The thylakoid membranes of vascular plants are differentiated into stacked granum and unstacked stroma regions. The formation of grana is triggered by the macrodomain formation of photosystem II and light-harvesting complex II (PSII-LHCII) and thus their lateral segregation from the photosystem I-light-harvesting complex I (PSI-LHCI) super-complexes and the ATP-synthase; which is then stabilized by stacking interactions of the adjacent PSII-LHCII enriched regions of the thylakoid membranes. The self-assembly and dynamics of this highly organized membrane system and the nature of forces acting between the PSII-LHCII macrodomains are not well understood. By using circular dichroism (CD) spectroscopy, small-angle neutron scattering (SANS) and transmission electron microscopy (TEM), we investigated the effects of Hofmeister salts on the organization of pigment-protein complexes and on the ultrastructure of thylakoid membranes. We found that the kosmotropic agent (NH4)2SO4 and the Hofmeister-neutral NaCl, up to 2 M concentrations, hardly affected the macro-organization of the protein complexes and the membrane ultrastructure. In contrast, chaotropic salts, NaClO4, and NaSCN destroyed the mesoscopic structures, the multilamellar organization of the thylakoid membranes and the chiral macrodomains of the protein complexes but without noticeably affecting the short-range, pigment-pigment excitonic interactions. Comparison of the concentration- and time-dependences of SANS, TEM and CD parameters revealed the main steps of the disassembly of grana in the presence of chaotropes. It begins with a rapid diminishment of the long-range periodic order of the grana membranes, apparently due to an increased stacking disorder of the thylakoid membranes, as reflected by SANS experiments. SANS measurements also allowed discrimination between the cationic and anionic effects-in stacking and disorder, respectively. This step is followed by a somewhat slower disorganization of the TEM ultrastructure, due to the gradual loss of stacked membrane pairs. Occurring last is the stepwise decrease and disappearance of the long-range chiral order of the protein complexes, the rate of which was faster in LHCII-deficient membranes. These data are interpreted in terms of a theory, from our laboratory, according to which Hofmeister salts primarily affect the hydrophylic-hydrophobic interactions of proteins, and the stroma-exposed regions of the intrinsic membrane proteins, in particular-pointing to the role of protein-water interface in the stacking interactions of granum thylakoid membranes.

Department of Physics Faculty of Science University of Ostrava Ostrava Czechia

Institute for Solid State Physics and Optics Wigner Research Centre for Physics Budapest Hungary

Institute of Biophysics Biological Research Centre Szeged Hungary

Institute of Plant Biology Biological Research Centre Szeged Hungary

Jülich Centre for Neutron Science at MLZ Forschungszentrum Jülich GmbH Garching Germany

Laboratory for Neutron Scattering and Imaging Paul Scherrer Institute Villigen PSI Switzerland

Neutron Scattering Division Oak Ridge National Laboratory Oak Ridge TN United States

Neutron Spectroscopy Department Centre for Energy Research Budapest Hungary

Zobrazit více v PubMed

Adam Z., Charuvi D., Tsabari O., Knopf R. R., Reich Z. (2011). Biogenesis of thylakoid networks in angiosperms: knowns and unknowns. Plant Mol. Biol. 76, 221–234.  10.1007/s11103-010-9693-5 PubMed DOI

Albanese P., Tamara S., Saracco G., Scheltema R. A., Pagliano C. (2020). How paired PSII–LHCII supercomplexes mediate the stacking of plant thylakoid membranes unveiled by structural mass-spectrometry. Nat. Commun. 11, 1361.  10.1038/s41467-020-15184-1 PubMed DOI PMC

Andersson B., Anderson J. M. (1980). Lateral heterogeneity in the distribution of chlorophyll–protein complexes of the thylakoid membranes of spinach chloroplasts. Biochim. Biophys. Acta 593, 427– 440.  10.1016/0005-2728(80)90078-X PubMed DOI

Austin J. R., Staehelin L. A. (2011). Three-dimensional architecture of grana and stroma thylakoids of higher plants as determined by electron tomography. Plant Physiol. 155, 1601–1611.  10.1104/pp.110.170647 PubMed DOI PMC

Barber J. (1982). Influence of surface charges on thylakoid structure and function. Ann. Rev. Plant Physiol. 33, 261–295.  10.1146/annurev.pp.33.060182.001401 DOI

