During hypoxia, the level of adenosine in the carotid bodies increases as a result of ATP catabolism and adenosine efflux via adenosine transporters. Using Ca2+ imaging, we found that adenosine, acting via A2A receptors, triggered a rise in cytoplasmic [Ca2+] ([Ca2+]i) in type I (glomus) cells of rat carotid bodies. The adenosine response could be mimicked by forskolin (but not its inactive analog), and could be abolished by the PKA inhibitor H89. Simultaneous measurements of membrane potential (perforated patch recording) and [Ca2+]i showed that the adenosine-mediated [Ca2+]i rise was accompanied by depolarization. Ni2+, a voltage-gated Ca2+ channel (VGCC) blocker, abolished the adenosine-mediated [Ca2+]i rise. Although adenosine was reported to inhibit a 4-aminopyridine (4-AP)-sensitive K+ current, 4-AP failed to trigger any [Ca2+]i rise, or to attenuate the adenosine response. In contrast, anandamide, an inhibitor of the TWIK-related acid-sensitive K+-1 (TASK-1) channels, triggered depolarization and [Ca2+]i rise. The adenosine response was attenuated by anandamide but not by tetraethylammonium. Our results suggest that adenosine, acting via the adenylate cyclase and PKA pathways, inhibits the TASK-1 K+ channels. This leads to depolarization and activation of Ca2+ entry via VGCC. This excitatory action of adenosine on type I cells may contribute to the chemosensitivity of the carotid body during hypoxia.
- O2 sensing
- A2A receptor
- protein kinase A
- TWIK-related acid-sensitive K+ channel
carotid bodies are peripheral chemoreceptors that detect lowering of arterial blood O2 level. During hypoxia, stimulation of chemoreceptors activates the carotid sinus afferent nerve terminals and leads to a reflex increase in respiratory rate (14, 32). It is generally accepted that the type I cell in carotid body is the primary oxygen sensor. Hypoxia inhibits O2-sensitive K+ channels [the large-conductance Ca2+-activated K+ (BK) channels and the TWIK-related acid-sensitive K+ (TASK)-like K+ channels] in carotid type I cells, resulting in depolarization and Ca2+ entry (32) via voltage-gated Ca2+ channels (VGCC) [L- and N-type (11, 39)]. The rise in cytoplasmic [Ca2+] ([Ca2+]i) triggers the release of multiple neurotransmitters, including dopamine, ACh, and ATP, from type I cells. There is increasing evidence that ATP has important paracrine and autocrine actions in carotid body. ATP has been shown to stimulate chemosensory discharge on the carotid sinus nerve terminals via P2X receptors (33, 48). On the other hand, our recent study (46) has shown that ATP, acting via P2Y1 receptors, inhibits the hypoxia-mediated [Ca2+]i rise in type I cells. We have also shown that ATP, acting via P2Y2 receptors, triggered Ca2+ release from the intracellular stores of the glial-like type II cells (45). After its release from type I cells, ATP is metabolized to adenosine in the extracellular space. Although hypoxia is known to elevate the extracellular level of adenosine in most tissues (9), a recent study (7) suggests that carotid bodies may be particularly sensitive to the hypoxia-mediated adenosine release. A moderate hypoxia challenge to rat carotid body increased adenosine release by ∼40% but failed to evoke any adenosine release in superior cervical ganglia and carotid arteries (7). About 40% of the increase in adenosine level in carotid body during hypoxia arises from the catabolism of ATP released from type I cells (7). In addition, hypoxia also stimulates adenosine efflux from carotid cells and arteries via the adenosine equilibrative transporters (7). Thus, during hypoxia, the local concentration of adenosine near type I cells is indeed elevated.
