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Vol. 19, Issue 1, 181-197, January 2008

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Distinct Roles for Two G–Gβ Interfaces in Cell Polarity Control by a Yeast Heterotrimeric G Protein

Department of Molecular Genetics and Microbiology, University of Massachusetts Medical School, Worcester, MA 01605

Submitted April 27, 2007; Revised October 9, 2007; Accepted October 22, 2007
Monitoring Editor: Daniel Lew

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

 
Saccharomyces cerevisiae mating pheromones trigger dissociation of a heterotrimeric G protein (Gβ) into G-guanosine triphosphate (GTP) and Gβ. The Gβ dimer regulates both mitogen-activated protein (MAP) kinase cascade signaling and cell polarization. Here, by independently activating the MAP kinase pathway, we studied the polarity role of Gβ in isolation from its signaling role. MAP kinase signaling alone could induce cell asymmetry but not directional growth. Surprisingly, active Gβ, either alone or with G-GTP, could not organize a persistent polarization axis. Instead, following pheromone gradients (chemotropism) or directional growth without pheromone gradients (de novo polarization) required an intact receptor–Gβ module and GTP hydrolysis by G. Our results indicate that chemoattractant-induced cell polarization requires continuous receptor–Gβ communication but not modulation of MAP kinase signaling. To explore regulation of Gβ by G, we mutated Gβ residues in two structurally distinct G–Gβ binding interfaces. Polarity control was disrupted only by mutations in the N-terminal interface, and not the Switch interface. Incorporation of these mutations into a Gβ–G fusion protein, which enforces subunit proximity, revealed that Switch interface dissociation regulates signaling, whereas the N-terminal interface may govern receptor–Gβ coupling. These findings raise the possibility that the Gβ heterotrimer can function in a partially dissociated state, tethered by the N-terminal interface.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

 
Cell polarization involves the asymmetric distribution of intracellular materials in a way that is usually directed toward localized instructional cues, which can be internal or external to the cell. External cues govern chemotaxis in cells such as Dictyostelium discoideum and mammalian neutrophils, such that a gradient of chemoattractant from a localized source stimulates cell polarization and directed cell movement along the gradient (Iijima et al., 2002; Bagorda et al., 2006; Franca-Koh et al., 2006). The ability to coordinate the direction of cell movement with the location of the external signal implies that the initial sensing of chemoattractant and at least some of the subsequent intracellular signal transduction events do not occur uniformly within the cell but instead occur in an asymmetric, spatially restricted manner.

The mating reaction of the budding yeast Saccharomyces cerevisiae provides a model system to study how cells generate an asymmetric response to polarization stimuli. Here, the two haploid cell types (a or ) secrete cell type-specific mating pheromones (a factor or factor, respectively), which serve as chemoattractants that stimulate mating responses in cells of the opposite type (Dohlman and Thorner, 2001). Each pheromone binds a specific G protein-coupled receptor (GPCR) (Ste2 or Ste3, which recognize factor and a factor, respectively) that couples to a heterotrimeric G protein (Gβ, composed of Gpa1 [G], Ste4 [Gβ], and Ste18 [G]), which is not cell type specific. Binding of pheromone to the receptor catalyzes exchange of guanosine triphosphate (GTP) for guanosine diphosphate (GDP) on the G subunit, causing dissociation of G-GTP from the Gβ dimer. Once released from the inhibitory G subunit, Gβ can trigger signaling through a mitogen-activated protein (MAP) kinase cascade; this involves membrane recruitment of the scaffold protein Ste5, which leads to activation of the Ste5-associated kinases, Ste11 (MAP kinase kinase kinase), Ste7 (MAP kinase kinase), and Fus3 (MAP kinase [MAPK]) (Pryciak and Huntress, 1998; Mahanty et al., 1999; van Drogen et al., 2000, 2001; Winters et al., 2005). These signaling events induce cell cycle arrest and the transcription of mating specific genes (Dohlman and Thorner, 2001).

Mating pheromones also regulate cell morphology. The actin cytoskeleton and membrane trafficking systems are rearranged to form a polarized mating projection, which then grows toward the partner in a process termed chemotropism (Segall, 1993; Arkowitz, 1999; Pruyne and Bretscher, 2000). These polarization events highlight another role for Gβ. Current evidence suggests that Gβ recruits the polarity proteins Far1 and Cdc24 (Valtz et al., 1995; Butty et al., 1998; Nern and Arkowitz, 1998, 1999) to the region of plasma membrane receiving the highest pheromone dose. Cdc24 is the guanine-nucleotide exchange factor (GEF) for the Rho-family GTPase Cdc42, which controls actin organization and cell polarity (Chant, 1999; Etienne-Manneville, 2004). Far1 is an adaptor protein that binds both Gβ and Cdc24 (Butty et al., 1998; Nern and Arkowitz, 1999). Hence, communication between Gβ and Far1/Cdc24 is thought to direct Cdc42 activity to the proper site to allow polarized growth along the gradient of pheromone. This process may be facilitated by additional interactions between G and Fus3 (Metodiev et al., 2002) or between Gβ and Rho1 (Bar et al., 2003).

