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Originally published In Press as doi:10.1074/jbc.M406450200 on August 2, 2004

J. Biol. Chem., Vol. 279, Issue 40, 41642-41649, October 1, 2004
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Role of Regulator of G Protein Signaling 2 (RGS2) in Ca2+ Oscillations and Adaptation of Ca2+ Signaling to Reduce Excitability of RGS2–/– Cells*

Xinhua Wang{ddagger}, Guojin Huang{ddagger}, Xiang Luo{ddagger}, Josef M. Penninger§, and Shmuel Muallem{ddagger}

From the {ddagger}Department of Physiology University of Texas Southwestern Medical Center at Dallas, Dallas, Texas 75390-9040 and the §Institute of Molecular Biotechnology (IMBA) of the Austrian Academy of Sciences, Dr. Bohrgasse 3-5, Vienna A-1030, Austria

Received for publication, June 9, 2004 , and in revised form, July 29, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Regulators of G protein signaling (RGS) proteins accelerate the GTPase activity of G{alpha} subunits to determine the duration of the stimulated state and control G protein-coupled receptor-mediated cell signaling. RGS2 is an RGS protein that shows preference toward G{alpha}q .To better understand the role of RGS2 in Ca2+ signaling and Ca2+ oscillations, we characterized Ca2+ signaling in cells derived from RGS2–/– mice. Deletion of RGS2 modified the kinetic of inositol 1,4,5-trisphosphate (IP3) production without affecting the peak level of IP3, but rather increased the steady-state level of IP3 at all agonist concentrations. The increased steady-state level of IP3 led to an increased frequency of [Ca2+]i oscillations. The cells were adapted to deletion of RGS2 by reducing Ca2+ signaling excitability. Reduced excitability was achieved by adaptation of all transporters to reduce Ca2+ influx into the cytosol. Thus, IP3 receptor 1 was down-regulated and IP3 receptor 3 was up-regulated in RGS2–/– cells to reduce the sensitivity for IP3 to release Ca2+ from the endoplasmic reticulum to the cytosol. Sarco/endoplasmic reticulum Ca2+ ATPase 2b was up-regulated to more rapidly remove Ca2+ from the cytosol of RGS2–/– cells. Agonist-stimulated Ca2+ influx was reduced, and Ca2+ efflux by plasma membrane Ca2+ was up-regulated in RGS2–/– cells. The result of these adaptive mechanisms was the reduced excitability of Ca2+ signaling, as reflected by the markedly reduced response of RGS2–/– cells to changes in the endoplasmic reticulum Ca2+ load and to an increase in extracellular Ca2+. These findings highlight the central role of RGS proteins in [Ca2+]i oscillations and reveal a prominent plasticity and adaptability of the Ca2+ signaling apparatus.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
G protein-coupled receptor (GPCR)1-evoked Ca2+ signaling is initiated by biochemical reactions at the receptor complex that generate Ca2+-releasing second messengers, which lead to Ca2+ fluxes into and out of the cytosol (1, 2). The biochemical complex is composed of a receptor, the heterotrimeric G protein Gq (and in some cases Gi), and the effector PLC{beta}. Ligand binding to GPCRs results in activation of G{alpha}q that, in turn, activates PLC{beta}. PLC{beta} hydrolyzes phosphatidylinositol bisphosphate to generate IP3. IP3 releases Ca2+ from the ER, which is followed by the activation of store-operated Ca2+ channels in the plasma membrane and Ca2+ influx. The increase in Ca2+ leads to activation of the plasma membrane Ca2+ ATPase (PMCA) and sarco/endoplasmic reticulum Ca2+ ATPase (SERCA) pumps to remove Ca2+ from the cytosol. The overall signal is a transient change in [Ca2+]i. In the case of an intense stimulation only a single Ca2+ transient is observed, whereas at weak physiological stimulus intensity the transient is repeated to generate [Ca2+]i oscillations (1, 2).

Because [Ca2+]i oscillations control virtually all cell functions from cell birth to cell death (1–3), the mechanisms for the generation of Ca2+ oscillations and the regulation of their amplitude and frequency are of major interest. Early work supported models in which the regulation of the IP3R Ca2+ release channels by cytoplasmic Ca2+ ([Ca2+]i) generates agonist-evoked [Ca2+]i oscillations. These models were based on the findings of the bell-shaped dependence of the IP3Rs on [Ca2+]i (4) and the generation of [Ca2+]i oscillations by nonhydrolyzable IP3 (5). However, the finding that weak agonist stimulation leads to oscillatory changes in IP3 concentration (6, 7) suggested that a primary mechanism of [Ca2+]i oscillations is the cyclical activation of PLC{beta}. Two mechanisms for the regulation of PLC{beta} were suggested; one is the direct regulation of PLC{beta} activity by [Ca2+]i (8), and the second is the regulation of PLC{beta} by regulation of the availability of activated G{alpha}q (9). [Ca2+]i can indeed regulate PLC{beta} activity to influence [Ca2+]i oscillations (10). However, in this model the receptor does not control any aspect of the oscillations after the initial stimulation, whereas it is well documented that stimulus intensity affects the amplitude and, in particular, the frequency of the oscillation (1, 2).

