Regulation of phospholipid distribution in the lipid bilayer by flippases and scramblases.
Journal
Nature reviews. Molecular cell biology
ISSN: 1471-0080
Titre abrégé: Nat Rev Mol Cell Biol
Pays: England
ID NLM: 100962782
Informations de publication
Date de publication:
08 2023
08 2023
Historique:
accepted:
09
03
2023
medline:
24
7
2023
pubmed:
28
4
2023
entrez:
27
4
2023
Statut:
ppublish
Résumé
Cellular membranes function as permeability barriers that separate cells from the external environment or partition cells into distinct compartments. These membranes are lipid bilayers composed of glycerophospholipids, sphingolipids and cholesterol, in which proteins are embedded. Glycerophospholipids and sphingolipids freely move laterally, whereas transverse movement between lipid bilayers is limited. Phospholipids are asymmetrically distributed between membrane leaflets but change their location in biological processes, serving as signalling molecules or enzyme activators. Designated proteins - flippases and scramblases - mediate this lipid movement between the bilayers. Flippases mediate the confined localization of specific phospholipids (phosphatidylserine (PtdSer) and phosphatidylethanolamine) to the cytoplasmic leaflet. Scramblases randomly scramble phospholipids between leaflets and facilitate the exposure of PtdSer on the cell surface, which serves as an important signalling molecule and as an 'eat me' signal for phagocytes. Defects in flippases and scramblases cause various human diseases. We herein review the recent research on the structure of flippases and scramblases and their physiological roles. Although still poorly understood, we address the mechanisms by which they translocate phospholipids between lipid bilayers and how defects cause human diseases.
Identifiants
pubmed: 37106071
doi: 10.1038/s41580-023-00604-z
pii: 10.1038/s41580-023-00604-z
pmc: PMC10134735
doi:
Substances chimiques
Lipid Bilayers
0
Phospholipids
0
Glycerophospholipids
0
Phosphatidylserines
0
Types de publication
Journal Article
Review
Research Support, Non-U.S. Gov't
Langues
eng
Sous-ensembles de citation
IM
Pagination
576-596Commentaires et corrections
Type : ErratumIn
Informations de copyright
© 2023. Springer Nature Limited.
Références
Vance, J. E. Phospholipid synthesis and transport in mammalian cells. Traffic 16, 1–18 (2015).
pubmed: 25243850
doi: 10.1111/tra.12230
Yang, Y., Lee, M. & Fairn, G. D. Phospholipid subcellular localization and dynamics. J. Biol. Chem. 293, 6230–6240 (2018).
pubmed: 29588369
doi: 10.1074/jbc.R117.000582
pmcid: 5925819
Bretscher, M. S. Asymmetrical lipid bilayer structure for biological membranes. Nat. N. Biol. 236, 11–12 (1972). This article reports the asymmetrical distribution of phospholipids in the plasma membrane by treating human erythrocytes or their membrane fraction with chemical reagents that specifically label amino groups.
doi: 10.1038/newbio236011a0
Tsuji, T. et al. Predominant localization of phosphatidylserine at the cytoplasmic leaflet of the ER, and its TMEM16K-dependent redistribution. Proc. Natl Acad. Sci. USA 116, 13368–13373 (2019).
pubmed: 31217287
doi: 10.1073/pnas.1822025116
pmcid: 6613088
Murate, M. et al. Transbilayer distribution of lipids at nano scale. J. Cell Sci. 128, 1627–1638 (2015).
pubmed: 25673880
Clarke, R. J., Hossain, K. R. & Cao, K. Physiological roles of transverse lipid asymmetry of animal membranes. Biochim. Biophy. Acta, Biomembr. 1862, 183382 (2020).
doi: 10.1016/j.bbamem.2020.183382
Lorent, J. H. et al. Plasma membranes are asymmetric in lipid unsaturation, packing and protein shape. Nat. Chem. Biol. 16, 644–652 (2020).
pubmed: 32367017
doi: 10.1038/s41589-020-0529-6
pmcid: 7246138
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).
pubmed: 26936867
doi: 10.1152/physrev.00020.2015
Kobayashi, T. & Menon, A. K. Transbilayer lipid asymmetry. Curr. Biol. 28, R386–R391 (2018).
pubmed: 29689220
doi: 10.1016/j.cub.2018.01.007
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).
pubmed: 33199908
doi: 10.1038/s41589-020-00688-0
pmcid: 7747298
Meca, J. et al. Avidity-driven polarity establishment via multivalent lipid–GTPase module interactions. EMBO J. 38, e99652 (2019).
pubmed: 30559330
doi: 10.15252/embj.201899652
Nagata, S., Suzuki, J., Segawa, K. & Fujii, T. Exposure of phosphatidylserine on the cell surface. Cell Death Differ. 23, 952–961 (2016).
pubmed: 26891692
doi: 10.1038/cdd.2016.7
pmcid: 4987739
Coleman, J. A., Quazi, F. & Molday, R. S. Mammalian P4-ATPases and ABC transporters and their role in phospholipid transport. Biochim. Biophys. Acta 1831, 555–574 (2013).
pubmed: 23103747
doi: 10.1016/j.bbalip.2012.10.006
Palmgren, M., Østerberg, J. T., Nintemann, S. J., Poulsen, L. R. & López-Marqués, R. L. Evolution and a revised nomenclature of P4 ATPases, a eukaryotic family of lipid flippases. Biochim. Biophys. Acta, Biomembr. 1861, 1135–1151 (2019).
pubmed: 30802428
doi: 10.1016/j.bbamem.2019.02.006
Jinek, M. et al. A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337, 816–821 (2012).
pubmed: 22745249
doi: 10.1126/science.1225829
pmcid: 6286148
Tanay, A. & Regev, A. Scaling single-cell genomics from phenomenology to mechanism. Nature 541, 331–338 (2017).
pubmed: 28102262
doi: 10.1038/nature21350
pmcid: 5438464
Cheng, Y., Grigorieff, N., Penczek, Pawel, A. & Walz, T. A primer to single-particle cryo-electron microscopy. Cell 161, 438–449 (2015).
pubmed: 25910204
doi: 10.1016/j.cell.2015.03.050
pmcid: 4409659
Jumper, J. et al. Highly accurate protein structure prediction with AlphaFold. Nature 596, 583–589 (2021).
pubmed: 34265844
doi: 10.1038/s41586-021-03819-2
pmcid: 8371605
Bretscher, M. S. Membrane structure: some general principles. Science 181, 622–629 (1973).
pubmed: 4724478
doi: 10.1126/science.181.4100.622
Leventis, P. A. & Grinstein, S. The distribution and function of phosphatidylserine in cellular membranes. Annu. Rev. Biophys. 39, 407–427 (2010).
pubmed: 20192774
doi: 10.1146/annurev.biophys.093008.131234
Seigneuret, M. & Devaux, P. ATP-dependent asymmetric distribution of spin-labeled phospholipids in the erythrocyte membrane: relation to shape changes. Proc. Natl Acad. Sci. USA 81, 3751–3755 (1984).
pubmed: 6587389
doi: 10.1073/pnas.81.12.3751
pmcid: 345297
Auland, M., Roufogalis, B., Devaux, P. & Zachowski, A. Reconstitution of ATP-dependent aminophospholipid translocation in proteoliposomes. Proc. Natl Acad. Sci. USA 91, 10938–10942 (1994). This work purifies an ATPase from human erythrocyte membranes, and reconstitutes its flippase activity in proteoliposomes.
pubmed: 7971987
doi: 10.1073/pnas.91.23.10938
pmcid: 45141
Tang, X., Halleck, M. S., Schlegel, R. A. & Williamson, P. A subfamily of P-type ATPases with aminophospholipid transporting activity. Science 272, 1495–1497 (1996). This article reports the identification of mammalian P4-type ATPase (flippase) that transports amino phospholipids.
pubmed: 8633245
doi: 10.1126/science.272.5267.1495
Palmgren, M. G. & Nissen, P. P-type ATPases. Annu. Rev. Biophys. 40, 243–266 (2011).
pubmed: 21351879
doi: 10.1146/annurev.biophys.093008.131331
Segawa, K. et al. Caspase-mediated cleavage of phospholipid flippase for apoptotic phosphatidylserine exposure. Science 344, 1164–1168 (2014). This article reports that a P4-ATPase, ATP11C, is complexed with its chaperone, CDC50A, works as the flippase at the plasma membrane and is cleaved by caspase 3 when cells undergo apoptosis.
pubmed: 24904167
doi: 10.1126/science.1252809
Segawa, K., Kurata, S. & Nagata, S. Human type IV P-type ATPases that work as plasma membrane phospholipid flippases, and their regulation by caspase and calcium. J. Biol. Chem. 291, 762–772 (2016).
pubmed: 26567335
doi: 10.1074/jbc.M115.690727
Wang, J. et al. Proteomic analysis and functional characterization of P4-ATPase phospholipid flippases from murine tissues. Sci. Rep. 8, 10795 (2018).
pubmed: 30018401
doi: 10.1038/s41598-018-29108-z
pmcid: 6050252
Coleman, J. A., Kwok, M. C. & Molday, R. S. Localization, purification, and functional reconstitution of the P4-ATPase Atp8a2, a phosphatidylserine flippase in photoreceptor disc membranes. J. Biol. Chem. 284, 32670–32679 (2009).
pubmed: 19778899
doi: 10.1074/jbc.M109.047415
pmcid: 2781682
Lee, S. et al. Transport through recycling endosomes requires EHD1 recruitment by a phosphatidylserine translocase. EMBO J. 34, 669–688 (2015).
