voltage ramp recordings, whole-cell currents were monitored using a voltage protocol that ramped from −100 to 100 mV (0.66 mV/ms). Voltage dependence of steady-state inactivation was assessed by a step protocol: (a) prepulse to −90 mV (300 ms); (b) stepping from −70 to +20 mV (10-mV intervals, 1.5 s each); (c) hyperpolarizing back to −90 mV (10 ms); and (d) stepping to a test voltage of 10 mV (200 ms). Whole-cell currents elicited by the test voltage were normalized to maximal current to plot %I/I_max versus voltage steps. Percentage of inactivation was calculated based on current amplitude at different time intervals (300, 600, 900, 1,200, and 1,500 ms) after peak current development. Inactivation time constants (τ) were obtained using Clampfit 10.3 software by standard exponential fitting of the inactivating segment of the current. To assess the voltage dependence of activation, isochronal tail currents were monitored. In particular, cells held at −60 mV were subjected to a prepulse (−90 mV, 300 ms), voltage steps (−80 to 40 mV, 10-mV intervals, 50 ms), and then a final hyperpolarizing test pulse (−90 mV, 200 ms) to evoke tail currents. Normalized tail currents (%I/I_max) were plotted versus the voltage step. Activation time constants (τ) were obtained by standard exponential fitting of the activating downward segment of the whole-cell Ba²⁺ currents.

Perforated patch-clamp electrophysiology was used to measure whole-cell K⁺ currents in isolated human smooth muscle cells. The bath solution contained (mM): 134 NaCl, 4 KCl, 2 MgCl₂, 2 CaCl₂, 10 glucose, and 10 HEPES, pH 7.4. The pipette solution contained (mM): 110 K aspartate, 30 KCl, 10 NaCl, 2 MgCl₂, 10 HEPES, and 0.05 EGTA, pH 7.2, with 200 µg/ml amphotericin B. Membrane currents were gap-free recorded while the cells were voltage clamped at −40 mV. As reported previously (Harraz et al., 2014) and to analyze the frequency and amplitude of spontaneous transient outward K⁺ currents (STOCs), the cutoff value was set to be three times the single-channel conductance.

A single complete arterial network of the cerebral vasculature was acquired using a micrograph of a resected brain biopsy. The micrograph was then manually segmented to obtain a network containing six orders (first–sixth) of branching arteries. This network was supplied by a single inflow vessel and contained 20 outflow vessels. To obtain a mathematical representation suitable for blood flow simulations, the network was discretized into 135 cylindrical vessel segments (213–1,700-mm long). Arterial diameter values were initially traced and set constant for each order according to average measured values (in µm): 600, 396, 261, 173, 114, and 75 for first, second, third, fourth, fifth, and sixth order vessels, respectively.

A previously developed theoretical model (Pries et al., 1990, 1994; Goldman and Popel, 2000) was used to calculate pressures and two-phase (erythrocyte and plasma) steady-state flows in the cerebral arterial network considered. This hemodynamic model implements conservation of blood volume flow (Q) and erythrocyte volume flow (QH_D) at each node joining two or three vessel segments, where H_D is the discharge (flow-averaged) hematocrit. These conservation equations are given below, where the sums are over all segments i connected to node j:

These equations are solved using the Poiseuille pressure–flow relation ΔP = 8ηLQ/(R⁴), where R and L are segment radius and length, respectively, and η is the blood viscosity. The model includes the following empirical properties of blood rheology: (a) the Fahraeus effect, which determines how tube (volume-averaged) hematocrit varies with vessel diameter; (b) the Fahraeus–Lindqvist effect, which determines how viscosity varies with diameter and hematocrit; and (c) the phase separation (or plasma skimming) effect, which determines how erythrocytes are distributed at diverging bifurcations. The model requires specification of hematocrit at the inlet segment and specification of pressures at the upstream ends of the inlet segments and downstream ends of the outlet segments. All flow simulations used an inlet hematocrit of 0.42, and inlet and outlet pressures of 75 and 40 mmHg, respectively, in accordance with earlier studies that monitored the arterial pressure drop in cerebral vascular networks (Faraci and Heistad, 1990).