The availability of activated G{alpha}q is regulated by the regulators of G protein signaling (RGS) proteins (11). The off reaction in the G protein cycle is the hydrolysis of GTP and the reassembly of the G{alpha}{beta}{gamma} heterotrimer. This reaction is accelerated by RGS proteins in a manner such that the continuous presence of agonist IP3 level oscillates to drive [Ca2+]i oscillations (9). An open question is which RGS protein dominates the regulation of Gq-coupled receptors in vivo. This is an important question in view of the promiscuity of RGS proteins toward G{alpha} subunits in vitro (11) and the expression of multiple RGS proteins in the same cell (12, 13). It is generally assumed that RGS2 regulates the activity of Gq because in vitro RGS2 shows some preference toward G{alpha}q (11, 14), although other RGS proteins can inhibit G{alpha}q activity as well as or better than RGS2 (11). In addition, the role of RGS2 in Ca2+ signaling and [Ca2+]i oscillations in vivo is not known. The availability of the RGS2–/– mice (15) allows direct examination of the roles of RGS2 in Ca2+ signaling in vivo. The RGS2–/– mice are viable and fertile, although they are immune compromised (15) and develop cardiac hypertrophy (16), probably as a result of hypertension (17). We show here that deletion of RGS2 increases the apparent affinity for agonist-stimulated [Ca2+]i signaling but without changing the dose response for agonist-stimulated peak IP3 production. Rather, deletion of RGS2 changes the kinetic of IP3 production to increase its steady-state level and, consequently, the frequency of [Ca2+]i oscillations.

Ca2+ signaling complexes are plastic, and the activity of their components adjusts to perturbations brought about by deletion or overexpression of their components. For example, overexpression of PMCA results in compensatory overexpression of SERCA pumps (18, 19), and partial deletion of SERCA2 results in up-regulation of PMCA4 (20). Furthermore, critical cellular activities such as Ca2+-triggered exocytosis (20) adapt to the change in the characteristics of the Ca2+ signal. Deletion of an RGS protein that regulates the accessibility of active G{alpha}q is expected to change PLC{beta} activity and IP3 production and is likely to lead to the adaptation of Ca2+ signaling to reduce excitability. The excitability of Ca2+ signaling is determined by the ability of IP3 to release Ca2+ from the ER, the contribution of Ca2+ influx to increase [Ca2+]i and augment Ca2+ release, and the capacity of SERCA and PMCA to remove Ca2+ from the cytosol. Ca2+ influx is likely to have a central role in determining the excitability of Ca2+ signaling complexes because it determines the duration and frequency of [Ca2+]i oscillations (21–23). Indeed, the regulation of many Ca2+-dependent cellular activities correlates with the activity of Ca2+ influx (24).

A detailed analysis of Ca2+ signaling in cells from RGS2–/– mice revealed the adaptation of all components of the Ca2+ signaling complex that leads to reduced excitability of Ca2+ signaling in RGS2–/– cells. These findings demonstrate the central role of RGS proteins in [Ca2+]i oscillations and the remarkable plasticity of Ca2+ signaling complexes.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Materials and Solutions—Fura2/AM was from Teff Laboratories, anti-PMCA 5F10 antibodies were from Sigma, anti-PLC{beta} isoforms were a generous gift from Dr. Paul Sternweis (University of Texas Southwestern Medical Center, Dallas, TX), anti-SJ1 was a generous gift from Dr. Pietro De Camilli (Yale University, New Haven, CT), anti-IP3R1 antibodies were a generous gift from Dr. Greg Mignery (Loyola University Chicago, IL), and anti-SERCA2b antibodies were a generous gift from Dr. Frank Wuytack (Katholieke Universiteit Leuven, Belgium). Anti-IP3R3 antibodies were from Transduction Laboratories, and anti-IP3 3-kinase was from Santa Cruz Biotechnology (Santa Cruz, CA). The standard perfusion solution A contained 140 mM NaCl, 5 mM KCl, 1 mM MgCl2, 1 mM CaCl2, 10 mM HEPES (pH 7.4 with NaOH), and 10 mM glucose. When supplemented with 10 mM pyruvate, 1 mg/ml bovine serum albumin, and 0.02% soybean trypsin inhibitor it was named PSA. The Ca2+-free medium was solution A without CaCl2 and with 0.2 mM EGTA.

Preparation of Pancreatic Acini—Acini were prepared as described previously (25). In brief, WT and RGS2–/– mice were anesthetized and killed by cervical dislocation. The pancreata were removed and digested with collagenase P in PSA solution. The acini were washed and, as needed, loaded with Fura2, suspended in PSA, and kept on ice until use.

Measurement of [Ca2+]i—Acini loaded with Fura2 were plated on polylysine-coated glass coverslips that form the bottom of a perfusion chamber. The acini were perfused with worm (37 °C) solution A, and agonists were delivered with the perfusate. Fura2 fluorescence was recorded using a Photon Technology International image equation and analysis system, and the fluorescence signals were calibrated as detailed previously (25).

