In nature, the intensity of light changes rapidly, spanning from 2000 µmol PHOTONS m-2 s-1 during the brightest sunlight, down to 200-500 µmol PHOTONS m-2 s-1, when shaded by clouds, followed by the period of dark, to encircle with the next sunrise of barely 10 µmol PHOTONS m-2 s-1. Plants, capable of performing limited movements, like leaf and chloroplast movements, respectively, have been forced to evolve different means of coping with the unpredictable nature. To better understand the acclimatization mechanisms, the intensity of light has been roughly divided into low-light and high-light conditions, shaped by the laboratory conditions of plant growth and the strength of the measuring apparatus available. Moreover, looking at the time scale, it is possible to differentiate between short-term and long-term acclimatization processes in plants. These artificial divisions are needed to simplify the most important process in photosynthesis in vascular plants: the rate and the distribution of excitation energy between two photosystems. Photosystem units are organized into large supercomplexes with peripherally attached antenna complexes, being further assembled into megacomplexes . Two types of peripheral antenna proteins associated to photosystem II (PSII) are known: the major Light-Harvesting Complex II (LHCII), which occurs as a trimeric complex containing the proteins Lhcb1, Lhcb2 and Lhcb3, and three minor monomeric complexes, namely Lhcb4 (CP29), Lhcb5 (CP26), and Lhcb6 (CP24). These peripheral complexes bind variable number of molecules of chlorophyll
2. The way the plant protects itself: Non-photochemical quenching
The non-photochemical quenching (NPQ) is a short-term response by which plants harmlessly dissipate excess excitation energy into heat under high-light conditions. NPQ is observed in all higher plants, in lower plants, green algae and diatoms . NPQ is also present in oceanic picophytoplankton species (see Chapter Kulk et al.). Basically, during the absorption of sunlight by light-harvesting complexes (LHCs) associated with reaction centres, a chlorophyll
2.1. ΔpH-dependent quenching
The process of qE is triggered by acidification of the thylakoid lumen under light-saturating conditions, which activates the interconversion of specific xanthophyll pigments (oxygenated carotenoids), that are mostly bound to the LHC proteins. For comparison, in cyanobacteria, strong blue-green or white light activates the orange carotenoid protein (OCP) which interacts with phycobilisome and dissipates the excess energy in the form of heat . The xanthophyll cycle in plants, green and brown algae consists of the pH-dependent conversion from violaxanthin first to antheraxanthin and then to zeaxanthin. In plants, the reactions towards zeaxanthin are catalysed by the enzyme violaxanthin de-epoxidase, while the relatively slow reactions towards violaxanthin are catalysed by the enzyme zeaxanthin epoxidase . The
Due to the slower kinetics of formation and relaxation of qE compared to the proton gradient, and taking into account that the light causes changes in charge distribution, which consequently alter the aggregation state of thylakoids, it was proposed that such a conformational change might accompany the qE event [6, 11]. Indeed, reports from at least three independent laboratories confirmed the structural rearrangement of the PSII-LHCII macro-organization during qE [12-14]. Time-correlated single photon counting (TCSPC) measurements revealed an additional far-red fluorescence component in the leaves of the high-light-adapted wild-type, the mutant unable to accumulate zeaxanthin at high-light (
Acidification of the thylakoid lumen also causes a conformational change in thylakoids that can be monitored at 535 nm (ΔA535). This change is most likely induced by protonation of the lumen-exposed carboxylate side chains in specific PSII proteins . Interestingly,
Although the main components for qE are known, the biophysical mechanism of de-excitation of the excited molecules of chlorophyll
Variations in NPQ capacities and processes in different organisms suggest a strong evolutionary pressure to obtain optimized photoprotection . For example, the LHCII proteins may have evolved from ancestors of contemporary stress-responsive proteins such as HLIP and ELIPS, which probably bind only carotenoids and are involved in photoprotection, while LHCII harvest light through high amounts of chlorophyll binding. In plants, PsbS binds pigments minimally and it is primarily involved in photoprotection, while the light-harvesting complex containing fucoxanthin (LHCF) in diatoms binds both chlorophyll and carotenoids in high amounts and contributes equally to the light harvesting and photoprotection. Under high-light conditions, the photosynthetic reaction centres can not accommodate all the electrons coming through electron transport chain, thus entering into the saturated, “closed” status. The already generated excitation energy becomes the burden for the photosynthetic membranes and has to be channeled safely, before destroying the reaction centres, especially the pigment core of PSII, the pair of the most potent oxidizers known to exist in nature, P680. If the vast energy is not diverted, P680 might rest in its prolonged oxidized state, P680+, and oxidize the neighbour protein amino-acids and pigments, ultimately leading to the destruction of the PSII protein D1. However, if P680 can not submit the electron to the oxidized plastoquinone, due to the increased number of already reduced plastoqinones, P680 could go into triplet state, interacting with atmospheric triplet oxygen and producing deleterious singlet oxygen. Both of these processes lead to the photoinhibition, the state in which the decreasing of the electron transport is observed . Photoinhibition has a specific signature that can be monitored by a lower oxygen production, analysis of the D1 protein level, formation of uncoupled chlorophyll and of triplet state of chlorophyll. However, photoinhibition provokes the formation of photoprotection processes, where most of the unwanted energy would be harmlessly dissipated as heat. This phenomenon leaves a palpable trace visible as a drop in the fluorescence intensity, measured by TCSPC. One of the approaches to study the origin of the quenching mechanism
