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Molecular Endocrinology, doi:10.1210/me.2003-0488
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Molecular Endocrinology 18 (12): 2967-2980
Copyright © 2004 by The Endocrine Society

Down-Regulation of Inositol 1,4,5-Trisphosphate Receptor in Cells Stably Expressing the Constitutively Active Angiotensin II N111G-AT1 Receptor

Mannix Auger-Messier, Guillaume Arguin, Benoit Chaloux, Richard Leduc, Emanuel Escher and Gaetan Guillemette

Department of Pharmacology, Faculty of Medicine, Université de Sherbrooke, Sherbrooke, Quebec, Canada J1H 5N4

Address all correspondence and requests for reprints to: Gaetan Guillemette, Ph.D., Department of Pharmacology, Faculty of Medicine, Université de Sherbrooke, 3001, 12th Avenue North, Sherbrooke, Quebec, Canada J1H 5N4. E-mail: Gaetan.Guillemette{at}USherbrooke.ca.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The diverse cellular changes brought about by the expression of a constitutively active receptor are poorly understood. QBI-human embryonic kidney 293A cells stably expressing the constitutively active N111G-AT1 receptor (N111G cells) showed elevated levels of inositol phosphates and frequent spontaneous intracellular Ca2+ oscillations. Interestingly, Ca2+ transients triggered with maximal doses of angiotensin II were much weaker in N111G cells than in wild-type cells. These blunted responses were observed independently of the presence or absence of extracellular Ca2+ and were also obtained when endogenous muscarinic and purinergic receptors were activated, revealing a heterologous desensitization process. The desensitized component of the Ca2+ signaling cascade was neither the G protein Gq nor phospholipase C. The intracellular Ca2+ store of N111G cells and their mechanism of Ca2+ entry also appeared to be intact. The most striking adaptive response of N111G cells was a down-regulation of their inositol 1,4,5-trisphosphate receptor (IP3R) as revealed by reduced IP3-induced Ca2+ release, lowered [3H]IP3 binding capacity, diminished IP3R immunoreactivity, and accelerated IP3R degradation involving the lysosomal pathway. Treatment with the inverse agonist EXP3174 reversed the desensitized phenotype of N111G cells. Down-regulation of IP3R represents a reversible adaptive response to protect cells against the adverse effects of constitutively active Ca2+-mobilizing receptors.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
THE AT1 RECEPTOR belongs to the G protein-coupled receptor (GPCR) superfamily and plays an active role in the renin-angiotensin system. The AT1 receptor mediates virtually all the known physiological actions of angiotensin II (Ang II), including vascular contraction, aldosterone secretion, sodium and water retention, neuronal activation, and cardiovascular cell growth and proliferation (1, 2). The AT1 receptor functions primarily through its productive coupling to the heterotrimeric guanyl nucleotide-binding regulatory protein (G protein) Gq/11, which activates phospholipase C, which in turn hydrolyzes membranous phosphatidylinositol 4,5-bisphosphate into inositol 1,4,5-trisphosphate (IP3) and diacylglycerol (3, 4). Whereas IP3 causes a rapid release of Ca2+ from intracellular stores upon activation of its receptor-channel (IP3R), diacylglycerol recruits and activates protein kinase C at the plasma membrane. IP3-induced Ca2+ release is generally followed by an increase in Ca2+ entry across the plasma membrane that can serve to replenish stores or contribute to Ca2+-dependent signaling. This entry of Ca2+ occurs through a poorly defined mechanism that is initiated by the depletion of Ca2+ stores, a process known as capacitative Ca2+ entry (5).

A GPCR able to adopt an active conformation in the absence of an agonist is said to be constitutively active. The AT1 receptor belongs to a large group of about 60 wild-type (WT) GPCRs exhibiting constitutive activity (6). The constitutive activity of the AT1 receptor was emphasized after its overexpression in COS-1 cells, which showed enhanced basal activity of phospholipase C (7). Recently, three independent studies simultaneously reported an important increase in the constitutive activity of the AT1 receptor when Asn111 (in the third transmembrane domain) was substituted for the smaller residues Ala or Gly (7, 8, 9). Regardless of the type of G protein they couple with, numerous other examples of constitutively active mutant (CAM) GPCRs have been reported in the literature, including A293E-{alpha}1B adrenergic receptor (10), M257Y-rhodopsin (11), and T279K-µ opioid receptor (12). It is believed that intramolecular interactions preferentially constrain GPCRs in the inactive conformation and that agonists or specific mutations relieve these constraints, thus privileging the active conformation (13). CAM-GPCRs may profoundly modify cell functions. Diseases such as retinitis pigmentosa (14) and Kaposi’s sarcoma (15) have been ascribed to CAM-GPCRs. More studies are needed to determine the effects of CAM-GPCRs on intracellular signaling mechanisms and cell functions.

In recent work, we noticed some refractoriness in the Ca2+ response of cells expressing the constitutively active N111G-AT1 receptor, suggesting that a desensitization process had developed as a consequence of the permanent activity of this receptor (16). In the study presented here, we selected a QBI-human embryonic kidney 293A clonal cell line stably expressing the N111G-AT1 receptor (N111G cells) to examine in greater detail the mechanism of intracellular Ca2+ regulation under basal conditions and after stimulation with different Ca2+ mobilizing agonists. We noted that agonist-induced intracellular Ca2+ release and subsequent Ca2+ entry activities were heterologously desensitized in N111G cells. This refractory state was caused mainly by a down-regulation of IP3R and could be reversed by a prolonged treatment with EXP3174, an inverse agonist of the AT1 receptor.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Pharmacological Properties of the N111G-AT1 Receptor
The constitutively active N111G-AT1 receptor and the AT1 receptor were stably transfected in QBI-HEK 293A cells, and representative clonal cell lines were analyzed for their functional properties (Table 1Go). In saturation binding studies, the AT1 receptor exhibited high- and low-affinity states for the agonist [125I]Ang II (0.5 ± 0.2 and 3.4 ± 0.9 nM, respectively) with expression levels (Bmax) of 0.4 ± 0.1 and 0.8 ± 0.3 pmol/mg of protein, respectively (Table 1Go). The high-affinity state was completely converted to the low-affinity state in the presence of guanyl nucleotides (data not shown). The N111G-AT1 receptor exhibited a single high-affinity state (0.9 ± 0.2 nM) that was not affected by the presence of guanyl nucleotides (data not shown) and its Bmax was 2.4 ± 0.6 pmol/mg of protein. Under basal conditions, N111G cells contained high levels of inositol phosphate (IP) that were at least 6-fold higher than those found in WT cells. These results confirm the constitutive activity of N111G-AT1 receptor. Upon stimulation with a high concentration of Ang II, N111G and WT cells accumulated comparable levels of IP (Table 1Go). These results are similar to those previously obtained after transient transfection of these receptors in COS-7 cells (16).


