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MEMBRANE TRANSPORTERS, ION CHANNELS, AND PUMPS

Nitric oxide induces [Ca²⁺]_i oscillations in pituitary GH₃ cells: involvement of I_DR and ERG K⁺ currents

Division of Pharmacology, Department of Neuroscience, School of Medicine, Federico II University of Naples, Naples, Italy

Submitted 12 May 2005 ; accepted in final form 2 September 2005

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION APPENDIX GRANTS REFERENCES

 
The role of nitric oxide (NO) in the occurrence of intracellular Ca²⁺ concentration ([Ca²⁺]_i) oscillations in pituitary GH₃ cells was evaluated by studying the effect of increasing or decreasing endogenous NO synthesis with L-arginine and nitro-L-arginine methyl ester (L-NAME), respectively. When NO synthesis was blocked with L-NAME (1 mM) [Ca²⁺]_i, oscillations disappeared in 68% of spontaneously active cells, whereas 41% of the quiescent cells showed [Ca²⁺]_i oscillations in response to the NO synthase (NOS) substrate L-arginine (10 mM). This effect was reproduced by the NO donors NOC-18 and S-nitroso-N-acetylpenicillamine (SNAP). NOC-18 was ineffective in the presence of the L-type voltage-dependent Ca²⁺ channels (VDCC) blocker nimodipine (1 µM) or in Ca²⁺-free medium. Conversely, its effect was preserved when Ca²⁺ release from intracellular Ca²⁺ stores was inhibited either with the ryanodine-receptor blocker ryanodine (500 µM) or with the inositol 1,4,5-trisphosphate receptor blocker xestospongin C (3 µM). These results suggest that NO induces the appearance of [Ca²⁺]_i oscillations by determining Ca²⁺ influx. Patch-clamp experiments excluded that NO acted directly on VDCC but suggested that NO determined membrane depolarization because of the inhibition of voltage-gated K⁺ channels. NOC-18 and SNAP caused a decrease in the amplitude of slow-inactivating (I_DR) and ether-à-go-go-related gene (ERG) hyperpolarization-evoked, deactivating K⁺ currents. Similar results were obtained when GH₃ cells were treated with L-arginine. The present study suggests that in GH₃ cells, endogenous NO plays a permissive role for the occurrence of spontaneous [Ca²⁺]_i oscillations through an inhibitory effect on I_DR and on I_ERG.

voltage-gated potassium channels; ether-à-go-go-related gene potassium channels; slow-inactivating outward currents; fast-inactivating outward currents

The burgeoning literature on the molecular mechanisms underlying [Ca²⁺]_i oscillations clearly reflects the crucial functional and physiological relevance ascribed to their occurrence (24, 46). To date, multiple lines of evidence have clearly established that two different systems take part in the modulation of [Ca²⁺]_i oscillations in pituitary cells: an intracellular oscillator and a plasma membrane oscillator (39). The former comprises the ryanodine- and the inositol 1,4,5-trisphosphate receptor (IP₃)-sensitive intracellular Ca²⁺ stores and their refilling channels and pumps, whereas the latter relies on the coordinated activity of different classes of plasma membrane voltage-gated and second messenger-activated ion channels. The genesis of spontaneous [Ca²⁺]_i oscillations is predominantly dependent on the activity of the plasma membrane oscillator in pituitary cells (12, 39). In particular, [Ca²⁺]_i oscillations appear to be a result of spontaneous action potentials (16) triggered by the opening of both L-type voltage-dependent Ca²⁺ channels (VDCC) and nonspecific cation channels (45). The ensuing massive influx of Ca²⁺ ions into the cytoplasm is followed by the activation of Ca²⁺-dependent, large-conductance K⁺ (BK) channels (25), which repolarize the cell by cooperating with other K⁺ channels belonging to the family of delayed rectifiers.

Clear evidence has been demonstrated for at least two other classes of voltage-gated K⁺ (Kv) channels involved in the appearance of [Ca²⁺]_i oscillations: the ether-à-go-go-related gene (ERG) channels (5, 35) and the channels maintaining so-called fast-inactivating outward currents (I_A) (21). These ion channels play a pivotal role in controlling resting potential and, hence, in regulating the tendency of the cell to undergo action potentials and [Ca²⁺]_i oscillations. In turn, the activity of these ion channels can be modulated by many well-characterized intracellular transduction pathways that provide the cells with further mechanisms for finely tuning [Ca²⁺]_i oscillations. It has been clearly demonstrated that there is a strict relationship between [Ca²⁺]_i oscillations and cAMP-PKA-dependent (18, 52), PKC-dependent (38), and PTK-dependent pathways (7). Intriguingly, many of the ion channels that play a role in the complex genesis of [Ca²⁺]_i oscillations are also regulated by nitric oxide (NO), a gaseous messenger that is generated from the enzymatic conversion of L-arginine into citrulline by nitric oxide synthase (NOS) (for review, see Ref. 3). In particular, NO is able to interfere both with ion channels responsible for triggering and maintaining the action potentials, such as TTX-sensitive voltage-dependent Na⁺ channels or VDCC, and with ion channels responsible for repolarizing the cell or for clamping the membrane to resting potential, such as ERG K⁺ channels or large- and small-conductance Ca²⁺-dependent K⁺ channels (for review, see Ref. 3).

