α-Adrenergic stimulation of cytosolic Ca²⁺ oscillations and exocytosis in identified rat corticotrophs

Abstract

1. The patch clamp technique was used in conjunction with a fluorescent Ca²⁺ indicator (indo-1, or indo-1FF) to measure simultaneously cytosolic Ca²⁺ concentration ([Ca²⁺]_i), ionic current and changes in membrane capacitance in single rat corticotrophs identified with the reverse haemolytic plaque assay.

2. Application of the adrenocorticotropin (ACTH) secretagogue noradrenaline (NA; norepinephrine), triggered [Ca²⁺]_i oscillation in corticotrophs via α-adrenergic receptors and the guanosine trisphosphate (GTP) binding protein-coupled phosphoinositide pathway.

3. Simultaneous measurement of [Ca²⁺]_i and capacitance shows that exocytosis was triggered during the first cycle of NA-induced [Ca²⁺]_i oscillation and the mean increase in cell membrane surface area was 1.4 ± 0.3 % (n = 6).

4. When Ca²⁺ was directly released from the inositol 1,4,5 trisphosphate (IP₃)-sensitive store via flash photolysis of caged IP₃, the mean increase in cell surface area was 1.5 ± 0.5 % (n = 6). Thus, NA-stimulated ACTH secretion in rat corticotrophs is closely coupled to intracellular Ca²⁺ release.

5. Large and rapid elevation of [Ca²⁺]_i (>15 μm) via flash photolysis of caged Ca²⁺ triggered two phases of exocytosis: a rapid exocytic burst that was complete in ∼100 ms and a slow burst that continued for many seconds.

6. The rapid exocytic burst reflected the exhaustion of a pool of readily releasable granules and, on average, increased the cell surface by 2.8 ± 0.1 % (n = 14).

7. We suggest that the relatively weak exocytic response in corticotrophs during intracellular Ca²⁺ release may be partially attributed to a smaller pool of readily releasable granules.

Secretion of the stress hormone adrenocorticotropin (ACTH) from the corticotrophs in the anterior pituitary gland is controlled by multiple hypothalamic hormones and neurotransmitters, including corticotropin-releasing hormone (CRH), arginine vasopressin (AVP), oxytocin (OT) and the catecholamines (reviewed by Antoni, 1986). CRH is the most potent ACTH secretagogue and it triggers a persistent membrane depolarization in rat corticotrophs via a protein kinase A dependent closure of a background K⁺ conductance (Lee & Tse, 1997). The membrane depolarization in turn activates voltage-gated Ca²⁺ entry and causes a sustained [Ca²⁺]_i elevation (Ritchie et al. 1996; Lee & Tse, 1997). In contrast, AVP activates the phospholipase C pathway in rat corticotrophs and results in a ‘transient and plateau’ pattern of [Ca²⁺]_i elevation in rat corticotrophs (Corcuff et al. 1993; Tse & Lee, 1998). The mechanism underlying the catecholamine-triggered ACTH secretion, however, is less well studied. In rat pituitary cells, both α- and β-adrenergic receptors have been implicated in the catecholamine-stimulated ACTH secretion (Giguere et al. 1981; Perkins et al. 1985). In the ACTH secreting tumour cell line AtT-20, catecholamine stimulated ACTH secretion via β₂-adrenergic receptors that were coupled to the adenylate cyclase pathway (Axelrod & Reisine, 1984). The difficulty in isolating and identifying corticotrophs from a heterogeneous population of anterior pituitary cells has been a major hindrance in the study of catecholamine action on corticotrophs.

Here we studied the mechanism underlying the noradrenaline (NA; norepinephrine) stimulated ACTH secretion in single rat corticotrophs which were identified with the reverse haemolytic plaque assay, using antibodies against ACTH. By measuring simultaneously ionic current and [Ca²⁺]_i in the same cell, we show that NA induces [Ca²⁺]_i oscillation which in turn causes periodic opening of apamin-sensitive Ca²⁺-activated K⁺ channels. The action of NA on [Ca²⁺]_i is mediated via the α-adrenergic receptors and a G-protein-coupled phosphoinositide pathway. Simultaneous measurement of [Ca²⁺]_i and exocytosis shows that NA triggers exocytosis in corticotrophs via release of intracellular Ca²⁺ from the IP₃-sensitive stores. The small number of readily releasable granules in the rat corticotroph may underlie the weak exocytic response during intracellular Ca²⁺ release.