It has been well documented that exogenous administration of adenosine to carotid body increases carotid sinus nerve discharge in cats (25, 26, 37) and rats (28, 41). This action has been suggested to involve a presynaptic excitatory action of adenosine on the A2A receptors of type I cells (27, 28, 41). Consistent with this, A2A receptors have been detected on type I cells of rat carotid body (13, 18), whereas A1 receptors are found to be expressed predominantly in the postsynaptic sites of the petrosal ganglia (13). However, inhibitory instead of excitatory action of adenosine on type I cells has been reported. Adenosine has been shown to reduce VGCC in the rat (via A2A receptors; Ref. 18) and rabbit type 1 cells (via A1 receptors; Ref. 36). Adenosine has also been shown to inhibit a 4-aminopyridine (4-AP)-sensitive K+ current in rat type I cells, but the membrane potential of type I cells was not affected by adenosine (41). This issue is further complicated by the observation that in cat carotid body, adenosine increased the release of ACh but decreased the hypoxia-mediated release of dopamine (12). Thus the mechanism underlying the excitatory action of adenosine on the carotid body remains elusive. In the current study, we found that adenosine, acting via the A2A receptors coupled to adenylate cyclase and PKA pathway, reduced the background TASK-like K+ current to trigger depolarization and [Ca2+]i rise in rat carotid type I cells. This action of adenosine on type I cells may underlie the excitatory effect of adenosine in carotid sinus nerve discharge.
The carotid bifurcation was removed from male Sprague-Dawley rats (age 6–7 wk) that had been euthanized with an overdose of halothane. The animals had been handled in accordance with the standards of the Canadian Council on Animal Care. Details of the cell dissociation procedure had been described in our previous studies (45, 46). Dissociated cells were plated onto glass coverslips and maintained in a medium containing F-12/Dulbecco's modified Eagle's medium (1:1), supplemented with 5% fetal calf serum, 50 U/ml penicillin G, and 50 μg/ml streptomycin (all from GIBCO). Cells were cultured at 37°C in an incubator circulated with 95% room air and 5% CO2 for 2–6 h before recordings.
Chemicals and solutions.
Fura-2 AM and indo-1 AM were obtained from Teflabs (Austin, TX). ZM-241385 is from Tocris (Ellisville, MO) and all other chemicals were obtained from Sigma-Aldrich (Oakville, ON, Canada). The standard bath solution contained (in mM) 117 NaCl, 4.5 KCl, 23 NaHCO3, 5 sucrose, 5 glucose, 2.5 CaCl2, and 1 MgCl2 (pH 7.4 when bubbled with 5% CO2). The pipette solution contained (in mM) 140 K+-gluconate, 10 K+-HEPES, 5 MgCl2, and 1 EGTA (pH 7.2). Amphotericin B (250 μg/ml) was included in the pipette solution for perforated patch recording. The cells were perfused with bath solution bubbled continuously with 5% CO2-95% air. Hypoxia was induced by perfusing the cells with bath solution bubbled with 5% CO2-95% N2. Under the hypoxic condition, the Po2 of the bath solution was ∼40 mmHg, as measured with a blood gas analyzer (RapidLab 348; Chiron Diagnostics, Toronto, ON, Canada). Hypercapnia was induced by perfusing the cells with bath solution bubbled with 20% CO2-20% O2-60% N2.
In all experiments involving electrophysiology, single cells were patch clamped with the perforated patch-clamp technique. Membrane potential was recorded using an EPC-7 patch-clamp amplifier that was controlled by a personal computer and the data-acquisition program pCLAMP version 6 (Axon Instruments, Foster City, CA). The pipettes were made from hematocrit glass (VWR Scientific Canada, London, ON, Canada) and the resistance was 20–30 MΩ during perforated patch recording. No correction for junction potential was applied in any of the experiments described here. All recordings were performed at room temperature (20–23°C). Values given in the text are means ± SD.
