Common activation mechanism of class A GPCRs

Essential revisions:

1) There are more than 800 different GPCRs in human body, 45 structures were resolved in class A, and only 4 pairs of single static structures have been analyzed in details in this report. While the analysis aiming to reveal a shared GPCR activation mechanism is a meaningful attempt, it is premature to claim a "universal" activation mechanism. It would be surprising if the activation elements involved in Gs coupling and Gi coupling receptors are entirely conserved, given that the dramatic difference in the outward movement in the cytoplasmic end of TM6 and that the coupling of GPCRs with different intracellular transducers is subjected to different conformational changes. At best we can hope that the proposed mechanism provides a framework and a common language for future structural/mechanistic analysis of GPCRs, and different GPCRs (especially GPCRs of the A family) may to variously degrees and in different aspects resemble proposed mechanism. The entire manuscript, especially the title and the Abstract, needs revision in recognition of this understanding.

We thank the reviewers for the comments. In the revised manuscript, we have reworked on the title, the Abstract and related contents to ensure the accuracy and clarity.

We discuss possible misunderstandings of our work below:

First, we analyzed 6 inactive vs. fully active pairs of structures in detail (listed in Author response table 1, inactive state is in orange) but not 4 pairs, as shown in Figure 2D, Figure 2—figure supplement 1, and Results paragraph two. These structures were picked based on their high quality. That is, they are all of high-resolution, no mutation or only one mutation in the transmembrane domain which is known not to affect receptor signaling. These 6 pairs were also selected for analysis by the previous research [Venkatakrishnan, et al. Nature 536.7617 (2016): 484.].

More importantly, beyond the analysis on the six receptors, we performed a class-wide RRCS comparison between the 142 inactive and 27 active state class A GPCR structures to identify residue pairs showing statistically significant different RRCS scores (P<0.001; two sample t-test) upon activation (Figure 2D, Figure 2—source data 2, Materials and methods). The combination of such an analysis and the across-class comparison could take the full advantage of tremendous successes of GPCR structural biology in the past decade, and only this exercise would give us the confidence to claim what we discovered are common to diverse class A GPCRs.

Second, the common activation pathway we discovered is a shared portion among G_s, G_i, and arrestin activation pathways of various receptors stimulated by various agonists. At the same time, receptors that do have their own receptor-, agonist-, and effector-specific pathways are not the focus of this study. Therefore, we totally agree that activation elements involved in G_s and G_i coupling are NOT entirely conserved (i.e., biased signaling), and we had avoided to use this term in the manuscript.

2) Within this four detailed analyzed structure, RHO actually doesn't have agonist. It is not justified to include RHO as an example for analysis.

We agree with the reviewers on the importance of structure selection to study receptor activation.

As listed in the Author response table 1, we used a high quality (2.9 Å resolution) structure (PDB: 3PQR) as the fully active state structure of Rho, which has all-trans-retinal (i.e., agonist) covalently bound.

In fact, we did investigate a low resolution (4.5 Å for the overall complex, in local resolution map, the receptor part has even a lower resolution) cryo-EM structure of the Rho-G_i complex (PDB: 6CMO), in which agonist is missing. It serves as a negative control for our analysis. We noticed that 6CMO has 8 pairs of residues that do not match the common activation pathway (i.e., 8 outliers out of 34 residues pairs), and we attributed these mismatches to the rather low resolution of the structure and probably the missing of agonist. The positive control is a 3.1 Å crystal structure of the Rho-mini-G_o complex (PDB: 6FUF), for which no outliers were observed (Figure 4, Figure 4—figure supplement 1 and Figure 4—figure supplement 2). These results reflect that our RRCS approach is dependent on the high-quality structures.

3) Can the authors based on their proposal provide explanation of the basis for GPCR coupling to different G proteins using the RRCS analysis?

This is a good question. After we developed the RRCS analysis, we have been trying to find possible applications for this method. In theory, this method could be used to analyze any system that have big or subtle but significant conformational changes. The limit is that it really relies on the accuracy of side chain conformations, thus high-resolution structures are the key.

We did apply the RRCS analysis to study G protein selectivity and failed to reach any meaningful conclusions. This may be caused by following reasons:

a) Lack of high resolution GPCR-G protein complex structures. Author response table 2 lists all available GPCR-G protein complexes to date (14 structures from 10 receptors). However, 6 of them have poor resolution (lines in gray) thus are not qualified for the RRCS analysis. For the other cryo-EM structures, carefully checking the local resolution for the GPCR-G protein interface but not directly use the overall resolution of the whole complex is need before doing any RRCS calculation, since in general, large G_β protein has the highest resolution in the complex. Hopefully, with more high-resolution complexes determined, RRCS analysis may help answer the question of G protein selectivity.

b) It is known that G-protein activation is a complex process^{1, 2} which may have several states. Taking NTSR1–G_i1 complexes for example, a canonical-state and a non-canonical state of G protein were determined showing different hNTSR1–G_i1 interfaces. The complexity of this dynamic process also increases the difficulty of RRCS analysis.

c) The G protein coupling region is much more diverse in sequence compared to intracellular half of the TM domain in which our common activation pathway locates. Thus, different receptors may recognize different positions of the G-protein through distinct residues, like multiple keys (receptors) opening the same lock (G protein) using non-identical cuts, as a previous study pointed out.³

4) The author always assumed that ligand binding is located above the highly conserved W6.48, but there are more and more new structures indicated that the ligand could actually diffuse as deep as below D2.50. This is represented by the structure of LTB4 receptor (Nat. Chem. Biol. 14: 262-269) as well as several others. The revision should discuss how a deeper ligand binding would affect their conclusion.

