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J. Biol. Chem., Vol. 279, Issue 34, 35287-35297, August 20, 2004

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Dominant-negative Inhibition of Pheromone Receptor Signaling by a Single Point Mutation in the G Protein Subunit^*

Yuh-Lin Wu, Shelley B. Hooks, T. Kendall Harden, and Henrik G. Dohlman¶

From the Department of Biochemistry and Biophysics and the Department of Pharmacology, University of North Carolina, Chapel Hill, North Carolina 27599-7260

Received for publication, May 3, 2004 , and in revised form, June 9, 2004.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
In yeast, two different constitutive mutants of the G protein subunit have been reported. Gpa1^Q323L cannot hydrolyze GTP and permanently activates the pheromone response pathway. Gpa1^N388D was also proposed to lack GTPase activity, yet it has an inhibitory effect on pheromone responsiveness. We have characterized this inhibitory mutant (designated G^ND) and found that it binds GTP, interacts with G protein subunits, and exhibits full GTPase activity in vitro. Although pheromone leads to dissociation of the receptor from wild-type G protein, the same treatment promotes stable association of the receptor with G^ND. We conclude that agonist binding to the receptor promotes the formation of a nondissociable complex with G^ND, and in this manner prevents activation of the endogenous wild-type G protein. Dominant-negative mutants may be useful in matching specific receptors and their cognate G proteins and in determining mechanisms of G protein signaling specificity.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Mammalian G protein-coupled receptors (GPCRs)¹ respond to a variety of signaling molecules, including hormones, neurotransmitters, odors, and light. These signals regulate diverse cellular processes, such as the control of blood pressure and heart rate, perception of pain, cell proliferation, inflammation, and platelet aggregation (1–3). Upon binding of an agonist to its cognate receptor, the G protein subunit transits from a GDP-bound to a GTP-bound state and liberates the G protein subunit complex. The dissociated (in the GTP-bound state) and subunits then activate a variety of downstream effectors. Regulators of G protein signaling reverse this process by binding to G-GTP and promoting GTP hydrolysis, after which the subunits reassociate and signaling terminates (4).

A prominent feature of G protein signaling is the tremendous diversity of the component proteins. Human genome analysis has revealed genes that encode several hundred candidate GPCRs, 16 subunits, 5 subunits, and 12 subunits (5). Further diversity results from alternative mRNA splicing and the potential of a given receptor to transduce signals to multiple G protein subunit subtypes and effectors. These signaling components do not assemble randomly; rather, one receptor typically activates only a subset of G protein heterotrimers, and the dissociated subunits activate only a subset of down-stream effectors (5).

Clearly, a major challenge is to define the coupling specificity of specific receptors, G proteins, and effectors. Bacterial toxins have long been used to perturb signaling mediated by susceptible G proteins. Cholera toxin catalyzes the ADP-ribosylation of G_s resulting in inactivation of its GTPase activity, thus maintaining G_s in the active GTP-bound state. Pertussis toxin catalyzes ADP-ribosylation of G_i, and this modification blocks G protein coupling to GPCRs (6). However, pertussis toxin does not modify all members of the G_i subfamily, and no toxins have been identified that modify members of the G_q and G₁₂ subfamilies. Thus, more general approaches are needed to analyze receptor and G protein coupling specificity.

An alternative approach to studying G protein signal specificity has been to mutate residues in G critical for GTPase activity. One early example of an activated G protein allele was described by Landis et al. (7), who showed that certain types of human pituitary tumors are associated with GTPase-deficient mutants of G_s. Another GTPase-deficient mutant replaces Gln-204 in G_i (Gln-277 in G_s and Gln-323 in Gpa1) (8–10). A crystal structure of G ^Q204L_i and additional biochemical analysis suggest that the catalytic Gln acts by stabilizing the trigonal-bipyramidal transition state and by helping to orient the hydrolytic water molecule (11, 12). This mutation is widely used to identify signaling pathways activated by G subunits, and we recently used this approach to show that the yeast G subunit Gpa1 directly activates the mating response pathway, in conjunction with G (13).

Another approach is to inactivate G protein function by using mutants that confer a dominant-negative effect on signaling (14). Dominant-negative mutants are proteins that disrupt the function of the endogenous wild-type protein when overexpressed. Thus, highly specific dominant-negative G proteins have tremendous potential for ascertaining the signaling specificity of diverse G proteins in complex systems. Although such mutants have been reported (15–21) for various G subunits, they have not yet been proved to be generally applicable to studying G protein signaling.

In contrast to the large number and variety of mammalian GPCRs, only two distinct G protein signaling systems exist in yeast. The first regulates the response to mating pheromones, which are secreted by haploid a and cell types in preparation for mating. Pheromone binding to receptors (e.g. Ste2 in a cells) triggers dissociation of G (Gpa1) from the G heterodimer (Ste4/Ste18). The dissociated subunits proceed to activate a mitogen-activated protein kinase cascade, resulting in new gene transcription, cell cycle arrest, and eventually cell fusion to form the a/ diploid (22). The second signaling pathway mediates the cellular response to glucose and environmental stressors such as high osmolarity and heat shock. Components of this pathway include the G protein Gpa2 working in conjunction with the putative glucose receptor Gpr1 (23, 24). Recent studies of Gpr1 signaling have identified two candidate G subunits, Gpb1 and Gpb2, and a candidate G, Gpg1 (25, 26). The Gpb1/2 proteins lack the seven WD40 repeats found in classical G proteins, but instead contain seven kelch repeats implicated in protein-protein interaction (26). The effector for this G protein has not been positively identified, and generally speaking much less is known about this pathway.

