Phospholipase Cδ1 Is a Guanine Nucleotide Exchanging Factor for Transglutaminase II (Gαh) and Promotes α1B-Adrenoreceptor-mediated GTP Binding and Intracellular Calcium Release

Effectors involved in G protein-coupled receptor signaling modulate activity of GTPases through GTPase-activating protein or guanine nucleotide exchanging factor (GEF). Phospholipase Cδ1 (PLCδ1) is an effector in tissue transglutaminase (TGII)-mediated α1B-adrenoreceptor (α1BAR) signaling. We investigated whether PLCδ1 modulates TGII activity. PLCδ1 stimulated GDP release from TGII in a concentration-dependent manner, resulting in an increase in GTPγS binding to TGII. PLCδ1 also inhibited GTP hydrolysis by TGII that was independent from the α1BAR. These results indicate that PLCδ1 is GEF for TGII and stabilizes the GTP·TGII complex. When GEF function of PLCδ1 was compared with that of the α1BAR, the α1BAR-mediated GTPγS binding to TGII was greater than PLCδ1-mediated binding and was accelerated in the presence of PLCδ1. Thus, the α1BAR is the prime GEF for TGII, and GEF activity of PLCδ1 promotes coupling efficacy of this signaling system. Overexpression of TGII and its mutants with and without PLCδ1 resulted in an increase in α1BAR-stimulated Ca2+ release from intracellular stores in a TGII-specific manner. We conclude that PLCδ1 assists the α1BAR function through its GEF action and is primarily activated by the coupling of TGII to the cognate receptors.

Phospholipase C (PLC) 1 ␦1 is a member of the PLC family that produces two second messengers, inositol 1,4,5-triphosphate (IP 3 ) and diacylglycerol by hydrolyzing phosphatidylinositol 4,5-bisphosphate (PIP 2 ) (1). Among PLCs, PLC␤ isozymes are stimulated by guanine nucleotide-binding proteins (G protein) G q and its family of proteins in response to activation of cell surface receptors. PLC␥ isozymes are activated by phosphorylation through growth factor receptors. A number of laboratories have reported that PLC␦1 is stimulated by a unique GTP-binding protein known as tissue transglutaminase (TGII, G␣ h ) (2)(3)(4)(5). TGII is a bifunctional enzyme, having GTPase and transglutaminase (TGase) activity (6,7) and is present in the plasma membrane, cytosol, and nucleus in a variety of tissues and cells (6). Exchange of GDP to GTP by TGII is facilitated by activation of the cell surface receptors (4 -9). These receptors include the ␣ 1B -adrenoreceptor (AR) (5,7,8), ␣ 1D AR (8), ␣-thromboxane receptor (9), and oxytocin receptor (4). The coupling of TGII with these receptors appears to be receptor subtype-specific (8,9). PLC␦1 is widely distributed and expressed highly in some tissues such as mouse heart (1,10). Stimulation of the enzyme by TGII involving ␣ 1B AR is modulated in a concerted fashion within the system. Thus, bimodal regulation of PLC␦1 activity has been observed depending on the Ca 2ϩ levels and occupancy of guanine nucleotide by TGII (3,11,12). PLC␦1 is stimulated with low concentrations of Ca 2ϩ by GTP␥S⅐TGII (11). However, activity of the enzyme is subsequently inhibited by increasing Ca 2ϩ concentrations where PLC␦1 is stimulated in the presence or absence of GDP. Similarly, Murthy et al. (3) have reported that GTP⅐TGII inhibits PLC␦1, while GDP⅐TGII stimulates the enzyme. The Ca 2ϩ dependence is not clearly defined in this study. The TGII-mediated PLC stimulation is also modulated by the level of TGII expression (12). At low levels of TGII expression, the ␣ 1B AR-stimulated PLC activity is increased, whereas the receptor-mediated PLC stimulation is inhibited when TGII is highly expressed. The PLC␦1 activity is also inhibited by IP 3 , competing with its substrate PIP 2 for a binding site known as the pleckstrin homology domain (13)(14)(15). Studies have also demonstrated that an increase in the intracellular concentration of Ca 2ϩ activates PLC␦1 (13,16,17), indicating that activation of PLC␦1 occurs secondarily in response to the receptor-mediated activation of other PLCs or Ca 2ϩ channels. A GTPase activating-protein (GAP) for the small GTPase RhoA (RhoGAP) also activates PLC␦1 by direct association (18). All of these observations suggest that the PLC␦1 activity is regulated by multiple factors.
