Atypical activation of the G protein Gα_q by the oncogenic mutation Q209P

Abstract

The causative role of G protein–coupled receptor (GPCR) pathway mutations in uveal melanoma (UM) has been well-established. Nearly all UMs bear an activating mutation in a GPCR pathway mediated by G proteins of the G_q/11 family, driving tumor initiation and possibly metastatic progression. Thus, targeting this pathway holds therapeutic promise for managing UM. However, direct targeting of oncogenic Gα_q/11 mutants, present in ∼90% of UMs, is complicated by the belief that these mutants structurally resemble active Gα_q/11 WT. This notion is solidly founded on previous studies characterizing Gα mutants in which a conserved catalytic glutamine (Gln-209 in Gα_q) is replaced by leucine, which leads to GTPase function deficiency and constitutive activation. Whereas Q209L accounts for approximately half of GNAQ mutations in UM, Q209P is as frequent as Q209L and also promotes oncogenesis, but has not been characterized at the molecular level. Here, we characterized the biochemical and signaling properties of Gα_q Q209P and found that it is also GTPase-deficient and activates downstream signaling as efficiently as Gα_q Q209L. However, Gα_q Q209P had distinct molecular and functional features, including in the switch II region of Gα_q Q209P, which adopted a conformation different from that of Gα_q Q209L or active WT Gα_q, resulting in altered binding to effectors, Gβγ, and regulators of G-protein signaling (RGS) proteins. Our findings reveal that the molecular properties of Gα_q Q209P are fundamentally different from those in other active Gα_q proteins and could be leveraged as a specific vulnerability for the ∼20% of UMs bearing this mutation.

Introduction

Uveal melanoma (UM)² is the second most frequent melanoma (after cutaneous melanoma). Despite progress in monitoring and detection, survival rates have not improved over the past few decades (1, 2). Regardless of tumor classification, UMs are treated by eye enucleation, plaque radiation, or tumor resection, which result in loss of vision (3, 4). For the ∼50% of UMs that are metastatic (5, 6), these treatments do not reduce the probability of mortality. Patients with metastatic UM have a grim median survival of less than 1 year (15% 5-year survival) (7–9). UM is largely insensitive to chemotherapeutics or immune checkpoint inhibitors that have shown efficacy in cutaneous melanomas (7, 9). This is easily explained by the fact that the oncogenic drivers of UM are different from those of cutaneous melanoma (i.e. ∼90% of UMs are caused by activating mutations in GNAQ or GNA11, the genes encoding the G proteins Gα_q and Gα₁₁, and not by mutations in BRAF) (10–14).

Heterotrimeric G proteins, such as G_q and G₁₁, are composed of a nucleotide-binding Gα subunit and an obligatory Gβγ heterodimer, which form a tight complex in the GDP-bound resting state (15). Upon ligand binding, GPCRs promote their activation by accelerating the exchange of GDP for GTP. In turn, Gα-GTP dissociates from Gβγ, allowing them both to regulate numerous downstream effectors. Cancer-associated mutations in Gα_q and Gα₁₁ affect residue Gln-209 in ∼95% of the cases, whereas mutations in Arg-183 are less frequent (11, 12). These residues are critical for the intrinsic GTPase activity of Gα, and their mutation is known to result in a constitutively active protein (16–18). Active Gα_q/11 can engage with multiple downstream effectors, which trigger various signaling pathways implicated in cell growth and oncogenic behavior. For instance, active Gα_q binds to some PLCβ isoforms, which ultimately promotes activation of the MAPK/ERK pathway via Ca²⁺/diacylglycerol/protein kinase C (19–21). Another major Gα_q-dependent mechanism relies on its direct binding and activation of a subfamily of RhoGEFs (22, 23), such as Trio, which ultimately activate TAZ/YAP to promote oncogenic transformation (24, 25). Recent evidence indicates that nearly all UMs bear one mutation within this GPCR-driven pathway (13, 14, 26). In addition to mutations in GNAQ or GNA11 found in ∼90% of the cases, there are also mutually exclusive mutations in CYSLTR2 (encoding the GPCR cysteinyl leukotriene receptor 2, CysLT2R) (27) and PLCB4 (encoding the G protein effector PLCβ4) (28), which operate directly upstream or downstream, respectively, of Gα_q/11. Interestingly, a similar pattern of mutually exclusive mutations in GNAQ, GNA11, CYSLTR2, and PLCB4 has been reported to occur in leptomeningeal melanocytic tumors (29–31), another type of noncutaneous melanoma that afflicts the central nervous system.

Given the insensitivity of UM to therapies used for other types of melanoma, targeting the signaling mechanisms triggered by Gα_q/11 has been in the limelight for the development of novel therapeutics for this type of cancer (32). Their suitability as targets is supported by many lines of evidence. For example, expression of the active G protein mutants in nontransformed cells is oncogenic (11, 12, 33). Similarly, mouse models in which activated Gα_q or Gα₁₁ are expressed in melanocytes develop metastatic UM (34, 35). Importantly from the standpoint of therapeutic targeting, genetic disruption of mutant Gα_q/11 or downstream signaling effectors in UM cells impairs proliferation and/or tumor growth in mice (24, 25, 36–38). Unfortunately, attempts to pharmacologically target signaling pathways activated downstream of mutant Gα_q/11 have not been successful, even when using multiple drugs in combination (7). A likely explanation for the inefficiency of these approaches in blunting UM is that Gα_q/11 activates a complex network of signaling effectors (14), such that targeting individual nodes of this network is insufficient to achieve therapeutic effects. Thus, direct inhibition of mutant Gα_q/11 may be required to completely inhibit all the network components required to promote UM and achieve sufficient efficacy and therapeutic benefit.

Direct targeting of oncogenic Gα_q/11 mutants is a reasonable therapeutic approach, although it presents challenges, as these mutants are predicted to closely resemble active Gα_q/11 WT. If so, strategies to inhibit mutant Gα_q/11 would be expected to also cause inhibition of Gα_q/11 WT, which could result in undesired side effects related to the major physiological functions of these G proteins (e.g. double Gα_q/Gα₁₁ knockout mice are nonviable (39)). Here, we present evidence that one of the most frequent Gα_q mutants in UM, Q209P, displays properties different from that of active Gα_q WT that could be leveraged to achieve specific targeting at the molecular level. Approximately 40–45% of UMs have mutations in residue Gln-209 of Gα_q, which are split evenly between Q209L and Q209P (11–13). Whereas Gα_q Q209L has been extensively characterized and used as a tool mutant to study G_q signaling for decades, Gα_q Q209P has not been adequately studied. Gα_q Q209P is not only as frequent as Gα_q Q209L in tumors, but it is also the driver mutation of many of the UM cell lines commonly used for cell biological and pharmacological experimentation, such as Mel270, OMM1.3 (also known as OMM2.3), OMM2.2, OMM2.5, and UPMM3 (25, 40). In fact, it has been shown that depletion of Gα_q Q209P and/or signaling components downstream of it from UM cells decreases proliferation and/or tumor growth (24, 36, 37). Motivated by the fundamental gap in knowledge about the molecular properties of Gα_q Q209P, we characterized this mutant by direct comparison with Gα_q Q209L, to discover that while leading to signaling hyperactivation, as expected, it possessed structural features different from other active Gα_q species, including active Gα_q WT and the Gα_q Q209L mutant. From a broader perspective, our findings reveal novel mechanistic insights into how G protein mutants lead to oncogenesis and into their possible suitability as direct targets for pharmacological intervention.