Mass Measurement of IP3—Acini in solution A were stimulated with the indicated carbachol concentration for 2–120 s. The reactions were stopped and the proteins precipitated by the addition of perchloric acid and by incubating the acini for at least 20 min on ice. IP3 was extracted with a mixture of 0.2 ml Freon and 0.2 ml of tri-n-octylamine, and IP3 was measured by a standard radioligand assay (26).

Western Blot Analysis—Microsomes were prepared from the brain, pancreas, and submandibular glands of WT and RGS2–/– mice by homogenization in medium composed of 20 mM HEPES, pH7.4, 150 mM NaCl, 20% glycerol, 1.5 mM MgCl2, 0.5% Triton X-100, 1 mM dithiothreitol, and 1 mM phenylmethylsulfonyl fluoride and supplemented with a protease inhibitor mixture (Roche Applied Science). Lysates were separated by SDS/PAGE, and proteins were detected by blotting with the desired antibodies.


   RESULTS AND DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
CONCLUSIONS
 REFERENCES
 
Deletion of RGS2 Increases Dynamic of IP3 Production— Because cells express multiple RGS proteins (12, 13), the most direct way to study the role of a specific RGS protein is deletion of the specific genes in mice. RGS2 is the RGS protein that preferentially activates G{alpha}q (14). Although deletion of RGS2 resulted in several physiological phenotypes (15–17), the only Ca2+ signaling effect reported is an increased responsiveness to the stimulation of P2 receptors (12). To determine the consequences of the deletion of RGS2 on Ca2+ signaling, we began by analyzing the effect of the deletion of RGS2 on the expression and activity of IP3 metabolizing enzymes. Preliminary experiments showed the signal/noise ratio obtained with extracts prepared from the pancreas or salivary glands was not adequate to quantify protein expression. A much more reproducible and quantifiable signal was obtained using brain extracts. Comparable quality of results in brain and secretory cells was obtained only for SERCA2b. Therefore, only data obtained with extracts prepared from brain microsomes are presented.

RGS proteins terminate GPCRs signaling by accelerating the GTPase activity of G{alpha} subunits to inhibit the activation of effector proteins, including PLC{beta}, by G{alpha} (11). The cells may adapt by down-regulation of PLC{beta} expression. This possibility was tested by Western blot analysis of PLC{beta} expression. Fig. 1A shows that the deletion of RGS2 had no discernible effect on the expression of PLC{beta}1, PLC{beta}2, or PLC{beta}3 isoforms. Once generated, IP3 is metabolized by IP3 5-phosphatases that remove the phosphate from the 5 position of many phosphoinositols, including IP3, and IP3 3-kinases that phosphorylate IP3 to 1,3,4,5-tetrakisphosphate (IP4). To date, eight 5-phosphatases that are grouped into three sub-families have been identified (27). We focused on synaptojanin 1 (SJ1), a member of the 5-phosphatase subfamily II. SJ1 is expressed in two major forms, an abundant and ubiquitously expressed SJ170 and a neuronal specific SJ145 (28). Fig. 1B shows that the deletion of RGS2 had no effect on the expression of SJ170. Interestingly, the deletion of RGS2 precipitously reduced the expression of SJ145. The significance of this finding to Ca2+ signaling in neurons is not known at present. However, because SJ1 can act on phosphatidylinositol 4,5-bisphosphate to regulate clathrin-mediated endocytosis of synaptic vesicles (29), it should be of interest to look at this activity and its behavioral consequences in the RGS2–/– mice. IP3 3-kinases exist as three isoforms, IP3KA, B, and C, which show tissue-specific expression (30). The 74-kDa IP3KB is ubiquitous and is associated with cellular membranes, including the plasma membrane (27, 30). Analysis of IP3KB expression showed that deletion of RGS2 had no effect on the expression of this enzyme (Fig. 1C).



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  		FIG. 1.
Expression of PLC{beta}, synaptojanin, and IP3KB in WT and RGS2–/– cells. Brain extracts from WT and RGS2–/– mice were used to blot for the expression of the indicated PLC{beta} isoforms (A), synaptojanin 1 (SJ1) (B), and IP3KB (C).

 
The fact that deletion of RGS2 had no effect on PLC{beta}, SJ170, and IP3KB expression raised the possibility that it might affect the dynamic of IP3 production. Measurement of the time course of IP3 production showed that this is indeed the case. Fig. 2A shows that stimulation of WT and RGS2–/– acini with 1 µM carbachol slowly increased IP3 levels that were the same in the two cell types. However, importantly, the steady-state level of IP3 after 60 and 120 s of stimulation was significantly higher in RGS2–/– acini. The effect of RGS2 deletion on the dynamic of IP3 production is further demonstrated in Fig. 2B, where the acini were stimulated with 1 mM carbachol. At this concentration IP3 production was transient, reaching a maximum level after 2–5 s of stimulation and then steadily reducing to stabilize at a new steady-state level. At a high agonist concentration it was possible to demonstrate that the deletion of RGS2 increased the initial rate of IP3 production and, again, the steady-state levels of IP3 at the longer stimulation periods. Experiments similar to those depicted in Fig. 2, A and B were used to determine the dose response for the peak and steady-state levels of IP3 production. Fig. 2C shows that deletion of RGS2 had no effect on the peak level attained at all carbachol concentrations. On the other hand, deletion of RGS2 increased the steady-state level of IP3 at all carbachol concentrations (Fig. 2D).