All known GTP binding subunits (G␣) of G proteins are GTPases, which hydrolyze GTP to GDP and orthophosphate (P i ). It is now recognized that a large number of regulators of G protein signaling (RGS) facilitate GTP hydrolysis by G␣ proteins (19 -21). Independent from the actions of these RGS proteins, certain effectors in the G protein-coupled receptor system modulate GTP hydrolysis by G␣ proteins acting as GAP or guanine nucleotide exchanging factor (GEF) (22)(23)(24)(25)(26). For ex-ample, PLC␤1 (22)(23)(24) and the ␥ subunit of cGMP phosphodiesterase (25) directly accelerates GTP hydrolysis by G␣ q and G␣ t , respectively. A recent study by Scholich et al. (26) has shown that adenylyl cyclase facilitates GTP binding to G s , thereby functioning as both GEF and GAP. These findings indicate that the effector molecules modulate their cognate GTPases to terminate or facilitate the signals.
To date, none of the known heterotrimeric G proteins stimulates PLC␦1 (1,27), and the mechanisms that regulate PLC␦1 activity remain complex and unclear. To further understand the characteristics and the interaction of PLC␦1 with TGII and its activation by TGII and the ␣ 1B AR, we investigated the roles of PLC␦1 in the modulation of TGII activities, including the ␣ 1B AR. Here, we report a distinct role of PLC␦1 in a coupling system involving the ␣ 1B AR and TGII. PLC␦1 displays two regulatory functions for TGII. One is a GEF function, and the other is the inhibition of GTP hydrolysis by TGII. The GEF function of PLC␦1 promotes the ␣ 1B AR-mediated GTP binding by TGII. Furthermore, our results also reveal that PLC␦1 is primarily activated by the activation of the ␣ 1B AR through TGII, resulting in Ca 2ϩ release from intracellular stores. Purification of Proteins-PLC␦1 was expressed in DH␣5 cells and purified as described (2). The purity of the PLC␦1 preparation was Ն90%, as judged by silver staining, and neither GTP␥S binding and TGase activity were observed. Guinea pig liver TGII was further purified using GTP-agarose as described (7,28). The purity of the TGII preparation was Ն95% as judged by silver staining, and PLC activity was not found in the TGII preparation as determined by measurement of PIP 2 hydrolysis (2). It should be noted that purified TGII was stable for Յ3-4 weeks in the presence of 10% glycerol at Ϫ80°C. The ␣ 1B AR was expressed in COS-1 cell (5) and partially purified in the presence of phentolamine by chromatography using heparin-agarose and wheat germ agglutinin-agarose as described (29). Contamination of TGII as well as other GTP-binding proteins was determined by measurement of TGase activity, direct photolabeling of GTPases with [␣-32 P]GTP (28), and immunoblotting with antibodies against TGII and G q/11␣ (5). G␣ q (the virus was kindly provided by Dr. Elliott Ross at Northern Texas University, Dallas, TX) was expressed in Sf9 cells. Since we have found that Sf9 cells do not express TGII as judged by immunoblotting and measurement of TGase activity, G␣ q was partially purified by one-step chromatography using Mono Q-Sepharose. Membrane (5 mg/ml) prepared from G␣ q -expressed Sf9 cells was extracted with 1% sodium cholate in 50 mM HEPES, pH 7.4, 50 mM NaCl, 3 mM 1,4-dithiothreitol, 3 mM EGTA, 1 mM EDTA, 5 mM MgCl 2 , 10 mM NaF, 30 M AlCl 3 as described (24,27). The extract was diluted 10-fold with the same buffer containing 0.02% sucrose monolaurate (SM) and loaded onto Mono Q-Sepharose, which was pre-equilibrated with the same buffer containing 0.02% SM. The column was washed with the buffer and eluted with 300 mM NaCl and 10% glycerol in the same buffer. The amount of G␣ q was determined by immunoblotting with G q/11␣ antibody. A single band with molecular mass of 42 kDa was detected with the antibody. Known concentrations of TGII were simultaneously immunoblotted to estimate the amounts of G␣ q . The protein preparations were aliquoted and stored at Ϫ80°C until use. For each experiment, the protein preparations in the elution buffer was loaded onto a dried Sephadex G-25 column (3 ml) to remove the salts (29). The dried column was preequilibrated with an assay buffer (25 mM HEPES, pH 7.4, 150 mM NaCl, 0.5 mM 1,4-dithiothreitol, and 1.5 mM MgCl 2 ). The recovery was ϳ40 -50%, as determined by immunoblotting with the TGII and G q/11␣ antibodies or TGase activity measurement for TGII. The recovery of the ␣ 1B AR was ϳ50%, as determined by binding ability of [ 3 H]prazosin (29). In addition, it should be noted that a nonhydrolyzable ATP analogue AppNHp (100 M) was included in the assay buffer throughout the study, since it has been reported that TGII also binds ATP and hydrolyzes it (30).

