Structural and functional consequences of reversible lipid asymmetry in living membranes


Maintenance of lipid asymmetry across the two leaflets of the plasma membrane (PM) bilayer is a ubiquitous feature of eukaryotic cells. Loss of this asymmetry has been widely associated with cell death. However, increasing evidence points to the physiological importance of non-apoptotic, transient changes in PM asymmetry. Such transient scrambling events are associated with a range of biological functions, including intercellular communication and intracellular signaling. Thus, regulation of interleaflet lipid distribution in the PM is a broadly important but underappreciated cellular process with key physiological and structural consequences. Here, we compile the mounting evidence revealing multifaceted, functional roles of PM asymmetry and transient loss thereof. We discuss the consequences of reversible asymmetry on PM structure, biophysical properties and interleaflet coupling. We argue that despite widespread recognition of broad aspects of membrane asymmetry, its importance in cell biology demands more in-depth investigation of its features, regulation, and physiological and pathological implications.

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Fig. 1: The interleaflet asymmetry of the PM bilayer is complex and dynamic.
Fig. 2: Functional roles of reversible PM lipid asymmetry.
Fig. 3: Effects of lipid scrambling on PM physical properties.
Fig. 4: Possible configurations for the interleaflet distribution of cholesterol in the PM.


  1. 1.

    Singer, S. J. & Nicolson, G. L. The fluid mosaic model of the structure of cell membranes. Science 175, 720–731 (1972).

    CAS  PubMed  Google Scholar 

  2. 2.

    Verkleij, A. J. et al. The asymmetric distribution of phospholipids in the human red cell membrane. A combined study using phospholipases and freeze-etch electron microscopy. Biochim. Biophys. Acta 323, 178–193 (1973). This study provides direct experimental evidence for the asymmetric distribution of different lipid classes in the plasma membrane of red blood cells.

  3. 3.

    Sessions, A. & Horwitz, A. F. Myoblast aminophospholipid asymmetry differs from that of fibroblasts. FEBS Lett. 134, 75–78 (1981).

    CAS  PubMed  Google Scholar 

  4. 4.

    Bevers, E. M. & Williamson, P. L. Getting to the outer leaflet: physiology of phosphatidylserine exposure at the plasma membrane. Physiol. Rev. 96, 605–645 (2016).

    CAS  PubMed  Google Scholar 

  5. 5.

    Lorent, J. H. et al. Plasma membranes are asymmetric in lipid unsaturation, packing and protein shape. Nat. Chem. Biol. 16, 644–652 (2020). This paper reports the detailed phospholipid compositions of the exo- and endoplasmic leaflets of RBC PMs, including chain unsaturation and distribution of rare lipids. It also presents evidence for differential packing/order of the two PM leaflets in both RBCs and cultured nucleated cells.

  6. 6.

    Makarova, M. & Owen, D. M. Asymmetry across the membrane. Nat. Chem. Biol. 16, 605–606 (2020).

    CAS  PubMed  Google Scholar 

  7. 7.

    Zachowski, A. Phospholipids in animal eukaryotic membranes: transverse asymmetry and movement. Biochem. J. 294, 1–14 (1993).

    CAS  PubMed  PubMed Central  Google Scholar 

  8. 8.

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

  9. 9.

    Gurtovenko, A. A. & Vattulainen, I. Lipid transmembrane asymmetry and intrinsic membrane potential: two sides of the same coin. J. Am. Chem. Soc. 129, 5358–5359 (2007).

    CAS  PubMed  Google Scholar 

  10. 10.

    Ma, Y. et al. A FRET sensor enables quantitative measurements of membrane charges in live cells. Nat. Biotechnol. 35, 363–370 (2017).

    CAS  PubMed  Google Scholar 

  11. 11.

    Entova, S., Billod, J. M., Swiecicki, J. M., Martín-Santamaría, S. & Imperiali, B. Insights into the key determinants of membrane protein topology enable the identification of new monotopic folds. eLife 7, e40889 (2018).

    PubMed  PubMed Central  Google Scholar 

  12. 12.

    Dowhan, W., Vitrac, H. & Bogdanov, M. Lipid-assisted membrane protein folding and topogenesis. Protein J. 38, 274–288 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  13. 13.