Barzda V., Garab G., Gulbinas V., Valkunas L. (1996). Evidence for long-range excitation energy migration in macroaggregates of the chlorophyll ab light-harvesting antenna complexes. Biochim. Biophys. Acta 1273, 231–236.  10.1016/0005-2728(95)00147-6 DOI

Ben-Shem A., Frolow F., Nelson N. (2003). Crystal structure of plant photosystem I. Nature 426, 630–635.  10.1038/nature02200 PubMed DOI

Bogár F., Bartha F., Násztor Z., Fábián L., Leitgeb B., Dér A. (2014). On the Hofmeister Effect: fluctuations at the protein–water interface and the surface tension. J. Phys. Chem. B 118, 8496–8504.  10.1021/jp502505c PubMed DOI

Bussi Y., Shimoni E., Weiner A., Kapon R., Charuvi D., Nevo R., et al. (2019). Fundamental helical geometry consolidates the plant photosynthetic membrane. Proc. Natl. Acad. Sci. U. S. A. 116, 22366–22375.  10.1073/pnas.1905994116 PubMed DOI PMC

Butt J.-H. (1991). Measuring electrostatic, van der Waals, and hydration forces in electrolyte solutions with an atomic force microscope. Biophys. J. 60, 1438–1444.  10.1016/S0006-3495(91)82180-4 PubMed DOI PMC

Cacace M. G., Landau E. M., Ramsden J. J. (1997). The Hofmeister series: salt and solvent effects on interfacial phenomena. Q. Rev. Biophys. 30, 241–277.  10.1017/s0033583597003363 PubMed DOI

Chow W. S., Kim E.-H., Horton P., Anderson J. M. (2005). Granal stacking of thylakoid membranes in higher plant chloroplasts: the physiochemical forces at work and the functional consequences that ensue. Photochem. Photobiol. Sci. 4, 1081–1090.  10.1039/b507310n PubMed DOI

Collins K. D., Washabaugh M. W. (1985). The Hofmeister effect and the behavior of water at interfaces. Q. Rev. Biophys. 18, 323–422.  10.1017/S0033583500005369 PubMed DOI

Cowley A. C., Fuller N. L., Rand R. P., Parsegian V. A. (1978). Measurement of repulsive forces between charged Phospholipid bilayers. Biochemistry 17, 3163–3168.  10.1021/bi00608a034 PubMed DOI

Cseh Z., Rajagopal S., Tsonev T., Busheva M., Papp E., Garab G. (2000). Thermooptic effect in chloroplast thylakoid membranes. Thermal and light stability of pigment arrays with different levels of structural complexity. Biochemistry 39, 15250–15257.  10.1021/bi001600d PubMed DOI

Daum B., Kühlbrand W. (2011). Electron tomography of plant thylakoid membranes. J. Exp. Bot. 62, 2393–2402.  10.1093/jxb/err034 PubMed DOI

Daum B., Nicastro D., Austin J., McIntosh J. R., Kühlbrandt W. (2010). Arrangement of Photosystem II and ATP synthase in Chloroplast membranes of spinach and pea. Plant Cell 22, 1299–1312.  10.1105/tpc.109.071431 PubMed DOI PMC

Dekker J. P., Boekema E. J. (2005). Supramolecular organization of thylakoid membrane proteins in green plants. Biochim. Biophys. Acta 1706, 12–39.  10.1016/j.bbabio.2004.09.009 PubMed DOI

Dér A., Ramsden J. J. (1998). Evidence for loosening of a protein mechanism. Naturwissenschaften 85, 353–355.  10.1007/s001140050515 DOI

Dér A., Kelemen L., Fábián L., Taneva S. G., Fodor E., Páli T., et al. (2007). Interfacial water structure controls protein conformation. J. Phys. Chem. B 111, 5344–5350.  10.1021/jp066206p PubMed DOI

Garab G., Mustardy L. (1999). Role of LHCII-containing macrodomains in the structure, function and dynamics of grana. Aust. J. Plant Physiol. 26, 647–649. 10.1071/PP99069 DOI

Garab G., van Amerongen H. (2009). Linear dichroism and circular dichroism in photosynthesis research. Photosynth. Res. 101, 135–146.  10.1007/s11120-009-9424-4 PubMed DOI PMC

Garab G., Faludi-Daniel A., Sutherland J. C., Hind G. (1988). Macroorganization of chlorophyll a/b light-harvesting complex in thylakoids and aggregates: information from circular differential scattering. Biochemistry 27, 2425–2430.  10.1021/bi00407a027 DOI