Materials-Fura
Reconstitution-Throughout the study, the proteins were reconstituted in phospholipid vesicles by a dilution method (31). Briefly, an appropriate amount of proteins was mixed with 0.2 mg of a clear phospholipid suspension (8 mg/ml in 0.2% SM solution) consisting of phosphatidylcholine, phosphatidylethanolamine, and phosphatidylserine (3:1:1, w/w/w). The concentrations of phospholipid mixture were 20 -30 g (final), and SM concentrations were ϳ0.008% (final). The reconstitution mixtures in the assay buffer were preincubated in an ice bath for 40 min throughout the study. For the studies involving ␣ 1B AR (150 pM/tube), the samples were preincubated in the presence of 1 ϫ 10 Ϫ5 M (Ϫ)-epinephrine or 1 ϫ 10 Ϫ4 phentolamine.
Preparation of Radiolabeled Guanine Nucleotide-bound TGII-A complex of [␥-32 P]GTP⅐TGII was prepared by incubating TGII (ϳ50 nM) with 50 M [␥-32 P]GTP (100,000 cpm/nM) in 300 l of the assay buffer. After incubation at room temperature for 20 min, unbound [␥-32 P]GTP and [ 32 P]P i was removed by a dried Sephadex G-25 column which was preequilibrated with the assay buffer. The amounts of [␥-32 P]GTP⅐TGII complex were determined by a nitrocellulose membrane filter assay (29). Commercial [ 3 H]GDP was lyophilized to remove ethanol and then reconstituted with water prior to use. The complex of [ 3 H]GDP⅐TGII was prepared by incubating TGII (50 nM) with [ 3 H]GDP (100 Ci) in 300 -500 l of the assay buffer at room temperature for 30 min. Unbound GDP was not removed since TGII has a low affinity for GDP (28).
Measurement of GTP Hydrolysis-Since we have found that GTP hydrolysis by TGII is temperature-sensitive, the reaction was performed at room temperature. Single turnover GTP hydrolysis was determined with the [␥-32 P]GTP⅐TGII complex (ϳ1 nM/tube) preparation. The complex was mixed with and without 4 nM PLC␦1 or with 4 nM heat-inactivated PLC␦1 (boiled for 20 min) in the assay buffer. At time 0, 100 M cold GTP was added to prevent the rebinding of radiolabeled guanine nucleotide. At the indicated time, the samples were transferred to an ice-water bath, and the amount of [␥-32 P]GTP⅐TGII remaining was determined by the nitrocellulose filter method. A standard GTPase activity was also performed by charcoal absorption method (29). Briefly, vesicles containing proteins were mixed with 2 M GTP plus 3 Ci of [␥-32 P]GTP in the assay buffer in a 100-l final volume. The reaction was performed at room temperature for 20 min and stopped by addition of ice-cold Norit A charcoal suspension (5%, w/v, 900 l) in 50 mM sodium phosphate buffer (pH 7.4). The reaction mixture was centrifuged for 20 min at 4°C, and a 700-l aliquot of the supernatants was withdrawn and recentrifuged under the same conditions. After a second centrifugation, the amount of [ 32 P]P i released in a 500-l aliquot was determined by a ␤-counter. To determine turnover, [ 35 S]GTP␥S binding was performed with the same samples at room temperature for 20 min.

Determination of [ 3 H]GDP Release and GTP␥S
Binding-Both experiments were carried out at 10°C throughout the study. [ 3 H]GDP⅐TGII (ϳ1 nM) was incubated with 4 nM PLC␦1 or Ca 2ϩ -bound PLC-␦1 or heat-inactivated PLC␦1. Ca 2ϩ -bound PLC␦1 was prepared by incubating of the enzyme with 30 M Ca 2ϩ at room temperature for 20 min. At this concentration of Ca 2ϩ , PLC␦1 was fully activated (2). The final concentration of Ca 2ϩ was adjusted to 5 M in the reaction mixtures. At time 0, 2 M GTP␥S was added to the reaction mixtures to prevent the rebinding of radiolabeled GDP. For the GTP␥S binding experiments, the reaction was started by adding 2 M [ 35 S]GTP␥S (1500 cpm/nM, final). Nonspecific binding was determined in the presence of 100 M GTP or in the presence of 1 ϫ 10 Ϫ4 M phentolamine when the ␣ 1B AR was reconstituted with TGII or G␣ q and PLC␦1. The time-dependent experiments were performed by transferring an aliquot (50 l) to the ice-water bath at the indicated time points. The amounts of GTP␥S binding by TGII or G␣ q and the remaining [ 3 H]GDP⅐TGII complex were determined by the nitrocellulose filter method (29).