    Kapus, A. & Janmey, P. Plasma membrane–cortical cytoskeleton interactions: a cell biology approach with biophysical considerations. Compr. Physiol. 3, 1231–1281 (2013).

    PubMed  Google Scholar 

  14. 14.

    McLaughlin, S. & Murray, D. Plasma membrane phosphoinositide organization by protein electrostatics. Nature 438, 605–611 (2005).

    CAS  PubMed  Google Scholar 

  15. 15.

    Platre, M. P. & Jaillais, Y. Anionic lipids and the maintenance of membrane electrostatics in eukaryotes. Plant Signal. Behav. 12, e1282022 (2017).

    PubMed  PubMed Central  Google Scholar 

  16. 16.

    Li, L., Shi, X., Guo, X., Li, H. & Xu, C. Ionic protein-lipid interaction at the plasma membrane: what can the charge do? Trends Biochem. Sci. 39, 130–140 (2014).

    CAS  PubMed  Google Scholar 

  17. 17.

    Swamy, M. J. et al. Coexisting domains in the plasma membranes of live cells characterized by spin-label ESR spectroscopy. Biophys. J. 90, 4452–4465 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  18. 18.

    Gupta, A., Korte, T., Herrmann, A. & Wohland, T. Plasma membrane asymmetry of lipid organization: fluorescence lifetime microscopy and correlation spectroscopy analysis. J. Lipid Res. 61, 252–266 (2020). Using fluorescent analogs, the authors report on the leaflet-specific biophysical properties (including packing and diffusion) of different lipid classes in the PM of nucleated cells. Their results point to a more fluid cytosolic leaflet and more tightly packed exoplasmic leaflet.

  19. 19.

    Hill, W. G. & Zeidel, M. L. Reconstituting the barrier properties of a water-tight epithelial membrane by design of leaflet-specific liposomes. J. Biol. Chem. 275, 30176–30185 (2000).

    CAS  PubMed  Google Scholar 

  20. 20.

    Williamson, P. & Schlegel, R. A. Back and forth: the regulation and function of transbilayer phospholipid movement in eukaryotic cells. Mol. Membr. Biol. 11, 199–216 (1994).

    CAS  PubMed  Google Scholar 

  21. 21.

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

    CAS  PubMed  Google Scholar 

  22. 22.

    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. USA 101, 4083–4088 (2004).

    CAS  PubMed  Google Scholar 

  23. 23.

    Lorent, J. H. et al. Structural determinants and functional consequences of protein affinity for membrane rafts. Nat. Commun. 8, 1219 (2017).

    PubMed  PubMed Central  Google Scholar 

  24. 24.

    Montigny, C., Lyons, J., Champeil, P., Nissen, P. & Lenoir, G. On the molecular mechanism of flippase- and scramblase-mediated phospholipid transport. Biochim. Biophys. Acta 1861, 767–783 (2016). 8 Pt B.

    CAS  PubMed  Google Scholar 

  25. 25.

    Daleke, D. L. Phospholipid flippases. J. Biol. Chem. 282, 821–825 (2007).

    CAS  PubMed  Google Scholar 

  26. 26.

    Kodigepalli, K. M., Bowers, K., Sharp, A. & Nanjundan, M. Roles and regulation of phospholipid scramblases. FEBS Lett. 589, 3–14 (2015).

    CAS  PubMed  Google Scholar 

  27. 27.

    Falzone, M. E. et al. Structural basis of Ca2+-dependent activation and lipid transport by a TMEM16 scramblase. eLife 8, e43229 (2019).

    PubMed  PubMed Central  Google Scholar 

  28. 28.

    Nagata, S., Suzuki, J., Segawa, K. & Fujii, T. Exposure of phosphatidylserine on the cell surface. Cell Death Differ. 23, 952–961 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  29. 29.

    Shlomovitz, I., Speir, M. & Gerlic, M. Flipping the dogma—phosphatidylserine in non-apoptotic cell death. Cell Commun. Signal. 17, 139 (2019).

    PubMed  PubMed Central  Google Scholar 

  30. 30.