Two different constitutive mutants in the yeast G subunit have been described. Gpa1^Q323L binds but does not hydrolyze GTP (13, 27). Gpa1^N388D was proposed to lack GTPase activity but paradoxically has an opposing or inhibitory effect on the pathway (28–30). We previously characterized the biochemical and physiological function of Gpa1^Q323L (13, 27), and here we characterize the inhibitory G^ND mutant. We report that G^ND binds and hydrolyzes GTP but is unable to dissociate effectively from receptors and therefore acts as a potent dominant-negative inhibitor of receptor signaling.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Strains, Media, and Plasmids—Established methods were used for the growth and genetic manipulation of bacteria and yeast (31). Escherichia coli strain DH 5 was used for plasmid maintenance and amplification. The strains of yeast Saccharomyces cerevisiae used in this study are as follows: YPH499 (MATa leu2-1 his3-200 trp1-63 ade2-101^oc lys2-801^am ura3-52), YPH501 (YPH499 MATa/) (32), YGS5 (YPH499 gpa1::hisG ste11^ts) (33), BY4741 (MATa his31 leu20 met150 ura30; from Research Genetics, Huntsville, AL), and a BY4741-derived gpa1 ste7 mutant strain (MATa ste7::KanMX gpa1::hisG, provided by Paul Flanary, University of North Carolina).

Yeast cells were grown in synthetic medium supplemented with adenine, amino acids, 2% glucose (SCD), or 2% galactose plus 0.2% sucrose (SCG) to express GAL1/10-inducible genes. Leucine, uracil, tryptophan, or histidine was omitted to maintain selection of plasmids as needed. Yeast cells were grown at 30 °C unless otherwise stated. The absence of GPA1 in strain YGS5 normally results in constitutive G signaling and growth arrest; however, these cells can be maintained at 34 °C due to inactivation of the temperature-sensitive ste11 mutant.

Several expression plasmids used in this study have been described previously: pRS315 (CEN, LEU2), pRS423 (2 µm, HIS3), pRS424 (2 µm, TRP1) (32), pAD4M (2 µm, LEU2, ADH1 promoter and terminator), and pAD4M-GST and pAD4M-GPA1-GST (34). pGAL^H (2 µm, LEU2) and pGAL^L (2 µm, LEU2) contain partially active GAL1/10 promoter sequences having 18–20 (pGAL^H) or 1%(pGAL^H) full activity (35) (provided by Ming Guo, Yale University). pRS315-GPA1 contains the GPA1 gene under the control of the native promoter.

Wild-type human G_i2 in pcDNA3.1 mammalian expression vector was obtained from the Guthrie cDNA Resource Center (Sayre, PA). The N270D mutation was introduced using a QuikChange mutagenesis kit (Stratagene, Alameda, CA). The mutant was amplified by PCR and ligated into pFastBacHta (Invitrogen) digested with BamHI and XhoI, which introduced an in-frame hexahistadine tag at the N terminus (His₆-G ^N270D_i2-pFastBacHta). Other Sf9 expression plasmids were described previously (36, 37).

GPA1^Q323L, GPA1^N388D, GPA2^N365D, and G ^N270D_i2 mutations were introduced using QuikChange. The primer sequences are as follows: 5'-TCGACGCTGGAGGCCTGCGTTCTGAACG-3' and 5'-CGTTCAGAACGCAGGCCTCCAGCGTCGA-3' for GPA1^Q323L; 5'-CGTTTATTTTGTTTTTAGATAAAATTGATTTGTTC-3' and 5'-GAACAAATCAATTTTATCTAAAAACAAAATAAACG-3' for GPA1^N388D; 5'-TCTGTCGTACTCTTTCTGGATAAAATCGACCTTTT TG-3' and 5'-CAAAAAGGTCGATTTTATCCAGAAAGAGTACGACAGA-3' for GPA2^N365D; 5'-TCCATCATCCTCTTCCTCGACAAGAAGGACCTGTTTG-3' and 5'-CAAACAGGTCCTTCTTGTCGAGGAAGAGGATGATGGA-3' for G ^N270D_i2. Each mutation was confirmed by sequencing analysis.

Expression of G (Ste4/Ste18) was under the control of the bidirectional GAL1/10 promoter in pRS424-GAL-STE4/STE18. This plasmid was constructed by combining a SalI-EcoRI digestion product containing the GAL1/10 promoter and STE4 (from pL19, provided by Malcolm Whiteway, University of Montreal) (38) with an EcoRI-SacI digestion product of STE18 amplified by PCR using the primers 5'-GGGAATTCTAGGATAGTAGCAATCGCA-3' and 5'-GAGGCTCTACGTAGCAAG-3' and a SalI-SacI digestion product of plasmid pRS424. Expression of STE2 or STE7 was achieved by PCR amplification and subcloning into the pYES2.1/V5-His-TOPO (2 µm, URA3, GAL1/10 promoter, CYC1 terminator; Invitrogen). Amplification primers used were 5'-CCCAAGCTTCCAGAATGTCTGATGCGGCTCCTTC-3' and 5'-CCCAAGCTTTAAATTATTATTATCTTCAGTC-3' for STE2; 5'-GCATCGGATCATATCTGTTT-3' and 5'-GCTGGAAAAAGAAGAGACTA-3' for STE7. A stretch of double-stranded DNA encoding three tandem repeats of the FLAG tag (sense, 5'-GATTATAAAGATGACGATGACAAGGATTATAAAGATGACGATGACAAGGATTATAAAGATGACGATGACAAG-3') in pYES2.1/V5-His-TOPO was digested with HindIII and ligated to HindIII-digested STE2 that had been PCR-amplified using the primers 5'-CCCAAGCTTCCAGAATGTCTGATGCGGCTCCTTC-3' and 5'-CCCAAGCTTTAAATTATTATTATCTTCAGTC-3', yielding pYES-STE2-FLAG.