Therefore, on the basis of these findings, we hypothesized that endogenous NO might interfere with the process of [Ca²⁺]_i oscillations in pituitary cells, because they express the enzyme NOS and spontaneously release NO (31). To test this hypothesis in the present study, we explored NO involvement in [Ca²⁺]_i oscillations in pituitary growth hormone GH₃ cells, a cell line derived from a radiation-induced rat pituitary tumor (44) that expresses high concentrations of the neuronal isoform of NOS (nNOS) (36, 49) and several classes of voltage-gated ion channels (17, 28, 50). Furthermore, large spontaneous [Ca²⁺]_i oscillations occur in 70% of fura-2 AM-loaded GH₃ cells, whereas the remaining cells are quiescent. To date, the factors responsible for determining whether a given GH₃ cell will spontaneously oscillate or remain quiescent remain totally undefined. Intriguingly, we report herein that endogenous NO is a major determinant of the occurrence of spontaneous [Ca²⁺]_i oscillations in GH₃ cells because of its inhibitory effect on a slow-inactivating component due to activation of delayed rectifiers (I_DR) and hyperpolarization-evoked deactivating K⁺ currents carried by ERG channels (I_ERG).

	MATERIALS AND METHODS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION APPENDIX GRANTS REFERENCES

 
Cell culture. GH₃ cells were obtained from Flow Laboratories (Irvine, Scotland) and grown in plastic dishes in Ham's F-10 medium (GIBCO/Invitrogen, Milan, Italy) supplemented with 15% horse serum (Flow Laboratories), 2.5% FCS (HyClone, Logan, UT), 100 U/ml penicillin, and 100 µg/ml streptomycin. The cells were cultured in a humidified 5% CO₂ atmosphere, and the culture medium was changed every 2 days. For microfluorometric and electrophysiological studies, the cells were plated on glass coverslips (Fisher, Springfield, NJ) coated with poly-L-lysine (30 µg/ml) (Sigma, St. Louis, MO) and used at least 12 h after being seeded.

[Ca²⁺]_i measurements and quantification of [Ca²⁺]_i oscillations. [Ca²⁺]_i was measured using single-cell computer-assisted video imaging (35). Briefly, GH₃ cells grown on glass coverslips were loaded with 5 µM fura-2 AM for 1 h at room temperature in normal Krebs solution containing (in mM) 5.5 KCl, 160 NaCl, 1.2 MgCl₂, 1.5 CaCl₂, 10 glucose, and 10 HEPES-NaOH, pH 7.4. At the end of the fura-2 AM loading period, the coverslips were placed into a perfusion chamber (Medical System, Greenvale, NY) mounted on the stage of an inverted Nikon Diaphot fluorescence microscope (Nikon, Torrance, CA). A 100-W Xe lamp (Osram, Berlin, Germany) with a computer-operated filter wheel bearing two different interference filters (340 and 380 nm) illuminated the microscopic field with UV light every 3 s, alternating the wavelengths at an interval of 500 ms. The light emitted by fura-2 AM-loaded cells was passed through a 400-nm dichroic mirror filtered at 510 nm and collected using an intensified camera (Photonic Science, Robertsbridge, UK). Images were digitized and analyzed using a Magiscan image processor (Applied Imaging, Dukesway, UK) driven by AutoLab software (RBR; Altair, Florence, Italy). Ratiometric values were automatically converted by the software into [Ca²⁺]_i using a preloaded calibration curve obtained in preliminary experiments (35).

Analysis of [Ca²⁺]_i oscillations. [Ca²⁺]_i oscillations were identified and their amplitude and frequency were determined using a computer program written in Java computer language. The details of the algorithm are reported in the APPENDIX. Briefly, for each single cell, the software calculated the [Ca²⁺]_i mean ± SD during the baseline recording interval before any drug addition; these values were used to define a cutoff to identify [Ca²⁺]_i oscillation, which was set at mean [Ca²⁺]_i ± 2 SD. Subsequently, the software identified as a single [Ca²⁺]_i oscillation each group of consecutive [Ca²⁺]_i values higher than this cutoff point, provided that this group was preceded and followed by at least one [Ca²⁺]_i value lower than the cutoff point. To quantify the effect of specific pharmacological treatments on the occurrence of [Ca²⁺]_i oscillations, the following parameters were determined: the oscillation peak, defined as the maximal [Ca²⁺]_i attained during a single [Ca²⁺]_i oscillation; the oscillation amplitude, defined as the difference between transient peak [Ca²⁺]_i and mean basal [Ca²⁺]_i; and the oscillation frequency, defined as the number of peaks divided by the duration of observation (21).

To evaluate the effect of the different drugs on the pattern of spontaneous [Ca²⁺]_i oscillations, we compared mean oscillation frequency and mean oscillation amplitude recorded before (i.e., control) and during treatment. In control experiments, no significant changes in either the frequency or the amplitude occurred after the addition of drug vehicles.

Nitrite detection. NO generation in GH₃ cells was determined using the Griess reaction assay (40, 42). Briefly, GH₃ cells plated in 100-mm petri dishes were incubated at 37°C in 4 ml of normal Krebs solution supplemented either with the appropriate concentration of the drugs under investigation or with vehicle. Incubation medium in 500-µl volumes was collected 300 s after incubation began. Griess reagent (500 µl; 1% sulfanilamide and 0.1% naphthyl ethylenediamine in 2% H₃PO₄) was then added to each sample. The tubes were mixed and left to stand for 10 min at room temperature. At the end of this time, the sample's absorbance was measured at 550 nm. A reference curve was prepared with NaNO₂ used as a standard to convert the absorbance values into nanomolar nitrite concentrations.

