(Received for publication, August 16, 1996, and in revised form, October 21, 1996)
From the Department of Biological Chemistry, The Johns Hopkins University, School of Medicine, Baltimore, Maryland 21205
The cAMP chemoattractant receptor, cAR1, of Dictyostelium transduces extracellular cAMP signals via G protein-dependent and G protein-independent mechanisms. While site-directed mutagenesis studies of G protein-coupled receptors have provided a host of information regarding the domains essential for various functions, many mechanistic and structural questions remain to be resolved. We therefore carried out polymerase chain reaction-mediated random mutagenesis over a large part of the cAR1 sequence (from TMIII through the proximal part of the cytoplasmic tail). We devised a rapid screen for loss-of-function mutations based on the essential role of cAR1 in the developmental program of Dictyostelium. Although there were an average of two amino acid substitutions per receptor, ~90% of the mutants were able to substitute for wild-type cAR1 when expressed in receptor null cells. About 2% were loss-of-function mutants that expressed wild-type levels of receptor protein. We used biochemical screens to select about 100 of these mutants and chose eight representative mutants for extensive characterization. These fell into distinct classes. One class had a conditional defect in cAMP binding that was reversed by high salt. Another large class had decreased affinity under all conditions. Curiously, the decreases were clustered into three discrete intervals. One of the most interesting class of mutants lost all capacity for signal transduction but was phosphorylated in response to agonist binding. This latter finding suggests that there are at least two activated states of cAR1 that can be recognized by different downstream effectors.
G protein1-coupled receptors (GPCR) constitute a large and diverse family of molecules that span the plasma membrane seven times and transduce extracellular signals using heterotrimeric G proteins (1, 2). Ligands for this family of receptors vary from small compounds like cAMP to large peptides (3-5). Despite the diversity of ligands, occupancy of these receptors typically triggers an analogous array of excitation and desensitization events (6). The occupied receptors activate heterotrimeric G proteins which, in turn, activate downstream effector molecules, such as adenylyl cyclases, ion channels, phosphodiesterases, and phospholipases (7, 8), and also mediate additional changes that do not require heterotrimeric G proteins (9). These include changes in plasma membrane ion permeability (10) and gene expression as well as processes that attenuate the receptor's ability to bind ligands and transduce signals (11-13).
Site-directed mutagenesis and chimeric receptor studies have provided information regarding the interactions of GPCRs with G proteins, the localization of ligand contact sites, and the location of domains important for activational isomerization (14, 15). In the case of adrenergic receptors, catecholamines are thought to interact with amino acid residues within a pocket formed by the transmembrane helices. The domains important in coupling to G proteins reside in small segments exiting from TMV, TMVI, and TMVII to the cytoplasmic loops (16, 17). Segments of the third intracellular loop, in particular, determine the specificity of G protein coupling (18, 19). From these studies, a general model has emerged; agonist binding rearranges interactions among the helices and transmits conformational changes to the segments exiting from membrane to cytoplasm (20). These changes are then detected by intracellular components of the signaling machinery. Despite these advances, much remains to be learned about the molecular determinants for ligand binding to G protein-coupled receptors and the sequence of events that follow.
We devised a genetic screen to isolate random mutations that alter the biological and biochemical properties of the chemoattractant receptors of Dictyostelium. This organism utilizes four homologous receptors (cAR1-cAR4), which vary in affinity for cAMP, to coordinate morphological transitions throughout a developmental program in which individual amoebae aggregate to form multicellular structures (21, 22). Cells clonally seeded on bacterial lawns divide and, by consuming the bacteria, create plaques. Within several days, the cells in the centers of these plaques are induced by starvation to aggregate and form macroscopic fruiting bodies (23). Cells lacking cAR1 or expressing only defective versions of the receptor fail to aggregate or differentiate (24) and therefore form smooth plaques, providing a convenient biological "read out" for function. This screen has allowed us to rapidly assess thousands of point mutations in the receptor and identify those affecting particular functions.