The pressure–diameter relationship measured in isolated human vessels under control conditions (see Fig. 1) was fit with a fourth-order polynomial, and this was used to adjust diameters from the initially constant values within each order given above based on the calculated pressures in each segment. Solutions were obtained iteratively, by alternating updates of pressures and flows with updates of diameters, until pressures, diameters, and flows became constant (steady state) throughout the model network. The same method was used for simulations in the presence of nifedipine, NNC, or Ni²⁺ (see Figs. 6 and 7), by using fourth-order polynomial fits to the measured variations in the pressure–diameter relationship.

Data are expressed as means ± SEM, and n = a/b indicates the number of vessels or cells (a) per the number of human subjects, rats, or mice (b). Where appropriate, paired or unpaired t tests, or one-way ANOVA was performed using SigmaPlot 12.5 to compare the effects of a given condition/treatment on arterial diameter, or whole-cell current. P-values ≤0.05 were considered statistically significant. Averaged I-V relationships were fit to the following Peak Gaussian third Parameter function: I(V) = I_max*exp[−0.5{(V − V_max)/b}²], where I_max is peak current (I), V_max is V at I_max, and b is the slope of the distribution. The voltage dependence of activation and steady-state inactivation were described with single Boltzmann distributions of the following forms: Activation: I(V) = I_max/(1 + exp[−(V − V₅₀)/k]); Inactivation: I(V) = I_max/(1 + exp[(V − V₅₀)/k]), where I_max is the maximal activatable current, V₅₀ is the voltage at which the current is 50% activated or inactivated, and k is the slope of the distribution.

Fig. S1 reports low and high voltage-activated currents in human cerebral arterial smooth muscle, and that high millilmolar Ba²⁺ masks the separation of the two current components. Fig. S2 illustrates the effects of high concentrations of Ba²⁺ as the charge carrier on the I-V plots recorded in human smooth muscle. Fig. S3 depicts the biophysical properties of the nifedipine-sensitive and -insensitive currents in human cerebral arterial smooth muscle. Fig. S4 shows kinetic analysis of inward currents before and after the application of nifedipine. Fig. S5 compares representative traces and reversal potentials of Ba²⁺ currents recorded in human, mouse, and rat cerebral arterial smooth muscle cells. Table S1 reports the demographic characteristics of the origin of tissues and cells used. Tables S2 and S3 show the primer sequences and validation parameters for the qPCR analysis.

Pial cerebral arteries, isolated from patients undergoing brain resection surgery (Fig. 1 A and Table S1), were cannulated and mounted in a myograph and pressurized from 20 to 100 mmHg to assess myogenic reactivity. In physiological Ca²⁺ (1.6 mM), human cerebral arteries gradually constricted to increases in intraluminal pressure (Fig. 1, B and C); this active response was abolished by removing extracellular Ca²⁺ (0 mM Ca²⁺/2 mM EGTA). Fig. 1 D illustrates that the percentage of myogenic tone increased progressively from 10% at 20 mmHg to 33% at 100 mmHg.

Given the preceding Ca²⁺-dependent observations, we screened for the α₁ pore–forming subunits of Ca_V channel subtypes (Ca_Vx.x). Beginning with qPCR, analysis of human cerebral arteries and isolated smooth muscle cells revealed that all Ca_V subunits were expressed in the former, whereas only Ca_V1.1, Ca_V1.2, Ca_V1.4, Ca_V3.2, and Ca_V3.3 were expressed in the latter (Fig. 2 A). When standardized to reference tissue (whole brain: Ca_V1.2, Ca_V1.3, Ca_V2.1, Ca_V2.2, Ca_V2.3, Ca_V3.1, Ca_V3.2, and Ca_V3.3; skeletal muscle: Ca_V1.1; retina: Ca_V1.4), Ca_V1.2 and Ca_V3.2 were enriched in whole arteries (Fig. 2 B), and Ca_V1.2, Ca_V3.2, and Ca_V3.3 displayed enriched expression in isolated smooth muscle cells (Fig. 2 C). Endothelial contamination of isolated smooth muscle samples was minimal as Von Willebrand factor mRNA levels decreased dramatically (∼96%) compared with whole arteries. Focusing on the enriched targets, Western blot analysis confirmed the protein expression of Ca_V1.2, Ca_V3.2, and Ca_V3.3 in human cerebral arteries (Fig. 2 D).