Transfection and Cell Culture-DNAs (10 g/dish) of TGII and its mutants inserted into a neomycin-resistant selection vector pcDNA3.1-Neo (Invitrogene) were transfected to hamster leiomyosarcoma (DDT1-MF2) using LipofectAMINE method provided by the man-ufacturer (Life Technologies, Inc.). DDT1-MF2 cell has been shown to express the ␣ 1B AR subtype only (32). TGII and its mutant expressed cells were selected using 500 g/ml G418 in Dulbecco's modified Eagle's medium (DMEM) containing 10% heat-inactivated fetal bovine serum and 100 g/ml penicillin, and 100 g/ml streptomycin. After completion of the selection, PLC␦1 DNA (10 g) inserted into a hygromycin selection vector pCEP4 (Invitrogene) was transfected into DDT1-MF2 cells expressing TGII and its mutants. The cells were selected with 500 g/ml hygromycin in a growth medium containing 300 g/ml G418. The established cells were maintained in the growth medium containing 300 g/ml G418 and 300 g/ml hygromycin.
Measurement of [Ca 2ϩ ] i -Cytosolic free Ca 2ϩ concentration ([Ca 2ϩ ] i ) was determined using the fluorescent Ca 2ϩ indicator Fura 2-AM as described by Xu et al. (33). DDT1-MF2 cells (1 ϫ 10 4 cells) were trypsinized and seeded on 35-mm glass culture dishes designed for fluorescence microscopy (Bioptech, Butler, PA). After the cells were incubated overnight in DMEM containing 10% heat-inactivated, the cells were further incubated in a serum-free DMEM for 24 h. The cells were then loaded with Fura 2-AM (2 M) at room temperature for 40 min in Krebs Ringer (KR) buffer (25 mM HEPES, pH 7.4, 125 mM NaCl, 5 mM KCl, 1.2 mM MgCl 2 , 2.5 mM CaCl 2 , 11 mM glucose) containing 0.2% bovine serum albumin. After washing cells three times with KR buffer, the cells were kept in the tissue culture incubator until use. Before measurement of [Ca 2ϩ ] i , the cells were washed three times with Ca 2ϩfree KR buffer containing 1 ϫ 10 Ϫ6 M propranolol and 1 ϫ 10 Ϫ7 M rawalscine. The ␣ 1B AR-mediated [Ca 2ϩ ] i was determined by addition of 1 ϫ 10 Ϫ5 M (Ϫ)-epinephrine (final). Five cells, which were within the beam light, were selected to collect data. The reason for the selection of multiple cells is to minimize the variability of outcome, since expression level of proteins is expected to vary from a cell to a cell. The culture dishes were placed in a temperature-regulated chamber (37°C). Fluorescence ratios were measured by an alternative wavelength time scanning method (dual excitation at 340 and 380 nm, emission at 500 nm). Estimation of [Ca 2ϩ ] i were achieved by comparing the cellular ratio with fluorescence ratios acquired using Fura 2 (free acid) in buffer containing known concentrations of Ca 2ϩ . [Ca 2ϩ ] i was calculated as described by Grynkiewicz et al. (34).

RESULTS AND DISCUSSION
Effects of PLC␦1 on GTPase Function of TGII-To assess whether PLC␦1 modulates GTPase activity of TGII, a single turnover hydrolysis of [␥-32 P]GTP by TGII was determined using a [␥-32 P]GTP⅐TGII preparation with and without PLC␦1 (Fig. 1A). In the absence of PLC␦1, the [␥-32 P]GTP hydrolysis by TGII was observed in a time-dependent manner and reached half-maximal GTP hydrolysis within 8 min. In contrast, when PLC␦1 was present, the [␥-32 P]GTP hydrolysis was extremely slow (ϳ21% hydrolysis for 30-min incubation). The [␥-32 P]GTP hydrolysis was not inhibited by heat-inactivated PLC␦1, indicating that the inhibition is caused by the interaction of PLC␦1 with TGII. The PLC␦1-mediated inhibition of GTP hydrolysis by TGII was further examined using the charcoal absorption method. Equimolar (4 nM) of TGII and PLC␦1 or heat-inactivated PLC-␦1 was mixed and incubated at room temperature for 20 min. The vesicles containing TGII alone or the heatinactivated PLC␦1 produced [ 32 P]P i with a turnover of 1.2-1.5 mol Ϫ1 min-1 . On the other hand, the vesicles containing TGII and PLC␦1 produced less [ 32 P]P i with a turnover of 0.24 mol Ϫ1 min Ϫ1 . This demonstrates that PLC␦1 inhibits GTP hydrolysis by TGII, showing that PLC␦1 is not GAP for TGII.