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  		Table 1. Binding and Functional Properties of Receptors Expressed in Clonal Cell Lines

 
Impaired Ca2+ Response in Single N111G Cells
The temporal patterns of intracellular Ca2+ signals in QBI-HEK 293A cells expressing the constitutively active N111G-AT1 receptor were compared with those elicited by the WT AT1 receptor. Under basal conditions (without agonist), the AT1 receptor did not generally elicit any fluctuations in intracellular Ca2+ concentrations (Fig. 1AGo). Upon stimulation with a relatively low concentration of Ang II (0.1 nM), the AT1 receptor elicited repetitive baseline-separated Ca2+ transients (Ca2+ oscillations) with a frequency of 32 ± 16 oscillations/h. Interestingly, under basal conditions, the N111G-AT1 receptor elicited spontaneous Ca2+ oscillations with a frequency of 18 ± 11 oscillations/h. Upon activation with 0.1 nM Ang II, N111G cells showed only a modest acceleration of the oscillatory rate, which increased by about 13 oscillations/h, barely reaching the oscillatory rate of WT cells stimulated with 0.1 nM Ang II (Fig. 1BGo). These results suggest that the N111G-AT1 receptor is less responsive than the AT1 receptor to a stimulation by Ang II. No significant difference was noted between the amplitude of the oscillations elicited in N111G cells (0.40 ± 0.02 fluorescence ratio unit) and those elicited in WT cells (0.41 ± 0.02 fluorescence ratio unit). Upon activation with a high dose of Ang II (1 µM), the AT1 receptor produced a single large Ca2+ transient (amplitude of 0.87 fluorescence ratio unit) that slowly declined toward a low level slightly above the resting level (Fig. 1CGo), whereas the N111G-AT1 receptor also produced a single Ca2+ transient but with a low amplitude of 0.66 fluorescence ratio unit (Fig. 1DGo). These results indicate that although the constitutively active N111G-AT1 receptor is capable of inducing spontaneous Ca2+ oscillations that can be accelerated upon activation with low doses of agonist, single cells expressing this receptor show some refractoriness in their Ca2+ response.



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  		Fig. 1. Spontaneous and Ang II-Induced Ca2+ Oscillations in Single Cells

Attached WT cells (A and C) or N111G cells (B and D) were loaded with fura 2-AM (0.1 µM) for 20 min at room temperature in HBSS, washed for 20 min at room temperature in HBSS, and mounted onto a videomicroscopy system. Fura 2 fluorescence in single cells was monitored under basal conditions for an initial period of 750 sec before adding either 0.1 nM Ang II (A and B) or 1 µM Ang II (C and D). These typical traces show variations of the fluorescence ratio (F334/F380) obtained at room temperature as described in Materials and Methods. Similar results were obtained with three different cell preparations.

 
Impaired Ca2+ Response in N111G Cell Populations
Despite their elevated IP content, the basal Ca2+ concentration in N111G cells (99 ± 13 nM) was not significantly different from that of WT cells (99 ± 22 nM). In N111G cells, however, 1 µM Ang II produced a low-amplitude Ca2+ transient (387 ± 97 nM; Fig. 2BGo) that was significantly lower than that produced in WT cells (790 ± 84 nM; Fig. 2AGo). Agonist-induced Ca2+ transients consist of two main components: the release of Ca2+ from intracellular stores and Ca2+ entry from the extracellular medium. To identify which of these two components could be responsible for the refractory state of N111G cells, we performed experiments in a nominally Ca2+-free medium. Under these conditions, WT cells maintained a stable low level of intracellular Ca2+ for at least 3 min (Fig. 2CGo). When extracellular Ca2+ was added to the medium, a very minor increase in the intracellular Ca2+ concentration was observed. According to the capacitative Ca2+ entry model (5), this minor increase could be due to the entry of Ca2+ resulting from a small leakage of the intracellular Ca2+ store. Interestingly, under the same conditions, N111G cells displayed a larger intracellular Ca2+ increase when extracellular Ca2+ was added (Fig. 2DGo). These results suggest that N111G cells have a dynamic capacitative Ca2+ entry activity under basal conditions. This is consistent with the constitutive activity of the N111G-AT1 receptor, which maintains an elevated level of IP3 in these cells (Table 1Go), thus causing a larger depletion of their intracellular Ca2+ store. In the absence of extracellular Ca2+, a high concentration of Ang II (1 µM) caused a robust intracellular Ca2+ transient (amplitude of 522 ± 66 nM) in WT cells that reflected a major depletion of their IP3-sensitive intracellular Ca2+ store (Fig. 2EGo). Under these conditions, the addition of extracellular Ca2+ caused a significant capacitative Ca2+ entry with a maximal amplitude of 197 ± 25 nM. Figure 2FGo shows that the IP3-induced Ca2+ release (amplitude of 306 ± 33 nM) and the capacitative Ca2+ entry (amplitude of 152 ± 4 nM) activities elicited by 1 µM Ang II in N111G cells were smaller than those observed in WT cells.



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  		Fig. 2. Ang II-Induced Ca2+ Release and Ca2+ Entry

Populations (1.25 x 106 cells per assay) of WT cells (A, C, and E) or N111G cells (B, D, and F) were loaded with fura 2-AM (5 µM) for 20 min at 37 C, washed by centrifugation, and resuspended either in an extracellular-like medium (A and B) or in a nominally Ca2+-free medium (C–F), and their intracellular Ca2+ concentration was monitored upon stimulation with 1 µM Ang II or upon addition of 1.8 mM CaCl2, as indicated. These experiments were performed at 37 C and [Ca2+]i variations were monitored with a Hitachi F-2000 spectrofluorometer as described in Materials and Methods. These typical traces are representative of at least three independent experiments done in duplicate.

 
Ang II dose-dependent effects on IP3-induced Ca2+ release and capacitative Ca2+ entry activities were evaluated with a protocol similar to that used in Fig. 2EGo. In WT cells, increasing concentrations of Ang II from 1 pM to 1 µM caused intracellular Ca2+ releases of increasing amplitude (Fig. 3AGo, solid circles). The threshold dose was approximately 30 pM Ang II, the maximal amplitude (522 ± 66 nM Ca2+) was obtained with 1 µM Ang II, and the EC50 (dose producing 50% of the maximal release) was 1.0 ± 0.1 nM. In N111G cells (Fig. 3Go, open circles), the dose-response curve revealed an EC50 of 0.7 ± 0.5 nM, not significantly different from that obtained in WT cells, but the maximal amplitude (306 ± 33 nM Ca2+) was significantly lower than that obtained in WT cells. Capacitative Ca2+ entry in WT cells showed a typical dose-response curve with an EC50 of 0.3 ± 0.1 nM and a maximal amplitude of 223 ± 49 nM (Fig. 3BGo, solid circles). In N111G cells (Fig. 3BGo, open circles), the dose-response curve for capacitative Ca2+ entry was very different, with a relatively significant entry (69 ± 16 nM Ca2+) under basal conditions and a maximal amplitude (153 ± 16 nM Ca2+) much lower than that obtained in WT cells. The EC50 was 0.3 ± 0.1 nM, but the maximal Ang II-induced Ca2+ entry (difference between basal and maximal amplitude) was only 84 nM Ca2+.