pubmed: 25595798
doi: 10.15252/embj.201489703
pmcid: 4365035
Cheng, M.-T. et al. Structural insights into the activation of autoinhibited human lipid flippase ATP8B1 upon substrate binding. Proc. Natl Acad. Sci. USA 119, e2118656119 (2022).
pubmed: 35349344
doi: 10.1073/pnas.2118656119
pmcid: 9168909
Dieudonné, T. et al. Autoinhibition and regulation by phosphoinositides of ATP8B1, a human lipid flippase associated with intrahepatic cholestatic disorders. eLife 11, e75272 (2022).
pubmed: 35416773
doi: 10.7554/eLife.75272
pmcid: 9045818
Martin, S. et al. Mutated ATP10B increases Parkinson’s disease risk by compromising lysosomal glucosylceramide export. Acta Neuropathol. 139, 1001–1024 (2020).
pubmed: 32172343
doi: 10.1007/s00401-020-02145-7
pmcid: 7244618
Best, J. T., Xu, P. & Graham, T. R. Phospholipid flippases in membrane remodeling and transport carrier biogenesis. Curr. Opin. Cell Biol. 59, 8–15 (2019).
pubmed: 30897446
doi: 10.1016/j.ceb.2019.02.004
pmcid: 6726550
Takatsu, H. et al. Phospholipid flippase activities and substrate specificities of human type IV P-type ATPases localized to the plasma membrane. J. Biol. Chem. 289, 33543–33556 (2014).
pubmed: 25315773
doi: 10.1074/jbc.M114.593012
Naito, T. et al. Phospholipid flippase ATP10A translocates phosphatidylcholine and is involved in plasma membrane dynamics. J. Biol. Chem. 290, 15004–15017 (2015).
pubmed: 25947375
doi: 10.1074/jbc.M115.655191
pmcid: 4463445
Miyata, Y., Yamada, K., Nagata, S. & Segawa, K. Two types of type IV P-type ATPases independently re-establish the asymmetrical distribution of phosphatidylserine in plasma membranes. J. Biol. Chem. 298, 102527 (2022).
pubmed: 36162506
doi: 10.1016/j.jbc.2022.102527
pmcid: 9597894
Segawa, K. et al. Phospholipid flippases enable precursor B cells to flee engulfment by macrophages. Proc. Natl Acad. Sci. USA 115, 12212–12217 (2018).
pubmed: 30355768
doi: 10.1073/pnas.1814323115
pmcid: 6275493
Kornberg, R. D. & McConnell, H. M. Inside–outside transitions of phospholipids in vesicle membranes. Biochemistry 10, 1111–1120 (1971). Using the vesicular membrane of a lipid bilayer, this article describes that the lateral movement of phospholipids in the layer is fast, whereas the inside-out transition is prolonged.
pubmed: 4324203
doi: 10.1021/bi00783a003
Bevers, E. M., Comfurius, P. & Zwaal, R. F. Changes in membrane phospholipid distribution during platelet activation. Biochim. Biophys. Acta 736, 57–66 (1983).
pubmed: 6418205
doi: 10.1016/0005-2736(83)90169-4
Miyanishi, M. et al. Identification of Tim4 as a phosphatidylserine receptor. Nature 450, 435–439 (2007).
pubmed: 17960135
doi: 10.1038/nature06307
Fadok, V. A. et al. Exposure of phosphatidylserine on the surface of apoptotic lymphocytes triggers specific recognition and removal by macrophages. J. Immunol. 148, 2207–2216 (1992). This article reports that PtdSer is exposed to the surface of apoptotic cells and is recognized by macrophages for engulfment.
pubmed: 1545126
doi: 10.4049/jimmunol.148.7.2207
Basse, F., Stout, J. G., Sims, P. J. & Wiedmer, T. Isolation of an erythrocyte membrane protein that mediates Ca
pubmed: 8663431
doi: 10.1074/jbc.271.29.17205
Bevers, E. M. & Williamson, P. L. Phospholipid scramblase: an update. FEBS Lett. 584, 2724–2730 (2010).
pubmed: 20302864
doi: 10.1016/j.febslet.2010.03.020
Suzuki, J., Umeda, M., Sims, P. J. & Nagata, S. Calcium-dependent phospholipid scrambling by TMEM16F. Nature 468, 834–838 (2010). This article reports that TMEM16F works as a Ca
pubmed: 21107324
doi: 10.1038/nature09583
Suzuki, J., Denning, D. P., Imanishi, E., Horvitz, H. R. & Nagata, S. Xk-related protein 8 and CED-8 promote phosphatidylserine exposure in apoptotic cells. Science 341, 403–406 (2013). This article reports that XKR8 at plasma membranes is cleaved at the C terminus to work as a scramblase, and the apoptotic cells require XKR8 to expose PtdSer to the cell surface.
pubmed: 23845944
doi: 10.1126/science.1236758
Schreiber, R. et al. Expression and function of epithelial anoctamins. J. Biol. Chem. 285, 7838–7845 (2010).
pubmed: 20056604
doi: 10.1074/jbc.M109.065367
pmcid: 2844227
Yang, H. et al. TMEM16F forms a Ca
pubmed: 23021219
doi: 10.1016/j.cell.2012.07.036
pmcid: 3582364
Almaça, J. et al. TMEM16 proteins produce volume-regulated chloride currents that are reduced in mice lacking TMEM16A. J. Biol. Chem. 284, 28571–28578 (2009).
pubmed: 19654323
doi: 10.1074/jbc.M109.010074
pmcid: 2781400
Martins, J. R. et al. Anoctamin 6 is an essential component of the outwardly rectifying chloride channel. Proc. Natl Acad. Sci. USA 108, 18168–18172 (2011).
pubmed: 22006324
doi: 10.1073/pnas.1108094108
pmcid: 3207678
Suzuki, J. et al. Calcium-dependent phospholipid scramblase activity of TMEM16 protein family members. J. Biol. Chem. 288, 13305–13316 (2013).
pubmed: 23532839
doi: 10.1074/jbc.M113.457937
pmcid: 3650369
Duran, C., Qu, Z., Osunkoya, A. O., Cui, Y. & Hartzell, H. C. ANOs 3–7 in the anoctamin/Tmem16 Cl
pubmed: 22075693
doi: 10.1152/ajpcell.00140.2011
Yu, K. et al. Identification of a lipid scrambling domain in ANO6/TMEM16F. eLife 4, e06901 (2015).
pubmed: 26057829
doi: 10.7554/eLife.06901
pmcid: 4477620
Scudieri, P. et al. Ion channel and lipid scramblase activity associated with expression of TMEM16F/ANO6 isoforms. J. Physiol. 593, 3829–3848 (2015).
pubmed: 26108457
doi: 10.1113/JP270691
pmcid: 4575572
Gyobu, S., Ishihara, K., Suzuki, J., Segawa, K. & Nagata, S. Characterization of the scrambling domain of the TMEM16 family. Proc. Natl Acad. Sci. USA 114, 6274–6279 (2017). This article reports that among ten members of the mouse TMEM16 family, seven members, including TMEM16E and TMEM16K, present at the ER have the potential to scramble phospholipids.
pubmed: 28559311
doi: 10.1073/pnas.1703391114
pmcid: 5474828
Alvadia, C. et al. Cryo-EM structures and functional characterization of the murine lipid scramblase TMEM16F. eLife 8, e44365 (2019). This article reports the tertiary structure of mouse TMEM16F with or without Ca
pubmed: 30785399
doi: 10.7554/eLife.44365
pmcid: 6414204
Feng, S. et al. Cryo-EM studies of TMEM16F calcium-activated ion channel suggest features important for lipid scrambling. Cell Rep. 28, 567–579 (2019).
pubmed: 31291589
doi: 10.1016/j.celrep.2019.06.023
pmcid: 6684876
Bushell, S. R. et al. The structural basis of lipid scrambling and inactivation in the endoplasmic reticulum scramblase TMEM16K. Nat. Commun. 10, 3956 (2019).
pubmed: 31477691
doi: 10.1038/s41467-019-11753-1
pmcid: 6718402
Watanabe, R., Sakuragi, T., Noji, H. & Nagata, S. Single-molecule analysis of phospholipid scrambling by TMEM16F. Proc. Natl Acad. Sci. USA 115, 3066–3071 (2018). In this work, a single molecule of TMEM16 dimer is integrated into the lipid bilayers in which phospholipids are asymmetrically distributed between the layers. The molecule scrambles phospholipids in response to Ca
pubmed: 29507235
doi: 10.1073/pnas.1717956115
pmcid: 5866571
Veshaguri, S. et al. Direct observation of proton pumping by a eukaryotic P-type ATPase. Science 351, 1469–1473 (2016).
pubmed: 27013734
doi: 10.1126/science.aad6429
pmcid: 5023152
Gyobu, S. et al. A role of TMEM16E carrying a scrambling domain in sperm motility. Mol. Cell Biol. 36, 645–659 (2016).
pubmed: 26667038
doi: 10.1128/MCB.00919-15
pmcid: 4751691
Marsault, R., Murgia, M., Pozzan, T. & Rizzuto, R. Domains of high Ca
pubmed: 9130702
doi: 10.1093/emboj/16.7.1575
pmcid: 1169761
Zayzafoon, M. Calcium/calmodulin signaling controls osteoblast growth and differentiation. J. Cell. Biochem. 97, 56–70 (2006).
pubmed: 16229015
doi: 10.1002/jcb.20675
Ehlen, H. W. et al. Inactivation of Anoctamin-6/Tmem16f, a regulator of phosphatidylserine scrambling in osteoblasts, leads to decreased mineral deposition in skeletal tissues. J. Bone Miner. Res. 28, 246–259 (2013).