    Hammill, A. K., Uhr, J. W. & Scheuermann, R. H. Annexin V staining due to loss of membrane asymmetry can be reversible and precede commitment to apoptotic death. Exp. Cell Res. 251, 16–21 (1999). Observations in this work provide one of the earliest examples that exposure of PS on the cell surface can be reversible and does not have to lead to cell death.

  31. 31.

    Shin, H. W. & Takatsu, H. Phosphatidylserine exposure in living cells. Crit. Rev. Biochem. Mol. Biol. 55, 166–178 (2020).

    CAS  PubMed  Google Scholar 

  32. 32.

    Brown, G. C. & Neher, J. J. Microglial phagocytosis of live neurons. Nat. Rev. Neurosci. 15, 209–216 (2014).

    CAS  PubMed  Google Scholar 

  33. 33.

    Neher, J. J. et al. Inhibition of microglial phagocytosis is sufficient to prevent inflammatory neuronal death. J. Immunol. 186, 4973–4983 (2011).

    CAS  PubMed  Google Scholar 

  34. 34.

    Neumann, B. et al. EFF-1-mediated regenerative axonal fusion requires components of the apoptotic pathway. Nature 517, 219–222 (2015).

    CAS  PubMed  Google Scholar 

  35. 35.

    Hisamoto, N. et al. Phosphatidylserine exposure mediated by ABC transporter activates the integrin signaling pathway promoting axon regeneration. Nat. Commun. 9, 3099 (2018).

    PubMed  PubMed Central  Google Scholar 

  36. 36.

    Scott-Hewitt, N. et al. Local externalization of phosphatidylserine mediates developmental synaptic pruning by microglia. EMBO J. 39, e105380 (2020).

  37. 37.

    Ruggiero, L., Connor, M. P., Chen, J., Langen, R. & Finnemann, S. C. Diurnal, localized exposure of phosphatidylserine by rod outer segment tips in wild-type but not Itgb5–/– or Mfge8–/– mouse retina. Proc. Natl. Acad. Sci. USA 109, 8145–8148 (2012).

    CAS  PubMed  Google Scholar 

  38. 38.

    Goodyear, R. J., Gale, J. E., Ranatunga, K. M., Kros, C. J. & Richardson, G. P. Aminoglycoside-induced phosphatidylserine externalization in sensory hair cells is regionally restricted, rapid, and reversible. J. Neurosci. 28, 9939–9952 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  39. 39.

    van den Eijnde, S. M. et al. Transient expression of phosphatidylserine at cell-cell contact areas is required for myotube formation. J. Cell Sci. 114, 3631–3642 (2001). In this study the authors show direct imaging of transient and punctate PS exposure on the surface of differentiating myoblasts during fusion into myotubes in both mouse embryos and cultured cells. The PS exposure is non-apoptotic in origin, and its onset and subsequent disappearance from the cell surface are coincident with the expression and reorganization of a developmental marker, suggesting involvement of scrambling in muscle development.

  40. 40.

    Jeong, J. & Conboy, I. M. Phosphatidylserine directly and positively regulates fusion of myoblasts into myotubes. Biochem. Biophys. Res. Commun. 414, 9–13 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. 41.

    Park, S. Y. et al. Stabilin-2 modulates the efficiency of myoblast fusion during myogenic differentiation and muscle regeneration. Nat. Commun. 7, 10871 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  42. 42.

    Tsuchiya, M. et al. Cell surface flip-flop of phosphatidylserine is critical for PIEZO1-mediated myotube formation. Nat. Commun. 9, 2049 (2018).

    PubMed  PubMed Central  Google Scholar 

  43. 43.

    de Vries, K. J., Wiedmer, T., Sims, P. J. & Gadella, B. M. Caspase-independent exposure of aminophospholipids and tyrosine phosphorylation in bicarbonate responsive human sperm cells. Biol. Reprod. 68, 2122–2134 (2003).

    PubMed  Google Scholar 

  44. 44.

    Rival, C. M. et al. Phosphatidylserine on viable sperm and phagocytic machinery in oocytes regulate mammalian fertilization. Nat. Commun. 10, 4456 (2019). The authors show not only that PS exposure plays a direct role in sperm–egg fusion but also that PS exposing mammalian sperm can fuse with myoblasts, likely through a PS recognition receptor similar to that on oocytes. This result points to a ubiquitous form of cell–cell fusion mediated through changes in PM lipid asymmetry.