2.2. How does the plant repair the damaged D1 protein?
The D1 repair cycle is regulated
3. Plastoquinone pool-inflicted plant responses
The most promising redox-active components are the pool of plastoquinone (PQ) and the PSI acceptor site (
State-transitions re-distribute excitation energy between two photosystems, which are electrochemically connected in series, through the supramolecular reorganisation of the photosynthetic membranes. The molecular complexes that cause the reorganisation are light-harvesting proteins, which collect light excitation and channel it to the reaction centres. Already mentioned LHCII is the prototype of large and abundant class of chloroplast transmembrane proteins that binds roughly half of the total chlorophyll in chloroplasts . The absorption spectra of LHCI is in the far red region owing to the chlorophyll
3.2. Stoichiometry adjustments
Photosystem stoichiometry adjustment is a long-term response that affects the relative amounts of the two photosystems by changing the expression of photosynthetic genes both in the chloroplast and in the nucleus . It was shown that the redox state of the PQ pool serves as a major signal in the regulation of LHCII photo-acclimation . In order to avoid excess excitation energy and to minimize oxidative damage, the LHCII protein level decreased by approximately 50% in
4. Immunophilins in photosynthesis
Immunophilins comprise a superfamily of conserved ubiquitous proteins consisting of two distinct subfamilies; cyclophilins and FKBPs (FK506/rapamycin-binding proteins), the targets of the immunosuppressive drugs cyclosporine A (CsA) and FK506/rapamycin, respectively . Despite the lack of structural similarity, all cyclophilins and FKBPs share a common enzymatic, so-called PPIase or rotamase activity, catalizing
Complex immunophilins all contain additional protein-protein interaction domains, such as leucine-zipper motifs and/or tetratricopeptide repeat domains. They may be constituents of supramolecular structures, as shown for the human or avian steroid receptor complex  or chaperone supercomplexes , and may be involved in hsp90-dependent signal transduction [49, 51, 52]. Various members of the immunophilins are involved in phosphorylation processes
Plants possess a substantial number of immunophilins which are localized in cytosol, chloroplasts, nucleus, mitochondria, or associated with secretory pathways. The majority of the cyclophilins are single-domain proteins, 23 isoforms in Arabidopsis . AtCYP20-3, AtCYP20-2, AtCYP26-2, AtCYP28 and AtCYP37 can be found in chloroplasts, mostly in thylakoid lumen. The multidomain cyclophilin isoforms possess unique domain arrangements, as exemplified by AtCYP38. AtCYP40 contains a C-terminal tetratricopeptide repeat module (TPR). Four additional multidomain cyclophilins contain RNA interaction domains, which implicates their involvement in nuclear RNA processing machinery.
Plant FKBPs encompass also 23 members in Arabidopsis and this family is one of the largest FKBP family identified to date. These FKBPs can also be divided into single and multidomain isoforms, consisting of 16 and 7 members, respectively . Interestingly, 11 single-domain FKBPs appear to be targeted to the thylakoid lumen. Thus, this chloroplast sub-compartment appears to have very important role in immunophilin function, or the processes taking place in lumen require specific activity of this protein family.
The dephosphorylation of D1, D2 and CP43 in spinach is catalysed by a cyclophilin-regulated PP2A-like protein phosphatase, which was found to be associated with, and regulated by, a cyclophilin-like protein, TLP40 . TLP40 is proposed to suppress phosphatase activity, when bound to the lumen-exposed epitope of the protein phosphatase, but to induce its activity, when released to the lumen . However, it remains to be seen if the Arabidopsis PP2C phosphatase PBCP  is also under the control of TLP40. Vener group demonstrated that D1 was not only vulnerable to light, but also to high temperature . Raising the temperature from 22 °C to 42 °C resulted in a very rapid dephosphorylation of the D1, D2 and CP43 proteins and in release of TLP40 from membrane into the thylakoid lumen. These events are proposed to trigger an accelerated repair of photodamaged PSII and to initiate other heat-shock responses in chloroplasts.
Higher plant thylakoid lumen prolineisomerases , or complex immunophilins, are found to be regulated by light and are responsive to various forms of environmental stress . The structure of TLP40 and its association with the thylakoid membrane system implicates diverse functions and involvement in the intracellular signaling networks . Binding of thylakoid membrane associated phosphatase involved in dephosphorylation of PSII core proteins might occur
5. Flow and partitioning of photosynthetic electrons
Photosynthetic apparatus has to be able to efficiently convert energy at low light and to avoid over-reduction and damage at excess light. This requires switching between different regulatory mechanisms which keep the cellular ATP pool almost at the constant level. Exposure of plants to higher light intensities than required for efficient photosynthesis results in saturation of photosynthetic electron transport (PET). The over-reduction of PET chain can lead to the formation of reactive oxygen species and may irreversibly damage the photosystems, as well as the cells and the whole organism [71, 72, 73]. To counteract and reduce the photoinhibitory damage, plants have developed several short- and long-term regulation mechanisms which include processes that modulate the structure and function of antenna complexes, including NPQ, alternative electron transport pathways and movement of chloroplasts, leaves and whole organisms away from intense light [74, 75]. Dissipation of excess light energy to heat in the antenna or in the reaction centre of PSII also counteracts the photoinhibitory damage. Retrograde signaling transduces information on the metabolic state of the organelle and induces many activities in the cytosol, nucleus and mitochondria, inducing alterations in nuclear gene expression of organelle-targeted proteins .
5.1. Linear, cyclic and pseudo-cyclic electron transfer routes
Various photosynthetic electron transfer routes become turned on and off, according to the need of the plant to adapt to wide-ranging quantities and qualities of light. Three major electron transfer pathways known are linear, cyclic and pseudo-cyclic electron transfer (LEF, CEF and PCEF, respectively). All three pathways are necessary for poised and sustained synthesis of ATP and NADPH and their interplay enables the flexibility of photosynthesis in meeting different metabolic demands .