Using patch-clamp electrophysiology, we then monitored inward currents in cerebral arterial smooth muscle. Freshly isolated human cells were bathed in 1.8 mM Ca²⁺ and 1.8 or 10 mM Ba²⁺; the first mimics physiological extracellular conditions, whereas the latter two accentuate charge flow through open Ca²⁺ channels. Stepping from −90 mV to voltage steps ranging from −50 to 40 mV, we observed a voltage-dependent inward current that was approximately fivefold less in 1.8 mM Ca²⁺ than in high [Ba²⁺] (Fig. 3, A and B). Fig. 3 C illustrates the voltage dependence of the whole-cell currents, with normalized I-V plots showing that Ba²⁺ evoked a depolarizing shift compared with Ca²⁺. Notably, normalized Ca²⁺ current (Fig. 3 C) exhibited a hyperpolarized threshold and an additional inward hump peaking at −30 mV, suggestive of a low voltage–activated current component (Fig. S1). The rightward shift associated with Ca²⁺ replacement could be further advanced by working with solutions containing 50 or 110 mM Ba²⁺ (Fig. S2). Given the small amplitude of native Ca²⁺ currents, subsequent recordings were performed in 10 mM Ba²⁺; step and ramp voltage protocols elicited robust inward Ba²⁺ currents (Fig. 3, D and E). Linear regression analysis (Fig. 3 F) indicated an inverse correlation between peak current amplitude and age of subjects (r² = 0.531; P < 0.0001; age range 19–70 yr; n = 45/13; six females). This correlation was particularly evident in female (r² = 0.6057; P < 0.0001; 19–58 yr; n = 19/6) rather than male (r² = 0.0001; P = 0.9566; 33–70 yr; n = 26/7) subjects.

The whole-cell inward Ba²⁺ current depicted in Fig. 3 reflected simultaneous influx through L-type (Ca_V1.2) and T-type (Ca_V3.2/Ca_V3.3) Ca²⁺ channels. To effectively delineate between the two types, nifedipine was applied at a concentration (200 nM) that fully abolishes L- but not T-type currents (Liao et al., 2007; Harraz and Welsh, 2013); ∼75% block was observed (Fig. 4 A). Note that the use of different prepulse voltages was not successful in the separation of low and high voltage-activated Ca²⁺ channel currents in rodent cerebral arterial smooth muscle (Kuo et al., 2010; Abd El-Rahman et al., 2013). To test whether the residual nifedipine-insensitive current was dominated by a T-type conductance, we applied 1 µM NNC 55–0396. This T-type blocker abolished the remaining current, and I-V plots showed that nifedipine- and NNC-sensitive current components peaked (V_max) at 19.7 ± 0.4 mV and 2.3 ± 0.9 mV, respectively (Fig. 4, B and C). Caution is warranted when using pharmacological T-type blockers given their reported off-target effects on L-type channels (Kuo et al., 2010; Abd El-Rahman et al., 2013; Harraz and Welsh, 2013). Experiments in Fig. 4 (D–F) confirmed the successful delineation as nifedipine evoked hyperpolarized shifts in the voltage dependence of: (a) steady-state inactivation (V_50-Inact, −35.4 ± 2.3 mV vs. control, −12.7 ± 0.9 mV); (b) activation (V_50-Act, −20.3 ± 1.3 mV vs. control, −5.2 ± 0.8 mV); and (c) peak window current (−33.4 mV vs. control −14.1 mV). Kinetic analysis additionally noted that the nifedipine-insensitive T-type component inactivated faster (Fig. 4, E and F), exhibited rapid activation (T-type: τ_Act, 2.1 ms; L-type: τ_Act, 7.5 ms), and deactivated slower (T-type: τ_Deact, 8.9 ms; L-type: τ_Deact, 3.2 ms) compared with subtracted nifedipine-sensitive current (Fig. S3) or total control current (Fig. S4). Linear regression analysis (Fig. 4 G) indicated a negative correlation between age and the amplitude of the L-type (r² = 0.7396; P < 0.0001; 36–70 yr; n = 19/6) but not for T-type current (r² = 0.0753; P = 0.2164; 33–70 yr; n = 22/7).

