Ca_V3.2 T-type Ca²⁺ channels in H₂S-mediated hypoxic response of the carotid body

Abstract

Arterial blood O₂ levels are detected by specialized sensory organs called carotid bodies. Voltage-gated Ca²⁺ channels (VGCCs) are important for carotid body O₂ sensing. Given that T-type VGCCs contribute to nociceptive sensation, we hypothesized that they participate in carotid body O₂ sensing. The rat carotid body expresses high levels of mRNA encoding the α_1H-subunit, and α_1H protein is localized to glomus cells, the primary O₂-sensing cells in the chemoreceptor tissue, suggesting that Ca_V3.2 is the major T-type VGCC isoform expressed in the carotid body. Mibefradil and TTA-A2, selective blockers of the T-type VGCC, markedly attenuated elevation of hypoxia-evoked intracellular Ca²⁺ concentration, secretion of catecholamines from glomus cells, and sensory excitation of the rat carotid body. Similar results were obtained in the carotid body and glomus cells from Ca_V3.2 knockout (Cacna1h^−/−) mice. Since cystathionine-γ-lyase (CSE)-derived H₂S is a critical mediator of the carotid body response to hypoxia, the role of T-type VGCCs in H₂S-mediated O₂ sensing was examined. Like hypoxia, NaHS, a H₂S donor, increased intracellular Ca²⁺ concentration and augmented carotid body sensory nerve activity in wild-type mice, and these effects were markedly attenuated in Cacna1h^−/− mice. In wild-type mice, TTA-A2 markedly attenuated glomus cell and carotid body sensory nerve responses to hypoxia, and these effects were absent in CSE knockout mice. These results demonstrate that Ca_V3.2 T-type VGCCs contribute to the H₂S-mediated carotid body response to hypoxia.

voltage-gated Ca²⁺ channels (VGCCs) are critical for Ca²⁺ influx and neurotransmission in the nervous system (2, 13). On the basis of their activation properties, VGCCs are divided into high- and low-voltage-activated channels. High-threshold VGCCs are further subdivided into L, N, P/Q, and R types, while low-voltage-activated Ca²⁺ channels are represented by T-type VGCCs (3, 6), designated Ca_V3.1, Ca_V3.2, and Ca_V3.3 channels, which contain pore-forming α_1G-, α_1H-, and α_1I-subunits, respectively (3, 29).

Carotid bodies are the sensory organs for monitoring arterial blood O₂ levels. Hypoxemia (i.e., reduced O₂ levels in blood) increases carotid body sensory nerve activity and triggers the chemosensory reflex, which is critical for maintaining cardiorespiratory functions during low O₂ (10). The carotid body consists of two cell types: 1) glomus (also called type I) cells, the primary O₂-sensing cells, and 2) type II cells (10). Sensory nerve excitation by hypoxia requires elevation of intracellular Ca²⁺ concentration ([Ca²⁺]_i) (1, 14, 30) and release of Ca²⁺-dependent neurotransmitter(s) from glomus cells (16, 30).

Previous studies have shown that VGCCs are critical for elevation of hypoxia-evoked [Ca²⁺]_i in glomus cells (10). Glomus cells express high-voltage-activated Ca²⁺ currents, including L-, N-, and P-type currents, as well as “blocker-resistant” Ca²⁺ currents (7, 18, 31), and L-type VGCCs mediate a substantial portion of the hypoxia-induced Ca²⁺ influx (1, 26). Relatively little information is available on the role of the T-type VGCC in the carotid body. A nickel-sensitive Ca²⁺ current was reported in rat glomus cells (7). Since nickel is known to block T-type VGCCs (11), this finding suggests that glomus cells also express T-type VGCCs. The rat carotid body expresses mRNA encoding the α_1G-subunit, and the Ca_V3.1 channel was proposed to contribute to hypoxia-evoked catecholamine (CA) efflux (5). However, the roles of the other T-type VGCC isoforms in the carotid body response to hypoxia have not been examined.

Recent studies have shown that dorsal root ganglion cells express T-type VGCCs and that their activation by H₂S contributes to hyperalgesia (17). Cystathionine-γ-lyase (CSE)-generated H₂S is required for VGCC-mediated [Ca²⁺]_i influx in glomus cells (14) and sensory nerve excitation of the carotid body (12, 20). Whether H₂S is required for the T-type VGCC-dependent carotid body response to hypoxia, however, is not known.