During non-cyclic electron transport or LEF, light drives the conversion of water to oxygen at the level of oxygen-evolving complex of PSII and NADP+ to NADPH on the stromal side of thylakoid membranes. The PET chain consists of PSII, the Cyt
Arnon, who discovered photophosphorylation in isolated chloroplasts in 1954, demonstrated that there is also CEF in the thylakoids, driven solely by PSI. CEF is a light-driven flow of electrons through a photosynthetic reaction centre with the electrons being transferred from PSI to Cyt
CEF around PSI occurs under conditions when acceptor limitation or ATP shortage results in a highly reduced PQ pool and contributes to the formation and maintenance of a pH gradient across a membrane but does not produce NADPH. The pH gradient generated may drive the production of ATP (cyclic photophosphorylation) or may regulate photosynthesis. When more light is absorbed than can be used for assimilation, the increased ∆pH is a switch to dissipate excess of the light absorbed by chlorophyll molecules of PSII . CEF is diminished when its components are completely reduced. Also, there is no CEF when its components are completely oxidized because there are no electrons to cycle . In an attempt to avoid these two extreme situations, kinetics, post-translational modifications [25, 26, 81] and redox control of reaction-centre gene expression  are all employed in maintaining a poised PQ pool. In spite of these control mechanisms, over-reduction easily occurs when the Calvin–Benson cycle is unable to use NADPH, usually due to the lack of ATP. The major physiological significance of CET most probably lies in additional availability of ATP.
LEF in chloroplasts produces a number of reduced components associated with PSI that may subsequently participate in reactions that reduce oxygen. When Fd transfers electrons to molecular oxygen instead of NADP+, PCEF linked with phosphorylation arises. O2 is directly reduced to superoxide radical in so-called Mehler reaction. Subsequently, two superoxides dismutate to form H2O2 and O2, which are by further redox processes converted to water. A reduced state of the FeS pool (Fd and PSI centres) promotes the Mehler reaction. The Mehler reaction leading to PCEF restores the redox poise when the PET chain is over-reduced, thereby allowing CEF to function and to generate ATP for the Calvin-Benson cycle, which will in turn oxidize NADPH and restore LEF . Under high light PCEF could also cover an increased energy demand, as long as the antioxidant systems for H2O2 removal is sufficiently active (ascorbate and GSH recycling).
A great number of contemporary research topics on photosynthesis aims at elucidating novel, alternative electron transfer routes as the safest pathways for channelling of unwanted electrons. Chlororespiration is a well-known respiratory process that can maintain a trans-thylakoid proton gradient, thus acting as an effective alternative electron sink in preventing over-reduction of the PQ pool and protecting RCIIs from photodamage under photoinhibitory light conditions . Two enzymes are important for chlororespiratory function: NADH dehydrogenase complex and nucleus-encoded plastid-localized terminal oxidase (PTOX), through which electrons from plastoquinol are transferred to molecular oxygen, forming water in the stroma . In oat leaves incubated at high temperature and under high-light intensities, the amounts of both enzymes were increased, suggesting that, under unfavourable conditions, chlororespiration can act as a protective mechanism .
Apart from the Mehler reaction and the chlororespiration, respectively, photorespiration is another efficient mechanism to adjust the ATP/NADPH ratio and to consume excess energy . Although the major part of energy supplied in the form of ATP and NADPH by the light reaction is consumed in the Calvin-Benson cycle, it is also needed for multiple anabolic processes in chloroplasts, such as synthesis of lipids and proteins and of many secondary metabolites. ATP/NADPH ratio in chloroplasts could be increased by indirect export of reducing equivalents in the form of malate through the malate valve . Malate can be used in numerous ways in cytoplasm and mitochondria, providing NADH for nitrate reduction and/or ATP for sucrose synthesis. Also, different stages of tissue growth require different ATP/NADPH ratios. The ATP/NADPH ratio that is available in the light can vary substantially because of the many possibilities for electron pathways and regulatory mechanisms . Therefore, the cooperation of different electron transport pathways enables optimization of ATP/NADPH stoichiometry . In example, sudden dark-to-light transition induces over-reduction of the PET chain in the first minute, which is relieved by electron transport to O2 , and by the rapid activation of the chloroplast NADP–MDH . Calculating the photosynthetic stoichiometries, it was estimated that ATP/NADPH ratio arising from LEF is about 1.28, which is not sufficient for driving the Calvin-Benson cycle [91, 77]. It is therefore obvious that the optimal operation of the Calvin-Benson cycle requires both LEF and CEF, whose tuning enables adjustment of ATP/NADPH to meet the cellular demands. Green alga
5.2. Electron partitioning at the ferredoxin hub
In photoautotrophic plants, ferredoxin (Fd) accepts one electron from the stromal side of PSI involving the subunits PsaC, PsaD and PsaE . Fd acts simultaneously as bottleneck and as a hub which distributes high-energy electrons to a multitude of enzymes involved in chloroplast metabolism. The hierarchy of electron distribution and subsequent regulation of channeling of photosynthetically derived electrons into different areas of chloroplast metabolism is still not defined. Electrons are preferentially directed to carbon assimilation, which requires NADPH, and so the majority of Fd is immediately oxidized by the enzyme ferredoxin:NADPH reductase (FNR), which is associated with the thylakoid membrane.
Besides its crucial metabolic role in reducing NADP+ and thioredoxin (TRX)
At least four Fd isoforms occur in plants . Fd1 and Fd2 are found in leaves, while Fd3 and Fd4 appear to play roles in non-photosynthetic metabolism [101, 100]. Fd as electron distributing hub is particularly suitable to provide information on the redox state of the system to be transmitted into the regulatory network. Fd contributes to the control of chloroplast energization by feeding electrons into the CEF pathway and thereby controls both phosphorylation potential and reductive power.