Past studies have shown that rat and mouse arterial smooth muscle express Ca_V1.2 (L), Ca_V3.1 (T), and Ca_V3.2 (T) subtypes (Kuo et al., 2010; Abd El-Rahman et al., 2013; Björling et al., 2013; Thuesen et al., 2014). Human cells displayed a similar expression profile, with the exception being that Ca_V3.1 was substituted by Ca_V3.3 (Fig. 2). Using patch-clamp electrophysiology, we compared the electrical properties of cerebral arterial Ca²⁺ channels between species using 10 mM Ba²⁺ as the charge carrier. Values of cell capacitance (pF), indicative of cell size, varied across species (Fig. 5 A). Inward current densities (pA/pF) were lower in human smooth muscle cells than in mouse or rat; however, normalized currents displayed comparable dependence on voltage (Fig. 5, B and C). Whole-cell currents inactivated faster in mouse smooth muscle cells, whereas the reversal potential of rat currents was depolarized relative to human or mouse (Fig. S5). 200 nM nifedipine elicited comparable blockade across species, and the residual insensitive currents displayed 10–20-mV hyperpolarized shifts consistent with T-type activity (Fig. 5, D and E). In all three species, L-type channel inhibition quickened inactivation and leftward shifted the activation/inactivation profiles (Fig. 5, F and G).

In rodent cerebral arteries, studies have shown that L- and T-type channels (Ca_V1.2 and Ca_V3.1) facilitate myogenic tone development at high and low intraluminal pressures, respectively (Abd El-Rahman et al., 2013; Björling et al., 2013). Both conductance types (L-type: Ca_V1.2; T-type: Ca_V3.3 substituting for Ca_V3.1) are present in human cerebral arteries and, as such, comparable experiments were conducted on pressurized human vessels to elucidate potential roles of Ca²⁺ channels. Similar to electrophysiology, L-type Ca²⁺ channel blockade preceded T-type suppression to minimize off-target effects. 200 nM nifedipine induced a vasodilator response in human vessels that was limited at low pressure and maximal at 80–100 mmHg. The additional application of a T-type blocker (1 µM NNC 55–0396) modestly dilated arteries in a pressure-dependent manner, with peak responses at 40–60 mmHg (Fig. 6, A–C). Data in Fig. 6 (B and C) displayed distinctive L- and T-type vasomotor trends, although statistical significance was not achieved because of limited observations (n = 3/3; P = 0.087–0.157). As such, the relative contribution of the two conductances to developed myogenic tone was dynamic, with T-type channels (Ca_V3.3) prevailing at lower pressures when the V_M was relatively hyperpolarized and L-type channels (Ca_V1.2) dominating at higher pressures when smooth muscle is strongly depolarized (Fig. 6 D).

To discriminate between the contributions of Ca_V3.2 and Ca_V3.3 to myogenic constriction, we examined the effects of a specific Ca_V3.2 blocker (50 µM Ni²⁺) on isolated human smooth muscle cells and arteries. At this concentration, Ni²⁺ was verified to selectively suppress Ca_V3.2 currents, but not other L- or T-type channels expressed in arterial smooth muscle or expression systems (Lee et al., 1999; Harraz and Welsh, 2013; Harraz et al., 2014). Starting with electrophysiology, we confirmed that 50 µM Ni²⁺ partially suppressed the nifedipine-insensitive Ca_V3.x current (Fig. 7, A and B). A kinetic analysis followed, and time constant measurements revealed that the Ni²⁺-sensitive current activated and inactivated faster (Ca_V3.2: τ_Act, 0.9 ± 0.1 ms; τ_Inact, 17.1 ± 3.3 ms) than the insensitive current (Ca_V3.3: τ_Act, 4.3 ± 0.3 ms; τ_Inact, 48.7 ± 9.5 ms), consistent with Ni²⁺ selectively blocking Ca_V3.2 (Fig. 7, C and D) (Lee et al., 1999; Perez-Reyes, 2003). We next applied 50 µM Ni²⁺ onto pressurized human arteries, and contrary to the vasodilation observed with a broad spectrum T-type blocker (NNC 55–0396; Fig. 6), Ni²⁺ enhanced myogenic constriction at pressures ≤60 mmHg (Fig. 7, E and F). Interestingly, Ni²⁺- and NNC 55–0396-induced vasomotor changes were pressure dependent but modestly different from one another, a finding that aligns with the voltage profiles of Ca_V3.2 and Ca_V3.3 (Fig. 7 G). The preceding Ni²⁺ observations suggest, in accordance with rat studies (Harraz et al., 2014), that Ca_V3.2 mediates a negative feedback response by downstream triggering of BK_Ca channels to initiate STOCs. Perforated patch-clamp electrophysiology indicated the presence of STOCs in human smooth muscle cells and showed that 50 µM Ni²⁺ significantly reduced the frequency (control, 1.30 ± 0.35 Hz; Ni²⁺, 0.69 ± 0.17 Hz) but not the amplitude (control, 6.87 ± 0.09 pA; Ni²⁺, 6.56 ± 0.25 pA) of these events (Fig. 7 H).

