Lipid Polymorphism of the Subchloroplast-Granum and Stroma Thylakoid Membrane-Particles. II. Structure and Functions
Jazyk angličtina Země Švýcarsko Médium electronic
Typ dokumentu časopisecké články, práce podpořená grantem
Grantová podpora
19-13637S
Grantová Agentura České Republiky
K 128679
Hungarian Scientific Research Fund
GINOP-2.3.2-15-2016-00058
National Research Development and Innovation Office of Hungary
CZ.02.1.01/0.0/0.0/16_019/0000797
SustES - Adaptation strategies for sustainable ecosystem services and food security under adverse environmental conditions
CZ.02.1.01/0.0/0.0/ 16_019/0000827
European Regional Development Fund
07359/2019/RRC
Silesian Region
SGS02/PřF/2021
Ostravská Univerzita v Ostravě
LM2015043
Ministry of Education, Youth and Science
LM2018123
Ministry of Education, Youth and Science
PubMed
34572012
PubMed Central
PMC8472583
DOI
10.3390/cells10092363
PII: cells10092363
Knihovny.cz E-zdroje
- Klíčová slova
- SAXS, bilayer, chlorophyll fluorescence, cryo-electron-tomography, electron microscopy, membrane energization, membrane networks, non-bilayer lipid phases, violaxanthin de-epoxidase,
- MeSH
- cirkulární dichroismus metody MeSH
- elektronová mikroskopie metody MeSH
- fotosyntéza genetika MeSH
- lipidy genetika MeSH
- magnetická rezonanční spektroskopie metody MeSH
- tylakoidy genetika MeSH
- Publikační typ
- časopisecké články MeSH
- práce podpořená grantem MeSH
- Názvy látek
- lipidy MeSH
In Part I, by using 31P-NMR spectroscopy, we have shown that isolated granum and stroma thylakoid membranes (TMs), in addition to the bilayer, display two isotropic phases and an inverted hexagonal (HII) phase; saturation transfer experiments and selective effects of lipase and thermal treatments have shown that these phases arise from distinct, yet interconnectable structural entities. To obtain information on the functional roles and origin of the different lipid phases, here we performed spectroscopic measurements and inspected the ultrastructure of these TM fragments. Circular dichroism, 77 K fluorescence emission spectroscopy, and variable chlorophyll-a fluorescence measurements revealed only minor lipase- or thermally induced changes in the photosynthetic machinery. Electrochromic absorbance transients showed that the TM fragments were re-sealed, and the vesicles largely retained their impermeabilities after lipase treatments-in line with the low susceptibility of the bilayer against the same treatment, as reflected by our 31P-NMR spectroscopy. Signatures of HII-phase could not be discerned with small-angle X-ray scattering-but traces of HII structures, without long-range order, were found by freeze-fracture electron microscopy (FF-EM) and cryo-electron tomography (CET). EM and CET images also revealed the presence of small vesicles and fusion of membrane particles, which might account for one of the isotropic phases. Interaction of VDE (violaxanthin de-epoxidase, detected by Western blot technique in both membrane fragments) with TM lipids might account for the other isotropic phase. In general, non-bilayer lipids are proposed to play role in the self-assembly of the highly organized yet dynamic TM network in chloroplasts.
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Dlouhý O., Javornik U., Zsiros O., Šket P., Karlický V., Špunda V., Plavec J., Garab G. Lipid polymorphism of the subchloroplast—granum and stroma thylakoid membrane—particles. I. 31P-NMR spectroscopy. Cells. 2021;10:2354. PubMed PMC
Mitchell P. Chemiosmotic coupling in oxidative and photosynthetic phosphorylation. Biol. Rev. Camb. Philos. Soc. 1966;41:445–502. doi: 10.1111/j.1469-185X.1966.tb01501.x. PubMed DOI
Douce R., Joyard J. Biosynthesis of Thylakoid Membrane Lipids. In: Ort D.R., Yocum C.F., Heichel I.F., editors. Oxygenic Photosynthesis: The Light Reactions. Springer; Dordrecht, The Netherlands: 1996. pp. 69–101.
