Abstract

G protein–coupled receptors (GPCRs) regulate a wide variety of physiological functions in response to structurally diverse ligands ranging from cations and small organic molecules to peptides and glycoproteins. For many GPCRs, structurally related ligands can have diverse efficacy profiles. To investigate the process of ligand binding and activation, we used fluorescence spectroscopy to study the ability of ligands having different efficacies to induce a specific conformational change in the human ₂-adrenoceptor (₂-AR). The 'ionic lock' is a molecular switch found in rhodopsin-family GPCRs that has been proposed to link the cytoplasmic ends of transmembrane domains 3 and 6 in the inactive state^{1, 2, 3}. We found that most partial agonists were as effective as full agonists in disrupting the ionic lock. Our results show that disruption of this important molecular switch is necessary, but not sufficient, for full activation of the ₂-AR.

Introduction

The Asp^3.49-Arg^3.50 pair (numbering according to the Ballesteros general scheme; see Methods) at the cytoplasmic end of transmembrane domain 3 (TM3) is part of the highly conserved (D/E)RY motif found in ₂-AR and other rhodopsin-family GPCRs, whereas the Glu^6.30 at the cytoplasmic end of TM6 is highly conserved in amine and opsin receptors. The ionic link between the Asp^3.49-Arg^3.50 pair and Glu^6.30 (also known as the ionic lock, Fig. 1) has been proposed to maintain GPCRs in the resting state^1–3 and was shown to be disrupted after photoactivation of rhodopsin⁴. Thus, disruption of the ionic lock represents a molecular switch that may be required for the activation of rhodopsin-family GPCRs. Previous biophysical studies on the ₂-AR suggest that agonist binding and activation occurs through at least one conformational intermediate, implying that more than one molecular switch is involved^{5, 6, 7}. These studies also show that structurally different agonists and partial agonists differ in their ability to induce specific conformational transitions.

Figure 1: Molecular model of the ₂-AR.

(a) Mutations I135W and A271C introduced for the bimane assay are shown in red and blue, respectively. The mutations introduced to remove endogenous cysteines are shown in green. The unreactive remaining cysteines are shown in yellow. The conserved positions involved in the ionic lock structure are outlined in orange. (b) Labeling of C271 with mBBr, showing the short linker introduced by the reaction. (c) Molecular model of the ₂-AR. The position of ₂-AR residues are followed by the Ballesteros general number¹⁸ in superscript, in the form X.YY, in which X refers to the TM segment and YY to the position relative to the most highly conserved amino acid in the TM segment, which is assigned an arbitrary position of 50. Bimane-labeled Cys271^6.33 and Trp135^3.54 are shown as solid spheres. In the inactive state, bimane is fluorescent (left). On activation, the movement of TM6 allows the rearrangement of the bimane and Trp135^3.54 side chains, resulting in bimane quenching and a decrease in fluorescence (right). (d) Detail of the environment of bimane. In the inactive state the ionic lock (yellow dots) between TM3 and TM6 (Asp130^3.49, Arg131^3.50 and Glu268^6.30) prevents direct contact between bimane and Trp135^3.54 (left). On activation, the ionic lock is broken and bimane can approach Trp135^3.54 (right).

