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调节膜蛋白的结构和功能ion by their lipid nano-environment

Nature Reviews Molecular Cell Biologyvolume24,pages107–122 (2023)Cite this article

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Abstract

Transmembrane proteins comprise ~30% of the mammalian proteome, mediating metabolism, signalling, transport and many other functions required for cellular life. The microenvironment of integral membrane proteins (IMPs) is intrinsically different from that of cytoplasmic proteins, with IMPs solvated by a compositionally and biophysically complex lipid matrix. These solvating lipids affect protein structure and function in a variety of ways, from stereospecific, high-affinity protein–lipid interactions to modulation by bulk membrane properties. Specific examples of functional modulation of IMPs by their solvating membranes have been reported for various transporters, channels and signal receptors; however, generalizable mechanistic principles governing IMP regulation by lipid environments are neither widely appreciated nor completely understood. Here, we review recent insights into the inter-relationships between complex lipidomes of mammalian membranes, the membrane physicochemical properties resulting from such lipid collectives, and the regulation of IMPs by either or both. The recent proliferation of high-resolution methods to study such lipid–protein interactions has led to generalizable insights, which we synthesize into a general framework termed the ‘functional paralipidome’ to understand the mutual regulation between membrane proteins and their surrounding lipid microenvironments.

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Fig. 1: Inter-relationships between membrane lipidomes, protein structure–function and collective membrane physical properties.
Fig. 2: How collective membrane properties can affect protein organization and function.
Fig. 3: Physical and compositional variations in subcellular membranes.
Fig. 4: Possible modes of lipid interactions with transmembrane proteins.
Fig. 5: The functional paralipidome.

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References

  1. Krogh, A., Larsson, B., von Heijne, G. & Sonnhammer, E. L. Predicting transmembrane protein topology with a hidden Markov model: application to complete genomes.J. Mol. Biol.305, 567–580 (2001).

    ArticleCASGoogle Scholar

  2. Overington, J. P., Al-Lazikani, B. & Hopkins, A. L. How many drug targets are there?Nat。牧师. Drug Discov.5, 993–996 (2006).

    ArticleCASGoogle Scholar

  3. Payandeh, J. & Volgraf, M. Ligand binding at the protein-lipid interface: strategic considerations for drug design.Nat。牧师. Drug Discov.20, 710–722 (2021).

    ArticleCASGoogle Scholar

  4. Harayama, T. & Riezman, H. Understanding the diversity of membrane lipid composition.Nat。牧师. Mol. Cell Biol.19, 281–296 (2018).

    ArticleCASGoogle Scholar

  5. Capolupo, L. et al. Sphingolipids control dermal fibroblast heterogeneity.Science376, eabh1623 (2022).

    ArticleCASGoogle Scholar

  6. Symons, J. L. et al. Lipidomic atlas of mammalian cell membranes reveals hierarchical variation induced by culture conditions, subcellular membranes, and cell lineages.Soft Matter17, 288–297 (2020).

    ArticleGoogle Scholar

  7. Levental, K. R. et al. omega-3 polyunsaturated fatty acids direct differentiation of the membrane phenotype in mesenchymal stem cells to potentiate osteogenesis.Sci. Adv.3, eaao1193 (2017).Demonstrates that membrane lipidomes can be comprehensively remodelled by exogenous, dietary fatty acids and that these effects can modulate stem cell differentiation.

    ArticleGoogle Scholar

  8. Han, X. Lipidomics for studying metabolism.Nat。牧师. Endocrinol.12, 668–679 (2016).

    ArticleCASGoogle Scholar

  9. Eiriksson, F. F. et al. Lipidomic study of cell lines reveals differences between breast cancer subtypes.PLoS One15, e0231289 (2020).

    ArticleCASGoogle Scholar

  10. Levental, K. R. et al. Lipidomic and biophysical homeostasis of mammalian membranes counteracts dietary lipid perturbations to maintain cellular fitness.Nat. Commun.11, 1339 (2020).

    ArticleCASGoogle Scholar

  11. Contreras, F. X. et al. Molecular recognition of a single sphingolipid species by a protein’s transmembrane domain.Nature481, 525–529 (2012).Demonstration of the remarkable specificity of binding between a single-pass transmembrane domain and a particular sub-species of sphingomyelin.

    ArticleCASGoogle Scholar

  12. Lemmon, M. A. Membrane recognition by phospholipid-binding domains.Nat。牧师. Mol. Cell. Biol.9, 99–111 (2008).

    ArticleCASGoogle Scholar

  13. Moravcevic, K., Oxley, C. L. & Lemmon, M. A. Conditional peripheral membrane proteins: facing up to limited specificity.Structure20, 15–27 (2012).

    ArticleCASGoogle Scholar

  14. Lee, A. G. How lipids affect the activities of integral membrane proteins.Biochim. Biophys. Acta1666, 62–87 (2004).

    ArticleCASGoogle Scholar

  15. Barrera, N. P., Zhou, M. & Robinson, C. V. The role of lipids in defining membrane protein interactions: insights from mass spectrometry.Trends Cell Biol.23, 1–8 (2013).

    ArticleCASGoogle Scholar

  16. Contreras, F. X., Ernst, A. M., Wieland, F. & Brugger, B. Specificity of intramembrane protein-lipid interactions.Cold Spring Harb. Perspect. Biol.3, a004705 (2011).

    ArticleGoogle Scholar

  17. Corradi, V. et al. Emerging diversity in lipid-protein interactions.Chem. Rev.119, 5775–5848 (2019).

    ArticleCASGoogle Scholar

  18. Sych, T., Levental, K. R. & Sezgin, E. Lipid-protein interactions in plasma membrane organization and function.Annu. Rev. Biophys.51, 135–156 (2022).

    ArticleGoogle Scholar

  19. Ernst, M. & Robertson, J. L. The role of the membrane in transporter folding and activity.J. Mol. Biol.433, 167103 (2021).

    ArticleCASGoogle Scholar

  20. Simunovic, M., Prevost, C., Callan-Jones, A. & Bassereau, P. Physical basis of some membrane shaping mechanisms.Philos. Trans. A Math. Phys. Eng. Sci.374, 20160034 (2016).

    Google Scholar

  21. Phillips, R., Ursell, T., Wiggins, P. & Sens, P. Emerging roles for lipids in shaping membrane-protein function.Nature459, 379–385 (2009).