We thank the reviewers for providing the valuable insights. We took this suggestion and added analysis for several crystal structures (PDB: 5X33, 6K1Q and 6BQH, Author response table 3), the ligand of which diffuses as deep as below D^2.50. These ligands are all inverse agonists that touch/occupy the Na⁺ pocket. By taking the space of the Na⁺, they prevented the collapse of Na⁺ pocket, as well as blocking the rotation of W^6.48, leading to lock the receptors to an inactive state.

These observations are consistent with our activation model and functional study of A_2AR mutants. As shown in Figure 6, Figure 6—source data 1, Figure 6—source data 2 and Figure 6—figure supplement 1, three mutations of A_2AR (L48^2x46R, N280^7x45R and N284^7x49K) that form slat bridges with D^2.50 and stabilize the inactive state were shown to abolish cAMP accumulation as expected.

Interestingly, for D^2.50, only loss-of-function disease mutations (Figure 7B) or constitutively inactivating mutations (Figure 7C) were observed for 24 receptors, implying that D^2.50 is indispensable for receptor activation. These finding are also supported by the NMR study of A_2AR⁴, which demonstrated the role of D52^2.50 as an allosteric link between the orthosteric drug binding site and the intracellular signaling surface⁴.

5) Many non-orthosteric ligand binding site have been captured in crystal structures (Sci Rep. 2019; 9: 6180; Front Pharmacol. 2018; 9: 128). How does the proposed activation mechanism provides new insight to these non-orthosteric ligand bindings.

We thank the reviewers for raising this point. Exactly as the recommended literature revealed, allosteric modulation of GPCR provides a promising potential of pharmacological intervention, due to the diversity in location, mechanism, and selectivity of allosteric ligands.

A list of reported allosteric modulators among determined structures of class A GPCRs are listed in Author response table 4, with PAMs labeled in green.

As we can see, the locations of binding site are quite diverse for GPCR allosteric modulators, and so do the mechanisms as one can expect.

Here is our brief analysis of the possible mechanisms. A) Directly altering the common activation pathway. For example, for allosteric modulators bound to the intracellular pocket of their receptor (PDB: 5X7D, 5LWE and 5T1A), they exert antagonism by preventing G protein coupling. B) Having receptor-, ligand-, and effector-specific pathways. An example is compound-6FA (in PDB 6N58), which binds to the extrahelical surface of β₂AR, but also directly interacts with G protein. C) A mixture of the above two mechanisms.

6) The activation mechanism and G protein specificity have been extensively discussed previously (e.g. Nature Structural and Molecular Biology 25(2):185-194 (2018); Nature 545: 317-322 (2017); Nature 536: 484-487 (2016); Nature Chemical Biology 14: 1059-1066(2018); Chem. Rev.20171171139-155). A summary of these discussions should be provided in an expanded Introduction, and the proposed mechanism should be contextualized with respect to the current understanding. Related, the citation of 19 references in one stroke is highly unconventional; this should be revised with a more nuanced account of the current structural understanding of GPCRs.

We thank the reviewer for the comments. We significantly rewrote the Introduction part to reflect the previous understanding of GPCR activation. Many more references are cited now.

7) Certain aspects of the proposed model have already been reported, and the authors in the revision should make a special effort in making sure that these reports are properly cited.

We thank the reviewer for this comment. We have greatly increased the citation and number of references to reflect previous findings on GPCR activation mechanism.

8) The description for the four layer elements in the activation pathway as "modular" is quite misleading. "Modular" means isolated and functionally independent, like classic DNA-binding domain and transcriptional activation domain in transcription factors, which can function separately and can be exchanged between transcriptional factors. In this paper, the modality of the four layers in GPCR activation has not been demonstrated. It is not clear whether layer elements can be exchanged between different receptors.

We changed the word modular to modular nature, we meant each layer has different function and can be activated/inactivated in a rather independent manner, as shown in our mutagenesis study (Figure 6B).

Analyzing residue pairs in each layer of the common activation pathway for intermediate structures (i.e., only agonist but not G protein bound structures) also supports the modular nature of the common activation pathway. As shown in Author response image 1, β₂AR (3PDS) only has layer 4 in the active state; 5-HT_2B (6DS0) has layers 2 and 3 partially activated, while leaving layers 1 and 4 not activated; A_2AR (2YDV) and LPA₆ (5XSZ) have layers 1-3 all partially activated, while leaving layer 4 untouched; and US28 (4XT1) has layers 2-4 in fully active state, while layer 1 is largely inactivated.