Results and discussion

Binding of signaling effectors to Gα_q Q209P is weaker than to Gα_q Q209L or GTP-bound Gα_q WT

To start characterizing the properties of Gα_q Q209P, we compared its ability to bind effectors with that of Gα_q Q209L or active Gα_q WT. First, we investigated binding to GRK2. GRK2 was the first protein co-crystallized with active Gα_q (41). Based on the atomic resolution structure of this complex, it was proposed, and subsequently confirmed, that GRK2 binding to Gα_q has effector-like properties. We expressed Gα_q WT, Gα_q Q209L, and Gα_q Q209P in HEK293T cells and carried out pulldowns with the RGS homology (RH) domain of GRK2 (aa 45–178) fused to GST. Cell lysis and pulldowns were carried out rapidly (∼1.5 h from lysis to protein complex elution) at 4 °C in a buffer without Mg²⁺ supplemented with EDTA to minimize GTP hydrolysis during the assay. As a positive control, we also included a condition in which lysates of cells expressing Gα_q WT were supplemented with Mg·AlF₄⁻, which mimics the GTP-bound transition state of the G protein that binds with high affinity to effectors. As expected, Gα_q WT loaded with Mg·AlF₄⁻ bound robustly to GRK2, whereas Gα_q WT in the absence of Mg·AlF₄⁻, presumably in its GDP-bound inactive conformation, did not (Fig. 1A). Also, as expected, Gα_q Q209L bound to GRK2 as much as Gα_q WT supplemented with Mg·AlF₄⁻, which is consistent with its predicted constitutive GTP-bound status. Surprisingly, Gα_q Q209P binding to GRK2 was weaker than Gα_q Q209L or active Gα_q WT, although consistently stronger than Gα_q WT in the absence of Mg·AlF₄⁻ (Fig. 1A).

Figure 1.

Gα_q Q209P does not bind to effectors as efficiently as Gα_q Q209L or active Gα_q WT. A, Gα_q Q209P binds to GRK2 less than Gα_q Q209L or active Gα_q WT (+Mg·AlF₄⁻) but more than inactive Gα_q WT (−Mg·AlF₄⁻). Lysates of HEK293T cells transfected with HA-Gα_q (WT or mutants, as indicated) were incubated with GST-GRK2^RH immobilized on GSH-agarose beads in the presence or absence of 10 mm MgCl₂, 30 μm AlCl₃, and 10 mm NaF (Mg·AlF₄⁻) as indicated. Resin-bound proteins (top panels) and aliquots of the lysates (bottom panels) were analyzed by Ponceau S staining and immunoblotting (IB) as indicated. One representative result of at least three independent experiments is shown. B, a helix-turn-helix in either GRK2 (purple), PLCβ3 (cyan) or p63RhoGEF (red) docks onto a groove of active Gα_q (gray) formed between the SwII region (green) and the α3 helix (orange). The structures of Gα_q bound to either GRK2 (PDB code 2BCJ), PLCβ3 (PDB code 3OHM), or p63RhoGEF (PDB code 2RGN) were superimposed using Gα_q, and the protein–protein interfaces are shown enlarged on the left. Only a single Gα_q is shown for clarity. C–E, Gα_q Q209P binds to RhoGEFs p63RhoGEF (C), Trio (D), or kalirin (E) less than Gα_q Q209L or active Gα_q WT (+Mg·AlF₄⁻) but more than inactive Gα_q WT (−Mg·AlF₄⁻). Pulldowns were carried out as in A using GST fusions of the DH/PHext domain of p63RhoGEF (aa 155–493), Trio (aa 1967–2296), or kalirin (aa 1901–2223). One representative result of at least three independent experiments is shown. F, Gα_q Q209P binds to PLCβ3 less than Gα_q Q209L or active Gα_q WT (+Mg·AlF₄⁻) but more than inactive Gα_q WT (−Mg·AlF₄⁻). Lysates of HEK293T cells co-transfected with HA-Gα_q (WT or mutants, as indicated) and FLAG-PLCβ3 were immunoprecipitated (IP) with FLAG antibodies in the presence or absence of 10 mm MgCl₂, 30 μm AlCl₃, and 10 mm NaF (Mg·AlF₄⁻) as indicated. Resin-bound proteins (top panels) and aliquots of the lysates (bottom panels) were analyzed by Ponceau S staining and immunoblotting (IB) as indicated. One representative result of at least three independent experiments is shown. G, binding of Gα_q Q209P to GRK2^RH in cells is diminished compared with Gα_q Q209L. Top left, schematic of the BRET assay used to monitor Gα_q binding to GRK2^RH. Nanoluciferase-fused GRK2^RH (GRK2^RH-Nluc, BRET donor) binds to active but not inactive Gα_q-V (BRET acceptor), which results in high or low BRET, respectively. Bottom left, representative immunoblots of lysates of HEK293T cells used in the BRET experiments shown on the right. Right, HEK293T cells were co-transfected with plasmids encoding GRK2^RH-Nluc and Gα_q-V (WT or mutants, as indicated) and BRET measured 24 h later. Results are mean ± S.E. (error bars) (n = 5; **, p < 0.01, Student's t test).

Prompted by these surprising results, we set out to investigate the binding properties of Gα_q Q209P with other G_q effectors. Gα_q works predominantly on two classes of effectors: a subfamily of PLC isoforms (PLCβs), and a subfamily of RhoGEFs composed of p63RhoGEF (also known as ARHGEF25), Trio (also known as ARHGEF23), and kalirin (also known as ARHGEF24). Both PLCβ3 and p63RhoGEF have been co-crystallized with Gα_q (42–44). The atomic resolution structures of these complexes reveal similarities with the GRK2–Gα_q complex. More specifically, one of the primary contacts of GRK2, p63RhoGEF, and PLCβ3 with Gα_q is mediated by a helix-turn-helix element that docks onto a conserved pocket between the α3 helix and switch II (SwII) region of the G protein (Fig. 1B). Consistent with these structural similarities, we found that the binding pattern of p63RhoGEF to different Gα_q species closely resembled that of GRK2 (i.e. Gα_q Q209P binding to p63RhoGEF (DH/PHext domain (45), aa 155–493) was weaker than that of Gα_q Q209L or Mg·AlF₄⁻-loaded Gα_q WT, although stronger than Gα_q WT in the absence of Mg·AlF₄⁻) (Fig. 1C). Similar results were obtained in pulldowns with the other two members of the p63RhoGEF subfamily, Trio and kalirin (Fig. 1, D and E), and in co-immunoprecipitations with PLCβ3 (Fig. 1F). The defect in Gα_q Q209P binding to effectors was not rescued by the addition of Mg·AlF₄⁻ (Fig. S1), suggesting that the diminished binding is not because the G protein is in a GDP-bound state.

To rule out that the weaker binding observed for Gα_q Q209P was due to the experimental conditions of our biochemical assays in vitro, we carried out bioluminescence resonance energy transfer (BRET) experiments in live cells to monitor the interaction between GRK2 and Gα_q (Fig. 1G). Briefly, the same domain of GRK2 used in the pulldowns above (RH domain) was fused to nanoluciferase to generate a BRET donor molecule (GRK2^RH-Nluc). The BRET acceptor consisted of a previously validated Gα_q construct internally tagged with Venus (Gα_q-V) (46). When co-expressed with GRK2^RH-Nluc in HEK293T cells, Gα_q-V Q209L led to a higher BRET signal than Gα_q-V WT, whereas Gα_q-V Q209P did not (Fig. 1G). This result confirms that Gα_q Q209P binding to GRK2 is weaker than Q209L in cells, much like what is observed in the in vitro pulldown experiments. In contrast to the results in pulldown experiments, the BRET assay failed to detect increased binding of Gα_q Q209P compared with Gα_q WT (Fig. 1G). A likely explanation for this is that nonspecific BRET due to random collision of donor and acceptor molecules restricted to the plane of a lipid membrane masks small signal increases due to weak interactions. Taken together, these findings demonstrate that the effector-binding properties of Gα_q Q209P are different from those of other active Gα_q proteins, showing diminished binding compared with GTP-bound Gα_q WT or the constitutively active mutant Q209L.