All [Ca2+]i measurements were performed at room temperature (20–23°C). In experiments involving only measurement of Ca2+ signal, [Ca2+]i was monitored with digital imaging using a Tillvision imaging system equipped with Polychrome II high-speed monochromator (Applied Scientific Instrument), as described previously (45, 46). Cells were loaded with fura-2 AM (2.5 μM) in standard bath solution at 37°C for 10 min and then washed with standard bath solution at 20–23°C for 15 min before being recorded. In experiments involving simultaneous measurement of [Ca2+]i and membrane potential, the electrophysiology rig was equipped with indo-1 fluorescence measurements. Therefore, cells were incubated with indo-1 AM (2.5 μM) instead of fura-2. The incubation procedure was similar to that described above for fura-2 AM. Details of the instrumentation and procedures of [Ca2+]i measurement with indo-1 were as described previously (20, 40). Because the cells were loaded with AM dyes, there was no correction for cell autofluorescence in this study. [Ca2+]i was calculated from the ratio of fluorescence (340 nm/380 nm for fura-2, 400 nm/500 nm for indo-1), using the following equation (15): where Rmin is the fluorescence ratio of Ca2+-free indicator and Rmax is the ratio of Ca2+-bound indicator. K* is a constant that was determined empirically. Calibrations for indo-1 or fura-2 measurements were determined from single cells dialyzed (via the whole cell pipette) with one of the three pipette solutions, as described previously (40). Rmin was measured in cells loaded with the following (in mM): 52 K+-aspartate, 10 KCl, 50 K+-EGTA, 50 K+-HEPES, and 0.1 indo-1 (or fura-2), pH 7.4; and Rmax was measured in cells loaded with (in mM) 136 K+-aspartate, 15 CaCl2, 50 K+-HEPES and 0.1 indo-1 (or fura-2), pH 7.4. K* was calculated from the equation above using the ratio values obtained from cells loaded with the following (in mM): 60 K+-aspartate, 50 K+-HEPES, 20 K-EGTA, 15 CaCl2, 0.1 indo-1 (or fura-2), pH 7.4, which had a calculated free [Ca2+] of 212 nM at 22°C (1).
Adenosine triggered [Ca2+]i elevation in type I cells via activation of A2A receptors.
Using the Ca2+ imaging technique, we examined the action of adenosine on [Ca2+]i of type I cells isolated from adult rat carotid bodies. The acutely dissociated cell preparation of rat carotid bodies was composed of type I, type II, and occasionally white blood cells. All three cell types were ovoid in shape. However, only type I cells contain catecholamines (45) and have Ca2+ response to hypoxia or hypercapnia challenge (3, 4, 46). Therefore, in the current study, we identified individual type I cells based on their Ca2+ response to hypoxia or hypercapnia challenge. To induce moderate hypoxia, the bath solution was bubbled with 5% CO2-95% N2 (Po2 of the bath solution was ∼40 mmHg). As shown in our previous study (46), such a hypoxia challenge could repetitively elicit [Ca2+]i rise in type I cells. Figure 1A shows that the application of adenosine (100 μM) to a hypoxia-sensitive type I cell also triggered a rise in [Ca2+]i. Note that in the example shown in Fig. 1A, the peak amplitude of the adenosine-mediated [Ca2+]i rise was slightly smaller than the hypoxia challenge but the duration of the Ca2+ signal was longer than those triggered by hypoxia. As shown later (Fig. 3), the longer duration of the adenosine-mediated Ca2+ signal was due to the activation of the adenylate cyclase pathway. The adenosine-mediated [Ca2+]i rise could be observed in 23 of the 29 cells that also exhibited a [Ca2+]i rise when challenged with hypoxia. In these 23 cells, the mean amplitude of the peak [Ca2+]i rise triggered by adenosine (100 μM) was 156 ± 91 nM, similar to the peak amplitude of Ca2+ signal triggered by moderate hypoxia in the same cells (184 ± 72 nM; n = 23). As described in discussion, the physiological concentration of adenosine in carotid body is at the micromolar range. Therefore, we examined whether the Ca2+ response could be triggered by lower concentrations of adenosine. Figure 1B shows that adenosine at 1 μM, but not 10 nM, could elicit [Ca2+]i rise. In 15 cells that responded to 100 μM adenosine, only 3 cells responded to 10 nM adenosine, but adenosine at 1 μM could elicit [Ca2+]i rise in 14 cells. In these 14 cells, the mean peak amplitude of the [Ca2+]i rise triggered by 1 μM adenosine was 130 ± 75 nM, similar to that triggered by 100 μM adenosine in the same batch of cells (134 ± 58 nM; n = 15). Thus adenosine at 1 μM was sufficient to trigger the maximum response in type I cells. We also examined whether the effect of adenosine and hypoxia on [Ca2+]i in type I cell was additive by comparing the Ca2+ response mediated by hypoxia alone and that evoked by a combination of adenosine and hypoxia in the same cell. In 9 of the 11 cells examined, the peak amplitude of the Ca2+ signal evoked by the combination of hypoxia and adenosine was slightly smaller than that triggered by hypoxia alone (mean decrease = 35 ± 6 nM) but larger than that triggered by adenosine alone (mean increase = 28 ± 9 nM).