pubmed: 22936354
doi: 10.1002/jbmr.1751
Fujii, T., Sakata, A., Nishimura, S., Eto, K. & Nagata, S. TMEM16F is required for phosphatidylserine exposure and microvesicle release in activated mouse platelets. Proc. Natl Acad. Sci. USA 112, 12800–12805 (2015).
pubmed: 26417084
doi: 10.1073/pnas.1516594112
pmcid: 4611630
Headland, S. E. et al. Neutrophil-derived microvesicles enter cartilage and protect the joint in inflammatory arthritis. Sci. Transl. Med. 7, 315ra190 (2015).
pubmed: 26606969
doi: 10.1126/scitranslmed.aac5608
pmcid: 6034622
Yang, X. et al. Bacterial endotoxin activates the coagulation cascade through Gasdermin D-dependent phosphatidylserine exposure. Immunity 51, 983–996 (2019).
pubmed: 31836429
doi: 10.1016/j.immuni.2019.11.005
Wu, N. et al. Critical role of lipid scramblase TMEM16F in phosphatidylserine exposure and repair of plasma membrane after pore formation. Cell Rep. 30, 1129–1140.e5 (2020).
pubmed: 31995754
doi: 10.1016/j.celrep.2019.12.066
pmcid: 7104872
Deisl, C., Hilgemann, D. W., Syeda, R. & Fine, M. TMEM16F and dynamins control expansive plasma membrane reservoirs. Nat. Commun. 12, 4990 (2021).
pubmed: 34404808
doi: 10.1038/s41467-021-25286-z
pmcid: 8371123
Gerke, V., Creutz, C. E. & Moss, S. E. Annexins: linking Ca
pubmed: 15928709
doi: 10.1038/nrm1661
Foltz, S. J., Cui, Y. Y., Choo, H. J. & Hartzell, H. C. ANO5 ensures trafficking of annexins in wounded myofibers. J. Cell Biol. 220, e202007059 (2021).
pubmed: 33496727
doi: 10.1083/jcb.202007059
pmcid: 7844426
Chandra, G. et al. Dysregulated calcium homeostasis prevents plasma membrane repair in Anoctamin 5/TMEM16E-deficient patient muscle cells. Cell Death Discov. 5, 118 (2019).
pubmed: 31341644
doi: 10.1038/s41420-019-0197-z
pmcid: 6639303
Griffin, D. A. et al. Defective membrane fusion and repair in Anoctamin5-deficient muscular dystrophy. Hum. Mol. Genet. 25, 1900–1911 (2016).
pubmed: 26911675
doi: 10.1093/hmg/ddw063
pmcid: 5062581
Petkovic, M., Oses-Prieto, J., Burlingame, A., Jan, L. Y. & Jan, Y. N. TMEM16K is an interorganelle regulator of endosomal sorting. Nat. Commun. 11, 3298 (2020).
pubmed: 32620747
doi: 10.1038/s41467-020-17016-8
pmcid: 7335067
Suzuki, J., Imanishi, E. & Nagata, S. Exposure of phosphatidylserine by Xk-related protein family members during apoptosis. J. Biol. Chem. 289, 30257–30267 (2014).
pubmed: 25231987
doi: 10.1074/jbc.M114.583419
pmcid: 4215210
Williamson, P. et al. Phospholipid scramblase activation pathways in lymphocytes. Biochemistry 40, 8065–8072 (2001).
pubmed: 11434775
doi: 10.1021/bi001929z
Schoenwaelder, S. M. et al. Two distinct pathways regulate platelet phosphatidylserine exposure and procoagulant function. Blood 114, 663–666 (2009).
pubmed: 19387006
doi: 10.1182/blood-2009-01-200345
Pervaiz, N. et al. Evolutionary history of the human multigene families reveals widespread gene duplications throughout the history of animals. BMC Evol. Biol. 19, 128 (2019).
pubmed: 31221090
doi: 10.1186/s12862-019-1441-0
pmcid: 6585022
Kawano, M. & Nagata, S. Lupus-like autoimmune disease caused by a lack of Xkr8, a caspase-dependent phospholipid scramblase. Proc. Natl Acad. Sci. USA 280, 2132–2137 (2018).
doi: 10.1073/pnas.1720732115
Suzuki, J., Imanishi, E. & Nagata, S. Xkr8 phospholipid scrambling complex in apoptotic phosphatidylserine exposure. Proc. Natl Acad. Sci. USA 113, 9509–9514 (2016).
pubmed: 27503893
doi: 10.1073/pnas.1610403113
pmcid: 5003272
Sakuragi, T., Kosako, H. & Nagata, S. Phosphorylation-mediated activation of mouse Xkr8 scramblase for phosphatidylserine exposure. Proc. Natl Acad. Sci. USA 33, 2907–2912 (2019).
doi: 10.1073/pnas.1820499116
Gadella, B. M. & Harrison, R. A. The capacitating agent bicarbonate induces protein kinase A-dependent changes in phospholipid transbilayer behavior in the sperm plasma membrane. Development 127, 2407–2420 (2000).
pubmed: 10804182
doi: 10.1242/dev.127.11.2407
Di Virgilio, F., Sarti, A. C., Falzoni, S., Marchi, E. D. & Adinolfi, E. Extracellular ATP and P2 purinergic signalling in the tumour microenvironment. Nat. Rev. Cancer 18, 601–618 (2018).
pubmed: 30006588
doi: 10.1038/s41568-018-0037-0
Kamata-Sakurai, M. et al. Antibody to CD137 activated by extracellular adenosine triphosphate is tumor selective and broadly effective in vivo without systemic immune activation. Cancer Discov. 11, 158–175 (2021).
pubmed: 32847940
doi: 10.1158/2159-8290.CD-20-0328
Ryoden, Y., Segawa, K. & Nagata, S. Requirement of Xk and Vps13a for the P2X7-mediated phospholipid scrambling and cell lysis in mouse T cells. Proc. Natl Acad. Sci. USA 119, e2119286119 (2022). This article reports that XK, a paralogue of XKR8, is complexed with VPS13A lipid transporter, and works as a scramblase in response to an unidentified signal from the ATP-engaged P2X7 receptor.
pubmed: 35140185
doi: 10.1073/pnas.2119286119
pmcid: 8851519
Kumar, N. et al. VPS13A and VPS13C are lipid transport proteins differentially localized at ER contact sites. J. Cell Biol. 217, 3625–3639 (2018).
pubmed: 30093493
doi: 10.1083/jcb.201807019
pmcid: 6168267
Puts, C. F. & Holthuis, J. C. Mechanism and significance of P4 ATPase-catalyzed lipid transport: lessons from a Na
pubmed: 19233312
doi: 10.1016/j.bbalip.2009.02.005
Pomorski, T. & Menon, A. K. Lipid flippases and their biological functions. Cell. Mol. Life Sci. 63, 2908–2921 (2006). This article proposes the credit card model for phospholipids to transport through the lipid bilayer, in which the hydrophilic head group of phospholipids passes the hydrophilic crevasse in the molecule.
pubmed: 17103115
doi: 10.1007/s00018-006-6167-7
Hiraizumi, M., Yamashita, K., Nishizawa, T. & Nureki, O. Cryo-EM structures capture the transport cycle of the P4-ATPase flippase. Science 365, 1149–1155 (2019). This article reports the tertiary structure of the human ATP8A1/CDC50A hetero complex in six intermediate conformations. The structure agrees with the credit card model proposed by Pomorski and Mennon (2006).
pubmed: 31416931
doi: 10.1126/science.aay3353
Kalienkova, V., Mosina, V. C. & Paulino, C. The groovy TMEM16 family: molecular mechanisms of lipid scrambling and ion conduction. J. Mol. Biol. 433, 166941 (2021).
pubmed: 33741412
doi: 10.1016/j.jmb.2021.166941
Falzone, M. E. et al. TMEM16 scramblases thin the membrane to enable lipid scrambling. Nat. Commun. 13, 2604 (2022).
pubmed: 35562175
doi: 10.1038/s41467-022-30300-z
pmcid: 9095706
Timcenko, M. et al. Structure and autoregulation of a P4-ATPase lipid flippase. Nature 571, 366–370 (2019). This article reports the first tertiary structure of P4-ATPase with yeast P4-ATPase and proposes a pathway for lipid transport.
pubmed: 31243363
doi: 10.1038/s41586-019-1344-7
Nakanishi, H. et al. Crystal structure of a human plasma membrane phospholipid flippase. J. Biol. Chem. 295, 10180–10194 (2020).
pubmed: 32493773
doi: 10.1074/jbc.RA120.014144
pmcid: 7383378
Nakanishi, H. et al. Transport cycle of plasma membrane flippase ATP11C by cryo-EM. Cell Rep. 32, 108208 (2020).
pubmed: 32997992
doi: 10.1016/j.celrep.2020.108208
Bai, L. et al. Autoinhibition and activation mechanisms of the eukaryotic lipid flippase Drs2p–Cdc50p. Nat. Commun. 10, 4142 (2019).
pubmed: 31515475
doi: 10.1038/s41467-019-12191-9
pmcid: 6742660
Timcenko, M. et al. Structural basis of substrate-independent phosphorylation in a P4-ATPase lipid flippase. J. Mol. Biol. 433, 167062 (2021).
pubmed: 34023399
doi: 10.1016/j.jmb.2021.167062
He, Y., Xu, J., Wu, X. & Li, L. Structures of a P4-ATPase lipid flippase in lipid bilayers. Protein Cell 11, 458–463 (2020).