At the outset, we expected to find mutants which failed to bind cAMP or were defective in coupling to G proteins or G protein-independent pathways. We found that most amino acid substitutions did not impair receptor function. Of those that did, we saw a large array of phenotypes that fell into multiple classes. In some mutants, the cAMP binding affinity was decreased conditionally at low ionic strength or, alternatively, decreased under all conditions to one of three discrete steps. In other mutants, cAMP binding and agonist-induced phosphorylation were intact, but signaling through G protein-dependent and G protein-independent pathways was defective. These mutant classes complement those of a companion study (50), which describes a series of mutants derived from random mutagenesis of the third intracellular loop.
The
1.3-kilobase pair cAR1 cDNA, EcoRI-EcoRI
fragment, was subcloned into the EcoRI site of pPL1 (25) in
BglII (5-end)-BamHI (3
-end) orientation to
construct pJK53. This plasmid was used as a template for PCR under
conditions designed to induce frequent misincorporation of nucleotides
(26). Briefly, 1 ng of pJK53 PCR-template was incubated with 30 ng each
of primers T3 and T7, 1 mM each of dCTP, dGTP, and dTTP,
0.2 mM dATP, 7 mM MgCl2, 0.5 mM MnCl2, and 5 units Taq
thermostable DNA polymerase (Perkin-Elmer) in 10 mM Tris,
pH 8.3, in a final volume of 100 µl. The reaction was carried out in
a thermal cycler (Perkin-Elmer) for 30 cycles with 94 °C, 1.5-min
denaturation, 45 °C, 2-min annealing, and 72 °C, 2-min extension.
The resulting PCR products were extracted with phenol-chloroform, then
digested simultaneously with BamHI, NdeI, and
BstXI. The mutagenized NdeI/BstXI
fragment (583 base pairs), encoding the amino acids from the third to
the seventh cAR1 transmembrane domains, was gel-purified and used to
replace the corresponding wild-type NdeI/BstXI
fragment of pJK53. The resulting mutant cAR1 cDNAs were excised
with BamHI/BstXI and directionally subcloned into
Dictyostelium expression vector, pMC34 (27).
Transformation of the
mutant cAR1 library into car1 or
car1
/car3
cells was
performed by electroporation as described previously (27). One day
after electroporation, nonselective medium was replaced with HL-5
containing G418 (Geneticin) (20 µg/ml), and for each transformation,
4 × 106 cells were placed into each well of a 24-well
plate. For the initial aggregation assay, approximately 150 Dictyostelium clones from each of the 24 wells were seeded
onto SM agar plates with lawn of Klebsiella aerogenes. After
5-6 days of growth and development, Dictyostelium plaques
that created aggregation-deficient "smooth" plaques were picked
into HL-5 with G418 (20 µg/ml). Plasmids DNA was recovered and
transformed into bacteria as described (27). Bacterially propagated
plasmids were then transformed by electroporation into
car1
/car3
or
car1
cells for subsequent phenotypic analysis.
The mutant plasmids were sequenced using automated sequencer.
The receptor electrophoretic mobility shift assay was performed as described elsewhere (28). Immunoblotting was performed using cAR1-specific rabbit polyclonal antiserum (29). Primary antibodies were detected using alkaline phosphatase-conjugated donkey anti-rabbit antibodies and chemiluminescence (Dupont).
DB Development AssayGrowth stage cells were washed once in development buffer (DB) (30), and 10 million cells were plated onto a 35-mm 1% DB agar plates and allowed to settle for 20 min. Excess buffer was drained, and cells were left undisturbed for 24 h at 22 °C to undergo development (30).
Adenylyl Cyclase AssayCells grown in shaking culture were
washed with DB, developed with 300 nM cAMP pulses every 6 min for 6 h, and shaken rapidly for 30 min in the presence of 5 mM caffeine to inhibit the cAMP oscillation and to obtain
cells with basal level of adenylyl cyclase activity (31). After
basalation, cells were washed 4 times with ice-cold phosphate manganese
buffer (10 mM KH2 PO4, pH 6.5, 2 mM MgSO4) to remove caffeine, resuspended to
8 × 107 cells/ml, and kept on ice until use. Cells
were transferred to 22 °C, stimulated with 103
M cAMP at time 0, and the amount of
[
-32P]ATP converted into [32P]cAMP in 1 min was measured as described previously (32).
