Structure and dynamics of GPCR signaling complexes

Abstract

G-protein-coupled receptors (GPCRs) relay numerous extracellular signals by triggering intracellular signaling through coupling with G proteins and arrestins. Recent breakthroughs in the structural determination of GPCRs and GPCR–transducer complexes represent important steps toward deciphering GPCR signal transduction at a molecular level. A full understanding of the molecular basis of GPCR-mediated signaling requires elucidation of the dynamics of receptors and their transducer complexes as well as their energy landscapes and conformational transition rates. Here, we summarize current insights into the structural plasticity of GPCR–G-protein and GPCR–arrestin complexes that underlies the regulation of the receptor’s intracellular signaling profile.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.

from$8.99

All prices are NET prices.

Fig. 1: G-protein-coupled receptor signal transduction.
Fig. 2: Receptor-mediated conformational changes in Gα.
Fig. 3: Conformational changes in arrestin-2 upon activation.
Fig. 4: Structure and interaction interface of the rhodopsin–arrestin-1 complex.

References

  1. 1.

    Kenakin, T. New concepts in pharmacological efficacy at 7TM receptors: IUPHAR review 2. Br. J. Pharmacol. 168, 554–575 (2013).

  2. 2.

    Zhang, D., Zhao, Q. & Wu, B. Structural studies of G protein-coupled receptors. Mol. Cells 38, 836–842 (2015).

  3. 3.

    Venkatakrishnan, A. J. et al. Diverse activation pathways in class A GPCRs converge near the G-protein-coupling region. Nature 536, 484–487 (2016).

  4. 4.

    Wu, F., Song, G., de Graaf, C. & Stevens, R. C. Structure and function of peptide-binding G protein-coupled receptors. J. Mol. Biol. 429, 2726–2745 (2017).

  5. 5.

    Shonberg, J., Kling, R. C., Gmeiner, P. & Löber, S. GPCR crystal structures: medicinal chemistry in the pocket. Bioorg. Med. Chem. 23, 3880–3906 (2015).

  6. 6.

    Manglik, A. & Kobilka, B. The role of protein dynamics in GPCR function: insights from the β2AR and rhodopsin. Curr. Opin. Cell Biol. 27, 136–143 (2014).

  7. 7.

    Latorraca, N. R., Venkatakrishnan, A. J. & Dror, R. O. GPCR dynamics: structures in motion. Chem. Rev. 117, 139–155 (2017).

  8. 8.

    Ye, L., Van Eps, N., Zimmer, M., Ernst, O. P. & Prosser, R. S. Activation of the A2A adenosine G-protein-coupled receptor by conformational selection. Nature 533, 265–268 (2016).

  9. 9.

    Liu, J. J., Horst, R., Katritch, V., Stevens, R. C. & Wüthrich, K. Biased signaling pathways in β2-adrenergic receptor characterized by 19F-NMR. Science 335, 1106–1110 (2012).

  10. 10.

    Okude, J. et al. Identification of a conformational equilibrium that determines the efficacy and functional selectivity of the μ-opioid receptor. Angew. Chem. Int. Edn Engl. 54, 15771–15776 (2015).

  11. 11.

    Manglik, A. et al. Structural insights into the dynamic process of β2-adrenergic receptor signaling. Cell 161, 1101–1111 (2015). NMR spectroscopy and DEER spectroscopy studies revealed the structural heterogeneity and dynamic character of the β2 adrenergic receptor.

  12. 12.

    DeVree, B. T. et al. Allosteric coupling from G protein to the agonist-binding pocket in GPCRs. Nature 535, 182–186 (2016).

  13. 13.

    Sounier, R. et al. Propagation of conformational changes during μ-opioid receptor activation. Nature 524, 375–378 (2015).

  14. 14.

    Rasmussen, S. G. F. et al. Crystal structure of the β2 adrenergic receptor-Gs protein complex. Nature 477, 549–555 (2011). This paper describes the first structure of a GPCR–G-protein complex, highlighting receptor-mediated conformational changes within the G protein that are important for nucleotide release.

  15. 15.

    Liang, Y.-L. et al. Phase-plate cryo-EM structure of a class B GPCR-G-protein complex. Nature 546, 118–123 (2017).

  16. 16.

    Carpenter, B., Nehmé, R., Warne, T., Leslie, A. G. W. & Tate, C. G. Structure of the adenosine A(2A) receptor bound to an engineered G protein. Nature 536, 104–107 (2016).