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To study the effect of different classes of ligands on the ionic lock of ₂-AR, we generated a modified receptor having a fluorescence reporter that monitors movement between the cytoplasmic ends of TM3 and TM6. According to a three-dimensional model of the ₂-AR (Fig. 1), Ala271^6.33 and Ile135^3.54 are close to each other (C-C distance 11 Å) yet separated by the ionic lock. These residues are well positioned for using the bimane-tryptophan technique, which relies on the quenching of bimane fluorescence by tryptophan at near-contact distances in the 5- to 15-Å range⁸. We mutated Ala271^6.33 to cysteine on a minimal-cysteine ₂-AR background (Fig. 1a) to generate ₂-AR–5-Cys271. Cys271^6.33 in ₂-AR–5-Cys271 was more reactive than any of the remaining cysteines in ₂-AR–5-Cys271 (Supplementary Fig. 1 online) and could be specifically labeled with monobromobimane (mBBr, Compound 1). We also mutated Ile135^3.54 to tryptophan to generate ₂-AR–5-Cys271+Trp135. Our ₂-AR model predicts that in the absence of ligand, the ionic lock keeps TM6 in a distorted conformation, markedly decreasing the helical twist at the level of the highly conserved proline kink at Pro288^6.50. The bimane on Cys271^6.33 is then restricted by the ionic lock and lies in a pocket formed by residues in TM2, TM6 and TM7. The quencher Trp135^3.54 is located one turn below Arg131^3.50. Thus, the ionic lock separates the fluorophore and the quencher, and no direct contact between the two is possible (Fig. 1c,d). When the ionic lock breaks upon receptor activation, TM6 is released and moves away from TM3. This rearrangement opens a crevice in the cytoplasmic region of the receptor. The fluorophore on Cys271^6.33 is no longer restricted by the ionic lock and can approach Trp135^3.54, which should result in decreased fluorescence intensity (Fig. 1c,d). This prediction is also consistent with a previous study of the homologous residues in rhodopsin using spin probes⁴.

The modified receptors ₂-AR–5-Cys271 and ₂-AR–5-Cys271+Trp135 had ligand-binding affinities (Supplementary Table 1 online) that are not notably different from those previously reported for wild-type ₂-AR (ref. 9). Moreover, labeling of Cys271 with bimane did not substantially alter agonist-binding affinity (Supplementary Fig. 2 online). We measured the fluorescence properties of bimane bound to Cys271^6.33 in the presence and absence of Trp135^3.54 by comparing bimane-labeled ₂-AR–5-Cys271 with bimane-labeled ₂-AR–5-Cys271+Trp135. The bimane spectra for both constructs showed maximal emission at 460 nm and were very similar in shape, having nearly the same emission peak (Fig. 2a,b). Thus, the tryptophan introduced at position 135^3.54 did not substantially perturb the microenvironment of the bimane label at Cys271^6.33. After normalizing for receptor concentration, we found the brightness of the bimane to be only marginally smaller for ₂-AR–5-Cys271+Trp135 than for ₂-AR–5-Cys271, which showed that there is no substantial quenching of bimane-Cys271^6.33 by Trp135^3.54 in the inactive state of ₂-AR, as predicted by our model.

Figure 2: Fluorescence intensity of the bimane-attached Cys271.

(a) Fluorescence emission spectra of the purified bimane-labeled receptors ₂-AR–5-Cys271 (blue line) and ₂-AR–5-Cys271+Trp135 (orange line) in detergent micelles. The excitation wavelength was 390 nm. The signal was corrected by dividing the fluorescence signal by the receptor concentration and normalizing to 100% for the construct ₂-AR–5-Cys271+Trp135. The amount of receptor used for measurements was typically around 10 fmol. (b) Emission scans of bimane fluorescence before (orange) and after (green) stimulation with the full agonist isoproterenol. The intensity was normalized to 100% for the maximal intensity in the basal condition. The emission scan shows a strong reduction in fluorescence intensity but no change in the wavelength of maximal emission. (c) Maximal bimane responses to the full agonists isoproterenol and epinephrine, the partial agonist salbutamol, the neutral antagonist alprenolol and the inverse agonist ICI-118,551. The bimane response was measured as the percentage of fluorescence intensity change over the basal signal, as measured in the absence of any ligands. The top bars show the responses for the control ₂-AR–5-Cys271. The bottom bars show the responses for ₂-AR–5-Cys271+Trp135. Data are presented as the mean s.e.m. from three to seven independent experiments. (d) Chemical structure of the ligands used for the experiment in c. Isoproterenol and epinephrine are catecholamine agonists, salbutamol is a noncatechol partial agonist, alprenolol is an antagonist and ICI-118,551 is an inverse agonist.