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  		Fig. 3. Dose-Response Curves for Ang II-Induced Ca2+ Release and Ca2+ Entry

Populations (1.25 x 106 cells per assay) of WT cells (solid circles) or N111G cells (open circles) were loaded with fura 2-AM (5 µM) for 20 min at 37 C, washed by centrifugation, and resuspended in a nominally Ca2+-free medium, and the release of intracellular Ca2+ was measured after stimulation with increasing concentrations of Ang II (A). Ca2+ entry was measured 3 min after Ang II stimulation by adding 1.8 mM CaCl2 to the medium (B). These experiments were performed at 37 C, and [Ca2+]i variation was monitored with a Hitachi F-2000 spectrofluorometer as described in Materials and Methods. Each point represents the maximal amplitude of the Ca2+ variation (nM) and is expressed as the mean ± SD of at least three independent experiments done in duplicate.

 
The Constitutive Activity of N111G-AT1 Receptor Increases Basal Ca2+ Entry
As previously argued, basal Ca2+ entry in N111G cells is likely due to the constitutive activity of the N111G-AT1 receptor. To support this hypothesis, cells were incubated in a nominally Ca2+ free medium for different periods of time before their capacitative Ca2+ entry after the addition of extracellular Ca2+ was assessed. Under these conditions, WT cells showed only a slight time-dependent increase in Ca2+ entry, likely due to a minor leakage of Ca2+ from the intracellular pool toward the exterior of the cells (Fig. 4AGo, solid circles). N111G cells showed a high time-dependent increase in Ca2+ entry that was consistent with an important leak of intracellular Ca2+ due to the constitutive activity of the N111G-AT1 receptor (Fig. 4AGo, open circles). EXP3174 is an inverse agonist known to block the constitutive activity of the N111G-AT1 receptor (7). In the presence of a saturating concentration of EXP3174, the time-dependent increase in capacitative Ca2+ entry was not significantly affected in WT cells but was completely blunted in N111G cells (Fig. 4BGo). The elevated basal Ca2+ entry in N111G cells is therefore likely due to the constitutive activity of the mutant AT1 receptor.



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  		Fig. 4. Capacitative Ca2+ Entry under Basal Conditions

Populations (1.25 x 106 cells per assay) of WT cells (solid circles) or N111G cells (open circles) were loaded with fura 2-AM (5 µM) for 20 min at 37 C, washed by centrifugation, and resuspended in a nominally Ca2+-free medium for varying periods of time (ranging from 5 to 16 min) before measurement of their Ca2+ entry activity after addition of 1.8 mM CaCl2 to the medium (A). With a similar protocol, WT cells (solid columns) or N111G cells (open columns) were pretreated for 6 min without (Control) or with 4 µM EXP3174 before measurement of their Ca2+ entry activity after addition of 1.8 mM CaCl2 to the medium (B). These experiments were performed at 37 C, and [Ca2+]i variations were monitored with a Hitachi F-2000 spectrofluorometer as described in Materials and Methods. Data are expressed as mean ± SD of triplicate values and are representative of three independent experiments.

 
Integrity of Internal Ca2+ Stores
Ang II dose-dependent curves revealed a diminished intracellular Ca2+ release and a diminished maximal amplitude of Ca2+ entry in N111G cells. The content of the intracellular Ca2+ pool has a strong influence on IP3-induced Ca2+ release and capacitative Ca2+ entry. In a nominally Ca2+-free medium, thapsigargin, a potent sarcoplasmic endoplasmic reticulum Ca2+-ATPase inhibitor, caused the same amount of Ca2+ to be released from the intracellular stores of WT cells (339 ± 30 nM) and N111G cells (340 ± 21 nM) (Fig. 5Go, left). Comparable results were obtained with the ionophore ionomycin, revealing that the total cellular Ca2+ content was similar in both cell types (data not shown). Interestingly, capacitative Ca2+ entry after depletion of intracellular stores with thapsigargin was very similar in both cell types (WT cells, 418 ± 30 nM; N111G cells, 466 ± 49 nM) (Fig. 5Go, right). After treatment with EXP3174, thapsigargin-induced Ca2+ release and subsequent capacitative Ca2+ entry in WT and N111G cells were not modified (data not shown). These results suggest that the intrinsic mechanisms responsible for capacitative Ca2+ entry are not modified in N111G cells.



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  		Fig. 5. Integrity of the Intracellular Ca2+ Stores

Populations (1.25 x 106 cells per assay) of WT cells (A) or N111G cells (B) were loaded with fura 2-AM (5 µM) for 20 min at 37 C, washed by centrifugation, and resuspended in a nominally Ca2+-free medium before being exposed successively to 1 µM thapsigargin and 1.8 mM CaCl2. These experiments were performed at 37 C, and [Ca2+]i variations were monitored with a Hitachi F-2000 spectrofluorometer as described in Materials and Methods. These typical traces are representatives of three independent experiments done in duplicate.

 
Heterologous Desensitization of Ca2+ Responses in N111G Cells
To verify whether the refractory state of N111G cells is an AT1 receptor-specific phenomenon, we analyzed the Ca2+ responses induced by carbachol (CCh) and ATP, two Ca2+-mobilizing agonists of endogenously expressed muscarinic and purinergic receptors, respectively. In the absence of extracellular Ca2+, maximal doses of CCh and ATP caused intracellular Ca2+ releases that were weaker in N111G cells (Fig. 6AGo, open columns) than in WT cells (Fig. 6AGo, solid columns). Likewise, the addition of extracellular Ca2+ to cells pretreated with CCh or ATP resulted in capacitative Ca2+ entries that were weaker in N111G cells than in WT cells (Fig. 6BGo). These results indicated that the refractory state of N111G cells is due to some factor(s) common to the mechanism of action of several Gq-coupled receptors. The reversibility of the phenomenon was assessed by pretreating the cells with the inverse agonist EXP3174. The deficits in CCh-induced intracellular Ca2+ release (Fig. 6CGo) and CCh-induced capacitative Ca2+ entry (Fig. 6DGo) were totally eliminated in N111G cells after a relatively long (at least 24 h) pretreatment with EXP3174. Losartan and L-158,809, two partial inverse agonists, could only partially reverse the desensitized state of N111G cells (data not shown).



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  		Fig. 6. EXP3174 Rescues the Ca2+ Release and Ca2+ Entry Activities of N111G Cells

Populations (1.25 x 106 cells per assay) of WT cells (solid columns) or N111G cells (open columns) were loaded with fura 2-AM (5 µM) for 20 min at 37 C, washed by centrifugation, and resuspended in a nominally Ca2+-free medium, and their intracellular Ca2+ releases were measured upon stimulation with 100 µM CCh or 100 µM ATP (A). Ca2+ entry was measured 3 min after stimulation with agonists by adding 1.8 mM CaCl2 to the medium (B). With a similar protocol, CCh-induced Ca2+ release (C) and Ca2+ entry (D) activities were measured after a pretreatment of the cells for varying periods of time (ranging from 0.1 to 48 h) with 4 µM EXP3174. These experiments were performed at 37 C, and [Ca2+]i variations were monitored with a Hitachi F-2000 spectrofluorometer as described in Materials and Methods. These data are expressed as mean ± SD of triplicate values and are representative of three independent experiments.