  45. 45.

    Naeini, M. B., Bianconi, V., Pirro, M. & Sahebkar, A. The role of phosphatidylserine recognition receptors in multiple biological functions. Cell. Mol. Biol. Lett. 25, 23 (2020).

    PubMed  PubMed Central  Google Scholar 

  46. 46.

    Martin, S. et al. Immunologic stimulation of mast cells leads to the reversible exposure of phosphatidylserine in the absence of apoptosis. Int. Arch. Allergy Immunol. 123, 249–258 (2000).

    CAS  PubMed  Google Scholar 

  47. 47.

    Rysavy, N. M. et al. Beyond apoptosis: the mechanism and function of phosphatidylserine asymmetry in the membrane of activating mast cells. Bioarchitecture 4, 127–137 (2014).

    PubMed  Google Scholar 

  48. 48.

    Ory, S. et al. Phospholipid scramblase-1-induced lipid reorganization regulates compensatory endocytosis in neuroendocrine cells. J. Neurosci. 33, 3545–3556 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  49. 49.

    MacKenzie, A. et al. Rapid secretion of interleukin-1beta by microvesicle shedding. Immunity 15, 825–835 (2001).

    CAS  PubMed  Google Scholar 

  50. 50.

    Elliott, J. I. et al. Membrane phosphatidylserine distribution as a non-apoptotic signalling mechanism in lymphocytes. Nat. Cell Biol. 7, 808–816 (2005).

    CAS  PubMed  Google Scholar 

  51. 51.

    Bleibaum, F. et al. ADAM10 sheddase activation is controlled by cell membrane asymmetry. J. Mol. Cell Biol. 11, 979–993 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  52. 52.

    Sommer, A. et al. Phosphatidylserine exposure is required for ADAM17 sheddase function. Nat. Commun. 7, 11523 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  53. 53.

    Fischer, K. et al. Antigen recognition induces phosphatidylserine exposure on the cell surface of human CD8+ T cells. Blood 108, 4094–4101 (2006).

    CAS  PubMed  Google Scholar 

  54. 54.

    Stowell, S. R. et al. Galectin-1 induces reversible phosphatidylserine exposure at the plasma membrane. Mol. Biol. Cell 20, 1408–1418 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  55. 55.

    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).

    CAS  PubMed  PubMed Central  Google Scholar 

  56. 56.

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

    CAS  PubMed  PubMed Central  Google Scholar 

  57. 57.

    Sezgin, E. et al. Measuring nanoscale diffusion dynamics in cellular membranes with super-resolution STED-FCS. Nat. Protoc. 14, 1054–1083 (2019).

    CAS  PubMed  Google Scholar 

  58. 58.

    Morrot, G. et al. Asymmetric lateral mobility of phospholipids in the human erythrocyte membrane. Proc. Natl. Acad. Sci. USA 83, 6863–6867 (1986).

    CAS  PubMed  Google Scholar 

  59. 59.

    el Hage Chahine, J. M., Cribier, S. & Devaux, P. F. Phospholipid transmembrane domains and lateral diffusion in fibroblasts. Proc. Natl. Acad. Sci. USA 90, 447–451 (1993).

    CAS  PubMed  Google Scholar 

  60. 60.

    Cogan, U. & Schachter, D. Asymmetry of lipid dynamics in human erythrocyte membranes studied with impermeant fluorophores. Biochemistry 20, 6396–6403 (1981).

    CAS  PubMed  Google Scholar 

  61. 61.

    Nikolova-Karakashian, M. N., Petkova, H. & Koumanov, K. S. Influence of cholesterol on sphingomyelin metabolism and hemileaflet fluidity of rat liver plasma membranes. Biochimie 74, 153–159 (1992).

    CAS  PubMed  Google Scholar 

  62. 62.

    Bogdanov, M. et al. Phospholipid distribution in the cytoplasmic membrane of Gram-negative bacteria is highly asymmetric, dynamic, and cell shape-dependent. Sci. Adv. 6, eaaz6333 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  63. 63.