Electrophysiology. K⁺ currents were recorded with the perforated-patch configuration of the whole cell technique using fire-polished borosilicate electrodes with a final resistance of 2.5–4 M back-filled with an internal solution containing nystatin (120–240 µg/ml) and (in mM) 140 KCl, 2 MgCl₂, 10 HEPES, 10 glucose, 10 EGTA, and 1 Mg²⁺-ATP, pH 7.4 (adjusted with KOH) (30). The cells, plated onto glass coverslips, were then placed into a perfusion chamber mounted on the stage of a Diaphot inverted microscope (Nikon, Torrance, CA).

Patch-clamp recordings were performed at room temperature (20–22°C) using a Digidata 1200 interface (Axon Instrument, Foster City, CA) and a commercially available amplifier (Axopatch 200A; Axon Instrument) driven by pCLAMP 6.0.4 software (Axon Instrument) run on a personal computer. Currents were filtered at 5 kHz. No compensation for pipette resistance or cell capacitance was performed, because in our experimental condition using 2.5- to 4-M electrodes, the series resistance was 6–8 M. In addition, most of the currents recorded were <0.5 nA, and therefore, in the absence of any compensation, the expected voltage error was <5 mV. Similarly, we did not compensate for cell capacitance, owing to the modest size of the cells (18.2 ± 0.8 pF; n = 40). Data were saved onto a computer disk for off-line analysis performed using Clampfit 6.0.4 (Axon Instrument) and SigmaPlot 5.0 software (Jandel Scientific, San Rafael, CA).

The outward and inward K⁺ currents were recorded using different extracellular solutions and stimulation protocols. For outward K⁺ current recordings, depolarizing voltage steps (300-ms duration) of increasing voltage from –80 mV to +40 mV preceded by conditioning pulses at –100 mV lasting 1.5 s were applied to GH₃ cells held at –80 mV and continuously perfused with a 5.4 mM KCl external solution containing (in mM) 150 NaCl, 5.4 KCl, 2 CaCl₂, 1 MgCl₂, and 10 HEPES, pH 7.4 (adjusted with NaOH). The two different components contributing to these outward K⁺ currents, I_A and I_DR, were separated by virtue of their different susceptibility to voltage-dependent inactivation. In particular, when the voltage of the depolarizing prepulse was set at –40 mV instead of –100 mV, I_A, which is extremely sensitive to steady-state inactivation, was virtually canceled, whereas I_DR was unaffected. The I_A component was thus obtained by subtracting the isolated I_DR component from the total K⁺ current.

Hyperpolarization-evoked deactivating K⁺ currents were evoked by applying a series of hyperpolarizing voltage steps (125-ms duration) of increasing amplitude (from 0 to –160 mV in –10-mV increments) to GH₃ cells held at 0 mV and perfused with a 54 mM KCl external solution containing (in mM) 96 NaCl, 54 KCl, 2 CaCl₂, 1 MgCl₂, and 10 HEPES, pH 7.4, adjusted with NaOH. To inactivate I_DR, each voltage step was preceded by a 10-s depolarizing prepulse to 0 mV. The currents evoked by this protocol consisted of at least two different components, a fast-inactivating current representing I_ERG that was blocked by 1 µM astemizole and a sustained, noninactivating current representing ill-defined contaminating currents resistant to the astemizole blockade. I_ERG was isolated by subtracting the currents that persisted after astemizole blockade from the total inwardly rectifying K⁺ currents elicited by the above-described protocol.

Ca²⁺ currents from GH₃ cells were recorded with an extracellular solution containing (in mM) 10 BaCl₂, 125 N-methyl-D-glucamine, 1 MgCl₂, and 10 HEPES, pH 7.3. The osmolarity of this solution was adjusted to 300 mosM with sucrose solution. The pipettes were filled with (in mM) 110 CsCl, 30 tetraethylammonium (TEA)-Cl, 2 MgCl₂, 10 EGTA, 8 glucose, 5 ATP, 15 creatine phosphate, 1 GTP, and 10 HEPES, pH 7.3. The holding potential (HP) of GH₃ cells was –80 mV. Ba²⁺ currents flowing through L-type VDCC were activated by ramping voltages from –80 mV to +40 mV (75-ms duration) with one pulse every 15 s.

Statistical analysis. All data are reported as means ± SE. Statistical analysis was performed using a paired Student's t-test or ANOVA followed by the Newman-Keuls post hoc test for unpaired data as appropriate. The threshold for statistical significance was set at P < 0.01.

Drugs and chemicals. Fura-2 AM and ryanodine were obtained from Calbiochem (La Jolla, CA). The NO donors (Z)-1-[N-(2-aminoethyl)-N-(2-ammonioethyl)amino]diazen-1-ium-1,2-diolate (DETA-NONOate, hereinafter referred to as NOC-18) and S-nitroso-N-acetylpenicillamine (SNAP) were purchased from Cayman Chemical (Ann Arbor, MI). Charybdotoxin (CTX) and paxillin (PAX) were obtained from Alomone Laboratories (Jerusalem, Israel). Astemizole was kindly provided by Janssen-Cilag (Rome, Italy). Xestospongin C was kindly provided by Dr. L. Santella (Laboratory of Cell Signaling, Stazione Zoologica A. Dohrn, Naples, Italy). All other chemicals were purchased from Sigma.

NOC-18, SNAP, L-arginine, and nitro-L-arginine methyl ester (L-NAME) were dissolved at the appropriate concentrations immediately before each experiment. Stock solutions of the other chemicals were prepared in DMSO and kept at –20°C. Appropriate dilutions were prepared daily.

	RESULTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION APPENDIX GRANTS REFERENCES

 
Effect of NOS substrate L-arginine, NOS inhibitor L-NAME, and NO donors NOC-18 and SNAP on [Ca²⁺]_i oscillations in GH₃ cells. As expected and as previously reported by our laboratory and others (35, 39, 47), the [Ca²⁺]_i in single fura-2 AM-loaded GH₃ cells showed significant oscillations over time (Fig. 1A). Interestingly, not all cells showed spontaneous [Ca²⁺]_i oscillations and, according to the criteria reported in MATERIALS AND METHODS, 30% of the entire cell population was quiescent (Fig. 1B).

View larger version (33K): [in this window] [in a new window]   Fig. 1. Effect of L-arginine, nitro-L-arginine methyl ester (L-NAME), (Z)-1-[N-(2-aminoethyl)-N-(2-ammonioethyl)amino]diazen-1-ium-1,2-diolate (DETA-NONOate, or NOC-18), and S-nitroso-N-acetylpenicillamine (SNAP) on nitric oxide (NO) generation and intracellular Ca2+ concentration ([Ca2+]i) oscillations in pituitary GH3 cells. A and B: time course of [Ca2+]i in two different fura-2 AM-loaded GH3 cells, which are representative of the group of spontaneously oscillating and quiescent cells. C: bar graph showing the extracellular nitrite concentrations measured in control (CTL) conditions and in the presence of L-NAME (1 mM), L-arginine (10 mM), NOC-18 (1 µM), or SNAP (1 µM). Each bar represents the mean ± SE of the values obtained for each experimental group in 3 different experimental sessions. *P < 0.01. D: effect of NO synthase (NOS) substrate L-arginine (10 mM) on a single quiescent fura-2 AM-loaded GH3 cell. This trace is representative of 150 cells studied in at least 3 different experimental sessions. E: effect of NOC-18 (1 µM) on a single quiescent fura-2 AM-loaded GH3 cell. This trace is representative of 171 cells studied in at least 3 different experimental sessions. F: effect of SNAP (1 µM) on a single quiescent fura-2 AM-loaded GH3 cell. This trace is representative of 100 cells studied at least in three different experimental sessions. G and H: effect of the NOS inhibitor L-NAME (1 mM) alone or in the presence of L-arginine (10 mM) on spontaneous [Ca2+]i oscillations. These traces are representative of 60 and 75 cells, respectively, studied in at least 3 different experimental sessions. I: effect of NOC-18 (1 µM) on L-NAME (1 mM)-induced inhibition of spontaneous [Ca2+]i oscillations in a single fura-2 AM-loaded GH3 cell. This trace is representative of 70 cells studied in at least 3 different experimental sessions. The drugs were introduced into the perfusion chamber and left therein for the time indicated by the bar.

Interestingly, when added after a 200 s treatment with L-NAME, L-arginine (10 mM) did not induce [Ca²⁺]_i oscillations (Fig. 1H). To confirm that L-NAME was affecting the occurrence of [Ca²⁺]_i oscillations in GH₃ cells by suppressing NO generation, we explored the ability of exogenous NO to rescue GH₃ cells from the effects of L-NAME. As shown in Fig. 1I, 1 µM NOC-18 was able to restore [Ca²⁺]_i oscillations in the presence of 1 mM L-NAME, further confirming that NO is responsible for maintaining [Ca²⁺]_i oscillations in GH₃ cells.

The observed effect of NO donors and inhibitors on [Ca²⁺]_i oscillations raised the important issue of determining whether this gaseous mediator was eliciting [Ca²⁺]_i oscillations by promoting the influx of Ca²⁺ ions from the extracellular solution or by regulating its release from the intracellular Ca²⁺ stores. To address this point, we first examined whether NOC-18 was able to induce [Ca²⁺]_i oscillations in the absence of extracellular Ca²⁺. When the drug was added to a perfusion chamber in a Ca²⁺-free solution in which Ca²⁺ ions were omitted from normal Krebs solution and 1.5 mM EGTA was added, NOC-18 was ineffective (Fig. 2A). This finding suggests that the effect of NOC-18 on [Ca²⁺]_i oscillations was critically dependent on Ca²⁺ influx from the extracellular solution. To establish whether this influx took place through L-type VDCC, which represent the privileged Ca²⁺ influx pathway for spontaneous [Ca²⁺]_i oscillations in GH₃ cells (8), we explored the consequences of the selective blockade of these channels with nimodipine on the ability of NOC-18 to induce [Ca²⁺]_i oscillations. At a concentration of 1 µM, this drug was able to prevent NO-induced [Ca²⁺]_i oscillations in GH₃ cells (Fig. 2B), strengthening the idea that NOC-18 promoted Ca²⁺ influx through L-type VDCC. The results of these experiments strongly suggest that intracellular Ca²⁺ stores are not essential to triggering NOC-18-induced oscillations, but these results do not provide information regarding the role that these stores play in maintaining this oscillatory phenomenon over time. To assess this point, we studied the effect of the selective blockade of either the ryanodine- or IP₃-sensitive stores, which are both activated in spontaneous [Ca²⁺]_i oscillations in the context of the so-called Ca²⁺-induced Ca²⁺ release (CICR) (39). In the presence either selective blocker of ryanodine stores, ryanodine (500 µM) (4) or the IP₃ receptor blocker xestospongin C (3 µM) (13), NOC-18 induced the appearance of large [Ca²⁺]_i oscillations; however, the oscillatory phenomenon was not maintained over time and rapidly faded (Fig. 2, C and D). Interestingly, the early induction of [Ca²⁺]_i oscillations was prevented completely by nimodipine (1 µM) (Fig. 2, E and F). These data suggest that NOC-18 promoted the early influx of Ca²⁺ ions through L-type VDCC but that the generation of sustained [Ca²⁺]_i oscillations critically required the simultaneous activation of ryanodine and IP₃ stores.

