Ca²⁺ oscillations in rat carotid body type 1 cells in normoxia and hypoxia

Abstract

We studied the mechanisms by which carotid body glomus (type 1) cells produce spontaneous Ca²⁺ oscillations in normoxia and hypoxia. In cells perfused with normoxic solution at 37°C, we observed relatively uniform, low-frequency Ca²⁺ oscillations in >60% of cells, with each cell showing its own intrinsic frequency and amplitude. The mean frequency and amplitude of Ca²⁺ oscillations were 0.6 ± 0.1 Hz and 180 ± 42 nM, respectively. The duration of each Ca²⁺ oscillation ranged from 14 to 26 s (mean of ∼20 s). Inhibition of inositol (1,4,5)-trisphosphate receptor and store-operated Ca²⁺ entry (SOCE) using 2-APB abolished Ca²⁺ oscillations. Inhibition of endoplasmic reticulum Ca²⁺-ATPase (SERCA) using thapsigargin abolished Ca²⁺ oscillations. ML-9, an inhibitor of STIM1 translocation, also strongly reduced Ca²⁺ oscillations. Inhibitors of L- and T-type Ca²⁺ channels (Ca_v; verapamil>nifedipine>TTA-P2) markedly reduced the frequency of Ca²⁺ oscillations. Thus, Ca²⁺ oscillations observed in normoxia were caused by cyclical Ca²⁺ fluxes at the ER, which was supported by Ca²⁺ influx via Ca²⁺ channels. Hypoxia (2–5% O₂) increased the frequency and amplitude of Ca²⁺ oscillations, and Ca_v inhibitors (verapamil>nifedipine>>TTA-P2) reduced these effects of hypoxia. Our study shows that Ca²⁺ oscillations represent the basic Ca²⁺ signaling mechanism in normoxia and hypoxia in CB glomus cells.

INTRODUCTION

Carotid body (CB) glomus cells are peripheral chemoreceptors that convert the arterial O₂ pressure signal to changes in cell membrane potential (E_m) that regulate the activity of the voltage-dependent Ca²⁺ channel (Ca_v) and [Ca²⁺]_i. The rise in [Ca²⁺]_i is critical for increasing the glomus cell secretory activity, and therefore, changes in [Ca²⁺]_i have been measured to indirectly assess the activity of glomus cells. Studies in isolated glomus cells have shown that the rise in mean [Ca²⁺]_i produced by graded levels of hypoxia follows a hyperbolic function (11, 34, 43). The averaged [Ca²⁺]_i response is very low at mild/moderate levels of hypoxia (Po₂ of perfusion solution; 35–100 mmHg), but increases steeply at severe levels of hypoxia (Po₂ < 30 mmHg) (8, 10, 31, 35, 47). Respiratory diseases such as chronic obstructive pulmonary disease, emphysema, and sleep apnea produce mild and moderate levels of hypoxia, where arterial Po₂ is reduced from a normal level of ∼100 mmHg to 72–78 mmHg (16, 22). Thus, it is important to understand how mild and moderate levels of hypoxia regulate the Ca²⁺ signaling processes in glomus cells.

In many cell types, agonists elevate [Ca²⁺]_i as a part of the signal transduction pathway to regulate cell function. In such cells, high agonist concentrations elicit a sustained increase in [Ca²⁺]_i, whereas low agonist concentrations elicit oscillations in [Ca²⁺]_i (26, 40). For example, a low-level stimulus increases the frequency of Ca²⁺ oscillations to regulate the cellular activity in various cell types such as pancreatic β-cells, adrenal cells, pituitary gonadotrophs, and airway smooth muscle cells (6, 17, 24, 26). Such Ca²⁺ oscillations are considered to be important for low-level signaling, for preventing desensitization, and for providing efficacy and specificity of information flow. In glomus cells, few and random fluctuations in averaged [Ca²⁺]_i have been observed in earlier studies (8, 33). Whether such fluctuations in [Ca²⁺]_i represent low-level Ca²⁺ oscillations and how they arise are not known. Search of the literature indicated no published studies on Ca²⁺ oscillations in glomus cells. In our attempts to better understand the nature of spontaneous Ca²⁺ fluctuations in glomus cells, we found that a large fraction of glomus cells generated relatively uniform low-frequency Ca²⁺ fluctuations, suggesting that Ca²⁺ signaling in glomus cells may also involve Ca²⁺ oscillations, similar to those in other cell types.