  17. 17.

    Zhang, Y. et al. Cryo-EM structure of the activated GLP-1 receptor in complex with a G protein. Nature 546, 248–253 (2017).

  18. 18.

    Kang, Y. et al. Crystal structure of rhodopsin bound to arrestin by femtosecond X-ray laser. Nature 523, 561–567 (2015).

  19. 19.

    Zhou, X. E. et al. Identification of phosphorylation codes for arrestin recruitment by G protein-coupled receptors. Cell 170, 457–469.e13 (2017).This paper describes the first high-resolution structure of a GPCR–arrestin complex and explores the role of receptor C-tail phosphate patterns in arrestin binding affinity.

  20. 20.

    Komolov, K. E. & Benovic, J. L. G protein-coupled receptor kinases: Past, present and future. Cell. Signal.  https://doi.org/10.1016/j.cellsig.2017.07.004 (2017). 

  21. 21.

    Downes, G. B. & Gautam, N. The G protein subunit gene families. Genomics 62, 544–552 (1999).

  22. 22.

    Simon, M. I., Strathmann, M. P. & Gautam, N. Diversity of G proteins in signal transduction. Science 252, 802–808 (1991).

  23. 23.

    Higashijima, T., Ferguson, K. M., Sternweis, P. C., Smigel, M. D. & Gilman, A. G. Effects of Mg2+ and the beta gamma-subunit complex on the interactions of guanine nucleotides with G proteins. J. Biol. Chem. 262, 762–766 (1987).

  24. 24.

    Kristiansen, K. Molecular mechanisms of ligand binding, signaling, and regulation within the superfamily of G-protein-coupled receptors: molecular modeling and mutagenesis approaches to receptor structure and function. Pharmacol. Ther. 103, 21–80 (2004).

  25. 25.

    Milligan, G. & Kostenis, E. Heterotrimeric G-proteins: a short history. Br. J. Pharmacol. 147, S46–S55 (2006). Suppl 1.

  26. 26.

    Smrcka, A. V. G protein βγ subunits: central mediators of G protein-coupled receptor signaling. Cell. Mol. Life Sci. 65, 2191–2214 (2008).

  27. 27.

    Khan, S. M. et al. The expanding roles of Gβγ subunits in G protein-coupled receptor signaling and drug action. Pharmacol. Rev. 65, 545–577 (2013).

  28. 28.

    Ross, E. M. & Wilkie, T. M. GTPase-activating proteins for heterotrimeric G proteins: regulators of G protein signaling (RGS) and RGS-like proteins. Annu. Rev. Biochem. 69, 795–827 (2000).

  29. 29.

    Kimple, A. J., Bosch, D. E., Giguère, P. M. & Siderovski, D. P. Regulators of G-protein signaling and their Gα substrates: promises and challenges in their use as drug discovery targets. Pharmacol. Rev. 63, 728–749 (2011).

  30. 30.

    Oldham, W. M. & Hamm, H. E. Structural basis of function in heterotrimeric G proteins. Q. Rev. Biophys. 39, 117–166 (2006).

  31. 31.

    Sprang, S. R. G protein mechanisms: insights from structural analysis. Annu. Rev. Biochem. 66, 639–678 (1997).

  32. 32.

    Noel, J. P., Hamm, H. E. & Sigler, P. B. The 2.2 Å crystal structure of transducin-alpha complexed with GTP gamma S. Nature 366, 654–663 (1993).

  33. 33.

    Van Eps, N. et al. Interaction of a G protein with an activated receptor opens the interdomain interface in the alpha subunit. Proc. Natl. Acad. Sci. USA 108, 9420–9424 (2011). In this paper, DEER distance measurements are used to demonstrate that G-protein coupling to an activated receptor induces domain separation of the AHD and the Ras domain that opens up an exit pathway for GDP.

  34. 34.

    Chung, K. Y. et al. Conformational changes in the G protein Gs induced by the β2 adrenergic receptor. Nature 477, 611–615 (2011).

  35. 35.

    Gao, Y. et al. Isolation and structure-function characterization of a signaling-active rhodopsin-G protein complex. J. Biol. Chem. 292, 14280–14289 (2017).

  36. 36.