Cells were developed as in the
adenylyl cyclase assay, resuspended in phosphate manganese buffer (see
above), and placed on ice. Ten minutes before each experiment, cells
were transferred to a rotary shaker (200 rpm) at 22 °C. At time 0, 107 M cAMP and 10 mM
dithiothreitol were added to induce actin polymerization, and at
various times, the reactions were stopped by transferring the aliquots
of cells into fixing solution (3.7% formaldehyde, 10 mM
Pipes, 0.1% Trion X-100, 20 mM
K2HPO4, 5 mM EGTA, 2 mM
MgCl2, and 250 nM tetramethylrhodamine B
isothiocyanate-phalloidin, pH 6.8) (33). After staining at least 1 h at room temperature, fixed cells were pelleted in a tabletop
centrifuge (14,000 rpm, 5 min), and the pellet was resuspended into 1 ml of methanol. Rhodamine fluorescence was detected with a Perkin-Elmer
fluorescence spectrophotometer (excitation at 540 nm, emission at 575 nm).
cAMP binding assays were performed in either phosphate buffer or in the presence of 3 M ammonium sulfate as described previously (11). Scatchard data were analyzed using the software program LIGAND.
GTP Inhibition of BindingCells were developed as before, washed once in phosphate buffer, and resuspended to 6 × 107/ml. Cell were then lysed through 5-µm filters (Millipore), and the lysates were pelleted (5 min, 10,000 rpm SS34 rotor). The membrane pellets were washed once and resuspended in phosphate buffer to a density of 8 × 107 cell equivalent/ml and kept on ice. The amount of cAMP binding to membrane measured with 2 nM [3H]cAMP either in the presence or absence of 100 µM GTP was then assessed by spinning the membrane through silicon oil as described previously (34).
Ca2+ Influx Assay45Ca2+ entry was followed for 40 s in the presence or absence of cAMP using a standard centrifugation assay. Values shown are the ratio of Ca2+ uptake into cAMP-treated cells/Ca2+ uptake into unstimulated cells (35).
Site-directed MutagenesisOligonucleotide-directed mutagenesis of cAR1 was performed in bacteriophage M13 as described elsewhere (27) to create cDNAs encoding mutants I-22 and II-22. The resulting cDNAs were subcloned into pMC34 (27).
We used PCR under conditions of decreased fidelity to
generate a population of mutant cAR1 cDNAs (26). Each receptor
contained an average of two amino acid substitutions within the region
extending from within the third transmembrane domain to the proximal
portion of the cytoplasmic carboxyl terminus (residues 88-264). Over
2000 mutant cDNAs were subcloned into a segregating
extrachromosomal plasmid that direct expression of the receptor
throughout growth and early development (27). The plasmid library was
electroporated into car1 cells (28), and about
4000 transformants were allowed to form plaques on bacterial lawns for
phenotypic screening and subsequent characterization (Fig.
1).
The results of a typical mutant screen are illustrated in Table IA. Of 1659 transformants analyzed, nearly 90% regained fully the ability to aggregate. Of the remaining 193 clones, 177 exhibited very weak aggregation, and 16 were completely aggregation-deficient. An aggregation-deficient phenotype could result from the expression of a dysfunctional cAR1 mutant or, alternatively, from low receptor protein expression. To distinguish these possibilities, we immunoblotted 116 nonaggregating or weakly aggregating clones for cAR1 expression. About two-thirds of the clones had little or no cAR1, while the remainder had wild-type levels of the protein (Table IB). We focused our studies on the latter type of clones isolated from four independent transformations and screens.
|
We carried out several tests to define the biochemical basis of the aggregation-deficient phenotype conferred by these mutant receptors. The initial test examined agonist-induced phosphorylation. Upon cAMP binding, serine residues (Ser-303 and Ser-304) in the carboxyl-terminal cytoplasmic domain of cAR1 undergo phosphorylation, resulting in a discrete mobility retardation on SDS-PAGE (11, 36). This alteration, detected by immunoblot, provides a convenient method for assessing agonist activated receptor-kinase coupling. At steady-state, the fraction of receptors in the slower mobility form is proportional to the concentration of the applied stimulus (37). When the assay is carried out at multiple concentrations of cAMP, the EC50 typically reflects the cAMP binding affinity (38).