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We next determined the effects of ligands with different structures and efficacies on bimane fluorescence intensity. We observed a 48%, 47% and 46% decrease in bimane intensity for the full agonists isoproterenol (Compound 2) and epinephrine (Compound 3) and the noncatechol partial agonist salbutamol (Compound 4), respectively (Fig. 2c,d). The antagonist alprenolol (Compound 5) and the inverse agonist ICI-118,551 (Compound 6) had very little effect on the bimane fluorescence intensity. The responses to agonists were specifically due to tryptophan quenching, as we obtained only small upward responses in the control construct ₂-AR–5-Cys271, in which Trp135^3.54 is absent (Fig. 2c). There was no substantial change in the wavelength of the emission peak after agonist treatment (Fig. 2b), suggesting a quenching interaction with no change in polarity around the bimane fluorophore.

The conformational transition detected by the quenching of bimane fluorescence (that is, the disruption of the ionic lock between TM3 and TM6; Fig. 1c,d) is therefore specific for agonists. To discover the structural determinants of agonists that are responsible for this specific conformational change, we measured the fluorescence response to a set of catechol and noncatechol agonists having different affinities and efficacies, ranging from full agonists to weak partial agonists. We determined the dose response for the catecholamine agonists isoproterenol, epinephrine, norepinephrine (Compound 7) and dopamine (Compound 8), as well as for the noncatecholamine partial agonists salbutamol and halostachine (Compound 9), and for catechol (Compound 10; Fig. 3). All catecholamine agonists gave the same maximal response in the bimane assay. This result was notable because dopamine is a weak partial agonist for the wild-type, unlabeled receptor. To confirm that dopamine is also a partial agonist for the bimane-labeled ₂-AR–5-Cys271+Trp135, we reconstituted the purified, labeled receptor with stimulatory G protein for adenylyl cyclase (G_s) and measured agonist-stimulated [-³⁵S]GTP binding. It should be noted that G protein activation by ₂-AR–5-Cys271+Trp135 is impaired relative to wild-type ₂-AR, most likely owing to a steric effect of the Trp135 mutation; nevertheless, we were able to detect a significant difference in [-³⁵S]GTP binding stimulated by dopamine and isoproterenol (P < 0.01; Fig. 4a). Thus, dopamine is as efficacious as isoproterenol at disrupting the ionic lock, yet this conformational change is not sufficient to induce full activation of the receptor.

Figure 3: Bimane response to catechol and noncatechol agonists.

(a) Dose-dependent bimane response of bimane-labeled ₂-AR–5-Cys271+Trp135 for the catecholamine agonists isoproterenol (black, filled circle), epinephrine (dark green, outlined circle), norepinephrine (dark cyan, filled square), and dopamine (blue, filled triangle) and for the noncatechol partial agonists salbutamol (orange, outlined square) and halostachine (red, outlined triangle). The chart also shows the absence of response to a saturating dose of catechol (1 mM, purple diamond, dotted line). The response is expressed as the percentage of the reponse to a saturating dose of isoproterenol. Data are presented as the mean s.e.m. from three to seven independent experiments. (b) Binding of bimane-labeled ₂-AR–5-Cys271+Trp135 to isoproterenol, epinephrine, norepinephrine, dopamine, salbutamol and halsotachine measured in competition with the radioligand [³H]DHA. The results are expressed as percentage of bound radioligand in the absence of competitor and presented as the mean s.e.m. of three independent experiments performed in triplicate. (c) Chemical structure of the ligands used for the experiments in a and b.

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Figure 4: Functional interactions of ₂-AR–5-Cys271+Trp135 with dopamine and catechol.