Garab G., Kieleczawa J., Sutherland J. C., Bustamante C., Hind G. (1991). Organization of pigment-protein complexes into macrodamains in the thylakoid membranes of wild-type and chlorophyll fo-less mutant of barley as revealed by circular dicroism. Photochem. Photobiol. 54, 273–281.  10.1111/j.1751-1097.1991.tb02016.x DOI

Garab G., Istokovics A., Butiuc A., Simidjiev I., Dér A. (1998). “Light-induced ion movements in thylakoid membranes and isolated LHC II,” in Photosynthesis: Mechanisms and Effects. Ed. Garab G. (Netherlands: Springer; ), 341–344.

Garab G., Cseh Z., Kovács L., Rajagopal S., Várkonyi Z., Wentworth M., et al. (2002). Light-induced trimer to monomer transition in the main light-harvesting antenna complex of plants:  Thermo-optic mechanism. Biochemistry 41, 15121–15129.  10.1021/bi026157g PubMed DOI

Garab G. (2014). Hierarchical organization and structural flexibility of thylakoid membranes. Biochim. Biophys. Acta 1837, 481–494.  10.1016/j.bbabio.2013.12.003 PubMed DOI

Garab G. (2016). Self-assembly and structural–functional flexibility of oxygenic photosynthetic machineries: personal perspectives. Photosynth. Res. 127, 131–150.  10.1007/s11120-015-0192-z PubMed DOI

Goodchild D., Highkin H. R., Boardman N. K. (1966). Fine structure of chloroplasts in a barley mutant lacking chlorophyll b. Exp. Cell Res. 43, 684–688.  10.1016/0014-4827(66)90045-0 PubMed DOI

Gunning B. E. S., Schwartz O. M. (1999). Confocal microscopy of thylakoid autofluorescence in relation to origin of grana and phylogeny in green algae Aust. J. Plant Physiol. 26, 695–708.  10.1071/PP99076 DOI

Harrison C. J., Morris J. L. (2018). The origin and early evolution of vascular plant shoots and leaves. Phil. Trans. R. Soc B 373, 20160496.  10.1098/rstb.2016.0496 PubMed DOI PMC

Hind G., Nakatani H., Izawa S. (1974). Light-dependent redistribution of ions in suspensions of chloroplast thylakoid membranes. Proc. Natl. Acad. Sci. U. S. A. 71, 1484–1488.  10.1073/pnas.71.4.1484 PubMed DOI PMC

Hind G., Wall J. S., Várkonyi Z., Istokovics A., Lambrev P. A., Garab G. (2014). Membrane crystals of plant light-harvesting complex II disassemble reversibly in light. Plant Cell Physiol. 55, 1296–1303.  10.1093/pcp/pcu064 PubMed DOI PMC

Horton P. (1999). Are grana necessary for regulation of light harvesting? Aust. J. Plant Physiol. 26, 659–669.  10.1071/PP99095 DOI

Iwai M., Pack C. G., Takenaka Y., Sako Y., Nakano A. (2013). Photosystem II antenna phosphorylation-dependent protein diffusion determined by fluorescence correlation spectroscopy. Sci. Rep. 3:2833.  10.1038/srep02833 PubMed DOI PMC

Izawa S., Good N. E. (1966). Effect of salts and electron transport on conformation of isolated chloroplasts. 2. Electron microscopy. Plant Physiol. 41, 544–552.  10.1104/pp.41.3.544 PubMed DOI PMC

Jakubauskas D., Kowalewska Ł., Sokolova A. V., Garvey C. J., Mortensen K., Jensen P. E., et al. (2019). Ultrastructural modeling of small angle scattering from photosynthetic membranes. Sci. Rep. 9, 19405.  10.1038/s41598-019-55423-0 PubMed DOI PMC

Janik E., Bednarska J., Zubik M., Puzio M., Luchowski R., Grudzinski W., et al. (2013). Molecular architecture of plant thylakoids under physiological and light stress conditions: A study of lipid light-harvesting complex II model membranes. Plant Cell 25, 2155–2170.  10.1105/tpc.113.113076 PubMed DOI PMC

Jennings R. C., Gerola P. D., Garlaschi F. M., Forti G. (1981). Effects of trypsin and cations on chloroplast membranes. Plant Physiol. 67, 212–215.  10.1104/pp.67.2.212 PubMed DOI PMC