In the present study, we characterized the kinetics of Ca²⁺ oscillations and investigated the roles of Ca²⁺ channels at the plasma membrane and endoplasmic reticulum (ER) in the generation of Ca²⁺ oscillations in normoxia. We studied the effects of different experimental conditions such as flow rate and perfusion temperature on the kinetics of Ca²⁺ oscillations. We then studied the effects of mild and moderate levels of hypoxia on the kinetics of Ca²⁺ oscillations and the role of Ca²⁺ channels in this process. Our results showed that low-frequency Ca²⁺ oscillations that occur in glomus cells in normoxia involve Ca²⁺ fluxes both at the ER and plasma membrane. Mild and moderate levels of hypoxia augmented the frequency and amplitude of Ca²⁺ oscillations by increasing Ca²⁺ influx via Ca_v. These findings show that Ca²⁺ oscillations most likely represent the physiological Ca²⁺ signaling processes that underlie glomus cell activity in normoxia as well as under mild and moderate levels of hypoxia. Some of the preliminary findings were presented at the Experimental Biology meeting (28).

METHODS

Animals and ethical approval.

All experiments were carried out according to the guidelines set by the Animal Care and Use Committee of Rosalind Franklin University under Institutional Animal Care and Use Committee protocol no. 18‐02, and conform to the principles and regulations as described by Grundy (20). The animal care and use program is Association for Assessment and Accreditation of Laboratory Animal Care‐accredited and approved by the Office of Laboratory Animal Welfare (A3279‐01). There are no ethical concerns. Rats (Sprague-Dawley, postnatal days 16–24) purchased from Envigo (Indianapolis, IN) were acclimatized in house for at least ∼1 wk. Standard laboratory diet (Teklad Global 19% Protein Extruded Rodent Diet, Harlan no. 2019S) and drinking water were provided.

Cell isolation.

Rats (male and female) were deeply anaesthetized by inhalation of isoflurane until cessation of breathing and decapitated with a sharp razor blade. Six to eight CBs from the region of the carotid artery bifurcation were removed from three to four rats and placed in ice-cold, low-Ca²⁺, low-Mg²⁺ phosphate-buffered saline (low-Ca²⁺/Mg²⁺ PBS: 137 mM NaCl, 2.8 mM KCl, 2 mM KH₂PO₄, 0.07 mM CaCl₂, and 0.05 mM MgCl₂, pH 7.4). Each CB was cut into three to four pieces and placed in a solution containing trypsin (0.4 mg/mL) and collagenase (0.4 mg/mL) in low-Ca²⁺/Mg²⁺ PBS and incubated at 37°C for 25 min. Following trituration using a fire-polished glass pipette, the dispersed cells were separated by centrifugation at low speed (1,200 g for 5 min) and suspended in CB growth medium. CB growth medium contained Ham’s F-12, 10% fetal bovine serum, 23 mM glucose, 4 mM Glutamax-I (l-alanyl glutamine), 10 kU penicillin, 10 kU streptomycin, and 300 μg/mL insulin. Cells were plated on to poly-d-lysine-pretreated glass coverslips and incubated at 37°C for ∼3 h in a humidified atmosphere of 95% air-5% CO₂. Cells were used the same day within 6 h after plating.

[Ca²⁺] measurement.

Isolated cells plated on glass coverslips were incubated with 2 μM fura-2 AM for 30 min at 37°C in culture medium. A coverslip with attached cells was placed in a recording chamber positioned on the stage of an inverted microscope (IX71; Olympus America Inc., Center Valley, PA). Fura-2 was alternately excited at 340 and 380 nm, and the emitted fluorescence was filtered at 510 nm and recorded using a charge-coupled device (CCD)-based imaging system running SimplePCI software (Hamamatsu Corp.). The cells in the recording chamber were continuously perfused with a solution containing (in mM) 117 NaCl, 5 KCl, 23 NaHCO₃, 1 MgCl₂, 1 CaCl₂ and 11 glucose (pH 7.3). Ratiometric data were calibrated by applying experimentally determined constants to the following equation: [Ca²⁺] = K_d × β × (R – R_min)/(R_max – R) (21). Values for R_max (12.0), R_min (0.1), and β (11.6) were determined in vitro, and a K_d value of 300 nM for fura-2 was assumed. Each experiment was performed using coverslips of cells prepared on different days, and no more than two coverslips were used for each day.

Hypoxia studies.