In our initial screen, we treated the mutant expressing clones with
three concentrations of cAMP, 0, 50 nM, and 100 µM for 15 min, sufficiently long to attain the steady
state. Since the EC50 is 35 nM (Fig.
2), at 50 nM the majority of wild-type cAR1 molecules are found in the slower mobility form. At 100 µM, even very low affinity receptors are expected to be
maximally occupied and phosphorylated (28). When 93 clones treated with
cAMP at these three concentrations were examined, the mutants fell into five categories based on the severity of the impairment (Table II). Eighteen of the clones (19.4%) exhibited a profile
identical to that seen with wild-type cAR1 (row a); the remaining 75 exhibited an altered profile. Of these, 60 (64.5%) responded fully to
100 µM but displayed only a partial or no response to 50 nM (rows b and c). The remaining 15 (16.1%) exhibited
little or no response even at 100 µM (rows d and e).
We carried out additional assays to further classify the mutants. We measured binding of [3H]cAMP to intact cells in phosphate buffer at 16 nM and in ammonium sulfate at 25 nM. Since the Kd of cAR1 under physiological conditions in phosphate buffer is 25-300 nM, the former assay is designed to quickly distinguish high from low affinity receptors. The ammonium sulfate assay is designed to measure the total number of binding sites, since addition of the salt reduces the Kd of wild-type cAR1 to about 8 nM (39). The cAMP-induced electrophoretic mobility shift profiles of the mutants corresponded with the amount of [3H]cAMP bound under physiological conditions. The amount of binding measured in the presence of ammonium sulfate also paralleled these data (data not shown).
We rescued the plasmids encoding eight representative mutants, one from
row a, three from the row d, and two from the rest to represent the
biochemical phenotypes observed in the initial screen (Table II),
retransformed them into car1 or
car1
/car3
cells, and
subjected them to more extensive characterization. With the exception
of the cAMP-induced electrophoretic mobility shift experiments which
were carried out in a car1
background, all of
the following studies were carried out in a
car1
/car3
background.
We chose these backgrounds for the following reason. Although the
car1
cells do not aggregate upon starvation
and therefore are adequate for initial identification of functionally
deficient receptors, some of the cAMP-mediated responses still persist
in the car1
cells (40, 41). On the other hand,
in the car1
/car3
cell, cAMP responsiveness is completely absent (41), and expression of
wild-type cAR1 restores all of the measured wild-type functions.
We next examined the full
concentration dependence of the cAMP-induced electrophoretic mobility
shift of the eight representative mutants (Fig. 2). On the basis of
these dose-response profiles and the cAMP binding analysis presented
below, the eight mutants fell into the following classes, I, II, and
IIIa-c. For wild-type cAR1, the EC50 was 35 nM, consistent with our previously published results (28,
38). Mutant I-21 exhibited a nearly wild-type concentration-dependence,
while mutants IIIa-21 and IIIa-22 and IIIb-21 and IIIb-22 required
about 5 and 100 times higher cAMP concentrations, respectively (Fig.
2). The two class IIIc mutants failed to show a mobility shift, even at
the highest cAMP concentrations (102 M)
tested. The electrophoretic mobility shift of three additional mutants
(IIIa-23, IIIb-23, and IIIc-23) also exhibited EC50
increases clustered around these 5-fold, 100-fold, and infinite ranges
(data not shown). Mutant II-21 displayed a dose dependence similar to that of the class IIIb mutants; however, it was placed in a separate class because of its unique binding properties, as described below. For
these 11 mutants, the full cAMP concentration dependence of this
response was consistent with that observed in the rapid two-dose screening assay (Table II). Therefore the electrophoretic mobility shift phenotypes exhibited in the initial screen, although crude, represented intrinsic, reproducible properties of the mutants, and Fig.