    Grishina, G. & Berlot, C. H. A surface-exposed region of G in which substitutions decrease receptor-mediated activation and increase receptor affinity. Mol. Pharmacol. 57, 1081–1092 (2000).

  37. 37.

    Warner, D. R. & Weinstein, L. S. A mutation in the heterotrimeric stimulatory guanine nucleotide binding protein α-subunit with impaired receptor-mediated activation because of elevated GTPase activity. Proc. Natl. Acad. Sci. USA 96, 4268–4272 (1999).

  38. 38.

    Yao, X.-Q. et al. Dynamic coupling and allosteric networks in the α subunit of heterotrimeric G proteins. J. Biol. Chem. 291, 4742–4753 (2016).

  39. 39.

    Ceruso, M. A., Periole, X. & Weinstein, H. Molecular dynamics simulations of transducin: interdomain and front to back communication in activation and nucleotide exchange. J. Mol. Biol. 338, 469–481 (2004).

  40. 40.

    Dror, R. O. et al. Structural basis for nucleotide exchange in heterotrimeric G proteins. Science 348, 1361–1365 (2015).

  41. 41.

    Markby, D. W., Onrust, R. & Bourne, H. R. Separate GTP binding and GTPase activating domains of a G alpha subunit. Science 262, 1895–1901 (1993). This elegant paper demonstrates that the AHD specific for heterotrimeric G proteins shows GTPase-activating protein (GAP) activity.

  42. 42.

    Aris, L. et al. Structural requirements for the stabilization of metarhodopsin II by the C terminus of the α subunit of transducin. J. Biol. Chem. 276, 2333–2339 (2001).

  43. 43.

    Schwindinger, W. F., Miric, A., Zimmerman, D. & Levine, M. A. A novel Gs alpha mutant in a patient with Albright hereditary osteodystrophy uncouples cell surface receptors from adenylyl cyclase. J. Biol. Chem. 269, 25387–25391 (1994).

  44. 44.

    Sullivan, K. A. et al. Identification of receptor contact site involved in receptor-G protein coupling. Nature 330, 758–760 (1987).

  45. 45.

    Oldham, W. M., Van Eps, N., Preininger, A. M., Hubbell, W. L. & Hamm, H. E. Mechanism of the receptor-catalyzed activation of heterotrimeric G proteins. Nat. Struct. Mol. Biol. 13, 772–777 (2006).

  46. 46.

    Conklin, B. R., Farfel, Z., Lustig, K. D., Julius, D. & Bourne, H. R. Substitution of three amino acids switches receptor specificity of Gq α to that of Gi α. Nature 363, 274–276 (1993).

  47. 47.

    Herrmann, R. et al. Rhodopsin-transducin coupling: role of the Galpha C-terminus in nucleotide exchange catalysis. Vision Res. 46, 4582–4593 (2006). This study describes the important role of the αN–β1 hinge region for rhodopsin-mediated nucleotide release in engaged cognate Gprotein transducin.

  48. 48.

    Iiri, T., Herzmark, P., Nakamoto, J. M., van Dop, C. & Bourne, H. R. Rapid GDP release from Gs α in patients with gain and loss of endocrine function. Nature 371, 164–168 (1994).

  49. 49.

    Posner, B. A., Mixon, M. B., Wall, M. A., Sprang, S. R. & Gilman, A. G. The A326S mutant of Gialpha1 as an approximation of the receptor-bound state. J. Biol. Chem. 273, 21752–21758 (1998).

  50. 50.

    Thomas, T. C., Schmidt, C. J. & Neer, E. J. G-protein alpha o subunit: mutation of conserved cysteines identifies a subunit contact surface and alters GDP affinity. Proc. Natl. Acad. Sci. USA 90, 10295–10299 (1993).

  51. 51.

    Sun, D. et al. Probing Gαi1 protein activation at single-amino acid resolution. Nat. Struct. Mol. Biol. 22, 686–694 (2015).

  52. 52.

    Kaya, A. I. et al. A conserved hydrophobic core in Gαi1 regulates G protein activation and release from activated receptor. J. Biol. Chem. 291, 19674–19686 (2016).

  53. 53.

    Kaya, A. I. et al. A conserved phenylalanine as a relay between the α5 helix and the GDP binding region of heterotrimeric Gi protein α subunit. J. Biol. Chem. 289, 24475–24487 (2014).

  54. 54.