In the present study we determined the expression and localization of T-type VGCCs in the carotid body, assessed their functional significance using pharmacological and genetic approaches, and elucidated the role of H₂S. Our results demonstrate that the Ca_V3.2 channel is the predominant T-type VGCC expressed in glomus cells and contributes to hypoxia-evoked Ca²⁺ influx and CA secretion, as well as sensory nerve excitation. Our results further demonstrate that CSE-derived H₂S is required for the Ca_V3.2 channel-dependent carotid body response to hypoxia.

METHODS

Experimental protocols were approved by the Institutional Animal Care and Use Committee of The University of Chicago. All experiments were performed on adult male Sprague-Dawley rats and age- and sex-matched Ca_V3.2 knockout (Cacna1h^−/−), CSE null (CSE^−/−), and C57BL/6J mice.

Real-time RT-PCR.

Carotid bodies were harvested from anesthetized rats (n = 4 rats). Two carotid bodies were pooled, and RNA was extracted using TRIzol and reverse-transcribed using SuperScript III reverse transcriptase. Real-time RT-PCR was performed using a MiniOpticon system (Bio-Rad Laboratories) with the SYBR GreenER two-step quantitative RT-PCR kit (catalog no. 11764-100, Invitrogen). Primer sequences for real-time RT-PCR amplification were as follows: GTAACCCGTTGAACCCCATT (forward) and CCATCCAATCGGTAGTAGCG (reverse) for 18S (151 aa; GenBank accession no. X_01117), 5′CTTTGACCTGCTGACACTCTG (forward) and GCCATTACAGTCTTGGTGCTCA (reverse) for Ca_V3.1 (185 aa; GenBank accession no. AF290212), ACTTGGCCATCGTCCTCCTA (forward) and ATGGTGGGATTGATGGGCAG (reverse) for Ca_V3.2 (101 aa; GenBank accession no. AF290213), and GACCCCAGAGCAGTGAGGAT (forward) and TACTTGCTGTCCACGATGCC (reverse) for Ca_V3.3 (155 aa; GenBank accession no. AF 290214). Relative mRNA quantification was calculated using the comparative threshold (C_T) method according to the following formula: 2^−ΔC_T, where ΔC_T is the difference between the threshold cycle of the given target cDNA. The C_T value was taken as a fractional cycle number at which the emitted fluorescence of the sample passes a fixed threshold above the baseline. mRNA levels were normalized to the 18S gene. For confirmation of the purity and specificity of all products, the template was omitted and a standard melting curve analysis was performed.

Immunocytochemistry.

Urethane (1.2 g/kg ip)-anesthetized rats (n = 4) were perfused transcardially with heparinized saline followed by 4% (vol/vol) paraformaldehyde. The carotid bifurcations were dissected and immersed in 25% sucrose solution in distilled water at 4°C for 16–24 h. The tissues were mounted in OCT compound, and 8- to 10-μm-thick sagittal sections were cut and processed for immunofluorescence. The tissue sections were treated with 20% normal goat serum and 0.2% Triton X-100 in PBS for 2 h and then incubated with polyclonal anti-α_1H (1:200 dilution; catalog no. ACC-025, Alomone Labs) and monoclonal anti-tyrosine hydroxylase (TH, 1:2,000 dilution; Sigma) antibodies in PBS with 1% normal goat serum and 0.2% Triton X-100. Control sections were treated with primary anti-α_1H antibody preincubated with α_1H control peptide (1:200 dilution; Alomone Labs). Immunostained regions were visualized with secondary antibodies fluorescently labeled with Alexa Red and Alexa Green (Molecular Probes). Sections mounted in Vectashield containing DAPI (Vector Labs) were analyzed using a fluorescence microscope (Eclipse E600, Nikon).

Western blot assay.

Dorsal root ganglia were harvested from urethane (1.2 g/kg ip)-anesthetized rats and homogenized in buffer containing 25 mM Tris·HCl, pH 7.6, 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS, 1 mM sodium orthovanadate, PMSF, and protease inhibitor cocktail. Tissue extracts containing equal amounts of proteins (1 mg/ml) were separated on a 10% SDS-polyacrylamide gel and transferred to a polyvinylpyrrolidone difluoride membrane. Membranes were probed with primary polyclonal anti-α_1H antibody (1:200 dilution; catalog no. ACC-025, Alomone Labs). Control membrane was probed with anti-α_1H antibody preincubated with equimolar α_1H control peptide (1:200 dilution) as described above. The immune complexes were visualized using an enhanced chemiluminescence detection system (Amersham).