(a) Dopamine and isoproterenol stimulated binding of [-³⁵S]GTP to purified G_s reconstituted with purified bimane-labeled ₂-AR–5-Cys271+Trp135. The data are representative of two independent experiments performed in triplicate, shown as mean s.e.m. (b,c) Competition binding analysis on purified bimane-labeled ₂-AR–5-Cys271+Trp135 reconstituted into phospholipid vesicles. We performed competition binding studies using 1 nM [³H]DHA together with the indicated concentrations of isoproterenol (b) or salbutamol (c) in the absence or presence of 10 mM catechol. Catechol competes with isoproterenol (b), but not with salbutamol (c), for binding to bimane-labeled ₂-AR–5-Cys271+Trp135. Binding experiments represent the average of three independent experiments performed in triplicate and are shown as mean s.e.m.

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Noncatechol partial agonists also triggered changes in bimane fluorescence, with salbutamol inducing 95% of the response observed for isoproterenol (whereas halostachine, a very weak partial agonist, induced 50% of the response seen for isoproterenol). For all of these ligands, the effector concentration for half-maximum response (EC₅₀) in the bimane assay was consistent with the affinity measured in competition binding assay (Fig. 3b).

Catechol was the only partial agonist that produced no quenching of bimane fluorescence. We previously showed that catechol (i) induces a conformational response in ₂-AR labeled with tetramethylrhodamine on Cys265^6.27 and (ii) is a weak partial agonist in a [-³⁵S]GTP binding assay on wild-type ₂-AR⁷. We also found that catechol competes for binding with catecholamines but not with noncatechol partial agonists or antagonists. We proposed that catechol occupies the same binding site as the catechol ring in catecholamines and is thereby able to activate another molecular switch, a rotamer toggle switch in TM6 that has been suggested to be involved in the activation of amine and opsin receptor families¹⁰. On binding, the aromatic catechol ring of catecholamines presumably has a direct interaction with the aromatic residues of the rotamer toggle switch, Trp286^6.48 and Phe290^6.52. Monte Carlo simulations suggest that rotamer configurations of Cys285^6.47, Trp286^6.48 and Phe290^6.52, the residues that comprise the rotamer toggle switch, are coupled and modulate the bend angle of TM6 around the highly conserved proline kink at Pro288^6.50, leading to the movement of the cytoplasmic end of TM6 on activation¹⁰. We found that a saturating concentration of catechol had no effect on the ionic lock (Fig. 3a), even though we could detect catechol binding to bimane-labeled ₂-AR–5-Cys271+Trp135 in a modified competition assay (Fig. 4b,c). Thus, catechol may be a weak agonist that triggers the rotamer toggle switch in TM6 but does not disrupt the ionic lock between TM3 and TM6. This proposal is consistent with the recent two-dimensional crystal structure of metarhodopsin I (meta-I). Aligning the electron density map of meta-I with the three-dimensional structure of inactive rhodopsin reveals no detectable rigid body movements or rotations of helices¹¹. Thus, in meta-I the ionic lock is still intact; yet there is evidence for a rearrangement around Trp265^6.48, adjacent to the retinal binding pocket. This rearrangement may involve changes in the rotameric position of Trp265^6.48 as well as a local distortion of the polypeptide backbone around Pro267^6.50 in TM6¹¹. Thus, in both rhodopsin and the ₂-AR, conformational changes close to the ligand binding pocket are not rigidly coupled to structural modifications around the ionic lock. However, it should be noted that we have not been able to detect catechol-stimulated [-³⁵S]GTP binding in bimane-labeled ₂-AR–5-Cys271+Trp135 reconstituted with Gs. Though this is likely due to the poor coupling efficiency of this modified receptor, we cannot exclude the possibility that the modifications made to ₂-AR to monitor the ionic lock altered the conformational response to catechol.