    Flock, T. et al. Universal allosteric mechanism for Gα activation by GPCRs. Nature 524, 173–179 (2015).

  55. 55.

    Alexander, N. S. et al. Energetic analysis of the rhodopsin-G-protein complex links the α5 helix to GDP release. Nat. Struct. Mol. Biol. 21, 56–63 (2014).

  56. 56.

    Herrmann, R., Heck, M., Henklein, P., Hofmann, K. P. & Ernst, O. P. Signal transfer from GPCRs to G proteins: role of the G alpha N-terminal region in rhodopsin-transducin coupling. J. Biol. Chem. 281, 30234–30241 (2006).

  57. 57.

    Franke, R. R., König, B., Sakmar, T. P., Khorana, H. G. & Hofmann, K. P. Rhodopsin mutants that bind but fail to activate transducin. Science 250, 123–125 (1990).

  58. 58.

    Muradov, K. G. & Artemyev, N. O. Coupling between the N- and C-terminal domains influences transducin-α intrinsic GDP/GTP exchange. Biochemistry 39, 3937–3942 (2000).

  59. 59.

    Zhang, Q., Okamura, M., Guo, Z.-D., Niwa, S. & Haga, T. Effects of partial agonists and Mg2+ ions on the interaction of M2 muscarinic acetylcholine receptor and G protein Galpha i1 subunit in the M2-Galpha i1 fusion protein. J. Biochem. 135, 589–596 (2004).

  60. 60.

    Seifert, R., Gether, U., Wenzel-Seifert, K. & Kobilka, B. K. Effects of guanine, inosine, and xanthine nucleotides on β(2)-adrenergic receptor/G(s) interactions: evidence for multiple receptor conformations. Mol. Pharmacol. 56, 348–358 (1999).

  61. 61.

    Selley, D. E., Sim, L. J., Xiao, R., Liu, Q. & Childers, S. R. mu-Opioid receptor-stimulated guanosine-5′-O-(gamma-thio)-triphosphate binding in rat thalamus and cultured cell lines: signal transduction mechanisms underlying agonist efficacy. Mol. Pharmacol. 51, 87–96 (1997).

  62. 62.

    Roberts, D. J., Lin, H. & Strange, P. G. Mechanisms of agonist action at D2 dopamine receptors. Mol. Pharmacol. 66, 1573–1579 (2004).

  63. 63.

    Gregorio, G. G. et al. Single-molecule analysis of ligand efficacy in β2AR-G-protein activation. Nature 547, 68–73 (2017). This study provides single-molecule insights into ligand-dependent allosteric regulation between the orthosteric binding site of the receptor and the nucleotide-binding pocket of the engaged G protein.

  64. 64.

    Harrison, C. & Traynor, J. R. The [35S]GTPgammaS binding assay: approaches and applications in pharmacology. Life Sci. 74, 489–508 (2003).

  65. 65.

    Toyama, Y. et al. Dynamic regulation of GDP binding to G proteins revealed by magnetic field-dependent NMR relaxation analyses. Nat. Commun. 8, 14523 (2017).

  66. 66.

    Goricanec, D. et al. Conformational dynamics of a G-protein α subunit is tightly regulated by nucleotide binding. Proc. Natl. Acad. Sci. USA 113, E3629–E3638 (2016).

  67. 67.

    Abdulaev, N. G. et al. The receptor-bound “empty pocket” state of the heterotrimeric G-protein alpha-subunit is conformationally dynamic. Biochemistry 45, 12986–12997 (2006).

  68. 68.

    Furness, S. G. B. et al. Ligand-dependent modulation of G protein conformation alters drug efficacy. Cell 167, 739–749.e11 (2016). This article describes ligand-dependent conformational differences in the calcitonin-receptor-coupled G protein that can modulate the GTP sensitivity of the ternary complex.

  69. 69.

    DeWire, S. M., Ahn, S., Lefkowitz, R. J. & Shenoy, S. K. β-arrestins and cell signaling. Annu. Rev. Physiol. 69, 483–510 (2007).

  70. 70.

    Nobles, K. N., Guan, Z., Xiao, K., Oas, T. G. & Lefkowitz, R. J. The active conformation of β-arrestin1: direct evidence for the phosphate sensor in the N-domain and conformational differences in the active states of β-arrestins1 and -2. J. Biol. Chem. 282, 21370–21381 (2007).

  71. 71.