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  		FIG. 2.
Kinetic of IP3 production in WT and RGS2–/– cells. A and B, pancreatic acini from WT and RGS2–/– mice were stimulated with 1 µM (A) or 1 mM (B) carbachol for the indicated times between 2 and 120 s, and the level of IP3 was determined. The single asterisk (*) indicates being statistically different from WT at p < 0.05, and the double asterisks (**) indicate being statistically different from WT at p < 0.01. C, pancreatic acini from WT ({blacksquare}) and RGS2–/– cells ({circ}) were stimulated for either 5 or 10 s with the indicated concentrations of carbachol, and IP3 levels were plotted as a function of carbachol concentration. D, pancreatic acini from WT ({blacksquare}) and RGS2–/– cells ({circ}) were stimulated for 120 s with the indicated concentrations of carbachol, and IP3 levels were plotted as a function of carbachol concentration.

 
The findings that the deletion of RGS2 increased the initial rate (Fig. 2B) and steady-state level (Fig. 2D) with no effect on the extent of IP3 production (Fig. 2C) have several implications. The increased rate of IP3 production indicates that in resting cells RGS2 exerts tonic inhibition on GPCR signaling. This conclusion is in line with previous observations demonstrating the constitutive inhibitory activity of recombinant RGS proteins when applied in vivo and the activation of Ca2+ signaling by scavenging RGS protein antibodies (9). The increased steady-state level of IP3 can result from the increased rate of IP3 production and the reduced rate of signal termination. However, it is significant that a more prominent increased steady state is observed at low agonist concentration (Fig. 2D) at which the IP3 level oscillates (6, 7). As was shown for [Ca2+]i oscillations (31), the increased frequency of IP3 oscillations will be translated to an increased steady-state when IP3 levels are measured in a cell population. Therefore, it is possible that the increased steady-state level of IP3 at low agonist concentrations observed in RGS2–/– cells may reflect the increased frequency of IP3 oscillations. The effect of RGS2 on steady-state levels of IP3 highlights the importance of RGS proteins in controlling IP3 production and [Ca2+]i signaling in vivo (9).

Deletion of RGS2 Increases Stimulus Intensity—An increased frequency of IP3 oscillations at a low agonist concentration predicts an increased frequency of [Ca2+]i oscillations, and increased steady-state IP3 levels at all agonist concentrations predict an increased apparent affinity for agonist-stimulated Ca2+ signaling. These predictions were tested by measuring [Ca2+]i in WT and RGS2–/– cells stimulated with carbachol and CCK that activate the Gq-coupled M3 and CCK receptors, respectively. Figs. 3, A and B show that the low concentrations of 0.05 and 0.1 µM carbachol had no effect on Ca2+ signaling in WT cells but triggered [Ca2+]i oscillations with increased frequency in RGS2–/– cells. Similarly, Fig. 3, D and E show that 0.5 and 2.5 pM CCK had no effect or induced low frequency [Ca2+]i oscillations in WT cells and higher frequency [Ca2+]i oscillations in RGS2–/– cells. Carbachol at 1 µM and CCK at 10 pM triggered [Ca2+]i oscillations in WT cells but a large Ca2+ transient in RGS2–/– cells. To determine the dose response for the agonists, at the end of each stimulation period the cells were exposed to a maximal agonist concentration to discharge the residual agonist-mobilizable Ca2+ pool and calculate the extent of Ca2+ mobilization at the lower agonist concentration. The results of these measurements are shown in Fig. 3, C and F, and indicate that the deletion of RGS2 decreased the EC50 for carbachol from 4.3 ± 0.5 to 0.48 ± 0.07 µM and the EC50 for CCK from 53 ± 5 to 7.3 ± 2.6 pM. Similar measurements with epinephrine stimulation of parotid duct cells revealed that the deletion of RGS2 also decreased the EC50 for epinephrine in parotid duct cells (not shown). These findings satisfy the two predictions that resulted from the change in the kinetic of agonist-stimulated IP3 production



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  		FIG. 3.
Dose response for an agonist-mediated [Ca2+]i increase in WT and RGS2–/– cells. Pancreatic acini from WT (A and D) and RGS2–/– mice (B and E) loaded with Fura2 were used to measure [Ca2+]i with different carbachol (A–C) or CCK (D–F) concentrations. At the end of each stimulation period with the various agonist concentrations, the acini were challenged with a maximal concentration of carbachol or CCK. The extent of Ca2+ mobilization was calculated from the residual Ca2+ mobilized by the maximal agonist concentration and plotted as a function of carbachol (C) or CCK (F) concentrations.