Gα_q Q209P activates downstream signaling in cells as efficiently as Gα_q Q209L

The findings above are puzzling because previous evidence indicates that G_q-dependent signaling in UM cells bearing the Q209P mutation is required to drive cell proliferation and/or tumor growth in mice. More specifically, it has been demonstrated that genetic disruption of G_q, of its signaling effectors like the RhoGEF Trio, or of other downstream components of the signaling cascade, such as YAP or RasGRP3, diminishes the proliferation and/or growth of tumor xenografts of UM cells with Q209P mutation, such as OMM1.3 (also known as OMM2.3) or Mel270 (24, 36, 37). These previous findings argue strongly that the Gα_q Q209P mutant is hyperactive and promotes oncogenic signaling in UM cancer cells.

To further substantiate this point, we investigated the ability of Gα_q Q209P to activate signaling in cells by comparing it side by side with the more thoroughly characterized Gα_q Q209L mutant. First, we looked at MAPK/ERK activation, which occurs downstream of G_q-mediated activation of PLC. We found that expression of Gα_q Q209L or Gα_q Q209P in HEK293T cells led to similar increases in phospho-ERK1/2 compared with cells expressing Gα_q WT or a vector control (Fig. 2A). Next, we investigated whether Gα_q Q209P was also capable of activating a different G_q-mediated pathway triggered by activation of RhoGEFs, as it has been previously shown that UM cells bearing the Q209P mutation also rely on this pathway to maintain their oncogenic properties (24, 37). For this, we used a luminescent reporter based on the serum response element (SRE) that monitors RhoGEF-dependent activity downstream of active G_q (42). We found that both Gα_q Q209L and Gα_q Q209P led to a robust and similar increase of SRE reporter activity compared with Gα_q WT or a vector control when expressed in HEK293T cells (Fig. 2B). One possible explanation for this result is that the signaling response is saturated at high levels of active Gα_q expression, thereby masking possible differences between the two mutants. However, we found that this is not the case because expression of lower amounts of Gα_q Q209L and Gα_q Q209P led to smaller increases of SRE reporter activity, but they were still similar for the two G protein mutants (Fig. S2). Taken together, these results demonstrate that Gα_q Q209P is constitutively active, leading to a hyperactivation of downstream signaling comparable with that observed for the other oncogenic mutant Q209L.

Figure 2.

Gα_q Q209P activates downstream signaling in cells like Gα_q Q209L. A, expression of Gα_q Q209L or Gα_q Q209P leads to similar levels of ERK1/2 phosphorylation. Lysates of HEK293T cells transfected with HA-Gα_q (WT or mutants, as indicated) were immunoblotted (IB) with the indicated antibodies. One representative result of at least three independent experiments is shown. B, expression of Gα_q Q209L or Gα_q Q209P leads to similar levels of SRE reporter activation. HEK293T cells were co-transfected with HA-Gα_q (WT or mutants, as indicated) and a firefly luciferase reporter driven by the SRE.L promoter and cultured overnight in medium with a low concentration of serum (0.5% FBS) before measuring luminescence. Results are mean ± S.E. (error bars) (n = 4).

Gα_q Q209P has impaired GTPase activity in vitro like Gα_q Q209L

To start dissecting the possible causes of the apparent discrepancy between signaling activity (Fig. 2, A and B) and effector binding (Fig. 1) observed when comparing the Q209P and Q209L mutants, we investigated their ability to hydrolyze nucleotides. Gln-209 in Gα_q corresponds to a residue with a conserved function in G protein-mediated GTP hydrolysis, and the oncogenicity of mutations at this residue is generally ascribed to constitutive G protein activity due to GTPase deficiency (16, 17). Whereas prior research indicates that Gα_q Q209L is GTPase-deficient, the GTPase function of Gα_q Q209P has not been studied before. To directly investigate GTPase activity in vitro, we purified a novel Gα_q variant, hereafter named Gα_q*, that expresses well in Escherichia coli. Gα_q* is a Gα_q/Gα_i1 chimera in which some residues of Gα_q not involved in interacting with G_q-specific binding partners or in nucleotide binding/hydrolysis have been replaced by the corresponding residues in Gα_i1 (Fig. S3A). His-tagged Gα_q* purified from E. coli binds GTPγS with a level of nucleotide occupancy similar to that previously reported for Gα_q purified from insect cells (47) and also binds robustly to several known Gα_q binding partners, such as effectors p63RhoGEF and GRK2, the GEF/chaperone Ric-8A, or the GAP GAIP (also known as RGS19), with the expected activation state dependence (Fig. S3). These results validate that His-Gα_q* recapitulates many of the properties of Gα_q. Next, we compared the GTPase activity of Gα_q* Q209P with that of Gα_q* WT and Gα_q* Q209L. Because the slow rate of nucleotide exchange of Gα_q precludes the efficient loading of nucleotide required to carry out GTPase assays under single-turnover conditions (48), we investigated the GTPase activity under multiple-turnover conditions (i.e. at steady state). As expected, Gα_q* Q209L displayed a lower rate of GTP hydrolysis than Gα_q* WT (Fig. 3). Moreover, Gα_q* Q209P also had diminished GTPase activity, which was comparable with that observed in Gα_q* Q209L (Fig. 3). These results are consistent with Gα_q* Q209P having a defect in GTPase activity comparable with that of Gα_q* Q209L.

Figure 3.

Gα_q Q209P has diminished GTPase activity. Purified Gα_q* Q209L or Gα_q* Q209P has similarly diminished GTPase activity compared with Gα_q* WT. Left, the steady-state GTPase activity of purified Gα_q* WT (black), Q209L (blue), or Q209P (red) was determined by measuring the production of [³²P]P_i at different times. Results are mean ± S.E. (error bars), n = 3. Right, Coomassie Blue–stained gel of an aliquot of the proteins used in one of the GTPase experiments.

The switch II region of Gα_q Q209P adopts a conformation different from other active Gα_q proteins

The results presented so far indicate that although Gα_q Q209P is GTPase-deficient and leads to signaling hyperactivation like Gα_q Q209L, it fails to recapitulate the strong binding to effectors observed for other active Gα_q proteins. We reasoned that whereas mutation of Gln-209 to any residue (including leucine) might be deleterious for GTPase activity, mutation to proline in particular might have a unique impact on the shape of the effector binding site and thereby account for the observed differences in protein–protein binding. There is a strong rationale for this reasoning based on available structural information. First, Gln-209 is located at the end of the SwII region (Fig. 4A). As its name indicates, the SwII alternates between different conformations, depending on the G protein activation status (49). Upon activation, the SwII region adopts a helical conformation that allows high-affinity binding of effectors to a newly formed pocket between SwII and α3 helix (Fig. 4A). Thus, we hypothesized that mutation of Gln-209 to proline, an amino acid with a cyclical side chain that imposes rotational constraints, might affect the conformation of the SwII region and distort the effector-binding site. To test this hypothesis, we carried out a well-established trypsin proteolysis assay that reports the conformation of the SwII (50). Briefly, when SwII adopts a helical conformation like that observed in active, GTP-bound Gα subunits, it becomes insensitive to trypsin digestion. This results in the formation of a trypsin-resistant species in which only an N-terminal fragment of the protein is cleaved off by trypsin, whereas G proteins in which the SwII region does not adopt such helical conformation, like GDP-bound Gα, are readily digested to low-molecular weight products (Fig. 4B). As expected, when lysates of HEK293T cells expressing Gα_q WT were incubated with trypsin, the G protein was readily digested, whereas supplementing the reaction with Mg·AlF₄⁻ to mimic the GTP-bound state resulted in the formation of a trypsin-resistant species (Fig. 4C). Trypsinization of lysates of cells expressing Gα_q Q209L also resulted in the formation of the trypsin-resistant species, consistent with the idea that this mutant is constitutively bound to GTP and mimics the canonical active conformation of Gα_q WT (Fig. 4C). In contrast, the Gα_q Q209P mutant was not trypsin-resistant under the same conditions (Fig. 4C), indicating that its SwII does not adopt the helical conformation that confers trypsin resistance to active Gα_q WT. Because SwII is one of the structural elements of Gα_q that makes direct contact with effectors and shapes the effector binding pocket, these findings help explain the diminished binding of effectors to Gα_q Q209P mutant compared with Gα_q Q209L or active Gα_q WT (Fig. 1).