The agonist-induced change in bimane fluorescence in ₂-AR–5-Cys271+Trp135 occurs with a t_1/2 of approximately 30 s in receptors reconstituted into phospholipid vesicles (data not shown). This is considerably slower than conformational changes observed in a modified ₂-AR containing a fluorescein arsenical hairpin (FlAsH) binder site in the third intracellular loop and having cyan fluorescent protein (CFP) fused to the carboxyl terminus^{12, 13}. This modified receptor was expressed in HEK293 human embryonic kidney cells and conformational changes were detected as changes in fluorescence resonance energy transfer (FRET) between FlAsH and CFP. There are several possible explanations for the observed differences in kinetics. The use of a large fluorescence reporter such as CFP does not allow one to determine the nature of the structural change that is responsible for the change in FRET; therefore, these FRET experiments may have been detecting a different conformational switch. Our previous studies provide evidence that agonist binding and activation occurs through a sequence of conformational changes having different rates^{6, 7}. However, the different rates observed could also be attributed to the fact that here we have studied purified receptors in the absence of G protein, whereas the FRET experiments were performed on receptors expressed in cell membranes that contained G proteins. Receptors form complexes with G proteins in the plasma membrane (an event called precoupling), and these complexes have a higher affinity for agonists than do receptors alone. Other factors that might contribute to differences between the results of our experiments using purified receptors and those of our experiments using receptors in cells could include the influence of the pH and salt gradients across the plasma membrane in living cells and the asymmetry of plasma membrane lipids. Finally, it is possible that the fluorescence changes we have observed reflect a conformational change that occurs following the disruption of the ionic lock and at a slower rate.

The structural basis of agonist efficacy in GPCRs is poorly understood. Using the evidence presented here and in recently published manuscripts^{6, 7}, we propose a model in which agonists stabilize partially or fully active states by using different chemical groups to activate different combinations of molecular switches that are not necessarily interdependent. In the unliganded, inactive state of a GPCR, the arrangement of transmembrane segments is stabilized by noncovalent interactions among side chains. Structurally distinct ligands are able to break different combinations of the basal state–stabilizing interactions either directly by binding to amino acids that are involved in these intramolecular interactions, or indirectly by stabilizing new intramolecular interactions. These ligand-specific conformational changes may be responsible for differential activation of the signaling cascades of the receptor. The affinity of a particular ligand is then dependent on the energy costs and gains associated with each disrupted and created interaction, whereas its efficacy is dependent on the ability to trigger the switches associated with activation. These molecular switches are normally activated by agonist binding^{10, 14}, but they are also revealed in constitutively active mutants, in which single point mutations in virtually any structural domain can lead to elevated basal activity¹⁵.

Based on our previous work⁷ and on studies presented here, we observe that structurally different agonists have different effects on the rotamer toggle switch and the ionic lock (Table 1). Dopamine represents the minimal structure required for activation of both the rotamer toggle switch and the ionic lock, yet dopamine is not a full agonist. Thus, additional switches must be triggered by interactions between ₂-AR and structural components on norepinephrine, epinephrine and isoproterenol that are lacking in dopamine. These structural components include the chiral -OH group and the amine substituent (Fig. 3c). The switches activated by these interactions may be involved in G protein activation as well as in other proteic interactions such as arrestin recruitment. A full understanding of these molecular switches will have direct repercussions in drug design, allowing better control of drug efficacy and affinity.

Table 1: Summary of the effect of different ligands on two molecular switches in the ₂-AR

Full table

Acknowledgments

This work was supported by grants from the US National Institutes of Health (Grant 5 RO1 NS28471 to B.K. and Grant R01 DA14896 to D.F.) and the Mather's Charitable Foundation (to B.K.).

Competing interests statement:

The authors declare no competing financial interests.

Received 20 December 2005; Accepted 16 May 2006; Published online 25 June 2006.

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1. Department of Molecular and Cellular Physiology, Stanford University School of Medicine, 279 Campus Drive, Stanford, Palo Alto, California 94305, USA.
2. Departments of Biochemistry and Molecular Biology, Oregon Health & Science University, 3181 Southwest Sam Jackson Park Drive, Portland, Oregon 97201-3098, USA.
3. These authors contributed equally to this work.

Correspondence to: Brian Kobilka¹ e-mail: kobilka@stanford.edu

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