Expression of G ^ND_i2 in Insect Cells—A baculovirus encoding human G ^N270D_i2 with an N-terminal hexahistidine tag was prepared and amplified using plasmid his₆-G ^N270D_i2-pFastBacHta according to the manufacturer's instructions (Invitrogen). Four liters of Sf9 cells at a density of 1.5 x 10⁶ cells/ml were infected with the virus at a multiplicity of infection of 2 and harvested 48 h after infection by centrifugation at 1,000 x g for 15 min at 4 °C. All subsequent steps were carried out at 4 °C. Cells were resuspended in 400 ml of ice-cold cell lysis buffer (20 mM HEPES, pH 8, 100 mM NaCl, 2 mM MgCl₂, 9.8 mM 2-mercaptoethanol, 0.01 mM GDP, 500 nM aprotinin, 10 µM leupeptin, 200 µM phenylmethylsulfonyl fluoride, 1 nM pepstatin, 10 µM L-1-tosylamido-2-phenylethyl chloromethyl ketone) and lysed by passage through a pressurized Emulsiflux (Avestin, Ottawa, Canada). The lysate was centrifuged at 500 x g for 15 min to remove intact cells and nuclei. The cleared lysate was further centrifuged at 150,000 x g for 35 min in an ultracentrifuge (Beckman). The resulting supernatant fraction was passed over an equilibrated 2.5-ml nickel-nitrilotriacetic acid (Ni-NTA)-agarose resin (Qiagen) column at a flow rate of 2 ml/min, and the flow-through was reapplied to the column. The column was washed with 10 ml of high salt wash (cell lysis buffer + 300 mM NaCl), and 10 ml of 10 mM imidizole in cell lysis buffer. His-G ^N270D_i2 was eluted with 10 ml of 150 mM imidazole in cell lysis buffer, diluted 1:4 in Buffer A (20 mM HEPES, pH 8, 2 mM MgCl₂, 1 mM dithiothreitol, 0.5 mM EDTA, 0.01 mM GDP, 500 nM aprotinin, 10 µM leupeptin, 200 µM phenylmethylsulfonyl fluoride, 1 nM pepstatin, 10 µM L-1-tosylamido-2-phenylethyl chloromethyl ketone), and loaded onto an equilibrated 1-ml HiTrap Q-Sepharose FPLC column (Amersham Biosciences) according to the manufacturer's recommendations. A linear gradient from 0 to 500 mM NaCl (30 ml) was used to elute proteins bound to the column. Five hundred microliter fractions were collected and immediately assayed for [³⁵S]GTPS binding and GTPase activity.

[³⁵S]GTPS Binding Assay—[³⁵S]GTPS binding assays were performed in triplicate in conical polypropylene tubes (Sarstedt) in a 100-µl reaction volume. Fifty microliters of each sample was added to 50 µl of 2x GTPS binding buffer (50 mM HEPES, pH 8, 4 mM MgCl₂, 1 mM EDTA, 5 µM GTPS, 500,000 cpm/reaction [³⁵S]GTPS) and incubated ina30 °C water bath for 60 min. Reactions were then transferred to ice and diluted with 5 ml of ice-cold stop buffer (20 mM Tris-HCl, pH 8, 25 mM MgCl₂, 120 mM NaCl). The diluted reactions were filtered over pre-wetted 0.45-µm type HA nitrocellulose filters (Millipore) using a vacuum manifold and washed twice with 5 ml of stop buffer. The filters were then added to scintillation vials with scintillant, and radioactivity was determined using a scintillation counter.

GTPase Activity—Steady state GTPase assays were performed in duplicate in conical polypropylene tubes in a 50-µl reaction volume. Twenty five microliters of each sample was added to 25 µlof2x GTPase buffer (40 mM HEPES, pH 8, 100 mM NaCl, 4 mM MgCl₂, 2 mM EDTA, 4 µM GTP, 625,000 cpm/reaction [-³²P]GTP) and incubated in a 30 °C water bath for 30 min. To terminate the reaction, the tubes were moved to an ice bath, and 950 µl of suspended 5% activated charcoal (Sigma) in 20 mM NaH₂PO₄ was added to each tube. The reaction tubes were subjected to centrifugation at 3,000 x g in a swinging bucket centrifuge for 10 min to collect the charcoal pellet. Six hundred microliters of the cleared supernatant were transferred to scintillation vials with scintillation fluid for quantification of radioactivity.

Pheromone Signaling Assays—Two outcomes of pheromone signaling were measured. The first was pheromone-dependent growth inhibition (halo assay) (39). Briefly, 100 µl of a saturated cell culture was mixed with 2 ml of water and 2 ml of 1% (w/v) dissolved agar (55 °C) and poured onto selective SCD or SCG agar plates. Synthetic -factor was spotted onto sterile paper disks and placed on the nascent lawn. The resulting zone of growth inhibition was recorded after 48 h.

The second assay was pheromone-dependent reporter transcription activity (39). In this method a pheromone-inducible promoter (FUS1) drives expression of a reporter enzyme (-galactosidase) (39). A saturated cell culture in selective SCD medium was diluted 1:200 in fresh SCD medium, allowed to grow overnight, washed, and resuspended in SCG medium to A_{600 nm} 0.6. After 4 h the cells were aliquoted (90 µl, in triplicate) to 96-well plates containing 10 µlof -factor and incubated for 90 min at 30 °C. -Galactosidase activity was measured by adding 20 µl of a freshly prepared solution of 83 µM fluorescein di--D-galactopyranoside (Marker Gene Technologies Inc.), 137.5 mM PIPES, pH 7.2, 2.5% Triton X-100 and incubating at 37 °C until a bright yellow color appeared. The reaction was stopped by the addition of 20 µlof1 M Na₂CO₃, and the fluorescence activity was monitored using an excitation of 485 nm and an emission of 530 nm.