5.3. TROL-FNR interaction influences the energy conductance
The last step of the photosynthetic electron transfer from Fd to NADP+ is catalyzed by FNR. There are two evolutionary conserved types of FNR in the chloroplasts of higher plants: predominantly, or exclusively, thylakoid membrane-bound isoproteins and ‘soluble’, non-tightly bound isoproteins [101, 102]. The membrane-bound FNR is supposed to be involved in electron transport, while the soluble enzyme provides protection against oxidative stress . Two chloroplast-type
NADP+ photoreduction activity of FNR was shown to be greater when the enzyme is associated with the thylakoid membrane and it has been proposed that binding of FNR to the thylakoid membrane regulates the enzyme activity [105, 106]. Subsequently, interactions of FNR and several photosynthetic protein complexes, such as Cyt
It was also demonstrated that FNR interacts specifically with two chloroplast proteins, Tic62 (62 kDa component of the translocon at the inner envelope of cloroplasts) and TROL (thylakoid rhodanese-like protein),
We have already mentioned that reducing equivalents could be exported in the form of malate through the malate valve to increase the ATP/NADPH ratio in chloroplasts . TROL-deficient plants grown under growth-light conditions show significant up-regulation of NADP-malic enzyme 2 that catalyses the oxidative decarboxylation of malate, producing pyruvate, carbon dioxide and NAD(P)H in cytosol . Therefore, the TROL knock-out Arabidopsis mutant lines (
6. Conclusions and perspectives
Photosynthesis is the crucial converter of sunlight into the chemical energy that is subsequently utilised to sustain life on Earth. However, without the layers of regulative pathways, it would be virtually impossible to discuss the efficient photosynthesis. Now, we are aware of the fact that more than 40-years-old “Z-scheme” has been constantly upgraded with many alternative routes allied with still not well-characterised electron sources and sinks. In addition, it seems that the thylakoid membranes are more flexible than anticipated, with PSII antenna involved directly in NPQ origin through detachment and aggregation.
One of the interesting domains in photosynthesis, but not enough explored, would be the involvement of lipids in a complex network of protein interactions. Lipids, especially phospholipids (phosphatidylglycerol; PG) and glycolipids (monogalactosyldiacylglycerol; MGDG, digalactosyldiacylglycerol; DGDG, and sulfoquinovosyldiacylglycerol; SQDG) are major building blocks of thylakoid membranes, providing the safe docking sites for proteins and protein complexes, respectively. It has been proposed that DGDG and PG are especially involved in the assembly and repair of PSII . Moreover, plants lacking almost 30% of the wild-type amount of PG, caused by the point mutation in one of the genes involved in PG synthesis pathway, displayed pale green leaves and somewhat reduced capacity for photosynthesis , while plants that accumulated only 10% of the wild-type PG amount were incapable of surviving on the growth medium without the addition of sucrose .
The other attractive domain would be retrograde signalling,
The rich past of exploring photosynthetic processes gave us a wealth of knowledge, but, it seems that we have just scratched the surface. In years to come, it would be interesting to fill the missing blanks and to develop some new, “out-of-the-box” perspectives.
Dekker JP, Boekema EJ. Supramolecular organization of thylakoid membrane proteins in green plants. Biochim Biophys Acta. 2005;1706(1-2):12-39.
Barros T, Kühlbrandt W. Crystallisation, structure and function of plant light-harvesting Complex II. Biochim Biophys Acta. 2009;1787(6):753-772.
Horton P, Ruban A. Molecular design of the photosystem II light-harvesting antenna: photosynthesis and photoprotection. J Exp Bot. 2005;56(411):365-373.
Müller P, Li XP, Niyogi KK. Non-photochemical quenching. A response to excess light energy. Plant Physiol. 2001;125(4): 1558-1566.
Szabó I, Bergantino E, Giacometti GM. Light and oxygenic photosynthesis: energy dissipation as a protection mechanism against photo-oxidation. EMBO Rep. 2005;6(7):629-634.
Ruban AV, Johnson MP, Duffy CD. The photoprotective molecular switch in the photosystem II antenna. Biochim Biophys Acta. 2012;1817(1):167-181.
Eberhard S, Finazzi G, Wollman FA. The dynamics of photosynthesis. Annu Rev Genet. 2008; 42:463-515.
Kirilovsky D, Kerfeld CA. The orange carotenoid protein in photoprotection of photosystem II in cyanobacteria. Biochim Biophys Acta. 2012;1817(1):158-166.
Xiao FG, Shen L, Ji HF. On photoprotective mechanisms of carotenoids in light harvesting complex. Biochem Biophys Res Commun. 2011;414(1):1-4.
Jahns P, Holzwarth AR. The role of the xanthophyll cycle and of lutein in photoprotection of photosystem II. Biochim Biophys Acta. 2012;1817(1):182-193.
Barber J. Influence of surface charges on thylakoid structure and function. Annu Rev Plant Physiol. 1982;33:261-295.
Holzwarth AR, Miloslavina Y, Nilkens M, Jahns P. Identification of two quenching sites active in the regulation of photosynthetic light-harvesting studied by time-resolved fluorescence. Chem Phys Lett. 2009;483(4-6):262-267.
Betterle N, Ballottari M, Zorzan S, de Bianchi S, Cazzaniga S, Dall'osto L, Morosinotto T, Bassi R. Light-induced dissociation of an antenna hetero-oligomer is needed for non-photochemical quenching induction. J Biol Chem. 2009;284(22):15255-15266.
Johnson MP, Goral TK, Duffy CD, Brain AP, Mullineaux CW, Ruban AV. Photoprotective energy dissipation involves the reorganization of photosystem II light-harvesting complexes in the grana membranes of spinach chloroplasts. Plant Cell. 2011;23(4):1468-1479.
Li Z, Wakao S, Fischer BB, Niyogi KK. Sensing and responding to excess light. Annu Rev Plant Biol. 2009;60:239-260.