In our experiments that used a recording chamber that was open to the atmosphere, we generated different levels of hypoxic solutions (ranging from approximately mild hypoxia to anoxia) by bubbling solutions with 5%CO₂-95%N₂ with 2 U/mL of glucose oxidase and 100 U/mL of catalase added to the solution (anoxia; see Refs. 5 and 29), 5% CO₂-1% O₂-94% N₂ (severe hypoxia), 5% CO₂-2% O₂-93% N₂ (moderate hypoxia), 5%CO₂-5%O₂-90%N₂ (mild hypoxia), or 5%CO₂-10%O₂-85%N₂. A reproducible level of [O₂] in the perfusion chamber was produced for each level of hypoxia, as judged by %O₂ measurements using an O₂ meter (ISO2; World Precision Instruments, Sarasota, FL). After a steady-state basal [Ca²⁺]_i was obtained with normoxic solution, the perfusion solution was switched to a desired hypoxic solution. The temperature of the perfusion solutions was maintained at 37.0 ± 0.3°C. In three trials, the mean O₂ levels in solutions in the recording chamber were 0.4% (3 mmHg) for anoxia, 1.6% (12 mmHg) for severe hypoxia, 2.3% (17 mmHg) for moderate hypoxia, and 5.2% (40 mmHg) for mild hypoxia. The O₂ meter was calibrated to 0% with solution bubbled with pure nitrogen for 60 min and to 21% with solution gassed with air for 30 min at 37°C.

Materials and data analysis.

2-APB, thapsigargin and ML-9 [1-(5-chloronaphthalenesulfonyl)-homopiperazine HCl] were from Tocris. Fura-2 AM was from Thermofisher Scientific. Catalase and glucose oxidase were from Sigma-Aldrich Co. All other chemicals for preparing perfusion solutions were from Sigma-Aldrich Co. Student’s t-test (for comparison of 2 sets of data) and one-way analysis of variance (comparison of 3 or more sets of data) were used for data analysis using PRISM software. Post hoc testing following analysis of variance was based on unpaired t-test with Bonferroni correction. P < 0.05 was considered significant. Data are presented as box plots when the total number of cells analyzed is large (n > 40). For other plots, means ± SD are shown. Box plots show four quartile groups (25% each) and median and mean (solid circle) values.

RESULTS

Spontaneous Ca²⁺ oscillations in normoxia.

In the first set of experiments, cells were perfused at 30°C, rather than at 37°C, to minimize the appearance of spontaneous Ca²⁺ fluctuations from the basal state so that [Ca²⁺]_i responses produced by high KCl and hypoxia could be identified clearly. In all six coverslips of cells tested, cells whose [Ca²⁺]_i increased in response to 20 mM KCl were also sensitive to severe hypoxia and anoxia (Fig. 1A). Expanded tracings of the sustained increase in [Ca²⁺]_i by anoxia are shown in Fig. 1B. Cells that did not respond to 20 mM KCl were insensitive to hypoxia and anoxia. Therefore, 20 mM KCl was used to identify hypoxia-sensitive type 1 (glomus) cells throughout the study.

Fig. 1.

Sustained increase in intracellular calcium concentration ([Ca²⁺]_i) produced by high KCl and anoxia in isolated glomus cells. A: [Ca²⁺]_i responses to 20 mM KCl, severe hypoxia, and anoxia. Perfusion temperature is 30°C. B: [Ca²⁺]_i response to anoxia at an expanded scale.

In isolated cells perfused with normoxic solution at a flow rate of 2 mL/min (velocity of ∼20 cm/min; see below) at 37°C, we observed variable levels of fluctuations in [Ca²⁺]_i in many cells. Fluctuations in basal [Ca²⁺]_i have been observed previously, but the signals were generally averaged, and thus their kinetic properties in individual cells remain poorly defined.

The kinetics of such fluctuations in basal [Ca²⁺]_i in individual cells were examined in detail. A representative recording from a coverslip of cells is shown in Fig. 2A (top tracing). The cell population could be divided into three types based on their response to KCl and appearance of Ca²⁺ fluctuations (Fig. 2, B–D). Cells that were not sensitive to 20 mM KCl did not show spontaneous Ca²⁺ fluctuations and were assumed to be mostly type II cells (Fig. 2B). In cells that responded to KCl, some cells showed no Ca²⁺ fluctuations (Fig. 2C), while others showed varying degrees of Ca²⁺ fluctuations in both frequency and amplitude. Ca²⁺ recordings in KCl-sensitive cells from two coverslips of cells are shown in Fig. 2D. Ca²⁺ fluctuations were present before application of 20 mM KCl, indicating that high KCl itself was not the trigger for the generation of Ca²⁺ oscillations.

Fig. 2.

Spontaneous Ca²⁺ oscillations in normoxia. A: all cells on a coverslip in normoxic solution containing 5 mM KCl (n = 34 cells). KCl (20 mM) is applied briefly at the beginning. B: cells that are insensitive to 20 mM KCl. C: cells that are sensitive to 20 mM KCl but without Ca²⁺ oscillations. D: cells that are sensitive to 20 mM KCl and show Ca²⁺ oscillations. Recordings are from two coverslips of cells. E: Ca²⁺ oscillations from individual cells from different coverslips at expanded scale. F: frequency and amplitude levels (8 coverslips; n = 160 cells).