Krumova S.B., Dijkema C., de Waard P., Van As H., Garab G., van Amerongen H. Phase behaviour of phosphatidylglycerol in spinach thylakoid membranes as revealed by P-31-NMR. Biochim. Biophys. Acta. 2008;1778:997–1003. doi: 10.1016/j.bbamem.2008.01.004. PubMed DOI
Garab G., Ughy B., de Waard P., Akhtar P., Javornik U., Kotakis C., Šket P., Karlický V., Materová Z., Špunda V., et al. Lipid polymorphism in chloroplast thylakoid membranes—as revealed by P-31-NMR and timeresolved merocyanine fluorescence spectroscopy. Sci. Rep. 2017;7 doi: 10.1038/s41598-017-13574-y. PubMed DOI PMC
Ughy B., Karlický V., Dlouhý O., Javornik U., Materová Z., Zsiros O., Šket P., Plavec J., Špunda V., Garab G. Lipid-polymorphism of plant thylakoid membranes. Enhanced non-bilayer lipid phases associated with increased membrane permeability. Physiol. Plant. 2019;166:278–287. doi: 10.1111/ppl.12929. PubMed DOI
Dlouhý O., Kurasová I., Karlický V., Javornik U., Šket P., Petrova N.Z., Krumova S.B., Plavec J., Ughy B., Špunda V., et al. Modulation of non-bilayer lipid phases and the structure and functions of thylakoid membranes: Effects on the water-soluble enzyme violaxanthin de-epoxidase. Sci. Rep. 2020;10:11959. doi: 10.1038/s41598-020-68854-x. PubMed DOI PMC
Garab G., van Amerongen H. Linear dichroism and circular dichroism in photosynthesis research. Photosynth. Res. 2009;101:135–146. doi: 10.1007/s11120-009-9424-4. PubMed DOI PMC
Lamb J.J., Rokke G., Hohmann-Marriott M.F. Chlorophyll fluorescence emission spectroscopy of oxygenic organisms at 77 K. Photosynthetica. 2018;56:105–124. doi: 10.1007/s11099-018-0791-y. DOI
Kalaji H.M., Jajoo A., Oukarroum A., Brestic M., Zivcak M., Samborska I.A., Cetner M.D., Lukasik I., Goltsev V., Ladle R.J. Chlorophyll a fluorescence as a tool to monitor physiological status of plants under abiotic stress conditions. Acta Physiol. Plant. 2016;38:102. doi: 10.1007/s11738-016-2113-y. DOI
Lazár D. Chlorophyll a fluorescence induction. Biochim. Biophys. Acta. 1999;1412:1–28. doi: 10.1016/S0005-2728(99)00047-X. PubMed DOI
Sipka G., Magyar M., Mezzetti A., Akhtar P., Zhu Q., Xiao Y., Han G., Santabarbara S., Shen J.-R., Lambrev P.H., et al. Light-adapted charge-separated state of photosystem II: Structural and functional dynamics of the closed reaction center. Plant Cell. 2021;33:1286–1302. doi: 10.1093/plcell/koab008. PubMed DOI PMC
Bailleul B., Cardol P., Breyton C., Finazzi G. Electrochromism: A useful probe to study algal photosynthesis. Photosynth. Res. 2010;106:179–189. doi: 10.1007/s11120-010-9579-z. PubMed DOI
Pabst G., Koschuch R., Pozo-Navas B., Rappolt M., Lohner K., Laggner P. Structural analysis of weakly ordered membrane stacks. J. Appl. Crystallogr. 2003;36:1378–1388. doi: 10.1107/S0021889803017527. DOI
Jakubauskas D., Mortensen K., Jensen P.E., Kirkensgaard J.J.K. Small-Angle X-Ray and Neutron Scattering on Photosynthetic Membranes. Front. Chem. 2021;9:82. doi: 10.3389/fchem.2021.631370. PubMed DOI PMC
Semenova G.A. The relationship between the transformation of thylakoid acyl lipids and the formation of tubular lipid aggregates visible on fracture faces. J. Plant Physiol. 1999;155:669–677. doi: 10.1016/S0176-1617(99)80081-9. DOI
Williams W.P. The Physical Properties of Thylakoid Membrane Lipids and Their Relation to Photosynthesis. In: Paul-André S., Norio M., editors. Lipids in Photosynthesis: Structure, Function and Genetics. Springer; Dordrecht, The Netherlands: 1998. pp. 103–118.
