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Regulation of Photosynthetic Electron Transport and Photoinhibition

Thomas Roach, Anja Krieger-Liszkay

Current Protein & Peptide Science · 2014 · Review

doi:10.2174/1389203715666140327105143 (opens in a new tab)

Abstract

Photosynthetic organisms and isolated photosystems are of interest for technical applications. In nature, photosynthetic electron transport has to work efficiently in contrasting environments such as shade and full sunlight at noon. Photosynthetic electron transport is regulated on many levels, starting with the energy transfer processes in antenna and ending with how reducing power is ultimately partitioned. This review starts by explaining how light energy can be dissipated or distributed by the various mechanisms of non-photochemical quenching, including thermal dissipation and state transitions, and how these processes influence photoinhibition of photosystem II (PSII). Furthermore, we will highlight the importance of the various alternative electron transport pathways, including the use of oxygen as the terminal electron acceptor and cyclic flow around photosystem I (PSI), the latter which seem particularly relevant to preventing photoinhibition of photosystem I. The control of excitation pressure in combination with the partitioning of reducing power influences the light-dependent formation of reactive oxygen species in PSII and in PSI, which may be a very important consideration to any artificial photosynthetic system or technical device using photosynthetic organisms.

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From PubMed Central, U.S. National Library of Medicine, as current on Oct 22, 2026. Paragraph numbers are ours, so an answer can point at one.

INTRODUCTION

In oxygenic photosynthesis light energy, with the help of light-harvesting antenna, is used to drive two specialized complexes called photosystem I (PSI) and photosystem II (PSII). The energy released from a captured photon triggers charge separation in PSI and PSII reaction centres, and subsequent electron transfer reactions, enabling electrons and protons to be taken from H2O and the release of O2. The released electrons are transported via a series of redox-active co-factors to reduce a final electron acceptor, such as NADP+. At the same time a proton gradient (ΔpH) is generated across the thylakoid membrane that provides, together with the electrochemical gradient (ΔΨ), the proton motive force required for the synthesis of ATP. Charge separation creates a positive charge at the donor side of the photosystems, which is reduced by plastocyanin in PSI and by electrons from the water-splitting complex in PSII. While the absorption of light energy by antenna systems is highly efficient (i.e., extinction coefficient of chl: ε: approx. 105 M-1 cm-1), and helped by a broad range in absorption wavelengths by chlorophyll a, b and a number of carotenoid molecules, energy is lost during charge separation, stabilization and onward electron transfer. In photosynthesis, further energy is lost during CO2 fixation, especially under suboptimal conditions. In an optimal environmental setting, the maximum conversion of solar energy to biomass is estimated at 6%, but only for the most efficient plants.The reaction centres of PSI and PSII convert photon energy into electrical potentials with very high efficiency (80 ± 15 % and 45 ± 10 %, respectively) when measured on a microsecond timescale, making them highly attractive as potential photovoltaic devices. On longer timescales, however, the energy conversion efficiency is largely reduced to about 40% for PSI. Technical applications are increasingly exploiting the efficiency of photosynthesis for solid-state devices mimicking photovoltaic cells. Photo-electric currents have been achieved with immobilized chloroplasts, thylakoid membranes, PSII or PSI core complexes and isolated reaction centres. One of the most promising current bio-photovoltaic without using elaborate or expensive surface chemistries is a PSI complex attached to a semiconductor, achieving a photocurrent density of 362 µA/cm2 and 0.5 V. Purified complexes, photosynthetic membranes or whole organisms have also been placed on electrodes for assembling biosensors (for review see), mainly for the detection of pollutants, but also as components for future H2 production devices. As PSI has a higher efficiency and is less prone to photoinhibion than PSII (see later), it could be more suitable for biomimetic devices (for recent reviews see).

Natural photosynthesis is a highly regulated process. Several mechanisms help to protect the photosystems against light-induced damage (photoinhibition) when photon flux densities exceed the photosynthetic capacity. Moreover, the intensity when light becomes excess depends on the environment. Hence, in unfavourable conditions light saturation occurs at lower intensities (Fig. 1). Excess energy that cannot be used to drive photosynthesis enhances the production of reactive oxygen species (ROS) and induces photooxidative damage. Although some regulatory mechanisms may only be important in a living organism, energy dissipation and alternative electron pathways could be relevant for improving the stability of technical devices based on the use of whole photosynthetic organisms like unicellular algae or of isolated photosystems. This review will cover the different levels that regulate photosynthesis in natural systems by using examples from higher plants and the model green alga Chlamydomonas reinhardtii. We start by covering the various pathways of light-induced production of ROS, major sources of ROS in plants (for reviews see), and then cover how this relates to photoinhibition. The review continues with how excess energy can be dissipated to heat or distributed between the photosystems for protection, but also for influencing the production ratio of ATP:NADPH. Furthermore, we will describe the various electron transport pathways and highlight their importance, from linear to pseudocyclic flow (also called the Mehler reaction) and cyclic, which seems particularly relevant to photoinhibition of PSI. The control of excitation pressure in combination with the partitioning of different electron transport pathways influences the light-dependent formation of ROS, thus is paramount in controlling the stability and longevity of the photosynthetic apparatus.