Myography experiments (Figs. 6 and 7) showed that human cerebral arterial Ca²⁺ channel subtypes elicited divergent vasomotor effects whereby Ca_V1.2/Ca_V3.3 augmented but Ca_V3.2 counterbalanced myogenic constriction. To conceptually ascertain how each channel subtype could influence cerebral blood flow, a computational model was developed using an arterial network map (lengths and basal diameters) generated from a photomicrograph of excised human brain tissue (see Materials and methods; Fig. 8, A–C). Intraluminal pressure within the virtual model varied, with inlet and outlet pressures set to 75 and 40 mmHg, respectively (Faraci and Heistad, 1990). The pressure–diameter relationship measured in isolated human vessels in the absence and presence of nifedipine, NNC 55–0936, or Ni²⁺ was fit with a fourth-order polynomial, and these percentage values were used to adjust the initially constant diameters within each segment (Fig. 8 D). This integration enabled the calculation of steady-state distributions of pressure, diameter, and blood flow for each arterial segment (135 in total; Fig. 8 E). Two-dimensional plots and color-coded virtual networks indicated that the suppression of Ca_V1.2 and to a lesser extent Ca_V3.3 increased network blood flow in a pressure/diameter-dependent manner. In contrast, Ca_V3.2 inhibition constricted arterial segments and decreased cerebral blood flow (Fig. 8 F).

This study delineated Ca_V channels in human cerebral arteries and determined subtype-specific contributions to arterial tone development. Experiments progressed from cells to tissues and used pressurized myography, PCR, Western blotting, electrophysiology, and computational modeling. Messenger RNA and protein analyses revealed the expression of L- and T-type α₁ pore–forming subunits in cerebral arterial smooth muscle. Patch-clamp recordings subsequently confirmed the presence of inward current divisible into L- and T-type components in isolated human smooth muscle cells. In pressurized human arteries, L- (Ca_V1.2) and T-type (Ca_V3.3) channel suppression dilated arteries in a pressure-dependent manner at higher and lower intraluminal pressures, respectively. Subtype-specific targeting of Ca_V3.2 paradoxically implicated this conductance in restraining arterial constriction via a mechanism involving BK_Ca channels. In a final set of experiments, computational modeling conceptually illustrated how each Ca_V subtype plays a distinctive role in setting cerebral blood flow. To summarize, this study reports for the first time the expression and divergent functions of three Ca²⁺ channel subtypes in human cerebral arteries.