The percentage of cells in each coverslip showing clearly identifiable Ca²⁺ fluctuations (>50 nM above the resting level) ranged from ∼30 to ∼85% (average of ∼65%; 32 coverslips of cells). Figure 2E shows examples of Ca²⁺ fluctuations in individual cells, illustrating the varying levels of frequency and amplitude. Each cell generated its own Ca²⁺ fluctuations with intrinsic frequency and amplitude that were generally uniform and sustained during the perfusion. Therefore, we will refer to such Ca²⁺ fluctuations as Ca²⁺ oscillations from hereon.

Figure 2F plots the frequency and amplitude of Ca²⁺ oscillations from individual cells from eight coverslips of cells (n = 160 cells). In cells showing Ca²⁺ oscillations, mean frequency and amplitude were 0.6 ± 0.1 min and 180 ± 42 nM (above the baseline of ∼100 nM), respectively. If all KCl-responding cells, including those that show no clearly identifiable Ca²⁺ oscillations, are taken into account, the mean frequency and amplitude levels would be significantly lower and depend on the coverslip of cells used for each experiment. Under our experimental conditions, the average duration of Ca²⁺ oscillations was ∼20 s (range 14–26 s; n = 16). These findings show that low-frequency Ca²⁺ oscillations are spontaneously generated in a substantial population of isolated glomus cells perfused at 37°C with a normoxic solution.

Temperature and flow rate effects on Ca²⁺ oscillations.

We observed a strong temperature dependence of Ca²⁺ oscillations in glomus cells, as illustrated in two coverslips of cells (Fig. 3A). At 24°C, Ca²⁺ oscillations were generally absent. As the temperature of the perfusion solution was increased to 30°C, a few cells started to show oscillations. The number of cells showing Ca²⁺ oscillations continued to increase as the perfusion temperature was elevated to 34°C and 37°C. Both the frequency and amplitude of Ca²⁺ oscillations were increased by heat (Fig. 3B). In the studies of Ca²⁺ oscillations described below, the perfusion temperature in the recording chamber was maintained at 37°C.

Fig. 3.

Perfusion temperature and Ca²⁺ oscillations. A: intracellular calcium concentration ([Ca²⁺]_i) recording in KCl-sensitive cells at different bath temperatures. B: frequency and amplitude of Ca²⁺ oscillations at four temperatures (4 coverslips of cells; n = 88) are all significantly different from each other (P < 0.01).

We also observed a significant effect of flow rate of the perfusion solution on the kinetics of Ca²⁺ oscillations. With the temperature of the standing solution maintained at ∼37°C using a heated recording chamber and zero flow, no Ca²⁺ oscillations were observed (Fig. 4A). Upon changing the perfusion to a rate of ∼2 ml/min while maintaining the temperature at 37°C, Ca²⁺ oscillations were quickly elicited in many cells. The estimated velocity of perfusion solution was ∼20 cm/min, a value calculated based on the cross-sectional area (2 mm deep × 5 mm wide) of ∼10 mm² for solution moving through the recording chamber. When the flow rate was switched from 1 to 3 mL/min, a rate-dependent increase in the frequency of Ca²⁺ oscillations was clearly present (Fig. 4B). Flow-dependent increase in oscillation frequency is shown in Fig. 4C. These findings show that the perfusion flow rate regulates the frequency of Ca²⁺ oscillations and suggest the possibility that a mechanism that senses mechanical force may be present in glomus cells. A perfusion flow rate of 2 mL/min was used for studies described below, unless indicated otherwise.

Fig. 4.

Flow rate and Ca²⁺ oscillations. A: intracellular calcium concentration ([Ca²⁺]_i) in KCl-sensitive cells on a coverslip at flow rates of 0 and 2 mL/min. Bath temperature is kept at 37°C. B: [Ca²⁺]_i in KCl-sensitive cells on a coverslip at flow rate of 1–3 mL/min. C: frequency of Ca²⁺ oscillations at 4 different flow rates (4 coverslips of cells; n = 53). All values were significantly different from each other (P < 0.01), except those between 2 and 3 mL/min (P > 0.05).

Inhibition of ER inositol (1,4,5)-trisphosphate receptor and store-operated Ca²⁺ entry on Ca²⁺ oscillations.

Ca²⁺ oscillation is known to occur as a result of Ca²⁺ release from and uptake by ER, which is modulated by other Ca²⁺ signals such as Ca²⁺ influx via Ca_v and store-operated Ca²⁺ entry (SOCE) (17). To determine the mechanism of Ca²⁺ oscillations in glomus cells, we first tested the effect of 2-APB that inhibits inositol (1,4,5)-trisphosphate (IP₃)-mediated Ca²⁺ release from ER and SOCE. Cells were perfused with normoxic solution at 37°C until a steady-state level of Ca²⁺ oscillations was achieved. In all five coverslips (n = 73 cells), switching to a solution that contained 100 μM 2-APB caused an immediate cessation of Ca²⁺ oscillations that fully recovered following washout of the drug (Fig. 5A), indicating that Ca²⁺ signaling by ER/SOCE is critical for the generation of Ca²⁺ oscillations.