View larger version (38K): [in this window] [in a new window]   Fig. 2. Effect of NOC-18 on [Ca2+]i oscillations in GH3 cells bathed in a Ca2+-free medium or exposed either to the L-type voltage-dependent Ca2+ channels (VDCC) blocker nimodipine, to the ryanodine receptor blocker ryanodine, or to the inositol 1,4,5-trisphosphate (IP3) receptor blocker xestospongin C. A: effect of NOC-18 (1 µM) on [Ca2+]i oscillations in a single quiescent GH3 cell bathed in Ca2+-free medium. Trace shown was obtained in a single cell that is representative of 56 more cells recorded in 3 different experimental sessions. B: effect of NOC-18 (1 µM) in the presence of nimodipine (1 µM) in a single quiescent GH3 cell that is representative of 100 cells recorded in 3 different experimental sessions. C: effect of NOC-18 (1 µM) on [Ca2+]i in a single quiescent GH3 cell treated with ryanodine (500 µM). Trace was obtained in a cell that is representative of 150 cells studied in at least 3 different experimental sessions. D: effect of NOC-18 (1 µM) on [Ca2+]i in a single quiescent GH3 cell treated with xestospongin C (3 µM). Trace is representative of 120 cells that were studied in at least 3 different experimental sessions. E and F: effect of nimodipine on [Ca2+]i oscillations induced by NOC-18 in the presence of ryanodine and/or xestospongin C. Traces are representative of 120 and 100 cells obtained in 3 different experimental sessions for the ryanodine and xestospongin C groups, respectively. The drugs were introduced into the perfusion chamber and left therein for the time indicated by the bar.

View larger version (18K): [in this window] [in a new window]   Fig. 3. Effect of NO donor NOC-18 on VDCC activity in GH3 cells. The superimposed traces show the Ba2+ currents elicited in the same cell using ramp depolarization and delivered accordingly to the protocol shown below the traces under control conditions and during exposure to 1 µM NOC-18 or to 500 µM Cd2+. Traces were obtained in a cell that is representative of 7 cells studied in at least 3 different experimental sessions.

View larger version (33K): [in this window] [in a new window]   Fig. 4. Effect of L-arginine, L-NAME, and NO donors NOC-18 and SNAP on outwardly rectifying K+ currents in GH3 cells. A: outwardly rectifying K+ currents elicited by the depolarization protocol (right) before (left) and after (middle) the addition of 1 µM NOC-18 to the perfusion medium in a single GH3 cell that is representative of a group of 12 recorded in 3 different experimental sessions. B: mean current-voltage (I-V) plots for the slow-inactivating component due to activation of delayed rectifiers (IDR; left) and fast-inactivating outward current component (IA; right) of the outward K+ currents elicited by the depolarization protocol (shown in A, right) and dissected as reported in MATERIALS AND METHODS. These plots were generated by averaging the values of maximal IDR and IA outward currents recorded in these cells in response to each depolarizing step. To allow comparison among groups, data were normalized for the maximal outward current recorded at +40 mV. *P < 0.01. C: time course of outward K+ currents recorded in GH3 cells exposed to 10 mM L-arginine (top; n = 9 in 3 different experimental sessions) or 1 µM NOC-18 (bottom; n = 9 in 3 different experimental sessions). The currents were measured at +40-mV depolarization potential at the end of the pulse for each time course experiment. D: bar graph showing mean ± SE of IDR %inhibition induced by drug addition in each cell of the groups treated with L-arginine, L-arginine plus L-NAME, SNAP, or NOC-18. *P < 0.01.

To determine whether the NO-induced inhibition of I_DR was cGMP dependent, we studied the effect of the membrane-permeable cGMP analog 8-bromoguanosine 3',5'-cyclic monophosphate (8-BrcGMP) on these currents. When this compound was added to the perfusion medium at the concentration of 150 µM, it entirely reproduced the effect of NOC-18. When outward K⁺ currents were evoked by step depolarization from –80 up to +40 mV after 1.5-s prepulses at –40 mV, 1 µM NOC-18 induced a 31 ± 3.8% inhibition of I_DR (n = 12), which was similar to that of 150 µM 8-BrcGMP (24 ± 4%; n = 4).

Because NO significantly slows down I_A inactivation (11), the inactivation kinetics of I_A elicited by step depolarization to +40 mV (duration, 300 ms; holding potential, –80 mV) was further determined in the presence and absence of 1 µM NOC-18. In both cases, I_A decayed with a biexponential time course and the drug did not induce any changes in either fast inactivation time (_fast, 10.7 ± 2 vs. 12 ± 2.6 ms; n = 9) or slow inactivation time (_slow, 202 ± 29 vs. 200 ± 20 ms; n = 9) (Fig. 5). The decay of the inactivating current obtained both before and after NOC-18 perfusion was fitted using a biexponential function Y = A_fast exp(–t/_fast) + A_slow exp(–t/_slow) + C, where A_fast and A_slow represent the absolute amplitudes of the fast and slow components, _fast and _slow are the intrinsic time constants of these components, and C is an offset factor with fast- and slow-inactivating components.

View larger version (15K): [in this window] [in a new window]   Fig. 5. Effect of NOC-18 on the inactivation kinetics of IA. A: IA isolated by subtraction as described in MATERIALS AND METHODS in a single GH3 cell before and after exposure to NOC-18 (1 µM). B: left, means ± SE (n = 9) of the fast inactivation time constant (fast) of fast-decaying component obtained under control conditions compared with that obtained after perfusion with NOC-18; right, means ± SE (n = 9) of slow inactivation time constant (slow) of the slow-decaying component. Decay of the inactivating current was fitted using the following biexponential function: Y = Afast exp(–t/fast) + Aslow exp(–t/slow) + C.