Because A2A receptors were reported to be present on type I cells of rat carotid body (13, 18), we examined the involvement of the A2A receptors in the adenosine-mediated Ca2+ signal with ZM-241385, a selective antagonist of A2A receptors (31). Figure 2A shows that applications of adenosine (100 μM) were able to trigger repetitively [Ca2+]i rises in control condition but failed to evoke any Ca2+ signal in the presence of ZM-241385 (10 μM). After the removal of ZM-241385, adenosine was able to trigger a robust Ca2+ signal in the same cell. In 34 of 38 cells examined, the adenosine (100 μM)-mediated Ca2+ signal was either reduced or abolished by ZM-241385 (10 μM). In these 34 cells, the peak amplitude of the [Ca2+]i rise elicited by adenosine alone was 153 ± 76 nM but reduced to 45 ± 35 nM in the presence of ZM-241385. Consistent with the involvement of A2A receptors, Fig. 2B shows that the adenosine response could be mimicked by the selective A2A receptor agonist, CGS-21680, but not by the A1 receptor agonist 2-chloro-N6-cyclopentyladenosine (CCPA) (17). In 13 cells examined, CGS-21680 (10 nM) evoked a [Ca2+]i rise in every cell (mean increase = 162 ± 50 nM) but CCPA (10 nM) failed to evoke any response. A2A receptors were reported to be coupled to the adenylate cyclase pathway (34) and A2A receptor activation has been shown to increase the cAMP content in rabbit (5) and rat carotid body (29). Therefore, we examined whether the activation of the adenylate cyclase by forskolin could mimic the adenosine response in type I cells. Figure 3A shows that application of forskolin (10 μM) triggered [Ca2+]i rise, which was similar to that triggered by adenosine (100 μM) in the same cell. In 15 cells, which exhibited adenosine response, forskolin triggered [Ca2+]i elevation in 14 cells and the mean peak [Ca2+]i rise triggered by forskolin (178 ± 86 nM) was similar to that triggered by adenosine in the same cells (179 ± 79 nM). In contrast, the inactive analog of forskolin, 1,9-dideoxyforskolin (10 μM), failed to elicit any [Ca2+]i rise in the adenosine-responsive cells (n = 17; Fig. 3B). Consistent with the involvement of the adenylate cyclase/PKA pathway, Fig. 3C shows that the adenosine response was abolished by H89 (10 μM), a selective inhibitor of PKA. A similarly robust inhibition by H89 was observed in 15 of 17 cells examined.
Adenosine response was mediated via membrane depolarization and inhibition of TASK-like K+ channels.
To examine whether the adenosine-mediated [Ca2+]i rise involves changes in membrane excitability, we simultaneously monitored [Ca2+]i and membrane potential (perforated-patch recording) in type I cells. Figure 4A shows an example of such an experiment. In the control condition, the resting membrane potential of this cell was around −38 mV. After the application of adenosine, the membrane potential depolarized to a plateau of around −20 mV, and there was some firing of action potentials. Note that the membrane depolarization was accompanied by [Ca2+]i rise (Fig. 4A). In the six cells examined, the resting membrane potential before adenosine application was −40 ± 3 mV (range −36 to −44 mV at 20–23°C). These values of resting membrane potential are comparable to those reported previously in adult rat type I cells [−32 ± 15 mV at room temperature (11)] and neonatal rat type I cells [range −39 to −63 mV at 35–37°C (3)]. In the presence of adenosine, the membrane potential depolarized (to a plateau) by 16 ± 5 mV and the mean increase in peak [Ca2+]i in these cells was 311 ± 213 nM. Application of the A2A receptor agonist CGS-21680 (10 nM) also caused membrane depolarization in type I cells (by 12 ± 9 mV; n = 7). Treatment with the PKA inhibitor H89 (10 μM) did not affect the membrane potential of type I cells (n = 3) but inhibited the CGS-21680-mediated depolarization (n = 4). Consistent with the notion that the A2A receptor-mediated membrane depolarization in turn leads to activation of voltage-gated Ca2+ entry, Fig. 4B shows that the CGS-21680-triggered [Ca2+]i rise could be reversibly inhibited by the VGCC blocker, Ni2+ (3 mM; n = 6). Note that we did not employ Cd2+, another common VGCC blocker, in this experiment because Cd2+ but not Ni2+ was reported to enter cells and cause quenching of the fluorescence of the Ca2+ indicator (38). In separate experiments, we also confirmed that Ni2+ did not affect the membrane potential of type I cells (n = 5).