2 and Table II reflect the distribution of observable phenotypes.
We next carried out a more extensive phenotypic
analysis of the eight mutants. Exponentially growing cells were washed
and spread on nutrient-free agar plates. Under these conditions, cells expressing wild-type cAR1 aggregate and form fruiting bodies within 24-48 h, while the vector-transformed cells remain as a monolayer (Fig. 3, A and B). The results for
one member of each class are illustrated in Fig. 3, C-G.
The members of classes I, II, and IIIc failed to restore any sign of
aggregation (Fig. 3, C, D, and G),
while the members of the remaining two subclasses, IIIa and IIIb,
mediated aggregation to varying degrees. Cells expressing either of the
two IIIa mutants nearly always aggregated and were capable of forming
slugs and fruiting bodies with a relatively wild-type morphology (Fig.
3E). Cells expressing either of the two IIIb receptors could
also aggregate, although a smaller fraction of cells participated.
These formed either loose mounds that went on to form tiny fruiting
bodies (Fig. 3F) or tight aggregates that transformed into
normal fruiting bodies. Thus, among the class III subclasses, there is
a correlation between the efficacy of the expressed receptor and the
resulting aggregation phenotype.
cAMP Binding Analysis
We next measured the affinity of
[3H]cAMP binding to the mutant receptors. Since the
members of a given class displayed similar properties, we chose one
from each of the classes for detailed binding analysis (I-21, II-21,
IIIa-21, IIIb-21, and IIIc-21). The resulting Scatchard data are
illustrated in Fig. 4 and Table III.
Wild-type cAR1 displayed two classes of binding sites as described previously. Most (about 93%) of the sites had an affinity of 500 nM, while a small fraction had an affinity of 30 nM in phosphate buffer. Since the lower affinity sites
constitute most of the binding sites, we concentrated our analysis on
them. As shown in Fig. 4A, the affinities of the class III
mutants, measured in phosphate buffer, corresponded to the relative
EC50 values of their electrophoretic mobility shift.
IIIa-21 had a 5-7-fold lower cAMP sensitivity than wild-type cAR1, as
assessed by both EC50 and Kd; IIIb-21
had a >10-fold decreased sensitivity reaching the detection limit of
the binding assay; and IIIc-21 did not show any detectable cAMP binding
(data not shown). The affinities of the class III mutants, measured in
ammonium sulfate, were also reduced in a corresponding pattern (Fig.
4C). The progressive decreases in cAMP efficacies and
binding affinities are highly correlated for the class III mutants. The
affinities and the EC50 of the electrophoretic mobility
shift of subclasses IIIa, IIIb, and IIIc were moderate, low, and
undetectable, respectively, as summarized in Table III.
|
For mutant II-21, while the binding affinity of the receptor measured in phosphate buffer and estimated from the EC50 of the electrophoretic mobility shift was very similar to those of IIIa and IIIb mutants (Fig. 4A, Table III), its affinity measured in ammonium sulfate was as high as that of wild-type (Fig. 4C, Table III). This striking discrepancy between the two affinities is also observed for cAR2, the receptor expressed exclusively in the muticellular stages of development (42, 43).
For mutant I-21, the EC50 for the electrophoretic mobility shift was slightly higher than that of wild-type cAR1 (Fig. 2). Yet, its binding affinity measured in phosphate buffer was at least 5-fold greater than wild-type (Fig. 4B). This greater affinity of I-21 persisted when the binding was measured in ammonium sulfate (Fig. 4D, Table III).
Activation of Adenylyl CyclaseTo assess the coupling of the
mutant receptors to G protein-mediated signaling pathways, we measured
the capacity of each to mediate the activation of the adenylyl cyclase
by extracellular cAMP (44). The cells were first induced to enter the
developmental program and express the components of the adenylyl
cyclase activation pathway. The expression of adenylyl cyclase was
confirmed for each cell line by immunoblot (Fig.