For chemotactic cells (Dictyostelium and neutrophils) and chemotropic cells (yeast), a gradient of chemoattractant normally serves as a spatial cue for the direction of polarization. However, these cells will still polarize when exposed to a uniform concentration of chemoattractant, implying the existence of "symmetry breaking" mechanisms that can generate asymmetric responses to symmetric signals (Sohrmann and Peter, 2003; Wedlich-Soldner and Li, 2003). In yeast, prior work reveals two ways in which a cell can polarize when pheromone is provided uniformly rather than as a gradient, which we will refer to as "default" and "de novo" polarization. Default polarization uses preexisting polarity information provided by the bud site selection proteins as a spatial cue, resulting in the formation of a mating projection at the presumptive bud site (Madden and Snyder, 1992; Dorer et al., 1995; Nern and Arkowitz, 1999). This default polarization is independent of Far1–Cdc24 communication, but it depends on bud site selection proteins such as Rsr1/Bud1 (Nern and Arkowitz, 1999). When these default sites are absent, cells can still polarize in a uniform field of pheromone, but now the axis of polarization forms de novo at random positions that bear no relationship to previous polarization sites. This de novo polarization is independent of bud site selection proteins, but it requires Far1–Cdc24 binding; when this binding is disrupted, cells initiate polarization but fail to organize a stable axis of polarized growth (Nern and Arkowitz, 1999, 2000). Thus, interactions that are required for asymmetric response to a gradient of pheromone are also required when asymmetric growth is initiated de novo.

To better understand how external signals couple with intracellular factors to generate an asymmetric response, we investigated the regulation of yeast Gβ activity in response to a gradient or uniform field of pheromone. We show that in addition to Gβ, the receptor and G are required not only to sense pheromone gradients during chemotropism but also for directionally persistent growth during de novo polarization. In addition, by using a series of mutations in Gβ (Ste4) to disrupt regulation of Gβ by G, we observed qualitatively different polarity phenotypes depending on which G-Gβ interaction interface is disrupted. Our results suggest that one G–Gβ interface controls signaling, whereas the other facilitates directional responses by allowing receptor–Gβ coupling.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

 
Yeast Strains
Yeast strains are listed in Table 1. P_GAL1-STE5N-CTM was integrated at the HIS3 locus by transformation with NheI-digested pPP1268 to create strains PPY1303, PPY1304, PPY1306, and PPY1307. P_GAL1-STE11N-STE7 was integrated at the HIS3 locus by transformation with NheI-digested pPP1270 to create strains PPY1309, PPY1310, PPY1311, PPY1312, PPY1313, PPY1314, PPY1951, and PPY1952.

Plasmid pPP2711 (CEN TRP1 GPA1) was created by polymerase chain reaction (PCR) amplification of GPA1 (–201 to +1707) and ligation as a SacI–KpnI fragment into pRS314 (Sikorski and Hieter, 1989). The GPA1 mutations Q323L and K21E R22E were transferred into pPP2711 as BsrGI–BstBI fragments from pRS316-GPA1-Q323L (Apanovitch et al., 1998) and YCplac22-GPA1-K21ER22E (Metodiev et al., 2002), to create pPP2802 and pPP2743, respectively. The GPA1 mutations E28K and E28A were generated by site-directed mutagenesis of YCpGPA1 (Stone and Reed, 1990) and pPP247 (Pryciak and Hartwell, 1996), creating plasmids pPP1501, pPP1502, pPP1503, and pPP1505. The P_GAL1-STE7 construct pPP2773 was made by transferring STE7 as a BamHI–PstI fragment from pG7 (Harris et al., 2001) into the CEN TRP1 P_GAL1 vector pPP449 (Pryciak and Huntress, 1998).

To create epitope-tagged forms of Ste4 and Gpa1, we first introduced unique NotI sites into their coding sequences. For STE4, the NotI site was added between a repeat of the first two codons (ATG GCA agc ggc cgc ATG GCA), and then a myc₁₃ NotI–NotI cassette was inserted to create pPP2838 (myc₁₃-STE4) and pPP2839 (myc₁₃-STE4-GPA1 fusion). Although these derivatives were functional in vivo, the myc₁₃ epitope caused reduced signaling in FUS1-lacZ and halo assays, and thus they were used only to test effects of Ste4 mutations on protein levels and Gpa1 coimmunoprecipitation. For GPA1, intact N and C termini of the protein are functionally important. Hence, the NotI site was added within a yeast-specific insert region (Lambright et al., 1996), overlapping codons 161–163 (AgC GGc CGc) and introducing a conservative residue change (T161S). An HA₃ NotI–NotI cassette was then inserted, creating pPP2775. These derivatives were fully functional in vivo.

Isolation of Ste4 Mutants

1. A genetic screen was conducted using a two-hybrid assay to isolate Ste4 mutants with specific defects in binding Gpa1. Libraries of ste4 mutants (made by error-prone PCR) were created in a two-hybrid activation-domain vector (pGAD424) as described previously (Pryciak and Hartwell, 1996), and they were transformed into a MATa two-hybrid reporter strain (L40). Transformants were mated by replica plating to a MAT strain (AMR70) harboring DNA-binding domain (DBD) fusions to either Gpa1 (pBTM-GPA1) or Ste18 (pBTM-STE18). Clones were identified that showed reduced interaction with Gpa1 but not with Ste18. Secondary screens tested interaction with Ste5 (pM276p16) and Far1 (pPP743). All sequenced isolates harbored mutations in residues that contact G in crystal structures (Lambright et al., 1996; Sondek et al., 1996). These isolates (and their mutations) were as follows: Gm3-E41 (L154S), Gm3-C7 (K126E), Gm5-B47 (K126E D370G), Gm5-E73 (L117R), Gm4-H32 (D224V), Gm4-J38 (D224E D402E), and Gm4-I22 (S164W K213R D272A V375A). The D272A mutation in Gm4-I22 was separated from the other mutations by swapping restriction fragments; although largely responsible for the Gpa1-binding defect, the single D272A mutant was not as severe as the original quadruple mutant, and therefore all assays reported here for D272A used the stronger quadruple mutant. The isolate Gm3-E41 (L154S) was briefly reported previously (Pryciak and Hartwell, 1996). Because this L154S mutant did not show the strong chemotropism defects seen with N-terminal interface mutants (see Results), and because both Leu154 and the neighboring Asn156 residue are predicted to contact Gpa1 (Lambright et al., 1996; Sondek et al., 1996), we made an additional mutant (L154R N156K) in an attempt to cause a more severe disruption. In fact, this mutant and the original L154S mutant were effectively indistinguishable, and so only the mutant with the more-drastic substitution (L154R N156K) is analyzed here. In addition, for comparison to the phenotypically strong mutants L117R and K126E, we made less drastic substitutions at the same positions (L117A, L117G, K126A, K126N).
   