Because a previous study (41) has shown that adenosine inhibited a 4-AP-sensitive K+ current in rat type I cells, we examined whether the inhibition of 4-AP-sensitive K+ current was involved in the adenosine-mediated [Ca2+]i rise. Figure 5A shows that the application of 4-AP (5 mM) to the type I cell did not trigger any [Ca2+]i rise, and adenosine (100 μM) was still able to trigger a rise in [Ca2+]i in the continued presence of 4-AP. In 31 cells examined, 4-AP (5 mM) could not trigger a [Ca2+]i rise in any of the cells but adenosine (100 μM) or CGS-21680 (20 nM) was still able to trigger a [Ca2+]i rise in the presence of 4-AP. In these cells, the mean [Ca2+]i rise evoked by adenosine or CGS-21680 in the presence of 4-AP was 187 ± 128 nM. Consistent with the previous studies (2, 41), we also found that application of 4-AP (5 mM) did not affect the membrane potential of type I cells (n = 5). These results suggest that the 4-AP-sensitive K+ current did not contribute significantly to the adenosine-mediated Ca2+ signal.
Hypoxia has been shown to inhibit the activities of BK channels in type I cells (35, 43, 44). To investigate the involvement of BK channels in the adenosine response, we examined whether the adenosine response could be affected by tetraethylammonium (TEA), which is a blocker of several voltage-dependent K+ channels, including the BK channels (30). In neonatal rat type I cells, TEA (10 mM) was reported to have no effect on the membrane potential or [Ca2+]i (4). In 4 of 7 adult rat type I cells examined, we found that TEA (10 mM) caused membrane depolarization (11 ± 5 mV; n = 4). When the Ca2+ response of the adult rat type I cells was examined (n = 35), TEA (10 mM) triggered [Ca2+]i elevations (e.g., Fig. 5B) in ∼60% of the cells. In ∼75% of the cells that responded to TEA, application of adenosine in the continued presence of TEA evoked a larger and more sustained [Ca2+]i rise (Fig. 5B). The time integral (for 200 s) of the Ca2+ signal (see Fig. 5B, for example) triggered in the presence of a combination of TEA and adenosine (23 ± 11 μM s; n = 14) was almost twofold larger than that triggered by TEA alone (12 ± 4 μM s; n = 14). For type I cells, which did not exhibit any rise in [Ca2+]i when challenged with TEA, application of adenosine could still trigger [Ca2+]i elevations (Fig. 5C; n = 15). Overall, this result suggests that the adenosine response did not involve any major contribution from TEA-sensitive K+ currents.
In rat type I cells, a TEA-insensitive background TASK-like K+ current has been shown to be an important mechanism underlying the hypoxia-mediated membrane depolarization and [Ca2+]i rise (2). Immunohistochemical study has revealed the presence of multiple TASK-like channels (including TASK-1, -2, and -3) in rat type I cells (47). Therefore, we examined whether anandamide, a selective TASK-1 K+ channel blocker (23), could affect the adenosine response. Anandamide at 3 μM was reported to inhibit ∼90% of the TASK-1 K+ channels but had no significant effect on the TASK-2 or TASK-3 channels (23). We found that application of anandamide (5 μM) resulted in robust [Ca2+]i rise in type I cells (Fig. 6A). Note that in the continued presence of anandamide, adenosine did not cause any further increase in [Ca2+]i (Fig. 6A; n = 27). Figure 6B shows that the anandamide-mediated [Ca2+]i rise in type I cells was accompanied by membrane depolarization and firing of action potentials. Application of adenosine did not cause any further increase in depolarization (Fig. 6B; n = 5). Thus the action of adenosine was blunted by anandamide. This result is consistent with the notion that a reduction of the TASK-1 background K+ current is the major mechanism underlying the adenosine-mediated Ca2+ signal in type I cells.