Carotid body sensory activity.

The protocol for recording sensory activity from ex vivo carotid bodies was essentially the same as that described previously (21). Briefly, the carotid body with the sinus nerve was placed in a recording chamber (250-μl volume) and superfused at a rate of 2.5 ml/min with warm (36°C) physiological saline (in mM: 125 NaCl, 5 KCl, 1.8 CaCl₂, 2 MgSO₄, 1.2 NaH₂PO₄, 25 NaHCO₃, 10 d-glucose, and 5 sucrose). The medium was bubbled with 21% O₂-5% CO₂. Po₂ (141 ± 2.8 mmHg), Pco₂ (35 ± 3 mmHg), and pH (7.36 ± 0.01) were determined by a blood gas analyzer (model ABL 5, Radiometer, Copenhagen, Denmark). To facilitate recording of clearly identifiable action potentials, the sinus nerve was treated with 0.1% collagenase for 5 min. Action potentials (2–4 active units) were recorded from one of the nerve bundles with a suction electrode, amplified (AC preamplifier, 100–3,000 Hz bandwidth; model P511K, Grass Instrument), displayed on an oscilloscope (model 5B12N, Tektronix), and stored in a computer via an analog-to-digital translation board (PowerLab/8P, ADInstruments). “Single” units were selected on the basis of the height and duration of the individual action potentials using a spike discrimination program (Spike Histogram Program, PowerLab, ADInstruments).

Primary cultures of glomus cells.

Primary cultures of glomus cells were prepared essentially as described elsewhere (14, 19). Briefly, carotid bodies were harvested from urethane (1.2 g/kg ip)-anesthetized rats and mice, and glomus cells were dissociated using a mixture of collagenase P (2 mg/ml; Roche), DNase (15 μg/ml; Sigma), and bovine serum albumin (3 mg/ml; Sigma) at 37°C for 20 min and then incubated for 15 min in medium containing DNase (30 μg/ml). Cells were plated on collagen (type VII; Sigma)-coated coverslips and maintained at 37°C in a 7% CO₂-20% O₂ incubator for 24 h. The growth medium consisted of F-12K medium (Invitrogen) supplemented with 1% fetal bovine serum, insulin-transferrin-selenium (ITS-X, Invitrogen), and 1% penicillin-streptomycin-glutamine (Invitrogen).

Measurement of [Ca²⁺]_i.

Glomus cells were incubated in Hanks' balanced salt solution (HBSS) with 2 μM fura 2-AM and 1 mg/ml albumin for 30 min and then washed in a fura 2-free solution for 30 min at 37°C. The coverslip was transferred to an experimental chamber for determination of changes in [Ca²⁺]_i. Background fluorescence at 340- and 380-nm wavelengths was obtained from an area of the coverslip that was devoid of cells. On each coverslip, 5–12 glomus cells were selected (identified by their characteristic clustering) and individual cells were imaged. Image pairs (one at 340 nm and the other at 380 nm) were obtained every 2 s by averaging 16 frames at each wavelength. Data were continuously collected throughout the experiment. Background fluorescence was subtracted from the individual wavelengths. The image obtained at 340 nm was divided by the image at 380 nm to obtain the ratiometric image. Ratios were converted to free [Ca²⁺]_i using calibration curves constructed in vitro by addition of fura 2 (50 μM free acid) to solutions containing known concentrations of Ca²⁺ (0–2,000 nM). The recording chamber was continually superfused with solution from gravity-fed reservoirs.

Measurement of CA secretion.