    ArticleCASGoogle Scholar

  22. Heberle, F. A. et al. Direct label-free imaging of nanodomains in biomimetic and biological membranes by cryogenic electron microscopy.Proc. Natl Acad. Sci. USA117, 19943–19952 (2020).

    ArticleCASGoogle Scholar

  23. Mitra, K., Ubarretxena-Belandia, I., Taguchi, T., Warren, G. & Engelman, D. M. Modulation of the bilayer thickness of exocytic pathway membranes by membrane proteins rather than cholesterol.Proc. Natl Acad. Sci. USA101, 4083–4088 (2004).

    ArticleCASGoogle Scholar

  24. Cornell, C. E., Mileant, A., Thakkar, N., Lee, K. K. & Keller, S. L. Direct imaging of liquid domains in membranes by cryo-electron tomography.Proc. Natl Acad. Sci. USA117, 19713–19719 (2020).

    ArticleCASGoogle Scholar

  25. Fischer, T. D., Dash, P. K., Liu, J. & Waxham, M. N. Morphology of mitochondria in spatially restricted axons revealed by cryo-electron tomography.PLoS Biol.16, e2006169 (2018).

    ArticleGoogle Scholar

  26. Andersen, O. S. & Koeppe, R. E. 2nd Bilayer thickness and membrane protein function: an energetic perspective.Annu. Rev. Biophys. Biomol. Struct.36, 107–130 (2007).

    ArticleCASGoogle Scholar

  27. Falzone, M. E. et al. TMEM16 scramblases thin the membrane to enable lipid scrambling.Nat. Commun.13, 2604 (2022).

    ArticleCASGoogle Scholar

  28. Kaiser, H.-J. et al. Lateral sorting in model membranes by cholesterol-mediated hydrophobic matching.Proc. Natl Acad. Sci. USA108, 16628–16633 (2011).

    ArticleCASGoogle Scholar

  29. Chadda, R. et al. Membrane transporter dimerization driven by differential lipid solvation energetics of dissociated and associated states.eLife10, e63288 (2021).Unique biochemical approaches are combined with simulations to show that IMP oligomerization is mediated by the energetics of TMD solvation by lipids rather than by direct lipid binding.

    ArticleCASGoogle Scholar

  30. Beaven, A. H. et al. Gramicidin A channel formation induces local lipid redistribution I: experiment and simulation.Biophys. J.112, 1185–1197 (2017).

    ArticleCASGoogle Scholar

  31. Sharpe, H. J., Stevens, T. J. & Munro, S. A comprehensive comparison of transmembrane domains reveals organelle-specific properties.Cell142, 158–169 (2010).

    ArticleCASGoogle Scholar

  32. Diaz-Rohrer, B., Levental, K. R. & Levental, I. Rafting through traffic: membrane domains in cellular logistics.Biochim. Biophys. Acta Biomembr.1838, 3003–3013 (2014).

    ArticleCASGoogle Scholar

  33. Prasad, R., Sliwa-Gonzalez, A. & Barral, Y. Mapping bilayer thickness in the ER membrane.Sci. Adv.6, aba5130 (2020).

    ArticleGoogle Scholar

  34. Zhemkov, V. et al. The role of sigma 1 receptor in organization of endoplasmic reticulum signaling microdomains.eLife10, e65192 (2021).

    ArticleCASGoogle Scholar

  35. King, C., Sengupta, P., Seo, A. Y. & Lippincott-Schwartz, J. ER membranes exhibit phase behavior at sites of organelle contact.Proc. Natl Acad. Sci. USA117, 7225–7235 (2020).

    ArticleCASGoogle Scholar

  36. Cornelius, F. Modulation of Na,K-ATPase and Na-ATPase activity by phospholipids and cholesterol. I. Steady-state kinetics.Biochemistry40, 8842–8851 (2001).

    ArticleCASGoogle Scholar

  37. Lee, A. G. Lipid-protein interactions in biological membranes: a structural perspective.Biochim. Biophys. Acta1612, 1–40 (2003).

    ArticleCASGoogle Scholar

  38. Botelho, A. V., Huber, T., Sakmar, T. P. & Brown, M. F. Curvature and hydrophobic forces drive oligomerization and modulate activity of rhodopsin in membranes.Biophys. J.91, 4464–4477 (2006).

    ArticleCASGoogle Scholar

  39. Jensen, M. O. & Mouritsen, O. G. Lipids do influence protein function — the hydrophobic matching hypothesis revisited.Biochim. Biophys. Acta Biomembr.1666, 205–226 (2004).A detailed discussion of experimental observations documenting how proteins perturb their lipid environments and, vice versa, how membrane properties like hydrophobic mismatch and curvature stress affect protein function.

    ArticleCASGoogle Scholar

  40. Cybulski, L. E. et al. Activation of the bacterial thermosensor DesK involves a serine zipper dimerization motif that is modulated by bilayer thickness.Proc. Natl Acad. Sci. USA112, 6353–6358 (2015).

    ArticleCASGoogle Scholar

  41. Simunovic, M., Evergren, E., Callan-Jones, A. & Bassereau, P. Curving cells inside and out: roles of BAR domain proteins in membrane shaping and its cellular implications.Annu. Rev. Cell Dev. Biol.35, 111–129 (2019).

    ArticleCASGoogle Scholar

  42. Cui, H., Lyman, E. & Voth, G. A. Mechanism of membrane curvature sensing by amphipathic helix containing proteins.Biophys. J.100, 1271–1279 (2011).

    ArticleCASGoogle Scholar

  43. Aimon, S. et al. Membrane shape modulates transmembrane protein distribution.Dev. Cell28, 212–218 (2014).

    ArticleCASGoogle Scholar

  44. Shurer, C. R. et al. Physical principles of membrane shape regulation by the glycocalyx.Cell177, 1757–1770.e21 (2019).Protein engineering and scanning electron microscopy reveal remarkable morphological transformations of living cell membranes induced by glycosylated IMPs.

    ArticleCASGoogle Scholar

  45. Stachowiak, J. C., Hayden, C. C. & Sasaki, D. Y. Steric confinement of proteins on lipid membranes can drive curvature and tubulation.Proc. Natl Acad. Sci. USA107, 7781–7786 (2010).

    ArticleCASGoogle Scholar

  46. Ernst, A. M. et al. S-palmitoylation sorts membrane cargo for anterograde transport in the Golgi.Dev. Cell47, 479–493.e7 (2018).