Figure 4.

The switch II region of Gα_q Q209P adopts a conformation different from other active Gα_q proteins. A, Gln-209 is located at the end of the helix formed by the SwII in active Gα_q that is involved in binding effectors. Left panels, surface view of the structure of active Gα_q with the SwII in green. The region in the box is shown enlarged on the right as a ribbon representation. Green, SwII; red, Gln-209; blue, GDP-AlF₄⁻. Right panels, surface view of the structure of Gα_q with the p63RhoGEF or PLCβ3 binding regions in cyan or purple, respectively, which overlap with the SwII. Red circle, position of Gln-209. B, diagram of limited proteolysis assay used to assess the conformational status of the SwII. The SwII becomes trypsin-resistant when it adopts a helical conformation, such as in active Gα_q WT, which results in the formation of a trypsin-resistant fragment of Gα in which only an N-terminal fragment of the protein has been cleaved off by trypsin. Mock gel depicts potential outcomes, depending on SwII conformation. C, the SwII of Gα_q Q209L but not of Gα_q Q209P adopts a conformation equivalent to that of active Gα_q WT (+Mg·AlF₄⁻) as determined by limited proteolysis with trypsin. Lysates of HEK293T cells transfected with HA-Gα_q (WT or mutants, as indicated) were incubated with trypsin in the presence or absence of 10 mm MgCl₂, 30 μm AlCl₃, and 10 mm NaF (Mg·AlF₄⁻) as indicated. The four lanes on the left are controls not treated with trypsin. The arrowhead indicates the position of full-length Gα_q, and the asterisk indicates the position of the N-terminally cleaved trypsin-resistant fragment of active Gα_q. One representative result of at least three independent experiments is shown.

Gα_q Q209P is constitutively dissociated from Gβγ in cells

A possible interpretation for the results obtained in the trypsin protection experiments described above is that Gα_q Q209P adopts a conformation analogous to that of GDP-bound Gα_q. This is unlikely because Gα_q Q209P binds to effectors better than inactive Gα_q (Fig. 1) and can also trigger signaling in cells (Fig. 2), which is largely incompatible with the properties of GDP-bound Gα. Moreover, Gα_q Q209P did not become trypsin-resistant upon the addition of Mg·AlF₄⁻ (Fig. S4), suggesting that the trypsin sensitivity is not because the G protein is in a GDP-bound state. Instead, it is more likely that the SwII of Gα_q Q209P adopts a unique conformation that represents neither the active nor the inactive conformation of Gα_q WT. To further substantiate this idea, we investigated the association of Gα_q Q209P with Gβγ. Gβγ binds preferentially to Gα subunits in the GDP-bound state by making extensive contacts with the SwII (Fig. 5A). For this, the SwII must adopt an inactive conformation, as formation of the characteristic helical conformation of GTP-bound Gα leads to Gβγ dissociation. Thus, Gβγ binding is sensitive to the conformational status of the SwII region of Gα. To investigate the association between Gα_q and Gβγ in cells, we used a previously described BRET-based assay (51, 52). Briefly, a fusion of the C-terminal domain of GRK3 fused to nanoluciferase (GRK3ct-Nluc) is used as the BRET donor, and Gβγ fused to Venus (V-Gβγ) serves as the BRET acceptor. Because GRK3ct and Gα_q share overlapping binding sites on Gβγ and can bind only in a mutually exclusive manner, BRET due to V-Gβγ binding to GRK3ct-Nluc reflects the dissociation of Gα–Gβγ heterotrimers (Fig. 5B, top).

Figure 5.

Gα_q Q209P is constitutively dissociated from Gβγ in cells and binds weakly to RGS proteins. A, Gβγ binds to the SwII region of Gα_q. Top, surface view of the structure of Gα_q with the SwII in green. Bottom, surface view of the structure of Gα_q with the Gβγ binding area in blue. Red circle, position of Gln-209. B, Gα_q Q209L and Gα_q Q209P are both constitutively dissociated from Gβγ, as observed for GPCR-activated Gα_q WT. Top, schematic of the assay used to monitor G protein activity with a BRET reporter of free Gβγ. Under resting conditions, Venus-tagged Gβγ (V-Gβγ, BRET acceptor) associates with inactive Gα_q, and BRET signal is low. Upon GPCR stimulation, V-Gβγ dissociates from active Gα and interacts with the C-terminal domain of GRK3 fused to nanoluciferase (GRK3ct-Nluc, BRET donor) inducing a BRET increase. Bottom, Gβγ dissociates from Gα_q WT upon GPCR stimulation, whereas Gα_q Q209L and Gα_q Q209P fail to associate efficiently with Gβγ under resting conditions, as indicated by the high BRET before agonist stimulation, and fail to undergo further dissociation upon GPCR stimulation. HEK293T cells transfected with plasmids encoding M3R, Venus(1–155)-Gγ₂ (VN-Gγ₂) and Venus(155–239)-Gβ₁ (VC-Gβ₁), HA-Gα_q (WT or mutants), and GRK3ct-Nluc were treated with carbachol (100 μm) and atropine (100 μm) as indicated. Twenty-four h after transfection, cells were harvested, and BRET was measured every 0.24 s. Results are mean ± S.E. (error bars) (shown only at 5-s intervals for clarity) of n = 3. Immunoblots of lysates of HEK293T cells, used to confirm the equal expression of Gα_q proteins, are shown below the graph. C and D, Gα_q Q209P binds to GAIP (C) or RGS2 (D) less than Gα_q Q209L or active Gα_q WT (+Mg·AlF₄⁻). Left panels, surface view of the structure of Gα_q with the RGS8 (C) or RGS2 (D) binding area in yellow or purple, respectively. Red circle, position of Gln-209. Right panels, lysates of HEK293T cells transfected with HA-Gα_q (WT or mutants, as indicated) were incubated with GST-GAIP (C) or GST RGS2 (D) immobilized on GSH-agarose beads in the presence or absence of 10 mm MgCl₂, 30 μm AlCl₃, and 10 mm NaF (Mg·AlF₄⁻) as indicated. Resin-bound proteins (Pulldown) and aliquots of the lysates were analyzed by Ponceau S staining and immunoblotting (IB) as indicated. One representative result of three independent experiments is shown.

Consistent with Gα–Gβγ heterotrimer dissociation upon G protein activation and previous observations using this assay system (51–53), agonist-induced activation of the G_q-coupled GPCR M3 muscarinic acetylcholine receptor (M3R) led to a rapid increase in BRET in HEK293T cells co-expressing the donor/acceptor pair and Gα_q WT. The subsequent addition of atropine, an antagonist of the M3 muscarinic receptor, caused the rapid decline of the BRET signal to basal levels, consistent with reassociation of Gα_q and V-Gβγ (Fig. 5B). When the same experiment was done in cells expressing Gα_q Q209L or Gα_q Q209P, the basal levels of BRET before agonist stimulation were as high as that observed for agonist-stimulated BRET in cells expressing Gα_q WT (Fig. 5B), indicating that Gα_q Q209L and Gα_q Q209P are constitutively dissociated from Gβγ. Consistent with constitutive dissociation of Gα and Gβγ, GPCR stimulation of cells expressing Gα_q Q209L and Gα_q Q209P failed to elicit a BRET response (Fig. 5B). The amount of Gα_q WT, Gα_q Q209L, and Gα_q Q209P in the cells used in these experiments was equal, ruling out the possibility that the BRET differences observed between Gα_q WT and the mutants were due to differential expression (Fig. 5B). There are two important points to be drawn from these observations. One is that the SwII of Gα_q Q209P adopts a unique conformation different from that of inactive Gα_q WT, as it cannot bind to Gβγ (Fig. 5B), and from that of other active Gα proteins, as it does not become trypsin-resistant (Fig. 4). The second point is that Gα_q Q209P exists preferentially in a monomeric, Gβγ-free form in cells.