A century ago, Bayliss (1902) first described the inherent ability of resistance arteries to constrict to elevated arterial pressure, a phenomenon known as the myogenic response. In the cerebral circulation, this myogenic response is essential to maintain brain perfusion, and a global rise in smooth muscle [Ca²⁺]_i is critical to the transduction process. Like most stimuli, elevated intraluminal pressure drives a rise in cytosolic [Ca²⁺]_i through the induction of arterial depolarization and subsequent activation of voltage-gated L-type Ca_V1.2 channels (Knot and Nelson, 1998). Although a dominant player, animal studies have noted other Ca²⁺ channels (T-type, Ca_V3.1/Ca_V3.2) in cerebral arteries (Kuo et al., 2010; Abd El-Rahman et al., 2013). Distinct from the “long-lasting” kinetics and high voltage activation profile of the “L”-type conductance, “T”-type channels are characteristically “transient” and activate at more hyperpolarized voltages (Perez-Reyes, 2003; Catterall, 2011). Although Ca²⁺ entry events through L-type channels (Ca_V1.2 sparklets) have been characterized in vascular smooth muscle (Navedo et al., 2005), Ca_V3.x sparklets remain to be investigated. They would likely display spatiotemporal characteristics distinct from Ca_V1.2 and which according to their single-channel properties would be more transient in nature and smaller in magnitude (Perez-Reyes, 2003). Recent studies in rodents have implied that T-type channels modulate myogenic tone, albeit in a manner unique and sometimes paradoxical to L-type channels (Chen et al., 2003; Kuo et al., 2010; Abd El-Rahman et al., 2013; Björling et al., 2013; Harraz et al., 2014). Despite the growing interest and therapeutic potential, human studies to date have failed to establish whether T-type channels are present in key vascular beds and if so, how they influence arterial tone.

Within this context, we began to isolate human cerebral arteries from patients undergoing resection surgery. Using vessel myography, we observed for the first time that human cerebral arteries constrict robustly to an increase in intraluminal pressure, with percentages of myogenic responses aligning with previous animal studies (Knot and Nelson, 1998; Cole and Welsh, 2011). The removal of extracellular Ca²⁺ abolished myogenic constriction, an observation consistent with elevated pressure evoking Ca²⁺ flux through Ca_V channels (Knot and Nelson, 1998). 10 genes encode for the α₁ pore–forming subunits of Ca²⁺ channels (Catterall, 2011), and molecular analyses showed robust mRNA expression of three subtypes (Fig. 2). Ca_V1.2 (L-type) and Ca_V3.2 (T-type) were present in isolated human smooth muscle cells prescreened for endothelial contamination. Messenger RNA of Ca_V3.3 was also observed while Ca_V3.1 was minimally expressed, a finding distinct from rodents (Kuo et al., 2010; Abd El-Rahman et al., 2013; Björling et al., 2013). Compared with Ca_V3.1, Ca_V3.3 channel displays slower activation/inactivation kinetics and activates at relatively depolarized potentials (Monteil et al., 2000; Frazier et al., 2001; Chemin et al., 2002). One could rationalize, in this regard, that Ca_V3.3 may be better suited than Ca_V3.1 to drive cellular functions requiring sustained Ca²⁺ flux (Chemin et al., 2002), such as smooth muscle contraction and arterial tone development. Western blot analysis confirmed protein expression of all three subtypes in whole arteries.

Having detected their message and protein expression, electrophysiological recordings followed to separate L- and T-type conductances in human cerebral arterial smooth muscle. Human cells demonstrated a small inward current in physiological Ca²⁺ solutions that displayed a voltage-dependent I-V profile consistent with Ca²⁺ channel activation (Fig. 3). Exchange of physiological Ca²⁺ for higher millimolar Ba²⁺ dramatically enhanced inward current, masked the low voltage–activated Ca²⁺ current component, and induced a depolarized shift in the voltage profile without destabilizing the cell. The latter shift is presumably responsible for masking the hyperpolarized current component and is reflective of a high divalent cation screening effect and barium’s preferential permeability through L-type channels (Perez-Reyes, 2003; Li et al., 2010). Using nifedipine at concentrations that fully block the smooth muscle splice variant of L-type Ca_V1.2 channels (Liao et al., 2007; Harraz and Welsh, 2013), a partial suppression of Ba²⁺ currents occurred. Note that nifedipine has been shown to lack an effect on Ca_V3.x in vascular smooth muscle or expression systems when applied in the submicromolar ranges (Harraz and Welsh, 2013; Harraz et al., 2014). Three lines of evidence subsequently confirmed that L- and T-type channels, respectively, carried the nifedipine-sensitive and -insensitive components. First, the nifedipine-insensitive current was actively blocked by T-type inhibitors. Second, the presumed L- and T-type currents displayed depolarized and hyperpolarized profiles, respectively, in accordance with conductances being high and low voltage activated. Third, the residual insensitive conductance activated and inactivated faster, in keeping with the transient nature of T-type channels (Catterall, 2011). Ca_V3.x currents recorded in human arterial smooth muscle display kinetics similar to those in heterologous expression systems. Note, however, they do display a depolarized voltage-dependent profile because of the use of high millimolar Ba²⁺ as the permeant recording ion (Perez-Reyes, 2003; Harraz and Welsh, 2013). Overall, these findings are unique in the vascular literature and are the first to show that multiple Ca_V conductances are not only present but also electrically active in native human arterial smooth muscle.