    ArticleCASGoogle Scholar

  47. Budin, I. et al. Viscous control of cellular respiration by membrane lipid composition.Science362, 1186–1189 (2018).巧妙的结合遗传学、生物物理学和垫子上hematical models to explain how membrane viscosity regulates cell growth via diffusion of electron carriers in the respiratory chain.

    ArticleCASGoogle Scholar

  48. Sinensky, M. Homeoviscous adaptation-a homeostatic process that regulates the viscosity of membrane lipids in Escherichia coli.Proc. Natl Acad. Sci. USA71, 522–525 (1974).

    ArticleCASGoogle Scholar

  49. Ernst, R., Ejsing, C. S. & Antonny, B. Homeoviscous adaptation and the regulation of membrane lipids.J. Mol. Biol.428, 4776–4791 (2016).

    ArticleCASGoogle Scholar

  50. Martiniere, A. et al. Homeostasis of plasma membrane viscosity in fluctuating temperatures.N. Phytol.192, 328–337 (2011).

    ArticleCASGoogle Scholar

  51. Behan-Martin, M. K., Jones, G. R., Bowler, K. & Cossins, A. R. A near perfect temperature adaptation of bilayer order in vertebrate brain membranes.Biochim. Biophys. Acta1151, 216–222 (1993).

    ArticleCASGoogle Scholar

  52. Ruiz, M. et al. Membrane fluidity is regulated by the C. elegans transmembrane protein FLD-1 and its human homologs TLCD1/2.eLife7, 40686 (2018).

    ArticleGoogle Scholar

  53. Covino, R. et al. A eukaryotic sensor for membrane lipid saturation.Mol. Cell63, 49–59 (2016).

    ArticleCASGoogle Scholar

  54. Ballweg, S. et al. Regulation of lipid saturation without sensing membrane fluidity.Nat. Commun.11, 756 (2020).

    ArticleCASGoogle Scholar

  55. Radanovic, T., Reinhard, J., Ballweg, S., Pesek, K. & Ernst, R. An emerging group of membrane property sensors controls the physical state of organellar membranes to maintain their identity.BioEssays40, e1700250 (2018).

    ArticleGoogle Scholar

  56. Halbleib, K. et al. Activation of the unfolded protein response by lipid bilayer stress.Mol. Cell67, 673–684.e8 (2017).Demonstration that proteins can sense membrane properties by compressing the bilayer to signal aberrant lipid compositions during cell stress.

    ArticleCASGoogle Scholar

  57. Cox, C. D., Bavi, N. & Martinac, B. Bacterial mechanosensors.Annu. Rev. Physiol.80, 71–93 (2018).

    ArticleCASGoogle Scholar

  58. Reddy, B., Bavi, N., Lu, A., Park, Y. & Perozo, E. Molecular basis of force-from-lipids gating in the mechanosensitive channel MscS.eLife8, e50486 (2019).

    ArticleCASGoogle Scholar

  59. Coste, B. et al. Piezo1 and Piezo2 are essential components of distinct mechanically activated cation channels.Science330, 55–60 (2010).

    ArticleCASGoogle Scholar

  60. Lin, Y. C. et al. Force-induced conformational changes in PIEZO1.Nature573, 230–234 (2019).

    ArticleCASGoogle Scholar

  61. Brohawn, S. G., Su, Z. & MacKinnon, R. Mechanosensitivity is mediated directly by the lipid membrane in TRAAK and TREK1 K+channels.Proc. Natl Acad. Sci. USA111, 3614–3619 (2014).

    ArticleCASGoogle Scholar

  62. Bavi, N. et al. The conformational cycle of prestin underlies outer-hair cell electromotility.Nature600, 553–558 (2021).

    ArticleCASGoogle Scholar

  63. Ge, J. et al. Molecular mechanism of prestin electromotive signal amplification.Cell184, 4669–4679 (2021).

    ArticleCASGoogle Scholar

  64. Dallos, P. et al. Prestin-based outer hair cell motility is necessary for mammalian cochlear amplification.Neuron58, 333–339 (2008).

    ArticleCASGoogle Scholar

  65. Sezgin, E., Levental, I., Mayor, S. & Eggeling, C. The mystery of membrane organization: composition, regulation and roles of lipid rafts.Nat。牧师. Mol. Cell Biol.18, 361–374 (2017).

    ArticleCASGoogle Scholar

  66. Lingwood, D. & Simons, K. Lipid rafts as a membrane-organizing principle.Science327, 46–50 (2010).

    ArticleCASGoogle Scholar

  67. Veatch, S. L. & Keller, S. L. Organization in lipid membranes containing cholesterol.Phys. Rev. Lett.89, 268101 (2002).

    ArticleGoogle Scholar

  68. Veatch, S. L. & Keller, S. L. Seeing spots: complex phase behavior in simple membranes.Biochim. Biophys. Acta1746, 172–185 (2005).

    ArticleCASGoogle Scholar

  69. Sodt, A. J., Sandar, M. L., Gawrisch, K., Pastor, R. W. & Lyman, E. The molecular structure of the liquid-ordered phase of lipid bilayers.J. Am. Chem. Soc.136, 725–732 (2014).

    ArticleCASGoogle Scholar

  70. Sodt, A. J., Pastor, R. W. & Lyman, E. Hexagonal substructure and hydrogen bonding in liquid-ordered phases containing palmitoyl sphingomyelin.Biophys. J.109, 948–955 (2015).

    ArticleCASGoogle Scholar

  71. Baumgart, T. et al. Large-scale fluid/fluid phase separation of proteins and lipids in giant plasma membrane vesicles.Proc. Natl Acad. Sci. USA104, 3165–3170 (2007).

    ArticleCASGoogle Scholar

  72. Lingwood, D., Ries, J., Schwille, P. & Simons, K. Plasma membranes are poised for activation of raft phase coalescence at physiological temperature.Proc. Natl Acad. Sci. USA105, 10005–10010 (2008).

    ArticleCASGoogle Scholar

  73. Levental, K. R. & Levental, I. Giant plasma membrane vesicles: models for understanding membrane organization.Curr. Top. Membr.75, 25–57 (2015).

    ArticleCASGoogle Scholar

  74. Veatch, S. L. et al. Critical fluctuations in plasma membrane vesicles.ACS Chem. Biol.3, 287–293 (2008).

    ArticleCASGoogle Scholar

  75. Levental, I. et al. Cholesterol-dependent phase separation in cell-derived giant plasma-membrane vesicles.Biochem. J.424, 163–167 (2009).