METHODS

Cell preparation

The anterior lobe of the pituitary gland was removed from male Sprague-Dawley rats (aged 5–6 weeks) killed with halothane in accordance with the standards of the Canadian Council on Animal Care. Anterior pituitary glands were dissociated enzymatically using collagenase and trypsin as previously described (Tse & Hille, 1994). Single corticotrophs were identified from the heterogeneous population by reverse haemolytic plaque assay (Smith et al. 1984), with procedures similar to those described previously (Lee & Tse, 1997; Tse & Lee, 1998). Briefly, the pituitary cells were suspended in Dulbecco's modified Eagle's medium (DMEM; Gibco) that contained 0.1 % (w/v) bovine serum albumin (BSA; Sigma). The pituitary cell suspension was then mixed with an equal volume of 12 % (v/v) sheep erythrocytes (Colorado Serum Co., Denver, CO, USA) in 0.9 % NaCl. The erythrocytes were previously conjugated with Staphylococcus aureus-derived protein A (Sigma), using 0.2 mg ml⁻¹ CrCl₃ as catalyst. The cell mixture was then plated onto a culture dish such that a monolayer of cells was attached to the bottom of the dish. Detail of the cell attachment procedure has been described previously (Tse & Hille, 1994). The cell layer was then incubated with 100 nm CRH (Peninsula Laboratories, Belmont, CA, USA, or a gift from Dr W. Vale, Salk institute, San Diego, CA, USA) and rabbit polyclonal antibodies to rat ACTH (1:20 dilution; gift from Dr R. J. Kemppainen, Auburn University, Auburn, AL, USA) for 3 h at 37°C. This was followed by a 30 min exposure to guinea-pig complement at 1:50 dilution. Individual corticotrophs were identified by the presence of a zone of lysed erythrocytes (a plaque) surrounding a pituitary cell. The cells were maintained under standard culture conditions in a DMEM supplemented with 10 % (v/v) horse serum, 50 u ml⁻¹ penicillin G and 50 μg ml⁻¹ streptomycin. Plaques remained identifiable for up to 5 days. Recordings were performed on cells cultured for 2–4 days after plaque formation.

Solutions

The standard bath solution contained (mm): 150 NaCl, 10 Na-Hepes, 8 glucose, 2.5 KCl, 2 CaCl₂ and 1 MgCl₂, pH 7.4. During capacitance measurements, apamin (0.4 μm) was included in the standard bath solution to inhibit the slow Ca²⁺-activated K⁺ (SK) current. The standard pipette solution contained (mm): 120 potassium aspartate, 20 KCl, 20 K-Hepes, 1 MgCl₂, 2 Na₂ATP, 0.1 Na₄GTP and 0.1 indo-1, pH 7.4. In experiments where extracellular Ca²⁺ was removed, Ca²⁺ was omitted from the standard bath solution and 2 mm MgCl₂ and 1 mm Na-EGTA were added. In experiments where [Ca²⁺]_i was elevated via flash photolysis of caged Ca²⁺, the pipette solution contained (mm): 70 caesium aspartate, 40 Cs-Hepes, 20 tetraethylammonium chloride, 0.1 Na₄GTP, 0.1 indo-1FF and 6.5–8 DM-nitrophen (∼90 % saturated with Ca²⁺), pH 7.4. The purity of DM-nitrophen varied slightly between batches and, therefore, the loading of each patch of DM-nitrophen with Ca²⁺ was determined empirically.

CRH (Peninsula Laboratories) was dissolved in 0.1 m acetic acid, lyophilized and kept at −20°C. Indo-1 (Calbiochem), indo-1FF (Texas Fluorescence Labs, Inc., Austin, TX, USA) and NA (Sigma) were kept as stock solutions in distilled water at −20°C. Heparin, DM-nitrophen, caged IP₃ and guanosine-5′-O-(2-thiodiphsophate) (GDP-β-S) were obtained from Calbiochem. Apamin was obtained from Sigma. Sodium ascorbate (30 μg ml⁻¹; Sigma) was added to the NA solution to reduce oxidation during the experiment. Except for the flash photolysis experiments, the bath was perfused continuously with control or drug solution by gravity. The time for a complete exchange of bath solution was ∼40–70 s.

Measurement of [Ca²⁺]_i

[Ca²⁺]_i was measured fluorometrically using the Ca²⁺ indicator indo-1 or indo-1FF. The Ca²⁺ indicator was dialysed into the cell via the whole-cell patch pipette. Details of the instrumentation and procedures of the [Ca²⁺]_i measurement were as described previously (Tse & Hille, 1994; Tse et al. 1994a). Briefly, indo-1 or indo-1FF in single corticotrophs was excited by 365 nm (band-pass filtered) light delivered from an HBO 100 W mercury lamp via a × 40, 1.3 NA UV fluor oil objective (Nikon). Photon counts were collected at 405 and 500 nm by two photomultiplier tubes (Hamamatsu H3460-04) and then translated into logic signals counted simultaneously by a CYCTM-10 counter card (Cyber Research Inc., Branford, CT, USA) in an IBM-compatible computer. [Ca²⁺]_i was calculated from the ratio (R) of fluorescence at 405 and 500 nm, using the equation of Grynkiewicz et al. (1985):