Sprague S.G., Staehelin L.A. Effects of reconstitution method on the structural organization of isolated chloroplast membrane-lipids. Biochim. Biophys. Acta. 1984;777:306–322. doi: 10.1016/0005-2736(84)90433-4. DOI
Daum B., Kuhlbrandt W. Electron tomography of plant thylakoid membranes. J. Exp. Bot. 2011;62:2393–2402. doi: 10.1093/jxb/err034. PubMed DOI
Bussi Y., Shimoni E., Weiner A., Kapon R., Charuvi D., Nevo R., Efrati E., Reich Z. Fundamental helical geometry consolidates the plant photosynthetic membrane. Proc. Natl. Acad. Sci. USA. 2019;116:22366–22375. doi: 10.1073/pnas.1905994116. PubMed DOI PMC
Mustárdy L.A., Janossy A.G.S. Evidence of helical thylakoid arrangement by scanning electron-microscopy. Plant Sci. Lett. 1979;16:281–284. doi: 10.1016/0304-4211(79)90039-7. DOI
Anderson J.M., Boardman N.K. Fractionation of photochemical systems of photosynthesis. I. Chlorophyll contents and photochemical activities of particles isolated from spinach chloroplasts. Biochim. Biophys. Acta. 1966;112:403–421. doi: 10.1016/0926-6585(66)90244-5. PubMed DOI
Peters F., Vanwielink J.E., Sang H., Devries S., Kraayenhof R. Studies on well coupled photosystem I-enriched subchloroplast vesicles—content and redox properties of electron-transfer components. Biochim. Biophys. Acta. 1983;722:460–470. doi: 10.1016/0005-2728(83)90062-2. DOI
Cuello J., Quiles M.J. Fractionation of thylakoid membranes into grana and stroma thylakoids. Methods Mol. Biol. 2004;274:1–9. doi: 10.1385/1-59259-799-8:001. PubMed DOI
Pavlovič A., Stolárik T., Nosek L., Kouřil R., Ilík P. Light-induced gradual activation of photosystem II in dark-grown Norway spruce seedlings. Biochim. Biophys. Acta-Bioenerg. 2016;1857:799–809. doi: 10.1016/j.bbabio.2016.02.009. PubMed DOI
Saga G., Giorgetti A., Fufezan C., Giacometti G.M., Bassi R., Morosinotto T. Mutation Analysis of Violaxanthin De-epoxidase Identifies Substrate-binding Sites and Residues Involved in Catalysis. J. Biol. Chem. 2010;285:23763–23778. doi: 10.1074/jbc.M110.115097. PubMed DOI PMC
Wacha A., Varga Z., Bota A. CREDO: A new general-purpose laboratory instrument for small-angle X-ray scattering. J. Appl. Crystallogr. 2014;47:1749–1754. doi: 10.1107/S1600576714019918. DOI
Wacha A. Optimized pinhole geometry for small-angle scattering. J. Appl. Crystallogr. 2015;48:1843–1848. doi: 10.1107/S1600576715018932. DOI
Severs N.J. Freeze-fracture electron microscopy. Nat. Protoc. 2007;2:547–576. doi: 10.1038/nprot.2007.55. PubMed DOI
Arshad R., Calvaruso C., Boekema E.J., Büchel C., Kouřil R. Revealing the architecture of the photosynthetic apparatus in the diatom Thalassiosira pseudonana. Plant Physiol. 2021 doi: 10.1093/plphys/kiab208. PubMed DOI PMC
Mastronarde D.N. Automated electron microscope tomography using robust prediction of specimen movements. J. Struct. Biol. 2005;152:36–51. doi: 10.1016/j.jsb.2005.07.007. PubMed DOI
Tang G., Peng L., Baldwin P.R., Mann D.S., Jiang W., Rees I., Ludtke S.J. EMAN2: An extensible image processing suite for electron microscopy. J. Struct. Biol. 2007;157:38–46. doi: 10.1016/j.jsb.2006.05.009. PubMed DOI
Frangakis A.S., Hegerl R. Noise reduction in electron tomographic reconstructions using nonlinear anisotropic diffusion. J. Struct. Biol. 2001;135:239–250. doi: 10.1006/jsbi.2001.4406. PubMed DOI
Kremer J.R., Mastronarde D.N., McIntosh J.R. Computer visualization of three-dimensional image data using IMOD. J. Struct. Biol. 1996;116:71–76. doi: 10.1006/jsbi.1996.0013. PubMed DOI