 
Down-Regulation of IP3R in N111G Cells
The IP3R is an intracellular Ca2+ channel that is a common component of the mechanism of action of all Gq-coupled receptors. The functional properties of the IP3R were assessed by fura 2 spectrofluorometry in saponin-permeabilized cells. Figure 7AGo shows that permeabilized WT cells could take up Ca2+ within their intracellular store by an ATP-dependent process, thus decreasing the ambient Ca2+ concentration to a low steady level. The addition of increasing doses of IP3 caused rapid, transient releases of increasing amounts of Ca2+ until a maximal effect (11.8 ± 1.4 nmol of Ca2+ released) was obtained. Note that the efficient Ca2+ reuptake after each IP3-induced Ca2+ release is consequent to the rapid degradation of IP3 under these experimental conditions. In permeabilized N111G cells, relatively high doses of IP3 were required to release sequestered Ca2+, and the maximal effect (7.4 ± 0.2 nmol of Ca2+ released) was lower than in WT cells (Fig. 7BGo). The dose-response curves shown in Fig. 7CGo indicate that the IP3-induced Ca2+ release activity of N111G cells (open circles) was clearly less efficient than that of WT cells (solid circles). Interestingly, after a 48-h pretreatment of N111G cells with EXP3174 (Fig. 7CGo, open squares), their IP3-induced Ca2+ release activity was not significantly different from that of WT cells. When the results of Fig. 7CGo were plotted as a percentage of maximal release under each condition, the three curves were superimposable with similar EC50s of 0.29 ± 0.03 µM (Fig. 7DGo). These results are consistent with a reduction of IP3Rs in N111G cells.



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  		Fig. 7. IP3-Induced Ca2+ Release Activity in Permeabilized Cells

Populations (20 x 106 cells per assay) of WT cells (A) or N111G cells (B) were permeabilized for 3 min at 37 C in a cytosol-like buffer supplemented with 50 µg/ml of saponin, 0.5 µM fura 2 acid, 20 U of creatine kinase, and 10 mM phosphocreatine. Ca2+ uptake [upon addition of 1 mM ATP (A)] was partially released with increasing concentrations of IP3 (ranging from 0.1 to 3 µM). The amount of Ca2+ released was calibrated by adding a known amount of exogenous Ca2+ [4 nmol CaCl2 (C)]. Maximal fluorescence was measured by adding a saturating concentration of Ca2+ [1.8 mM CaCl2 (S)]. Panel C shows the dose-response curves for IP3-induced Ca2+ releases from WT cells (solid circles), N111G cells (open circles) and EXP3174-treated (4 µM for 48 h) N111G cells (open squares). Panel D shows the results of panel C represented as percent of maximal release. These experiments were performed at 37 C, and ambient [Ca2+] variations were monitored with a Hitachi F-2000 spectrofluorometer as described in Materials and Methods. These typical traces are representative of at least three independent experiments done in duplicate and summarized as mean ± SD values in panels C and D.

 
To directly assess the pharmacological properties of IP3Rs expressed in our clonal cell lines, [3H]IP3 binding studies were performed after permeabilization of the cells with saponin. The typical dose-displacement curves shown in Fig. 8Go, clearly indicate that the IP3 binding activity of N111G cells (open circles) was weaker than that of WT cells (solid circles). In both cell types, however, [3H]IP3 binding was inhibited in a similar fashion by increasing concentrations of unlabeled IP3. A Scatchard analysis of the data (inset) showed that the binding affinity of N111G cells (19.2 ± 1.3 nM) was not significantly different from that of WT cells (20.3 ± 1.7 nM). The maximal binding capacity, however, was significantly lower in N111G cells (Bmax of 0.38 pmol/mg of protein) than in WT cells (Bmax of 0.86 pmol/mg of protein).



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  		Fig. 8. [3H]IP3 Binding to Permeabilized Cells

WT cells (solid circles) or N111G cells (open circles) were permeabilized for 10 min at 37 C in an intracellular-like medium supplemented with saponin (50 µg/ml) and then incubated (20 x 106 cells per tube) for 15 min at 0 C with approximately 2 nM [3H]IP3 (15,000 cpm) and increasing concentrations of unlabeled IP3 (ranging from 0.1 nM to 1 µM) as described in Materials and Methods. The Kd and Bmax values were calculated from the Scatchard analysis of the data (inset). These data expressed as the means ± SEM of duplicate values are representative of three independent experiments.

 
The level of expression of type III IP3R (IP3RIII), an abundant subtype in QBI-HEK 293A cells, was directly assessed by immunoblot analysis with a selective anti-IP3RIII antibody. Figure 9AGo (upper panel) shows that IP3RIII migrates on SDS-PAGE as a single sharp band with a molecular mass of approximately 230 kDa. The intensity of this band was higher in extracts from WT cells (lanes 1 and 2) than in extracts from N111G cells (lanes 3 and 4). A 48-h pretreatment of cells with EXP3174 did not modify the intensity of this band in extracts from WT cells (lanes 5 and 6) but significantly increased the intensity of the band in extracts from N111G cells (lanes 7 and 8). The densitometric analysis shown at Fig. 9BGo clearly indicates that N111G cells expressed less IP3RIII than WT cells and that the level of expression of IP3RIII was significantly increased by about 2-fold after a treatment of N111G cells with the inverse agonist EXP3174. These results suggest that the refractory state of N111G cells was due, at least in part, to a down-regulation of IP3R.



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  		Fig. 9. Immunoblot Analysis of IP3RIII

WT cells and N111G cells were grown for 48 h in the absence (lanes 1–4) or in the presence of 4 µM EXP3174 (lanes 5–8). In panel A, proteins from WT cells lysates (lanes 1, 2, 5, and 6) and from N111G cells lysates (lanes 3, 4, 7, and 8) (1 x 105 cells per lane) were resolved on a 7% polyacrylamide gel and immunoblotted with an anti-IP3RIII antibody (upper panel) or with an antiactin antibody (lower panel) as described in Materials and Methods. Bands corresponding to IP3RIII (~230 kDa) and actin (42 kDa) are identified with black and white arrows, respectively. Panel B shows the densitometric analysis of the IP3RIII that was performed as described in Materials and Methods and expressed as means ± SEM of relative units of integrated peaks (ratio of IP3RIII/actin densities). These results are representatives of at least three independent experiments. *, P < 0.05 compared with respective EXP3174-untreated cells.