Fig. 5.

Effects of 2-aminoethoxydiphenyl borate (2-APB), thapsigargin, and ML-9 on Ca²⁺ oscillations. A: reversible inhibition of Ca²⁺ oscillations by 2-APB. B: slow and irreversible inhibition of Ca²⁺ oscillations by thapsigargin. C: reversible inhibition of Ca²⁺ oscillations by ML-9.

Thapsigargin (1 μM), an irreversible inhibitor of ER Ca²⁺-ATPase, produced a gradual and complete inhibition of Ca²⁺ oscillations in all four coverslips (n = 63 cells; Fig. 5B). Application of ML-9 (50 μM), an agent that interferes with STIM1 translocation and thereby inhibits SOCE (41), produced a reversible block of Ca²⁺ oscillations in normoxia in all four coverslips (n = 66 cells; Fig. 5C). Therefore, these findings indicate that Ca²⁺ oscillations in glomus cells are mediated by ER/SOCE-dependent mechanisms, similar to those found in other secretory cells.

Ca²⁺ influx and Ca²⁺ oscillations in normoxia.

In many cell types that exhibit Ca²⁺ oscillations, Ca²⁺ influx via the plasma membrane regulates the frequency of Ca²⁺ oscillations by altering “the Ca²⁺ load” in the cell (42). In glomus cells showing Ca²⁺ oscillations, lowering [Ca²⁺]_o from 1 to ∼0 mM quickly abolished Ca²⁺ oscillations in all cells and the addition of 1 mM Ca²⁺ restored Ca²⁺ oscillations (Fig. 6A), indicating that Ca²⁺ influx across the plasma membrane is necessary to sustain the basal level of Ca²⁺ oscillations in normoxia. Nifedipine (1 μM) that blocks the L-type Ca_v produced a strong reversible inhibition of both frequency and amplitude of Ca²⁺ oscillations (Fig. 6B). Verapamil (100 μM), a nonspecific Ca_v inhibitor, reversibly abolished Ca²⁺ oscillations, supporting the critical role of Ca²⁺ influx via Ca_v in supporting Ca²⁺ oscillations (Fig. 6C). A summary of the effects of nifedipine on the frequency and amplitude of Ca²⁺ oscillations is shown in Fig. 6D.

Fig. 6.

Ca²⁺ influx and Ca²⁺ oscillations. A: removal of external Ca²⁺ on intracellular calcium concentration ([Ca²⁺]_i) oscillations. B: nifedipine effect on Ca²⁺ oscillations. C: cessation of Ca²⁺ oscillations by verapamil (3 coverslips; n = 43 cells). D: nifedipine effects on frequency and amplitude of Ca²⁺ oscillations (means ± SD; 6 coverslips of cells). *Significantly different (P < 0.01).

Interestingly, TTA-P2 (a T-type Ca_v inhibitor; see Ref. 14) also caused a significant inhibition of the frequency and amplitude of Ca²⁺ oscillations (Fig. 7, A and B). The inhibitory effect of TTA-P2 on Ca²⁺ oscillations was as strong as that of nifedipine (Fig. 7, C and D). To be sure that the inhibition of Ca²⁺ oscillations by TTA-P2 was not due to block of the L-type Ca_v, we tested the effect of TTA-P2 on FPL-64176-induced increase in [Ca²⁺]_i. FPL-64176 is a potent activator of the L-type Ca_v and elicits a strong, nifedipine-sensitive increase in [Ca²⁺]_i (27). As predicted, nifedipine produced a 92 ± 5% inhibition of FPL-induced increase in [Ca²⁺]_i (3 coverslips; n = 48 cells; Fig. 7E). However, TTA-P2 produced no significant effect in any of the three coverslips (n = 52 cells), showing that TTA-P2 (20 μM) did not block the L-type Ca_v (Fig. 7F). ML-218, another T-type Ca_v inhibitor, also reduced the frequency of basal Ca²⁺ oscillations by 62 ± 6% (n = 5 coverslips; 78 cells). These results indicate that Ca²⁺ influx via both L- and T-type Ca_v occurs in resting isolated glomus cells and is important for the generation of basal Ca²⁺ oscillations in normoxia.

Fig. 7.