Dobrikova A.G., Várkonyi Z., Krumova S.B., Kovács L., Kostov G.K., Todinova S.J., Busheva M.C., Taneva S.G., Garab G. Structural Rearrangements in chloroplast thylakoid membranes revealed by differential scanning calorimetry and circular dichroism spectroscopy. Thermo-optic effect. Biochemistry. 2003;42:11272–11280. doi: 10.1021/bi034899j. PubMed DOI
Lambrev P.H., Varkonyi Z., Krumova S., Kovacs L., Miloslavina Y., Holzwarth A.R., Garab G. Importance of trimer-trimer interactions for the native state of the plant light-harvesting complex II. Biochim. Biophys. Acta-Bioenerg. 2007;1767:847–853. doi: 10.1016/j.bbabio.2007.01.010. PubMed DOI
Berthold D.A., Babcock G.T., Yocum C.F. A highly resolved, oxygen-evolving photosystem-II preparation from spinach thylakoid membranes—electron-paramagnetic-res and electron-transport properties. FEBS Lett. 1981;134:231–234. doi: 10.1016/0014-5793(81)80608-4. DOI
Morosinotto T., Segalla A., Giacometti G.M., Bassi R. Purification of structurally intact grana from plants thylakoids membranes. J. Bioenerg. Biomembr. 2010;42:37–45. doi: 10.1007/s10863-009-9261-3. PubMed DOI
Miloslavina Y., Lambrev P.H., Jávorfi T., Várkonyi Z., Karlický V., Wall J.S., Hind G., Garab G. Anisotropic circular dichroism signatures of oriented thylakoid membranes and lamellar aggregates of LHCII. Photosynth. Res. 2012;111:29–39. doi: 10.1007/s11120-011-9664-y. PubMed DOI
Nellaepalli S., Zsiros O., Toth T., Yadavalli V., Garab G., Subramanyam R., Kovacs L. Heat- and light-induced detachment of the light harvesting complex from isolated photosystem I supercomplexes. J. Photochem. Photobiol. B-Biol. 2014;137:13–20. doi: 10.1016/j.jphotobiol.2014.04.026. PubMed DOI
Tian L.J., Xu P.Q., Chukhutsina V.U., Holzwarth A.R., Croce R. Zeaxanthin-dependent nonphotochemical quenching does not occur in photosystem I in the higher plant Arabidopsis thaliana. Proc. Natl. Acad. Sci. USA. 2017;114:4828–4832. doi: 10.1073/pnas.1621051114. PubMed DOI PMC
Krumova S.B., Várkonyi Z., Lambrev P.H., Kovács L., Todinova S.J., Busheva M.C., Taneva S.G., Garab G. Heat- and light-induced detachment of the light-harvesting antenna complexes of photosystem I in isolated stroma thylakoid membranes. J. Photochem. Photobiol. B-Biol. 2014;137:4–12. doi: 10.1016/j.jphotobiol.2014.04.029. PubMed DOI
Akhtar P., Lingvay M., Kiss T., Deák R., Bóta A., Ughy B., Garab G., Lambrev P.H. Excitation energy transfer between Light-harvesting complex II and Photosystem I in reconstituted membranes. Biochim. Biophys. Acta-Bioenerg. 2016;1857:462–472. doi: 10.1016/j.bbabio.2016.01.016. PubMed DOI
Xu P.Q., Tian L.J., Kloz M., Croce R. Molecular insights into Zeaxanthin-dependent quenching in higher plants. Sci. Rep. 2015;5:13679. doi: 10.1038/srep13679. PubMed DOI PMC
Akhtar P., Dorogi M., Pawlak K., Kovács L., Bóta A., Kiss T., Garab G., Lambrev P.H. Pigment Interactions in Light-harvesting Complex II in Different Molecular Environments. J. Biol. Chem. 2015;290:4877–4886. doi: 10.1074/jbc.M114.607770. PubMed DOI PMC
Garab G.I., Horváth G., Faludi-Dániel A. Resolution of fluorescence bands in greening chloroplasts of maize. Biochem. Biophys. Res. Commun. 1974;56:1004–1009. doi: 10.1016/S0006-291X(74)80288-3. PubMed DOI
Williams W.P., Brain A.P.R., Dominy P.J. Induction of nonbilayer lipid phase separations in chloroplast thylakoid membranes by compatible co-solutes and its relation to the thermal-stability of photosystem-II. Biochim. Biophys. Acta. 1992;1099:137–144. doi: 10.1016/0005-2728(92)90210-S. DOI