    Diaz-Rohrer, B. B., Levental, K. R., Simons, K. & Levental, I. Membrane raft association is a determinant of plasma membrane localization. Proc. Natl. Acad. Sci. USA 111, 8500–8505 (2014).

    CAS  PubMed  Google Scholar 

  64. 64.

    Heberle, F. A. et al. Direct label-free imaging of nanodomains in biomimetic and biological membranes by cryogenic electron microscopy. Proc. Natl. Acad. Sci. USA 117, 19943–19952 (2020).

    CAS  PubMed  Google Scholar 

  65. 65.

    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. USA 117, 19713–19719 (2020).

    CAS  PubMed  Google Scholar 

  66. 66.

    Demo, S. D. et al. Quantitative measurement of mast cell degranulation using a novel flow cytometric annexin-V binding assay. Cytometry 36, 340–348 (1999).

    CAS  PubMed  Google Scholar 

  67. 67.

    Smrz, D., Dráberová, L. & Dráber, P. Non-apoptotic phosphatidylserine externalization induced by engagement of glycosylphosphatidylinositol-anchored proteins. J. Biol. Chem. 282, 10487–10497 (2007).

    CAS  PubMed  Google Scholar 

  68. 68.

    Pyrshev, K. A., Klymchenko, A. S., Csúcs, G. & Demchenko, A. P. Apoptosis and eryptosis: striking differences on biomembrane level. Biochim. Biophys. Acta Biomembr. 1860, 1362–1371 (2018).

    CAS  PubMed  Google Scholar 

  69. 69.

    Le Roux, A. L., Quiroga, X., Walani, N., Arroyo, M. & Roca-Cusachs, P. The plasma membrane as a mechanochemical transducer. Phil. Trans. R. Soc. Lond. B 374, 20180221 (2019).

    Google Scholar 

  70. 70.

    Lacoste, D. & Bassereau, P. in Liposomes, Lipid Bilayers and Model Membranes: From Basic Research to Application. p271 (CRC Press, Taylor & Francis Group, 2014).

  71. 71.

    Shi, Z., Graber, Z. T., Baumgart, T., Stone, H. A. & Cohen, A. E. Cell membranes resist flow. Cell 175, 1769–1779 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  72. 72.

    Dumitru, A. C. et al. Nanoscale membrane architecture of healthy and pathological red blood cells. Nanoscale Horiz. 3, 293–304 (2018).

    CAS  PubMed  Google Scholar 

  73. 73.

    Hossein, A. & Deserno, M. Spontaneous curvature, differential stress, and bending modulus of asymmetric lipid membranes. Biophys. J. 118, 624–642 (2020). This paper formalizes the concept of differential stress, i.e., stress caused by suboptimal packing of the lipids in the two membrane leaflets, and demonstrates the dramatic effects that it can have on the membrane’s mechanical properties.

  74. 74.

    Doktorova, M. & Weinstein, H. Accurate in silico modeling of asymmetric bilayers based on biophysical principles. Biophys. J. 115, 1638–1643 (2018). This study discusses the importance of ensuring the absence of differential stress, i.e., the zero-tension state, in the two bilayer leaflets when comparing asymmetric to symmetric membranes in the pursuit of mechanistic insights into interleaflet coupling effects in model systems.

  75. 75.

    Steinkühler, J., Sezgin, E., Urbančič, I., Eggeling, C. & Dimova, R. Mechanical properties of plasma membrane vesicles correlate with lipid order, viscosity and cell density. Commun Biol 2, 337 (2019).

    PubMed  PubMed Central  Google Scholar 

  76. 76.

    Keller, H., Lorizate, M. & Schwille, P. PI(4,5)P2 degradation promotes the formation of cytoskeleton-free model membrane systems. ChemPhysChem 10, 2805–2812 (2009).

    CAS  PubMed  Google Scholar 

  77. 77.

    Baumgart, T. et al. Large-scale fluid/fluid phase separation of proteins and lipids in giant plasma membrane vesicles. Proc. Natl. Acad. Sci. USA 104, 3165–3170 (2007).