Jia H., Liggins J., Chow W. (2014). Entropy and biological systems: Experimentally-investigated entropy-driven stacking of plant photosynthetic membranes. Sci. Rep. 4, 4142.  10.1038/srep04142 PubMed DOI PMC

Johnson M. P., Goral T. K., Duffy C. D. P., Brain A. P. R., Mullineaux C. W., Ruban A. V. (2011). Photoprotective energy dissipation involves the reorganization of photosystem II light-harvesting complexes in the grana membranes of spinach chloroplasts. Plant Cell 23, 1468–1479.  10.1105/tpc.110.081646 PubMed DOI PMC

Kana R., Govindjee (2016). Role of ions in the regulation of light-harvesting. Front. Plant Sci. 7, 1849.  10.3389/fpls.2016.01849 PubMed DOI PMC

Keller D., Bustamante C. (1986). Theory of the interaction of light with large inhomogeneous molecular aggregates. II. Psi-type circular dichroism. J. Chem. Phys. 84, 2972–2980.  10.1063/1.450278 DOI

Khoroshyy P., Dér A., Zimányi L. (2013). Effect of Hofmeister Cosolutes on the photocycle of photoactive yellow protein at moderately Alkaline PH. J. Photochem. Photobiol. B Biol. 120, 111–119.  10.1016/j.jphotobiol.2012.12.014 PubMed DOI

Kirchhoff H., Lenhert S., Buchel C., Chi L., Nield J. (2008). Probing the organization of photosystem II in photosynthetic membranes by atomic force microscopy. Biochemistry 47, 431–440.  10.1021/bi7017877 PubMed DOI

Kouřil R., Dekker J. P., Boekema E. J. (2012). Supramolecular organization of photosystem II in green plants. Biochim. Biophys. Acta 1817, 2–12.  10.1016/j.bbabio.2011.05.024 PubMed DOI

Kovacs B., Saftics A., Biro A., Kurunczi S., Szalontai B., Kakasi B., et al. (2018). Kinetics and structure of self-assembled flagellin monolayers onhydrophobic surfaces in the presence of Hofmeister salts: experimental measurement of the protein interfacial tension at the nanometer scale. Phys. Chem. C 122, 21375–21386.  10.1021/acs.jpcc.8b05026 DOI

Kowalewska L., Mazur R., Suski S., Garstka M., Mostowska A. (2016). Three-dimensional visualization of the tubular-lamellar transformation of the internal plastid membrane network during runner bean chloroplast biogenesi ). Plant Cell 28, 875–891.  10.1105/tpc.15.01053 PubMed DOI PMC

Krumova S. B., Laptenok S. P., Kovács L., Tóth T., van Hoek A., Garab G., et al. (2010). Digalactosyl-diacylglycerol-deficiency lowers the thermal stability of thylakoid membranes. Photosynth. Res. 105, 229–242.  10.1007/s11120-010-9581-5 PubMed DOI PMC

Lambrev P. H., Akhtar P. (2019). Macroorganisation and flexibility of thylakoid membranes. Biochem. J. 476, 2981–3018.  10.1042/BCJ20190080 PubMed DOI

Lo Nostro P., Ninham B. W. (2012). Hofmeister Phenomena: an update on ion specificity in Biology. Chem. Rev. 112, 2286.  10.1021/cr200271j PubMed DOI

lsraelachvili J. N., Pashley R. M. (1983). Molecular layering of water at surfaces and origin of repulsive hydration forces. Nature 306, 249–250.  10.1038/306249a0 DOI

Majee A., Bier M., Blossey R., Podgornik R. (2019). Charge regulation radically modifies electrostatics in membrane stacks. Phys. Rev. 100, 1–6.  10.1103/PhysRevE.100.050601 PubMed DOI

Melander W., Horváth C. (1977). Salt Effects on Hydrophobic interactions in precipitation and chromatography of Proteins: an interpretation of the Lyotropic series. Arch. Biochem. Biophys. 183, 200–215.  10.1016/0003-9861(77)90434-9 PubMed DOI

Miller K. R., Staehelin L. A. (1976). Analysis of the thylakoid outer surface. Coupling factor is limited to unstacked membrane regions. J. Cell Biol. 68, 30–47.  10.1083/jcb.68.1.30 PubMed DOI PMC