(1)

Where R_min is the fluorescence ratio of Ca²⁺-free indicator and R_max is the ratio of Ca²⁺-bound indicator. K* is a constant that was determined empirically. Calibrations were determined from groups of single corticotrophs (n = 5–8) dialysed with one of the three pipette solutions as described previously (Lee & Tse, 1997; Tse et al. 1997). For indo-1 measurement, R_min was measured in cells loaded with (mm): 52 potassium aspartate, 10 KCl, 50 K-EGTA, 0.1 indo-1 and 50 K-Hepes, pH 7.4; and R_max was measured in cells loaded with (mm): 136 potassium aspartate, 15 CaCl₂, 0.1 indo-1 and 50 K-Hepes, pH 7.4. K* was calculated from eqn (1) using R values obtained from cells loaded with (mm): 60 potassium aspartate, 50 K-Hepes, 20 K-EGTA, 15 CaCl₂, 0.1 indo-1, pH 7.4, which had a calculated free Ca²⁺ concentration of 212 nm at 24°C (Blinks et al. 1982). For all indo-1 measurements reported here, the values of R_min, R_max and K* were 0.403, 5.05 and 2.62 μm, respectively. In experiments involving flash photolysis of Ca-DM-nitrophen, [Ca²⁺]_i was measured with the low affinity Ca²⁺ indicator indo-1FF. For these measurements, R_max was measured in cells loaded with (mm): 109 potassium aspartate, 25 CaCl₂, 10 DM-nitrophen, 0.1 indo-1FF and 40 K-Hepes, pH 7.4. K* was calculated from eqn (1) using R values obtained from cells loaded with (mm): 70 diglycolic acid, 15 potassium aspartate, 50 K-Hepes, 10 CaCl₂ and 0.1 indo-1FF, pH 7.4, which had a free Ca²⁺ concentration of 24 μm at 24°C (measured with a Ca²⁺ electrode). For all indo-1FF measurements reported here, the values of R_max and K* were 2.56 and 79.2 μm, respectively. R_min was obtained individually for each cell as the mean of five to ten measurements taken immediately before the flash and the values ranged from 0.21 to 0.25.

Electrophysiological recording

Membrane current was recorded with the whole-cell, gigaseal method (Hamill et al. 1981) using an EPC-9 patch clamp amplifier. Changes in membrane capacitance (ΔC_m) were measured with a dual phase lock-in amplifer by superimposing an 800 Hz sinusoid of 30 mV peak-to-peak amplitude onto the holding potential as previously described (Tse & Hille, 1994). Values of [Ca²⁺]_i, currents and ΔC_m were first recorded on VCR tapes with a Neuro Data PCM recorder (Neuro Data Instruments Corp., New York) and digitized later. The pipettes were made from haematocrit glass (VWR Scientific Canada Ltd, London, Ontario, Canada) and the resistance was 2–4 MΩ after filling and 5–10 MΩ during whole-cell recording. Recordings were done at room temperature (22–25°C). A −10 mV junction potential was corrected throughout.

Values given are means ±s.e.m. Unless indicated otherwise, the time in each figure shows the time after establishment of the whole-cell configuration.

RESULTS

NA triggers [Ca²⁺]_i oscillations via stimulation of α-adrenergic receptors

Figure 1 shows the effect of NA on [Ca²⁺]_i and ionic current in a single identified rat corticotroph. The cell was voltage clamped at −50 mV to prevent activation of voltage gated Ca²⁺ channels. Application of NA (1 μm) triggered a [Ca²⁺]_i oscillation. Each cycle of the [Ca²⁺]_i oscillation was accompanied by a rhythmic outward current. This current was completely abolished by apamin (0.4 μm), a selective inhibitor of the small conductance Ca²⁺-activated K⁺ (SK) channels (n = 5), indicating that the oscillatory [Ca²⁺]_i in turn causes periodic opening of SK channels. Note that the amplitude of the [Ca²⁺]_i elevation varied between cycles of oscillation (Fig. 1). In ten cells examined, peak [Ca²⁺]_i during a cycle of the oscillation ranged from 0.2 to 3 μm. The frequency of the NA-induced [Ca²⁺]_i oscillation also varied between cells even though the same concentration of NA was employed in all our experiments (see Fig. 2 for example). Following the removal of NA, the [Ca²⁺]_i oscillation typically continued for one to three cycles (Figs 1 and 4). The delay between NA removal and the stopping of the [Ca²⁺]_i oscillation is in part due to the time required for a complete exchange of bath solution (∼60 s). In addition, since the stimulation of NA receptors leads to the activation of the phospholipase C (PLC) pathway and the generation of IP₃ (see below), the time required for the shutdown of the PLC pathway and degradation of IP₃ may also contribute to this delay. A similar phenomenon has also been observed in pituitary gonadotrophs where the [Ca²⁺]_i oscillation can continue for many cycles following agonist removal (Tse & Hille, 1992).