Binding of RGS proteins to Gα_q Q209P is weaker than to Gα_q Q209L

We reasoned that binding of Gα_q Q209P to RGS proteins might be impaired because they also utilize the SwII as an important contact point. In fact, crystal structures of RGS8 and RGS2 in complex with Gα_q (54, 55) have revealed that, despite adopting different poses, both RGS proteins make contact with Gln-209 and several adjacent residues in the SwII (Fig. 5, C and D). Consistent with our expectation, we found that both GAIP (also known as RGS19) and RGS2, representing the two G protein binding poses of RGS proteins (54), bound to Gα_q Q209P less than to Gα_q Q209L or Gα_q WT activated with Mg·AlF₄⁻, but slightly more than inactive Gα_q WT (Fig. 5, C and D). Once again, adding Mg·AlF₄⁻ did not rescue the defect in RGS binding of Gα_q Q209P (Fig. S5), suggesting that the diminished binding is not because the G protein is in a GDP-bound state. These results indicate that Gα_q Q209P does not bind to RGS proteins, a family of negative regulators of G_q signaling.

Mechanistic model and discussion

Our findings using in vitro and cell-based reconstitution systems reveal that the oncogenic Gα_q Q209P mutant is an active G protein with atypical properties. Although we can only speculate about the mechanism by which this mutant leads to signaling hyperactivation in the context of uveal melanoma, our findings provide a framework to try to explain how two mutants with different biochemical properties, Gα_q Q209L and Gα_q Q209P, result in similar oncogenicity (Fig. 6). Both Gα_q Q209L and Gα_q Q209P are GTPase-deficient and trigger constitutive activation of downstream signaling, which explains well their ability to promote cell phenotypic traits that support oncogenic transformation. The fact that the frequency of Q209P mutations in uveal melanoma tumors is similar to that of Q209L mutations supports the idea that both of them provide a similar adaptive advantage during cancer development through abnormal signaling hyperactivation. This is in good agreement with previous observations indicating that both Gα_q Q209L- and Gα_q Q209P-dependent signaling in UM-derived cell lines are required for cell proliferation and/or tumor growth in mice (24, 25, 36–38). The main difference between Gα_q Q209P and other active Gα_q species (i.e. Gα_q Q209L and GTP-bound Gα_q WT) is that it has unique structural properties that impact its ability to bind different interacting partners. More specifically, our results show that the SwII of Gα_q Q209P adopts a conformation different from that of Gα_q Q209L or Gα_q WT that results in diminished binding to effectors as well as to Gβγ or RGS proteins, the two main protein classes that function as negative regulators of Gα_q activity in cells. It is unlikely that the pro-oncogenic properties of Gα_q Q209P arise from engagement to an effector different from those investigated in Fig. 1 for two reasons. One is that the effectors investigated in Fig. 1 represent not only the most widely and best characterized direct effectors of Gα_q, but also the ones that lead to pro-oncogenic signaling in UM cell lines (24, 25, 36, 37). The second one is that the mode of engagement of effectors on Gα subunits is highly conserved, invariably involving the α3 helix/SwII groove (43, 56). This is consistent with our finding that five different effectors engage similarly with Gα_q Q209P (Fig. 1), making it very unlikely that another putative effector would engage Gα_q Q209P in a completely different manner.

Figure 6.

Working model for the mechanisms leading to the constitutive signaling activity of Gα_q Q209L and Gα_q Q209P in cancer. In the absence of GPCR stimulation, the low signaling activity of Gα_q WT (top) is ensured by two mechanisms: 1) low affinity for effectors (blue arrow) and 2) the action of negative regulators, such as Gβγ and RGS protein, which diminishes coupling to effectors. In Gα_q Q209L (middle), the SwII (dark green) adopts an active conformation equivalent to that of GTP-bound Gα_q that causes 1) high-affinity binding to effectors (red arrow) and 2) dissociation from Gβγ, which lead to constitutive signaling activity. Nevertheless, this mutant still binds with high affinity to RGS proteins, which are potent inhibitors of effector binding. In Gα_q Q209P (bottom), the SwII (dark green) adopts a conformation different from that of Gα_q Q209L or GTP-bound Gα_q that causes 1) moderate affinity binding to effectors (orange arrow), 2) dissociation from Gβγ, and 3) impaired binding to RGS proteins. We speculate that the moderate affinity binding to effectors combined with the loss of Gβγ-mediated blockade of the effector-binding site and the lack of RGS-mediated antagonism for effector binding might be sufficient to account for the signaling hyperactivation reported in cells bearing this mutation (11, 24, 25, 37).

Then, how can Gα_q Q209P lead to hyperactive signaling and oncogenic transformation if its binding to effectors is diminished compared with GTP-bound Gα_q WT? First, Gα_q Q209P binding to effectors is stronger than that of inactive Gα_q WT, so it still retains some ability to engage and activate effectors. Second, Gα_q Q209P is constitutively dissociated from Gβγ and has diminished binding to RGS proteins, both of which are negative regulators of Gα_q in cells. A frequently overlooked function of Gβγ is to prevent the spurious action of Gα on its effectors by competitive binding (57, 58). The importance of this function is further highlighted by evidence that Gβγ can bind to GTP-bound Gα_q at physiological concentrations and antagonize binding of effectors such as PLCβ (48, 59). The role of RGS proteins as negative regulators of G protein signaling is well-documented. This function is not only mediated by their GAP activity, to which GTPase-deficient Gln-209 mutants are presumably insensitive (60), but also by their ability to antagonize the binding of effectors to G proteins. In fact, it has been reported that RGS proteins are potent inhibitors of signaling activation by the Gα_q Q209L oncogenic mutant (61, 62). Thus, the moderately higher affinity of Gα_q Q209P for effectors compared with inactive Gα_q might work concurrently with the relief from Gβγ- and RGS-mediated antagonism to attain effective levels of downstream signaling (Fig. 6).

This model shares similarities with the recently proposed mode of action for another Gα oncogenic mutant (i.e. Gα_s R201C). Hu and Shokat (63) have recently reported that this mutant might exist in a GDP-bound yet active state that accounts for its pro-oncogenic properties in cells. In the presence of Gβγ, only GDP-bound Gα_s R201C and not GDP-bound Gα_s WT can activate a downstream effector (i.e. adenylyl cyclase) (63). Thus, although GDP-bound Gα_s R201C is a weaker activator of adenylyl cyclase than GTP-bound Gα_s, it leads to efficient signaling due to its inability to bind Gβγ. We propose that insensitivity to Gβγ antagonism might contribute similarly to enhancing Gα_q Q209P–dependent signaling.