View larger version (39K): [in this window] [in a new window]   Fig. 6. Effect of L-arginine, L-NAME, the NO donors NOC-18 and SNAP, and astemizole on hyperpolarization-evoked deactivating K+ currents carried by ERG channels (IERG) in GH3 cells. A, top, left to right, inwardly rectifying K+ currents elicited in a single GH3 cell that is representative of a group of 9 under the hyperpolarization protocol (shown at right under astemizole traces), in control conditions, after 1 µM NOC-18, after washout of NOC-18 (IT), and after addition of 1 µM astemizole (IR). A, bottom, left to right, IERG isolated by subtraction as described in MATERIALS AND METHODS, in control conditions, after 1 µM NOC-18 addition, and after washout of NOC-18. B: mean I-V plots for residual component (top) and IERG (bottom) obtained in control conditions, after NOC-18 addition, and after NOC-18 washout. These plots were generated by averaging the values of maximal currents of both sustained and ERG K+ components recorded after prepulses at 0 mV in response to each hyperpolarizing step. To allow comparison between groups, each membrane potential value recorded was normalized to the corresponding maximum value recorded at –160 mV under control conditions. *P < 0.01. C: effect of L-arginine on inwardly rectifying K+ currents elicited by the hyperpolarization protocol (inset), under control conditions, after perfusion with 10 mML-arginine, after washout of L-arginine, and after perfusion with 1 µM astemizole (IR). D: bar graph showing %IERG inhibition in presence of L-arginine with or without L-NAME, NOC-18, and SNAP. Each bar represents the mean ± SE of the maximal inward current amplitude expressed as a percentage of control (n = 9 in 3 different experimental sessions). *P < 0.01.

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION APPENDIX GRANTS REFERENCES

These results indicate that NO generation is a limiting factor for [Ca²⁺]_i oscillations in GH₃ cells. They also raise the possibility that quiescent cells may not display oscillations, because they do not generate enough NO to sustain this process, owing to either a lower expression of nNOS or an inefficient L-arginine influx system. Therefore, NO appears to be one of the factors that determines whether a given GH₃ cell displays spontaneous [Ca²⁺]_i oscillations. Although previous reports suggested that NO could play a role in [Ca²⁺]_i oscillations in different cell preparations, such as cultured glial cells (48), pancreatic -cells (20), or rat cardiomyocytes (53), we have demonstrated in the present study that the extent of endogenous NO synthesis is responsible for determining whether a given cell is quiescent or spontaneously active.

That NO can trigger [Ca²⁺]_i oscillations could be of physiological relevance because the release of anterohypophyseal hormones is controlled by changes in the amplitude or frequency of [Ca²⁺]_i oscillations, which also regulate the transcription of the genes encoding these molecules. For example, it was previously demonstrated that [Ca²⁺]_i oscillations can modulate prolactin (PRL) secretion (5, 8) and that the amount of growth hormone released by single somatomammotrophs is strictly dependent on the amplitude and frequency of spontaneous [Ca²⁺]_i oscillations (19). Furthermore, Villalobos et al. (47) demonstrated that the expression of a PRL luciferase reporter gene monitored at the single-cell level was inversely related to the occurrence of [Ca²⁺]_i oscillations in primary cultures of rat somatomammotroph cells. In pituitary cells, [Ca²⁺]_i oscillations not only control the transcription of anterohypophyseal hormones but also behave as a gene transcription regulator of wider significance involved, for example, in controlling c-fos transcription via the serum response element (SRE) in the mouse corticotroph cell line AtT-20 (23). In particular, the efficiency of SRE-dependent c-fos expression and the transcription factor cAMP-response element-binding protein (CREB)-dependent c-fos expression has been shown to be critically dependent on the shape and duration of [Ca²⁺]_i oscillations in these cells (9). Because we found that NO induced an increase in [Ca²⁺]_i oscillation frequency, it is tempting to speculate that this gaseous mediator could control PRL release and gene transcription via changes in the [Ca²⁺]_i oscillatory pattern. The present findings suggest also that the factors that induce nNOS expression could promote the same effect on the oscillatory pattern in GH₃ cells via NO. As it stands, the factors regulating nNOS gene expression have been determined only partially. It is known, however, that nNOS expression is regulated by neuropeptides, namely, pituitary adenylate cyclase-activating polypeptide (15) and gonadotropin-releasing hormone (14), and by steroid hormones, namely, estrogens (10), androgens (33), and glucocorticoids (34). Intriguingly, we recently observed that in GH₃ cells, PRL is a strong inducer of both nNOS expression (36) and [Ca²⁺]_i oscillations (Secondo A, Pannellione A, Cataldi M, Sirabella R, Formisano L, Di Renzo GF, and Annunziato L, unpublished results).

The results reported herein indicate that the ability of NO to sustain [Ca²⁺]_i oscillations relies, at least in part, on its inhibitory effect on Kv channels. This finding is in accord with a wealth of experimental evidence supporting both the central role of these channels in regulating the tendency of GH₃ cells to undergo [Ca²⁺]_i oscillations (28) and their susceptibility to be regulated by NO (for review, see Ref. 3). By using specific electrophysiological approaches, different components of K⁺ currents, I_A, I_DR, and I_ERG were isolated and their susceptibility to NO was investigated in GH₃ cells.