    ArticleCASGoogle Scholar

  76. Levental, K. R. et al. Polyunsaturated lipids regulate membrane domain stability by tuning membrane order.Biophys. J.110, 1800–1810 (2016).

    ArticleCASGoogle Scholar

  77. Toulmay, A. & Prinz, W. A. Direct imaging reveals stable, micrometer-scale lipid domains that segregate proteins in live cells.J. Cell Biol.202, 35–44 (2013).

    ArticleCASGoogle Scholar

  78. Rayermann, S. P., Rayermann, G. E., Cornell, C. E., Merz, A. J. & Keller, S. L. Hallmarks of reversible separation of living, unperturbed cell membranes into two liquid phases.Biophys. J.113, 2425–2432 (2017).

    ArticleCASGoogle Scholar

  79. Raghupathy, R. et al. Transbilayer lipid interactions mediate nanoclustering of lipid-anchored proteins.Cell161, 581 - 594(2015)。

    ArticleCASGoogle Scholar

  80. Owen, D. M., Williamson, D. J., Magenau, A. & Gaus, K. Sub-resolution lipid domains exist in the plasma membrane and regulate protein diffusion and distribution.Nat. Commun.3, 1256 (2012).

    ArticleGoogle Scholar

  81. Sanchez, S. A., Tricerri, M. A. & Gratton, E. Laurdan generalized polarization fluctuations measures membrane packing micro-heterogeneity in vivo.Proc. Natl Acad. Sci. USA109, 7314–7319 (2012).

    ArticleCASGoogle Scholar

  82. Kinoshita, M. et al. Raft-based sphingomyelin interactions revealed by new fluorescent sphingomyelin analogs.J. Cell Biol.216, 1183–1204 (2017).

    ArticleCASGoogle Scholar

  83. Komura, N. et al. Raft-based interactions of gangliosides with a GPI-anchored receptor.Nat. Chem. Biol.12, 402–410 (2016).

    ArticleCASGoogle Scholar

  84. Stone, M. B., Shelby, S. A., Nunez, M. F., Wisser, K. & Veatch, S. L. Protein sorting by lipid phase-like domains supports emergent signaling function in B lymphocyte plasma membranes.eLife6, e19891 (2017).

    ArticleGoogle Scholar

  85. Eggeling, C. et al. Direct observation of the nanoscale dynamics of membrane lipids in a living cell.Nature457, 1159–1162 (2009).

    ArticleCASGoogle Scholar

  86. Levental, I., Levental, K. R. & Heberle, F. A. Lipid rafts: controversies resolved, mysteries remain.Trends Cell Biol.30, 341–353 (2020).

    ArticleCASGoogle Scholar

  87. Sezgin, E. et al. Elucidating membrane structure and protein behavior using giant plasma membrane vesicles.Nat. Protoc.7, 1042–1051 (2012).

    ArticleCASGoogle Scholar

  88. Levental Diaz-Rohrer, B, B。,西蒙斯,k和k·RLevental, I. Membrane raft association is a determinant of plasma membrane localization.Proc. Natl Acad. Sci. USA111, 8500–8505 (2014).

    ArticleCASGoogle Scholar

  89. Lorent, j . h . et al。结构性因素和functional consequences of protein affinity for membrane rafts.Nat. Commun.8, 1219 (2017).

    ArticleGoogle Scholar

  90. Levental, I., Lingwood, D., Grzybek, M., Coskun, U. & Simons, K. Palmitoylation regulates raft affinity for the majority of integral raft proteins.Proc. Natl Acad. Sci. USA107, 22050–22054 (2010).

    ArticleCASGoogle Scholar

  91. Levental, I., Grzybek, M. & Simons, K. Greasing their way: lipid modifications determine protein association with membrane rafts.Biochemistry49, 6305–6316 (2010).

    ArticleCASGoogle Scholar

  92. Castello-Serrano, I., Lorent, J. H., Ippolito, R., Levental, K. R. & Levental, I. Myelin-associated MAL and PLP Are unusual among multipass transmembrane proteins in preferring ordered membrane domains.J. Phys. Chem. B124, 5930–5939 (2020).

    ArticleCASGoogle Scholar

  93. Sengupta, P. et al. A lipid-based partitioning mechanism for selective incorporation of proteins into membranes of HIV particles.Nat. Cell Biol.21, 452–461 (2019).

    ArticleCASGoogle Scholar

  94. Ono, A. & Freed, E. O. Plasma membrane rafts play a critical role in HIV-1 assembly and release.Proc. Natl Acad. Sci. USA98, 13925–13930 (2001).

    ArticleCASGoogle Scholar

  95. Mitchell, D. C., Straume, M., Miller, J. L. & Litman, B. J. Modulation of metarhodopsin formation by cholesterol-induced ordering of bilayer lipids.Biochemistry29, 9143–9149 (1990).

    ArticleCASGoogle Scholar

  96. Huang, S. K. et al. Allosteric modulation of the adenosine A2A receptor by cholesterol.Elife11, e73901 (2021).

    ArticleGoogle Scholar

  97. Tikku, S. et al. Relationship between Kir2.1/Kir2.3 activity and their distributions between cholesterol-rich and cholesterol-poor membrane domains.Am. J. Physiol. Cell Physiol.293, C440–C450 (2007).

    ArticleCASGoogle Scholar

  98. Murata, M. et al. VIP21/caveolin is a cholesterol-binding protein.Proc. Natl Acad. Sci. USA92, 10339–10343 (1995).

    ArticleCASGoogle Scholar

  99. Parton, R. G. & Simons, K. The multiple faces of caveolae.Nat。牧师. Mol. Cell Biol.8, 185–194 (2007).

    ArticleCASGoogle Scholar

  100. Parton, R. G., Kozlov, M. M. & Ariotti, N. Caveolae and lipid sorting: shaping the cellular response to stress.J. Cell Biol.219, e201905071 (2020).

    ArticleGoogle Scholar

  101. Ortegren, U. et al. Lipids and glycosphingolipids in caveolae and surrounding plasma membrane of primary rat adipocytes.Eur. J. Biochem.271, 2028–2036 (2004).