pubmed: 32303992
doi: 10.1007/s13238-020-00712-y
pmcid: 7251018
Bai, L. et al. Transport mechanism of P4 ATPase phosphatidylcholine flippases. eLife 9, e62163 (2020).
pubmed: 33320091
doi: 10.7554/eLife.62163
pmcid: 7773333
Post, R. L., Hegyvary, C. & Kume, S. Activation by adenosine triphosphate in the phosphorylation kinetics of sodium and potassium ion transport adenosine triphosphatase. J. Biol. Chem. 247, 6530–6540 (1972).
pubmed: 4263199
doi: 10.1016/S0021-9258(19)44725-X
Albers, R. W. Biochemical aspects of active transport. Annu. Rev. Biochem. 36, 727–756 (1967).
pubmed: 18257736
doi: 10.1146/annurev.bi.36.070167.003455
Vestergaard, A. L. et al. Critical roles of isoleucine-364 and adjacent residues in a hydrophobic gate control of phospholipid transport by the mammalian P4-ATPase ATP8A2. Proc. Natl Acad. Sci. USA 111, E1334–E1343 (2014).
pubmed: 24706822
doi: 10.1073/pnas.1321165111
pmcid: 3986137
Bai, L. et al. Structural basis of the P4B ATPase lipid flippase activity. Nat. Commun. 12, 5963 (2021).
pubmed: 34645814
doi: 10.1038/s41467-021-26273-0
pmcid: 8514546
Segawa, K. et al. A sublethal ATP11A mutation associated with neurological deterioration causes aberrant phosphatidylcholine flipping in plasma membranes. J. Clin. Invest. 131, e148005 (2021). This article reports that a patient suffering neurological disorder carries a de novo dominant point mutation in ATP11A which causes flipping PtdCho in addition to PtdSer.
pubmed: 34403372
doi: 10.1172/JCI148005
pmcid: 8439608
Baldridge, R. D. & Graham, T. R. Two-gate mechanism for phospholipid selection and transport by type IV P-type ATPases. Proc. Natl Acad. Sci. USA 110, E358–E367 (2013).
pubmed: 23302692
doi: 10.1073/pnas.1216948110
pmcid: 3562821
Brunner, J. D., Lim, N. K., Schenck, S., Duerst, A. & Dutzler, R. X-ray structure of a calcium-activated TMEM16 lipid scramblase. Nature 516, 207–212 (2014). This article reports the first tertiary structure of TMEM16 homodimer with fungus TMEM16. The structure has a rhomboidal architecture with a trans-bilayer hydrophilic groove at the periphery.
pubmed: 25383531
doi: 10.1038/nature13984
Kalienkova, V. et al. Stepwise activation mechanism of the scramblase nhTMEM16 revealed by cryo-EM. eLife 8, e44364 (2019).
pubmed: 30785398
doi: 10.7554/eLife.44364
pmcid: 6414200
Falzone, M. E. et al. Structural basis of Ca
pubmed: 30648972
doi: 10.7554/eLife.43229
pmcid: 6355197
Le, T. et al. An inner activation gate controls TMEM16F phospholipid scrambling. Nat. Commun. 10, 1846 (2019).
pubmed: 31015464
doi: 10.1038/s41467-019-09778-7
pmcid: 6478717
Lee, B.-C. et al. Gating mechanism of the extracellular entry to the lipid pathway in a TMEM16 scramblase. Nat. Commun. 9, 3251 (2018).
pubmed: 30108217
doi: 10.1038/s41467-018-05724-1
pmcid: 6092359
Khelashvili, G. et al. Membrane lipids are both the substrates and a mechanistically responsive environment of TMEM16 scramblase proteins. J. Comput. Chem. 41, 538–551 (2020).
pubmed: 31750558
doi: 10.1002/jcc.26105
Ishihara, K., Suzuki, J. & Nagata, S. Role of Ca
pubmed: 27227820
doi: 10.1021/acs.biochem.6b00176
Bethel, N. P. & Grabe, M. Atomistic insight into lipid translocation by a TMEM16 scramblase. Proc. Natl Acad. Sci. USA 113, 14049–14054 (2016). This article reports the molecular dynamics simulation analysis of TMEM16, and proposes that phospholipids move through the groove of the protein using hydrophilic residues as ‘stepping stones’.
pubmed: 27872308
doi: 10.1073/pnas.1607574113
pmcid: 5150362
Jiang, T., Yu, K., Hartzell, H. C. & Tajkhorshid, E. Lipids and ions traverse the membrane by the same physical pathway in the nhTMEM16 scramblase. eLife 6, e28671 (2017).
pubmed: 28917060
doi: 10.7554/eLife.28671
pmcid: 5628016
Malvezzi, M. et al. Out-of-the-groove transport of lipids by TMEM16 and GPCR scramblases. Proc. Natl Acad. Sci. USA 115, E7033–E7042 (2018).
pubmed: 29925604
doi: 10.1073/pnas.1806721115
pmcid: 6065010
Khelashvili, G., Kots, E., Cheng, X., Levine, M. V. & Weinstein, H. The allosteric mechanism leading to an open-groove lipid conductive state of the TMEM16F scramblase. Commun. Biol. 5, 990 (2022).
pubmed: 36123525
doi: 10.1038/s42003-022-03930-8
pmcid: 9484709
Jojoa-Cruz, S. et al. Cryo-EM structure of the mechanically activated ion channel OSCA1.2. eLife 7, e41845 (2018).
pubmed: 30382939
doi: 10.7554/eLife.41845
pmcid: 6235563
Liu, X., Wang, J. & Sun, L. Structure of the hyperosmolality-gated calcium-permeable channel OSCA1.2. Nat. Commun. 9, 5060 (2018).
pubmed: 30498218
doi: 10.1038/s41467-018-07564-5
pmcid: 6265326
Maity, K. et al. Cryo-EM structure of OSCA1.2 from Oryza sativa elucidates the mechanical basis of potential membrane hyperosmolality gating. Proc. Natl Acad. Sci. USA 116, 14309–14318 (2019).
pubmed: 31227607
doi: 10.1073/pnas.1900774116
pmcid: 6628804
Ballesteros, A., Fenollar-Ferrer, C. & Swartz, K. J. Structural relationship between the putative hair cell mechanotransduction channel TMC1 and TMEM16 proteins. eLife 7, e38433 (2018).
pubmed: 30063209
doi: 10.7554/eLife.38433
pmcid: 6067890
Pan, B. et al. TMC1 forms the pore of mechanosensory transduction channels in vertebrate inner ear hair cells. Neuron 99, 736–753 (2018).
pubmed: 30138589
doi: 10.1016/j.neuron.2018.07.033
pmcid: 6360533
Jeong, H. et al. Structures of the TMC-1 complex illuminate mechanosensory transduction. Nature 610, 796–803 (2022).
pubmed: 36224384
doi: 10.1038/s41586-022-05314-8
pmcid: 9605866
Ballesteros, A. & Swartz, K. J. Regulation of membrane homeostasis by TMC1 mechanoelectrical transduction channels is essential for hearing. Sci. Adv. 8, eabm5550 (2022).
pubmed: 35921424
doi: 10.1126/sciadv.abm5550
pmcid: 9348795
Sakuragi, T. et al. The tertiary structure of the human Xkr8–Basigin complex that scrambles phospholipids at plasma membranes. Nat. Struct. Mol. Biol. 28, 825–834 (2021). This article reports the tertiary structure of the human XKR8–Basigin complex and proposes a phospholipid entry site and hydrophilic path for scrambling phospholipids.
pubmed: 34625749
doi: 10.1038/s41594-021-00665-8
pmcid: 8500837
Straub, M. S., Alvadia, C., Sawicka, M. & Dutzler, R. Cryo-EM structures of the caspase activated protein XKR9 involved in apoptotic lipid scrambling. eLife 10, e69800 (2021).
pubmed: 34263724
doi: 10.7554/eLife.69800
pmcid: 8298096
Jung, H. H., et al. McLeod neuroacanthocytosis syndrome. National Library of Medicine https://www.ncbi.nlm.nih.gov/books/NBK1354/ (2021).