Heat Shock Assay—A saturated cell culture in selective SCD medium was diluted 1:20 into fresh SCD medium and incubated for 48 h. Cell cultures were then transferred to glass tubes and placed in a 50 °C water bath for 45 min. Heat-shocked (50 °C) and nonheat-shocked (30 °C) cells were diluted and plated on selective SCG agar plates. Surviving cell colonies were counted after 2–3 days (23).

Gpa1 and Gpa1^N388D Expression Assay—To compare expression of Gpa1 versus Gpa1^N388D, protein concentration was examined in the gpa1 deletion YGS5 strain and in the diploid YPH501 strain, which normally does not express the receptor or G protein. To assess regulation by pheromone, -factor (2.5 µM) was added at A_{600 nm} 0.6 and incubation continued for an additional 2 h. Cells were grown to mid-log phase in SCG (A_{600 nm} 1.0), and cell growth was stopped by addition of 10 mM (final concentration) NaN₃. Cells were harvested by centrifugation at 1,000 x g for 10 min. Cells were washed once with 10 mM NaN₃ and resuspended in phosphate-buffered saline, pH 7.3. Cells were then lysed by vortexing with glass beads four times for 1 min each and then centrifuged at 10,000 x g for 30 s. The resulting supernatant was collected and mixed with SDS-PAGE sample buffer (60 mM Tris-HCl, pH 6.8, 10% glycerol, 14.4 mM 2-mercaptoethanol, 10 µg/ml bromphenol blue, 4% SDS) and heated at 100 °C for 10 min. The samples were allowed to cool and subjected to immunoblotting analysis (see below). To compare stability of Gpa1 and Gpa1^N388D, cycloheximide was added (10 µg/ml final concentration) for various times prior to harvesting the cells.

Co-purification of Gpa1 with G—YPH501 cells expressing receptor (Ste2), G (Ste4/Ste18), and either G (Gpa1) or G^ND (Gpa1^N388D) fused to glutathione S-transferase (GST) or GST alone were grown in selective SCG medium. All the following procedures were carried out at 4 °C. After termination of cell growth, 50 A_{600 nm} units of cells were resuspended in purification lysis buffer (40 mM triethanolamine, pH 7.2, 2 mM EDTA, 150 mM NaCl, 2 mM dithiothreitol, 0.2 mM 4-[2-aminoethyl]benzenesulfonyl fluoride HCl, 15 µg/ml leupeptin, 20 µg/ml pepstatin, 1 mM benzamidine, 10 µg/ml aprotinin, 100 µM glycerol 2-phosphate, 0.5 mM sodium orthovanadate). Cells were split into two equal portions, and each portion was resuspended in 1 ml of lysis buffer containing 3 mM MgCl₂ and 10 µM GDP (condition 1, "–AlF^–₄") or 3 mM MgCl₂,10 µM GDP, 30 µM AlCl₃, and 10 mM NaF (condition 2, "+AlF^–₄"). Cells were lysed by vortexing with glass beads four times for 1 min each. The resultant lysates were harvested by centrifugation at 1,000 x g for 10 min and solubilized by addition of Triton X-100 (1% final concentration) and rocking for 1 h. The samples were then centrifuged at 1,000 x g for 10 min, and the resulting supernatant was mixed with 100 µl of a 30% slurry of glutathione-Sepharose 4B (Amersham Biosciences) in the appropriate lysis buffer (condition 1 or condition 2) and incubated for 2 h. The glutathione-Sepharose 4B was centrifuged at 10,000 x g for 10 min and washed three times with 1 ml of phosphate-buffered saline. The bound proteins were eluted by heating at 100 °C in SDS-PAGE sample buffer.

Co-immunoprecipitation of Receptor (Ste2) and Gpa1—YPH501 cells expressing G (Ste4/Ste18), Gpa1, or Gpa1^N388D and receptor (Ste2) tagged or nontagged with the FLAG epitope were grown in selective SCG medium to A_{600 nm} 1.0. All the following procedures were conducted at 4 °C. Twenty five A_{600 nm} units of cells were resuspended in IP lysis buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 0.1% Triton X-100, 5 mM EDTA, 10% glycerol, 100 µg/ml phenylmethylsulfonyl fluoride, 1 mM dithiothreitol, 1x of protease inhibitor mixture (catalog number 1873580; Roche Applied Science)), followed by vortexing with glass beads four times for 1 min each. Cell lysates were harvested by centrifugation at 1,000 x g for 10 min, and the supernatant was subjected to 1 h of rocking to liberate membrane-bound proteins. The samples were then centrifuged at 10,000 x g for 10 min, and the resulting supernatant was incubated with 40 µl of a 50% slurry of EZview^TM Red anti-FLAG M2 affinity gel (Sigma). After2hof gentle agitation, the gel was centrifuged at 10,000 x g for 30 s and washed three times with IP lysis buffer. Elution of FLAG-tagged protein was achieved by incubating the gel with 15 µg of 3x FLAG peptide in 50 µl of elution buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 5 mM EDTA) with gentle shaking for 30 min. Supernatant was harvested by centrifugation at 8,000 x g for 30 s and subjected to immunoblotting assay.