    CAS  PubMed  Google Scholar 

  78. 78.

    Skinkle, A. D., Levental, K. R. & Levental, I. Cell-derived plasma membrane vesicles are permeable to hydrophilic macromolecules. Biophys. J. 118, 1292–1300 (2020). This study provides experimental evidence for the presence of pores of varying sizes on the surface of giant plasma-membrane vesicles, calling into question their ability to retain and maintain the native lipid asymmetry of the cell PM.

  79. 79.

    Skotland, T. & Sandvig, K. The role of PS 18:0/18:1 in membrane function. Nat. Commun. 10, 2752 (2019).

    PubMed  PubMed Central  Google Scholar 

  80. 80.

    Koyama-Honda, I. et al. High-speed single-molecule imaging reveals signal transduction by induced transbilayer raft phases. J. Cell Biol. 219, e202006125 (2020). This study presents a clear example of interleaflet communication in the PM of nucleated cells whereas clustering of GPI-anchored proteins on the exoplasmic leaflet initiates a signaling cascade by recruiting lipidated proteins to the cytosolic leaflet right under the GPI clusters.

  81. 81.

    Fujimoto, T. & Parmryd, I. Interleaflet coupling, pinning, and leaflet asymmetry-major players in plasma membrane nanodomain formation. Front. Cell Dev. Biol. 4, 155 (2017).

    PubMed  PubMed Central  Google Scholar 

  82. 82.

    Abe, M. et al. A role for sphingomyelin-rich lipid domains in the accumulation of phosphatidylinositol-4,5-bisphosphate to the cleavage furrow during cytokinesis. Mol. Cell. Biol. 32, 1396–1407 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  83. 83.

    Róg, T. et al. Interdigitation of long-chain sphingomyelin induces coupling of membrane leaflets in a cholesterol dependent manner. Biochim. Biophys. Acta 1858, 281–288 (2016).

    PubMed  Google Scholar 

  84. 84.

    Gupta, A., Muralidharan, S., Torta, F., Wenk, M. R. & Wohland, T. Long acyl chain ceramides govern cholesterol and cytoskeleton dependence of membrane outer leaflet dynamics. Biochim. Biophys. Acta Biomembr. 1862, 183153 (2020).

    CAS  PubMed  Google Scholar 

  85. 85.

    Sarmento, M. J., Hof, M. & Šachl, R. Interleaflet coupling of lipid nanodomains—insights from in vitro systems. Front. Cell Dev. Biol. 8, 284 (2020).

    PubMed  PubMed Central  Google Scholar 

  86. 86.

    Seo, S., Murata, M. & Shinoda, W. Pivotal role of interdigitation in interleaflet interactions: implications from molecular dynamics simulations. J. Phys. Chem. Lett. 11, 5171–5176 (2020).

    CAS  PubMed  Google Scholar 

  87. 87.

    Marrink, S. J. et al. Computational modeling of realistic cell membranes. Chem. Rev. 119, 6184–6226 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  88. 88.

    Bennett, W. F., MacCallum, J. L., Hinner, M. J., Marrink, S. J. & Tieleman, D. P. Molecular view of cholesterol flip-flop and chemical potential in different membrane environments. J. Am. Chem. Soc. 131, 12714–12720 (2009).

    CAS  PubMed  Google Scholar 

  89. 89.

    Liu, S. L. et al. Orthogonal lipid sensors identify transbilayer asymmetry of plasma membrane cholesterol. Nat. Chem. Biol. 13, 268–274 (2017).

    CAS  PubMed  Google Scholar 

  90. 90.

    Courtney, K. C. et al. C24 sphingolipids govern the transbilayer asymmetry of cholesterol and lateral organization of model and live-cell plasma membranes. Cell Reports 24, 1037–1049 (2018).

    CAS  PubMed  Google Scholar 

  91. 91.

    Steck, T. L. & Lange, Y. Transverse distribution of plasma membrane bilayer cholesterol: picking sides. Traffic 19, 750–760 (2018).

    CAS  PubMed  Google Scholar 

  92. 92.