    ArticleGoogle Scholar

  102. Rothberg, K. G. et al. Caveolin, a protein component of caveolae membrane coats.Cell68, 673–682 (1992).

    ArticleCASGoogle Scholar

  103. Epand, R. M., Sayer, B. G. & Epand, R. F. Caveolin scaffolding region and cholesterol-rich domains in membranes.J. Mol. Biol.345, 339–350 (2005).

    ArticleCASGoogle Scholar

  104. Sinha, B. et al. Cells respond to mechanical stress by rapid disassembly of caveolae.Cell144, 402–413 (2011).

    ArticleCASGoogle Scholar

  105. Marsh, D. Protein modulation of lipids, and vice-versa, in membranes.Biochim. Biophys. Acta1778, 1545–1575 (2008).

    ArticleCASGoogle Scholar

  106. Gonen, T. et al. Lipid-protein interactions in double-layered two-dimensional AQP0 crystals.Nature438, 633–638 (2005).Ground-breaking study using electron diffraction from two-dimension crystals of a membrane protein, AQP0, providing direct evidence for lipid–protein interaction and the water conduction mechanism.

    ArticleCASGoogle Scholar

  107. Jost, P. C., Griffith, O. H., Capaldi, R. A. & Vanderkooi, G. Evidence for boundary lipid in membranes.Proc. Natl Acad. Sci. USA70, 480–484 (1973).

    ArticleCASGoogle Scholar

  108. Brotherus, j . r . et al。Lipid-protein多个本ding equilibria in membranes.Biochemistry20, 5261–5267 (1981).

    ArticleCASGoogle Scholar

  109. 阅读l . I。Brophy, p . j . &沼泽,d .交换rates at the lipid-protein interface of myelin proteolipid protein studied by spin-label electron spin resonance.Biochemistry27, 46–52 (1988).

    ArticleCASGoogle Scholar

  110. Marius, P., Alvis, S. J., East, J. M. & Lee, A. G. The interfacial lipid binding site on the potassium channel KcsA is specific for anionic phospholipids.Biophys. J.89, 4081–4089 (2005).

    ArticleCASGoogle Scholar

  111. McAuley, K. E. et al. Structural details of an interaction between cardiolipin and an integral membrane protein.Proc. Natl Acad. Sci. USA96, 14706–14711 (1999).

    ArticleCASGoogle Scholar

  112. Belrhali, H. et al. Protein, lipid and water organization in bacteriorhodopsin crystals: a molecular view of the purple membrane at 1.9 A resolution.Structure7, 909–917 (1999).

    ArticleCASGoogle Scholar

  113. Zhou, Y., Morais-Cabral, J. H., Kaufman, A. & MacKinnon, R. Chemistry of ion coordination and hydration revealed by a K+channel-Fab complex at 2.0 A resolution.Nature414, 43–48 (2001).

    ArticleCASGoogle Scholar

  114. Duncan, A. L., Song, W. & Sansom, M. S. P. Lipid-dependent regulation of ion channels and G protein-coupled receptors: insights from structures and simulations.Annu. Rev. Pharmacol. Toxicol.60, 31–50 (2020).

    ArticleCASGoogle Scholar

  115. Hansen, S. B., Tao, X. & MacKinnon, R. Structural basis of PIP2 activation of the classical inward rectifier K+channel Kir2.2.Nature477, 495–498 (2011).

    ArticleCASGoogle Scholar

  116. McLaughlin, S., Wang, J., Gambhir, A. & Murray, D. PIP(2) and proteins: interactions, organization, and information flow.Annu. Rev. Biophys. Biomol. Struct.31, 151–175 (2002).

    ArticleCASGoogle Scholar

  117. Rosenhouse-Dantsker, A., Noskov, S., Durdagi, S., Logothetis, D. E. & Levitan, I. Identification of novel cholesterol-binding regions in Kir2 channels.J. Biol. Chem.288, 31154–31164 (2013).

    ArticleCASGoogle Scholar

  118. Romanenko, V. G. et al. Cholesterol sensitivity and lipid raft targeting of Kir2.1 channels.Biophys. J.87, 3850–3861 (2004).

    ArticleCASGoogle Scholar

  119. Duncan, A. L., Corey, R. A. & Sansom, M. S. P. Defining how multiple lipid species interact with inward rectifier potassium (Kir2) channels.Proc. Natl Acad. Sci. USA117, 7803–7813 (2020).

    ArticleCASGoogle Scholar

  120. Zaydman, M. A. et al. Kv7.1 ion channels require a lipid to couple voltage sensing to pore opening.Proc. Natl Acad. Sci. USA110, 13180–13185 (2013).

    ArticleCASGoogle Scholar

  121. Hilgemann, D. W. & Ball, R. Regulation of cardiac Na+,Ca2+exchange and KATP potassium channels by PIP2.Science273, 956–959 (1996).

    ArticleCASGoogle Scholar

  122. Hedger, G. et al. Lipid-loving ANTs: molecular simulations of cardiolipin interactions and the organization of the adenine nucleotide translocase in model mitochondrial membranes.Biochemistry55, 6238–6249 (2016).

    ArticleCASGoogle Scholar

  123. Hite, R. K., Butterwick, J. A. & MacKinnon, R. Phosphatidic acid modulation of Kv channel voltage sensor function.eLife3, e04366 (2014).

    ArticleGoogle Scholar

  124. Zocher, M., Zhang, C., Rasmussen, S. G., Kobilka, B. K. & Muller, D. J. Cholesterol increases kinetic, energetic, and mechanical stability of the human beta2-adrenergic receptor.Proc. Natl Acad. Sci. USA109, E3463–E3472 (2012).

    ArticleCASGoogle Scholar

  125. Cherezov, V. et al. High-resolution crystal structure of an engineered human beta2-adrenergic G protein-coupled receptor.Science318, 1258–1265 (2007).

    ArticleCASGoogle Scholar

  126. Liu, W. et al. Structural basis for allosteric regulation of GPCRs by sodium ions.Science337, 232–236 (2012).

    ArticleCASGoogle Scholar

  127. Guixa-Gonzalez, R. et al. Membrane cholesterol access into a G-protein-coupled receptor.Nat. Commun.8, 14505 (2017).

    ArticleCASGoogle Scholar

  128. Manna, M. et al. Mechanism of allosteric regulation of beta2-adrenergic receptor by cholesterol.eLife5, e18432 (2016).

    ArticleGoogle Scholar

  129. Rouviere, E., Arnarez, C., Yang, L. & Lyman, E. Identification of two new cholesterol interaction sites on the A2A adenosine receptor.Biophys. J.113, 2415–2424 (2017).