Immunoblot Detection—Protein samples in SDS-PAGE sample buffer were resolved by SDS-PAGE, transferred to nitrocellulose membrane, and probed with antibodies against Gpa1 (1:1,000 dilution) (40), FLAG (1: 3,000; Sigma), GST (1:1,500; from Joan Steitz, Yale University), Ste4 (1:2,000; from Duane Jenness, University of Massachusetts), or G_i2 (1:2,500; Qiagen). Antibodies were detected using secondary antibodies such as horseradish peroxidase-conjugate goat anti-mouse IgG or anti-rabbit IgG (Bio-Rad). The signal was detected by the ECL system (Amersham Biosciences) according to the manufacturer's instructions.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
G^ND Can Bind and Hydrolyze GTP—Gpa1^N388D is a potent inhibitor of pheromone signaling. Other investigators have suggested that this mutation impairs GTPase function and proposed that the inhibition of signaling might occur through activation of a "desensitization effector" (28–30, 41). However, the GTPase activity of the ND mutant has not been measured in any system. A previous attempt to purify recombinant Gpa1^N388D from bacteria yielded a product unable to hydrolyze GTP, but which was also unable to bind to GTP or G (28). These findings suggest that the mutant protein is unstable and loses activity during purification. As an alternative strategy we attempted to purify the analogous mutant form of G_i2 expressed in insect cells. G_i is the closest mammalian homologue to Gpa1; both proteins have nearly identical guanine nucleotide binding pockets, and a direct comparison revealed that they have very similar kinetic properties in vitro (27).

As shown in Fig. 1, we were able to purify small quantities of the mutant protein from insect cells. We generated a baculovirus encoding human G ^N270D_i2 fused at the N terminus to a hexahistidine affinity tag (his₆-G ^N270D_i2). Sf9 cell infection with the virus resulted in heterologous expression of the mutant protein, although at significantly lower levels than that observed following infection with a similar virus encoding wild-type G_i2 (data not shown). Lysates were separated by high speed centrifugation into particulate and soluble fractions, and the soluble fraction was passed over an Ni-NTA affinity column. After extensive washing the his₆-G ^N270D_i2 was eluted with 150 mM imidazole. Fractions were then resolved by SDS-PAGE and visualized by staining with Coomassie Blue as well as by immunoblotting and detection with anti-pentahistidine antibodies. Both detection methods revealed a single prominent band migrating at 40.5 kDa, which is the predicted molecular mass of his₆-G ^N270D_i2 (data not shown).

View larger version (45K): [in this window] [in a new window]   FIG. 1. Nucleotide binding and GTPase activity of purified G N270Di2. Recombinant his6-Gi2N270D was purified from Sf9 cell lysates by Ni-NTA affinity chromatography followed by ion exchange chromatography, as described under "Experimental Procedures." A, His6-Gi2N270D was eluted from an ion exchange column using a 0–500 mM NaCl linear gradient. Purity was assessed by Coomassie Blue staining of 12.5% acrylamide SDS-PAGE gels. Fraction numbers are provided above each lane, and the position of molecular weight standards (kDa) are indicated on the left. B, elution fractions were resolved by SDS-PAGE, transferred to nitrocellulose, and probed with anti-pentahistidine antibody to detect the presence of his6-Gi2N270D. numbers are Fraction provided above each lane. C, elution fractions were assayed for [35S]-GTPS binding and GTPase activities. Open bars, [35S]GTPS binding. Samples (in triplicate) of each fraction were incubated with 2.5 µM [35S]GTPS for 60 min at 30 °C. The reactions were filtered over nitrocellulose, and the retained radioactivity was quantitated by scintillation counting to determine GTPS binding activity. Hatched bars, GTPase activity. Samples (in duplicate) of each eluate fraction were incubated with 2 µM [32P]GTP for 30 min at 30 °C. Activated charcoal was added to each reaction and collected by centrifugation, and radioactivity in the supernatant was quantitated by scintillation counting to determine free phosphate. Fraction numbers are indicated below each bar. Error bars, mean ± S.E. Note that a wider range of fractions are included in B and C than in A.

The Q-Sepharose eluate was assayed for [³⁵S]GTPS binding and GTPase activities. As shown in Fig. 1C, substantially more [³⁵S]GTPS binding and GTPase activity was observed in the first peak (fractions 32–36) than in the second peak, suggesting that the his₆-G_i2^N270D present in the later fractions was inactive. Even in this early peak the active form of the protein (determined by GTPS binding) was 15% of the estimated total concentration of G ^N270D_i2 (determined by protein staining). Wild-type his-G_i2 purified under the same conditions eluted as a single discrete peak at 200 mM NaCl, and these fractions exhibited nearly stoichiometric [³⁵S]GTPS binding (data not shown).

Significant loss of [³⁵S]GTPS binding activity of G ^N270D_i2 occurred as a function of time and temperature. For this reason all purification steps were carried out at 4 °C, and all assays of [³⁵S]GTPS binding and GTPase activities were performed within 10 h of cell lysis. Despite these precautions, it appeared that much of the protein was inactive, further suggesting that the protein is unstable (data not shown).

Finally, we observed that co-expression with G₁ and G₂ increased the proportion of G ^N270D_i2 associated with the membrane fraction, suggesting that the mutant protein interacts with G dimers and is recruited to the membrane by this interaction (data not shown). We have also co-expressed untagged G ^N270D_i2 and pentahistidine-tagged G₂ with G₁ and purified the heterotrimeric complex by Ni-NTA and ion exchange chromatography. We found that G ^N270D_i2 co-eluted with his₆-G in this procedure, further suggesting that the mutant associates with G (data not shown).