    Gurevich, V. V. & Gurevich, E. V. Overview of different mechanisms of arrestin‐mediated signaling. Curr. Protoc. Pharmacol. 67, 2.10.1–2.10.9 (2014).

  72. 72.

    Peterson, Y. K. & Luttrell, L. M. The diverse roles of arrestin scaffolds in G protein-coupled receptor signaling. Pharmacol. Rev. 69, 256–297 (2017).

  73. 73.

    Kohout, T. A., Lin, F. S., Perry, S. J., Conner, D. A. & Lefkowitz, R. J. beta-Arrestin 1 and 2 differentially regulate heptahelical receptor signaling and trafficking. Proc. Natl. Acad. Sci. USA 98, 1601–1606 (2001).

  74. 74.

    Gurevich, E. V., Benovic, J. L. & Gurevich, V. V. Arrestin2 and arrestin3 are differentially expressed in the rat brain during postnatal development. Neuroscience 109, 421–436 (2002).

  75. 75.

    Gurevich, E. V., Benovic, J. L. & Gurevich, V. V. Arrestin2 expression selectively increases during neural differentiation. J. Neurochem 91, 1404–1416 (2004).

  76. 76.

    Scott, M. G. H. et al. Differential nucleocytoplasmic shuttling of β-arrestins. Characterization of a leucine-rich nuclear export signal in β-arrestin2. J. Biol. Chem. 277, 37693–37701 (2002).

  77. 77.

    Oakley, R. H., Laporte, S. A., Holt, J. A., Caron, M. G. & Barak, L. S. Differential affinities of visual arrestin, beta arrestin1, and beta arrestin2 for G protein-coupled receptors delineate two major classes of receptors. J. Biol. Chem. 275, 17201–17210 (2000).

  78. 78.

    Srivastava, A., Gupta, B., Gupta, C. & Shukla, A. K. Emerging functional divergence of β-arrestin isoforms in GPCR function. Trends Endocrinol. Metab. 26, 628–642 (2015).

  79. 79.

    Hirsch, J. A., Schubert, C., Gurevich, V. V. & Sigler, P. B. The 2.8 A crystal structure of visual arrestin: a model for arrestin’s regulation. Cell 97, 257–269 (1999).

  80. 80.

    Han, M., Gurevich, V. V., Vishnivetskiy, S. A., Sigler, P. B. & Schubert, C. Crystal structure of β-arrestin at 1.9 A: possible mechanism of receptor binding and membrane translocation. Structure 9, 869–880 (2001).

  81. 81.

    Zhan, X., Gimenez, L. E., Gurevich, V. V. & Spiller, B. W. Crystal structure of arrestin-3 reveals the basis of the difference in receptor binding between two non-visual subtypes. J. Mol. Biol. 406, 467–478 (2011).

  82. 82.

    Sutton, R. B. et al. Crystal structure of cone arrestin at 2.3A: evolution of receptor specificity. J. Mol. Biol. 354, 1069–1080 (2005).

  83. 83.

    Gurevich, V. V. & Gurevich, E. V. Structural determinants of arrestin functions. Prog. Mol. Biol. Transl. Sci. 118, 57–92 (2013).

  84. 84.

    Granzin, J., Stadler, A., Cousin, A., Schlesinger, R. & Batra-Safferling, R. Structural evidence for the role of polar core residue Arg175 in arrestin activation. Sci. Rep. 5, 15808 (2015).

  85. 85.

    Kim, Y. J. et al. Crystal structure of pre-activated arrestin p44. Nature 497, 142–146 (2013).

  86. 86.

    Shukla, A. K. et al. Structure of active β-arrestin-1 bound to a G-protein-coupled receptor phosphopeptide. Nature 497, 137–141 (2013).

  87. 87.

    Shukla, A. K. et al. Visualization of arrestin recruitment by a G-protein-coupled receptor. Nature 512, 218–222 (2014). This study provides direct evidence by single-particle EM for the existence of two binding modes between arrestin and receptor.

  88. 88.

    Chen, Q. et al. Structural basis of arrestin-3 activation and signaling. Nat. Commun. 8, 1427 (2017).

  89. 89.

    Ostermaier, M. K., Peterhans, C., Jaussi, R., Deupi, X. & Standfuss, J. Functional map of arrestin-1 at single amino acid resolution. Proc. Natl. Acad. Sci. USA 111, 1825–1830 (2014).