Kotakis C., Akhtar P., Zsiros O., Garab G., Lambrev P.H. Increased thermal stability of photosystem II and the macro-organization of thylakoid membranes, induced by co-solutes, associated with changes in the lipid-phase behaviour of thylakoid membranes. Photosynthetica. 2018;56:254–264. doi: 10.1007/s11099-018-0782-z. DOI
Mustardy L., Garab G. Granum revisited. A three-dimensional model—where things fall into place. Trends Plant Sci. 2003;8:117–122. doi: 10.1016/S1360-1385(03)00015-3. PubMed DOI
Peters F., van Spanning R., Kraayenhof R. Studies on well coupled photosystem I-enriched subchloroplast vesicles—optimization of ferredoxin-mediated cyclic phosphorylation and electric-potential generation. Biochim. Biophys. Acta. 1983;724:159–165. doi: 10.1016/0005-2728(83)90037-3. DOI
Latowski D., Akerlund H.E., Strzalka K. Violaxanthin de-epoxidase, the xanthophyll cycle enzyme, requires lipid inverted hexagonal structures for its activity. Biochemistry. 2004;43:4417–4420. doi: 10.1021/bi049652g. PubMed DOI
Goss R., Lohr M., Latowski D., Grzyb J., Vieler A., Wilhelm C., Strzalka K. Role of hexagonal structure-forming lipids in diadinoxanthin and violaxanthin solubilization and de-epoxidation. Biochemistry. 2005;44:4028–4036. doi: 10.1021/bi047464k. PubMed DOI
Rockholm D.C., Yamamoto H.Y. Violaxanthin de-epoxidase—Purification of a 43-kilodalton lumenal protein from lettuce by lipid-affinity precipitation with monogalactosyldiacylglyceride. Plant Physiol. 1996;110:697–703. doi: 10.1104/pp.110.2.697. PubMed DOI PMC
Simionato D., Basso S., Zaffagnini M., Lana T., Marzotto F., Trost P., Morosinotto T. Protein redox regulation in the thylakoid lumen: The importance of disulfide bonds for violaxanthin de-epoxidase. FEBS Lett. 2015;589:919–923. doi: 10.1016/j.febslet.2015.02.033. PubMed DOI
Turner D.C., Gruner S.M. X-ray-diffraction reconstruction of the inverted hexagonal (HII) phase in lipid water-systems. Biochemistry. 1992;31:1340–1355. doi: 10.1021/bi00120a009. PubMed DOI
Kirkensgaard J.J.K., Holm J.K., Larsen J.K., Posselt D. Simulation of small-angle X-ray scattering from thylakoid membranes. J. Appl. Crystallogr. 2009;42:649–659. doi: 10.1107/S0021889809017701. DOI
Unnep R., Zsiros O., Hörcsik Z., Markó M., Jajoo A., Kohlbrecher J., Garab G., Nagy G. Low-pH induced reversible reorganizations of chloroplast thylakoid membranes—As revealed by small-angle neutron scattering. Biochim. Biophys. Acta-Bioenerg. 2017;1858:360–365. doi: 10.1016/j.bbabio.2017.02.010. PubMed DOI
Stock D., Leslie A.G.W., Walker J.E. Molecular architecture of the rotary motor in ATP synthase. Science. 1999;286:1700–1705. doi: 10.1126/science.286.5445.1700. PubMed DOI
Kirchhoff H., Haase W., Wegner S., Danielsson R., Ackermann R., Albertsson P.A. Low-light-induced formation of semicrystalline photosystem II arrays in higher plant chloroplast. Biochemistry. 2007;46:11169–11176. doi: 10.1021/bi700748y. PubMed DOI
Simidjiev I., Stoylova S., Amenitsch H., Javorfi T., Mustárdy L., Laggner P., Holzenburg A., Garab G. Self-assembly of large, ordered lamellae from non-bilayer lipids and integral membrane proteins in vitro. Proc. Natl. Acad. Sci. USA. 2000;97:1473–1476. doi: 10.1073/pnas.97.4.1473. PubMed DOI PMC
Holm J.K. Structure and Structural Flexibility of Thylakoid Membranes. University of Roskilde; Roskilde, Denmark: 2004.