To determine whether PLC␦1 influences exchange of GDP to GTP by TGII, GDP release from TGII was determined (Fig. 1,  B and C). To evaluate whether Ca 2ϩ -bound or unbound PLC␦1 exhibits this effect on TGII, PLC␦1 preincubated with Ca 2ϩ (Ca 2ϩ -PLC␦1) was also tested. A [ 3 H]GDP⅐TGII complex was reconstituted with various concentrations of PLC␦1. The results revealed that the GDP release from TGII was accelerated as a function of PLC␦1 concentration (Fig. 1B) GEF action of PLC␦1 for TGII was further examined by determining GTP␥S binding to TGII (Fig. 2, A and B). Consistent with the observations that PLC␦1 stimulated GDP release, the GTP␥S binding of TGII was increased as a function of PLC␦1 concentration ( Fig. 2A). At a 1:2 ratio of TGII versus PLC␦1, the GTP␥S binding to TGII reached a plateau. PLC␦1 alone showed no GTP␥S binding activity, indicating that the increased GTP␥S binding is caused by the interaction of TGII with PLC␦1. When the GTP␥S binding by TGII was determined as a function of incubation time with and without PLC␦1, the GTP␥S binding in the presence of PLC␦1 was higher (ϳ3-fold) than TGII alone and reached a maximum within 11 min (Fig.  2B). In the presence of Ca 2ϩ -PLC␦1, the GTP␥S binding was similar to TGII alone, again demonstrating that Ca 2ϩ -PLC␦1 does not stimulate GTP␥S binding to TGII. A slight increase of basal GTP␥S binding by TGII was also observed in the presence of PLC␦1, probably due to interaction of the enzyme with TGII during preincubation.
It has also been shown that conformational changes in TGII modulate TGII activity, which are induced by the activators  (35,36). Thus, Ca 2ϩ -bound TGII can not function as GTPase, and GTP-bound TGII does not exhibit TGase activity. To determine whether PLC␦1 induces GTPase form of TGII, the TGase activity was determined in the presence of various concentrations of PLC␦1 or heat-inactivated PLC␦1 (Fig. 2C). The reaction was started with Ca 2ϩ to prevent the activation of PLC␦1 and TGase of TGII. The results showed that Ca 2ϩmediated TGase activation was inhibited in a concentrationdependent manner by PLC␦1. Heat-inactivated PLC␦1 was unable to inhibit TGase activity, demonstrating that the inhibition of TGase activity is due to the interaction TGII with PLC␦1. In addition, to discern whether the cross-linking of proteins caused the decrease in the enzyme activity, samples treated under the same conditions were subjected to immunoblotting with TGII and PLC␦1 antibodies. Cross-linking of TGII-PLC␦1 or TGII⅐TGII or PLC␦1-PLC␦1 was not observed (data not shown). Taken together, these data clearly show that PLC␦1 is GEF for TGII and that the interaction of TGII with PLC␦1 induces a conformational change in TGII to become GTPase.
PLC␦1 Is a Helper of ␣ 1B AR Function That Stabilizes the GTPase Conformation of TGII-G protein-coupled receptors are GEFs for their cognate G proteins. Since the ␣ 1B AR couples with both TGII and G␣ q (7, 8) but G␣ q does not stimulate PLC␦1 activity (27), we first evaluated the specificity of GEF function of PLC␦1 for TGII. The ␣ 1B AR was reconstituted with either TGII or G␣ q in the presence and absence of PLC␦1, and GTP␥S binding by TGII or G␣ q was determined (Fig. 3A). The results revealed that, although the ␣ 1B AR was able to activate GTP␥S binding to both TGII and G␣ q , GEF activity of PLC␦1 was specific for TGII. Thus, in the presence of PLC␦1, the ␣ 1B AR-mediated GTP␥S binding to TGII was further increased ϳ58%, whereas the level of the receptor-mediated GTP␥S binding to G␣ q did not change. Moreover, PLC␦1 again stimulated GTP␥S binding by TGII, but not by G␣ q . This observation is also consistent with the finding that PLC␦1 is not activated by G␣ q (27). Since GTP hydrolysis by TGII was inhibited in the presence of PLC␦1, we also determined whether the inhibition of GTP hydrolysis occurs in the presence of the ␣ 1B AR (Fig. 3B).