Gpa1^N388D Is a Receptor- and G Subtype-selective Dominant-negative Mutant—The data presented in Fig. 1 indicate that G^ND binds and hydrolyzes GTP normally. This makes it unlikely that Gpa1^N388D functions by activating any effector protein, because effectors recognize only the GTP-bound form of G. We therefore considered whether the mutant functions as a dominant negative. Dominant-negative mutants will, when overexpressed, inhibit the function of the endogenous wild-type protein (14). In this scenario, the Gpa1^N388D phenotype could result from interference with receptor-G protein coupling, from inhibition of G protein subunit dissociation, or from both.

As an initial test of the model we asked whether Gpa1^N388D specifically inhibits the signaling activity of the pheromone receptor Ste2. Two distinct GPCR signaling pathways exist in yeast. In haploid cells, the primary function of Ste2 is to modulate the activity of Gpa1 (G) and the Ste4/Ste18 dimer (G) (22). Activation of this pathway leads to growth arrest and mating. Although less well characterized, a putative glucose receptor Gpr1 modulates the activity of Gpa2 (G), Gpb1 or Gpb2 (G-like proteins), and Gpg1 (G). Activity of this pathway allows the cell to adapt to cell stress (42).

We first compared pheromone signaling in wild-type cells that express Gpa1^N388D or the analogous mutant form of Gpa2, Gpa2^N365D. Both G mutants were expressed using the attenuated galactose-inducible promoters, GAL^H (20% of wild-type activity) and GAL^L (1% of wild-type activity). Two outcomes of pheromone signaling were measured. The first was pheromone-mediated cell growth arrest (halo assay). Pheromone spotted onto a filter disk produces a zone of growth arrest the size of which correlates with pheromone sensitivity. As shown in Fig. 2, modest overexpression of Gpa1^N388D inhibited the response to pheromone, as described previously (30). Inhibition was dependent on the expression level of Gpa1^N388D as the GAL^H promoter conferred more effective inhibition (Fig. 2, bottom panel) than GAL^L (top panel). Inhibition was absent when expression was repressed by growth in glucose. Expression of wild-type Gpa1, wild-type Gpa2, or Gpa2^N365D also did not affect the halo response (Fig. 2 and data not shown).

View larger version (80K): [in this window] [in a new window]   FIG. 2. Gpa1N388D inhibits pheromone-dependent growth arrest. To measure pheromone-dependent growth arrest, wild-type cells (strain BY4741) were transformed with wild-type GPA1 or GPA1N388D expressed from an attenuated GAL1/10 promoter having 1 (GALL) or 20% (GALH) of wild-type activity. Cells were plated onto medium containing either galactose (to induce expression) or glucose (to repress expression) and exposed to filter disks spotted with -factor pheromone (clockwise from top left, 75, 25, and 8 µg). The resulting zone of growth inhibition was documented after 2–3 days.

View larger version (26K): [in this window] [in a new window]   FIG. 3. Gpa1N388D inhibits pheromone-induced transcription. To measure pheromone-dependent transcription, wild-type cells (strain BY4741) were co-transformed with a reporter plasmid containing the pheromone-responsive FUS1 promoter fused to lacZ and a second plasmid containing GPA1N388D (A), GPA2N365D (B), or no insert (Vector, A and B) expressed from either the GALL or GALH promoter. Cells were grown in the presence of galactose and analyzed for -galactosidase activity following treatment with the indicated concentrations of -factor. -Galactosidase activity units are arbitrary. Each point is an average of three measurements, and the data shown are representative of three independent experiments. Error bars, S.E.

View larger version (16K): [in this window] [in a new window]   FIG. 4. Gpa2N365D inhibits recovery from heat shock. Wild-type cells (strain YPH499) were transformed with a plasmid containing no insert (Vector), wild-type GPA1, wild-type GPA2, GPA1N388D, or GPA2N365D under the control of the GALH promoter. Cells in liquid medium were placed in either a 50 °C water bath for 45 min (A) or maintained at the normal growth temperature of 30 °C (B) and then plated onto solid medium. Colonies were counted after 2–3 days. Heat shock recovery was expressed as the relative survival rate of each transformed strain and expressed relative to the vector control group (defined as 100%). Each value is an average of three measurements, and the data shown are representative of four independent experiments. Error bars, S.E.

View larger version (46K): [in this window] [in a new window]   FIG. 5. Signal inhibition by Gpa1N388D requires receptor coupling activity. To determine whether coupling to receptors is required for Gpa1N388D activity, wild-type cells (strain BY4741) were transformed with plasmid pAD4M containing GPA1N388D-GST, GPA1-GST, or GST alone. Fusion to GST preserves G binding but blocks activation by receptors, as discussed above. A, cells were plated onto selective medium and exposed to disks spotted with -factor pheromone (clockwise from top, 75, 25, 8, and 0 µg). The resulting zone of growth inhibition was documented after 2–3 days. B, total lysates were prepared from the cells used in A, resolved by 7.5% SDS-PAGE, and subjected to immunoblotting with anti-GST antibodies (Ab). Arrows indicate full-length protein specifically detected by the antibody. Other bands are presumed to be degradation products.

To test this aspect of the model, Gpa1^N388D was expressed in the absence of wild-type Gpa1 (i.e. a gpa1 mutant strain). Cells lacking GPA1 are normally not viable due to constitutive release of G leading to cell division arrest (43, 44). However, the cells used here also do not express the downstream kinase gene STE7 and are viable. Thus, signaling can be monitored by growth arrest and reporter transcription assays following induction of STE7 expression from a plasmid.