CA secretion from glomus cells was monitored by amperometry as described previously (14, 19). Briefly, recordings were made from adherent cells that were under constant perfusion with a flow rate of ∼1.0 ml/min (∼80-μl chamber volume). The electrode was held at +700 mV vs. a ground electrode using an NPI VA-10 amplifier to oxidize CAs. The amperometric signal was low-pass-filtered at 2 kHz (8-pole Bessel; Warner Instruments) and sampled into a computer at 10 kHz using a 16-bit analog-to-digital converter (National Instruments). Records with root-mean-square noise >1.0 pA were not analyzed. Amperometric spike features, quantal size, and kinetic parameters were analyzed using a series of macros written in Igor Pro (WaveMetrics). The number of secretory events and the amount of CA secreted per event were analyzed in each experiment, and data are presented as total CAs secreted. All experiments were performed at ambient temperature (23 ± 2°C), and the solutions consisted of (in mM) 137.93 NaCl, 1.26 CaCl₂, 0.4 MgCl₂·6H₂O, 0.49 MgSO₄·7H₂O, 5.33 KCl, 0.44 KH₂PO₄, 0.34 Na₂HPO₄·7H₂O, 5.56 dextrose, and 20 HEPES at pH 7.35 with an osmolarity of 300 mosM. Control normoxic solutions were equilibrated with room air (Po₂ ∼146 mmHg). For hypoxia challenge, solutions were degassed and equilibrated with 1% O₂ balanced with 99% N₂, which resulted in final medium Po₂ of ∼30 mmHg as measured by a blood gas analyzer.

Drugs and chemicals.

All stock solutions were made fresh before the experiments. Mibefradil (Sigma-Aldrich), nifedipine (Sigma-Aldrich), TTA-A2 ([2-(4-cyclopropylphenyl)-N-((1R)-1-[5-[(2,2,2-trifluoroethyl)oxo]-pyridin-2-yl]ethyl)acetamide]; Alomone Labs), and NaHS (Sigma-Aldrich) were obtained from commercial sources. Mibefradil and nifedipine stock solutions were made in HBSS. TTA-A2 stock solution was made in DMSO (Sigma-Aldrich; 0.002% final concentration of DMSO), and the pH of final solution was adjusted to ∼7.4. Stock solutions of NaHS (30 mM) were prepared in HBSS, with pH adjusted to ∼7.4, and were kept on ice. Desired concentrations were added to glomus cell culture plates or to the reservoirs containing medium irrigating the ex vivo carotid body preparation.

Analysis of data.

Average data are presented as means ± SE. Statistical significance was assessed by Student's t-test for statistical comparisons between two groups and one-way ANOVA followed by Tukey's test for multiple-group comparisons. P < 0.05 was considered significant.

RESULTS

T-type VGCC expression in the carotid body.

We first determined the expression of mRNAs encoding α_1H-, α_1G-, and α_1I-subunits of T-type VGCC isoforms in the rat carotid bodies by real-time quantitative PCR. Rat carotid body expressed mRNAs encoding α_1H- α_1G-, and α_1I-subunits, and α_1H mRNA was more abundant than α_1G and α_1I mRNA (P < 0.001; Fig. 1A). Localization of α_1H protein was determined by immunocytochemistry. Specificity of the anti-α_1H antibody was determined by Western blot assay of the rat dorsal root ganglion, which expresses α_1H (28). Immunoblot assay showed a single discrete protein band with a molecular weight corresponding to that of α_1H, and this protein was absent following preincubation of the antibody with the control α_1H peptide (Fig. 1B). Carotid body sections were stained with anti-α_1H and -TH antibodies, the latter of which served as a marker of glomus cells (32). α_1H-like immunoreactivity was seen predominantly in glomus cells, as evidenced by its colocalization with TH-positive cells (Fig. 1C).

Fig. 1.

T-type voltage-gated Ca²⁺ channel (VGCC) expression in rat carotid body. A: real-time RT-PCR analysis of mRNAs encoding α_1H-, α_1G-, and α_1I-subunits of T-type VGCCs in rat carotid bodies. Data were normalized to 18S mRNA and are expressed as fold change. Values are means ± SE from 4 individual experiments performed in triplicate. ***P < 0.001 vs. α_1H. B: immunoblots of rat dorsal root ganglion with anti-α_1H antibodies with (right) and without (left) α_1H control peptide (Pept). Arrow points to a protein band corresponding to molecular weight of α_1H. C: α_1H protein expression in rat carotid body glomus cells. Carotid bodies were stained with antibodies specific for α_1H (red) or tyrosine hydroxylase (TH, green), a marker of glomus cells. Individual staining and merged images are shown. Top and middle: immunostaining with and without α_1H control peptide (Pept+ and Pept−, respectively). Bottom: enlarged (×8) images of glomus cells enclosed in white rectangles. Scale bars = 100 μm.