2. Another screen was designed to isolate Ste4 mutants with reduced mating efficiency but normal pathway signaling. A ste4 mutant library was created in a P_GAL1 vector (pYES-R) as described previously (Pryciak and Hartwell, 1996), and it was transformed into a MATa ste4 FUS1-lacZ strain. Transformants were mated (on galactose medium) to a mixed lawn of pheromone-producing and pheromoneless MAT cells, and screened for clones showing reduced mating only with the pheromone-producing MAT partner. The same clones were also screened for intact FUS1-lacZ induction (on galactose plates), by using a filter-based β-galactosidase assay (Bartel and Fields, 1995). Five isolates with the strongest mating defects were sequenced. One had multiple mutations and was not pursued further. Each of the others harbored mutations at Lys126 (three were K126E, one was K126N). One isolate, called m3-D10 (T76S K126E) was described previously (Pryciak and Hartwell, 1996). Because their mutations and phenotypes were similar to other K126E mutants (see 1), further analysis of these isolates is not presented.
   
3. A Ste4 mutant with Gpa1-binding defects, W136R L138F, which was isolated in a previous screen (Whiteway et al., 1994), was also studied here. This mutation was transferred by PCR from its original context (CEN HIS3 P_GAL1-STE4^{Hpl 21–1}) into pPP268, and then it was transferred to other contexts in parallel with the new mutations.
   
4. Site-directed mutagenesis was used to target Ste4 residues that were predicted to contact Gpa1 but that were not uncovered in unbiased screens. Structural data predict that G is contacted by 16 Gβ residues that lie in nine regions (Lambright et al., 1996; Sondek et al., 1996). The mutations identified from genetic screens (see 1–3) affected six of these nine regions. Therefore, the other three regions (composed of 6 residues) were mutated, using drastic residue changes to disrupt Gpa1 binding as severely as possible. These included the triple mutation N92G K94E S96A, the double mutation T181R Y183N, and the single mutation W411R. The T181R Y183N mutant was defective in all assays, and it was defective at binding all partners tested (including Ste18), suggesting it may be misfolded, and so it was not pursued further. The other mutations had only weak effects on Gpa1 binding (see Results), consistent with their absence from genetic screens.
   
5. Additional mutants used as controls were isolated in screens for Ste4 mutants with Ste18-binding defects (e.g., Gm3-F30 [D97G K380E]) (Pryciak and Hartwell, 1996) or with defects in binding Ste5 and Far1 (e.g., Gm3-J70 [L49P]). The D97G K380E mutation disrupts binding to the G subunit Ste18 and hence to Gβ-binding partners; each mutation lies close to a predicted G-contact residue (W100 and L379) (Sondek et al., 1996), although it is not known which mutation disrupts Ste18 binding (or whether both contribute). The L49P mutant was isolated in a screen to be described elsewhere, but it harbors a mutation identical to a mutation found previously (Dowell et al., 1998).
   

Microscopy
For de novo and default polarization assays, transformants were grown in selective raffinose media, and expression of P_GAL1-driven constructs was induced by addition of galactose to 2%; where indicated, 10 µM factor was also added simultaneously. After 2- to 8-h induction, cells were visualized without fixation. Differential labeling of old and new cell surface growth with fluorophore-conjugated concanavalin A (ConA) used methods described previously (Nern and Arkowitz, 2000; Matheos et al., 2004). Cells were grown first in synthetic raffinose medium, labeled with 0.1 mg/ml fluorescein isothiocyanate (FITC)-ConA, induced with 2% galactose ± 10 µM factor for 4 h, and then labeled with 0.05 mg/ml tetramethylrhodamine B isothiocyanate (TRITC)-ConA.

Mating Assays
For patch mating assays, a cells were patched directly onto a lawn of partner PT2 cells, mated overnight at 30°C, and then diploids were selected by replication to minimal media. After an immediate (1°) replica was made, more dilute replicas were made by repeating the replication of the master mating plate twice more, using a fresh velvet each time, to create 2° and 3° replicas (Harris et al., 2001). Replicas were incubated for 2 d at 30°C. For pheromone confusion assays, factor was spread on one plate to give a final concentration of 30 µM (unless indicated otherwise) and allowed to dry before the PT2 lawn was spread.

Quantitative mating assays followed previous methods (Pryciak and Huntress, 1998; Lamson et al., 2002). In brief, plasmid-transformed cells were grown overnight in selective raffinose medium, and then 0.5–5 x 10⁶ a cells were mixed with an equal number of wild-type cells (strain PT2) and collected onto filters. The filters were placed on SC/raffinose/galactose plates and incubated for 6–7 h (unless indicated otherwise) at 30°C. Afterward, filters were suspended in phosphate-buffered saline (PBS), and serial dilutions were plated on minimal media to select for diploids. Mating efficiency was calculated as the percentage of input haploid cells (before mating) that formed diploid colonies (after mating). Where indicated, factor was spread on the mating plate (to give a final concentration of 30 µM) before filters were added.

To analyze zygote formation, mated cells were suspended in PBS, sonicated, and fixed in 5% formaldehyde. Cells were imaged by differential interference contrast microscopy (DIC) and green fluorescent protein (GFP) microscopy.