5B). Intact cells were then stimulated with a
sufficiently high concentration of cAMP (103
M) to saturate the binding sites of even the lowest
affinity receptors. We verified the activation of each mutant receptor by demonstrating that the expressed receptors shifted to the slower electrophoretic mobility form after cAMP stimulation. The
car1
/car3
cells
transformed with empty vector exhibited no measurable activation of
adenylyl cyclase (Fig. 5A), while those expressing wild-type cAR1 exhibited a rapid, transient rise in activity. The mutant receptors capable of restoring aggregation competence (IIIa-21 and
IIIb-21) mediated an essentially wild-type adenylyl cyclase response.
In contrast, mutants I-21 and II-21 failed to activate adenylyl
cyclase. This observation offers an explanation for the complete
inability of these two mutants to restore aggregation competence (Fig.
3, C and D). It also suggests that mutants I-21 and II-21 are impaired in G protein activation despite the fact that
they bind cAMP and display agonist-induced phosphorylation.
Actin Polymerization Assay
As a second method to assess the ability of each mutant receptor to activate G protein-dependent signaling pathways, we monitored cAMP-stimulated actin polymerization (45). The actin polymerization assays and the adenylyl cyclase assays described above were performed in parallel as to allow a reliable comparison between the results. When the stimulus was applied, a rapid and transient increase in F-actin was observed in cells transformed with wild-type cAR1; vector control transformants showed no response (Fig. 5C). The mutants that restored aggregation (IIIa-21, IIIb-21) responded fully, while those that did not rescue the phenotype (I-21 and II-21) showed no agonist-induced F-actin response.
Separation of Two Mutations in Mutant II-21As noted above,
mutant II-21 displayed very low affinity in phosphate buffer but high
affinity in ammonium sulfate, which is reminiscent of cAR2. Unlike
cAR2, however, which can activate cAMP-mediated adenylyl cyclase and
actin polymerization (reviewed in Kim et al. (46)), II-21
did not transduce either signals (Fig. 5, A and
C). Sequence analysis revealed that II-21 had two substitutions (R160S and H191P) in the second extracellular loop and in
the third intracellular loop, respectively (Fig.
6A). We separated these two mutations by
site-directed mutagenesis. The resulting mutants split the phenotype of
II-21. H191P failed to transduce signals but displayed higher affinity
than cAR1 (Fig. 6, C and D) and therefore was
classified as I-22; R160S retained the binding characteristics of the
original II-21 mutant but effectively transduced signals (Fig. 6,
C and D, and Table IV) and was
thus classified as II-22. The cAMP binding affinities of mutant I-22 measured in both phosphate buffer (38 and 250 nM) and
ammonium sulfate (6 nM) were higher than wild-type (Fig. 6,
C and D, and Table III). The EC50 of
the electrophoretic mobility shift was similar to that of wild-type.
I-22 failed to rescue development and only weakly activated adenylyl
cyclase and actin polymerization (Table IV). For II-22, the
EC50 of the electrophoretic mobility shift was 500 nM and the Kd in phosphate buffer was 5 µM; however, its affinity, measured in ammonium sulfate,
was similar to wild-type (3 and 100 nM). This mutant was
able to effectively activate adenylyl cyclase and rescue development
(Table IV).
|
To correlate the observed functional
defects exhibited by these mutant receptors with the predicted
structure of cAR1 (30), we sequenced each of the mutant cDNAs
characterized in Table IV. The predicted topological positions of each
of the amino acid substitutions observed for each mutant are
illustrated in Fig. 7. Mutant I-21 has a single
substitution, I108N, at the interface of TMIII and the second
intracellular loop. The class III receptors have substitutions
distributed throughout the mutagenized region. IIIa-21 possesses a
single substitution, N210S, at the base of TMVI in the interface with
the cytoplasmic loop. IIIa-22 has two substitutions, I171V and Y259H,
at the base of the helices TMV and TMVII, respectively. IIIb-21 has a
single substitution, N229D, that introduces a negatively charged
residue into the proximal half of the third extracellular loop. IIIb-22
possesses 4 amino acid substitutions, L107S, F164Y, N197I, and I208Y,
between the distal end of TMIII and the middle of TMVI. IIIc-21 has two
substitutions, S90R and V189A, which introduce a positively charged
residue at the top of helix TMIII and a conservative change in the
third intracellular loop. IIIc-22 also contains two conservative
substitutions, T97A and V136T, one in the middle of TMIII and another
at the base of TMV.