 
It is important to note that most of the results obtained with our N111G cell clone (weaker intracellular Ca2+ release, weaker capacitative Ca2+ entry, and lower expression of IP3RIII than in WT cells) were repeated with a heterogenous population of G-418-resistant N111G-AT1 receptor-transfected cells (Fig. 10Go). Therefore, the down-regulation of IP3R appears to be a common cellular mechanism in response to the expression of a constitutively active AT1 receptor and not a mere epiphenomenon observed with an eccentric cell clone.



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  		Fig. 10. Ca2+ Responses and IP3RIII Expression in Heterogenous Populations of G-418-Resistant Transfected Cells

Heterogenous populations (1.25 x 106 cells per assay) of G-418-resistant WT- (A and B) or N111G-AT1 receptor-transfected cells (C and D) were loaded with fura 2-AM (5 µM) for 20 min at 37 C, washed by centrifugation, and resuspended in a nominally Ca2+-free medium, and their intracellular Ca2+ concentration was monitored upon stimulation with 100 µM CCh or upon addition of 1.8 mM CaCl2, as indicated. These experiments were performed at 37 C, and [Ca2+]i variations were monitored with a Hitachi F-2000 spectrofluorometer as described in Materials and Methods. These typical traces are representative of at least three independent experiments done in triplicate. In panel E, proteins from heterogenous populations of G-418-resistant WT- (lanes 1 and 2) or N111G-AT1 receptor-transfected cells lysates (lanes 3 and 4) were resolved on a 7% polyacrylamide gel and immunoblotted with an anti-IP3RIII antibody (upper panel) or with an antiactin antibody (lower panel) as described in Materials and Methods. Bands corresponding to IP3RIII (~230 kDa) and actin (42 kDa) are identified with black and white arrows, respectively. This result is representative of three independent experiments.

 
Mechanism of IP3RIII Down-Regulation in N111G Cells
Different mechanisms could be responsible for the down-regulation of IP3RIII in N111G cells. RNA analyses by various cycles of RT-PCR revealed that the mRNA levels for IP3RIII were identical in WT and N111G cells, indicating no change in the expression and the stability of gene product (Fig. 11AGo). With a pulse-chase labeling approach, the rates of synthesis and degradation of IP3RIII were analyzed. Metabolically labeled IP3RIII appeared within about 30 min and gradually increased for 4 h (Fig. 11BGo). Within this time period, the rate of appearance of metabolically labeled IP3RIII did not differ between WT and N111G cells, indicating no change in the rate of synthesis of IP3RIII. At the end of a 6-h pulse labeling, a 24-h chase showed that the level of metabolically labeled IP3RIII declined more rapidly in N111G cells than in WT cells (Fig. 11CGo). Measurements made at 8, 12, 16, 20, and 24 h demonstrated different disappearance rates in both cell types. After 24 h, pulse-labeled IP3RIII had declined by 24% in WT cells and by 83% in N111G cells. In the presence of the inverse agonist EXP3174, the decline of pulse-labeled IP3RIII in N111G cells was substantially reduced (data not shown). These results indicate that the down-regulation of IP3RIII in N111G cells is via a protein degradation pathway. Immunoblot assays after preincubation of cells for different times with the protein synthesis inhibitor cycloheximide further suggested that IP3RIII was degraded more rapidly in N111G cells than in WT cells (Fig. 11DGo). Again, in the presence of EXP3174, the degradation of immunoreactive IP3RIII was reduced (data not shown). At the end of a 6-h pulse labeling, an 8-h chase in the presence of cycloheximide revealed an important decline of the metabolically labeled IP3RIII in N111G cells (Fig. 11EGo). Addition of chloroquine or NH4Cl (two lysosomal activity inhibitors), during the chase period, offered good protection against degradation whereas lactacystin and N-acetyl-Leu-Leu-Nle-CHO (ALLN) (two proteasomal activity inhibitors) offered only weak protection. These results suggest that the main degradation pathway for the down-regulation of IP3RIII in N111G cells involves the lysosome.



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  		Fig. 11. Mechanism of IP3RIII Down-Regulation in N111G Cells

RT-PCR analysis (from cycles 25 to 50) of IP3RIII mRNA extracted from WT and N111G cells is shown in panel A. In panels B, C, and E, WT and N111G cells were pulse labeled with Expre35S35S-Protein labeling mix (50 µCi) for 0.5 to 6 h. Cell lysates were immunoprecipitated with anti-IP3RIII antibody and resolved on a 7% polyacrylamide gel. The gel was dried and subjected to fluorography for at least 5 d. In panel C, WT and N111G cells were pulse labeled for 6 h and subsequently chased for 1–24 h. In panel D, WT and N111G cells were treated with cycloheximide (50 µg/ml) for indicated times, and IP3RIII from cell lysates were revealed by immunoblot analysis as described in Fig. 9Go. In panel E, WT and N111G cells were pulsed for 6 h, and the indicated inhibitors were applied to the cells together with cycloheximide (50 µg/ml) during an 8-h chase. Final concentrations of the inhibitors were as follows: chloroquine (ChQ, 200 µM), NH4Cl (5 mM), lactacystin (LC, 10 µM), and ALLN (30 µM). Bands corresponding to IP3RIII (~230 kDa) were identified with black arrows and actin control with white arrow. These typical results are representative of at least three independent experiments.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
It is expected that the expression of a mutant receptor with constitutive activity would perturb cell homeostasis either by causing exaggerated downstream activities or by mediating adaptive refractory processes. In the study presented here, we showed that QBI-HEK 293A cells expressing the constitutively active N111G-AT1 receptor exhibited a refractory state in their Ca2+ signaling mechanism. Despite their spontaneous Ca2+ oscillatory activity and their basal capacitative Ca2+ entry activity, N111G cells had weaker responses than WT cells to low and high concentrations of Ang II. When the different phases of their Ang II-induced Ca2+ response were analyzed, N111G cells displayed a lower release of intracellular Ca2+ and weaker capacitative Ca2+ entry than WT cells. Because thapsigargin released the same amount of intracellular Ca2+ and triggered a similar Ca2+ entry in both cell types, the refractory state of N111G cells can be attributed neither to a lower content of their intracellular Ca2+ pool nor to the desensitization of a component of their capacitative Ca2+ entry mechanism.