Horton P, Ruban AV, Rees D, Pascal AA, Noctor G, Young AJ. Control of the light-harvesting function of chloroplast membranes by aggregation of the LHCII chlorophyll-protein complex. FEBS Lett. 1991;292(1-2):1-4.
Li XP, Björkman O, Shih C, Grossman AR, Rosenquist M, Jansson S, Niyogi KK. A pigment-binding protein essential for regulation of photosynthetic light harvesting. Nature. 2000;403(6768):391-395.
Goral TK, Johnson MP, Duffy CD, Brain AP, Ruban AV, Mullineaux CW. Light-harvesting antenna composition controls the macrostructure and dynamics of thylakoid membranes in Arabidopsis. Plant J. 2012;69(2):289-301.
Ahn TK, Avenson TJ, Ballottari M, Cheng YC, Niyogi KK, Bassi R, Fleming GR. Architecture of a charge-transfer state regulating light harvesting in a plant antenna protein. Science. 2008;320(5877):794-797.
Belgio E, Johnson MP, Jurić S, Ruban AV. Higher plant photosystem II light-harvesting antenna, not the reaction center, determines the excited-state lifetime-both the maximum and the nonphotochemically quenched. Biophys J. 2012;102(12):2761-2771.
Tikkanen M, Nurmi M, Kangasjärvi S, Aro EM. Core protein phosphorylation facilitates the repair of photodamaged photosystem II at high light. Biochim Biophys Acta. 2008;1777(11):1432-1437.
Vener AV. Environmentally modulated phosphorylation and dynamics of proteins in photosynthetic membranes. Biochim Biophys Acta. 2007;1767(6):449-457.
Kato Y, Sakamoto W. Protein quality control in chloroplasts: a current model of D1 protein degradation in the photosystem II repair cycle. J Biochem. 2009;146(4):463-469.
Kato Y, Sun X, Zhang L, Sakamoto W. Cooperative D1 Degradation in the Photosystem II Repair Mediated by Chloroplastic Proteases in Arabidopsis. Plant Physiol. 2012;159(4):1428-1439.
Allen JF, Forsberg J. Molecular recognition in thylakoid structure and function. Trends Plant Sci. 2001;6(7):317-326.
Wollman FA. State-transitions reveal the dynamics and flexibility of the photosynthetic apparatus. EMBO J. 2001;20(14):3623-3630.
Mullineaux CW, Emlyn-Jones D. State-transitions: an example of acclimation to low-light stress. J Exp Bot. 2004;56(411):389-393.
Allen JF, Pfannschmidt T. Balancing the two photosystems: photosynthetic electron transfer governs transcription of reaction centre genes in chloroplasts. Philos Trans R Soc Lond B Biol Sci. 2000; 355(1402):1351-1359.
Pfannschmidt T, Bräutigam K, Wagner R, Dietzel L, Schröter Y, Steiner S, Nykytenko A. Potential regulation of gene expression in photosynthetic cells by redox and energy state: approaches towards better understanding. Ann Bot. 2009;103(4):599-607.
Iwai M, Yokono M, Inada N, Minagawa J. Live-cell imaging of photosystem II antenna dissociation during state transitions. Proc Natl Acad Sci USA. 2010;107(5):2337-2342.
Depège N, Bellafiore S, Rochaix JD. Role of chloroplast protein kinase Stt7 in LHCII phosphorylation and state transition in Chlamydomonas. Science. 2003;299(5612):1572-1575.
Bellafiore S, Barneche F, Peltier G, Rochaix JD. State-transitions and light adaptation require chloroplast thylakoid protein kinase STN7. Nature. 2005;433(7028):892-895.
Shapiguzov A, Ingelsson B, Samol I, Andres C, Kessler F, Rochaix JD, Vener AV, Goldschmidt-Clermont M. The PPH1 phosphatase is specifically involved in LHCII dephosphorylation and state-transitions in Arabidopsis. Proc Natl Acad Sci USA. 2010;107(10):4782-4787.
Pribil M, Pesaresi P, Hertle A, Barbato R, Leister D. Role of plastid protein phosphatase TAP38 in LHCII dephosphorylation and thylakoid electron flow. PLoS Biol. 2010;8(1):e1000288.
Samol I, Shapiguzov A, Ingelsson B, Fucile G, Crèvecoeur M, Vener AV, Rochaix JD, Goldschmidt-Clermont M. Identification of a Photosystem II Phosphatase Involved in Light Acclimation in Arabidopsis. Plant Cell. 2012;24(6):2596-2609.
Bonardi V, Pesaresi P, Becker T, Schleiff E, Wagner R, Pfannschmidt T, Jahns P, Leister D. Photosystem II core phosphorylation and photosynthetic acclimation require two different protein kinases. Nature. 2005;437(7062):1179-1182.
Oelze ML, Kandlbinder A, Dietz KJ. Redox regulation and overreduction control in the photosynthesizing cell: complexity in redox regulatory networks. Biochim Biophys Acta. 2008;1780(11):1261-1272.
Schreiber SL. Chemistry and biology of the immunophilins and their immunosuppressive ligands. Science. 1991;251 (4991):283-287.
Fischer G, Wittmann-Liebold B, Lang K, Kiefhaber T, Schmid FX. Cyclophilin and peptidyl-prolyl cis-transisomerase are probably identical proteins. Nature. 1989;337(6206):476-478.
Takahashi N, Hayano T, Suzuki M. Peptidyl-prolyl cis-transisomerase is the cyclosporin A-binding protein cyclophilin. Nature. 1989;337(6206):473-475.
Rudd KE, Sofia HJ, Koonin EV, Plunkett GI, Lazar S, Rouviere PE. A new family of peptidyl-prolyl isomerases. Trends Biochem Sci. 1995;20(1):12-14.