Figure 1. NA triggers [Ca²⁺]_i oscillation and rhythmic outward current.

[Ca²⁺]_i and ionic current were measured simultaneously from a single rat corticotroph in whole-cell configuration. The cell was voltage clamped at −50 mV. Application of NA (1 μm) induced [Ca²⁺]_i oscillation which was accompanied by rhythmic outward current. Application of the bee venom apamin (0.4 μm) completely abolished the outward current without affecting the [Ca²⁺]_i oscillations.

Figure 2. The NA-induced [Ca²⁺]_i oscillation is mediated via α-adrenergic receptors.

A, NA could trigger [Ca²⁺]_i oscillation in the presence of the β-adrenergic blocker propranolol but the NA response was abolished by the-α-adrenergic blocker phentolamine. B, [Ca²⁺]_i oscillations could be produced by the α-adrenergic agonist phenylephrine. C, the NA response was abolished by the α-adrenergic blocker, prazosin. In all experiments shown here, the membrane potentials were held at −50 mV.

Figure 4. The NA-induced [Ca²⁺]_i oscillation is independent of extracellular Ca²⁺.

[Ca²⁺]_i fell slightly when the external saline was switched from one containing 2 mm Ca²⁺ to a Ca²⁺-free solution that included 1 mm EGTA. Application of NA could still trigger a robust [Ca²⁺]_i oscillation. The membrane potential was held at −50 mV.

We investigated the receptors involved in the NA response with selective inhibitors of α- and β-adrenergic receptors. Figure 2A shows that application of the β-adrenergic receptor blocker propranolol (10 μm) failed to prevent NA (1 μm) from triggering a [Ca²⁺]_i oscillation. But the NA-induced [Ca²⁺]_i oscillation was completely abolished when the bath solution was subsequently switched to one containing 1 μm NA and 10 μm phentolamine (an α-adrenergic receptor blocker) (n = 4). When the [Ca²⁺]_i oscillation was first elicited by NA (1 μm), and the cell was then exposed to propranolol (10 μm) in the continued presence of NA, the NA-induced Ca²⁺ response persisted (n = 3). Subsequent addition of phentolamine (10 μm) abolished the Ca²⁺ response. Figure 2B shows that the NA-induced [Ca²⁺]_i oscillations could be mimicked by the application 10 μm phenylephrine, an α-adrenergic receptor agonist (n = 3). Consistent with the involvement of α-adrenergic receptors, the NA-induced [Ca²⁺]_i oscillations could be abolished by prazosin, an α-adrenergic receptor antagonist (Fig. 2C; n = 3).

The signalling pathway underlying the NA-induced [Ca²⁺]_i oscillation

Among the α-adrenergic receptors, the α₁ subtype in many cells is coupled to the GTP binding protein G_q, which in turn activates the phosphoinositide pathway. To examine the involvement of G-protein in the NA-induced [Ca²⁺]_i oscillation, we prevented the activation of G-protein in corticotrophs by including 2 mm GDP-β-S (a non-hydrolysable analogue of GDP) in the pipette solution. In five cells recorded with GDP-β-S, NA failed to trigger [Ca²⁺]_i elevation (Fig. 3A). The involvement of the IP₃ receptor was investigated with heparin, a competitive blocker of IP₃ receptors. Corticotrophs were recorded with 100 μm heparin in the pipette solution. The cell was dialysed for 5 min to allow for the equilibration of the heparin with the cytosol. Note that in control cells (e.g. Figs 2A and 4), the NA response was not affected by the time of whole-cell dialysis. Figure 3B shows that application of NA (1 μm) failed to trigger [Ca²⁺]_i elevation in cells loaded with heparin (n = 4). Interestingly, the NA-induced [Ca²⁺]_i oscillation could not be completely mimicked by intracellular IP₃. When we bypassed the NA receptors and directly released Ca²⁺ from the IP₃-sensitive store via flash photolysis of caged IP₃, only a transient Ca²⁺ elevation was observed (Fig. 3C; n = 4). A second flash delivered ∼80 s later, could still trigger robust [Ca²⁺]_i elevation, suggesting that the intracellular Ca²⁺ store was not depleted. These results suggest that NA acts via the G-protein-coupled phosphoinositide pathway to trigger Ca²⁺ release from the IP₃-sensitive store. Consistent with this, NA could trigger [Ca²⁺]_i oscillation in the absence of extracellular Ca²⁺ (Fig. 4; n = 4). Note that the removal of extracellular Ca²⁺ typically resulted in a reduction of the resting [Ca²⁺]_i, presumably reflecting a passive Ca²⁺ influx across the plasma membrane when extracellular Ca²⁺ was present.

Figure 3. The NA-induced [Ca²⁺]_i oscillation is mediated via the G-protein-coupled phosphoinositide pathway.