Relatively few studies have examined the underlying effects of the expression of a constitutively active Gq-coupled receptor on cellular Ca2+ homeostasis. We previously observed that COS-7 cells expressing the N111G-AT1 receptor produced a diminished intracellular Ca2+ transient in response to Ang II (16). In HEK 293 cells stably transfected with the N111A-AT1 receptor, Ang II-induced intracellular Ca2+ transients were no different from those obtained with cells expressing the WT AT1 receptor (9). In our hands, the N111A-AT1 receptor had relatively weak constitutive activity (18% of maximal WT activity) that may not be strong enough to cause a significant adaptive response (16). The expression of a constitutively active mutant TRH receptor caused a desensitization of the Ca2+ response to TRH (homologous) and also to CCh (heterologous) in AtT20 cells (17). The authors provided evidence that the desensitization process was dependent on the availability of extracellular Ca2+ and on the activity of protein kinase C, but no specific Ca2+ handling component or intracellular target was identified. The GTPase-deficient Q212L-G16{alpha} which constitutively activates phospholipase C, creates a cell environment that may be similar theoretically to that created by a constitutively active Gq-coupled GPCR. NIH-3T3 cells stably transfected with Q212L-G16{alpha} had a desensitized Ca2+ response to ATP (18). These cells did not show any capacitative Ca2+ entry under basal conditions, and their intracellular Ca2+ store was partially depleted. These different adaptive responses between Q212L-G16{alpha} cells and N111G cells could be directly related to phenotypic differences between the two cell types. They could also be related to the fact that a specific GPCR may activate several downstream effectors through direct interactions with G proteins and also with diverse adaptors or scaffolding proteins (arrestins, A-kinase anchoring proteins, InaD, Homer, Janus kinase, etc.; for review see Ref.19) whereas a specific G protein is known to interact with a more restricted set of downstream partners. Xenopus oocytes expressing the constitutively active Kaposi’s sarcoma-associated herpes virus GPCR showed homologous and heterologous (elicited by TRH or acetylcholine) desensitizations of their Ca2+ response accompanied with an impaired response to IP3 injection (20). The main cause of this impairment appeared to be a depletion of their thapsigargin-sensitive intracellular Ca2+ pool. In HEK 293 cells expressing a WT TRH receptor, preexposure to a high concentration of TRH caused an adaptive response mainly due to the depletion of their intracellular Ca2+ pool (21). Again, this adaptive response was very different from that of N111G cells, which showed a significant capacitative Ca2+ entry activity under basal conditions and no apparent depletion of their intracellular Ca2+ pool. N111G cells therefore possess an efficient store-operated Ca2+ influx mechanism that contributes adequately to the refilling of their intracellular Ca2+ pool.

The refractory state of N111G cells was also revealed when their endogenous muscarinic (CCh) and purinergic (ATP) receptors were stimulated. These results indicated that the diminished agonist-induced Ca2+ release activity observed in N111G cells is a heterologous desensitization phenomenon that is not directly related to the specific properties of the N111G-AT1 receptor but rather to a deficient component downstream from the receptor. The G protein Gq and phospholipase C are located immediately downstream from the receptor. Agonist-dependent desensitization of Gq{alpha} or phospholipase C have been observed in different cell types (22, 23, 24, 25). However, rat 1 fibroblasts expressing a constitutively active {alpha}1B-adrenergic receptor did not show any desensitization of phospholipase C or any down-regulation of Gq{alpha} unless they were chronically stimulated with phenylephrine (24). These studies suggest that very strong stimulations are necessary to desensitize Gq{alpha} and phospholipase C. Because we showed that, in response to a high dose of Ang II, N111G cells could produce the same maximal amount of IP as WT cells, it is unlikely that the activities of the G protein Gq and of phospholipase C are depressed in these cells. This interpretation is supported by immunoblot studies showing similar amounts of the G protein G{alpha}q and of phospholipase Cß3 in WT and N111G cells (data not shown).

The next component of the Ca2+ signaling mechanism downstream from phospholipase C is the IP3R. We showed that IP3 released less Ca2+ from permeabilized N111G cells than from permeabilized WT cells, suggesting that the refractory state of N111G cells could be related to the function of IP3R. This possibility was supported by binding studies showing a diminished amount of IP3R in N111G cells. Our immunoblot analysis further showed that IP3RIII is less abundant in N111G cells than in WT cells. Agonist-induced down-regulation of IP3R was first demonstrated in SH-SY5Y neuroblastoma cells activated with high concentrations of CCh (26). Other studies showed that chronic activation of different Gq-coupled receptors causes a down-regulation of IP3R within different cell types (27, 28, 29, 30). Agonist-induced IP3R down-regulation occurs by a mechanism involving the proteasome pathway (31, 32). Interestingly, the mechanism responsible for the down-regulation of IP3RIII in N111G cells appears to involve primarily the lysosome and to a minor extent the proteasome. This difference could be related to the fact that the N111G-AT1 receptor is chronically producing a submaximal stimulation whereas agonist-induced IP3R down-regulation was obtained with supramaximal and relatively acute doses of agonists. Further studies are needed to clarify that question. Nonetheless, the N111G cells represent another interesting model with which to study the degradation pathways for IP3R.

In conclusion, it is known that prolonged elevations of intracellular Ca2+ may be very deleterious for cells (33). The survival of cells expressing a constitutively active Ca2+-mobilizing receptor is therefore dependent on some desensitization of their Ca2+ signaling pathway. Our results, obtained with a N111G cell clone and also with a heterogenous population of G-418-resistant N111G-AT1 receptor-transfected cells, indicated that the IP3RIII is down-regulated in these cells, which nonetheless maintain a normal Ca2+ concentration under basal conditions and can still respond, although less efficiently than WT cells, to Ang II and other Ca2+-mobilizing stimuli. Interestingly, long-term treatments with the inverse agonist EXP3174 restored the CCh-induced Ca2+ transient, IP3-induced Ca2+ release and the level of expression of IP3RIII in N111G cells. The time course of these recovery responses was consistent with de novo protein synthesis and with the metabolic turnover of IP3R (30, 34). Whereas the activation of IP3R plays a fundamental role in cellular Ca2+ responses, its down-regulation appears to be a key mechanism to protect cells against the deleterious effects of chronic Ca2+ elevations.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Materials
The cDNA clones encoding AT1 receptor and N111G-AT1 receptor, both with a N-terminal FLAG epitope, were constructed in our laboratory as described previously (16). QBI-HEK 293A cells were from Qbiogene (Carlsbad, CA). DMEM, fetal bovine serum (FBS), geneticin (G-418 sulfate), lipofectamine, Met/Cys-free DMEM, and penicillin-streptomycin-glutamine were from Life Technologies (Gaithersburg, MD). EN3HANCE Reagent, [3H]IP3 (23 Ci/mmol), myo-[3H]inositol (65 Ci/mmol), and Expre35S35S-Protein labeling mix (1175 Ci/mmol) were from PerkinElmer Corp. (Boston, MA). IP3 was from Alexis Biochemicals (San Diego, CA). Ang II, ATP, bacitracin, BSA, chloroquine diphosphate salt, creatine phosphokinase, phosphocreatine, poly-L-lysine hydrobromide, saponin, and thapsigargin were from Sigma-Aldrich (Oakville, Ontario, Canada). AG 1-X8 resin was from Bio-Rad Laboratories (Mississauga, Ontario, Canada). ALLN, CCh, fura 2 (free acid), fura 2-AM, and lactacystin were from Calbiochem (San Diego, CA). Proteases inhibitors cocktail (Complete) and Expand High Fidelity DNA polymerase were from Roche Molecular Biochemicals (Laval, Quebec, Canada). Moloney murine leukemia virus reverse transcriptase was from Promega Corp. (Mississauga, Ontario, Canada). TRIzol reagent was from Invitrogen Life Technologies (Burlington, Ontario, Canada). Immobilon-P polyvinylidene fluoride transfer membranes were from Millipore Corp. (Bedford, MA). Mouse anti-IP3RIII antibody (recognizing an N-terminal epitope) was from BD Biosciences Transduction Laboratories (Mississauga, Ontario, Canada). Mouse antiactin antibody was from Chemicon International (Temecula, CA). AMDEX sheep antimouse IgG antibody coupled to horseradish peroxidase and ECL Plus Western blotting detection reagents were from Amersham Biosciences (Piscataway, NJ). EXP3174 is a generous gift previously obtained from DuPont Merck Pharmaceutical Co. (Wilmington, DE). [125I]Ang II (1000 Ci/mmol) was prepared with IODO-GEN (Pierce Chemical Co., Rockford, IL) according to the method of Fraker and Speck (35) in an acetic acid buffer (pH 5.4) and purified by HPLC on a C-18 column (Waters Corp., Mississauga, Ontario, Canada) as previously reported (36). The specific radioactivities of the radiolabeled peptides were determined by self-displacement and saturation binding experiments as described previously (37).