    Bruckner, R. J., Mansy, S. S., Ricardo, A., Mahadevan, L. & Szostak, J. W. Flip-flop-induced relaxation of bending energy: implications for membrane remodeling. Biophys. J. 97, 3113–3122 (2009). This paper presents experimental evidence for the ability of cholesterol to quickly redistribute between the two leaflets of a bilayer and thus act as a buffer for deformation-induced membrane tension in osmotically challenged model systems.

  93. 93.

    Miettinen, M. S. & Lipowsky, R. Bilayer membranes with frequent flip-flops have tensionless leaflets. Nano Lett. 19, 5011–5016 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  94. 94.

    Nyholm, T. K. M., Jaikishan, S., Engberg, O., Hautala, V. & Slotte, J. P. The affinity of sterols for different phospholipid classes and its impact on lateral segregation. Biophys. J. 116, 296–307 (2019). The authors measure and compare the partitioning of cholesterol and fluorescent cholesterol analogs in different phospholipid environments. They report a link between a lower affinity for unsaturated PLs and higher propensity for lateral segregation and observe striking differences between the partitioning behavior of cholesterol and its fluorescent analogs that may help explain some of the conflicting results on cholesterol distribution in cell PMs reported in the literature.

  95. 95.

    Wassall, S. R. & Stillwell, W. Polyunsaturated fatty acid-cholesterol interactions: domain formation in membranes. Biochim. Biophys. Acta 1788, 24–32 (2009).

    CAS  PubMed  Google Scholar 

  96. 96.

    Marsh, D. Handbook of Lipid Bilayers 2nd edn (CRC Press, 2013).

  97. 97.

    Doktorova, M. et al. Cholesterol promotes protein binding by affecting membrane electrostatics and solvation properties. Biophys. J. 113, 2004–2015 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  98. 98.

    Leeb, F. & Maibaum, L. Spatially resolving the condensing effect of cholesterol in lipid bilayers. Biophys. J. 115, 2179–2188 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  99. 99.

    Evans, E., Rawicz, W. & Smith, B. A. Back to the future: mechanics and thermodynamics of lipid biomembranes. Faraday Discuss. 161, 591–611 (2013).

    CAS  PubMed  Google Scholar 

  100. 100.

    Chakraborty, S. et al. Reassessment of membrane mechanics: how cholesterol stiffens unsaturated lipid membranes. Proc. Natl. Acad. Sci. USA 117, 21896–21905 (2020).

    CAS  PubMed  Google Scholar 

  101. 101.

    Kollmitzer, B., Heftberger, P., Rappolt, M. & Pabst, G. Monolayer spontaneous curvature of raft-forming membrane lipids. Soft Matter 9, 10877–10884 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  102. 102.

    Sodt, A. J., Venable, R. M., Lyman, E. & Pastor, R. W. Nonadditive compositional curvature energetics of lipid bilayers. Phys. Rev. Lett. 117, 138104 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  103. 103.

    Renooij, W., Van Golde, L. M., Zwaal, R. F., Roelofsen, B. & Van Deenen, L. L. Preferential incorporation of fatty acids at the inside of human erythrocyte membranes. Biochim. Biophys. Acta 363, 287–292 (1974).

    CAS  PubMed  Google Scholar 

  104. 104.

    Rawyler, A., van der Schaft, P. H., Roelofsen, B. & Op den Kamp, J. A. Phospholipid localization in the plasma membrane of Friend erythroleukemic cells and mouse erythrocytes. Biochemistry 24, 1777–1783 (1985).

    CAS  PubMed  Google Scholar 

  105. 105.

    Van der Schaft, P. H., Roelofsen, B., Op den Kamp, J. A. & Van Deenen, L. L. Phospholipid asymmetry during erythropoiesis. A study on Friend erythroleukemic cells and mouse reticulocytes. Biochim. Biophys. Acta 900, 103–115 (1987).

    PubMed  Google Scholar 

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Funding for I.L. was provided by the US National Institutes of Health/National Institute of General Medical Sciences (R35 GM134949, R01 GM124072, R21 AI146880), the Volkswagen Foundation (93091), and the Human Frontiers Science Program (RGP0059/2019). M.D. was supported by grant F32 GM134704. J.L.S. was supported by grant T32 GM008280.

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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).

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