A, inhibition of the NA response by GDP-β-S. The cell was recorded with 2 mm GDP-β-S in the pipette solution. B, inhibition of the NA-triggered [Ca²⁺]_i oscillation by heparin, a competitive blocker of IP₃ receptors. The cell was recorded with 100 μm heparin in the pipette solution. C, intracellular IP₃ partially mimics the NA response. The cell was recorded with 10 μm caged IP₃ in the pipette solution. Two UV flashes were delivered at the time indicated by the arrows. A transient [Ca²⁺]_i elevation was elicited following each flash. The holding potential was −50 mV in all the experiments shown here.

NA triggers exocytosis via intracellular Ca²⁺ release

To examine whether the NA-induced [Ca²⁺]_i elevation can trigger secretion, we employed the high temporal resolution capacitance measurement (Gillis, 1995) to measure exocytosis in single corticotrophs. The capacitance measurement monitors increases in cell membrane surface area resulting from exocytosis of secretory granules. Figure 5A shows a simultaneous measurement of [Ca²⁺]_i and exocytosis in a corticotroph during NA stimulation. The cell was voltage clamped at −70 mV (DC), a potential at which most voltage-gated channels, including Ca²⁺channels, were closed. A short application of NA triggered two Ca²⁺ waves in this cell. The first [Ca²⁺]_i wave (shown in Fig. 5A) was accompanied by an increase in membrane capacitance, reflecting exocytosis. After the peak of this Ca²⁺ wave, the membrane capacitance began to fall, reflecting net endocytosis. Within ∼20 s, the membrane capacitance returned completely to the basal level. The maximum increase in capacitance in this cell was 50 fF, corresponding to an increase of ∼1.5 % in membrane surface area (initial cell membrane capacitance = 3.34 fF). About 30 s following the first Ca²⁺ wave, a second small (∼0.5 μm) Ca²⁺ wave was elicited (data not shown) but it was not accompanied by any increase in capacitance. A similar pattern of exocytosis and endocytosis was observed in five other cells. In all the cells examined here, no further increase in capacitance could be detected during subsequent cycles of [Ca²⁺]_i oscillation. However, as shown in Fig. 2, the peak of the Ca²⁺ waves tended to be smaller after the first wave. In this batch of cells, the amplitude of the subsequent cycles of NA-induced [Ca²⁺]_i oscillations was small (<0.6 μm). Thus, it is possible that the small Ca²⁺ rise is insufficient to trigger exocytosis. Nevertheless, the mean [Ca²⁺]_i during the first cycle of NA-induced [Ca²⁺]_i oscillation was 1.43 ± 0.20 μm and the accompanying increase in membrane surface area was 1.4 ± 0.3 % (capacitance increase = 57.7 ± 10.7 fF; mean initial cell capacitance = 4.43 ± 0.90 pF; n = 6).

Figure 5. NA-triggered exocytosis is closely coupled to intracellular Ca²⁺ release.

A, the NA-induced [Ca²⁺]_i elevation was accompanied by exocytosis. Time course of changes in [Ca²⁺]_i and membrane capacitance (ΔC_m) during NA challenge. In this cell, NA application triggered two Ca²⁺ waves. Only the first Ca²⁺ wave is shown here. B, exocytosis triggered by Ca²⁺ release from IP₃-sensitive stores. The pipette solution contained 10 μm caged IP₃. Two UV flashes were delivered at the times indicated by the arrowheads. During the first [Ca²⁺]_i elevation, exocytosis (increase in ΔC_m) was elicited and it was followed by robust endocytosis (decrease in ΔC_m). The rate of endocytosis appeared to increase following the second Ca²⁺ elevation. In all the experiments described here, the cell was voltage clamped at −70 mV (DC) to shut off most voltage-gated channels, including the Ca²⁺ channels. The Ca²⁺-activated K⁺ channel was blocked by 0.4 μm apamin.