T-type VGCCs contribute to hypoxia-evoked Ca²⁺ influx in glomus cells.

Hypoxia increases Ca²⁺ influx in glomus cells (10). The role of T-type VGCCs in hypoxia (Po₂ ∼30 mmHg)-induced Ca²⁺ influx was determined in rat glomus cells. Changes in [Ca²⁺]_i were monitored in the presence of increasing concentrations of mibefradil, a pan T-type VGCC blocker (15, 23). Hypoxia increased [Ca²⁺]_i, and this effect was reduced by mibefradil in a dose-dependent manner, with a maximum response at 1 μM (49 ± 14% reduction, P < 0.01; Fig. 2A). However, sodium cyanide (NaCN, 3 μM)-induced Ca²⁺ influx was unaffected by 1 μM mibefradil (Fig. 2B). Increasing mibefradil concentration to 10 μM not only inhibited hypoxia, but also NaCN-evoked Ca²⁺ influx (Fig. 2, A and B), suggesting that >1 μM mibefradil exerts nonselective action. Consequently, further studies were performed with 1 μM mibefradil, which selectively affected hypoxia-evoked Ca²⁺ influx.

Fig. 2.

T-type VGCC blockers attenuate hypoxia-induced elevation of intracellular Ca²⁺ concentration ([Ca²⁺]_i) in glomus cells of rat carotid body. A: [Ca²⁺]_i response of glomus cells treated with 0.01–10 μM mibefradil (Mib), a blocker of T-type VGCCs, to hypoxia (Po₂ ∼30 mmHg). Changes in [Ca²⁺]_i (Δ[Ca²⁺]_i) reflect response to hypoxia minus baseline levels. Values are means ± SE of number of cells shown in parentheses. **P < 0.01 vs. untreated controls. B: [Ca²⁺]_i response to 3 μM NaCN. Δ[Ca²⁺]_i represents response to NaCN minus baseline levels in the presence of vehicle (Veh) or mibefradil. Values are means ± SE of number of cells shown in parentheses. **P < 0.01 vs. Veh; ns, not significantly different (P > 0.05). C and E: representative examples of [Ca²⁺]_i response of glomus cells to hypoxia (Hx; Po₂ ∼30 mmHg) in the presence of vehicle or 1 μM mibefradil (C) or 25 μM TTA-A2 (TTA, E). Horizontal bars show duration of challenges with hypoxia or vehicle, mibefradil, or TTA-A2. D and F: [Ca²⁺]_i response to hypoxia presented as Δ[Ca²⁺]_i, i.e., response to hypoxia minus baseline levels in the presence of vehicle or 1 μM mibefradil (D) or 25 μM TTA-A2 (F). Wash, 5 min after washout of mibefradil or TTA-A2. Values are means ± SE of number of cells shown in parentheses. **P < 0.01 vs. Veh.

The effects of 1 μM mibefradil were completely reversed within 5 min after washout of the drug (Fig. 2, C and D). The residual hypoxia-induced [Ca²⁺]_i elevation following mibefradil was blocked by 80 ± 11% (P < 0.01) in the presence of 5 μM nifedipine, an L-type VGCC blocker. To further establish the role of T-type VGCCs, we examined the effects of TTA-A2, another selective T-type VGCC blocker (8, 9, 24). On the basis of preliminary studies, we tested the effects of 25 μM TTA-A2. As shown in Fig. 2, E and F, TTA-A2 reversibly attenuated [Ca²⁺]_i responses to hypoxia (41 ± 16% reduction, P < 0.01).

T-type VGCCs contribute to hypoxia-evoked CA secretion and carotid body sensory nerve activity.

Hypoxia releases CAs from glomus cells in a Ca²⁺-dependent manner (10). CA secretion from individual glomus cells was monitored by amperometry in the presence of either 1 μM mibefradil or 25 μM TTA-A2. Hypoxia-evoked CA secretion was markedly reduced in the presence of mibefradil or TTA-A2 (Fig. 3).

Fig. 3.