In the presence of PLC␦1, the rate of GTP hydrolysis was inhibited ϳ77% with the samples containing TGII and PLC␦1 and ϳ41% with the samples containing all three components. These results again indicate that PLC␦1 functions as a GTP hydrolysis-inhibiting factor (GHIF) and that there is a stable association of TGII with PLC␦1.
To understand the mechanism of GEF activity of the ␣ 1B AR versus PLC␦1 for TGII, the ␣ 1B AR, TGII, and PLC␦1 were reconstituted, and the GTP␥S binding activity of TGII was assessed under various conditions (Fig. 4). The ␣ 1B AR-mediated GTP␥S binding to TGII was evident, reaching a plateau within 6 min (Fig. 4A). When PLC␦1 was present, the receptormediated GTP␥S binding was further increased (ϳ47% at 2 min) and reached a plateau within 4 min. PLC␦1-mediated GTP␥S binding to TGII was slow compared with ␣ 1B AR-mediated GTP␥S binding in both the presence and absence of PLC␦1. These data indicate that the ␣ 1B AR is the prime GEF for TGII and that PLC␦1 functions secondarily. Although TGII alone showed no GTP␥S binding at time zero, when the receptor and/or PLC␦1 were present, the basal level of GTP␥S binding by TGII was increased. The order of the basal GTP␥S binding was ␣ 1B AR ϩ TGII ϩ PLC␦1 Ͼ ␣ 1B AR ϩ TGII Ͼ TGII ϩ PLC␦1. To further understand the role of PLC␦1, the receptor and TGII were reconstituted with various concentrations of PLC␦1, and GTP␥S binding by TGII was determined at 2 and 4 min (Fig. 4B). At the 2-min time point, GTP␥S binding was increased as a function of PLC␦1 concentration. At the 4-min time point, GTP␥S binding was reached maximum at 1:1 ratio of TGII and PLC␦1, and a further increase in the concentration of PLC␦1 did not increase the ␣ 1B AR-mediated GTP␥S binding, probably due to the limited amounts of TGII. When the TGII concentrations were varied at fixed amounts of PLC␦1, GTP␥S binding by TGII was increased as a function of TGII concentration (Fig. 4C). Although maximum coupling efficacy was observed at 1:1 ratio of TGII and PLC␦1, a further increase of TGII concentration resulted in a decrease in the coupling efficacy. These results indicate that a level of each component governs the activation of GTP binding to TGII and that the ␣ 1B AR and PLC␦1 induce GTPase conformation of TGII in a concerted way. The sequence of conformational changes of TGII would be: basal state of TGII, which can function as GTPase and TGase; the second state, GTPase conformation that is induced by the receptor and can reverse to the basal state; the third state, GTPase conformation induced by the receptor and stabilized by interacting with PLC␦1.
Overexpression of PLC␦1 Enhances [Ca 2ϩ ] by Activation of the ␣ 1B AR-The role of PLC␦1 in facilitating coupling of the ␣ 1B AR with TGII was further investigated using DDT1-MF2 cells stably expressing PLC␦1 without and with wild-type TGII (wtTG) and its mutants (Fig. 5). A TGII mutant ( C-S TG), which lacks TGase activity by mutation of Cys 277 to Ser at TGase active site (37), was utilized to delineate GTPase versus TGase activity of TGII. Moreover, if PLC␦1 acts as a stabilizer of GTPase conformation of TGII through its GEF/GHIF activity, wtTG would provide the same result as C-S TG does. To evaluate a specific interaction among ␣ 1B AR, TGII, and PLC␦1, two TGII mutants were utilized; m3TG in which an ␣ 1B AR interaction site on TGII was mutated (5), and ⌬L656 (⌬␦1TG) in which a PLC␦1 interaction site was deleted (38). Proteins were highly expressed, and the expression levels were comparable with each other (Fig. 5, A and B). It should be noted that a fast mobility of the m3TG on SDS-PAGE was also observed when the mutant was expressed in COS-1 cell (5). The reason is not clearly understood. However, differences in an apparent molecular weight were observed with TGIIs from different species, indicating that the mobility of TGII on SDS-PAGE gel is greatly affected by the primary structure of the enzyme (39). The slow mobility of ⌬␦1TG is expected, because of the deletion of 30 amino acid residues from C terminus (38). The coupling among ␣ 1B AR, TGII, and PLC␦1 was assessed by measuring [Ca 2ϩ ] i in a Ca 2ϩ -free buffer (Fig. 5C). The control cells (vector) transfected with vectors (pcDNA3.1 and pCEP4) displayed an increase in the level of [Ca 2ϩ ] i in response to activation of the ␣ 1B AR with (Ϫ)-epinephrine. The ␣ 1 -agonist-evoked peak increase in [Ca 2ϩ ] i was further increased