 
The results in Figs. 1, 2, 3 indicate that in vivo RGS2 is a key regulator of Gq-dependent Ca2+ signaling. The combined effects of RGS2 deletion on the kinetic of IP3 production and [Ca2+]i oscillations at a low agonist concentration provide the first direct evidence for the importance of RGS2 in controlling IP3 and, therefore, [Ca2+]i oscillations in vivo and further support the conclusion that RGS proteins have a primary role in the generation of agonist-evoked [Ca2+]i oscillations (9). Another important observation is that the deletion of RGS2 affected Ca2+ signaling by all GPCRs examined and in both pancreatic acinar cells and parotid gland duct cells, including the M3, CCK, bombesin, and {alpha}-adrenergic receptors. In vivo RGS proteins display receptor specificity (32), with the receptor recognition domain residing at the N terminus of RGS proteins (33). Interestingly, of all RGS proteins examined, only RGS2 showed similar potency for all receptors (32). This result is consistent with the present finding that Ca2+ signaling was similarly affected by all of the Gq-coupled receptors examined. Evidently, if RGS proteins control Ca2+ signaling in a receptor-specific manner, other RGS proteins that participate in Ca2+ signaling will fulfill this function. Indeed, multiple RGS proteins are found in a single cell (12), including in a single pancreatic acinar cell (13), and the results in Fig. 4 provide direct evidence that RGS proteins other than RGS2 can regulate Ca2+ signaling in pancreatic acini.



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  		FIG. 4.
Rapid termination of the stimulated state in WT and RGS2–/– cells. WT (A) and RGS2–/– acini (B) were stimulated with 1 mM carbachol (Carb) for the indicated duration, and at the [Ca2+]i zenith stimulation was terminated with 10 µM atropine. In panel C the reduction in [Ca2+]i from the time of atropine addition in the two cell types was rescaled and presented an expanded time scale. The averaged times to a 50% reduction in [Ca2+]i after rescaling is given in the columns in seconds (Sec).

 
Termination of Cell Stimulation in RGS2–/– Cells—An effect of RGS2 deletion on the kinetic of IP3 production and Ca2+ signaling raises the question of how the termination of cell stimulation was affected in RGS2–/– cells and whether other RGS proteins participate in Ca2+ signaling. The finding that IP3 levels in RGS2–/– cells remained transient rather than showing continuous accumulation even at low agonist concentration and that maximal levels of IP3 were the same in WT and RGS2–/– cells indicate that not all negative controls of GPCR-dependent Ca2+ signaling were eliminated in RGS2–/– cells. This indication is further demonstrated in Fig. 4, in which the effect of RGS2 deletion on the termination of cell stimulation was examined. In these experiments the cells were stimulated for 3–5 s with 1 mM carbachol to completely discharge the ER Ca2+ pool, and stimulation was rapidly and completely terminated by washing away the carbachol and adding the antagonist atropine. We showed previously that this leads to rapid termination of cell stimulation, hydrolysis of IP3, and re-uptake of Ca2+ into the ER to reduce [Ca2+]i bake to basal levels (34, 35). Although the rate of [Ca2+]i reduction after the addition of atropine mostly reflects the rate on Ca2+ uptake into the ER, it can be used to determine whether the stimulated state is actively terminated by a negative regulatory mechanism such as that exerted by RGS proteins. Fig. 4 shows that the addition of atropine at the [Ca2+]i zenith increased the rate of [Ca2+]i reduction in both WT and RGS2–/– cells, but the rate in RGS2–/– cells was 1.34 ± 0.05-fold (n = 7) slower that in WT cells. The fast rate of [Ca2+]i reduction in WT and RGS2–/– cells leads to two important conclusions. First, as expected, RGS2 does regulate the rate of signal termination of Gq-mediated Ca2+ signaling in vivo. Second, other RGS proteins must participate in the termination of Gq-mediated Ca2+ signaling, because atropine did rapidly terminate Ca2+ signaling in RGS2–/– cells. In fact, the small increase in the rate of [Ca2+]i reduction in the RGS2–/– cells suggest that other RGS proteins have a major role in terminating Gq-mediated Ca2+ signaling. Although at present we cannot exclude the possibility that other RGS proteins were recruited to the Ca2+ signaling complex as a result of the deletion of RGS2, our findings do indicate that RGS proteins other than RGS2 can communicate with Gq in the Ca2+ signaling complex to efficiently terminate Ca2+ signaling. These can be any of the RGS proteins found in single pancreatic acinar cells (13) that stimulate the GTPase activity of G{alpha}q (11).