In summary, our findings reveal a fundamental difference in the molecular properties of Gα_q Q209P compared with other active Gα_q proteins, including the other most frequent Gα_q mutation in uveal melanoma, Q209L. The unique structural features of Gα_q Q209P could be leveraged as a specific vulnerability of uveal melanomas bearing this mutation, which account for ∼20% (11–13), and to overcome current limitations for the treatment of this type of cancer. To date, targeting signaling pathways downstream of G_q in uveal melanoma has not been effective for therapeutic purposes (63). On the other hand, although it is logical to think that targeting mutant Gα_q directly might be more efficacious than targeting downstream signaling nodes, it has been an area underexplored due to the assumption that it could lead to concurrent blockade of Gα_q WT function and subsequent undesired side effects. Thus, the safety of cell-permeable peptide-like inhibitors of G_q recently shown to effectively blunt proliferation of uveal melanoma cells bearing activating mutations in Gα_q remains to be established (64). However, we show that contrary to other mutants like Gα_q Q209L, Gα_q Q209P could be specifically inhibited without affecting Gα_q WT if targeting the unique structural features of the mutant proves to be feasible. Based on our results, it is likely that the α3 helix/SwII groove of Gα_q in the Q209P mutant is different than in other active Gα_q species. Identifying molecules that specifically bind to this putative pocket would disrupt the binding and activation of effectors downstream of the Q209P mutant. Although disruption of protein–protein interactions is challenging, there are now dozens of different protein–protein interactions that have been targeted by small molecules, and many of them have shown promising therapeutic effects, even entering clinical trials (65–69). An atomic resolution structure of Gα_q Q209P might provide additional information on the molecular basis for its unique properties while establishing a framework to explore its druggability.

Experimental procedures

Reagents and antibodies

Unless otherwise indicated, all chemical reagents were obtained from Sigma or Fisher Scientific. E. coli DH5α strain was purchased from New England Biolabs, and the BL21(DE3) strain was purchased from Life Technologies. PfuUltra DNA polymerase was purchased from Agilent. Carbachol (catalog no. AC-10824) was obtained from Acros Organics, and atropine (catalog no. A10236) was from Alfa Aesar. Leupeptin (catalog no. L-010), pepstatin A (catalog no. P-020), and aprotinin (catalog no. A-655) were from Gold Biotechnology. [γ-³²P]GTP was from PerkinElmer Life Sciences. Mouse mAb raised against hemagglutinin (HA) tag (clone 12CA5) was obtained from Roche Applied Science. Mouse monoclonal antibodies raised against α-tubulin (T6074) and FLAG tag (F1804) were from Sigma. Rabbit polyclonal antibody raised against nanoluciferase (Nluc) was kindly provided by Dr. Lance Encell (Promega). Rabbit polyclonal antibodies raised against Gα_q (E-17) were purchased from Santa Cruz Biotechnology, Inc. Rabbit antibodies for phospho-ERK1/2 (Thr-202/Tyr-204) (catalog no. 4370) and total ERK1/2 (catalog no. 9102) were obtained from Cell Signaling. Goat anti-rabbit Alexa Fluor 680 and goat anti-mouse or IRDye 800 secondary antibodies were from Life Technologies and LI-COR, respectively.

Plasmids

pcDNA3.1-Gα_q-Venus (Gα_q-V, Venus inserted into the b/c loop of Gα_q (46)), pcDNA3.1-Venus(1–155)-Gγ₂ (VN-Gγ₂), and pcDNA3.1-Venus(155–239)-Gβ₁ (VC-Gβ₁) were kindly provided by N. Lambert (Augusta University, Augusta, GA) (51). pcDNA3.1-masGRK3ct-Nluc (52, 70) was a gift from K. Martemyanov (Scripps Research Institute, Jupiter, FL). pcDNA3.1-FLAG-PLCβ3 was a kind gift from T. Filtz (Oregon State University). pcDNA3-Gα_q-HA (internally tagged) was kindly provided by P. Wedegaertner (Thomas Jefferson University) (71). The plasmid encoding human M3R 3×HA-tagged (N terminus, pcDNA3.1–3×HA-M3R, catalog no. MAR030TN00) was purchased from the cDNA Resource Center (Bloomsburg University). pGL3-SRE.L and pRL-TK plasmids were a present from R. Neubig (Michigan State University). The plasmid construct for the expression of the RH domain of GRK2 (GRK2^RH) fused to Nluc (pcDNA3.1-masGRK2^RH-Nluc) was generated by PCR amplification of residues 45–178 of bovine GRK2 and ligation into the HindIII/BamHI sites of pcDNA3.1-masGRK3ct-Nluc, which results in the replacement of GRK3ct by GRK2^RH. The plasmid used for the bacterial expression of GST-GRK2^RH (pGEX-2T-GRK2(45–178)) was kindly provided by P. Wedegaertner (Thomas Jefferson University) (72). Gα_q* sequence was cloned into the pET28a plasmid for the expression of His-Gα_q* in bacteria. A plasmid for the expression of GST-p63RhoGEF^DH/PHext(155–493) in bacteria (pLIC-GST-p63RhoGEF (residues 155–493, DH/PHext domain)) and the empty pLIC-GST plasmid (a ligation-independent cloning (LIC) plasmid) derived from pMCSG7 (73) were kindly provided by J. Sondek (University of North Carolina, Chapel Hill, NC) (45). Plasmids for the expression of GST-Trio^DH/PHext (residues 1967–2296, DH/PHext domain), GST-kalirin^DH/PHext (residues 1901–2223, DH/PHext domain), and GST-RGS2 (residues 72–203, RGS box) in bacteria were generated by PCR amplification and insertion into pLIC-GST via a LIC procedure (74). Cloning of the plasmid encoding GST-GAIP (rat, pGEX-KG-GAIP) has been described previously (75). All point mutations were generated by following the manufacturer's instructions (QuikChange II, Agilent).

Protein expression and purification

GST-GRK2^RH, GST-p63RhoGEF^DH/PHext, GST-Trio^DH/PHext, GST-kalirin^DH/PHext, GST-GAIP, and GST-RGS2 were expressed in BL21(DE3) E. coli transformed with the corresponding plasmids by overnight induction at 23 °C with 1 mm isopropyl-β-d-1-thio-galactopyranoside. GST-Ric-8A (aa 12–412) protein induction was carried out as described previously (76). Protein purification was carried out following protocols described previously (77). Briefly, bacteria pelleted from 1 liter of culture were resuspended in 25 ml of buffer (50 mm NaH₂PO₄, pH 7.4, 300 mm NaCl, 10 mm imidazole, 1% (v/v) Triton X-100 supplemented with protease inhibitor mixture (1 μm leupeptin, 2.5 μm pepstatin, 0.2 μm aprotinin, 1 mm phenylmethylsulfonyl fluoride)). After sonication (four 20-s sonication cycles with 1-min intervals), lysates were centrifuged at 12,000 × g for 20 min at 4 °C. The soluble fraction (supernatant) of the lysate was used for affinity purification on GSH-agarose resin (Pierce) and eluted with 50 mm Tris-HCl, pH 8, 100 mm NaCl, 30 mm reduced GSH. All purified GST-tagged proteins were dialyzed overnight at 4 °C against PBS. All protein samples were aliquoted and stored at −80 °C.

His-Gα_q* was purified by adapting a protocol described by Waldo et al. (43). His-Gα_q* expression in BL21(DE3) E. coli was induced overnight with 1 mm isopropyl-β-d-1-thio-galactopyranoside at 23 °C. Bacteria were pelleted, lysed by sonication, and cleared from insoluble material as described above but using a different lysis buffer (20 mm HEPES, pH 8.0, 300 mm NaCl, 5 mm MgCl₂, 10 mm β-mercaptoethanol, 15 mm imidazole, 10% glycerol, 50 μm GDP, 30 μm AlCl₃, and 10 mm NaF supplemented with protease inhibitor mixture (1 μm leupeptin, 2.5 μm pepstatin, 0.2 μm aprotinin, 1 mm phenylmethylsulfonyl fluoride)). The soluble fraction of the lysate was incubated with HisPur cobalt resins (Pierce, catalog no. 89964) for 90–120 min at 4 °C with rotation, followed by four cycles of washes with ∼20 resin volumes of lysis buffer and centrifugation (2,000 × g, 2 min), and eluted with lysis buffer supplemented with imidazole to obtain a final concentration of 500 mm. The eluate was supplemented with 100 mm EDTA and incubated on ice for 20 min before passing it over a Superdex 200 HR10/30 gel filtration column using a running buffer consisting of 20 mm HEPES, pH 8.0, 100 mm NaCl, 1 mm MgCl₂, 2 mm DTT, 2% glycerol, and 10 μm GDP. In some cases, proteins were concentrated using an Amicon centrifugal filter with a molecular mass cut-off of 10 kDa. The typical yield was 0.5 mg of protein/liter of culture. Purified His-Gα_q* was aliquoted and stored at −80 °C.