In this study, we used random mutagenesis in conjunction with a
biological screen and biochemical analyses to dissect the functions of
a G protein-coupled receptor. We were thereby able to broadly survey
the types of functional defects that mutations can create. An advantage
of this approach over site-directed mutagenesis is the larger number of
mutants that can be analyzed and the lack of bias in correlating
mutations with expected defects. Nearly 2000 receptor mutants we
screened possessed an average of two amino acid substitutions. Despite
the extent of mutagenesis, nearly 90% of all mutants were able to
rescue the aggregation-deficient phenotype of
car1 cells. These mutant receptors must have
been properly folded, transported to the plasma membrane, and deployed
at the cell surface to display nearly wild-type properties of binding,
signal transduction, and desensitization. Of the 12% of mutant
receptors which failed to restore aggregation, or did so weakly, about
one-third expressed wild-type levels of cAR1 protein. Using the
cAMP-induced electrophoretic mobility shift of cAR1 as a rapid screen,
we demonstrated that most of these mutants retained some extent of cAMP
binding and receptor-kinase interaction. These findings suggest that
there is a great range of substitutions within the cAR1 sequence that can be accommodated with little effect on functions. This supports and
extends our previous observations on cAR1 third intracellular mutants
and on a series of random chimeras between cAR1 and cAR2 (27, 28).
Other investigators have reported that over 80% of randomly mutated G
protein-coupled yeast pheromone receptors, bearing substitutions within
their third intracellular loops, are capable of restoring pheromone
responsiveness to receptor null cells (47).
On the basis of their biological and biochemical phenotypes, the mutants obtained in this study can be grouped into several classes. Class III mutants, which constitute the largest group obtained in our screen, are specifically defective in ligand binding affinity. Particularly intriguing is the fact that the cAMP sensitivities of these mutants, as assessed by cAMP binding and the cAMP concentration dependence of receptor phosphorylation, were not distributed in a smooth continuum. Rather, these mutants fell into discrete subclasses, each possessing a narrow range of sensitivity with values of 250 nM (class IIIa), 5 µM (class IIIb), and infinity (class IIIc). This pattern of sensitivities raises the possibility that the deficits exhibited by class III mutants are modular; they might represent the selective disruption of one or more cAMP contact sites or, alternatively, of interactions of cAR1 with one or more cAR-associated proteins.
The IIIa mutants have conservative replacements in or at the base of the putative transmembrane helices. IIIa-22 contains a single substitution at the base of TMVI, while IIIa-21 contains two substitutions. Based on the importance of TMVII in receptor function shown in GPCRs, it is possible that the Y259H change at the base of TMVII is the residue that is responsible for the phenotype of this mutant receptor. IIIb-21 has a negatively charged residue in the distal portion of extracellular loop III, while IIIb-22 has four substitutions at four distinct regions and is difficult to interpret. Of the two substitutions in IIIc-21, the residue mutated within the third intracellular loop, V189, has been substituted with A, L, and F without significant effect in other studies (27, 50). The second mutation introduces a positively charged residue at the beginning of TMIII (S90R) and could have a more disruptive effect on receptor function. In contrast, the drastic phenotypic impairment of IIIc-21 is more difficult to understand since both substitutions are relatively conservative (T97A, V136T).