by ϳ59% when PLC␦1 (vector plus PLC␦1) was expressed, demonstrating that PLC␦1 increases the coupling efficacy of this signaling system. Since the experiments were performed in Ca 2ϩ -free buffer, the increase in [Ca 2ϩ ] i is due to the release of Ca 2ϩ from an intracellular store that is likely mediated by IP 3 formed in response to PLC␦1 activation. Expression of wtTG or C-S TG resulted in an increase in peak [Ca 2ϩ ] i that was ϳ74% and ϳ63% greater than that observed in vector-transfected cells, respectively. The cells coexpressing wtTG and C-S TG with PLC␦1 exhibited ϳ23% increase in peak [Ca 2ϩ ] i compared with wtTG or C-S TG alone. The reason for this limited increase of [Ca 2ϩ ] i is probably due to a limited number of the ␣ 1B AR, since the receptor is the prime GEF in this signaling system (see Fig. 4). The ␣ 1 -agonistmediated Ca 2ϩ release was due to the coupling of TGII with the ␣ 1B AR and PLC␦1, because both m3TG-and ⌬␦1TG-expressing cells showed an increase in peak [Ca 2ϩ ] i , which was ϳ53% less than the cells expressing wtTG or C-S TG. Moreover, the peak [Ca 2ϩ ] i was 20% less than the control cell (vector plus PLC␦1), and coexpression of PLC␦1 with these mutants did not significantly increase [Ca 2ϩ ] i . Although a residual stimulation of Ca 2ϩ release in activation of the ␣ 1B AR is most likely due to the incomplete blocking of the interaction among these three proteins, it is also possible that the increase in [Ca 2ϩ ] i in these cells is due to the coupling of the ␣ 1B AR with other G proteins such as the G q family of proteins. Preincubation of the cells with the ␣ 1 -antagonist prazosin or nonspecific PLC inhibitor U73122 completely abolished the ␣ 1 -agonist-mediated increase in [Ca 2ϩ ] i (data not shown). To assess endoplasmic reticulum Ca 2ϩ content, we treated the cells with thapsigargin (an inhibitor of endoplasmic reticulum Ca 2ϩ pump, which stimulates Ca 2ϩ release) at the end of the experiments (Fig. 5D). The thapsigargin-induced release of [Ca 2ϩ ] i was the lowest in the cells coexpressing of PLC␦1 with wtTG or C-S TG and correlated with the amounts of [Ca 2ϩ ] i release induced by the activation of the ␣ 1B AR with the cells expressing TGII and its mutants without or with PLC␦1. These data further demonstrate that the increase in [Ca 2ϩ ] i is caused by release from the intracellular Ca 2ϩ stores.
To date, an effector protein acting as both GEF and GHIF for a GTPase has not been described in either a heterotrimeric or a monomeric GTPase signaling system. Our studies on the roles of PLC␦1 in regulation of TGII activities reveal that PLC␦1 exhibits GEF and GHIF activities for GTPase function of TGII. Evidence for the GEF function of PLC␦1 is that the FIG. 4. GTP␥S binding to TGII mediated by the ␣ 1B AR in the presence and absence of PLC␦1. A, GTP␥S binding to TGII in the presence and absence of the ␣ 1B AR and/or PLC␦1. Indicated proteins (150 pM ␣ 1B AR, 4 nM TGII, and PLC-␦1) in the figure were reconstituted in phospholipid vesicles. At indicated time, an aliquot (50 l) was transferred to a tube in an ice-water bath. Nonspecific GTP␥S binding was determined in the presence 100 M GTP or 1 ϫ 10 Ϫ4 M phentolamine. Standard error was 7-10% of the specific GTP␥S binding. B, rate of the ␣ 1B AR-mediated GTP␥S binding to TGII in the presence of various concentrations of PLC␦1. The samples were incubated at 10°C for 2 or 4 min. C, effects of TGII level on the coupling efficacy involving ␣ 1B AR and PLC␦1. The samples were incubated at 10°C for 4 min. All experiments were performed three times in duplicate, and the specific GTP␥S binding is shown in the means Ϯ S.E. from one of the representative experiments. enzyme facilitates GDP release from TGII and stimulation of GTP␥S binding (Figs. 1 and 2). The inhibition of TGase activity by PLC␦1 and the GHIF activity of the PLC␦1 suggest that the interaction of PLC␦1 with TGII induces and stabilizes GTPase conformation of TGII. The GEF/GHIF activity of PLC␦1 displays independently from the ␣ 1B AR. However, when the ␣ 1B AR is present, the receptor is the prime GEF (Fig. 4). This conclusion is based on the observations that (i) the ␣ 1B ARmediated GTP␥S binding is not additively enhanced in the presence of PLC␦1, (ii) PLC␦1 increases the rate of GTP␥S binding mediated by the receptor, and (iii) PLC␦1-mediated GTP␥S binding to TGII is slow as compared with that mediated by the ␣ 1B AR.