Author response image 1

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9) Compared to the diverse residue types in the Class A GPCR activation pathways, residues involved in class B GPCR activation appeared to be much more conserved as presented in the recent PTH1R receptor. Discussing the difference and similarity of class A and class B GPCR activation will broaden the interests and impact of the paper.

We thank the reviewers for this suggestion. Actually, the idea of applying RRCS on other allosteric systems including class B GPCRs is on our to-do-list. However, we have not done so due to the following reasons:

a) Different from class A GPCRs, only one receptor (GLP-1R) has been determined with both inactive and fully active state structures, while other receptors only have either antagonist-bound inactive structures or G protein-bound active structures.

b) The active state structures of class B GPCRs are all EM structures, their resolutions are generally low compared to that of crystal structures. As mentioned before, the RRCS analysis highly relies on the quality of side chain conformations. More class B structures with resolution higher than 3.0 Å are needed.

c) The primary results of RRCS on class B GPCRs revealed that RRCS could describe the well-known conformational changes including the HETX motif (Nature. 2017;546(7657):248-253⁵) very well (Author response image 2).

Author response image 2

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The comparison of the activation mechanism between classes A and B GPCRs have been frequently discussed⁶. The most notable conformational change in class B GPCRs is the sharp kink and unwinding in the middle of TM6, as observed for GLP-1R⁵/CTR⁷/CGRP⁸/PTH1R⁹. Another notable difference is the signal initiation and transduction pathway: class A has PIF/CWxP/Na⁺ pocket/DRY/NPxxY motifs, while class B has totally different conserved motifs. Despite these differences, they both couple G protein in similar orientation, highlighting the converge of signal transduction in the cytoplasmic region.

10) The authors introduced biological mutation experiment to support their ideas. It should be made clear that a positive result can only show the involvement of the mutated residue in GPCR activation, but cannot confirm a residue-residue interaction. With this in mind, the claims associated with the mutation data should be re-calibrated.

Thank the reviewer for raising this point. We were very careful when making mechanistic interpretations of the successfully predicted mutants, we always used the words “likely”, “would”, and “probably” to avoid any confirmation of residue-residue interactions, unless such interactions were reported by NMR studies.

11) The paper will be much strengthened if the mutagenesis experiments performed on A2aR are also performed for one of representative Gi coupled receptors. (Probably not the whole set of mutants but the representative CAM and CIM for a Gi coupled receptor.)

Thank the reviewer for the suggestion. We expanded our mutagenesis studies to two additional receptors: G_s-coupled 5-HT₇ and G_i-coupled 5-HT_1B. The three mutations on the G_s-coupled 5-HT₇ were designed and validated by Dr. Nevin Lambert (who designed the mutations by directly following our designs on A_2AR reported in the preprint of this manuscript. We did not interact with each other before this paper was submitted to the preprint server). The four mutations on the G_i-coupled 5-HT_1B were validated by ourselves. All the mutations are remarkably consistent with what we discovered in A_2AR. We added a new figure (Figure 6—figure supplement 2) to show the new mutagenesis data.

12) To confirm the predicted residue-residue interactions, it would be highly desirable to use NMR to show correlated signal changes of the residue pairs.

This is an excellent suggestion. Exactly, NMR study could provide comprehensive and precise information on residue contacts changes upon receptor activation. Although we were unable to perform NMR study due to 2-month time limit and lack of expertise, we collected and analyzed published NMR data, which covered several regions in the common activation pathway, including residues at 5.57, 3.40, 5.58, 7.53, the Na⁺ pocket, TM6 and DRY motif. The following is a brief summary of our analysis.

Isogai et al. in the NMR study of turkey β₁-adrenergic receptor (β₁AR)¹⁰ demonstrated that chemical shifts of the labeled V226^5.57 correlate linearly with ligand efficacies of the G protein pathway, thereby proving the involvement of residue 5.57 in receptor activation. The same NMR study also showed the participation of three residues (V129^3.40, A227^5.58, and L343^7.53) in receptor activation through NMR response to ligand binding and point mutations (V129^3.40I, A227^5.58Y, and L343^7.53Y) These results provide experimental evidence at high resolution of an extensive signal transduction network that connects the ligand binding site to the intracellular sides of TM5, TM6, and TM7.

Notably, 5.57, 3.40, 5.58 and 7.53 are nodes in our common activation pathway. Exactly as shown in Figure 5 and Figure 5—figure supplement 1, receptor activation involves the elimination of TM3-TM6 contacts, formation of TM3-TM7 and TM5-TM6 contacts, reflecting the outward movement of the cytoplasmic end of TM6 away from TM3, the inward movement of TM7 towards TM3 and the repacking of TM5 and TM6.

By using stable isotope NMR study for A_2A adenosine receptor (A_2AR)⁴,Eddy et al. observed the interplay of the ‘‘toggle switch’’ W246^6.48 with the allosteric center at D52^2.50. The observation is modelled in Figures 3 and 5 of our manuscript.

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