Pheromone Response, β-Galactosidase, and Two-Hybrid Assays
Halo assays of growth arrest were performed by plating cells on –Trp-Ura plates and overlaying with filters containing 20 µl of 1 mM factor (Lamson et al., 2002). Plates were incubated for 2 d at 30°C.

FUS1-lacZ transcriptional induction assays were performed as described previously (Pryciak and Huntress, 1998; Lamson et al., 2002). Induction by pathway-activating constructs under control of the GAL1 promoter was measured 4 h after addition of 2% galactose ± 10 µM factor to cultures grown in raffinose medium. For FUS1-lacZ assays comparing mutant derivatives of P_GAL1-STE4 in ste4 cells, or STE4 derivatives in ste4 ste7 + P_GAL1-STE7 cells, cultures in raffinose medium were pretreated with 2% galactose for 1 h and then treated for 2 h with 2% galactose ± 10 µM factor.

β-Galactosidase and two-hybrid assays were performed as described previously (Pryciak and Hartwell, 1996; Lamson et al., 2002).

Cell Lysates and Protein Analysis
To test coimmunoprecipitation of myc₁₃-Ste4 with HA₃-Gpa1, clarified cell lysates were prepared by glass bead lysis, incubated with mouse anti-myc (9E10; Santa Cruz Biotechnology, Santa Cruz, CA) or mouse anti-hemagglutinin (HA) (HA.11; Covance Research Products, Princeton, NJ) antibodies and precipitated with protein G-agarose beads by using methods described previously (Lamson et al., 2002). After separation by SDS-polyacrylamide gel electrophoresis (PAGE) and transfer to polyvinylidene difluoride membranes, blots were probed with rabbit anti-myc (A-14; Santa Cruz Biotechnology) or rabbit anti-HA (HA.11; Covance Research Products). To analyze steady-state levels of Ste4 and Ste4-Gpa1 fusion proteins, crude whole-cell extracts were prepared by pelleting 1.5 ml of culture (OD₆₆₀ = 0.5), resuspension in 45 µl of SDS-PAGE loading buffer supplemented with protease inhibitors (Lamson et al., 2002), and boiling for 10 min. After pelleting cell debris for 2 min, 10 µl was analyzed by SDS-PAGE, blotted, and probed with rabbit anti-myc (A-14; Santa Cruz Biotechnology).

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

 
Separating the Polarity Role of Gβ from Its Signaling Role
The Gβ dimer normally performs two roles in the mating pathway: it activates MAP kinase cascade signaling and it regulates proteins that control cell polarity. Our goal in this work was to study the polarity role of the receptor–Gβ module in isolation from its requirement in activating the MAP kinase cascade. Therefore, we used a variety of methods to activate signaling independent of pheromone and Gβ, and then studied how perturbing Gβ function affects chemotropism and cell polarization. This strategy is an extension of one used previously in which overexpression of the transcription factor Ste12 was used to study the mating role of various MAP kinase pathway components while bypassing their role in transcriptional induction (Schrick et al., 1997). Here, we used several newer reagents such as membrane-targeted versions of Ste5 (Ste5N-CTM and Ste5N-Sec22), which can promote robust MAP kinase cascade signaling, wild-type levels of mating, and normal polarized morphogenesis (Pryciak and Huntress, 1998; Harris et al., 2001). To ensure that communication between Gβ and Ste5 was severed, these reagents used a truncated form of Ste5 that lacks the Gβ-binding site (Ste5N), and all assays were performed in strains where the genomic STE5 locus was deleted (ste5). For comparison, we also used a constitutively active form of Ste11 (Ste11N) (Cairns et al., 1992; Neiman and Herskowitz, 1994) and overexpressed Ste12 (Dolan and Fields, 1990; Schrick et al., 1997). When signaling output was measured by induction of a transcriptional reporter construct (FUS1-lacZ), we found that some bypass methods caused stronger activation than others, but all were independent of Ste4 (Gβ) and pheromone (Figure 1A, right).

View larger version (68K): [in this window] [in a new window]   Figure 1. Chemotropism role of Gβ and receptor is separate from signaling. (A) Chemotropism requires Ste4 even when its signaling role is bypassed. Strains PPY858 (ste5) and PPY886 (ste5 ste4) harbored galactose-inducible forms of Ste5 (pPP452), Ste5N-CTM (pPP513), Ste5N-Sec22 (pPP524), Ste11N (pPP575), or Ste12 (pPP741 or pPP271). Chemotropic proficiency (left) was assessed by patch matings performed in the absence (–) or presence (+) of exogenous factor. The mating results using Ste5 derivatives show the more-dilute 2° replica, whereas the others show the 1° replica (see Materials and Methods). Results were similar in both a and cells, and in both W303 and 381G strain backgrounds (unpublished observations). Transcriptional activation of FUS1-lacZ (right) is shown for the same strains and plasmids after galactose induction ± factor. Bars, mean ± SD (n = 3). To emphasize that transcription levels were not the primary determinant of mating efficiency, results are shown using Ste12 overexpression constructs that yield high (hi) or low (lo) levels of transcriptional induction (due to different vector contexts). (B) Ste4 can perform its chemotropism role without Ste5 and Ste20. Quantitative matings were performed ± exogenous factor. Strains PPY863 and PPY842 harbored galactose-inducible Ste11N (pRD-STE11-H3) or Ste12 (pNC252). Bars, mean ± range (n = 2). (C) An intact receptor–Gβ module is required for chemotropism. Signaling was activated by galactose-inducible constructs (pPP513, pPP575, and pPP741) in the indicated strains (PPY858, PPY979, PPY978, PPY886, and PPY989). Chemotropism was monitored by patch or quantitative mating assays in the absence (–) or presence (+) of factor. Bars, mean ± SD (for Ste5N-CTM; n = 4) or mean ± range (n = 2). Transcriptional activation of FUS1-lacZ by the same constructs after galactose induction ± factor was similar in all strains (data not shown). (D) Zygote formation. Strains from C, harboring galactose-inducible GFP-Ste5N-CTM (pPP513), were mated with PT2 partner cells for 5.5 h. Representative fields of DIC and fluorescence (GFP) images are shown. We counted 500 cells for each mixture, and the percentage that were zygotes is shown.