Adaptation of ER Ca2+ Uptake and Ca2+ Release in RGS2–/– Cells—Resting [Ca2+]i levels were the same in WT (58 ± 5) and RGS2–/– (55 ± 4) cells. By contrast, Fig. 5, a and d show that maximal stimulation of WT and RGS2–/– cells incubated in Ca2+-containing medium increased [Ca2+]i to 465 ± 50 and 630 ± 65 nM (n = 20), respectively. A higher [Ca2+]i increase in RGS2–/– cells can be due to increased Ca2+ release from the ER, increased Ca2+ entry, or both. To distinguish the contribution of each pathway, the cells were stimulated in Ca2+-free medium. The results in Fig. 5, b and e indicate that the higher [Ca2+]i increase in RGS2–/– cells was due to higher Ca2+ release from the ER, because in Ca2+-free medium carbachol increased [Ca2+]i to 450 ± 40 and 605 ± 55 nM (n = 5) in WT and RGS2–/– cells, respectively. Furthermore, the addition of external Ca2+ to stimulated cells incubated in Ca2+-free medium resulted in a lower increase in [Ca2+]i in RGS2–/– than in WT cells. Higher Ca2+ release requires higher Ca2+ content in the ER of RGS2–/– cells. The Western blot analysis in Fig. 5g shows that the deletion of RGS2 resulted in a compensatory increase in SERCA2b expression by 1.87 ± 0.13-fold.



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  		FIG. 5.
Reduced Ca2+ influx and excitability of RGS2–/– cells. As indicated by the bars, WT (a–c) and RGS2–/– cells (d–f) were stimulated with 1 mM carbachol, inhibited with 10 µM atropine (Atro), and restimulated with 10 nM CCK. In panels a and d the acini were incubated in media containing 5, 0, and 5 mM Ca2+ (b, c, e, and f) as indicated. In panels c and f the experiments were done with 25 µM CPA. In panels g and h the expression of SERCA2b was analyzed by Western blot.

 
In agreement with the results in Fig. 4, the addition of atropine rapidly reduced [Ca2+]i to the basal level in WT and RGS2–/– cells. Another consistent difference between the two cell types was that after the addition of atropine, [Ca2+]i oscillated for at least 5 min in WT but not in RGS2–/– cells (Fig. 5, a, b, d, and e), and the oscillations required incubating the cells with 5 mM external Ca2+ during carbachol stimulation and the incubation with atropine (not shown) and active SERCA pumps (Fig. 5, c and f). The requirement for high external Ca2+ during cell stimulation and active SERCA pumps to observe the oscillations suggest that overloading the ER with Ca2+ may have sensitized the Ca2+ release process. The lack of [Ca2+]i oscillations in RGS2–/– cells under the same conditions, despite the SERCA pump overexpression and ER Ca2+ overload, suggests that the Ca2+ release mechanism in RGS2–/– cells was modified to reduce their excitability. This possibility was tested directly by measuring the potency of IP3 to release Ca2+ from the ER of WT and RGS2–/– cells.

Adaptation of IP3-mediated Ca2+ Release in RGS2–/– Cells—Fig. 6 shows an analysis of expression of IP3Rs and IP3-mediated Ca2+ release in WT and RGS2–/– cells. The deletion of RGS2 down-regulated expression of IP3R1 by ~25%, whereas it up-regulated the expression of IP3R3 by ~30% (Fig. 6, A and B). This resulted in a reduced potency for IP3 to trigger Ca2+ release from ~0.23 to 0.48 µM (Figs. 6, C–E). These findings are consistent with the modified properties of Ca2+ release and may explain the reduced excitability of Ca2+ release in RGS2–/– cells. Thus, it seems that the entire ER Ca2+ homeostasis was modified in RGS2–/– cells. The adaptation of IP3-mediated Ca2+ release will function to reduce Ca2+ release by the elevated steady-state levels of IP3 in the RGS2–/– cells (Figs. 1 and 2), and the up-regulation of SERCA2b will speed up removal of [Ca2+]i between the [Ca2+]i spikes in RGS2–/– cells.



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  		FIG. 6.
IP3-mediated Ca2+ release in WT and RGS2–/– cells. The expression of IP3R1 and IP3R3 was analyzed by Western blot (A and B). IP3-mediated Ca2+ release was measured in WT (C) or RGS2–/– (D) pancreatic acini permeabilized with streptolysin-O. After the cells reduced Ca2+ to ~45 nM, Ca2+ release was initiated by successive additions of 0.1 µM IP3 (small arrows), and full store discharge was attained by a final addition of 4 µM IP3 (large arrow). The results are summarized in panel E for WT ({blacksquare}) and RGS2–/– cells ({circ}).