I_A is involved in the occurrence of spontaneous [Ca²⁺]_i oscillations in GH₃ cells as suggested by Kushmerick et al. (21), who found that these channels are open at voltages equal to resting GH₃ potentials (–40 mV) and that their blockade with the Phoneutria nigriventer toxin Tx3-1 induces [Ca²⁺]_i spiking in these cells. However, we can exclude the notion that the [Ca²⁺]_i oscillation promoting effect of NO was exerted at the level of this current. In fact, in the present study, I_A was completely insensitive to the NO donor NOC-18. This finding is at variance with the results of Ciorba et al. (11), who showed that NO affected I_A inactivation by oxidizing a critical methionine residue in the ball chain domain of the channel. However, it should not be overlooked that these results were obtained using millimolar concentrations of the NO donor NOR-3 and DETA-NO, whereas in our study, the NO concentration was maintained in the low micromolar range. Furthermore, Ciorba et al. (11) studied Drosophila ShB/C recombinant channels (corresponding to Kv1.1 according to the new nomenclature), whereas Kv1.4 channels are expressed in GH₃ cells (50).

Although NO had no effect on I_A, it markedly inhibited I_DR and I_ERG. This finding was suggested by the experimental evidence that L-arginine induced a decrease in the amplitude of both these currents and that this drug was ineffective when the cells were preincubated with the specific NOS-inhibitor L-NAME. In addition, I_DR and I_ERG were also inhibited by NOC-18 and SNAP, two chemically unrelated NO donors.

As Charles et al. (8) pointed out, I_DR in GH₃ cells are carried by two different classes of K⁺ channels, I_DR and BK channels, both of which are sensitive to NO (1). These two types of K⁺ channels have been observed in GH₃ cells using electrophysiology (8, 22, 25, 37) and RT-PCR (50, 51). However, at least under our recording conditions, the contribution of BK channels to I_DR seems to be negligible, because CTX and PAX, two specific blockers of these channels, did not reduce the amplitude of these currents and failed to induce [Ca²⁺]_i oscillations in quiescent GH₃ cells (data not shown). Therefore, in our system, the effect of NOC-18 on outward K⁺ currents can be ascribed entirely to the inhibition of I_DR, which in GH₃ cells is carried by several channel subtypes, some of which are TEA sensitive, such as Kv1.4 and Kv2.1, and others that are TEA resistant, such as Kv1.2 and Kv1.5 (50). Given that NOC-18 was ineffective on TEA-resistant currents, we can conclude that its effect was exerted entirely on TEA-sensitive I_DR.

The observed reduction in I_DR induced by NO could provide a reasonable explanation for the NO effect on [Ca²⁺]_i oscillations. In fact, as shown by Sand et al. (32) and Charles et al. (8), the pharmacological blockade of I_DR is able to induce [Ca²⁺]_i oscillations in quiescent GH₃ cells by causing membrane depolarization. In fact, TEA evokes a dramatic increase in the amplitude of [Ca²⁺]_i oscillations; this effect is likely caused by prolongation of the action potential and a subsequent increase in the action potential-induced influx of Ca²⁺ (8).

An additional explanation for the ability of NO to induce [Ca²⁺]_i oscillations derives from the observation that NO induced a significant decrease in I_ERG. Previous studies firmly established that the inhibition of these channels leads to the appearance of [Ca²⁺]_i oscillations in GH₃ cells (35). This finding has been demonstrated, for example, in the case of TRH (6)- and second-generation antihistamine (35)-induced [Ca²⁺]_i oscillations. Interestingly, the finding that exogenous NO does reduce ERG channel activity is in agreement with our observations while testing the effect of NO donors on ERG channels heterologously expressed in Xenopous oocytes (29, 42).

In conclusion, our experiments involving Kv channels suggest that NOC-18 can promote membrane depolarization by blocking I_ERG and can delay membrane repolarization by inhibiting I_DR. These considerations provide a plausible mechanism for its ability to promote [Ca²⁺]_i oscillations because the changes in membrane potential determined by the effect of NO on Kv channels are expected to promote the opening of VDCCs, which represent the main Ca²⁺ influx mechanism involved in the genesis of the oscillatory process. The role of VDCCs as the immediate effectors in the promotion of NO is further strengthened by the results of the experiments that we performed using Ca²⁺-free media or adding the L-type Ca²⁺ channel blocker nimodipine to the extracellular solution. These experimental maneuvers both determined the loss of NO's ability to induce [Ca²⁺]_i oscillations, implying that Ca²⁺ influx from the extracellular milieu is required, and that this influx actually takes place through L-type VDCC. We can reasonably exclude the idea that NO acted directly on this class of ion channels because, we did not observe NOC-18 to affect the amplitude of inward Ba²⁺ currents elicited by depolarizing ramp protocols. These results are in accord with the evidence reported by Andric et al. (2), who showed that NO does not modify the activity of L-type Ca²⁺ channels in primary cultures of somatomammotrophs. The results of the experiments that we performed using selective blockers of either the ryanodine (ryanodine) or IP₃ stores (xestospongin C) suggest that while L-type Ca²⁺ channel activation is essential as a trigger for NO-induced [Ca²⁺]_i oscillations, the maintenance of the oscillatory process over time also requires Ca²⁺ discharge from the intracellular Ca²⁺ stores. NO-induced [Ca²⁺]_i oscillations prematurely disappeared in cells treated with ryanodine or with xestospongin C. These results are in line with current ideas regarding the genesis of [Ca²⁺]_i oscillations in pituitary cells under the assumption that CICR takes part in the oscillatory phenomenon because the intracellular Ca²⁺ stores are recruited as a consequence of the increase in [Ca²⁺]_i as determined by the activation of the plasma membrane oscillator (39).