    ArticleCASGoogle Scholar

  130. Lee, J. Y. & Lyman, E. Predictions for cholesterol interaction sites on the A2A adenosine receptor.J. Am. Chem. Soc.134, 16512–16515 (2012).

    ArticleCASGoogle Scholar

  131. Nelson, L. D., Johnson, A. E. & London, E. How interaction of perfringolysin O with membranes is controlled by sterol structure, lipid structure, and physiological low pH: insights into the origin of perfringolysin O-lipid raft interaction.J. Biol. Chem.283, 4632–4642 (2008).

    ArticleCASGoogle Scholar

  132. D’Avanzo, N., Hyrc, K., Enkvetchakul, D., Covey, D. F. & Nichols, C. G. Enantioselective protein-sterol interactions mediate regulation of both prokaryotic and eukaryotic inward rectifier K+channels by cholesterol.PLoS One6, e19393 (2011).

    ArticleGoogle Scholar

  133. Gutorov, R. et al. Modulation of transient receptor potential C channel activity by cholesterol.Front. Pharmacol.10, 1487 (2019).

    ArticleCASGoogle Scholar

  134. Elkins, M. R. et al. Cholesterol-binding site of the influenza M2 protein in lipid bilayers from solid-state NMR.Proc. Natl Acad. Sci. USA114, 12946–12951 (2017).

    ArticleCASGoogle Scholar

  135. Barrett, P. J. et al. The amyloid precursor protein has a flexible transmembrane domain and binds cholesterol.Science336, 1168–1171 (2012).

    ArticleCASGoogle Scholar

  136. Marlow, B., Kuenze, G., Li, B., Sanders, C. R. & Meiler, J. Structural determinants of cholesterol recognition in helical integral membrane proteins.Biophys. J.120, 1592–1604 (2021).

    ArticleCASGoogle Scholar

  137. Romer, W. et al. Shiga toxin induces tubular membrane invaginations for its uptake into cells.Nature450, 670–675 (2007).

    ArticleGoogle Scholar

  138. Flores, A. et al. Gangliosides interact with synaptotagmin to form the high-affinity receptor complex for botulinum neurotoxin B.Proc. Natl Acad. Sci. USA116, 18098–18108 (2019).

    ArticleCASGoogle Scholar

  139. Ewers, H. et al. GM1 structure determines SV40-induced membrane invagination and infection.Nat. Cell Biol.12, 11–18 (2010).

    ArticleCASGoogle Scholar

  140. Coskun, U., Grzybek, M., Drechsel, D. & Simons, K. Regulation of human EGF receptor by lipids.Proc. Natl Acad. Sci. USA108, 9044–9048 (2010).

    ArticleGoogle Scholar

  141. Song, W., Yen, H. Y., Robinson, C. V. & Sansom, M. S. P. State-dependent lipid interactions with the A2a receptor revealed by MD simulations using in vivo-mimetic membranes.Structure27, 392–403.e3 (2019).

    ArticleCASGoogle Scholar

  142. Laganowsky, A. et al. Membrane proteins bind lipids selectively to modulate their structure and function.Nature510, 172–175 (2014).Native mass spectrometry used to demonstrate that lipids can remain bound to proteins through the extraction and ionization process and be identified directly by their molecular mass.

    ArticleCASGoogle Scholar

  143. Cong, X. et al. Determining membrane protein-lipid binding thermodynamics using native mass spectrometry.J. Am. Chem. Soc.138, 4346–4349 (2016).

    ArticleCASGoogle Scholar

  144. Patrick, J. W. et al. Allostery revealed within lipid binding events to membrane proteins.Proc. Natl Acad. Sci. USA115, 2976–2981 (2018).

    ArticleCASGoogle Scholar

  145. Gupta, K. et al. The role of interfacial lipids in stabilizing membrane protein oligomers.Nature541, 421–424 (2017).

    ArticleCASGoogle Scholar

  146. Yang, L. & Lyman, E. Local enrichment of unsaturated chains around the A2A adenosine receptor.Biochemistry58, 4096–4105 (2019).

    ArticleCASGoogle Scholar

  147. Tanford, C. Extension of the theory of linked functions to incorporate the effects of protein hydration.J. Mol. Biol.39, 539–544 (1969).

    ArticleCASGoogle Scholar

  148. Salari, R., Joseph, T., Lohia, R., Henin, J. & Brannigan, G. A streamlined, general approach for computing ligand binding free energies and its application to GPCR-bound cholesterol.J. Chem. Theory Comput.14, 6560–6573 (2018).

    ArticleCASGoogle Scholar

  149. Corey, R. A., Stansfeld, P. J. & Sansom, M. S. P. The energetics of protein-lipid interactions as viewed by molecular simulations.Biochem. Soc. Trans.48, 25–37 (2020).

    ArticleCASGoogle Scholar

  150. Salas-Estrada, L. A., Leioatts, N., Romo, T. D. & Grossfield, A. Lipids alter rhodopsin function via ligand-like and solvent-like interactions.Biophys. J.114, 355–367 (2018).

    ArticleCASGoogle Scholar

  151. Leonard, A. N. & Lyman, E. Activation of G-protein-coupled receptors is thermodynamically linked to lipid solvation.Biophys. J.120, 1777–1787 (2021).Conceptual underpinning of the functional paralipidome model, demonstrating via computational and theoretical analysis, how lipid nano-environments could influence protein conformational equilibria.

    ArticleCASGoogle Scholar

  152. Huber, T., Botelho, A. V., Beyer, K. & Brown, M. F. Membrane model for the G-protein-coupled receptor rhodopsin: hydrophobic interface and dynamical structure.Biophys. J.86, 2078–2100 (2004).

    ArticleCASGoogle Scholar

  153. van ‘t Klooster, J. S. et al. Periprotein lipidomes of Saccharomyces cerevisiae provide a flexible environment for conformational changes of membrane proteins.eLife9, e57003 (2020).Demonstration of the remarkable specificity of binding between a single-pass transmembrane domain and a particular sub-species of sphingomyelin.

    ArticleGoogle Scholar

  154. Gupta, K. et al. Identifying key membrane protein lipid interactions using mass spectrometry.Nat. Protoc.13, 1106–1120 (2018).