Despite their diverse demographic origins, Ca²⁺ channel currents were resolved in all tested cells. With that said, an intriguing inverse relationship between current amplitude and donor age existed (Fig. 3), similar to reports in animal models (Georgeon-Chartier et al., 2013; Fukuda et al., 2014). This relationship was sexually dimorphic and manifested exclusively in females. These observations suggest that Ca²⁺ channel expression and/or activity may be under the regulation of sex hormones (Bowles et al., 2004; Ullrich et al., 2007); however, caution is warranted given the limited sample size and the pathological state of the human subjects. Further analysis showed that the preceding age–current correlation stemmed primarily from variations in L-type activity (Fig. 4), a finding that indicates that the efficacy of calcium channel blockers in the treatment of constrictive disorders may be altered or compromised in the elderly (Kumar and Hall, 2003; Fukuda et al., 2014).

Recent rodent studies have suggested that both L- and T-type channels play a direct role in elevating cytosolic [Ca²⁺]_i and myogenic tone, albeit at specific V_Ms (Abd El-Rahman et al., 2013; Björling et al., 2013). In particular, the L-type Ca_V1.2 channels appear to dominate at elevated pressures when vessels are depolarized. In comparison, the low voltage–activated T-type Ca_V3.1 channels are thought to facilitate tone development at lower pressures when smooth muscle resides in a hyperpolarized state (Abd El-Rahman et al., 2013; Björling et al., 2013). Human L- and T-type channels could operate similarly, and to pursue this hypothesis, a two-step approach was instituted. Starting with a comparative electrical analysis, findings in Fig. 5 illustrate that human, rat, and mouse smooth muscle cells all exhibited L- and T-type current components, with human cells displaying smaller amplitudes. No substantive difference was observed in activation/steady-state inactivation voltage profiles, although inactivation kinetics were faster in mouse smooth muscle. Given these similarities, we next pressurized human cerebral arteries from 20 to 100 mmHg and performed vessel myography under control conditions and in the presence of L-type (nifedipine) and T-type (NNC 55–0396) channel blockers. In a pressure-dependent manner, L- and T-type blockers evoked vasodilation in myogenically responsive arteries (Fig. 6). L- and T-type channels dominated myogenic tone at higher and lower pressures, respectively, in accordance with the physiological window currents of Ca_V1.2 and Ca_V3.x channels. Reduced Ca²⁺ flux through Ca_V3.3 channels is most likely responsible for the vasodilator response to NNC 55–0396, as findings below note that selective Ca_V3.2 blockade induces arterial constriction. Our observations are the first to invoke a role for T-type channels in human cerebral arteries, and arguably this conductance may dominate Ca²⁺ entry in distal arteries where intraluminal pressures are ∼30–45% less than systemic (Faraci and Heistad, 1990). If true, it follows that constrictive disorders in smaller arteries may be more effectively treated by simultaneous L- and T-type channel suppression, a view consistent with clinical findings (Beltrame et al., 2004; Ogura et al., 2012; Sugano et al., 2013).