The findings here demonstrate that in the majority of rat carotid type I cells (identified by their Ca2+ response to hypoxia or hypercapnia), adenosine triggered a rise in [Ca2+]i. In ∼20% of the hypoxia-responding cells, adenosine failed to elicit any [Ca2+]i rise. Because the adenosine-mediated [Ca2+]i rise was dependent on depolarization and activation of VGCC (Fig. 4), it is possible that the adenosine-mediated depolarization in some type I cells may be insufficient to trigger significant VGCC activation and Ca2+ entry. We also found that adenosine at 1 or 100 μM triggered a similar increase in [Ca2+]i, suggesting that a maximal response could be achieved by 1 μM adenosine. The amount of adenosine released from the carotid body was estimated to be ∼100 pmol per carotid body during normoxia and increased by ∼34% during 10 min of moderate hypoxia (7). Although the concentration of adenosine in the carotid body has not been measured, the level of adenosine in rat hippocampal slice during hypoxia was found to reach ∼5 μM (8). Thus it is probable that the local concentration of adenosine near type I cells during hypoxia may also reach the micromolar range. Adenosine at concentrations 1 to 100 μM has been reported to increase the chemoreceptor discharge in rat carotid body as well as inhibiting a 4-AP-sensitive K+ current in type I cells (41). Interestingly, the same study showed that adenosine did not affect the membrane potential of type I cells (41). In contrast, our current finding shows that the adenosine-mediated [Ca2+]i rise was accompanied by depolarization and firing of action potentials (Fig. 4A). One possible explanation for this discrepancy is that perforated patch-clamp recording was employed in the present study. Because the adenosine response involved inhibition of TASK-like channels (Fig. 6), and these K+ channels were reported to be regulated by cytosolic factors (42), it is possible that these modulations might be lost during whole cell recording (41). Our experiments also show that the 4-AP-sensitive K+ current has no significant contribution to the adenosine-triggered Ca2+ signal as application of 4-AP did not trigger any [Ca2+]i rise in type I cells and the mean peak [Ca2+]i rise evoked by adenosine in the presence of 4-AP was similar to that of the control cells (Fig. 5A).
Consistent with previous studies that show the presence of A2A receptors on type I cells of rat carotid bodies (13, 18), we found that the adenosine-triggered [Ca2+]i rise in type I cells was mediated via A2A receptors because the response could be mimicked by the A2A receptor agonist, CGS-21680 (Fig. 2B), and inhibited by ZM-241385, an A2A receptor antagonist (Fig. 2A). A2A receptors are known to stimulate PKA activity via Gs protein and adenylate cyclase (34). Inhibition of PKA by H89 abolished the adenosine-mediated Ca2+ signal (Fig. 3C). H89 has been reported to have PKA-independent actions, such as inhibition of sarcoplasmic reticulum Ca2+-ATPase and L-type Ca2+ current (16), reduction of Kv1.3 channels (6), and translocation of epithelia Na+ channels (24). Nevertheless, our finding that forskolin, an activator of adenylate cyclase (but not the inactive analog of forskolin), could mimic the adenosine-mediated [Ca2+]i rise in type I cells (Fig. 3, A and B) further implicated the involvement of PKA in the adenosine response. Overall, our results suggest that stimulation of A2A receptor in rat type I cell activates PKA that in turn inhibits the TASK-like K+ channels and leads to depolarization. Consistent with an inhibitory effect of PKA on TASK-like K+ channels, activation of PKA by forskolin and IBMX has been shown to reduce TASK-like K+ current by ∼40% (21, 22). Inhibition of PKA via GABAB receptor activation in rat type I cells has been shown to activate TASK-like K+ channels and cause hyperpolarization (10).