As predicted by the model, Gpa1^N388D blocked cell division arrest in pheromone-treated cells (those nearest the source of pheromone) (Fig. 6A) (45). Cells grew poorly at the perimeter of the halo, where pheromone concentrations are reduced (Fig. 6A). Cells expressing wild-type Gpa1 exhibited a more typical growth arrest phenotype. Cells closest to pheromone underwent cell division arrest, whereas those further away continued to grow. This pattern of signaling supports the model that Gpa1^N388D can inhibit signaling, but only upon receptor activation. This is in contrast to wild-type Gpa1, which promotes signaling upon receptor activation.

View larger version (54K): [in this window] [in a new window]   FIG. 6. Pheromone-dependent inhibition of constitutive signaling by Gpa1N388D in a gpa1 mutant strain. A, gpa1 ste7 mutant strain was transformed with plasmids containing STE7 under the control of the GAL1/10 promoter and either wild-type GPA1 or GPA1N388D under the control of the GALH promoter. Cells were plated onto galactose-containing medium and exposed to -factor pheromone (counter-clockwise from top, 75, 25, 8, and 0 µg). The resulting zone of pheromone-dependent cell growth was recorded after 2 days. B, the same strain was transformed with plasmids containing STE7 (GAL1/10 promoter), the FUS1-lacZ reporter, and either GPA1N388D (GALH promoter) or wild-type GPA1 regulated by its native promoter (pRS315-GPA1). Each point is an average of three measurements, and the data shown are representative of three independent experiments. Error bars, S.E.

Pheromone Stimulation Increases Gpa1^N388D Stability and Expression—The above results suggest that pheromone triggers the association of Gpa1^N388D with G and receptor. Thus, we investigated the mechanism by which pheromone treatment can unmask the apparent ability of Gpa1^N388D to sequester G. One possibility is that pheromone is required for stable expression of Gpa1^N388D (47). Indeed, purification of G_i2^N270D or Gpa1^N388D yielded protein that was unstable in vitro (28). Thus we considered whether Gpa1^N388D is also unstable in vivo.

To determine whether Gpa1^N388D is unstable, we monitored its rate of loss in cells following treatment with the protein synthesis inhibitor cycloheximide. As shown in Fig. 7A, overexpressed wild-type Gpa1 protein was quite stable, with almost no change after a 90-min treatment with cycloheximide. In contrast, Gpa1^N388D was expressed at much lower levels, and expression was undetectable after only 30 min of translation inhibition (Fig. 7A). These data indicate that the mutant is unstable.

View larger version (34K): [in this window] [in a new window]   FIG. 7. Pheromone-dependent expression of Gpa1N388D. A, to measure Gpa1N388D expression, a gpa1 mutant (strain YGS5) was transformed with a plasmid containing either no insert (Vec), wild-type GPA1, or GPA1N388D under the control of the GALH promoter. Cells were grown to mid-log phase in galactose medium and then treated with the protein synthesis inhibitor cycloheximide (CHX) for the indicated times. Whole cell lysates were resolved by 7.5% SDS-PAGE and subjected to immunoblotting with anti-Gpa1 antibodies. B, the same cells as in A were grown for 18 h in the absence or presence of 2.5 µM -factor, as indicated (o/n -MF). Another addition of -factor was made 2 h prior to collecting the cells, as indicated (-MF). Whole cell lysates were resolved by 7.5% SDS-PAGE and subjected to immunoblotting with anti-Gpa1 antibodies (Ab). Arrows indicate protein specifically detected by the antibody (Ab). Data are representative of four experiments.

In Vivo Reconstitution of Receptor, G, and Gpa1^N388D— The data presented above indicate that pheromone-occupied receptor slows the degradation of Gpa1^N388D. Therefore, we asked whether binding to G also contributes to stable expression of the mutant protein. These experiments were conducted in diploid cells, which normally lack the receptor and G protein subunits. Through heterologous expression of each component, alone or in combination, we determined the relative contribution of each to stabilized expression of Gpa1^N388D.

As shown in Fig. 8 (top panel), Gpa1 and Gpa1^N388D proteins were barely detectable when expressed alone. The abundance of both the mutant and wild-type protein was enhanced by co-expression of the receptor and was further enhanced by co-expression of both the receptor and G (bottom panel). Most surprising, pheromone treatment in this case did not appear to affect the expression of either the wild-type or mutant protein. This could be due to overexpression of the signaling proteins, which might dampen signaling efficiency or reflect the absence in diploids of another required signaling component such as the haploid-specific proteins Ste5, Far1, Sst2, or Fus3. Nevertheless, these results support our hypothesis that receptor helps to stabilize the expression of Gpa1^N388D. These data also reveal a contribution of G to G stability.

View larger version (34K): [in this window] [in a new window]   FIG. 8. Receptor- and G-dependent expression of Gpa1N388D. To determine the relative contribution of receptor and G to stable Gpa1N388D expression, wild-type diploid cells (strain YPH501) were transformed with plasmids containing no insert (Vec), wild-type GPA1, or GPA1N388D, alone or in combination with plasmids that express receptor (STE2), G (STE4/18), or both, as indicated. Whole cell lysates were resolved by 7.5% SDS-PAGE and subjected to immunoblotting with anti-Gpa1 antibodies (Ab). The arrows indicate the protein specifically detected by the Gpa1 antibody. Data are representative of four experiments.

View larger version (49K): [in this window] [in a new window]   FIG. 9. GTPase activity is not required for Gpa1N388D function. To determine whether GTP hydrolase activity is required for Gpa1N388D function, a second mutation (Q323L) was introduced into GPA1N388D. The gpa1 ste7 mutant strain was transformed with a plasmid containing either GPA1N388D or GPA1N388D/Q323L under the control of the GALH promoter and analyzed by using the growth inhibition assay (-factor pheromone, counter-clockwise from top, 75, 25, and 8 µg) (A) and the transcription activation assay (B) as described above. Each point is an average of three measurements, and the data shown are representative of three independent experiments. Error bars, S.E.