The class II mutants exhibit a markedly increased EC50 of electrophoretic mobility shift and a parallel reduction in cAMP binding affinity under physiological conditions. However, in the presence of 3 M ammonium sulfate, they have high affinity similar to cAR1 and thus have properties similar to those of wild-type cAR2, expressed during multicellular development (42, 43). These effects of ammonium sulfate on cAMP-binding are not fully understood. We have previously explored the cAMP binding properties of cAR2 by constructing a series of cAR1-cAR2 chimeras and mapping a domain of cAR1 which, when inserted into cAR2, conferred high affinity binding under physiological conditions. This domain is the carboxyl-terminal half of the second extracellular loop, which includes five amino acid differences between the two receptor subtypes (28). There are two extra negatively charged residues in cAR2, resulting in a net negative charge, compared to a net positive charge in cAR1 (48). Based on these observations, we proposed that the second extracellular loop plays an unique role in modulating ligand binding. In this study we found that the removal of a positive charge (R160S) in the same domain results in a similar conditional reduction of affinity. Thus, the changes in the net charge of the second extracellular loop appears to modulate access to an "intrinsic" binding site and thereby lower the apparent affinity, in effect acting as a "gatekeeper" region rather than as an actual contact site. As mentioned earlier, mutant IIIb-21 has a substitution, N229D, that results in a net charge reduction in the third extracellular loop. Unlike class II mutants, however, IIIb-21 exhibited reduced binding affinities in both phosphate buffer and high salt assays. Thus, the conditional "gatekeeper" function we propose is very specific to the second extracellular loop.
The mutants I-21 (I108N) and I-22 (H191P) comprise a novel and
interesting class. These mutants are completely incapable of restoring
aggregation to
car1/car3
cells. This
deficiency cannot be explained by impaired cAMP binding (the affinities
of these receptors actually exceed that of wild-type receptor) or by
defects in agonist-induced phosphorylation, which was barely affected.
More likely, the inability of these mutants to restore aggregation is
due to their failure to mediate signal transduction responses
including, adenylyl cyclase, actin polymerization, and Ca2+
influx. The first two of these deficits suggest an impairment in G
protein activation. As with other GPCRs, the ligand binding affinity of
cAR1 is modulated by G protein interaction. The exceptionally high cAMP
binding affinity of the class I mutants and its insensitivity to
exogenously supplied GTP to reduce that affinity (Table IV) suggest
that they may be able to associate with but unable to activate and
release heterotrimeric G proteins. A similar mutant of rhodopsin locked
in association with the GDP-bound form of transducin has been described
(49).
Why do these mutant receptors fail to carry out certain G protein-independent responses such as ligand-induced Ca2+ influx? If they are locked in a nonproductive complex with heterotrimeric G protein, this might sterically impede G protein independent signaling as well. A second possibility is that the specific residues mutated in I-21 and I-22 might serve as contact sites shared by multiple effectors, such as G proteins, and those mediating Ca2+ influx and activation of other G protein independent signal processes. We favor the third possibility that the catalytic G protein coupling and Ca2+ influx may all require the formation of a specific activated receptor conformation distinct from that recognized by the receptor kinase. Further evidence for multiple activated states is presented in a companion study (50) where signaling mutants impaired in multiple pathways as well as in a specific function are isolated and characterized. Combined with the class I mutants of this study, these observations lead us to propose that G protein-coupled receptors can undergo multiple activational isomerization steps. Upon agonist binding, the receptor first undergoes conformational changes, which allow its interaction with a receptor-specific kinase. Then it undergoes additional changes, which allow it to interact with G proteins and with factor(s) mediating G protein-independent signaling processes.
The distribution of mutants among the different functional classes was not definitively established. However, from the initial screens, it is clear that most fall into classes IIIa and IIIb, as nearly two-thirds exhibited impaired phosphorylation at low but not high cAMP concentrations. This observation suggests that there are many more ways to selectively disrupt cAMP binding affinity than transmembrane signaling. This may reflect a relatively larger number of amino acid residues influencing ligand binding. Our observations also indicate that cAR1 can be extensively substituted and still functionally and phenotypically replace wild-type receptors. This sequence flexibility may account for the ability of GPCRs of widely different sequences to perform a similar set of biochemical functions and explain why during the course of evolution, changes in ligand binding affinity and specificity occur more readily than changes in signaling functions. Moreover, those binding alterations may occur in quantitatively discrete steps, as in the class III mutants of this study. Such modular affinity alterations could account for the polymorphisms in hormonal and pharmacological responsiveness exhibited by the human population.
We thank the Genetic Resources CORE Facility at Johns Hopkins University for sequencing the mutant cDNA sequences.