During α-adrenergic receptor stimulation, the activation of phospholipase C generates both IP₃ and diacylglycerol. Thus, protein kinase C (PKC) may be activated by diacylglycerol during NA stimulation. Although it is not clear whether PKC has a role in the α-adrenergic stimulation of ACTH secretion, PKC had been postulated to be essential for the ACTH secretion evoked by AVP, which also utilizes the phospholipase C pathway (Bilezikjian et al. 1987; Koch & Lutz-Bucher, 1991). To test whether NA can trigger exocytosis without PKC activation, we bypassed the receptor-coupled pathway and directly released Ca²⁺ from the IP₃-sensitive store via flash photolysis of caged IP₃. A single corticotroph was recorded with 10 μm caged IP₃ in the pipette solution. Figure 5B shows that shortly after the delivery of an ultraviolet (UV) flash, [Ca²⁺]_i rose rapidly to ∼2 μm and was accompanied by a clear increase in membrane capacitance, suggesting that the NA-triggered exocytosis in pituitary corticotrophs is closely coupled to intracellular Ca²⁺ release from the IP₃-sensitive store. The increase in capacitance reached a maximum at 90 fF within 2 s and it was followed by endocytosis. In this cell, the retrieval of membrane continued to occur even after the capacitance was restored to the basal value. To examine whether additional exocytosis can be triggered during subsequent [Ca²⁺]_i elevation, a second UV flash was delivered at ∼65 s following the first UV flash to photolyse more caged IP₃. Although [Ca²⁺]_i rose to ∼1 μm following the second UV flash, only a further reduction in membrane capacitance was triggered. Similar experiments were performed in five other cells. In all cells examined here, exocytosis could be detected only during the first UV flash. The mean [Ca²⁺]_i elevation following the first UV flash was 1.88 ± 0.14 μm and it was accompanied by an addition of 1.5 ± 0.5 % to the membrane surface area (capacitance increase = 77.5 ± 27.4 fF; mean cell initial capacitance = 5.73 ± 0.78 pF; n = 6). Although the mean [Ca²⁺]_i elevation following the second UV flash was 1.47 ± 0.21 μm, similar to that triggered during the first cycle of NA-induced [Ca²⁺]_i oscillation, no further increase in capacitance was detected.

Size of the readily releasable pool of granules

The results above show that the first Ca²⁺ wave from the release of intracellular Ca²⁺ typically increased the cell surface area of single corticotrophs by 1.5 %. Interestingly, the amplitude of this exocytic response (normalized for cell surface area) in corticotrophs is ∼3-fold smaller than that elicited by the first cycle of [Ca²⁺]_i oscillation in pituitary gonadotrophs (Tse et al. 1993; Tse et al. 1997). In many endocrine cells, including gonadotrophs (Tse et al. 1997), melanotrophs (Thomas et al. 1993) and chromaffin cells (Neher & Zucker, 1993), rapid rise in [Ca²⁺]_i beyond the tens of micromolar range triggered multiple kinetically distinct components of exocytosis. The fastest kinetic component (exocytic burst) reflects the depletion of a pool of granules that have reached the last station of the exocytic pathway (readily releasable pool of granules). We have previously shown that the pool of most readily releasable granules in gonadotrophs can be exhausted during the first Ca²⁺ wave of intracellular Ca²⁺ release (Tse et al. 1997). Thus, it is possible that the weak exocytic response observed in corticotrophs is related to a smaller number of granules that are readily available for release. To test this idea, we investigated the size of the pool of most readily releasable granules in corticotrophs. Figure 6 shows two typical exocytic responses in corticotrophs when [Ca²⁺]_i was rapidly elevated via flash photolysis of caged Ca-DM-nitrophen. Shortly after the UV flash, [Ca²⁺]_i rose rapidly beyond 20 μm and triggered two kinetically distinct component of exocytosis. Within ∼100 ms after the flash, a fast exocytic burst added ∼100 fF to the cell membrane capacitance. The fast exocytic burst either came to a halt (Fig. 6A) or was followed by a rapid endocytosis (Fig. 6B). In both types of exocytic response, the fast burst was followed by a slow component of exocytosis that continued for many seconds.

Figure 6. Exocytic response to a rapid and large increase in [Ca²⁺]_i triggered by flash photolysis of caged Ca²⁺.

A, two kinetically distinct components of exocytosis were triggered by the rapid and large jump in [Ca²⁺]_i. A fast exocytic burst increased the membrane capacitance by 100 fF and it was followed by a slower component. B, the fast exocytic burst was followed by rapid endocytosis in some cells. Nevertheless, a slower component can still be detected. In these experiments, the pipette contained DM-nitrophen and the cell was held at −70 mV (DC). The Ca²⁺-activated K⁺ channel was blocked by 0.4 μm apamin. To allow sufficient time for DM-nitrophen to diffuse into the cell, recording typically started ∼5 min after establishment of the whole cell configuration. The time ‘zero’ denotes the time of delivery of the UV flash.

Since the fast exocytic burst reflects the exhaustion of the most readily releasable pool, the size of this pool can be measured from the amplitude of the burst. Figure 7A plots the amplitude of the fast exocytic burst measured from twenty-five corticotrophs. In these experiments, various [Ca²⁺]_i values in individual corticotrophs were achieved by altering the intensity of the UV flash. Note that the amplitude of the fast exocytic burst is dependent on [Ca²⁺]_i. At high [Ca²⁺]_i (>10 μm), the amplitude of the fast exocytic burst appeared to saturate at ∼120 fF. Since corticotrophs vary in size (range, 2.5–8 pF), we normalized the amplitude of the fast exocytic burst to the initial capacitance of individual cells in Fig. 7A. The normalized data were then plotted as increases in cell surface area in Fig. 7B. Note that at [Ca²⁺]_i beyond 15 μm, the increase in the cell surface area reached a maximum, suggesting that a pool of most readily releasable granules in corticotrophs was exhausted during the fast exocytic burst. The average of all the data points from cells with [Ca²⁺]_i > 15 μm is 2.8 ± 0.1 % (n = 14). Thus, the exhaustion of the pool of most readily releasable granules in corticotrophs should contribute to an average increase of ∼2.8 % of the cell surface area. For an average sized corticotroph of 4 pF, the pool size will be ∼110 fF. Since the average diameter of the ACTH-containing granules in rat corticotrophs was ∼200 nm (Childs, 1992), each granule should contribute a capacitance of ∼1.3 fF. Therefore, approximately eighty-five granules are most readily releasable in a single rat corticotroph.