Effect of T-type VGCC blockers on hypoxia-evoked catecholamine (CA) secretion from glomus cells. Glomus cells were isolated from carotid bodies harvested from anesthetized rats, and CA secretion was monitored by amperometry. A and C: hypoxia (Po₂ ∼30 mmHg)-evoked CA secretion from rat glomus cells in the presence of vehicle or 1 μM mibefradil or 25 μM TTA-A2. Horizontal bars show duration of challenges with hypoxia or vehicle, mibefradil, or TTA-A2. B and D: total number of CA molecules secreted during hypoxia. Values are means ± SE of number of cells shown in parentheses. **P < 0.01 vs. Veh.

We then tested the effects of T-type VGCC blockers on the carotid body sensory nerve response to hypoxia. These experiments utilized an ex vivo carotid body preparation to exclude the confounding influence of T-type VGCC blockers on blood pressure in intact rats. Hypoxia resulted in an increase in sensory nerve activity, which was markedly attenuated in the presence of mibefradil (1 μM) or TTA-A2 (25 μM) in a reversible manner (Fig. 4).

Fig. 4.

Effect of T-type Ca²⁺ channel blockers on carotid body sensory response to hypoxia. Carotid bodies were harvested from anesthetized rats, and sensory nerve activity was recorded ex vivo. A and C: representative examples of sensory nerve responses [impulses (imp)/s] to hypoxia (Po₂ ∼30 mmHg) in the presence of vehicle or 1 μM mibefradil (A) or 25 μM TTA-A2 (C) and 5 min after washout of drugs. Horizontal bars show duration of challenges. Inset: superimposed action potentials of a “single” fiber from which data were derived. B and D: effects of 1 μM mibefradil or TTA-A2 on sensory nerve response to hypoxia. Data are presented as change (Δimp/s), i.e., stimulus-evoked activity minus baseline activity. Values are means ± SE of number of experiments shown in parentheses. **P < 0.01 vs. Veh.

Selective contribution of Ca_V3.2 to the glomus cell and carotid body response to hypoxia.

Since glomus cells express α_1H, the pore-forming subunit of the Ca_V3.2 subtype of T-type VGCC (Fig. 1), we sought to determine the selective role of Ca_V3.2 in the carotid body response to hypoxia. Studies were performed on mice with homozygous deletion of the α_1H allele (Cacna1h^−/−), which exhibit attenuated Ca_V3.2 T-type Ca²⁺ current (4).

Hypoxia-evoked [Ca²⁺]_i elevation and CA secretion were significantly reduced in glomus cells from Cacna1h^−/− mice compared with wild-type (WT) controls (Fig. 5). On average, [Ca²⁺]_i and CA secretory responses were attenuated by 63 ± 14.6% and 55 ± 4.0%, respectively, in cells from Cacna1h^−/− mice (P < 0.01; Fig. 5, B and D). Similarly, carotid bodies from Cacna1h^−/− mice exhibited a markedly attenuated sensory nerve response to hypoxia compared with WT controls (P < 0.01; Fig. 6, A and B). However, NaCN-evoked sensory nerve excitation was comparable in carotid bodies from Cacna1h^−/− and WT mice (Fig. 6C).

Fig. 5.

Glomus cell [Ca²⁺]_i and CA secretory responses to hypoxia in wild-type (WT) and Ca_V3.2 knockout (Cacna1h^−/−) mice. A: representative examples of [Ca²⁺]_i responses of glomus cells from WT and Cacna1h^−/− mice to hypoxia (Po₂ ∼30 mmHg). B: [Ca²⁺]_i responses to hypoxia, presented as change (Δ), i.e., stimulus-evoked response minus baseline levels. Values are means ± SE of number of cells shown in parentheses. C: representative examples of hypoxia (Po₂ ∼30 mmHg)-evoked CA secretion from glomus cells from WT and Cacna1h^−/− mice. Horizontal bars in A and C show duration of hypoxic challenge. D: total number of CA molecules secreted during hypoxia. Values are means ± SE of number of cells shown in parentheses. **P < 0.01 vs. WT.

Fig. 6.