Staehelin L.A. Chloroplast Structure and Supramolecular Organization of Photosynthetic Membranes. In: Staehelin L.A., Arntzen C.J., editors. Photosynthesis III: Photosynthetic Membranes and Light Harvesting Systems. Springer; Berlin/Heidelberg, Germany: 1986. pp. 1–84.
Gounaris K., Sen A., Brain A.P.R., Quinn P.J., Williams W.P. The formation of non-bilayer structures in total polar lipid extracts of chloroplast membranes. Biochim. Biophys. Acta. 1983;728:129–139. doi: 10.1016/0005-2736(83)90445-5. DOI
Oszlánczi A., Bóta A., Klumpp E. Influence of aminoglycoside antibiotics on the thermal behaviour and structural features of DPPE-DPPG model membranes. Colloids Surf. B Biointerfaces. 2010;75:141–148. doi: 10.1016/j.colsurfb.2009.08.027. PubMed DOI
Dekker J.P., Boekema E.J. Supramolecular organization of thylakoid membrane proteins in green plants. Biochim. Biophys. Acta Bioenerg. 2005;1706:12–39. doi: 10.1016/j.bbabio.2004.09.009. PubMed DOI
Daum B., Nicastro D., Il J.A., McIntosh J.R., Kuhlbrandt W. Arrangement of Photosystem II and ATP Synthase in Chloroplast Membranes of Spinach and Pea. Plant Cell. 2010;22:1299–1312. doi: 10.1105/tpc.109.071431. PubMed DOI PMC
Watts A. NMR of Lipids. In: Roberts G.C.K., editor. Encyclopedia of Biophysics. Springer; Berlin/Heidelberg, Germany: 2013. pp. 1727–1738.
Kouřil R., Oostergetel G.T., Boekema E.J. Fine structure of granal thylakoid membrane organization using cryo electron tomography. Biochim. Biophys. Acta-Bioenerg. 2011;1807:368–374. doi: 10.1016/j.bbabio.2010.11.007. PubMed DOI
van Eerden F.J., de Jong D.H., de Vries A.H., Wassenaar T.A., Marrink S.J. Characterization of thylakoid lipid membranes from cyanobacteria and higher plants by molecular dynamics simulations. Biochim Biophys Acta. 2015;1848:1319–1330. doi: 10.1016/j.bbamem.2015.02.025. PubMed DOI
Kirchhoff H. Chloroplast ultrastructure in plants. New Phytol. 2019;223:565–574. doi: 10.1111/nph.15730. PubMed DOI
Seddon J.M., Templer R.H. Chapter 3—Polymorphism of Lipid-Water Systems. In: Lipowsky R., Sackmann E., editors. Handbook of Biological Physics. Volume 1. North-Holland; Amsterdam, The Netherlands: 1995. pp. 97–160.