Park, J.-S., Hu, Y., Hollingsworth, N. M., Miltenberger-Miltenyi, G. & Neiman, A. M. Interaction between VPS13A and the XK scramblase is important for VPS13A function in humans. J. Cell Sci. 135, jcs260227 (2022).
pubmed: 35950506
doi: 10.1242/jcs.260227
Guillén-Samander, A. et al. A partnership between the lipid scramblase XK and the lipid transfer protein VPS13A at the plasma membrane. Proc. Natl Acad. Sci. USA 119, e2205425119 (2022).
pubmed: 35994651
doi: 10.1073/pnas.2205425119
pmcid: 9436381
Segawa, K. & Nagata, S. An apoptotic ‘eat me’ signal: phosphatidylserine exposure. Trends Cell Biol. 25, 649–650 (2015).
doi: 10.1016/j.tcb.2015.08.003
Whitlock, J. M. & Chernomordik, L. V. Flagging fusion: phosphatidylserine signaling in cell–cell fusion. J. Biol. Chem. 296, 100411 (2021).
pubmed: 33581114
doi: 10.1016/j.jbc.2021.100411
pmcid: 8005811
Wood, W. et al. Mesenchymal cells engulf and clear apoptotic footplate cells in macrophageless PU.1 null mouse embryos. Development 127, 5245–5252 (2000).
pubmed: 11076747
doi: 10.1242/dev.127.24.5245
Nagasaka, A., Kawane, K., Yoshida, H. & Nagata, S. Apaf-1-independent programmed cell death in mouse development. Cell Death Differ. 17, 931–941 (2010).
pubmed: 19960021
doi: 10.1038/cdd.2009.186
Nagata, S., Hanayama, R. & Kawane, K. Autoimmunity and the clearance of dead cells. Cell 140, 619–630 (2010).
pubmed: 20211132
doi: 10.1016/j.cell.2010.02.014
deCathelineau, A. M. & Henson, P. M. The final step in programmed cell death: phagocytes carry apoptotic cells to the grave. Essays Biochem. 39, 105–117 (2003).
pubmed: 14585077
doi: 10.1042/bse0390105
Muñoz, L. E., Lauber, K., Schiller, M., Manfredi, A. A. & Herrmann, M. The role of defective clearance of apoptotic cells in systemic autoimmunity. Nat. Rev. Rheumatol. 6, 280–289 (2010).
pubmed: 20431553
doi: 10.1038/nrrheum.2010.46
Surh, C. D. & Sprent, J. T-cell apoptosis detected in situ during positive and negative selection in the thymus. Nature 372, 100–103 (1994).
pubmed: 7969401
doi: 10.1038/372100a0
Ren, Y. & Savill, J. Apoptosis: the importance of being eaten. Cell Death Differ. 5, 563–568 (1998).
pubmed: 10200510
doi: 10.1038/sj.cdd.4400407
Segawa, K., Suzuki, J. & Nagata, S. Constitutive exposure of phosphatidylserine on viable cells. Proc. Natl Acad. Sci. USA 108, 19246–19251 (2011).
pubmed: 22084121
doi: 10.1073/pnas.1114799108
pmcid: 3228483
Davies, L. C., Jenkins, S. J., Allen, J. E. & Taylor, P. R. Tissue-resident macrophages. Nat. Immunol. 14, 986–995 (2013).
pubmed: 24048120
doi: 10.1038/ni.2705
pmcid: 4045180
Yanagihashi, Y., Segawa, K., Maeda, R., Nabeshima, Y.-I. & Nagata, S. Mouse macrophages show different requirements for phosphatidylserine receptor Tim4 in efferocytosis. Proc. Natl Acad. Sci. USA 114, 8800–8805 (2017).
pubmed: 28768810
doi: 10.1073/pnas.1705365114
pmcid: 5565444
Lemke, G. How macrophages deal with death. Nat. Rev. Immunol. 36, 1–11 (2019).
Nishi, C., Toda, S., Segawa, K. & Nagata, S. Tim4- and MerTK-mediated engulfment of apoptotic cells by mouse resident peritoneal macrophages. Mol. Cell. Biol. 34, 1512–1520 (2014).
pubmed: 24515440
doi: 10.1128/MCB.01394-13
pmcid: 3993587
Hanayama, R. et al. Identification of a factor that links apoptotic cells to phagocytes. Nature 417, 182–187 (2002).
pubmed: 12000961
doi: 10.1038/417182a
Hanayama, R., Tanaka, M., Miwa, K. & Nagata, S. Expression of developmental endothelial locus-1 in a subset of macrophages for engulfment of apoptotic cells. J. Immunol. 172, 3876–3882 (2004).
pubmed: 15004195
doi: 10.4049/jimmunol.172.6.3876
Kourtzelis, I. et al. DEL-1 promotes macrophage efferocytosis and clearance of inflammation. Nat. Immunol. 20, 40–49 (2019).
pubmed: 30455459
doi: 10.1038/s41590-018-0249-1
Proto, J. D. et al. Regulatory T cells promote macrophage efferocytosis during inflammation resolution. Immunity 49, 666–677 (2018).
pubmed: 30291029
doi: 10.1016/j.immuni.2018.07.015
Park, D. et al. BAI1 is an engulfment receptor for apoptotic cells upstream of the ELMO/Dock180/Rac module. Nature 450, 430–434 (2007).
pubmed: 17960134
doi: 10.1038/nature06329
Park, S. Y. et al. Rapid cell corpse clearance by stabilin-2, a membrane phosphatidylserine receptor. Cell Death Differ. 15, 192–201 (2008).
pubmed: 17962816
doi: 10.1038/sj.cdd.4402242
Das, S. et al. Brain angiogenesis inhibitor 1 (BAI1) is a pattern recognition receptor that mediates macrophage binding and engulfment of Gram-negative bacteria. Proc. Natl Acad. Sci. USA 108, 2136–2141 (2011).
pubmed: 21245295
doi: 10.1073/pnas.1014775108
pmcid: 3033312
Harris, E. N. & Cabral, F. Ligand binding and signaling of HARE/Stabilin-2. Biomolecules 9, 273 (2019).
pubmed: 31336723
doi: 10.3390/biom9070273
pmcid: 6681266
Galluzzi, L. et al. Molecular mechanisms of cell death: recommendations of the Nomenclature Committee on Cell Death 2018. Cell Death Differ. 25, 486–541 (2018).
pubmed: 29362479
doi: 10.1038/s41418-017-0012-4
pmcid: 5864239
Shan, B., Pan, H., Najafov, A. & Yuan, J. Necroptosis in development and diseases. Genes Dev. 32, 327–340 (2018).
pubmed: 29593066
doi: 10.1101/gad.312561.118
pmcid: 5900707
Gong, Y.-N. et al. ESCRT-III acts downstream of MLKL to regulate necroptotic cell death and its consequences. Cell 169, 286–300 (2017).
pubmed: 28388412
doi: 10.1016/j.cell.2017.03.020
pmcid: 5443414
Rühl, S. et al. ESCRT-dependent membrane repair negatively regulates pyroptosis downstream of GSDMD activation. Science 362, 956–960 (2018).
pubmed: 30467171
doi: 10.1126/science.aar7607
Di Virgilio, F., Ben, D. D., Sarti, A. C., Giuliani, A. L. & Falzoni, S. The P2X7 receptor in infection and inflammation. Immunity 47, 15–31 (2017).
pubmed: 28723547
doi: 10.1016/j.immuni.2017.06.020
Aswad, F., Kawamura, H. & Dennert, G. High sensitivity of CD4
pubmed: 16116196
doi: 10.4049/jimmunol.175.5.3075
Ryoden, Y. & Nagata, S. The XK plasma membrane scramblase and the VPS13A cytosolic lipid transporter for ATP-induced cell death. BioEssays 44, e2200106 (2022).
pubmed: 35996795
doi: 10.1002/bies.202200106
Thiagarajan, P., Parker, C. J. & Prchal, J. T. How do red blood cells die? Front. Physiol. 12, 655393 (2021).
pubmed: 33790808
doi: 10.3389/fphys.2021.655393
pmcid: 8006275
Yoshida, H. et al. Phosphatidylserine-dependent engulfment by macrophages of nuclei from erythroid precursor cells. Nature 437, 754–758 (2005). At the final stage of definitive erythropoiesis, pyrenocytes (nuclei surrounded by plasma membranes) are separated from reticulocytes. This article reports that the pyrenocytes expose PtdSer for engulfment by macrophages soon after the division from reticulocytes.
pubmed: 16193055
doi: 10.1038/nature03964
Toda, S., Segawa, K. & Nagata, S. MerTK-mediated engulfment of pyrenocytes by central macrophages in erythroblastic islands. Blood 123, 3963–3971 (2014).
pubmed: 24659633
doi: 10.1182/blood-2014-01-547976
Ball, J. B., Green-Fulgham, S. M. & Watkins, L. R. Mechanisms of microglia-mediated synapse turnover and synaptogenesis. Prog. Neurobiol. 218, 102336 (2022).
pubmed: 35940391
doi: 10.1016/j.pneurobio.2022.102336
Frost, J. L. & Schafer, D. P. Microglia: architects of the developing nervous system. Trends Cell Biol. 26, 587–597 (2016).
pubmed: 27004698
doi: 10.1016/j.tcb.2016.02.006
pmcid: 4961529
Li, T. et al. A splicing isoform of GPR56 mediates microglial synaptic refinement via phosphatidylserine binding. EMBO J. 39, e104136 (2020).
pubmed: 32452062
doi: 10.15252/embj.2019104136
pmcid: 7429740
Scott-Hewitt, N. et al. Local externalization of phosphatidylserine mediates developmental synaptic pruning by microglia. EMBO J. 39, e105380 (2020). Microglia eliminate supernumerary synapses generated during the development of the nervous network, which is called pruning. This article reports that the synapses to be eliminated expose PtdSer as a neuronal ‘eat me’ signal.
pubmed: 32657463
doi: 10.15252/embj.2020105380
pmcid: 7429741
Kurematsu, C. et al. Synaptic pruning of murine adult-born neurons by microglia depends on phosphatidylserine. J. Exp. Med. 219, e20202304 (2022).
pubmed: 35297954
doi: 10.1084/jem.20202304
pmcid: 9195048
Sapar, M. L. et al. Phosphatidylserine externalization results from and causes neurite degeneration in Drosophila. Cell Rep. 24, 2273–2286 (2018).
pubmed: 30157423
doi: 10.1016/j.celrep.2018.07.095
pmcid: 6174084
Pereira, M. et al. Common signalling pathways in macrophage and osteoclast multinucleation. J. Cell. Sci. 131, jcs216267 (2018).
pubmed: 29871956
doi: 10.1242/jcs.216267
Petrany, M. J. & Millay, D. P. Cell fusion: merging membranes and making muscle. Trends Cell Biol. 29, 964–973 (2019).