 
Adaptation of Ca2+ Influx in RGS2–/– Cells—Another aspect of cellular Ca2+ homeostasis is Ca2+ influx and efflux across the plasma membrane. The standard Ca2+ removal and re-addition protocol was used to assay for Ca2+ influx and showed a lower response in RGS2–/– cells (Fig. 5, a, b, d, and e). However, the reduced response can be due to the higher SERCA2b pump expression in RGS2–/– cells that can rapidly remove the Ca2+ entering the cells and establish a lower [Ca2+]i plateau. In addition, increased PMCA activity in RGS2–/– cells can contribute to the lower apparent [Ca2+]i increase due to the addition of external Ca2+. In the protocol in Fig. 5, c and f, we isolated the Ca2+ influx activity. The acini were incubated in Ca2+-free medium, stimulated with 1 mM carbachol to maximally deplete the stores, and incubated with the SERCA pump inhibitor cyclopiazonic acid (CPA) (25 µM). This concentration of CPA was maintained thereafter. After the return of [Ca2+]i to the basal level the cells were incubated with atropine to terminate the stimulated state and minimize the contribution of PMCA. The cells were then exposed to 5 mM Ca2+, which increased [Ca2+]i to 275 ± 30 nM in WT cells and to 185 ± 20 nM (n = 7) in RGS2–/– cells. These results clearly indicate that another adaptation of Ca2+ signaling in RGS2–/– cells is the down-regulation of agonist-activated Ca2+ influx.

Adaptation of PMCA Activity in RGS2–/– Cells—Next, we tested whether PMCA protein and activity underwent adaptation in RGS2–/– cells. The Western blot analysis in Fig. 7A shows that PMCA protein expression was unaltered in RGS2–/– cells. Two protocols were used to assay PMCA activity. The first is based on the measurement of [Ca2+]i in single cells or acini and is shown in Fig. 7B. We reasoned that the activation of PMCA limits the increase of [Ca2+]i in response to the addition of external Ca2+ to the agonist-stimulated cell and that PMCA activity can be recovered by the termination of cell stimulation with an antagonist. PMCA activity can then be read from the rate and extent of the [Ca2+]i increase induced by the antagonist. As an additional control, the cells can be re-stimulated with a second agonist that should result in the reactivation of PMCA, Ca2+ efflux, and a reduction in [Ca2+]i. The results of this protocol with WT and RGS2–/– cells are shown in Fig. 7B. Cells incubated in Ca2+-free medium were stimulated with carbachol, and SERCA pumps were inhibited with CPA. While stimulated, the cells were exposed to 5 mM Ca2+, and after the stabilization of [Ca2+]i, stimulation was terminated by incubating the cells with atropine. The inhibition of SERCA pumps ensured that the Ca2+ increase triggered by atropine is due to the unstimulation of PMCA. Unstimulation of PMCA resulted in a larger increase in [Ca2+]i in RGS2–/– cells. In WT cells the addition of atropine increased [Ca2+]i by 52 ± 8 nM and in RGS2–/– cells by 96 ± 11 nM (n = 12). To further demonstrate that the atropine-induced increase in [Ca2+]i is due to the stimulation of PMCA, the cells were re-stimulated with CCK, which reduced [Ca2+]i back to the level set by carbachol stimulation.



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  		FIG. 7.
PMCA activity in WT and RGS2–/– cells. The expression of PMCA was analyzed by Western blot (a). To estimate PMCA activity in single cells and acini (b), the cells were incubated in Ca2+-free medium and treated with 1 mM carbachol (Carb) and 25 µM CPA. The cells were then exposed to media containing 5 mM Ca2+ and, after stabilization of [Ca2+]i, PMCA activity was uncovered by inhibiting the stimulated state with atropine. The cells were then stimulated with CCK to reactivate PMCA. The average PMCA activity was measured in cell suspension in WT and RGS2–/– cells (c), and the columns show the average of three experiments.

 
The second assay of PMCA activity is based on the measurement of the extracellular Ca2+ of a large number of cells or acini suspended in lightly buffered Ca2+-free medium, as detailed previously (31, 36). The stimulation of pancreatic acini with carbachol resulted in Ca2+ efflux, which was monitored as an increase in Fura2 fluorescence in the extracellular medium. Fig. 7C shows that after an initial delay, extracellular Ca2+ increased at a rate of 33 ± 4 nM/min in WT acini and 48 ± 5 nM/min in RGS2–/– acini, confirming the results obtained in acini in Fig. 7B. The results in Fig. 7 used an established protocol and a new protocol to confirm (36) the activation of PMCA by GPCRs and to show the adaptation of PMCA activity in RGS2–/– cells. Hence, as was found for the ER membrane, Ca2+ homeostasis across the plasma membrane was adapted to account for the deletion of RGS2–/–.

Reduced Excitability in RGS2–/– Cells—To examine the overall effect of adaptation of the ER and plasma membrane Ca2+ homeostasis on Ca2+ signaling under physiological conditions, we tested the response of agonist-evoked [Ca2+]i oscillations to increased extracellular Ca2+ in WT and RGS2–/– cells. Extracellular Ca2+ is essential, as in the case of CCK, or obligatory, as in the case of carbachol-evoked [Ca2+]i oscillations (2). Fig. 8A shows that in WT pancreatic acini stimulated with 0.25 µM carbachol, increasing external Ca2+ from 1 to 3 to 7.5 mM increased the frequency of [Ca2+]i oscillations by ~1.8 ± 0.2 and 2.7 ± 0.3-fold, respectively (Fig. 8C). The [Ca2+]i oscillations evoked by 0.25 µM carbachol in RGS2–/– cells incubated with 1 mM CaCl2 occurred at a frequency 1.9 ± 0.03-fold higher than that measured in WT cells. However, in contrast to the findings in WT cells, increasing external Ca2+ from 1 to 3 mM had no effect on the [Ca2+]i oscillations in RGS2–/– cells, and further increasing external Ca2+ to 7.5 mM increased the frequency of the oscillations by only 1.3 ± 0.2-fold.