Chernomordik L. Non-bilayer lipids and biological fusion intermediates. Chem. Phys. Lipids. 1996;81:203–213. doi: 10.1016/0009-3084(96)02583-2. PubMed DOI
Jouhet J. Importance of the hexagonal lipid phase in biological membrane organization. Front Plant Sci. 2013;4:494. doi: 10.3389/fpls.2013.00494. PubMed DOI PMC
Yang Y., Yao H.W., Hong M. Distinguishing Bicontinuous Lipid Cubic Phases from Isotropic Membrane Morphologies Using P-31 Solid-State NMR Spectroscopy. J. Phys. Chem. B. 2015;119:4993–5001. doi: 10.1021/acs.jpcb.5b01001. PubMed DOI PMC
Mustárdy L., Buttle K., Steinbach G., Garab G. The Three-Dimensional Network of the Thylakoid Membranes in Plants: Quasihelical Model of the Granum-Stroma Assembly. Plant Cell. 2008;20:2552–2557. doi: 10.1105/tpc.108.059147. PubMed DOI PMC
Austin J.R., II, Staehelin L.A. Three-Dimensional Architecture of Grana and Stroma Thylakoids of Higher Plants as Determined by Electron Tomography. Plant Physiol. 2011;155:1601–1611. doi: 10.1104/pp.110.170647. PubMed DOI PMC
de Kruijff B. Biomembranes—Lipids beyond the bilayer. Nature. 1997;386:129–130. doi: 10.1038/386129a0. PubMed DOI
Brown M.F. Curvature Forces in Membrane Lipid-Protein Interactions. Biochemistry. 2012;51:9782–9795. doi: 10.1021/bi301332v. PubMed DOI PMC
Ball W.B., Neff J.K., Gohil V.M. The role of nonbilayer phospholipids in mitochondrial structure and function. FEBS Lett. 2018;592:1273–1290. doi: 10.1002/1873-3468.12887. PubMed DOI PMC
Zhang M., Mileykovskaya E., Dowhan W. Gluing the respiratory chain together - Cardiolipin is required for supercomplex formation in the inner mitochondrial membrane. J. Biol. Chem. 2002;277:43553–43556. doi: 10.1074/jbc.C200551200. PubMed DOI
Pfeiffer K., Gohil V., Stuart R.A., Hunte C., Brandt U., Greenberg M.L., Schagger H. Cardiolipin stabilizes respiratory chain supercomplexes. J. Biol. Chem. 2003;278:52873–52880. doi: 10.1074/jbc.M308366200. PubMed DOI
Heinemeyer J., Braun H.P., Boekema E.J., Kouřil R. A structural model of the cytochrome c reductase/oxidase supercomplex from yeast mitochondria. J. Biol. Chem. 2007;282:12240–12248. doi: 10.1074/jbc.M610545200. PubMed DOI
Schuurmans J.J., Veerman E.C.I., Francke J.A., Torrespereira J.M.G., Kraayenhof R. Temperature-dependence of energy-transducing functions and inhibitor sensitivity in chloroplasts. Plant Physiol. 1984;74:170–175. doi: 10.1104/pp.74.1.170. PubMed DOI PMC
Tikhonov A.N., Vershubskii A.V. Temperature-dependent regulation of electron transport and ATP synthesis in chloroplasts in vitro and in silico. Photosynth. Res. 2020;146:299–329. doi: 10.1007/s11120-020-00777-0. PubMed DOI
Garab G., Lohner K., Laggner P., Farkas T. Self-regulation of the lipid content of membranes by non-bilayer lipids: A hypothesis. Trends Plant Sci. 2000;5:489–494. doi: 10.1016/S1360-1385(00)01767-2. PubMed DOI
Gasanov S.E., Kim A.A., Yaguzhinsky L.S., Dagda R.K. Non-bilayer structures in mitochondrial membranes regulate ATP synthase activity. Biochim. Biophys. Acta. 2018;1860:586–599. doi: 10.1016/j.bbamem.2017.11.014. PubMed DOI PMC
Morelli A.M., Ravera S., Calzia D., Panfoli I. An update of the chemiosmotic theory as suggested by possible proton currents inside the coupling membrane. Open Biol. 2019;9:180221. doi: 10.1098/rsob.180221. PubMed DOI PMC
Role of isotropic lipid phase in the fusion of photosystem II membranes