Gething MJ, Sambrook J. Protein folding in the cell. Nature. 1992;355(6355):33-45.
Kieffer LJ, Seng TW, Li W, Osterman DG, Handschumacher RE, Bayney R. Cyclophilin-40, a protein with homology to the P59 component of the steroid receptor complexx. Cloning of the cDNA and further characterization. J Biol Chem. 1993;268(17):12303-12310.
Fruman DA, Burakoff SJ, Bierer BE. Immunophilins in protein folding and immunosuppression. FASEB J. 1994;8(6):391-400.
Matouschek A, Rospert S, Schmid K, Click BS, Schatz G. Cyclophilin catalyzes protein folding in yeast mitochondria. Proc Natl Acad Sci USA. 1995;92(14):6319-6323.
Freskgård PO, Bergenhem N, Jonson BH, Svensson M, Carlsson U. Isomerase and chaperone activity of prolyl isomerase in folding of carbonic anhydrase. Science. 1992;258(5081):466-468.
Chang HC, Lindquist S. Conservation of Hsp90 macromolecular complex in Sacharomyces cerevisiae. J Biol Chem. 1994;269(40):24983-24988.
Johnson JL, Toft DO. A novel chaperone complex for steroid receptors involving heat shock proteins, immunophilins, and p23. J Biol Chem. 1994;269(40):24989-24994.
Duina AA, Chang HC, Marsh JA, Lindquist S, Gaber RF. A cyclophilin function in Hsp90-dependent signal transduction. Science. 1996;274(5293):1713-1715.
Freeman BC, Toft DO, Morimoto RI. Molecular chaperone machines: Chaperone activities of the cyclophilin Cyp-40 and the steroid aporeceptors-associated protein p23. Science. 1996;274(5293):1718-1720.
Coss MC, Stephens RM, Morison DK, Winterstein D, Smith LM, Simek SL. The immunophilin FKBP65 forms an association with the serine/threonine kinase c-Raf-1. Cell Growth Diff. 1998;9(1):41-48.
Pratt WB. The hsp90-based chaperone system-involvement in signal transduction from variety of hormone and growth factor receptors. Proc Soc Exp Biol Med. 1998;217(4):420-434.
Owens-Grillo JK, Hoffmann K, Hutchison KA, Yem AW, Deibel MR Jr, Handschumacher RE, Pratt WB. The cyclosporin A-binding immunophilin CyP-40 and the FK506-binding immunophilin hsp56 bind to a common site on hsp90 and exist in independent cytosolic heterocomplexes with the untransformed glucocorticoid receptor. J Biol Chem. 1995;270(35):20479-20484.
Schreiber SL. Immunophilin-sensitive protein phosphatase action in cell signaling pathways. Cell. 1992;70(3):365-368.
Wilson LK, Benton BM, Zhou S, Thorner J, Martin GS. The yeast immunophilin Fpr3 is a physiological substrate of the tyrosine-specific phosphoprotein phosphatas Ptp1. J Biol Chem. 1995;270(42):25185-25193.
Silverstein AM, Galigniana MD, Mei-Shya C, Owens-Grillo JK, Chinkers M, Pratt WB. Protein phosphatase 5 is a major component of glucocorticoid receptor-hsp90 complex with properties of an FK606-binding immunophilin. J Biol Chem. 1997;271(26):16224-16230.
Romano P, Gray J., Horton P, Luan S. Plant immunophilins: functional versatility beyond protein maturation. New Phytologist. 2005;166(3):753-769.
He Z, Li L, Luan S. Immunophilins and parvulins. Superfamily of peptidyl prolyl isomerases in Arabidopsis. Plant Physiol. 2004;134(4):1248-1267.
Vener AV, Rokka A, Fulgosi H, Andersson B, Herrmann RG. A cyclophilin-regulated PP2A-like protein phosphatase in thylakoid membranes of plant chloroplasts. Biochemistry. 1999;38(45):14955-14965.
Rokka A, Aro EM, Herrmann RG, Andersson B, Vener AV. Dephosphorylation of photosystem II reaction center proteins in plant photosynthetic membranes as an immediate response to abrupt elevation of temperature. Plant Physiol. 2000;123(4):1525-1536.
Luan S, Albers MW, Schreiber SL. Light-regulated, tissue-specific immunophilins in a higher plant. Proc Natl Acad Sci USA. 1994;91(3):984-988.
Fulgosi H, Vener AV, Altschmied L, Hermann RG, Andersson B. A novel multi-functional chloroplast protein: identification of 1 40 kDaimmunophilin-like protein located in the thylakoid lumen. EMBO J. 1998;17(6):1577-1587.
Baena-Gonzàlez E, Aro EM. Biogenesis, assembly and turnover of photosystem II units. Philos Trans R Soc Lond B Biol Sci. 2002;357(1426):1451–1460.
Aro EM, Suorsa M, Rokka A, Allahverdiyeva Y, Paakkarinen V, Saleem A, Battchikova N, Rintamäki E. Dynamics of photosystem II: A proteomic approach to thylakoid protein complexes. J Exp Bot. 2005;56(411):347-356.
Shapiguzov A, Edvardsson A, Vener AV. Profound redox sensitivity of peptidyl-prolylisomerase activity in arabidopsis thylakoid lumen. FEBS Lett. 2006;580(15):3671-3676.
Edvardsson A, Shapiguzov A, Petersson UA, Schroder WP, Vener AV. Immunophilin AtFKBP13 sustains all peptidyl–prolylisomerase activity in the thylakoid lumen from Arabidopsis thaliana deficient in AtCYP20-2. Biochemistry. 2007;46(33):9432-9442.
Sirpiö S, Khrouchtchova A, Allaverdiyeva Y, Hansson M, Fristedt R, Vener AV, Scheller HV, Jensen PE, Haldrup A, Aro EM. AtCYP38 ensures early biogenesis, correct assembly and sustenance of photosystem II. Plant J. 2008;55(4):639-651.