In vitro protein-binding assays with GST-fused proteins

For pulldowns of proteins expressed in mammalian cells, ∼2 × 10⁶ HEK293T cells were seeded on 100-mm dishes and transfected the following day using the calcium phosphate method with plasmids encoding the following constructs (DNA amounts in parenthesis): Gα_q-HA (WT or mutants, 3 μg) or Gα_q-V (WT or mutants, 3 μg). Cell medium was changed 6 h after transfection. Twenty-four h after transfection, cells were lysed on ice with lysis buffer (20 mm Hepes, pH 7.2, 125 mm K(CH₃COO), 0.4% (v/v) Triton X-100, 1 mm DTT, 10 mm β-glycerophosphate, 30 μm GDP, and 0.5 mm Na₃VO₄ supplemented with a protease inhibitor (SigmaFAST, catalog no. S8830)). For some conditions, the lysis buffer was supplemented with 30 μm AlCl₃, 10 mm NaF, and 10 mm MgCl₂ (Mg·AlF₄⁻ condition). The cell lysates were cleared (14,000 × g, 10 min) and used as the source of soluble protein ligands for immobilized GST-fused proteins in pulldown assays. The following GST-fused proteins were immobilized on GSH-agarose beads for 90 min at room temperature in PBS (amounts per condition indicated in parenthesis): GST-GRK2^RH (5 μg), GST-p63RhoGEF^DH/PHext (7.5 μg), GST-Trio^DH/PHext (5 μg), GST-kalirin^DH/PHext (5 μg), GST-GAIP (7.5 μg), and GST-RGS2 (7.5 μg). Beads were washed twice with PBS and resuspended in 400 μl of binding buffer (50 mm Tris-HCl, pH 7.4, 100 mm NaCl, 0.4% (v/v) Nonidet P-40, 5 mm EDTA, 30 μm GDP, 2 mm DTT), which in some cases was supplemented with 30 μm AlCl₃, 10 mm NaF, and 10 mm MgCl₂ (Mg·AlF₄⁻ condition). After the addition of cleared HEK293T cell lysates (∼400 μg of total protein), tubes were incubated for 90 min at 4 °C with constant rotation. Beads were rapidly washed four times with 1 ml of wash buffer (4.3 mm Na₂HPO₄, 1.4 mm KH₂PO₄, pH 7.4, 137 mm NaCl, 2.7 mm KCl, 0.1% (v/v) Tween 20, 30 μm GDP, 5 mm EDTA, 1 mm DTT), which in some cases was supplemented with 30 μm AlCl₃, 10 mm NaF, and 10 mm MgCl₂ (Mg·AlF₄⁻ condition). Resin-bound proteins were eluted with Laemmli sample buffer by incubation at 37 °C for 10 min. Proteins were separated by SDS-PAGE, transferred to PVDF membranes, stained with Ponceau S, and immunoblotted with the corresponding antibodies.

For pulldowns of purified His-Gα_q* (Fig. S1), the following GST-fused proteins were immobilized on GSH-agarose beads by incubation at room temperature for 90 min in PBS (amounts indicated in parenthesis): GST-GRK2^RH (10 μg), GST-GAIP (10 μg), GST-p63RhoGEF^DH/PHext (10 μg), and GST-Ric-8A (10 μg). Beads were washed twice with PBS and resuspended in 300 μl of binding buffer (50 mm Tris-HCl, pH 7.4, 100 mm NaCl, 0.4% (v/v) Nonidet P-40, 5 mm EDTA, 10 mm MgCl₂, 1 mm DTT) supplemented with 30 μm GDP or 30 μm GDP plus 30 μm AlCl₃ and 10 mm NaF (GDP·AlF₄⁻ condition) as indicated. After the addition of 1.7 μg of purified His-Gα_q*, tubes were incubated 4 h at 4 °C with constant rotation. Beads were washed four times with 1 ml of wash buffer (4.3 mm Na₂HPO₄, 1.4 mm KH₂PO₄, pH 7.4, 137 mm NaCl, 2.7 mm KCl, 0.1% (v/v) Tween 20, 10 mm MgCl₂, 5 mm EDTA, 1 mm DTT) supplemented with 30 μm GDP or 30 μm GDP plus 30 μm AlCl₃ and 10 mm NaF (GDP·AlF₄⁻ condition) as indicated. Resin-bound proteins were eluted with Laemmli sample buffer by incubation at 37 °C for 10 min. Proteins were separated by SDS-PAGE, transferred to PVDF membranes, stained with Ponceau S, and immunoblotted with the corresponding antibodies.

Immunoprecipitation

Approximately 2 × 10⁶ HEK293T cells were seeded on 100-mm dishes and transfected the day after using the calcium phosphate method with plasmids encoding Gα_q-HA (WT or mutants, 3 μg) and FLAG-PLCβ3 (6 μg). Cell medium was changed 6 h after transfection. Twenty-four hours after transfection, cells were lysed on ice with 500 μl of lysis buffer (20 mm Hepes, pH 7.2, 125 mm K(CH₃COO), 0.4% (v/v) Triton X-100, 1 mm DTT, 10 mm β-glycerophosphate, 30 μm GDP, and 0.5 mm Na₃VO₄ supplemented with a protease inhibitor (SigmaFAST, catalog no. S8830)). For some conditions, the lysis buffer was supplemented with 30 μm AlCl₃, 10 mm NaF, and 10 mm MgCl₂ (Mg·AlF₄⁻ condition). Cell lysates were cleared (14,000 × g, 10 min) and incubated with 2 μg of anti-FLAG antibodies (Sigma, F1804) immobilized on Protein G–agarose beads (Thermo Scientific, ∼35 μl of beads preblocked with 5% BSA) for 90 min at 4 °C with rotation. Beads were washed three times (4.3 mm Na₂HPO₄, 1.4 mm KH₂PO₄, pH 7.4, 137 mm NaCl, 2.7 mm KCl, 0.1% (v/v) Tween 20, 30 μm GDP, 5 mm EDTA, 1 mm DTT), and proteins were eluted by adding Laemmli sample buffer and boiling for 5 min. For some conditions, the wash buffer was supplemented with 30 μm AlCl₃, 10 mm NaF, and 10 mm MgCl₂ (Mg·AlF₄⁻ condition). Eluted proteins were separated by SDS-PAGE and analyzed by immunoblotting.