pubmed: 31648852
doi: 10.1016/j.tcb.2019.09.002
pmcid: 7849503
Gerbaud, P. & Pidoux, G. An overview of molecular events occurring in human trophoblast fusion. Placenta 36, S35–S42 (2015).
pubmed: 25564303
doi: 10.1016/j.placenta.2014.12.015
Das, M. et al. Phosphatidylserine efflux and intercellular fusion in a BeWo model of human villous cytotrophoblast. Placenta 25, 396–407 (2004).
pubmed: 15081634
doi: 10.1016/j.placenta.2003.11.004
Satouh, Y. & Ikawa, M. New insights into the molecular events of mammalian fertilization. Trends Biochem. Sci. 43, 818–828 (2018).
pubmed: 30170889
doi: 10.1016/j.tibs.2018.08.006
pmcid: 6162164
Helming, L., Winter, J. & Gordon, S. The scavenger receptor CD36 plays a role in cytokine-induced macrophage fusion. J. Cell Sci. 122, 453–459 (2009).
pubmed: 19155290
doi: 10.1242/jcs.037200
pmcid: 2714432
Verma, S. K. et al. Cell-surface phosphatidylserine regulates osteoclast precursor fusion. J. Biol. Chem. 293, 254–270 (2018).
pubmed: 29101233
doi: 10.1074/jbc.M117.809681
Jeong, J. & Conboy, I. M. Phosphatidylserine directly and positively regulates fusion of myoblasts into myotubes. Biochem. Biophys. Res. Commun. 414, 9–13 (2011).
pubmed: 21910971
doi: 10.1016/j.bbrc.2011.08.128
pmcid: 3195849
Rival, C. M. et al. Phosphatidylserine on viable sperm and phagocytic machinery in oocytes regulate mammalian fertilization. Nat. Commun. 10, 4456 (2019).
pubmed: 31575859
doi: 10.1038/s41467-019-12406-z
pmcid: 6773685
Martens, S. & McMahon, H. T. Mechanisms of membrane fusion: disparate players and common principles. Nat. Rev. Mol. Cell Biol. 9, 543–556 (2008).
pubmed: 18496517
doi: 10.1038/nrm2417
Deneke, V. E. & Pauli, A. The fertilization enigma: how sperm and egg fuse. Annu. Rev. Cell Dev. Biol. 37, 391–414 (2021).
pubmed: 34288709
doi: 10.1146/annurev-cellbio-120219-021751
Szondy, Z. et al. Involvement of phosphatidylserine receptors in the skeletal muscle regeneration: therapeutic implications. J. Cachexia Sarcopenia Muscle 13, 1961–1973 (2022).
pubmed: 35666022
doi: 10.1002/jcsm.13024
pmcid: 9397555
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).
pubmed: 11707515
doi: 10.1242/jcs.114.20.3631
Tsuchiya, M. et al. Cell surface flip-flop of phosphatidylserine is critical for PIEZO1-mediated myotube formation. Nat. Commun. 9, 2049 (2018).
pubmed: 29799007
doi: 10.1038/s41467-018-04436-w
pmcid: 5967302
Grifell-Junyent, M. et al. CDC50A is required for aminophospholipid transport and cell fusion in mouse C2C12 myoblasts. J. Cell Sci. 135, jcs258649 (2022).
pubmed: 34664668
doi: 10.1242/jcs.258649
Ochiai, Y., Suzuki, C., Segawa, K., Uchiyama, Y. & Nagata, S. Inefficient development of syncytiotrophoblasts in the Atp11a-deficient mouse placenta. Proc. Natl Acad. Sci. USA 119, e2200582119 (2022). Trophoblasts in the placenta fuse to form syncytiotrophoblasts at the maternal–fetal interface. This article reports that the flippase-deficient trophoblasts fail to fuse, probably because of the constitutive exposure of PtdSer.
pubmed: 35476530
doi: 10.1073/pnas.2200582119
pmcid: 9170144
Middel, V. et al. Dysferlin-mediated phosphatidylserine sorting engages macrophages in sarcolemma repair. Nat. Commun. 7, 12875 (2016).
pubmed: 27641898
doi: 10.1038/ncomms12875
pmcid: 5031802
Croissant, C., Carmeille, R., Brévart, C. & Bouter, A. Annexins and membrane repair dysfunctions in muscular dystrophies. Int. J. Mol. Sci. 22, 5276 (2021).
pubmed: 34067866
doi: 10.3390/ijms22105276
pmcid: 8155887
Neumann, B. et al. EFF-1-mediated regenerative axonal fusion requires components of the apoptotic pathway. Nature 517, 219–222 (2015).
pubmed: 25567286
doi: 10.1038/nature14102
Hisamoto, N. et al. Phosphatidylserine exposure mediated by ABC transporter activates the integrin signaling pathway promoting axon regeneration. Nat. Commun. 9, 3099 (2018).
pubmed: 30082731
doi: 10.1038/s41467-018-05478-w
pmcid: 6079064
Bevers, E. M., Comfurius, P., van Rijn, J. L., Hemker, H. C. & Zwaal, R. F. Generation of prothrombin-converting activity and the exposure of phosphatidylserine at the outer surface of platelets. Eur. J. Biochem. 122, 429–436 (1982). This article reports that the activated platelets expose PtdSer, to which blood clotting factors bind and are activated to generate thrombin for blood clotting.
pubmed: 7060583
doi: 10.1111/j.1432-1033.1982.tb05898.x
Zwaal, R. F., Comfurius, P. & Bevers, E. M. Lipid–protein interactions in blood coagulation. Biochim. Biophys. Acta 1376, 433–453 (1998).
pubmed: 9805008
doi: 10.1016/S0304-4157(98)00018-5
Peschon, J. J. et al. An essential role for ectodomain shedding in mammalian development. Science 282, 1281–1284 (1998).
pubmed: 9812885
doi: 10.1126/science.282.5392.1281
Sommer, A. et al. Phosphatidylserine exposure is required for ADAM17 sheddase function. Nat. Commun. 7, 11523 (2016).
pubmed: 27161080
doi: 10.1038/ncomms11523
pmcid: 4866515
Elliott, J. I. et al. Membrane phosphatidylserine distribution as a non-apoptotic signalling mechanism in lymphocytes. Nat. Cell Biol. 7, 808–816 (2005).
pubmed: 16025105
doi: 10.1038/ncb1279
Kirkin, V. et al. The Fas ligand intracellular domain is released by ADAM10 and SPPL2a cleavage in T-cells. Cell Death Differ. 14, 1678–1687 (2007).
pubmed: 17557115
doi: 10.1038/sj.cdd.4402175
Schulte, M. et al. ADAM10 regulates FasL cell surface expression and modulates FasL-induced cytotoxicity and activation-induced cell death. Cell Death Differ. 14, 1040–1049 (2007).
pubmed: 17290285
doi: 10.1038/sj.cdd.4402101
Bleibaum, F. et al. ADAM10 sheddase activation is controlled by cell membrane asymmetry. J. Mol. Cell Biol. 11, 979–993 (2019).
pubmed: 30753537
doi: 10.1093/jmcb/mjz008
pmcid: 6927242
Patton, S. & Keenan, T. W. The milk fat globule membrane. Biochim. Biophys. Acta 415, 273–309 (1975).
pubmed: 1101969
doi: 10.1016/0304-4157(75)90011-8
Hanayama, R. & Nagata, S. Impaired involution of mammary glands in the absence of milk fat globule EGF factor 8. Proc. Natl Acad. Sci. USA 102, 16886–16891 (2005).
pubmed: 16275924
doi: 10.1073/pnas.0508599102
pmcid: 1277971
Théry, C., Ostrowski, M. & Segura, E. Membrane vesicles as conveyors of immune responses. Nat. Rev. Immunol. 9, 581–593 (2009).
pubmed: 19498381
doi: 10.1038/nri2567
Nakai, W. et al. A novel affinity-based method for the isolation of highly purified extracellular vesicles. Sci. Rep. 6, 33935 (2016).
pubmed: 27659060
doi: 10.1038/srep33935
pmcid: 5034288
Wei, X. et al. Surface phosphatidylserine is responsible for the internalization on microvesicles derived from hypoxia-induced human bone marrow mesenchymal stem cells into human endothelial cells. PLoS ONE 11, e0147360 (2016).
pubmed: 26808539
doi: 10.1371/journal.pone.0147360
pmcid: 4726621
Puhm, F., Boilard, E. & Machlus, K. R. Platelet extracellular vesicles. Arterioscler. Thromb. Vasc. Biol. 41, 87–96 (2020).
pubmed: 33028092
pmcid: 7769913
Sims, P., Wiedmer, T., Esmon, C., Weiss, H. & Shattil, S. Assembly of the platelet prothrombinase complex is linked to vesiculation of the platelet plasma membrane. Studies in Scott syndrome: an isolated defect in platelet procoagulant activity. J. Biol. Chem. 264, 17049–17057 (1989).
pubmed: 2793843
doi: 10.1016/S0021-9258(18)71457-9
Mercer, J. & Helenius, A. Vaccinia virus uses macropinocytosis and apoptotic mimicry to enter host cells. Science 320, 531–535 (2008). This article reports that the infection of the vaccinia virus, an enveloped virus, is promoted by PtdSer exposed on the surface of the virus particles.
pubmed: 18436786
doi: 10.1126/science.1155164
Morizono, K. & Chen, I. S. Y. Role of phosphatidylserine receptors in enveloped virus infection. J. Virol. 88, 4275–4290 (2014).
pubmed: 24478428
doi: 10.1128/JVI.03287-13
pmcid: 3993771
Li, M. et al. TIM-family proteins inhibit HIV-1 release. Proc. Natl Acad. Sci. USA 111, E3699–E3707 (2014).