Fu A, He Z, Sun Cho H, Lima A, Buchanan BB, Luan S. A chloroplast cyclophilin functions in the assembly and maintenance of photosystem II in Arabidopsis thaliana. Proc Natl Acad Sci USA. 2007;104(40):15947-15952.
Lepeduš H, Tomašić A, Jurić S, Katanić Z, Cesar V, Fulgosi H. Photochemistry of PSII in CYP38 Arabidopsis thaliana Mutant. Food Technol Biotechnol. 2009;47(3):275-280.
Schreiber U, Bilger W, Neubauer C Schreiber U, Bilger W, Neubauer C. Chlorophyll fluorescence as a nonintrusive indicator for rapid assessment of in vivo photosynthesis. In: Schulze E-D, Caldwell MM. (eds.) Ecophysiology of Photosynthesis. Vol 100. Springer: Berlin Heidelberg New York; 1994. p49-70.
Asada K. Production and scavenging of reactive oxygen species in chloroplasts and their functions. Plant Physiol. 2006;141(2):391–396.
Sonoike K. Photoinhibition of photosystem I. Physiol Plant. 2011;142(1):56–64.
Vass I. Molecular mechanisms of photodamage in the Photosystem II complex. Biochim Biophys Acta. 2012;1817(1):209–217.
Baker NR, Harbinson J, Kramer DM. Determining the limitations and regulation of photosynthetic energy transduction in leaves. Plant Cell Environ. 2007;30(9):1107–1125.
Rochaix JD. Reprint of: Regulation of photosynthetic electron transport. Biochim Biophys Acta. 2011; 1807(8):375–383.
Barajas-López JD, Blanco NE, Strand A. Plastid-to-nucleus communication, signals controlling the running of the plant cell. Biochim Biophys Acta. 2013;1833(2):425-437.
Allen JF. Cyclic, pseudocyclic and noncyclic photophosphorylation: new links in the chain. Trends Plant Sci. 2003;8(1):15–19.
Vojta L, Fulgosi H. Energy Conductance from Thylakoid Complexes to Stromal Reducing Equivalents, Advances in Photosynthesis - Fundamental Aspects, Dr Mohammad Najafpour (Ed.), ISBN: 978-953-307-928-8, InTech, 2012, Available from: http://www.intechopen.com/books/advances-in-photosynthesisfundamental-aspects/energy-conductance-from-thylakoid-complexes-to-stromal-reducing-equivalents
Heber U, Walker D. Concerning a dual function of coupled cyclic electron transport in leaves. Plant Physiol. 1992;100(4):1621-1626.
Whatley FR. Photosynthesis by isolated chloroplasts: the early work in Berkeley. Photosynth Res. 1995;46(1-2):17–26.
Finazzi G, Rappaport F, Furia A, Fleischmann M, Rochaix JD, Zito F, Forti G. Involvement of state transitions in the switch between linear and cyclic electron flow in Chlamydomonasreinhardtii. EMBO Rep. 2002;3(3):280–285.
Pfannschmidt T, Nilsson A, Allen JF. Photosynthetic control of chloroplast gene expression. Nature. 1999;39:625-628.
Ort DR, Baker NR. A photoprotective role for O(2) as an alternative electron sink in photosynthesis? Curr Opin Plant Biol. 2002;5(3):193-198.
Ivanov AG, Sane PV, Hurry V, Oquist G, Huner NP. Photosystem II reaction centre quenching: mechanisms and physiological role. Photosynth Res. 2008;98(1-3):565-574.
Quiles MJ. Stimulation of chlororespiration by heat and high light intensity in oat plants. Plant Cell Environ. 2006;29(8):1463-1470.
Scheibe R, Dietz KJ. Reduction-oxidation network for flexible adjustment of cellular metabolism in photoautotrophic cells. Plant Cell Environ. 2012;35(2):202-216.
Scheibe R. Malate valves to balance cellular energy supply. Physiol Plant. 2004;120(1):21-26.
Kramer DM, Evans JR. The importance of energy balance in improving photosynthetic productivity. Plant Physiol. 2011;155(1):70-78.
Steiger HM, Beck E. Formation of hydrogen peroxide and oxygen dependence of photosynthetic CO2 assimilation by intact chloroplasts. Plant Cell Physiol. 1981;22: 561-576.
Scheibe R, Stitt M. Comparison of NADP-malate dehydrogenase activation, QA reduction and O2 evolution in spinach leaves. Plant Physiol Biochem. 1988;26:473–481.
Joliot P, Joliot A. Cyclic electron transfer in plant leaf. Proc Natl Acad Sci USA. 2002;99(15):10209-10214.
Hemschemeier A, Happe T. Alternative photosynthetic electron transport pathways during anaerobiosis in the green alga Chlamydomonas reinhardtii. Biochim Biophys Acta. 2011;1807(8):919-926.
Sétif P, Fischer N, Lagoutte B, Bottin H, Rochaix JD. The ferredoxin docking site of photosystem I. Biochim Biophys Acta. 2002;1555(1-3):204-209.
Zhang L, Happe T, Melis A. Biochemical and morphological characterization of sulfur-deprived and H2-producing Chlamydomonas reinhardtii (green alga). Planta. 2002;214(4):552-561.
Tolleter D, Ghysels B, Alric J, Petroutsos D, Tolstygina I, Krawietz D, Happe T, Auroy P, Adriano JM, Beyly A, Cuiné S, Plet J, Reiter IM, Genty B, Cournac L, Hippler M, Peltier G. Control of hydrogen photoproduction by the proton gradient generated by cyclic electronflow in Chlamydomonas reinhardtii. Plant Cell. 2011;23(7):2619-2630.