Miloslavina Y., Lambrev P. H., Jávorfi T., Várkonyi Z., Karlicky V., Wall J. S., et al. (2012). Anisotropic circular dichroism signatures of oriented thylakoid membranes and lamellar aggregates of LHCII. Photosynth. Res. 111, 29–39.  10.1007/s11120-011-9664-y PubMed DOI

Mullineaux C. W. (2005). Function and evolution of grana. Trends Plant Sci. 10, 521–525.  10.1016/j.tplants.2005.09.001 PubMed DOI

Murakami S., Packer L. (1970). Protonation and chloroplast membrane structure. J. Cell Biol. 47, 332–351.  10.1083/jcb.47.2.332 PubMed DOI PMC

Mustárdy L., Garab G. (2003). Granum revisited. A three-dimensional model–where things fall into place. Trends Plant Sci. 8, 117–122.  10.1016/S1360-1385(03)00015-3 PubMed DOI

Mustárdy L., Buttle K., Steinbach G., Garab G. (2008). The three-dimensional network of the thylakoid membranes in plants: quasihelical model of the granum-stroma assembly. Plant Cell 20, 2552–2557.  10.1105/tpc.108.059147 PubMed DOI PMC

Nagy G., Garab G. (2020). Neutron scattering in photosynthesis research. Recent advances and perspectives for testing crop plants. Photosynt. Res. [published online ahead of print, 2020 Jun 2].  10.1007/s11120-020-00763-6 PubMed DOI PMC

Nagy G., Posselt D., Kovács L., Holm J. K., Szabó M., Ughy B., et al. (2011). Reversible membrane-reorganizations during photosynthesis in vivo - revealed by small-angle neutron scattering. Biochem. J. 436, 225–230.  10.1042/BJ20110180 PubMed DOI

Nagy G., Kovács L., Ünnep R., Zsiros O., Almásy L., Rosta L., et al. (2013). Kinetics of structural reorganizations in multilamellar photosynthetic membranes monitored by small-angle neutron scattering. Eur. Phys. J. E 36, 69.  10.1140/epje/i2013-13069-0 PubMed DOI

Násztor Z., Bogár F., Dér A. (2016). The interfacial tension concept, as revealed by fluctuations. Curr. Opin. Colloid Interface Sci. 23, 29–40.  10.1016/j.cocis.2016.05.007 DOI

Násztor Z., Dér A., Bogár F. (2017). Ion-induced alterations of the local hydration environment elucidate Hofmeister effect in a simple classical model of Trp-cage miniprotein. J. Mol. Model. 23, 298.  10.1007/s00894-017-3471-0 PubMed DOI

Neagu A., Neagu M., Dér A. (2001). Fluctuations and the Hofmeister Effect. Biophys. J. 81, 1285–1294.  10.1016/S0006-3495(01)75786-4 PubMed DOI PMC

Nevo R., Charuvi D., Tsabari O., Reich Z. (2012). Composition, architecture and dynamics of the photosynthetic apparatus in higher plants. Plant J. 70, 157–176.  10.1111/j.1365-313X.2011.04876.x PubMed DOI

Paolillo D. J., Jr (1970). The three-dimensional arrangement of intergranal lamellae in chloroplasts. J. Cell Sci. 6, 243–225. PubMed

Petrova N., Stoichev S., Paunov M., Todinova S., Taneva S. G., Krumova S. (2019). Structural organization, thermal stability, and excitation energy utilization of pea thylakoid membranes adapted to low light conditions. Acta Physiol. Plant 41, 188.  10.1007/s11738-019-2979-6 DOI

Posselt D., Nagy G., Kirkensgaard J. J. K., Holm J. K., Aagaard T. H., Timmins P., et al. (2012). Small-angle neutron scattering study of the ultrastructure of chloroplast thylakoid membranes — Periodicity and structural flexibility of the stroma lamellae. Biochim. Biophys. Acta 1817, 1220–1228.  10.1016/j.bbabio.2012.01.012 PubMed DOI

Puthiyaveetil S., Kirchhoff H., Höhner R. (2016). Structural and functional dynamics of the thylakoid membrane system in. Chloroplasts: Current Research and Future Trend. Ed. Kirchhoff H. (Norfolk, UK: Caister Academic Press; ).  10.21775/9781910190470.03 DOI

Puthiyaveetil S., van Oort B., Kirchhoff H. (2017). Surface charge dynamics in photosynthetic membranes and the structural consequences. Nat. Plants 3, 17020.  10.1038/nplants.2017.20 PubMed DOI