Gβ activates MAP kinase cascade signaling via interactions with Ste5 and Ste20 (Whiteway et al., 1995; Inouye et al., 1997; Feng et al., 1998; Leeuw et al., 1998; Pryciak and Huntress, 1998). To unequivocally determine whether these interactions are dispensable for the chemotropic role of Gβ, we performed quantitative assays of chemotropic mating in ste5 ste20 cells, by using activated Ste11 (Ste11N) or excess Ste12 to induce signaling or transcription. Indeed, these cells were proficient at chemotropism, whereas strains that also lacked Ste4 (ste4 ste5 ste20) were not (Figure 1B). Thus, the chemotropism role of Gβ does not require it to interact with Ste5 or Ste20. These findings provided a framework from which to further probe the chemotropism and polarity functions of the receptor–Gβ module without concern for their effects on MAP kinase pathway signaling.

The Pheromone Receptor and All Three Gβ Subunits Are Required for Chemotropism
The ability of Gβ to mediate chemotropism without regulating MAP kinase cascade signaling is consistent with the fact that Gβ interacts directly with polarity proteins via Far1 (Butty et al., 1998; Nern and Arkowitz, 1998, 1999). However, because chemotropism is a directional phenomenon, it would be logical that Gβ–Far1 binding could help guide polarization in the proper direction only if Gβ was activated in a spatially asymmetric manner, congruent with the pheromone gradient. Because Gβ activation is regulated by the receptor and G subunit, we directly compared the requirement for the receptor and all three G protein subunits in chemotropic mating assays. As mentioned above, MAP kinase signaling, transcription, or both were activated independent of Gβ, so that genetic perturbation of the receptor–Gβ module would affect only chemotropism. We found that only the cells with an intact receptor–Gβ module could use pheromone gradients to increase their mating success, whereas cells lacking the receptor (ste5 ste2) or any one of the G protein subunits (ste5 gpa1, ste5 ste4, or ste5 ste18) could not (Figure 1C). It is important to note that under these signaling bypass conditions the receptor–Gβ module can only contribute to mating success when pheromone gradients are intact, whereas it performs no detectable role when gradients are absent (i.e., the wild-type and null alleles of each component are indistinguishable).

Microscopic analysis (Figure 1D) confirmed that the intact receptor–Gβ module allowed cells to locate and fuse with mating partners, as judged by the formation of dumbbell-shaped zygotes, although the mating-defective cells could still form polarized mating projections. Therefore, under these conditions the receptor-Gβ module is not required for polarization per se, but for properly guiding cell polarization toward a mating partner. Note that these findings are consistent with the expectation that polarization in the "correct" direction (i.e., toward the source of pheromone) should require spatial regulation of Gβ activity, and so they do not necessarily imply that the receptor and/or G perform separate polarization functions.

Free Gβ Is Insufficient for De Novo Polarization
To study the polarity function of the receptor–Gβ module in a setting where cells do not have to detect the direction of a localized stimulus, we assayed de novo polarization in response to a uniform field of pheromone. Polarization was restricted to the de novo pathway by using rsr1 mutant strains, in which the default pathway is inactivated (Nern and Arkowitz, 1999, 2000). First, we tested whether pheromone had a role in de novo polarization beyond activating the MAP kinase cascade. Pathway signaling was activated by expressing either Ste5N-CTM (P_GAL1-STE5N-CTM) or an activated form of Ste11 fused to Ste7 (P_GAL1-STE11N-STE7), which permits normal mating morphology by reducing cross-activation of other signaling pathways (Harris et al., 2001). Despite being able to trigger default polarization (i.e., in RSR1 cells), pathway activation by Ste5N-CTM or Ste11N-Ste7 could not trigger de novo polarization (i.e., in rsr1 cells) (Figure 2A). This inability was not due to interference from excess MAP kinase pathway signaling, as cells harboring these activators (ste5 + P_GAL1-STE5N-CTM or P_GAL1-STE11N-STE7) could undergo de novo polarization when pheromone was added (Figure 2B). Importantly, we found that de novo polarization requires Gβ activity, as cells lacking the Gβ subunit (ste4 ste5 + P_GAL1-STE11N-STE7) did not polarize even when pheromone was added (Figure 2B). Notably, however, communication between Gβ and Ste5 was not required, because pheromone could stimulate polarization in cells lacking Ste5 (ste5 + P_GAL1-STE11N-STE7) (Figure 2B). As expected, transcriptional induction levels and kinetics in these strains were uninfluenced by the factors that governed polarization, such as RSR1, STE4, or pheromone (Figure 2, A and D). Therefore, as with chemotropism, the ability of pheromone and Gβ to regulate de novo polarization is separable from any regulatory effects on the MAP kinase cascade.