Activation of A2A receptors has been reported to decrease the voltage-gated Ca2+ current in rat type I cells (18) and PC-12 cells (19). Note that our observation that adenosine triggered [Ca2+]i rise in type I cells does not necessarily contradict an inhibitory action of adenosine on voltage-gated Ca2+ current. Application of adenosine typically depolarized the membrane potential of type I cells by ∼15 mV (e.g., from around −38 to −23 mV in Fig. 4A). At −20 mV, the inhibition of voltage-gated Ca2+ current by adenosine was very small (18). Thus adenosine could still trigger a robust [Ca2+]i rise at this potential. Our result also suggests that adenosine does not cause additional increase in the peak amplitude of the hypoxia-induced Ca2+ signal. At least two factors may contribute to this observation. First, the TASK-like channel is a common target for both hypoxia and adenosine. If hypoxia already inhibited most of the TASK-like channels, the effect of adenosine and hypoxia would not be additive. Second, with increasing depolarization during the combined challenge, the inhibitory action of adenosine on voltage-gated Ca2+ current would become more prominent, thus preventing further increase in the amplitude of the Ca2+ signal.
Our result contradicts a previous study by Kobayashi et al. (18), which reported that adenosine did not evoke any [Ca2+]i rise in rat type I cells. Two experimental conditions may contribute to this discrepancy. First, there may be a difference in the sensitivity of type I cells. In the study of Kobayashi et al. (18), cells that were cultured overnight were used and severe hypoxia (evoked by the O2 scavenger, sodium dithionate) was needed to evoke Ca2+ signal in type I cells. We found that there is a drastic decline in the Ca2+ response of rat type I cells to hypoxia after 24 h of culture. Therefore, in our study, only cells cultured for 2–6 h were employed. Under this condition, even a moderate hypoxia (∼40 mmHg) can evoke a [Ca2+]i rise in type I cells. Second, Ca2+ indicators such as fura-2 also act as a Ca2+ buffer. Thus excessive loading of fura-2 will increase the cytosolic Ca2+ buffering capacity and lead to a decrease in the amplitude of the Ca2+ signal. In our study, we loaded the cells with 2.5 μM fura-2 AM at 37°C for 10 min. In contrast, in the study of Kobayashi et al. (18), cells were loaded with 5 μM fura-2 AM at 37°C over a 40-min time period. Because the amplitude of the adenosine-mediated Ca2+ signal is small (∼150 nM), it is possible that a reduction in the sensitivity of type I cells in conjunction with an increase in cytosolic Ca2+ buffer may mask the [Ca2+]i rise triggered by adenosine in the latter study.
It has been suggested that adenosine may contribute to the carotid body chemosensitivity to modest hypoxia (7). In this study, we found that adenosine triggered depolarization and [Ca2+]i rise in type I cells and the Ca2+ response was comparable to that triggered by moderate hypoxia (∼40 mmHg). This raises the possibility that near the threshold level of tissue Po2 for activation of carotid sinus nerve discharge, the rise in adenosine level may play a significant role in the stimulation of type I cells and thus in turn increase the carotid sinus nerve discharge. We (46) have shown previously that ATP (released from type I cells) acts as a negative regulator to inhibit the hypoxia-mediated Ca2+ signal in type I cells by causing membrane hyperpolarization. The catabolism of extracellular ATP into adenosine, in conjunction with the hypoxia-mediated adenosine efflux, may activate the A2A receptors and trigger membrane depolarization, thus opposing the inhibitory action of ATP on type I cells. In view of this, adenosine may be important in helping type I cells to recover from the negative feedback action of ATP.
This work is supported by grants from the Canadian Institute of Health Research and the Alberta Heritage Foundation for Medical Research, where A. Tse and F. W. Tse are senior scholars.
Present address for J. Xu: National Institute of Neurological Disorders and Stroke, 35 Convent Dr., Bethesda, MD 20892.
↵* F. Xu and J. Xu have contributed equally to this work.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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