Robinson D. R., Jencks W. P. (1965). The effect of concentrated salt solutions on the activity coefficient of acetyltetraglycine ethyl ester. J. Am. Chem. Soc 87, 2470–2479.  10.1021/ja01089a029 PubMed DOI

Rojdestvenski I., Ivanov A. G., Cottam M. G., Borodich A., Huner N. P. A., Öquist G. (2002). Segregation of photosystems in thylakoid membranes as a critical phenomenon. Biophys. J. 82, 1719–1730.  10.1016/S0006-3495(02)75524-0 PubMed DOI PMC

Simidjiev I., Barzda V., Mustárdy L., Garab G. (1997). Isolation of lamellar aggregates of the light-harvesting chlorophylla/b proteincomplex of photosystem II with long-range chiral order and structural flexibility. Anal. Biochem. 250, 169–175.  10.1006/abio.1997.2204 PubMed DOI

Simidjiev I., Stoylova S., Amenitsch H., Jávorfi T., Mustárdy L., Laggner P., et al. (2000). Self-assembly of large, ordered lamellae from nonbilayer lipids and integral membrane-proteins in vitro. Proc. Natl. Acad. Sci. U. S. A. 97, 1473–1476.  10.1073/pnas.97.4.1473 PubMed DOI PMC

Solymosi K., Keresztes Á. (2012). Plastid structure, diversification and interconversions II. Land plants. Curr. Chem. Biol. 6, 187–204.  10.2174/2212796811206030003 DOI

Solymosi K. (2012). Plastid structure, diversification and interconversions I. Algae. Curr. Chem. Biol. 6, 167–186.  10.2174/2212796811206030002 DOI

Standfuss J., van Scheltinga A. C. T., Lamborghini M., Kühlbrandt W. (2005). Mechanisms of photoprotection and nonphotochemical quenching in pea light-harvesting complex at 2.5 Å resolution. EMBO J. 24, 919–928.  10.1038/sj.emboj.7600585 PubMed DOI PMC

Szalontai B., Nagy G., Krumova S., Fodor E., Páli T., Taneva S. G., et al. (2013). Hofmeister ions control protein dynamics. Biochim. Biophys. Acta 1830, 4564–4572.  10.1016/j.bbagen.2013.05.036 PubMed DOI

Tanford C. (1979). Interfacial Free Energy and the Hydrophobic Effect. Proc. Natl. Acad. Sci. U. S. A. 76, 4175–4176.  10.1073/pnas.76.9.4175 PubMed DOI PMC

Tóth T. N., Rai N., Solymosi K., Zsiros O., Schröder W. P., Garab G., et al. (2016). Fingerprinting the macro-organisation of pigment–protein complexes inplant thylakoid membranes in vivo by circular-dichroism spectroscopy. Biochim. Biophys. Acta 1857, 1479–1489.  10.1016/j.bbabio.2016.04.287 PubMed DOI

Ünnep R., Nagy G., Markó M., Garab G. (2014). Monitoring thylakoid ultrastructural changes in vivo using small-angle neutron scattering. Plant Physiol. Biochem. 81, 197–207.  10.1016/j.plaphy.2014.02.005 PubMed DOI

Wood W. H. J., MacGregor-Chatwin C., Barnett S. F. H., Mayneord G. E., Huang X., Hobbs J. K., et al. (2018). Dynamic thylakoid stacking regulates the balance between linear and cyclic photosynthetic electron transfer. Nat. Plants 4, 116–127.  10.1038/s41477-017-0092-7 PubMed DOI

Zer H., Vink M., Keren N., Dilly-Hatrwig H., Paulsen H., Herrmann R. G., et al. (1999). Regulation of thylakoid protein phosphorylation at the substrate level: reversible light-induced conformational changes expose the phosphorylation site of the light-harvesting complex II. Proc. Natl. Acad. Sci. U. S. A. 96, 8277–8282.  10.1073/pnas.96.14.8277 PubMed DOI PMC

Zsiros O., Nagy V., Párducz Á, Nagy G., Ünnep R., El-Ramady H., et al. (2019). Effects of selenate and red Se-nanoparticles on the photosynthetic apparatus of Nicotiana tabacum. Photosynth. Res. 139, 449–460.  10.1007/s11120-018-0599-4 PubMed DOI

Accumulation of geranylgeranylated chlorophylls in the pigment-protein complexes of Arabidopsis thaliana acclimated to green light: effects on the organization of light-harvesting complex II and photosystem II functions