T-type Ca_v inhibitors and Ca²⁺ oscillations in normoxia. A: effects of TTA-P2 (10 μM) and nifedipine (1 μM) on Ca²⁺ oscillations. B: effects of TTA-P2 (20 μM) and nifedipine (1 μM). C: effects of nifedipine (1 μM) and TTA-P2 (20 μM). D: TTA-P2 effects on frequency and amplitude of Ca²⁺ oscillations (means ± SD; 6 coverslips of cells). *Significantly different (P < 0.01). E: nifedipine block of L-type Ca_v activated by FPL64176. F: lack of effect of TTA-P2 on L-type Ca_v activated by FPL64176. Ca_V, voltage-dependent Ca²⁺ channel.

Mild/moderate hypoxia and Ca²⁺ oscillations.

Mild and moderate levels of hypoxia (2–5% O₂) generally produce a very small increase in basal [Ca²⁺]_i when Ca²⁺ signals from multiple cells are averaged (11, 34, 47). We tested the possibility that mild and moderate levels of hypoxia modulate Ca²⁺ signaling by regulating Ca²⁺ oscillations in individual glomus cells. Cells were perfused with normoxic solution at 37°C and then perfused with solution containing 2% or 5% O₂. Both levels of hypoxia increased the frequency of Ca²⁺ oscillations without causing a sustained elevation of basal [Ca²⁺]_i (Fig. 8, A and B). We used 21% O₂ for normoxia, although this level of O₂ is higher than that found inside the CB. In the CB microvasculature, Po₂ measured using different techniques ranges from ∼20 to ∼60 mmHg (4, 13, 30, 48). The frequency of Ca²⁺ oscillations at 10% O₂ was not significantly different from those observed at 21% O₂ (P > 0.20, 4 coverslips, 68 cells; Fig. 8D). In support of this finding, earlier studies have found very little or no effect of 50–60 mmHg Po₂ on averaged [Ca²⁺]_i (11, 43, 46).

Fig. 8.

Hypoxia and Ca²⁺ oscillations. A–C: Ca²⁺ oscillations in response to hypoxia (1–5% O₂). D: plot of frequency as a function of %O₂ (4–6 coverslips for each level of O₂). *Significantly different from that at 21% O₂ (P < 0.01). E–G: inhibition of Ca²⁺ oscillations by 1 μM nifedipine, 20 μM TTA-P2, and 100 μM verapamil under hypoxic conditions. H: frequency effects of nifedipine (5 coverslips; n = 72), TTA-P2 (5 coverslips; n = 66), and verapamil (3 coverslips; n = 48). *Significantly different from control (P < 0.01).

In cells that were quiescent in normoxia, hypoxia (2 or 5% O₂) elicited Ca²⁺ oscillations, and the number of cells responsive to hypoxia on a coverslip increased progressively as the severity of hypoxia was increased from 5% to 1% O₂. At 1% O₂, some cells responded with an increase in Ca²⁺ oscillations, whereas others showed sustained elevation of basal [Ca²⁺]_i. These findings show a wide range in glomus cell sensitivity to hypoxia. When only those cells that showed Ca²⁺ oscillations were analyzed and plotted, it was clear that mild and moderate levels of hypoxia produced a significant increase the frequency of Ca²⁺ oscillations (Fig. 8D). Such a phenomenon is difficult to detect when [Ca²⁺]_i signals are averaged from multiple cells, as done in earlier studies.

Nifedipine markedly reduced the frequency of Ca²⁺ oscillations augmented by hypoxia (2%O₂; Fig. 8, E and H), indicating that hypoxia enhanced Ca²⁺ influx via the L-type Ca_v. However, TTA-P2 (20 μM) produced only a transient inhibition and failed to produce a significant effect overall, suggesting that hypoxia perhaps activated the T-type Ca_v only transiently at the beginning, which is not enough to keep the inhibition sustained (Fig. 8, F and H). ML-218 (20 μM) also failed to produce a significant inhibition of Ca²⁺ oscillations under hypoxia (2%O₂) in three coverslips of cells. Verapamil, an inhibitor of all isoforms of Ca_v, abolished Ca²⁺ oscillations under hypoxia (2%O₂), showing that Ca²⁺ influx is critical for hypoxia (2%O₂)-induced increase in Ca²⁺ oscillations (Fig. 8G). These findings indicate that the L-type Ca_v plays a major role in hypoxia (2% O₂)-induced increase in Ca²⁺ oscillations. Under hypoxic conditions (2%O₂), the frequency of Ca²⁺ oscillations was also strongly reduced by 2-APB (100 μM, 92 ± 6% inhibition, 4 coverslips) and ML-9 (50 μM, 76 ± 9% inhibition, 2 coverslips; n = 48 cells), showing that SOCE/ER play a critical role in the generation of Ca²⁺ oscillations in hypoxia.