Figure 7. Size of the pool of readily releasable granules.

A, plot of the amplitude of the fast exocytic burst (ΔC_m) versus[Ca²⁺]_i. Initially, ΔC_m increases with [Ca²⁺]_i but appears to reach a maximum at ∼100–150 fF, reflecting the depletion of the pool of readily releasable granules at high [Ca²⁺]_i. B, the exhaustion of the readily releasable pool of granules causes an average increase of 2.8 % of the cell surface area. In this plot, the amplitude of the fast exocytic burst from each cell was normalized as the percentage increase of its initial membrane capacitance. The increase in cell surface area reached a maximum at [Ca²⁺]_i > 15 μm. The dotted line at 2.8 % was the average of all data points from cells with [Ca²⁺]_i > 15 μm. The experimental condition was the same as in Fig. 6. Each data point was generated from 2–6 cells.

DISCUSSION

NA-induced [Ca²⁺]_i oscillation

Our results show that in rat pituitary corticotrophs, NA induces a [Ca²⁺]_i oscillation that in turn stimulates a rhythmic apamin-sensitive Ca²⁺-activated K⁺ current (Fig. 1). Similar agonist-induced [Ca²⁺]_i oscillations have been observed in a variety of cell types, including the pituitary gonadotrophs, which exhibit a robust [Ca²⁺]_i oscillation and a rhythmic apamin-sensitive Ca²⁺-activated K⁺ current in response to their natural stimulating hormone, gonadotropin-releasing hormone (GnRH) (Tse & Hille, 1992). Our result also shows that the NA-induced Ca²⁺ response in corticotrophs is mediated via the α-adrenergic receptors (Fig. 2). An α-adrenergic receptor-induced [Ca²⁺]_i oscillation has also be observed in rat hepatocytes (Sanchez-Bueno et al. 1993). Among the α-adrenergic receptors, the α₁ subtype is coupled to the G-protein G_q and the phospholipase C pathway. The α₂ subtype, on the other hand, is coupled to the G-proteins G_i and G_o. Since the NA-induced [Ca²⁺]_i elevation involves the phosphoinositide pathway (Fig. 3), it is likely that the α₁-adrenergic receptor subtype is primarily involved. The frequency of the NA-induced [Ca²⁺]_i oscillation in corticotrophs (0.02–0.08 s⁻¹) is relatively slow when compared with the GnRH response in gonadotrophs (0.2–0.3 s⁻¹; Hille et al. 1994) but is similar to that triggered by phenylephrine in hepatocytes (Sanchez-Bueno et al. 1993).

In this study, flash photolysis of caged IP₃ triggers only a transient Ca²⁺ rise in corticotrophs (Fig. 3C). If a maintained or oscillatory level of IP₃ was essential for the generation of the [Ca²⁺]_i oscillation, such conditions could not be met by the flash photolysis of caged IP₃ as the UV flash caused only a transient rise in IP₃ concentration. However, our previous study (Tse & Lee, 1998) suggests that the requirement for a maintained IP₃ level for [Ca²⁺]_i oscillation in corticotrophs is unlikely. When the concentration of IP₃ in corticotrophs was maintained at a high level via the continuous supply of IP₃ (10 μm) from the whole-cell pipette, only a ‘transient and plateau’ pattern of [Ca²⁺]_i elevation was elicited. At present, it is not clear whether an oscillatory level of IP₃ is essential for [Ca²⁺]_i oscillation in corticotrophs. In pituitary gonadotrophs, however, robust [Ca²⁺]_i oscillations could be elicited when the level of IP₃ was maintained via a continuous supply of either IP₃ (Tse et al. 1995) or a non-hydrolysable analogue of IP₃ from the whole-cell pipette (Stojilkovic et al. 1993). It is also possible that other than releasing Ca²⁺ from IP₃-sensitive stores, NA may have additional action on corticotrophs to elicit [Ca²⁺]_i oscillation. One of these possibilities involves PKC, which is activated by diacylglycerol during NA stimulation. However, the involvement of PKC is unlikely because other ACTH secretagogues such as AVP and OT, which also stimulate the phosphoinositide pathway in corticotrophs, trigger a ‘transient and plateau’ pattern of [Ca²⁺]_i elevation (Leong, 1988; Link et al. 1992; Corcuff et al. 1993; Tse & Lee, 1998). Another possibility involves the activation of a cAMP-dependent pathway via β-adrenergic receptors. Since a NA-induced [Ca²⁺]_i oscillation persisted in the presence of propranolol, a β-adrenergic blocker, the critical involvement of the cAMP pathway can be ruled out. In gonadotrophs, the robust [Ca²⁺]_i oscillation has been partially attributed to a rapid Ca²⁺ uptake into the intracellular stores and the presence of a large intracellular Ca²⁺ reserve in the IP₃-sensitive stores (Tse et al. 1994a,b). Thus, a possible mechanism underlying the NA-induced oscillation may involve an increase in the activity of the intracellular Ca²⁺ pumps such that cytosolic Ca²⁺ can be pumped rapidly back to the intracellular stores. Further experiments will be needed to test these possibilities.