View larger version (39K): [in this window] [in a new window]   Figure 2. De novo polarization requires pheromone and G, in addition to Gβ. (A) Polarization was examined in RSR1 or rsr1 cells, after signaling was activated by factor (strains PPY398 and PPY1259) or by galactose induction of PGAL1-STE5N-CTM (strains PPY1303 and PPY1306) or PGAL1-STE11N-STE7 (strains PPY1309 and PPY1312). Right, the same strains were transformed with a FUS1-lacZ reporter (p3058), and transcriptional induction (mean ± SD; n = 4) was monitored 0, 2, and 4 h after addition of galactose with (i and ii) or without (iii–vi) factor. (B) Pheromone, Ste4, and Gpa1 are required for de novo polarization. The indicated rsr1 strains harboring PGAL1-STE5N-CTM or PGAL1-STE11N-STE7 (PPY1307, PPY1313, PPY1314, and PPY1952) were treated with galactose ± factor. Congenic RSR1 strains (PPY1304, PPY1310, PPY1311, and PPY1951) show that the galactose-inducible constructs can activate polarization by the default pathway. Right, old cell wall was labeled with FITC-ConA (green), and new cell wall formed during the period of mating pathway activation was labeled with TRITC-ConA (red). (C) Free Gβ is not sufficient for de novo polarization. Strains PPY794, PPY1228, PPY1248, and PPY1380 harboring PGAL1-STE4 (pGT-STE4) were induced with galactose ± factor. At right, old and new cell wall was labeled as in B. (D) Conditions affecting de novo polarization do not affect transcription. The eight strains in B were transformed with reporter plasmid p3058, and FUS1-lacZ induction (mean ± SD; n = 4) was monitored 2–4 h after addition of galactose ± factor.

To follow the pattern of cell surface growth, we used fluorophore-conjugated ConA to differentially label cell wall formed before versus during the period of mating pathway activation. It was previously shown that when de novo polarization failed due to disruption of Far1–Cdc24 interaction, cells could initiate polarized growth, but the polarization axis could not be maintained; consequently, it wandered about a broad region of the cell periphery (Nern and Arkowitz, 2000). Similarly, we observed that when cells failed de novo polarization due to absence of pheromone, Gpa1, or Ste4, they still showed asymmetric cell surface growth, but it was broadly distributed across one hemisphere (Figure 2, B and C, right). Thus, consistent with the requirement for Fus3 (Matheos et al., 2004), our results suggest that activation of the MAP kinase cascade is sufficient to initiate asymmetric growth, even in the absence of G or Gβ. Nevertheless, organization of a well-focused and persistent polarization axis requires both Far1–Cdc24 interaction and an intact receptor–Gβ module.

Gβ Mutants Reveal Distinct Roles for Two G–Gβ Binding Interfaces
To further investigate the requirement for the intact receptor–Gβ module in chemotropism and de novo polarization, we used a series of Gβ mutants to disrupt regulation of Gβ by the G subunit. Crystal structures of mammalian Gβ heterotrimers reveal two contact surfaces between G and Gβ, termed the "switch interface" and the "N-terminal interface" (Wall et al., 1995; Lambright et al., 1996). The switch (Sw) interface involves a region of G that undergoes a conformational switch upon GTP binding, whereas the N-terminal (Nt) interface involves an N-terminal helix of G that protrudes away from the globular GTPase domain (Figure 3A). The G and Gβ residues contacting one another in each interface are well conserved between mammalian and yeast counterparts (Lambright et al., 1996; Sondek et al., 1996); in Gβ these include 16 residues in nine regions. Here, we studied a series of Ste4 (Gβ) mutations that collectively affect all positions predicted to contact G. These mutants were derived from multiple sources; briefly, mutations in six of the nine G-contact regions were identified in three different genetic screens, and the remaining regions were changed by site-directed mutagenesis (see Materials and Methods).

View larger version (42K): [in this window] [in a new window]   Figure 3. Ste4 mutations in G–Gβ binding interfaces. (A) Model for orientation of the G protein-coupled receptor rhodopsin, the heterotrimeric G protein transducin, and the membrane. Adapted from Hamm, 2001; copyright Proceedings of the National Academy of Sciences USA. (B) Effects of Ste4 mutations on Gpa1 binding. Mutated Ste4 residues are listed alongside their homologous residues in mammalian Gβ1. Activation domain fusions (AD-Ste4) were expressed in a gpa1 ste11 two-hybrid tester strain (PPY1158), with DNA-binding domain fusions to Gpa1 (pPP247) or to the Gβ-binding regions of Ste5 (pPP305) or Far1 (pPP743). Binding was detected by growth on –His plates ± 3-aminotriazole (3-AT; an inhibitor of His3), or by quantitative β-galactosidase assay (bars, mean ± SD; n = 4). To reveal the full range of effects on Gpa1 binding, quantitative assays used AD–Ste4 fusions expressed from either a strong promoter (derivatives of pPP643) or a weak promoter (derivatives of pPP268); plate growth assays used pPP643 derivatives. As controls (at bottom), two additional Ste4 mutations outside the G–Gβ interfaces (see Materials and Methods) were analyzed in parallel; these disrupt binding to Ste5 and Far1, but not to Gpa1. (C) Ste4 mutations cause deregulated signaling. To avoid constitutive growth arrest, the Ste4 mutants were tested in ste4 ste7 cells (PPY1662) harboring PGAL1-STE7 (pPP2773). FUS1-lacZ induction (mean ± SD; n = 4) was measured 2 h after addition of galactose ± factor. Ste4 was expressed from the native STE4 promoter (derivatives of pPP2968; vector = pRS316). (D) Ste4 mutations disrupt coimmunoprecipitation with Gpa1. HA-tagged Gpa1 (pPP2775) or vector (pRS314) was coexpressed with myc-tagged Ste4 (derivatives of pPP2838; vector = pRS316) in strain PPY1230 (gpa1 ste4 ste5). Cell extracts were analyzed by immunoprecipitation (IP) and immunoblotting (blot).