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  		FIG. 8.
Reduced excitability in RGS2–/– cells. Pancreatic acini from WT (A) and RGS2–/– cells (B) in medium containing 1 mM Ca2+ were stimulated with 0.25 µM carbachol (Carb). As indicated by the bars, the cells were then exposed to media containing 3, 7.5, and 0 mM. The frequencies of [Ca2+]i oscillations at each external Ca2+ concentration are summarized in panel C.

 

   CONCLUSIONS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
REFERENCES
 
The present findings reveal a central role for RGS proteins in general and RGS2 in particular in the regulation of IP3 production and [Ca2+]i oscillations. The frequency and amplitude of [Ca2+]i oscillations are regulated by all components of the Ca2+ signal, including IP3 production and Ca2+ fluxes across the ER and plasma membranes (2). However, the finding of agonist-stimulated oscillations in IP3 concentration (6, 7) suggests that such oscillation are the primary generator of [Ca2+]i oscillations. This allows control of [Ca2+]i oscillation frequency and amplitude by the agonist, as observed in many cells (1, 2). As the negative regulators of GPCRs signaling, RGS proteins play a central role in determining the duration of the stimulated state and, thus, can play a central role in controlling [Ca2+]i oscillations. In a previous work we showed that cyclical activation and inactivation of RGS protein activity generates [Ca2+]i oscillations in pancreatic acini and salivary gland cells (9). The present work extends these findings in three ways. First, this work demonstrates a central role for RGS2 in regulating [Ca2+]i oscillations in vivo. Second, the present findings provides independent evidence for the role of RGS proteins and the kinetic of IP3 production in triggering [Ca2+]i oscillations. Thus, deletion of RGS2 increased the steady-state concentration of IP3 that can result from an increased frequency of oscillations in the concentration of IP3 (Fig. 2) and, consequently, increased the frequency of [Ca2+]i oscillations. Third, persistent rapid termination of Ca2+ signaling by the antagonist atropine indicates that RGS proteins in addition to RGS2 play a critical role in controlling Ca2+ signaling in native cells in vivo.

The results presented here reveal broad plasticity of the Ca2+ signaling complexes, as observed in the overall adaptation of all Ca2+ transporting pathways to reduce the excitability of Ca2+ signaling in RGS2–/– cells. Hence, removal of a central negative regulatory mechanism at the receptor complex resulted in the acceleration of IP3 production and higher steady-state level of IP3 to increase the responsiveness of all the Gq-coupled receptor complexes tested (M3, CCK, bombesin, and {alpha}-adrenergic). In response, the cellular Ca2+ transporting machinery adapted to reduced excitability of Ca2+ signaling. Reduced excitability was accomplished by restricting Ca2+ influx into the cytosol. This was achieved by reduced sensitivity of Ca2+ release to IP3 and the ER Ca2+ load and reduced Ca2+ influx across the plasma membrane to restrict Ca2+ release to the cytosol, and increased SERCA and PMCA activity to rapidly clear Ca2+ from the cytosol. The combined effects is the buffering of changes in [Ca2+]i to prevent a large fluctuation in [Ca2+]i oscillations, as observed in the reduced responsiveness to perturbations such as a large increase in external Ca2+. Considering the central role of [Ca2+]i in regulating its own concentration by regulating the activity of many components of the Ca2+ signal such as PLC{beta}, IP3Rs, store-operated Ca2+ channels, and PMCA activity (2), it seems that the most efficient way to regulate the excitability of Ca2+ signaling is to regulate Ca2+ fluxes to the cytosol. In addition, [Ca2+]i regulates virtually all cellular activities (1–3). Adaptation to deletion of RGS2 by buffering [Ca2+]i and reduced excitability is also expected to guard against hyperactivation of Ca2+-regulated cellular processes.


   FOOTNOTES
 
* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed: The University of Texas Southwestern Medical Center at Dallas, 5323 Harry Hines Blvd., Dallas, TX 75390-9040. Tel.: 214-648-2593; Fax: 214-648-8879; E-mail: SHMUEL.MUALLEM{at}utsouthwestern.edu.

1 The abbreviations used are: GPCR, G protein-coupled receptor; PLC{beta}, phospholipase C {beta}; IP3, inositol 1,4,5-trisphosphate; IP3R, IP3 receptor; IP3KB, IP3 3-kinase isoform B; RGS, regulator of G protein signaling; SJ, synaptojanin; PMCA, plasma membrane Ca2+ ATPase; ER, endoplasmic reticulum; SERCA, sarco/endoplasmic reticulum Ca2+ ATPase pump; CCK, cholecystokinin; CPA, cyclopiazonic acid; WT, wild type. Back



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 

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