DISCUSSION

Slow and random fluctuations in mean [Ca²⁺]_i have been observed in glomus cells previously, but it is not known whether they represent true Ca²⁺ oscillations. Our study shows for the first time that a large percentage of isolated glomus cells are able to generate spontaneous, relatively uniform Ca²⁺ oscillations at a relatively low frequency in normoxia. Our data show that these Ca²⁺ oscillations are produced by Ca²⁺ release/uptake processes at the ER membrane and are sustained by Ca²⁺ influx via the Ca²⁺ channels at the plasma membrane. Thus, the mechanisms of Ca²⁺ oscillations in glomus cells are similar to those found in other secretory and contracting cells such as pancreatic β-cells, adrenal cells, and airway smooth muscle cells (7, 15, 17, 18, 24). We also show that mild and moderate levels of hypoxia increase the frequency and amplitude of Ca²⁺ oscillations, as opposed to a sustained elevation of [Ca²⁺]_i produced by severe levels of hypoxia. Thus, Ca²⁺ oscillations represent the basic Ca²⁺ signaling mechanism in glomus cells. Thus, mild and moderate levels of hypoxia augment glomus cell activity most likely by altering the kinetics of Ca²⁺ release/uptake by the ER to regulate Ca²⁺ oscillations.

Because the elevation of [Ca²⁺]_i is critical for the secretory activity of glomus cells (34, 36, 44), one could speculate that Ca²⁺ oscillations in normoxia are probably important for basal transmitter secretion that occurs in normoxia and that the increased frequency of Ca²⁺ oscillations produced by mild levels of hypoxia augments the secretory response rapidly without desensitization. Although we have not directly determined how changes in frequency and amplitude of Ca²⁺ oscillations affect the secretory activity of glomus cells, we speculate that a strong relationship exists between Ca²⁺ oscillations and transmitter secretion as in other cell types that exhibit Ca²⁺ oscillations.

General property of spontaneous Ca²⁺ oscillations in isolated glomus cells in normoxia.

Under our experimental conditions in which the cells are perfused with normoxic solution at a constant flow rate (∼2 ml/min) and temperature (37°C), spontaneous Ca²⁺ oscillations occurred in ∼65% of glomus cells, with varying frequency among individual cells (0–3 Hz). The frequency of Ca²⁺ oscillations was highly dependent on perfusion temperature, with almost no oscillations present at 24° and prominent oscillations at 37°C. This heat dependence of oscillation frequency is not surprising and has been observed in other types of cells (23, 25).

The frequency of Ca²⁺ oscillations was also dependent on flow rate within the perfusion chamber. Interestingly, no Ca²⁺ oscillations were observed at zero-flow rate even at 37°C, suggesting that mechanical perturbation could be affecting Ca²⁺ signaling mechanisms in glomus cells. Because glomus cells are not in direct contact with the flowing blood in the CB in vivo, we suspect that a pressure-sensing mechanism such as that present in baroreceptors could exist in glomus cells. Abudara and Eyzaguirre (3) first described a mechanical sensitivity of glomus cells. They reported that stirring the fluid bathing cultured rat glomus cells elicited Ca²⁺ spikes that were abolished in Ca²⁺-free external solution. Mechanical stimulation by touching the cells with the tip of the pipette also elicited Ca²⁺ spikes (3). These earlier findings and our recording of flow-sensitive Ca²⁺ oscillations support the notion that glomus cell activity may be regulated by pressure. The ability of glomus cells to sense pressure in an in vivo system will need further investigation but offers an interesting possibility of blood pressure regulation of CB activity. In this regard, hypertension-associated CB hyperactivity reported in a number of studies could involve pressure-induced changes in Ca²⁺ oscillations that in turn control glomus cell activity (1, 32, 37).

The frequency and amplitude of spontaneous Ca²⁺ oscillations observed in individual glomus cells in normoxia were generally stable during ∼30 min of perfusion. Ca²⁺ oscillations were observed in both single isolated cells as well as in cells within clusters. Although cells in each cluster seemed to be attached to each other, Ca²⁺ oscillations frequently occurred independently such that the frequency of Ca²⁺ oscillations in one cell was different from that of another within a cluster. The wide ranges of frequency and amplitude of Ca²⁺ oscillations in isolated glomus cells indicate that the basal excitability and Ca²⁺ signaling kinetics also vary widely among glomus cells. However, this could be an artifact caused by cell isolation, and Ca²⁺ oscillations in vivo could occur in a highly orchestrated process. It seems plausible that glomus cells within a lobule of the carotid body in vivo are connected by gap junctions and, therefore, fire at the same time. Future studies using ex vivo CB preparations should help identify the presence of Ca²⁺ oscillations as well as their kinetics in different regions of intact CB.

Role of Ca²⁺ channels in Ca²⁺ oscillations.