Despite similar behavior in binding and signaling assays, mutations in the two G–Gβ interfaces had opposite effects on polarity control. Mutations in the Nt interface disrupted chemotropism, because the cells were largely insensitive to the presence or absence of pheromone gradients (Figure 4A). This phenotype is consistent with deregulated Gβ activity. Surprisingly, however, the Sw interface mutants were chemotropism proficient (Figure 4A), even though by signaling and binding criteria they seemed to be as strongly dissociated from Gpa1 as the Nt interface mutants. Although it was possible that the individual Sw mutations have a less disruptive effect on Gpa1 interaction than the Nt mutations, two further observations suggested that the explanation was not this simple. First, chemotropism remained intact even when multiple Sw interface mutations were combined, such as W136R L138F L154R N156K (WL/RF + LN/RK; Figure 4A), as well as D224V D272A, L154S D272A, and W136R L138F D272A (unpublished observations). Second, by making less drastic substitutions at Nt interface residues (i.e., K126A, K126N, L117A, and L117G), we could detect residual Gpa1 binding for several of these new mutants (as well as the original K126E); yet, they all showed a stronger defect in chemotropic mating than the Sw mutants WL/RF and LN/RK (Figure 4B).

View larger version (64K): [in this window] [in a new window]   Figure 4. Ste4 mutations in the Nt interface, but not the Sw interface, disrupt polarity control. (A) Chemotropism proficiency was assessed by patch mating assays performed in the absence (–) or presence (+) of exogenous factor. PPY867 (ste4 ste5) harbored PGAL1-STE5N-SEC22 (pPP524) and either vector (pRS316) or Ste4 variants expressed from the native STE4 promoter (derivatives of pPP2968). The effects of the Ste4 mutations on Gpa1 binding are summarized for reference (see Figure 3B). (B) Analysis of additional Nt interface mutants. Left, two-hybrid binding assays (as in Figure 3B) using AD–Ste4 fusions (derivatives of pPP643; vector=pPP636) and DBD-Gpa1 (pPP247) in PPY1158. (Note: in quantitative assays using the lacZ reporter, the Gpa1 interaction signal of Ste4-K126A was measurably reduced to 27% of Ste4-WT.) Right, quantitative assays of Ste4 function during chemotropic mating. PPY867 (ste4 ste5) harbored PGAL1-STE5N-SEC22 (pPP524) and either vector (pRS316) or Ste4 variants (derivatives of pPP2968). As Ste4 affects mating under these conditions only when gradients are intact (see Figures 1, A–C, and 4A), these assays were conducted only in the absence of exogenous factor. Results (mean ± SD; n = 4) were normalized to the average mating efficiency of wild-type Ste4. (C) Default and de novo polarization phenotypes. To avoid constitutive growth arrest, Ste4 mutants were expressed from the GAL1 promoter (derivatives of pPP1151) in PPY794 (ste4 RSR1) or PPY1248 (ste4 rsr1), and cell morphology was examined after induction with galactose ± factor.

Suppression of an Nt Interface Mutant by a Compensatory Mutation in G
First, we wanted to determine whether the stronger phenotype of the Nt mutants was truly a consequence of disrupted interaction between Gβ and G, rather than between Gβ and some other protein involved in cell polarity. Although these Ste4 mutants could still bind Far1 (Figure 3B), in principle we could not rule out effects on binding to other, unknown partners. Therefore, we attempted to restore G–Gβ binding via a compensatory mutation in Gpa1. One of the Nt interface mutations (Ste4-K126E) involves a residue that, based on mammalian Gβ structures, is expected to form an ion pair between Lys126 in Ste4 and Glu28 in Gpa1 (Figure 5A). The Ste4 mutation changes Lys126 to Glu (K126E), thereby reversing the charge. To make a compensatory charge-reversal mutation in Gpa1, we changed Glu28 to Lys (E28K). This Gpa1–E28K mutation, but not a control mutation (Gpa1-E28A), restored measurable binding with Ste4-K126E (Figure 5B), although not to wild-type levels. Also, the Gpa1 E28K and E28A mutations each reduced binding to wild-type Ste4, although by a mild degree that was most noticeable when Ste4 was expressed at lower levels from a weak promoter (Figure 5B). It was not entirely surprising that Gpa1-E28K only partially restored binding to Ste4-K126E and that Ste4-K126E caused a stronger binding defect than Gpa1-E28K, because the mammalian Gβ residue homologous to Ste4 Lys126 contacts G not only through this ion pair but also through hydrogen bonding and van der Waals interactions (Lambright et al., 1996). Nevertheless, these binding effects were enough to confer informative phenotypes in mating assays.

View larger version (24K): [in this window] [in a new window]   Figure 5. Allele-specific suppression of a Ste4 Nt interface mutant. (A) Red side chains highlight an ion pair predicted to form between Ste4(Gβ)-K126 and Gpa1(G)-E28 in the Nt interface, based on homologous residues (Gβ-K89 and G-E16) in transducin (Lambright et al., 1996). Structural rendering of Gβ used Protein Data Bank coordinate set #1GOT and Cn3D version 4.1 software (National Center for Biotechnology Information/National Library of Medicine). (B) Two-hybrid assay showing that the Gpa1-E28K mutation partially restores interaction with Ste4-K126E. DBD fusions to Gpa1 derivatives (pPP247, pPP1502, and pPP1505) were coexpressed in PPY762 with AD–Ste4 fusions under control of either a strong or weak promoter (pPP636, pPP643, pPP1121, or pPP249). Bars, mean ± SD (n = 3). (C) Gpa1-E28K partially suppresses the chemotropism defect of Ste4-K126E. Patch mating assays used ste4 gpa1 ste5 cells (PPY1230) harboring PGAL1-STE5N-SEC22 (pPP524) and the indicated combinations of GPA1 (YCpGPA1, pPP1501, pPP1503), and STE4 (derivatives of pPP226). (D) Quantitative mating assay measuring suppression of Ste4-K126E by Gpa1-E28K. Cells were as in panel C, mated to partner strain PT2 in the absence of exogenous factor. Mating frequencies from eight independent 7-h mating experiments were expressed relative to the average of the fully wild-type combination (Gpa1-WT + Ste4-WT). Bars, mean ± SEM (n = 8). For Ste4-K126E matings, the difference between Gpa1-WT and Gpa1-E28K (means, 4.1 vs. 18.7%) was ranked highly significant (p < 0.0005) by a two-tailed unpaired Student's t test.