Studying T-type channels in native cells has been consistently limited by the lack of subtype-specific modulators. One exception is Ni²⁺, a divalent cation that preferentially blocks Ca_V3.2 channels when applied in the low micromolar range (Lee et al., 1999; Harraz et al., 2014). In rat cerebral arterial smooth muscle and HEK cells, Ni²⁺ abolished Ca_V3.2, but not other high or low voltage–activated Ca²⁺ currents. Previous experiments have additionally shown that Ni²⁺ lacked any effects on cells and arteries where Ca_V3.2 channels had been genetically ablated (Harraz and Welsh, 2013; Harraz et al., 2014). Exploiting this tool, electrophysiological experiments followed by kinetic analysis confirmed that 50 µM Ni²⁺ inhibited Ca_V3.2 but not Ca_V3.3 channels in human cerebral arterial smooth muscle cells. In paradoxical contrast to the vasodilation induced by a broad-spectrum T-type blocker, Ni²⁺ enhanced myogenic constriction at pressures ≤60 mmHg (Fig. 7). Such results suggest a distinctive role of Ca_V3.2 channels, perhaps involving a vasodilator conductance sensitive to Ca²⁺ and/or voltage. In this regard, recent rodent literature has argued that Ca_V3.2 can trigger Ca²⁺ release from the sarcoplasmic reticulum in the form of Ca²⁺ sparks, and that the subsequent generation of Ca²⁺ sparks elicits hyperpolarization via BK_Ca channels (Chen et al., 2003; Harraz et al., 2014). To address this possibility, we monitored BK_Ca-mediated STOC activity in human cells, and in support of our hypothesis, Ni²⁺ inhibited these transient hyperpolarizing currents. It is worth noting that previous whole-cell and whole arterial recordings argued against a direct modulatory role of Ni²⁺ on K⁺ channels when applied at low micromolar concentrations (Harraz et al., 2014). Our findings challenge the stereotypic view that Ca²⁺ channels only facilitate myogenic constriction. They further highlight the view that if T-type modulators are to be used therapeutically to treat cardiovascular disorders (Giordanetto et al., 2011), subtype specificity should be thoughtfully considered.

With Ca_V1.2, Ca_V3.2, and Ca_V3.3 channels distinctively influencing arterial diameter, logic dictates that unique blood flow responses should ensue. Although it is impractical to monitor cerebral blood flow in small human arteries, such responses can be predicted in a virtual arterial network (Pries et al., 1990, 1994; Goldman and Popel, 2000). Using architectural and vasomotor data from human arteries, we consequently designed a computational model that mimics a native arterial network. Findings in Fig. 8 highlight that Ca_V1.2 or Ca_V3.3 inhibition dilated virtual cerebral arteries and increased network blood flow. Ca_V1.2 blockade elicited a particularly profound response in larger arteries, vessels that harbor higher intraluminal pressures. Subtype-specific suppression of Ca_V3.2 alternatively constricted arterial segments, diminishing network cerebral blood flow. Such predictions conceptually emphasize the differential significance of Ca²⁺ channels in matching blood flow with brain metabolic activity.

This study identified Ca_V channels in human cerebral arteries and ascertained how each conductance influences myogenic tone. Molecular and electrophysiological analyses demonstrated that α₁ subunits of L- (Ca_V1.2) and T-type (Ca_V3.2 and Ca_V3.3) channels are robustly expressed in human cerebral arterial smooth muscle. In myogenically active human arteries, Ca_V1.2 inhibition induced vasodilation at high intraluminal pressures, whereas Ca_V3.3 blockade induced modest dilation at lower pressures when vessels resided in a hyperpolarized state. Paradoxically, cerebral arterial Ca_V3.2 channels appear to attenuate myogenic tone through the downstream activation of BK_Ca channels.

We thank Dr. Bjorn Hald, Dr. Christophe Altier, and Dr. Michael Walsh for critical comments on the preliminary figures.

This work was supported by an operating grant from the Canadian Institutes of Health Research (CIHR; MOP-69088 to D.G. Welsh). D.G. Welsh is an Alberta Innovates-Health Solutions (AIHS) senior scholar and holds a Canada Research Chair. O.F. Harraz is a Vanier Scholar (CIHR) and was supported by salary studentships from AIHS and Achievers in Medical Sciences (AIMS scholarship). D. Goldman was partially supported by a Discovery grant from the Natural Sciences and Engineering Research Council of Canada (NSERC, R4081A03). A. Zechariah was supported by AIHS and Eyes High postdoctoral fellowships. A.M. Hashad was a Vanier Scholar and a recipient of an AIHS studentship.

The authors declare no competing financial interests.

Richard L. Moss served as editor.