The finding that spontaneous Ca²⁺ oscillations do not occur in Ca²⁺-free external solution indicates that Ca²⁺ influx is necessary to keep the Ca²⁺ signaling machinery that generates Ca²⁺ oscillations working. Glomus cells express both T- and L-type Ca_v (11, 31, 38). Strong inhibition of basal Ca²⁺ oscillations in normoxia by inhibitors of L- (nifedipine, verapamil) and T-type Ca_v (TTA-P2, ML-218) shows that Ca²⁺ influx via both types of Ca_v occurs in normoxia and contributes critically to the Ca²⁺ oscillatory mechanism. Strong inhibition of hypoxia-induced increase in Ca²⁺ oscillations by nifedipine and a weak inhibition by TTA-P2 and ML-218 indicate that Ca²⁺ influx via the L-type Ca_v is the predominant pathway by which mild and moderate levels of hypoxia increase Ca²⁺ oscillations. Thus, Ca²⁺ influx via L- and T-type Ca_v can reduce the frequency of Ca²⁺ oscillations in normoxia, but the L-type Ca_v inhibitor has a greater effect under hypoxic conditions.

It is known that the T-type Ca_v is activated by mild depolarization, involved in pacemaker role in neurons and cardiac cells, and critical for generating low-level spikes and oscillatory firing in many neurons (12, 19, 39). T-type Ca_v can also open near resting E_m and regulate basal Ca²⁺ signaling (39, 49). Thus, the relative overall contributions of L- and T-type Ca_v to the generation of Ca²⁺ oscillations in glomus cells probably depend on the E_m of the cell found under normoxic and hypoxic conditions.

Mechanism of Ca²⁺ oscillations in glomus cells.

Based on our findings, we propose that basal Ca²⁺ influx via both L- and T-type Ca_v maintains a physiological level of cytosolic “Ca²⁺ load” that allows Ca²⁺ release and uptake by ER to occur at a low frequency in normoxia. SOCE exists in glomus cells (45), and thus its activity is expected to correlate with Ca²⁺ oscillations to help sustain the increase in Ca²⁺ oscillation frequency produced by hypoxia. When glomus cells depolarize in response to mild and moderate levels of hypoxia, the resulting increase in Ca²⁺ influx probably elevates the cytosolic “Ca²⁺ load,” thereby shifting the ER release/uptake process to a higher frequency state. Therefore, glomus cells possess Ca²⁺ signaling mechanisms that are similar to those described in other cell types. There is some evidence indicating that glomus cells can spontaneously depolarize and hyperpolarize (8). Such events would cause Ca_v to open and close in an oscillatory manner to help produce and sustain Ca²⁺ oscillations. Therefore, we suspect that oscillations in cell E_m and [Ca²⁺]_i are functionally linked, but this needs to be confirmed in the future.

Limitations of our study in isolated cells.

Although Ca²⁺ oscillations occur in glomus cells in isolated cells, the presence of such Ca²⁺ oscillations in vivo is not known. Although unlikely, enzymatic or mechanical dissociation could have caused Ca²⁺ oscillations following isolation. We were able to observe Ca²⁺ oscillations in a few CB slices, and therefore, we suspect that glomus cells in vivo can also generate Ca²⁺ oscillations, possibly with frequency and amplitude that are different from those in isolated cells. Electrical coupling of pairs of rat glomus cells in culture for 3–8 days has previously been demonstrated (2). In our studies, we used cells within 6 h after isolation, and electrical coupling may not be fully established between cells. In intact CB, Ca²⁺ oscillations may propagate from one cell to the next due to the presence of gap junctions, resulting in identical oscillation frequency within a lobule of the carotid body. The sensitivity of isolated glomus cells to hypoxia may be different from those in vivo due to the presence of various circulatory transmitters and hormones that regulate the excitability of the carotid body. At present, we also do not know the effect of changes in frequency and amplitude of Ca²⁺ oscillations on the secretory activity of glomus cells as well as on the firing of carotid sinus afferent nerve fibers. Future studies are clearly necessary to address these questions by identifying and studying Ca²⁺ oscillations and nerve fiber activity in perfused and pressurized ex vivo preparations, which should help better understand the biological relevance and importance of Ca²⁺ oscillations in CB function.

GRANTS

This work was funded by National Heart, Lung, and Blood Institute Grants HL-111497 (to D. Kim) and HL-142906 (to C. White) and by a grant from Rosalind Franklin University of Medicine and Science.

DISCLOSURES

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

AUTHOR CONTRIBUTIONS

D.K. and C.W. conceived and designed research; D.K., J.O.H., and C.W. performed experiments; D.K., J.O.H., and C.W. analyzed data; D.K., J.O.H., and C.W. interpreted results of experiments; D.K. and J.O.H. prepared figures; D.K. drafted manuscript; D.K. and C.W. edited and revised manuscript; D.K., J.O.H., and C.W. approved final version of manuscript.