Munekage Y, Hojo M, Meurer J, Endo T, Tasaka M, Shikanai T. PGR5 is involved in cyclic electron flow around photosystem I and is essential for photoprotection in Arabidopsis. Cell. 2002;110(3):361–371.
Burrows PA, Sazanov LA, Svab Z, Maliga P, Nixon PJ. Identification of a functional respiratory complex in chloroplasts through analysis of tobacco mutants containing disrupted plastid ndh genes. EMBO J. 1998;17(4):868–876.
Sazanov LA, Burrows PA, Nixon PJ. The chloroplast Ndh complex mediates the dark reduction of the plastoquinone pool in response to heat stress in tobacco leaves. FEBS Lett. 1998 ;429(1):115–118.
Shikanai T, Endo T, Hashimoto T, Yamada Y, Asada K, Yokota A. Directed disruption of the tobacco ndhB gene impairs cyclic electron flow around photosystem I. Proc Natl Acad Sci USA. 1998;95(16):9705–9709.
Hanke GT, Hase T. Variable photosynthetic roles of two leaf-type ferredoxins in arabidopsis, as revealed by RNA interference. Photochem Photobiol. 2008;84(6):1302-1309.
Hanke GT, Okutani S, Satomi Y, Takao T, Suzuki A, Hase T. Multiple iso-proteins of FNR in Arabidopsis: evidence for different contributions to chloroplast function and nitrogen assimilation. Plant Cell Environ. 2005;28(9):1146–1157.
Okutani S, Hanke GT, Satomi Y, Takao T, Kurisu G, Suzuki A, Hase T. Three maize leaf ferredoxin:NADPH oxidoreductases vary in subchloroplast location, expression, and interaction with ferredoxin. Plant Physiol. 2005;139(3):1451-1459.
Rodriguez RE, Lodeyro A, Poli HO, Zurbriggen M, Peisker M, Palatnik JF, Tognetti VB, Tschiersch H, Hajirezaei MR, Valle EM, Carrillo N. Transgenic tobacco plants overexpressing chloroplastic ferredoxin-NADP(H) reductase display normal rates of photosynthesis and increased tolerance to oxidative stress. Plant Physiol. 2007;143(2):639-649.
Hanke GT, Endo T, Satoh F, Hase T. Altered photosynthetic electron channelling into cyclic electron flow and nitrite assimilation in a mutant of ferredoxin:NADP(H) reductase. Plant Cell Environ. 2008;31(7):1017-1028.
Forti G, Bracale M. Ferredoxin-ferredoxin NADP reductase interaction. FEBS Lett. 1984;166(1):81–84.
Nakatani S, Shin M. The reconstituted NADP photoreducing system by rebinding of the large form of ferredoxin-NADP reductase to depleted thylakoid membranes. Arch Biochem Biophys. 1991;291(2):390–394.
Clark RD, Hawkesford MJ, Coughlan SJ, Bennett J, Hind J. Association of ferredoxin-NADP+ oxidoreductase with the chloroplast cytochrome b-f complex. FEBS Lett. 1984;174(1):137–142.
Zhang H, Whitelegge JP, Cramer WA. Ferredoxin: NADP+ oxidoreductase is a subunit of the chloroplast cytochrome b6f complex. J Biol Chem. 2001;276(41):38159–38165.
Andersen B, Scheller HV, Moller BL. The PSI E subunit of photosystem I binds ferredoxin:NADP+ oxidoreductase. FEBS Lett. 1992;311(2):169–173.
Quiles MJ, Cuello J. Association of ferredoxin-NADP oxidoreductase with the chloroplastic pyridine nucleotide dehydrogenase complex in barley leaves. Plant Physiol. 1998;117(1):235–244.
Twachtmann M, Altmann B, Muraki N, Voss I, Okutani S, Kurisu G, Hase T, Hanke GT. N-Terminal Structure of Maize Ferredoxin:NADP+ Reductase Determines Recruitment into Different Thylakoid Membrane Complexes. Plant Cell. 2012;24(7):2979-2991.
Benz JP, Stengel A, Lintala M, Lee YH, Weber A, Philippar K, Gugel IL, Kaieda S, Ikegami T, Mulo P, Soll J, Bölter B. ArabidopsisTic62 and ferredoxin-NADP(H) oxidoreductase form light-regulated complexes that are integrated into the chloroplast redox poise. Plant Cell. 2009;21(12):3965–3983.
Jurić S, Hazler-Pilepić K, Tomašić A, Lepeduš H, Jeličić B, Puthiyaveetil S, Bionda T, Vojta L, Allen JF, Schleiff E, Fulgosi H. Tethering of ferredoxin:NADP+ oxidoreductase to thylakoid membranes is mediated by novel chloroplast protein TROL. Plant J. 2009;60(5):783–794.
Vojta L, Horvat L, Fulgosi H. Balancing chloroplast redox status – regulation of FNR binding and release. Periodicum biologorum. 2012;114(1):25–31.
Stengel A, Benz P, Balsera M, Soll J, Bölter B. TIC62 redox-regulated translocon composition and dynamics. J Biol Chem. 2008;283(11):6656–6667.
Mizusawa N, Wada H. The role of lipids in photosystem II. Biochim Biophys Acta. 2012;1817(1):194-208.
Xu C, Härtel H, Wada H, Hagio M, Yu B, Eakin C, Benning C. The pgp1 mutant locus of Arabidopsis encodes a phosphatidylglycerophosphate synthase with impaired activity. Plant Physiol. 2002;129:594–604.
Hagio M, Sakurai I, Sato S, Kato T, Tabata S, Wada H. Phosphatidylglycerol is essential for the development of thylakoid membranes in Arabidopsis thaliana. Plant Cell Physiol. 2002;43(12):1456-1464.