pubmed: 25136083
doi: 10.1073/pnas.1404851111
pmcid: 4156686
Rood, J. E., Maartens, A., Hupalowska, A., Teichmann, S. A. & Regev, A. Impact of the Human Cell Atlas on medicine. Nat. Med. 28, 2486–2496 (2022).
pubmed: 36482102
doi: 10.1038/s41591-022-02104-7
Liou, A. Y., Molday, L. L., Wang, J., Andersen, J. P. & Molday, R. S. Identification and functional analyses of disease-associated P4-ATPase phospholipid flippase variants in red blood cells. J. Biol. Chem. 294, 6809–6821 (2019).
pubmed: 30850395
doi: 10.1074/jbc.RA118.007270
pmcid: 6497962
Siggs, O. M. et al. The P4-type ATPase ATP11C is essential for B lymphopoiesis in adult bone marrow. Nat. Immunol. 12, 434–440 (2011).
pubmed: 21423172
doi: 10.1038/ni.2012
pmcid: 3079768
Siggs, O. M., Schnabl, B., Webb, B. & Beutler, B. X-linked cholestasis in mouse due to mutations of the P4-ATPase ATP11C. Proc. Natl Acad. Sci. USA 108, 7890–7895 (2011).
pubmed: 21518881
doi: 10.1073/pnas.1104631108
pmcid: 3093471
Yabas, M. et al. Mice deficient in the putative phospholipid flippase ATP11C exhibit altered erythrocyte shape, anemia, and reduced erythrocyte life span. J. Biol. Chem. 289, 19531–19537 (2014).
pubmed: 24898253
doi: 10.1074/jbc.C114.570267
pmcid: 4094063
Yabas, M. et al. ATP11C is critical for the internalization of phosphatidylserine and differentiation of B lymphocytes. Nat. Immunol. 12, 441–449 (2011).
pubmed: 21423173
doi: 10.1038/ni.2011
pmcid: 3272780
Arashiki, N. et al. ATP11C is a major flippase in human erythrocytes and its defect causes congenital hemolytic anemia. Haematologica 101, 559–565 (2016).
pubmed: 26944472
doi: 10.3324/haematol.2016.142273
pmcid: 5004368
Brain, M. C., Pihl, C., Robertson, L. & Brown, C. B. Evidence for a mechanosensitive calcium influx into red cells. Blood Cell. Mol. Dis. 32, 349–352 (2004).
doi: 10.1016/j.bcmd.2004.01.005
Boas, F. E., Forman, L. & Beutler, E. Phosphatidylserine exposure and red cell viability in red cell aging and in hemolytic anemia. Proc. Natl Acad. Sci. USA 95, 3077–3081 (1998).
pubmed: 9501218
doi: 10.1073/pnas.95.6.3077
pmcid: 19697
Li, T. et al. Phospholipid-flippase chaperone CDC50A is required for synapse maintenance by regulating phosphatidylserine exposure. EMBO J. 40, e107915 (2021).
pubmed: 34585770
doi: 10.15252/embj.2021107915
pmcid: 8561630
Mühle, C. et al. Characterization of acid sphingomyelinase activity in human cerebrospinal fluid. PLoS ONE 8, e62912 (2013).
pubmed: 23658784
doi: 10.1371/journal.pone.0062912
pmcid: 3642176
Kornhuber, J., Rhein, C., Müller, C. P. & Mühle, C. Secretory sphingomyelinase in health and disease. Biol. Chem. 396, 707–736 (2015).
pubmed: 25803076
doi: 10.1515/hsz-2015-0109
Pater, J. A. et al. Autosomal dominant non-syndromic hearing loss maps to DFNA33 (13q34) and co-segregates with splice and frameshift variants in ATP11A, a phospholipid flippase gene. Hum. Genet. 141, 431–444 (2022).
pubmed: 35278131
doi: 10.1007/s00439-022-02444-x
pmcid: 9035003
Okamoto, S. et al. The N- or C-terminal cytoplasmic regions of P4-ATPases determine their cellular localization. Mol. Biol. Cell 31, 2115–2124 (2020).
pubmed: 32614659
doi: 10.1091/mbc.E20-04-0225
pmcid: 7530900
Feenstra, B. et al. Common variants associated with general and MMR vaccine-related febrile seizures. Nat. Genet. 46, 1274–1282 (2014).
pubmed: 25344690
doi: 10.1038/ng.3129
pmcid: 4244308
Wang, T. A. et al. TMEM16C is involved in thermoregulation and protects rodent pups from febrile seizures. Proc. Natl Acad. Sci. USA 118, e202334211 (2021).
Zanni, E. D., Gradogna, A., Picco, C., Scholz-Starke, J. & Boccaccio, A. TMEM16E/ANO5 mutations related to bone dysplasia or muscular dystrophy cause opposite effects on lipid scrambling. Hum. Mutat. 41, 1157–1170 (2020).
pubmed: 32112655
doi: 10.1002/humu.24006
Zwaal, R. F. & Schroit, A. J. Pathophysiologic implications of membrane phospholipid asymmetry in blood cells. Blood 89, 1121–1132 (1997).
pubmed: 9028933
doi: 10.1182/blood.V89.4.1121
Marconi, C. et al. A novel missense mutation in ANO5/TMEM16E is causative for gnathodiaphyseal dyplasia in a large Italian pedigree. Eur. J. Hum. Genet. 21, 613–619 (2013).
pubmed: 23047743
doi: 10.1038/ejhg.2012.224
Savarese, M. et al. Next generation sequencing on patients with LGMD and nonspecific myopathies: findings associated with ANO5 mutations. Neuromuscul. Disord. 25, 533–541 (2015).
pubmed: 25891276
doi: 10.1016/j.nmd.2015.03.011
pmcid: 4502439
Tsutsumi, S. et al. The novel gene encoding a putative transmembrane protein is mutated in gnathodiaphyseal dysplasia (GDD). Am. J. Hum. Genet. 74, 1255–1261 (2004).
pubmed: 15124103
doi: 10.1086/421527
pmcid: 1182089
Xu, J. et al. Genetic disruption of Ano5 in mice does not recapitulate human ANO5-deficient muscular dystrophy. Skelet. Muscle 5, 43 (2015).
pubmed: 26693275
doi: 10.1186/s13395-015-0069-z
pmcid: 4685631
Sui, T. et al. Development of muscular dystrophy in a CRISPR-engineered mutant rabbit model with frame-disrupting ANO5 mutations. Cell Death Dis. 9, 609 (2018).
pubmed: 29789544
doi: 10.1038/s41419-018-0674-y
pmcid: 5964072
Millington-Burgess, S. L. & Harper, M. T. Gene of the issue: ANO6 and Scott syndrome. Platelets 31, 964–967 (2020).
pubmed: 31746257
doi: 10.1080/09537104.2019.1693039
Castoldi, E., Collins, P. W., Williamson, P. L. & Bevers, E. M. Compound heterozygosity for 2 novel TMEM16F mutations in a patient with Scott syndrome. Blood 117, 4399–4400 (2011).
pubmed: 21511967
doi: 10.1182/blood-2011-01-332502
Boisseau, P. et al. A new mutation of ANO6 in two familial cases of Scott syndrome. Br. J. Haematol. 180, 750–752 (2016).
pubmed: 27879994
doi: 10.1111/bjh.14439
Rosing, J. et al. Impaired factor X and prothrombin activation associated with decreased phospholipid exposure in platelets from a patient with a bleeding disorder. Blood 65, 1557–1561 (1985).
pubmed: 3995186
doi: 10.1182/blood.V65.6.1557.bloodjournal6561557
Brooks, M. B. et al. A TMEM16F point mutation causes an absence of canine platelet TMEM16F and ineffective activation and death-induced phospholipid scrambling. J. Thromb. Haemost. 13, 2240–2252 (2015).
pubmed: 26414452
doi: 10.1111/jth.13157
Das, S. et al. NGEP, a prostate-specific plasma membrane protein that promotes the association of LNCaP cells. Cancer Res. 67, 1594–1601 (2007).
pubmed: 17308099
doi: 10.1158/0008-5472.CAN-06-2673
Wahlström, G. et al. The variant rs77559646 associated with aggressive prostate cancer disrupts ANO7 mRNA splicing and protein expression. Hum. Mol. Genet. 31, 2063–2077 (2022).
pubmed: 35043958
doi: 10.1093/hmg/ddac012
pmcid: 9239746
Renaud, M. et al. Autosomal recessive cerebellar ataxia type 3 due to ANO10 mutations: delineation and genotype–phenotype correlation study. JAMA Neurol. 71, 1305–1310 (2014).
pubmed: 25089919
doi: 10.1001/jamaneurol.2014.193
Balreira, A. et al. ANO10 mutations cause ataxia and coenzyme Q
pubmed: 25182700
doi: 10.1007/s00415-014-7476-7
pmcid: 4221650
Vermeer, S. et al. Targeted next-generation sequencing of a 12.5 Mb homozygous region reveals ANO10 mutations in patients with autosomal-recessive cerebellar ataxia. Am. J. Hum. Genet. 87, 813–819 (2010).
pubmed: 21092923
doi: 10.1016/j.ajhg.2010.10.015
pmcid: 2997370
Kramer, J. & Hawley, R. S. The spindle-associated transmembrane protein Axs identifies a membranous structure ensheathing the meiotic spindle. Nat. Cell Biol. 5, 261–263 (2003).
pubmed: 12646877
doi: 10.1038/ncb944
Lee, S., Russo, D. & Redman, C. M. The Kell blood group system: Kell and XK membrane proteins. Semin. Hematol. 37, 113–121 (2000).
pubmed: 10791880
doi: 10.1016/S0037-1963(00)90036-2
Peikert, K., et al. VPS13A disease. National Library of Medicine. https://www.ncbi.nlm.nih.gov/books/NBK1387/ (2023).