Cell Culture
QBI-HEK 293A clonal cell lines were cultured in complete DMEM (supplemented with 10% heat-inactivated FBS, 2 mM L-glutamine, 100 IU/ml penicillin, 100 µg/ml streptomycin, and 0.4 mg/ml G-418) at 37 C in a humidified atmosphere containing 5% CO2 and 95% air. To establish clonal cell lines expressing either the AT1 receptor or the N111G-AT1 receptor, QBI-HEK 293A cells at 60–70% confluence were transfected with 4 µg of the cDNA constructs and 25 µl of lipofectamine. Cells were grown for 36 h before selection with 0.8 mg/ml G-418 for 2 wk. Most of these G-418-resistant cells were conserved as a heterogenous population of stably transfected cells. Some of the G-418-resistant cells were seeded at a density of 0.5 cell per well into 96-well plates. G-418-resistant clones were amplified and tested for AT1 receptor expression with a [125I]Ang II binding assay as previously described (16).

Dynamic Video Imaging of Cytosolic Ca2+
Fluorescence from fura 2-loaded cells was monitored as previously described (38). Briefly, QBI-HEK 293A cells were allowed to attach to a glass coverslip (no. 1) coated with poly-L-lysine and to grow in complete DMEM for 36–48 h before being washed twice with a HEPES-buffered physiological saline solution (HBSS: 20 mM HEPES at pH 7.4; 120 mM NaCl; 5.3 mM KCl; 0.8 mM MgSO4; 1.8 mM CaCl2; and 11.1 mM dextrose). The coverslips were clamped into a Teflon circular open-bottom chamber, and cells were incubated with 0.2 µM fura 2-AM for 20 min at room temperature in the dark. Cells were then washed and bathed in fresh HBSS for 20 min to ensure complete hydrolysis of the fura 2-AM before mounting the Teflon chamber onto the stage of a Axiovert inverted microscope (Carl Zeiss, Thornwood, NY) fitted with an Attofluor Digital Imaging and Photometry System (Attofluor Inc., Rockville, MD). The system allows data acquisition from up to 99 user-defined variably sized regions of interest per field of view. Fluorescence from isolated fura 2-loaded cells was monitored by videomicroscopy using alternating excitation wavelengths of 334 and 380 nm and recording emitted fluorescence at 510 nm. All experiments were done at room temperature, and the data are expressed as fura 2 fluorescence ratio (F334/F380). Data acquisition was typically at 3-sec intervals and lasted for 1600 sec.

Cytosolic [Ca2+] Measurement
QBI-HEK 293A cells (1.25 x 106 cells grown in 10-cm dishes for 24–40 h) were detached by a brief trypsin/EDTA treatment, resuspended in complete DMEM, and washed by centrifugation for 4 min at 100 x g before being incubated with 5 µM fura 2-AM in an extracellular-like medium (ECM) (15 mM HEPES at pH 7.4; 140 mM NaCl; 5 mM KCl; 1 mM MgCl2; 10 mM dextrose; 1.8 mM CaCl2; and 0.1% BSA) for 20 min at 37 C. After a wash by centrifugation, cells were resuspended in ECM and incubated for 20 min at 37 C to ensure complete hydrolysis of the fura 2-AM. Cells were then centrifuged again and resuspended in 2 ml of ECM or of nominally Ca2+-free ECM (which was identical in composition except for the omission of CaCl2). Cells suspension was gently stirred in a quartz cuvette maintained at 37 C while [Ca2+]i was monitored on a F-2000 spectrofluorometer (Hitachi Scientific Instruments, Inc., Hialeah, FL) with alternative excitation wavelengths of 340 and 380 nm and with emission wavelength of 510 nm to measure changes in intracellular fura 2 fluorescence intensity (F). At the end of each recording, maximal fluorescence ratio (Rmax) and minimal fluorescence ratio (Rmin) were determined by adding successively 0.1% Triton X-100 and 10 mM EGTA to the cell suspensions. The following equation from Grynkiewicz et al. (39) was used to relate the intensity ratios to Ca2+ levels:

where R represents the fluorescence intensity ratio F{lambda}1 (340 nm)/F{lambda}2 (380 nm). KD is the Ca2+ dissociation constant of the indicator (224 nM). Drugs were added in small volumes (<20 µl) of concentrated stocks (dissolved either in water or dimethylsulfoxide).

Intracellular IP Accumulation Measurement
QBI-HEK 293A cells (5 x 105 cells per well into six-well plates) were labeled for 18–24 h in inositol-free DMEM containing 15 µCi/ml of myo-[3H]inositol. Cells were then stimulated with 100 nM Ang II for 20 min at 37 C in Medium 199 containing 25 mM HEPES at pH 7.4, 10 mM LiCl, and 0.1% BSA. Incubations were stopped by addition of ice-cold perchloric acid [5% (vol/vol)]. Water-soluble IP were then extracted with an equal volume of a 1:1 mixture of 1,1,2-trichlorotrifluoroethane and tri-n-octylamine. The samples were vigorously mixed and centrifuged at 15,000 x g for 15 min at 4 C. The upper phase was applied to an AG 1-X8 resin column, and the IP were sequentially eluted by addition of ammonium formate/formic acid solutions of increasing ionic strength. Radioactive content from each sample was evaluated with an LS 6800 liquid scintillation counter (Beckman, Fullerton, CA).

IP3-Induced Ca2+ Release
QBI-HEK 293A cells (20 x 106 cells grown in 15-cm dishes) were detached by a brief trypsin/EDTA treatment, resuspended in 10 ml DMEM, and washed by centrifugation. After rinsing with 5 ml of a cytosol-like buffer (20 mM Tris/HCl at pH 7.4; 110 mM KCl; 10 mM NaCl; 5 mM KH2PO4; and 2 mM MgCl2), cells were resuspended in 2 ml of permeabilization buffer composed of the cytosol-like buffer supplemented with 50 µg/ml of saponin, 0.5 µM fura 2 acid, 20 U of creatine kinase, and 10 mM phosphocreatine. After 3 min of permeabilization, less than 10% of the cells excluded Trypan Blue. Ca2+ uptake (upon ATP addition) and release (upon IP3 addition) were monitored at 37 C with a Hitachi F-2000 spectrofluorometer as previously described (40).