    ArticleCASGoogle Scholar

  155. Corradi, V. et al. Lipid-protein interactions are unique fingerprints for membrane proteins.ACS Cent. Sci.4, 709–717 (2018).

    ArticleCASGoogle Scholar

  156. Denisov, I. G. & Sligar, S. G. Nanodiscs for structural and functional studies of membrane proteins.Nat. Struct. Mol. Biol.23, 481–486 (2016).

    ArticleCASGoogle Scholar

  157. Dorr, J. M. et al. Detergent-free isolation, characterization, and functional reconstitution of a tetrameric K+ channel: the power of native nanodiscs.Proc. Natl Acad. Sci. USA111, 18607–18612 (2014).

    ArticleGoogle Scholar

  158. Flores, J. A. et al. Connexin-46/50 in a dynamic lipid environment resolved by CryoEM at 1.9 A.Nat. Commun.11, 4331 (2020).Ultra-high-resolution structures of native connexin channels, which span two bilayers, reveal novel details of IMP–membrane interactions, including remarkable ordering of outer leaflet lipids.

    ArticleCASGoogle Scholar

  159. Cuevas Arenas, R. et al. Fast collisional lipid transfer among polymer-bounded nanodiscs.Sci. Rep.7, 45875 (2017).

    ArticleCASGoogle Scholar

  160. Stepien, P. et al. Complexity of seemingly simple lipid nanodiscs.Biochim. Biophys. Acta Biomembr.1862, 183420 (2020).

    ArticleCASGoogle Scholar

  161. Javanainen, M. et al. Reduced level of docosahexaenoic acid shifts GPCR neuroreceptors to less ordered membrane regions.PLoS Comput. Biol.15, e1007033 (2019).

    ArticleCASGoogle Scholar

  162. Sodt, A. J., Beaven, A. H., Andersen, O. S., Im, W. & Pastor, R. W. Gramicidin a channel formation induces local lipid redistribution II: a 3D continuum elastic model.Biophys. J.112, 1198–1213 (2017).

    ArticleCASGoogle Scholar

  163. Lundbaek, j ., Collingwood, s。Ingolfsson, H. I., Kapoor, R. & Andersen, O. S. Lipid bilayer regulation of membrane protein function: gramicidin channels as molecular force probes.j . r . Soc。接口7, 373–395 (2010).

    ArticleCASGoogle Scholar

  164. McGraw, C., Yang, L., Levental, I., Lyman, E. & Robinson, A. S. Membrane cholesterol depletion reduces downstream signaling activity of the adenosine A2A receptor.Biochim. Biophys. Acta Biomembr.1861, 760–767 (2019).

    ArticleCASGoogle Scholar

  165. Gutierrez, M. G., Mansfield, K. S. & Malmstadt, N. The functional activity of the human serotonin 5-HT1A receptor is controlled by lipid bilayer composition.Biophys. J.110, 2486–2495 (2016).

    ArticleCASGoogle Scholar

  166. Kumar, G. A. et al. A molecular sensor for cholesterol in the human serotonin1A receptor.Sci. Adv.7, eabh2922 (2021).

    ArticleCASGoogle Scholar

  167. Barbotin, A. et al. z-STED imaging and spectroscopy to investigate nanoscale membrane structure and dynamics.Biophys. J.118, 2448–2457 (2020).

    ArticleCASGoogle Scholar

  168. Nadezhdin, K. D. et al. Structural mechanism of heat-induced opening of a temperature-sensitive TRP channel.Nat. Struct. Mol. Biol.28, 564–572 (2021).

    ArticleCASGoogle Scholar

  169. Doktorova, M., Symons, J. L. & Levental, I. Structural and functional consequences of reversible lipid asymmetry in living membranes.Nat. Chem. Biol.16, 1321–1330 (2020).

    ArticleCASGoogle Scholar

  170. Lorent, j . h . et al。Plasma membranes are asymmetric in lipid unsaturation, packing and protein shape.Nat. Chem. Biol.16, 644–652 (2020).Experiments and computational modelling detail the lipidomic and biophysical asymmetry of a mammalian plasma membrane, while bioinformatics shows that membrane asymmetry is reflected in asymmetry of protein TMDs, which is conserved throughout Eukaryota.

    ArticleCASGoogle Scholar

  171. Tsuji, T. et al. Predominant localization of phosphatidylserine at the cytoplasmic leaflet of the ER, and its TMEM16K-dependent redistribution.Proc. Natl Acad. Sci. USA116, 13368–13373 (2019).

    ArticleCASGoogle Scholar

  172. Malvezzi, M. et al. Ca2+-dependent phospholipid scrambling by a reconstituted TMEM16 ion channel.Nat. Commun.4, 2367 (2013).

    ArticleGoogle Scholar

  173. van Meer, G., Voelker, D. R. & Feigenson, G. W. Membrane lipids: where they are and how they behave.Nat。牧师. Mol. Cell Biol.9, 112–124 (2008).

    ArticleGoogle Scholar

  174. Akimzhanov, A. M. & Boehning, D. Rapid and transient palmitoylation of the tyrosine kinase Lck mediates Fas signaling.Proc. Natl Acad. Sci. USA112, 11876–11880 (2015).

    ArticleCASGoogle Scholar

  175. Martin, B. R., Wang, C., Adibekian, A., Tully, S. E. & Cravatt, B. F. Global profiling of dynamic protein palmitoylation.Nat. Methods9, 84–89 (2011).

    ArticleGoogle Scholar

  176. Martin, B. R. & Cravatt, B. F. Large-scale profiling of protein palmitoylation in mammalian cells.Nat. Methods6, 135–138 (2009).

    ArticleCASGoogle Scholar

  177. Veatch, S. L., Gawrisch, K. & Keller, S. L. Closed-loop miscibility gap and quantitative tie-lines in ternary membranes containing diphytanoyl PC.Biophys. J.90, 4428–4436 (2006).

    ArticleCASGoogle Scholar

  178. Feigenson, G. W. & Buboltz, J. T. Ternary phase diagram of dipalmitoyl-PC/dilauroyl-PC/cholesterol: nanoscopic domain formation driven by cholesterol.Biophys. J.80, 2775–2788 (2001).

    ArticleCASGoogle Scholar

  179. Sezgin, E. et al. Partitioning, diffusion, and ligand binding of raft lipid analogs in model and cellular plasma membranes.Biochim. Biophys. Acta1818, 1777–1784 (2012).