In conclusion, endogenous NO generates spontaneous [Ca²⁺]_i oscillations in GH₃ cells by virtue of its ability to inhibit the activity of classic delayed rectifiers, a class of channels that halts membrane depolarization, and that of ERG K⁺ channels, which control membrane resting potential.

	APPENDIX

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION APPENDIX GRANTS REFERENCES

 
[Ca²⁺]_i oscillations were identified and their parameters (i.e., frequency and amplitude) were determined using an algorithm software written by Dr. Paolo Fiore according to the following criteria:

Cutoff.

The cutoff value m was defined as the mean of basal levels of [Ca²⁺]_i ± 2 SD.

[Ca²⁺]_i oscillation.

Considering a bidimensional space in which the x-axis represents time and the y-axis represents [Ca²⁺]_i, a sequence S of bidimensional points (t₁,c₁),(t₂,c₂)...(t_n,c_n) ordered by increasing time values is defined as a single oscillation (using the line equation c = m to define the cutoff), provided that it can be divided into three subsets of points , , and having no point in common and each containing at least one point of S and for which the following conditions are verified:

Cases A, B, and C represented above satisfy the conditions described in the previous definitions, and represent examples of different kinds of [Ca²⁺]_i oscillations (one straight peak, one bicuspidate peak, and one large peak).

Oscillation peak.

The oscillation peak () was defined as the point of S with the maximum value of [Ca²⁺]_i.

Oscillation amplitude.

The oscillation amplitude was defined as the difference between the oscillation peak and m.

Mean oscillation amplitude.

Mean oscillation amplitude was calculated dividing the sum of the amplitudes of each oscillation by the number of oscillations.

Oscillatory frequency:

The oscillatory frequency was calculated as the number of oscillations divided by the time period of observation.

Algorithm.

Step 1: The software calculates, for each cell, mean [Ca²⁺]_i ± SD during the baseline recording interval before any drug addition.

Step 2: The software calculates, for each single cell, the cutoff.

Step 3: The software examines, for each cell, the time period after drug addition and identifies [Ca²⁺]_i oscillation as follows:

Step 3.1: The interval to be examined is set by choosing the pair of extreme points [(t_s,c_s),(t_e,c_e)], where t_s stands for t-start, t_e for t-end, c_s for c-start, and c_e for c-end.

Step 3.2: Starting from the point (t_s,c_s), the software looks for the first sequence of points ([t_s,c_s),(t_z,c_z)] that satisfies the criteria to identify a [Ca²⁺]_i oscillation and chooses the one where t_z has the smallest value satisfying the conditions t_s < t_z t_e.

Step 3.3:

Step 3.3.1: If t_z exists, go to Step 3.3.1. If it does not exist, go to Step 3.3.2. The sequence of points [(t_s,c_s), (t_z,c_z)] is identified as a [Ca²⁺]_i oscillation. The maximal value of [Ca²⁺]_i of the interval [(t_s,c_s),(t_z,c_z)] is defined as the oscillatory peak.

Step 3.3.2: The interval [(t_s,c_s),(t_e,c_e)] will be considered as [Ca²⁺]_i oscillation if it satisfies the criteria for [Ca²⁺]_i oscillation only in the case in which this interval occurs at the end of the acquisition period when cannot be defined. The maximal value of [Ca²⁺]_i of set [(t_s,c_s),(t_z,c_z)] is the oscillatory peak. Step 3 ends.

Step 3.4: If t_z < t_e, the software keeps searching for a [Ca²⁺]_i oscillation by reducing the interval to the [(t_z,c_z),(t_e,c_e)] interval and restarts from Step 3.2.

Step 3.5: If t_z = t_e, Step 3 ends.

Step 4: For each cell, the software examines the interval before drug addition in a manner similar to Step 3.

Step 5: For each cell, the software counts the number of [Ca²⁺]_i oscillations after and before drug addition.

Step 6: For each cell, the software calculates oscillatory frequency after and before drug addition.

Step 7. For each cell, the software calculates the amplitude of each [Ca²⁺]_i oscillation.

Step 8. For each cell, the software calculates the mean oscillation amplitude.

	GRANTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION APPENDIX GRANTS REFERENCES

 
The present study was supported by grants from the Italian Ministry of Health, Programma Speciale art. 12bis comma 6, D. Lgs. 229/99; Special Project "Alzheimer 2001/2004," COFIN-MIUR 2002, COFIN 2004, PNR-FIRB RBNE01E7YX_007 2001, Regione Campania GEAR, and Ricerca Finalizzata Ministero della Salute legge 502/92 "Geni Vulnerabiltà e di Riparazione DNA" (all to L. Annunziato), and by POP and Legge 41 form Regione Campania, Italian Ministry of Health, Programma Speciale art. 12bis comma 6, D Lgs. 229/99.

	ACKNOWLEDGMENTS

 
We thank Vincenzo Grillo for technical support, Dr. Ilaria Staiano for help with cell cultures, and Dr. Paola Merolla for editorial revision. We are also indebted to Dr. Paolo Fiore for writing the software used for [Ca²⁺]_i oscillation detection and to Dr. Luigia Santella (Laboratory of Cell Signaling, Stazione Zoologica A. Dohrn, Naples, Italy) for the generous gift of xestospongin C.

	FOOTNOTES

 

Address for reprint requests and other correspondence: L. Annunziato, Division of Pharmacology, Dept. of Neuroscience, School of Medicine, Federico II Univ. of Naples, via Sergio Pansini 5, 80131 Naples, Italy (e-mail: lannunzi{at}unina.it)

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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