[3H]IP3 Binding Assay
QBI-HEK 293A cells were permeabilized in a binding buffer (25 mM Tris/HCl at pH 8.5; 110 mM KCl; 10 mM NaCl; 5 mM KH2PO4; 1 mM EDTA) supplemented with 50 µg/ml saponin for 10 min at 37 C. Cells (20 x 106 cells/0.5 ml) were then incubated for 15 min at 0 C in the presence of approximately 2 nM [3H]IP3 (15,000 cpm) and increasing concentrations of unlabeled IP3 (ranging from 0.1 nM to 1 µM). Nonspecific binding was determined in the presence of 2 µM IP3. Incubations were terminated by centrifugation at 15,000 x g for 5 min at 4 C. The pellets were solubilized with 1% Triton X-100, and the receptor-bound radioactivity was evaluated with a Beckman LS 6800 liquid scintillation counter.

Electrophoresis and Immunoblotting
QBI-HEK 293A cells (107 cells/ml) were solubilized for 1 h at 4 C in solubilization buffer (50 mM Tris/HCl at pH 7.4; 150 mM NaCl; 1 mM EDTA; 1% Triton X-100; and the proteases inhibitors cocktail Complete 1X). Insoluble material was precipitated by centrifugation at 35,000 x g for 30 min at 4 C. The supernatant was mixed with 1 ml of 2x Laemmli’s buffer (60 mM Tris/HCl at pH 6.8; 10% glycerol; 2% sodium dodecyl sulfate, 125 mM dithiothreitol; and 0.3% bromophenol blue) and heated for 5 min at 95 C. Samples were loaded onto a 7% polyacrylamide gel that was subjected to a constant current of 15 mA for 105 min. Proteins were electrotransferred to a polyvinylidene fluoride membrane at a constant current of 0.5 A for 24 h at 4 C. Blotted membranes were incubated for 2 h at room temperature in PBS-T (PBS containing 0.1% Tween-20) supplemented with 5% nonfat dried milk. Blots were then incubated overnight at 4 C with either the anti-IP3RIII antibody or the antiactin antibody. After extensive washing with PBS-T, the blots were incubated for 1 h at room temperature with a peroxidase-conjugated secondary antibody, and after extensive washing with PBS-T, the immunostained bands were revealed with ECL Plus according to the manufacturer’s instruction on a BioMax ML film (Eastman Kodak, Rochester, NY). Autoradiograms were digitized on a Hewlett Packard Scan Jet 5100c (Hewlett-Packard Co., Palo Alto, CA) and integrated peak areas were determined using the gel analysis Quantity One software (version 4.2; Bio-Rad Laboratories).

RT-PCR
Total RNA was isolated from WT and N111G cells using TRIzol reagent according to the manufacturer’s instruction. cDNA synthesis was carried out using Moloney murine leukemia virus reverse transcriptase with random hexamers as primers according to the manufacturer’s instruction. PCR was performed using Expand High Fidelity DNA Polymerase with standard buffer. For each reaction, 1 µg of cDNA template was used, and cycling conditions consisted of 94 C for 30 sec, 55 C for 1 min, and 72 C for 2 min for 50 cycles of PCR carried out with the iCycler system (Bio-Rad Laboratories). Approximately 5 µl of product were collected every five cycles (beginning at cycle 25) and run on 0.5x Tris borate/EDTA-2% agarose gel. The specific oligonucleotide primer sets used for amplification of IP3RIII (nucleotides 1975–2534) were as follows: sense, 5'-TACCCCAAGAGCTCATCTGCAAG-3'; and antisense, 5'-ACTTGTTCTTCTTGTCATCTGGGG-3'. The separated PCR fragments were visualized using the Gel Doc system from Bio-Rad.

Metabolic Labeling and Immunoprecipitation
Metabolic labeling experiments were performed on WT and N111G cells (1.5 x 106 cells in 6-cm dishes). Cells were incubated for 1 h with Met/Cys-free DMEM supplemented with 2 mM L-glutamine, 100 IU/ml penicillin, 100 µg/ml streptomycin, and 0.4 mg/ml G-418 before adding 50 µCi of Expre35S35S-Protein labeling mix for the indicated periods of time (pulse). Chase was done by replacing pulse labeling medium for DMEM without FBS supplemented with different inhibitors as indicated. After washing twice, cell lysates were prepared as described above and immunoprecipitated with anti-IP3RIII antibody (5 µl) on wet protein A/G-plus agarose beads (50 µl; Santa Cruz Biotechnology, Inc., Santa Cruz, CA) for 16 h, at 4 C, under constant rotation. The agarose beads were sedimented by centrifugation at 5000 x g for 2 min, and the immune complexes were washed once with ice-cold solubilization buffer before resuspension in 45 µl of 1x Laemmli buffer. Labeled proteins were resolved by SDS-PAGE on a 7% polyacrylamide gel. Separated proteins were fixed before the gels were treated with EN3HANCE for 1 h, dried for 2 h at 60 C under a vacuum, and exposed (for at least 5 d) on a BioMax MS film with an intensifying screen. Integrated peak areas were determined using the gel analysis Quantity One software.


   FOOTNOTES
 
This work was supported by the Canadian Institutes of Health Research (CIHR). M.A.-M. is recipient of a studentship from CIHR. G.A. is recipient of a studentship from Natural Sciences and Engineering Research Council of Canada. R.L. is a Senior Scholar from the Fonds de la Recherche en Santé du Québec (FRSQ). E.E. is recipient of a J.C. Edwards Chair in cardiovascular research. This work is part of the Ph.D. thesis of M.A.-M.

M.A.-M. and G.A. contributed equally to this work and should both be considered first authors.

Abbreviations: ALLN, N-Acetyl-Leu-Leu-Nle-CHO; Ang II, angiotensin II; AT1 receptor, angiotensin II type-1 receptor; CAM, constitutively active mutant; CCh, carbachol; ECM, extracellular-like medium; FBS, fetal bovine serum; GPCR, G protein-coupled receptor; HBSS, HEPES-buffered physiological saline solution; HEK, human embryonic kidney; IP, inositol phosphate; IP3, inositol 1,4,5-trisphosphate; IP3R, inositol 1,4,5-trisphosphate receptor; IP3RIII, type III IP3R; PBS-T, PBS containing 0.1% Tween 20; WT, wild-type.

Received for publication December 18, 2003. Accepted for publication August 18, 2004.


   REFERENCES
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 ABSTRACT
 INTRODUCTION
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 DISCUSSION
 MATERIALS AND METHODS
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