    ArticleCASGoogle Scholar

  180. Kaiser, H. J. et al. Order of lipid phases in model and plasma membranes.Proc. Natl Acad. Sci. USA106, 16645–16650 (2009).

    ArticleCASGoogle Scholar

  181. Gerl, M. J. et al. Quantitative analysis of the lipidomes of the influenza virus envelope and MDCK cell apical membrane.J. Cell Biol.196, 213–221 (2012).

    ArticleCASGoogle Scholar

  182. Sampaio, J. L. et al. Membrane lipidome of an epithelial cell line.Proc. Natl Acad. Sci. USA108, 1903–1907 (2011).

    ArticleCASGoogle Scholar

  183. Franck, J. M., Chandrasekaran, S., Dzikovski, B., Dunnam, C. R. & Freed, J. H. Focus: two-dimensional electron-electron double resonance and molecular motions: the challenge of higher frequencies.J. Chem. Phys.142, 212302 (2015).

    ArticleGoogle Scholar

  184. Kuerschner, L. et al. Polyene-lipids: a new tool to image lipids.Nat. Methods2, 39–45 (2005).

    ArticleCASGoogle Scholar

  185. Ward, A. E., Ye, Y., Schuster, J. A., Wei, S. & Barrera, F. N. Single-molecule fluorescence vistas of how lipids regulate membrane proteins.Biochem. Soc. Trans.49, 1685–1694 (2021).

    ArticleCASGoogle Scholar

  186. McIntosh, A. L. et al. Fluorescence techniques using dehydroergosterol to study cholesterol trafficking.Lipids43, 1185–1208 (2008).

    ArticleCASGoogle Scholar

  187. Thiele, C., Hannah, M. J., Fahrenholz, F. & Huttner, W. B. Cholesterol binds to synaptophysin and is required for biogenesis of synaptic vesicles.Nat. Cell Biol.2, 42–49 (2000).

    ArticleCASGoogle Scholar

  188. Hoglinger, D. et al. Trifunctional lipid probes for comprehensive studies of single lipid species in living cells.Proc. Natl Acad. Sci. USA114, 1566–1571 (2017).

    ArticleGoogle Scholar

  189. Schuhmacher, M. et al. Live-cell lipid biochemistry reveals a role of diacylglycerol side-chain composition for cellular lipid dynamics and protein affinities.Proc. Natl Acad. Sci. USA117, 7729–7738 (2020).

    ArticleCASGoogle Scholar

  190. Romero, L. O. et al. Dietary fatty acids fine-tune Piezo1 mechanical response.Nat. Commun.10, 1200 (2019).

    ArticleGoogle Scholar

  191. Chakrapani, S., Cordero-Morales, J. F. & Perozo, E. A quantitative description of KcsA gating I: macroscopic currents.J. Gen. Physiol.130, 465–478 (2007).

    ArticleCASGoogle Scholar

  192. Garten, M. et al. Whole-GUV patch-clamping.Proc. Natl Acad. Sci. USA114, 328–333 (2017).

    ArticleCASGoogle Scholar

  193. Newport, T. D., Sansom, M. S. P. & Stansfeld, P. J. The MemProtMD database: a resource for membrane-embedded protein structures and their lipid interactions.Nucleic Acids Res.47, D390–D397 (2019).

    ArticleCASGoogle Scholar

  194. Kimura, Y. et al. Surface of bacteriorhodopsin revealed by high-resolution electron crystallography.Nature389, 206–211 (1997).

    ArticleCASGoogle Scholar

  195. Reichow, S. L. & Gonen, T. Lipid-protein interactions probed by electron crystallography.Curr. Opin. Struct. Biol.19, 560–565 (2009).

    ArticleCASGoogle Scholar

  196. Han, X., Yang, K. & Gross, R. W. Multi-dimensional mass spectrometry-based shotgun lipidomics and novel strategies for lipidomic analyses.Mass. Spectrom. Rev.31, 134–178 (2012).

    ArticleCASGoogle Scholar

  197. Laganowsky, A., Reading, E., Hopper, J. T. & Robinson, C. V. Mass spectrometry of intact membrane protein complexes.Nat. Protoc.8, 639–651 (2013).

    ArticleCASGoogle Scholar

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Acknowledgements

Major acknowledgement goes to K. Levental for insightful feedback and for generating the beautiful and informative figures for the manuscript. We also acknowledge the members of the Levental and Lyman labs for critical reading and enlightening discussions that have polished the ideas in the manuscript. This work was supported by NIH/National Institute of General Medical Sciences (GM134949, GM124072, GM120351), the Volkswagen Foundation (grant 93091) and the Human Frontiers Science Program (RGP0059/2019).

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  1. Department of Molecular Physiology and Biological Physics, Center for Molecular and Cell Physiology, University of Virginia, Charlottesville, VA, USA

    Ilya Levental

  2. Department of Physics and Astronomy, Department of Chemistry and Biochemistry, University of Delaware, Newark, DE, USA

    Ed Lyman

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Glossary

Unfolded protein response

A conserved cellular response to various stresses to the protein folding and secretory systems; can also be induced by lipid perturbations.

MscL and MscS families

Bacterial mechanosensitive channels that open in response to membrane tension, for example, induced by osmotic shock.

PIP2

Phosphatidylinositol 4,5-bisphosphate, a highly charged and tightly regulated lipid type that can associate tightly with integral membrane proteins through both electrostatic and stereospecific interactions.

Sphingolipids

Lipid with sphingoid backbones rather than glycerol. Lipids of this class (for example, sphingomyelin) are common components of eukaryote plasma membranes, interact well with sterols and bind strongly to some proteins.

Native mass spectrometry

A mass spectrometric technique capable of measuring molecular weights of large macromolecules (that is, proteins and their complexes) without fragmentation.

Shotgun lipidomics

A mass spectrometric technique for identifying and quantifying the lipid components of a complex sample (for example, cell membrane) without prior chromatographic separation.

Nanodiscs

An experimental construct containing an integral membrane protein, lipids and a scaffold that solubilizes them. The scaffold can be a protein (MSP) or synthetic polymer (for example, styrene maleic acid).

Gramicidin A

An antibiotic peptide that assembles to form transmembrane pores, dependent on its membrane environment.

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Levental, I., Lyman, E. Regulation of membrane protein structure and function by their lipid nano-environment.Nat Rev Mol Cell Biol24, 107–122 (2023). https://doi.org/10.1038/s41580-022-00524-4

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