  90. 90.

    Lally, C. C. M., Bauer, B., Selent, J. & Sommer, M. E. C-edge loops of arrestin function as a membrane anchor. Nat Commun. 8, 14258 (2017).

  91. 91.

    Scheerer, P. & Sommer, M. E. Structural mechanism of arrestin activation. Curr. Opin. Struct. Biol. 45, 160–169 (2017).

  92. 92.

    Gimenez, L. E., Vishnivetskiy, S. A., Baameur, F. & Gurevich, V. V. Manipulation of very few receptor discriminator residues greatly enhances receptor specificity of non-visual arrestins. J. Biol. Chem. 287, 29495–29505 (2012).

  93. 93.

    Peterson, S. M. et al. Elucidation of G-protein and β-arrestin functional selectivity at the dopamine D2 receptor. Proc. Natl. Acad. Sci. USA 112, 7097–7102 (2015).

  94. 94.

    Prokop, S. et al. Differential manipulation of arrestin-3 binding to basal and agonist-activated G protein-coupled receptors. Cell. Signal. 36, 98–107 (2017).

  95. 95.

    Hanson, S. M. et al. Differential interaction of spin-labeled arrestin with inactive and active phosphorhodopsin. Proc. Natl. Acad. Sci. USA 103, 4900–4905 (2006).

  96. 96.

    Charest, P. G., Terrillon, S. & Bouvier, M. Monitoring agonist-promoted conformational changes of beta-arrestin in living cells by intramolecular BRET. EMBO Rep. 6, 334–340 (2005).

  97. 97.

    Shukla, A. K. et al. Distinct conformational changes in beta-arrestin report biased agonism at seven-transmembrane receptors. Proc. Natl. Acad. Sci. USA 105, 9988–9993 (2008).

  98. 98.

    Nuber, S. et al. β-Arrestin biosensors reveal a rapid, receptor-dependent activation/deactivation cycle. Nature 531, 661–664 (2016).

  99. 99.

    Lee, M.-H. et al. The conformational signature of β-arrestin2 predicts its trafficking and signalling functions. Nature 531, 665–668 (2016).

  100. 100.

    Kim, M. et al. Conformation of receptor-bound visual arrestin. Proc. Natl. Acad. Sci. USA 109, 18407–18412 (2012).

  101. 101.

    Zhuo, Y., Vishnivetskiy, S. A., Zhan, X., Gurevich, V. V. & Klug, C. S. Identification of receptor binding-induced conformational changes in non-visual arrestins. J. Biol. Chem. 289, 20991–21002 (2014).

  102. 102.

    Kirchberg, K. et al. Conformational dynamics of helix 8 in the GPCR rhodopsin controls arrestin activation in the desensitization process. Proc. Natl. Acad. Sci. USA 108, 18690–18695 (2011).

  103. 103.

    Sommer, M. E., Hofmann, K. P. & Heck, M. Distinct loops in arrestin differentially regulate ligand binding within the GPCR opsin. Nat. Commun. 3, 995 (2012).

  104. 104.

    Zhuang, T. et al. Involvement of distinct arrestin-1 elements in binding to different functional forms of rhodopsin. Proc. Natl. Acad. Sci. USA 110, 942–947 (2013).

  105. 105.

    Tobin, A. B., Butcher, A. J. & Kong, K. C. Location, location, location...site-specific GPCR phosphorylation offers a mechanism for cell-type-specific signalling. Trends Pharmacol. Sci. 29, 413–420 (2008).

  106. 106.

    Butcher, A. J. et al. Differential G-protein-coupled receptor phosphorylation provides evidence for a signaling bar code. J. Biol. Chem. 286, 11506–11518 (2011).

  107. 107.

    Nobles, K. N. et al. Distinct phosphorylation sites on the β(2)-adrenergic receptor establish a barcode that encodes differential functions of β-arrestin. Sci. Signal. 4, ra51 (2011). The authors identify distinct phosphorylation patterns caused by distinct GRK isoforms and couple them to distinct arrestin BRET responses and signaling outcomes.

  108. 108.

    Inagaki, S. et al. G protein-coupled receptor kinase 2 (GRK2) and 5 (GRK5) exhibit selective phosphorylation of the neurotensin receptor in vitro. Biochemistry 54, 4320–4329 (2015).

  109. 109.