Carotid body (CB) sensory response to hypoxia in WT and Cacna1h^−/− mice. A: representative examples of carotid body sensory nerve responses to hypoxia (Po₂ ∼30 mmHg) in WT and Cacna1h^−/− mice. Horizontal bars show duration of hypoxic challenge. Inset: superimposed action potentials of a “single” fiber from which data were derived. B and C: carotid body sensory nerve response to hypoxia (B) or 3 μM NaCN (C) presented as change (Δ), i.e., stimulus-evoked activity minus baseline activity. Values are means ± SE of number of experiments shown in parentheses. **P < 0.01 vs. WT.

H₂S is required for the Ca_V3.2-dependent carotid body response to hypoxia.

Recent studies reported that CSE-derived H₂S is a critical mediator of the carotid body response to hypoxia (12, 14, 20, 27). We hypothesized that the Ca_V3.2-dependent carotid body response to hypoxia requires H₂S. This possibility was tested in WT and Cacna1h^−/− mice. In WT mice, NaHS, a H₂S donor, like hypoxia, increased [Ca²⁺]_i in glomus cells and carotid body sensory nerve activity, and these responses were markedly reduced in glomus cells and carotid bodies from Cacna1h^−/− mice (Fig. 7). To assess the selective role of CSE-derived H₂S, studies were performed on WT and CSE^−/− mice. Hypoxia increased [Ca²⁺]_i in glomus cells, and this response was attenuated by TTA-A2 in glomus cells from WT, but not CSE^−/−, mice (Fig. 8, A and B). To determine whether genetic deletion of CSE impaired T-type VGCC function, glomus cells from CSE^−/− mice were challenged with NaHS. In response to NaHS, [Ca²⁺]_i increased, and this effect was inhibited by TTA-A2 in a reversible manner (Fig. 8, C and D). Hypoxia-evoked carotid body sensory nerve excitation was reduced by TTA-A2 in carotid bodies from WT, but not CSE^−/−, mice (Fig. 9).

Fig. 7.

Glomus cell and carotid body responses to NaHS in WT and Cacna1h^−/− mice. A: representative examples of [Ca²⁺]_i responses to 50 μM NaHS and hypoxia (Po₂ ∼30 mmHg) in glomus cells from WT and Cacna1h^−/− mice. B: [Ca²⁺]_i responses to 50 μM NaHS and hypoxia in glomus cells from WT and Cacna1h^−/− mice, analyzed as change (Δ), i.e., stimulus-evoked response minus baseline levels. Values are means ± SE of number of cells shown in parentheses. C: representative examples of carotid body sensory nerve responses to 50 μM NaHS and hypoxia (Po₂ ∼30 mmHg) in WT and Cacna1h^−/− mice. Horizontal bars in A and C show duration of exposure to NaHS or hypoxia. D: carotid body sensory nerve response to NaHS or hypoxia, analyzed as change (Δ), i.e., stimulus-evoked activity minus baseline activity. Values are means ± SE of number of cells shown in parentheses. **P < 0.01, *P < 0.05 vs. WT.

Fig. 8.

Effect of TTA-A2 on [Ca²⁺]_i responses of glomus cells from WT and cystathionine-γ-lyase (CSE) null (CSE^−/−) mice to hypoxia and H₂S. A and C: representative examples of [Ca²⁺]_i responses to hypoxia (A) or 50 μM NaHS (C) in the presence of vehicle or 25 μM TTA-A2 in glomus cells from WT and CSE^−/− mice. Horizontal bars show duration of exposure to NaHS, hypoxia, vehicle, and TTA-A2. B and D: [Ca²⁺]_i responses of glomus cells from WT and CSE^−/− mice to hypoxia (B) or 50 μM NaHS (D), presented as change (Δ), i.e., stimulus-evoked response minus baseline levels. Values are means ± SE of number of cells shown in parentheses. **P < 0.01 vs. Veh.

Fig. 9.

Effect of TTA-A2 on carotid body sensory nerve responses to hypoxia in WT and CSE^−/− mice. A: representative examples of sensory nerve responses to hypoxia (Po₂ ∼30 mmHg) in WT and CSE^−/− mice in the presence of vehicle or TTA-A2. Horizontal bars show duration of hypoxia challenge, vehicle, or TTA-A2. Inset: superimposed action potentials of a “single” fiber from which data were derived. B: carotid body sensory response to hypoxia analyzed as change (Δ), i.e., stimulus-evoked activity minus baseline activity. Values are means ± SE of number of experiments shown in parentheses. **P < 0.01 vs. Veh.