NA triggers exocytosis in corticotrophs

Results from this study demonstrate that the first wave of NA-induced [Ca²⁺]_i oscillation is accompanied by an increase in membrane capacitance (reflecting exocytosis) in corticotrophs (Fig. 5A). However, no increase in capacitance could be detected in the subsequent waves of NA-induced [Ca²⁺]_i oscillation. Similarly, consecutive flash photolysis of caged IP₃ (∼60 s apart) triggered exocytosis only during the first [Ca²⁺]_i elevation, although the amplitude of the second [Ca²⁺]_i elevation was comparable to that triggered during the first wave of NA-induced [Ca²⁺]_i oscillation (Fig. 5B). In contrast, the initial one to three cycles of [Ca²⁺]_i oscillation in gonadotrophs (induced by GnRH or flash photolysis of caged IP₃) are often accompanied by significant exocytosis (Tse et al. 1993; Tse et al. 1997). In gonadotrophs, a single large Ca²⁺ wave (2–3 μm) can often exhaust the entire pool of most readily releasable granules. Therefore, we examined whether the weak exocytic response in corticotrophs may be related to the size of its readily releasable pool. Our results (Fig. 7) indicate that approximately eighty-five granules are most readily releasable in a corticotroph. The exhaustion of this pool of granules will increase the cell surface area of the corticotroph by 2.8 %. Interestingly, this value is only about half of what is reported in other types of pituitary cells. In gonadotrophs (Tse et al. 1997) and melanotrophs (Thomas et al. 1993), the depletion of the readily releasable pool added 4.7 and 5.3 %, respectively, to the cell membrane surface area. This result suggests that the weak exocytic response in corticotrophs may be partially attributed to the smaller number of readily releasable granules. Indeed, ultrastructural studies (reviewed by Childs, 1992) have suggested that among the anterior pituitary cells, corticotrophs have relatively few granules. However, the cumulative increase in surface area during NA or flash photolysis of caged IP₃ in a corticotroph was only ∼1.5 %. Thus, only about half of the most readily releasable granules were released during the first Ca²⁺wave. Since the capacitance measurement detects changes in membrane surface area, some of the smaller and perhaps slower increase in capacitance during subsequent [Ca²⁺]_i elevations may be masked by the robust endocytosis in these cells (e.g. Figs 5B and 6B). Nevertheless, the mean transient increase in capacitance stimulated by NA was 57.7 ± 10.7 fF when the average peak [Ca²⁺]_i was 1.43 ± 0.2 μm. This value closely resembled that obtained during the AVP-induced biphasic Ca²⁺ elevation where the cumulative increase in capacitance was 49 ± 13 fF at a [Ca²⁺]_i of 1.76 ± 0.15 μm (Tse & Lee, 1998).

In this study, we have shown that the NA-induced Ca²⁺ response in corticotrophs was mediated via the α-adrenergic receptor-coupled phosphoinositide pathway. The generation of IP₃ triggered intracellular Ca²⁺ release and the rise in [Ca²⁺]_i is sufficient to act as the initial trigger of exocytosis. However, during NA stimulation, PKC as well as adenylate cyclase may also be activated. While these two messenger pathways do not contribute significantly to the NA-induced Ca²⁺ response or the initial trigger of exocytosis, their involvement in ACTH secretion cannot be ruled out totally. In pituitary melanotrophs, PKC has been shown to enhance Ca²⁺-dependent secretion (Lee, 1996). On the other hand, cAMP has been shown to enhance Ca²⁺-dependent secretion in pituitary lactotrophs (Sikdar et al. 1990), melanotrophs (Lee, 1996) and pancreatic β cells (Ämmälä et al. 1993). Thus, it is possible that both PKC and cAMP may have a role in modulating Ca²⁺-dependent exocytosis in corticotrophs.