    Ren, X.-R. et al. Different G protein-coupled receptor kinases govern G protein and beta-arrestin-mediated signaling of V2 vasopressin receptor. Proc. Natl. Acad. Sci. USA 102, 1448–1453 (2005).

  110. 110.

    Bouzo-Lorenzo, M. et al. Distinct phosphorylation sites on the ghrelin receptor, GHSR1a, establish a code that determines the functions of ß-arrestins. Sci. Rep. 6, 22495 (2016).

  111. 111.

    Prihandoko, R. et al. Distinct phosphorylation clusters determine the signaling outcome of free fatty acid receptor 4/G protein-coupled Receptor 120. Mol. Pharmacol. 89, 505–520 (2016).

  112. 112.

    Mendez, A. et al. Rapid and reproducible deactivation of rhodopsin requires multiple phosphorylation sites. Neuron 28, 153–164 (2000).

  113. 113.

    Vishnivetskiy, S. A. et al. Regulation of arrestin binding by rhodopsin phosphorylation level. J. Biol. Chem. 282, 32075–32083 (2007).

  114. 114.

    Yang, F. et al. Phospho-selective mechanisms of arrestin conformations and functions revealed by unnatural amino acid incorporation and (19)F-NMR. Nat. Commun. 6, 8202 (2015). An elegant study that reveals how distinct receptor C-tail phosphorylation patterns imprint specific arrestin conformations and dynamics.

  115. 115.

    Gurevich, V. V. & Benovic, J. L. Visual arrestin interaction with rhodopsin. Sequential multisite binding ensures strict selectivity toward light-activated phosphorylated rhodopsin. J. Biol. Chem 268, 11628–11638 (1993).

  116. 116.

    Kumari, P. et al. Core engagement with β-arrestin is dispensable for agonist-induced vasopressin receptor endocytosis and ERK activation. Mol. Biol. Cell 28, 1003–1010 (2017).

  117. 117.

    Cahill, T. J. III et al. Distinct conformations of GPCR-β-arrestin complexes mediate desensitization, signaling, and endocytosis. Proc. Natl. Acad. Sci. USA 114, 2562–2567 (2017).

  118. 118.

    Kumari, P. et al. Functional competence of a partially engaged GPCR-β-arrestin complex. Nat. Commun. 7, 13416 (2016).

  119. 119.

    Jung, S.-R., Kushmerick, C., Seo, J. B., Koh, D.-S. & Hille, B. Muscarinic receptor regulates extracellular signal regulated kinase by two modes of arrestin binding. Proc. Natl. Acad. Sci. USA 114, E5579–E5588 (2017).

  120. 120.

    Peterhans, C., Lally, C. C. M., Ostermaier, M. K., Sommer, M. E. & Standfuss, J. Functional map of arrestin binding to phosphorylated opsin, with and without agonist. Sci. Rep. 6, 28686 (2016).

  121. 121.

    Richardson, M. D. et al. Human substance P receptor lacking the C-terminal domain remains competent to desensitize and internalize. J. Neurochem. 84, 854–863 (2003).

  122. 122.

    Jala, V. R., Shao, W.-H. & Haribabu, B. Phosphorylation-independent beta-arrestin translocation and internalization of leukotriene B4 receptors. J. Biol. Chem. 280, 4880–4887 (2005).

  123. 123.

    Granzin, J. et al. Crystal structure of p44, a constitutively active splice variant of visual arrestin. J. Mol. Biol. 416, 611–618 (2012).

  124. 124.

    Eichel, K., Jullié, D. & von Zastrow, M. β-Arrestin drives MAP kinase signalling from clathrin-coated structures after GPCR dissociation. Nat. Cell Biol. 18, 303–310 (2016).

Download references

Acknowledgements

The authors thank N. R. Latorraca for critical reading of the manuscript and insightful suggestions. This work was supported by National Institutes of Health grants R01NS028471 and R01GM083118 (B.K.K.), the German Academic Exchange Service (DAAD) (D.H.) and the American Heart Association Postdoctoral fellowship (17POST33410958) (M.M.). B.K.K. is supported by the Chan Zuckerberg Biohub.

Author information

Correspondence to Brian K. Kobilka.

Ethics declarations

Competing interests

B.K.K. is a cofounder and consultant for ConfometRx, Inc.

Additional information

Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Rights and permissions

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Further reading