Differences between Nt and Sw Interface Mutants Are Not Due to Altered Gpa1–Fus3 Interaction
Gpa1 can interact with the MAPK Fus3 via a docking motif in its N terminus, and mutations in this motif (Gpa1-K21E R22E, denoted Gpa1-EE) disrupt Fus3 binding and reduce mating (Metodiev et al., 2002). This raised the possibility that an intact Nt interface is required mainly to allow proper interaction between the Gpa1 N terminus and Fus3. To test this notion, we used the Gpa1-EE mutant to disrupt interaction between Gpa1 and Fus3, and we found that the Nt and Sw mutations in Ste4 still showed their characteristic phenotypic differences (Figure 6A). We also tested the Ste4 mutants in cells lacking Fus3 (ste4 fus3). Here, mating by the Sw interface mutants was not as efficient as that of wild-type Ste4, but it was still more efficient than that of the Nt interface mutants (Figure 6A). Thus, both approaches suggest that the role of the Nt interface, and the different behavior of the Nt versus Sw mutants, cannot be explained by indirect effects on the Gpa1–Fus3 interaction.

View larger version (61K): [in this window] [in a new window]   Figure 6. Qualitative differences between Ste4 Nt and Sw mutants are independent of Gpa1–Fus3 interaction. (A) Patch mating assays used the following strain and plasmid combinations. Left, strain PPY1228 (ste4 gpa1) harbored the indicated Gpa1 derivatives (pPP2711 or pPP2743) and PGAL1-STE4 plasmids (derivatives of pPP1151). Center right, strain PPY1230 (ste4 gpa1 ste5) harbored Gpa1 derivatives (pPP2711 or pPP2743) plus native STE4 plasmids (derivatives of pPP226) and either PGAL1-STE5N-CTM (pPP479) or PGAL1-STE5N-SEC22 (pPP1175). Far right, strain PPY1937 (fus3 ste4) harbored the indicated PGAL1-STE4 plasmids. (B) Pheromone confusion assay showing that fus3 cells are chemotropically proficient. Strains, from top to bottom: PPY577, PPY824, PPY827, and PPY836. Similar results were also seen in S288C and W303 strain backgrounds (unpublished observations). (C) Quantitative mating assay of strains PPY663, PPY817, PPY498, and PPY820 expressing galactose-inducible Ste12 (pPP271). Cells were mated for 18 h, ± exogenous factor. Bars, mean ± SD (n = 3).

Fusion of Gβ to G Suggests a Role for the Nt Interface in Receptor Coupling
Finally, we addressed whether maintenance of the Nt interface was necessary for coupling of the heterotrimer (Gβ) to the receptor. Existing models for the coupling between GPCRs and heterotrimeric G proteins predict that the Nt interface lies tangential to the membrane (Hamm, 1998, 2001). In addition, the N terminus of G has been implicated in receptor recognition (Taylor et al., 1994; Itoh et al., 2001; Cabrera-Vera et al., 2003). These facts suggested that the Nt interface mutations may not only disrupt interaction between Gβ and G, but they may also disrupt the way Gβ interacts with the receptor. However, the ability to test this notion was hindered by the fact that the Ste4 mutations caused constitutive signaling, which obscured whether the receptor might still exert some regulatory control over the G protein. To circumvent this difficulty, we took advantage of a previously described Ste4–Gpa1 fusion protein (Klein et al., 2000), with the rationale that forced association to Gpa1 may inhibit constitutive signaling by the Ste4 mutants and thus allow us to assay receptor coupling.

Starting with the prior Ste4–Gpa1 fusion construct, we replaced the original GAL1 promoter with the native STE4 promoter, and then compared its function with wild-type (unfused) polypeptides. By multiple assays, we found this Gβ–G fusion (Ste4-Gpa1) to function in a manner that was virtually indistinguishable from separate Gβ and G polypeptides. This included growth arrest (Figure 7A, left), regulation by the RGS-family protein Sst2 (Figure 7A, right), and pheromone-induced transcription (Figure 7B). Furthermore, the Gβ–G fusion was able to mediate total mating levels and chemotropic mating behavior that was similar to the wild-type heterotrimer (Figure 7, C and D). Thus, fusion of Gβ to G does not interfere with Gβ function in either signaling or gradient detection.

View larger version (42K): [in this window] [in a new window]   Figure 7. Phenotypes of Ste4–Gpa1 fusion proteins. (A) A Ste4–Gpa1 fusion expressed from the native STE4 promoter functions indistinguishably from separate Ste4 and Gpa1 polypeptides. Halo assays show growth arrest of ste4 gpa1 cells (PPY1228) or ste4 gpa1 sst2 cells (PPY1942) harboring Gpa1 and Ste4 as separate (YCplac22-GPA1-WT + pPP226) or fused (pPP1340 + pRS314 vector) polypeptides. (B) FUS1-lacZ assays (mean ± SD; n = 4) showing that the Ste4-Gpa1 fusion retains a normal dose response to pheromone. PPY1663 (ste4 gpa1) harbored