DISCUSSION

The results of the present study demonstrate that carotid bodies express T-type VGCCs and establish their functional significance in O₂ sensing. We found a higher abundance of α_1H mRNA in carotid bodies from Sprague-Dawley rats as opposed to relatively high levels of α_1G mRNA expression in Wistar rat carotid bodies as reported previously (5). Our data further demonstrate that much of the α_1H protein is localized to glomus cells, the primary O₂-sensing cells in the carotid body. The following findings demonstrate the functional significance of T-type VGCCs in carotid body O₂ sensing. 1) Mibefradil and TTA-A2, the two established blockers of T-type VGCCs (15, 23, 24), effectively reduced hypoxia-induced Ca²⁺ influx in glomus cells by ∼50%, and these effects were reversible and selective to hypoxia, because Ca²⁺ influx by NaCN was unaffected. 2) T-type VGCC blockers were equally effective in reducing hypoxia-evoked CA secretion from glomus cells, as well as carotid body sensory nerve excitation. Previous studies reported that L-type VGCCs are the major contributors to Ca²⁺ influx by hypoxia in glomus cells (1, 14). Consistent with this possibility, we found that nifedipine, an L-type VGCC blocker, abolished ∼80% of the residual hypoxia-evoked Ca²⁺ influx following mibefradil. Together, these observations demonstrate that hypoxia-induced voltage-gated Ca²⁺ influx in glomus cells requires high-threshold (L-type) and low-threshold (T-type) VGCCs.

Mice with genetic absence of α_1H (Cacna1h^−/−) exhibited markedly attenuated glomus cell responses to hypoxia as well as carotid body sensory nerve excitation by hypoxia, whereas carotid body responses to NaCN were unaltered in Cacna1h^−/− mice. Furthermore, α_1H is predominantly expressed in glomus cells. Since α_1H constitutes the pore-forming subunit of Ca_V3.2, these observations demonstrate that Ca_V3.2 is the major T-type VGCC that contributes to hypoxia-evoked voltage-gated Ca²⁺ influx, neurotransmitter secretion, and carotid body sensory nerve excitation.

The following observations suggest that CSE-derived H₂S is required for Ca_V3.2-dependent carotid body responses to hypoxia. 1) Like hypoxia, NaHS, a H₂S donor, increased Ca²⁺ influx in glomus cells and carotid body sensory nerve activity, and these effects were absent in Cacna1h^−/− mice. 2) The inhibitory effects of TTA-A2 on glomus cell and carotid body sensory nerve responses were strikingly absent in CSE^−/− mice. The lack of TTA-A2 effect in CSE^−/− mice is unlikely due to compromised Ca_V3.2 function, because a H₂S donor still increased Ca²⁺ influx, which could be attenuated by TTA-A2. We previously reported that L-type VGCCs contribute in part to the H₂S-mediated hypoxic sensing by the carotid body (14). The current findings suggest that, in addition to L-type VGCCs, H₂S-dependent hypoxia sensing also requires Ca_V3.2, analogous to their proposed role in nociception (17). However, the effects of CSE-derived H₂S on glomus cell Ca_V3.2 currents remain to be studied.

In summary, the present study employing pharmacological and genetic approaches establishes a role for T-type VGCCs, especially Ca_V3.2, in the carotid body sensory response to acute hypoxia. The carotid body response to hypoxia is augmented by chronic intermittent hypoxia (CIH), mimicking the O₂ saturation profiles encountered with recurrent apnea (22). Given that CIH upregulates Ca_V3.2 in adrenal medullary chromaffin cells (25), it would be of interest to examine in future studies whether T-type VGCCs contribute to CIH-evoked heightened hypoxic sensitivity of the carotid body.

GRANTS

This research was supported by National Heart, Lung, and Blood Institute Grant HL-90554.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

V.V.M. and N.R.P. are responsible for conception and design of the research; V.V.M., Y.-J.P., G.Y., and J.N. performed the experiments; V.V.M., Y.-J.P., G.Y., and J.N. analyzed the data; V.V.M. and A.P.F. interpreted the results of the experiments; V.V.M., Y.-J.P., and J.N. prepared the figures; V.V.M. and N.R.P. drafted the manuscript; A.P.F., G.K.K., and N.R.P. edited and revised the manuscript; N.R.P. approved the final version of the manuscript.