The observation that overexpression of PLC␦1 results in elevation of the ␣ 1B AR-mediated Ca 2ϩ release from the intracellular Ca 2ϩ stores is consistent with findings that PLC␦1 is the effector in TGII-mediated signaling pathway (2)(3)(4)(5)40). Furthermore, overexpression of wtTG and C-S TG substantially enhances the ␣ 1B AR-mediated Ca 2ϩ release, and the TGII mutants m3TG and ⌬␦1TG greatly reduce the level of the ␣ 1B ARevoked Ca 2ϩ release with or without overexpression of PLC␦1. Interestingly, the increase in [Ca 2ϩ ] i was somewhat small when PLC␦1 was coexpressed with wtTG or C-S TG (Fig. 5C). Although the reason for the limited Ca 2ϩ release is probably due the limited number of cognate receptors, other mechanisms may be involved. Thus, the PLC␦1 activity is positively and negatively regulated by TGII depending on the Ca 2ϩ level, expression level of TGII, and binding of guanine nucleotides (3,11,12). PLC␦1 can also be inhibited by its metabolite IP 3 (13)(14)(15). Since coexpression of TGII with PLC␦1 increases basal IP 3 formation (5), all of these factors would reduce the interaction capability of PLC␦1 with TGII. There were no significant differences in peak [Ca 2ϩ ] i between wtTG-and C-S TG-expressing cells, indicating that at the initiation of coupling of these three molecules, the increased Ca 2ϩ level in cell does not affect the GTP binding by TGII. These results also support the find-ings that Ca 2ϩ -unbound PLC␦1 is GEF for TGII (Figs. 1C and 2B). Our results also indicate that PLC␦1 is catalytically activated by GTP⅐TGII, since the level of the endogenous PLC␦1 is sufficient to increase [Ca 2ϩ ] i maximally when wtTG as well as C-S TG was highly expressed (Fig. 5C).
Effectors such as PLC␤1 and cGMP phosphodiesterase as well as RGS proteins terminate GTPase function to prevent further activation of the effector themselves (19 -26). In contrast, PLC␦1 facilitates the signaling involving the ␣ 1B AR and TGII through its GEF/GHIF activity for TGII when the cognate receptors are present. This novel role of PLC␦1 is probably necessary to promote the production of second messengers and to overcome the nature of its regulation by multiple factors. The coupling efficacy of the ␣ 1B AR with TGII in respect to the activation of the cognate PLC is poor as compared with that with G␣ q (7,8,12). In addition, two mechanisms for the regulation of PLC␦1 activity have been studied intensely: (i) a supporting role of other Ca 2ϩ -mobilizing receptor systems and (ii) an effector role in the receptor⅐TGII coupling system. Our results clearly support the latter mechanism; PLC␦1 is activated at the basal Ca 2ϩ level in an intact cell by the ␣ 1B AR through TGII. Operation of these two mechanisms probably depends on whether the cells express the cognate receptors and TGII. It has been reported that PLC␦1 is activated by the capacitative Ca 2ϩ entry, following bradykinin stimulation in rat pheochromocytoma (PC-12) cell, which does not express TGII (17). In these regards, further study is required to establish physiological relevance. Cell lysates (150 g) were subjected to immunoblotting using a TGII antibody, followed by SDS-PAGE (10% gel). B, cell lysate (150 g) was used to determine PLC␦1 by immunoblotting, followed by SDS-PAGE (10% gel). C, effects of PLC␦1 on the ␣ 1B ARmediated Ca 2ϩ release in cells expressing PLC␦1 with and without wtTG and its mutants. D, thapsigargin-induced Ca 2ϩ release. Concentration of thapsigargin was 5 M, and level of [Ca 2ϩ ] with the vector cells was taken as 100%. The data presented were obtained from the experiments shown in panel C.