Zhu, X. et al. Giant axon formation in mice lacking Kell, XK, or Kell and XK animal models of McLeod neuroacanthocytosis syndrome. Ame. J. Pathol. 184, 800–807 (2014).
doi: 10.1016/j.ajpath.2013.11.013
Yamashita, Y., Suzuki, C., Uchiyama, Y. & Nagata, S. Infertility caused by inefficient apoptotic germ cell clearance in Xkr8-deficient male mice. Mol. Cell. Biol. 40, e00402–e00419 (2020).
pubmed: 31712393
doi: 10.1128/MCB.00402-19
pmcid: 6965033
Hanayama, R. et al. Autoimmune disease and impaired uptake of apoptotic cells in MFG-E8-deficient mice. Science 304, 1147–1150 (2004).
pubmed: 15155946
doi: 10.1126/science.1094359
Miyanishi, M., Segawa, K. & Nagata, S. Synergistic effect of Tim4 and MFG-E8 null mutations on the development of autoimmunity. Int. Immunol. 24, 551–559 (2012).
pubmed: 22723547
doi: 10.1093/intimm/dxs064
Kaneshiro, N. et al. Lipid flippase dysfunction as a therapeutic target for endosomal anomalies in Alzheimer’s disease. iScience 25, 103869 (2022).
pubmed: 35243232
doi: 10.1016/j.isci.2022.103869
pmcid: 8857600
Devaux, P. F. Is lipid translocation involved during endo- and exocytosis? Biochimie 82, 497–509 (2000).
pubmed: 10865135
doi: 10.1016/S0300-9084(00)00209-1
Hu, Y. et al. Scramblase TMEM16F terminates T cell receptor signaling to restrict T cell exhaustion. J. Exp. Med. 213, 2759–2772 (2016).
pubmed: 27810927
doi: 10.1084/jem.20160612
pmcid: 5110022
Baker, R. W. & Hughson, F. M. Chaperoning SNARE assembly and disassembly. Nat. Rev. Mol. Cell. Biol. 17, 465–479 (2016).
pubmed: 27301672
doi: 10.1038/nrm.2016.65
pmcid: 5471617
Shin, H.-W. & Takatsu, H. Substrates of P4-ATPases: beyond aminophospholipids (phosphatidylserine and phosphatidylethanolamine). FASEB J. 33, 3087–3096 (2019).
pubmed: 30509129
doi: 10.1096/fj.201801873R
Levano, K. et al. Atp8a1 deficiency is associated with phosphatidylserine externalization in hippocampus and delayed hippocampus-dependent learning. J. Neurochem. 120, 302–313 (2012).
pubmed: 22007859
doi: 10.1111/j.1471-4159.2011.07543.x
McMillan, H. J. et al. Recessive mutations in ATP8A2 cause severe hypotonia, cognitive impairment, hyperkinetic movement disorders and progressive optic atrophy. Orphanet J. Rare Dis. https://doi.org/10.1186/s13023-018-0825-3 (2018).
doi: 10.1186/s13023-018-0825-3
pubmed: 30012219
pmcid: 6048855
Guissart, C. et al. ATP8A2-related disorders as recessive cerebellar ataxia. J. Neurol. 267, 203–213 (2020).
pubmed: 31612321
doi: 10.1007/s00415-019-09579-4
Zhu, X. et al. Mutations in a P-type ATPase gene cause axonal degeneration. PLoS Genet. 8, e1002853 (2012).
pubmed: 22912588
doi: 10.1371/journal.pgen.1002853
pmcid: 3415440
Klomp, L. W. et al. Characterization of mutations in ATP8B1 associated with hereditary cholestasis. Hepatology 40, 27–38 (2004).
pubmed: 15239083
doi: 10.1002/hep.20285
Stapelbroek, J. M. et al. ATP8B1 is essential for maintaining normal hearing. Proc. Natl Acad. Sci. USA 106, 9709–9714 (2009).
pubmed: 19478059
doi: 10.1073/pnas.0807919106
pmcid: 2700994
Vogt, G. et al. Biallelic truncating variants in ATP9A cause a novel neurodevelopmental disorder involving postnatal microcephaly and failure to thrive. J. Med. Genet. 59, 662–668 (2022).
pubmed: 34379057
doi: 10.1136/jmedgenet-2021-107843
Mattioli, F. et al. Biallelic truncation variants in ATP9A are associated with a novel autosomal recessive neurodevelopmental disorder. NPJ Genom. Med. 6, 94 (2021).
pubmed: 34764295
doi: 10.1038/s41525-021-00255-z
pmcid: 8586153
Meguro, M. et al. A novel maternally expressed gene, ATP10C, encodes a putative aminophospholipid translocase associated with Angelman syndrome. Nat. Genet. 28, 19–20 (2001).
pubmed: 11326269
doi: 10.1038/ng0501-19
Dhar, M. S., Yuan, J. S., Elliott, S. B. & Sommardahl, C. A type IV P-type ATPase affects insulin-mediated glucose uptake in adipose tissue and skeletal muscle in mice. J. Nutr. Biochem. 17, 811–820 (2006).
pubmed: 16517145
doi: 10.1016/j.jnutbio.2006.01.002
Real, R. et al. ATP10B and the risk for Parkinson’s disease. Acta Neuropathol. 140, 401–402 (2020).
pubmed: 32556962
doi: 10.1007/s00401-020-02172-4
pmcid: 7540943
Roland, B. P. et al. Yeast and human P4-ATPases transport glycosphingolipids using conserved structural motifs. J. Biol. Chem. 294, 1794–1806 (2019).
pubmed: 30530492
doi: 10.1074/jbc.RA118.005876
Sigruener, A. et al. Lipidomic and metabolic changes in the P4-type ATPase ATP10D deficient C57BL/6J wild type mice upon rescue of ATP10D function. PLoS ONE 12, e0178368 (2017).
pubmed: 28542499
doi: 10.1371/journal.pone.0178368
pmcid: 5444826
Charlesworth, G. et al. Mutations in ANO3 cause dominant craniocervical dystonia: ion channel implicated in pathogenesis. Am. J. Hum. Genet. 91, 1041–1050 (2012).
pubmed: 23200863
doi: 10.1016/j.ajhg.2012.10.024
pmcid: 3516598
Jun, I. et al. ANO9/TMEM16J promotes tumourigenesis via EGFR and is a novel therapeutic target for pancreatic cancer. Br. J. Cancer 117, 1798–1809 (2017).
pubmed: 29024940
doi: 10.1038/bjc.2017.355
pmcid: 5729472
Li, C., Cai, S., Wang, X. & Jiang, Z. Identification and characterization of ANO9 in stage II and III colorectal carcinoma. Oncotarget 6, 29324–29334 (2015).
pubmed: 26317553
doi: 10.18632/oncotarget.4979
pmcid: 4745729
Chrysanthou, A., Ververis, A. & Christodoulou, K. ANO10 function in health and disease. Cerebellum https://doi.org/10.1007/s12311-022-01395-3 (2022).
doi: 10.1007/s12311-022-01395-3
pubmed: 35648332
pmcid: 10126014
Chung, J. et al. PI4P/phosphatidylserine countertransport at ORP5- and ORP8-mediated ER–plasma membrane contacts. Science 349, 428–432 (2015).
pubmed: 26206935
doi: 10.1126/science.aab1370
pmcid: 4638224
Pinot, M. et al. Polyunsaturated phospholipids facilitate membrane deformation and fission by endocytic proteins. Science 345, 693–697 (2014).
pubmed: 25104391
doi: 10.1126/science.1255288
Corbalán-García, S. & Gómez-Fernández, J. C. Classical protein kinases C are regulated by concerted interaction with lipids: the importance of phosphatidylinositol-4,5-bisphosphate. Biophys. Rev. 6, 3–14 (2013).
pubmed: 28509956
doi: 10.1007/s12551-013-0125-z
pmcid: 5427809
Zhou, Y. et al. Membrane potential modulates plasma membrane phospholipid dynamics and K-Ras signaling. Science 349, 873–876 (2015).
pubmed: 26293964
doi: 10.1126/science.aaa5619
pmcid: 4687752
Yeung, T. et al. Membrane phosphatidylserine regulates surface charge and protein localization. Science 319, 210–213 (2008).
pubmed: 18187657
doi: 10.1126/science.1152066
Yeung, T. et al. Receptor activation alters inner surface potential during phagocytosis. Science 313, 347–351 (2006).
pubmed: 16857939
doi: 10.1126/science.1129551
Fairn, G. D., Hermansson, M., Somerharju, P. & Grinstein, S. Phosphatidylserine is polarized and required for proper Cdc42 localization and for development of cell polarity. Nat. Cell Biol. 13, 1424–1430 (2011).
pubmed: 21964439
doi: 10.1038/ncb2351
Liu, X. et al. Inflammasome-activated gasdermin D causes pyroptosis by forming membrane pores. Nature 535, 153–158 (2016).
pubmed: 27383986
doi: 10.1038/nature18629
pmcid: 5539988
Deng, W. et al. Streptococcal pyrogenic exotoxin B cleaves GSDMA and triggers pyroptosis. Nature 602, 496–502 (2022).
pubmed: 35110732
doi: 10.1038/s41586-021-04384-4
pmcid: 9703647