View larger version (41K): [in this window] [in a new window]   FIG. 10. G binds preferentially to Gpa1N388D in the GDP-bound conformation. To determine whether Gpa1N388D binds to G in a guanine nucleotide-dependent manner, diploid cells (strain YPH501) were transformed with plasmids containing STE2 (receptor), STE4/18 (G), and either wild-type GPA1-GST, GPA1N388D-GST, or GST alone. The cells were disrupted in the presence of GDP (–) or GDP and AlF–4 (+), as indicated. Detergent-solubilized lysates were then immobilized on glutathione-Sepharose, washed, and eluted with SDS-PAGE sample buffer. Samples of total cell lysates (Applied) and retained proteins (Bound) were resolved by 10% SDS-PAGE and subjected to immunoblotting with antibodies (Ab) against GST or Ste4, as indicated. Data shown are representative of three independent experiments. Arrows indicate proteins specifically detected by the indicated antibodies.

The receptor was fused to a triple-FLAG epitope tag (Ste2-FLAG) and co-expressed with either wild-type Gpa1 or Gpa1^N388D as well as Ste4/18. The receptor and any associated G protein were then immunoprecipitated and resolved by gel electrophoresis and immunoblotting. As shown in Fig. 11, treatment with pheromone caused diminished binding of receptor to wild-type Gpa1. In contrast, pheromone treatment enhanced the interaction of receptor with Gpa1^N388D. Even without pheromone treatment, the receptor appeared to have a higher affinity for Gpa1^N388D than wild-type Gpa1 (Fig. 11). These data indicate that receptor activation by pheromone not only stabilizes Gpa1^N388D expression but also actively promotes assembly of a receptor-Gpa1^N388D-G complex. This complex would preclude access of endogenous wild-type Gpa1 and therefore inhibit pheromone signal propagation in a dominant-negative manner.

View larger version (53K): [in this window] [in a new window]   FIG. 11. Pheromone-dependent complex formation of Gpa1N388D and receptor. To determine whether Gpa1N388D forms a stable complex with the receptor, diploid cells (strain YPH501) were transformed with plasmids containing either tagged or untagged STE2, STE4/18 (G), and either wild-type or mutant GPA1 and then treated with 2.5 µM -factor (-MF), as indicated. Detergent-solubilized cell lysates were immunoprecipitated with anti-FLAG antibodies. Samples of total cell lysates (Applied) and purified proteins (IP) were resolved by 7.5% SDS-PAGE and subjected to immunoblotting with antibodies that detect Ste2 (FLAG Ab) and Gpa1. Data shown are representative of three independent experiments. Arrows indicate the proteins specifically detected by the indicated antibodies.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Dominant-negative mutants have long been used to study a variety of signaling proteins, most notably monomeric G proteins such as Ras (49). At least three dominant-negative Ras mutants have been identified (50–52). Extensive biochemical analysis of the most widely used mutant, RasN17 (Ser-17 Asn mutation), revealed that it competes with normal Ras for binding to guanine nucleotide exchange factors. More specifically, the mutant assumes an unactivable "dead-end" complex with the exchange factor, thereby preventing it from binding to the endogenous wild-type protein. This mechanism of action is analogous to the one proposed here, in which Gpa1^N388D is thought to act by competing with normal Gpa1 for binding to the receptor, thereby preventing activation of the pathway.

Although less widely used, dominant-negative mutations of heterotrimeric G proteins have also been described. The earliest report was from Osawa and Johnson (15), who showed that G ^G226T_s could partially inhibit -adrenergic receptor-promoted stimulation of cAMP synthesis. Simon and co-workers (16, 53) described two other dominant-negative mutants, G ^S47C_o and G ^S48C_i, and showed that these mutants lack GTP binding activity but retain G binding function. Another dominant-negative G_s mutant was constructed using multiple substitutions (17) including A366S, which decreases affinity for GDP and causes the protein to spend more time in the empty state (54), as well as G226A and E268A, two substitutions that impair binding to GTP and the conformational changes required for dissociation of G (55, 56). More recently, Berlot and co-workers (57, 58) have described a dominant-negative G_s mutant that combines G226A and A366S with multiple substitutions in the 3 5 loop region that increase receptor affinity, decrease receptor-mediated activation, and impair activation of adenylyl cyclase. Expression of this mutant at close to wild-type levels blocked signaling from the luteinizing hormone receptor to G_s by up to 97% (21).

Perhaps the best characterized dominant-negative G mutants are variants of G_o,G₁₁, and G₁₆ that were engineered to bind xanthine nucleotides instead of guanine nucleotides. Xanthine monophosphate is an intermediate in the biosynthesis of guanosine monophosphate. However, the cellular abundance of xanthine diphosphate and xanthine triphosphate is negligible, so the xanthine nucleotide-binding G proteins remain in the empty (nucleotide-free) state. Because it is the nucleotide-free form of G that has highest affinity for agonist-bound receptors, stable association with the mutant G protein makes the receptor unavailable to activate the endogenous wild-type G protein (18–20).

Gpa1^N388D was originally reported to have no activity based on its inability to rescue a gpa1 mutant (41) but was later shown to promote recovery from pheromone-induced growth arrest (30). Thus, Gpa1^N388D was long known to have properties of a dominant-negative mutant, but not recognized as such in part because of the earlier conclusion that Gpa1^N388D is incapable of binding to G. The evidence for lack of G binding, albeit negative, is as follows: (i) Gpa1^N388D failed to prevent constitutive signaling in a cell lacking the GPA1 gene (30, 41); (ii) Gpa1^N388D displayed no binding to Ste4 in the two-hy