BRET

HEK293T cells were grown at 37 °C, 5% CO₂ in Dulbecco's modified Eagle's medium supplemented with 10% FBS, 100 units/ml penicillin, 100 μg/ml streptomycin, and 1% l-glutamine. HEK293T cells were seeded on 6-well plates (∼400,000 cells/well) coated with gelatin and after 1 day transfected using the calcium phosphate method. For the detection of Gα_q-V binding to GRK2^RH-Nluc, the following constructs were used (DNA amounts per well in parenthesis): Gα_q-V (0.5 μg, WT or mutants) and mas-GRK2^RH-Nluc (0.05 μg). For the detection of Gα_q-V association with GRK3ct-Nluc, we followed previously described protocols (53, 78, 79). For these transfections, the following constructs were used (DNA amounts per well in parenthesis): Gα_q (0.5 μg), VC-Gβ₁ (0.2 μg), VN-Gγ₂ (0.2 μg), mas-GRK3ct-Nluc (0.2 μg), and M3R (0.2 μg). Cell medium was changed 6 h after transfection. Approximately 16–24 h after transfection, cells were washed and gently scraped in warm PBS, centrifuged (5 min at 550 × g), and resuspended in Tyrode's solution (140 mm NaCl, 5 mm KCl, 1 mm MgCl₂, 1 mm CaCl₂, 0.37 mm NaH₂PO₄, 24 mm NaHCO₃, 10 mm HEPES, pH 7.4, 0.1% glucose) at a concentration of ∼10⁶ cells/ml. 25,000 cells were added to a white opaque 96-well plate (Opti-Plate, PerkinElmer Life Sciences) and mixed with the nanoluciferase substrate Nano-Glo (Promega, final dilution 1:200) for 2 min before measuring. A POLARstar OMEGA plate reader (BMG Labtech) was used to measure luminescence signals at 460 ± 20 and 528 ± 10 nm at 28 °C, and BRET signals were calculated as the ratio between the emission intensity at 528 ± 10 nm over the emission intensity at 460 ± 20 nm. For the kinetic experiments, BRET measurements were performed every 0.24 s. An aliquot of cells from each experiment was processed for subsequent immunoblot analysis as follows. Cells were centrifuged (1 min at 14,000 × g) and resuspended on ice with lysis buffer (20 mm Hepes, pH 7.2, 5 mm Mg(CH₃COO)₂, 125 mm K(CH₃COO), 0.4% (v/v) Triton X-100, 1 mm DTT, 10 mm β-glycerophosphate, and 0.5 mm Na₃VO₄ supplemented with a protease inhibitor mixture (SigmaFAST, catalog no. S8830)). Lysates were cleared by centrifugation (14,000 × g, 10 min, 4 °C) and boiled for 5 min in Laemmli sample buffer before protein separation by SDS-PAGE and immunoblotting.

ERK1/2 phosphorylation

HEK293T cells were seeded on 6-well plates (∼200,000 cells/well) coated with gelatin and after 1 day were transfected using the calcium phosphate method with plasmids encoding Gα_q-HA WT or mutants (0.5 μg/well) or with an empty vector. Cell medium was changed 6 h after transfection. Approximately 16–24 h after transfection, cells were scraped in PBS, centrifuged (30 s at 10,000 × g), and resuspended on ice with lysis buffer (20 mm Hepes, pH 7.2, 5 mm Mg(CH₃COO)₂, 125 mm K(CH₃COO), 0.4% (v/v) Triton X-100, 1 mm DTT, 10 mm β-glycerophosphate, and 0.5 mm Na₃VO₄ supplemented with a protease inhibitor mixture (SigmaFAST, catalog no. S8830)). Lysates were cleared by centrifugation (14,000 × g, 10 min, 4 °C) and boiled for 5 min in Laemmli sample buffer prior to protein separation by SDS-PAGE and immunoblotting.

SRE reporter activation

These experiments were performed using a luciferase reporter assay as described by Lutz et al. (42) with minor modifications. HEK293T cells were seeded on 6-well plates (∼350,000 cells/well) coated with gelatin and after 1 day were transfected using the calcium phosphate method with plasmids encoding Gα_q-HA (WT or mutants, 1 μg/well in Fig. 2 or 0.1 μg/well in Fig. S2) or with an empty vector, along with the luciferase reporter plasmids pGL3-SRE.L and pRL-TK (0.5 μg of each per well). Cell medium was changed 6 h after transfection by medium containing a reduced concentration of FBS (0.5% (v/v) final). Approximately 16–24 h after transfection, cells were harvested for measurement of firefly and Renilla luciferase activity using the Dual-Glo® Luciferase Assay System (Promega, catalog no. E2920). Approximately one-sixth of the transfected cells were used for each 96-well condition in this assay using the Passive Lysis Buffer (Promega, catalog no. E1941). SRE activity–related counts (firefly) were normalized by the counts obtained for the Renilla luciferase under the control of a constitutive promoter, and then results were expressed as -fold activation compared with control cells transfected with an empty plasmid instead of Gα_q.

GTPase activity

This assay was performed as described previously (47). Briefly, reactions were started by mixing equal volumes of 200 nm His-Gα_q* (WT or mutants) and 1 μm [γ-³²P]GTP (∼50 cpm/fmol) at 20 °C in assay buffer (50 mm Na-HEPES, pH 7.5, 100 mm NaCl, 0.1 mm EDTA, 0.3 mm MgCl₂, 1 mm DTT, 0.2 m (NH₄)₂SO₄, 0.05% (w/v) C₁₂E₁₀). Duplicate aliquots (25 μl) were removed at 0, 5, 10, 15, 20, 30, and 45 min, and reactions were stopped by mixing with 975 μl of ice-cold 5% (w/v) activated charcoal in 20 mm H₃PO₄, pH 3. Samples were then centrifuged for 10 min at 10,000 × g, and 500 μl of the resultant supernatant were scintillation-counted to quantify released [³²P]P_i, which was subsequently converted to moles.

Limited proteolysis

HEK293T cells were seeded on 100-mm dishes (∼2 × 10⁶ cells/dish) coated with gelatin and after 1 day were transfected using the calcium phosphate method with plasmids encoding Gα_q-HA WT or mutants (3 μg/dish). Cell medium was changed 6 h after transfection. Approximately 16–24 h after transfection, cells were scraped in PBS, centrifuged (5 min at 550 × g) and resuspended on ice with 2 ml of lysis buffer (20 mm Hepes, pH 7.2, 5 mm Mg(CH₃COO)₂, 125 mm K(CH₃COO), 0.4% (v/v) Triton X-100, 30 μm GDP, 1 mm DTT, 10 mm β-glycerophosphate, and 0.5 mm Na₃VO₄ supplemented with a protease inhibitor mixture (SigmaFAST, catalog no. S8830)). For some conditions, the lysis buffer was supplemented with 30 μm AlCl₃, 10 mm NaF, and 10 mm MgCl₂ (Mg·AlF₄⁻ condition). Ten μl of the cleared lysate (14,000 × g, 10 min) were used for each condition. Digestions were started by the addition of 100 ng of trypsin (or an equivalent volume of PBS) to each tube and incubated for 20 min at 30 °C. Reactions were stopped by the addition of Laemmli sample buffer and boiling followed by protein separation by SDS-PAGE and immunoblotting.

Immunoblotting

Proteins separated by SDS-PAGE were transferred to PVDF membranes. Membranes were blocked for 30 min with 5% (w/v) nonfat milk (or 5% (w/v) BSA for antibodies for phosphoproteins) and then incubated overnight at 4 °C with primary antibodies. The primary antibodies were used at the following dilutions: Gα_q, 1:1,000; tubulin, 1:2,500; FLAG, 1:1,000; HA, 1:250; nanoluciferase, 1:1,000; phospho-ERK1/2, 1:1,000, total ERK1/2, 1:1,000. Following primary antibody incubation, membranes were washed with PBS + 0.1% Tween 20 and incubated in secondary antibodies (goat anti-rabbit Alexa Fluor 680 and goat anti-mouse or IRDye 800) at 1:10,000 dilution. IR imaging of Western blots was performed according to the manufacturer's protocols using an Odyssey IR Imaging System (LI-COR Biosciences). Images were processed using ImageJ software (National Institutes of Health) and assembled for presentation using Photoshop and Illustrator software (Adobe).

Statistical analyses

Each experiment was performed at least three times unless otherwise indicated. The data shown are presented as means with error bars representing the S.E. or as one representative result of each biological replicate (as indicated in the figure legends). Statistical significance between various conditions was assessed with Student's t test. p < 0.05 was considered significant.

Author contributions

M. M., A. Leyme, A. M., A. Luebbers, P. P. P., and M. G.-M. conducted experiments. M. M., A. Leyme, A. M., and M. G.-M. designed experiments. M. M., A. L., A. M., and M. G.-M. analyzed data. Z. C. and S. R. S. provided critical reagents. M. M. and M. G.-M. wrote the manuscript. M. G.-M. conceived and supervised the project.