GENERATION OF REACTIVE OXYGEN SPECIES AND PHOTOINHIBITION

Excitation of pigments and electron transfer reactions in an oxygen-rich environment inevitably leads to photooxidative damage. This is visible to the eye by a bleaching of the chlorophyll leading to pale green or even whitish leaves under extreme light conditions, especially when plants adapted to shade are suddenly exposed to high light intensities. Light-induced damage of the photosynthetic apparatus is caused by excessive production of ROS such as singlet oxygen (1O2), superoxide (O2•-), hydrogen peroxide (H2O2) and hydroxyl radicals (HO•) (Fig. 2).

Among the ROS, 1O2 and HO• are the most reactive species that are able to oxidize lipids, proteins and nucleic acids. Although ROS are important signalling molecules in photosynthetic organisms, high production rates saturate antioxidant defences, lead to oxidative damage and ultimately reduce growth and plant fitness. The up regulation of antioxidant defences is part of an acclimation of photosynthetic organisms to very high light intensities. Moreover, in non-living devices the production of ROS for signalling purposes is unnecessary, allowing ROS production to be minimised.

1O2 is produced by the reaction of excited chlorophyll in its triplet state (3Chl) with 3O2 (molecular oxygen is in a triplet state in its ground state). In the reaction centre of PSII, 3Chl is generated by charge recombination of the primary radical pair (P680+ Phe-), with pheophytin (Phe) being the primary electron acceptor and P680 the primary chlorophyll electron donor. When light absorption exceeds the capacity of photosynthetic electron transport, the probability of 1O2 generation increases. The pathway of charge recombination depends on the energetic of the electron acceptors of PSII (for details see). Charge recombination between the primary quinone electron acceptor (QA) and P680+ can proceed via an indirect pathway and the repopulation of the primary radical pair or directly into the ground state of P680. The indirect pathway leads to the formation of 3Chl and 1O2 while the direct pathway is safe (for a more detailed description see). A regulation mechanism has been described by which the yield of 1O2 production is lowered in PSII with an inactive water-splitting complex. According to this mechanism, 1O2 generation in PSII is controlled by a regulation of midpoint potential of QA. The yield of 1O2 formation is lowered when QA is in its so-called high potential form, i.e., when the midpoint redox potential of QA is shifted to a more positive value. This shift in the midpoint potential allows a direct recombination of P680+QA- to its ground state without repopulating the primary radical pair P680+Phe-. PSII centres with high potential QA have been observed under different physiological conditions in vivo: 1) In green algae prior to photoactivation (the light-dependent assembly of the Mn cluster), and 2) in leaves of higher plants under high light conditions. The dependence of the amount of 1O2 generation in PSII upon the midpoint potential QA has been demonstrated by electron paramagnetic resonance (EPR) spectroscopy in vitro using a spin probe and in vivo by a specific fluorescence dye in Chlamydomonas. It has been shown that the yield of 1O2 generation correlates with the loss of the D1 protein, one of the main subunits of the PSII reaction centre. Although chlorophyll-containing light hravesting complex (LHC) of photosystems contain many more chlorophylls, they are less susceptible to damage by 1O2. In native systems, these antennas are well protected against 1O2 formation by nearby carotenoids, including xanthophylls, which efficiently quench 3Chl.

Photoinhibition and degradation of the D1 protein takes place over a large range of light intensities, although a net loss of PSII activity is only observed at high light intensities since the repair of PSII is very efficient in vivo. However, at very low light intensities when the secondary PSII quinone electron acceptor (QB) is only semi-reduced, photodamage and D1 loss can also take place. For example, PSII photoinhibition caused by charge recombination reactions and 1O2 generation has been observed in green algae at very low light intensities and after excitation of PSII in isolated thylakoid membranes by single turnover flashes. It is not only the midpoint potential of the redox couple QA/QA- that influences the probability of the non-radiative pathway of charge recombination, but also the midpoint potential of the redox couple Phe/Phe-. Interestingly, cyanobacteria have two genes for distinct D1 proteins, a main subunit of the PSII reaction centre, with different amino acids at position D1-130. Special D1-E130 proteins are expressed only during high light conditions, and are thought to shift the redox potential of Phe to enhance charge recombination via the safe non-radiative pathways, thereby lowering the yield of 1O2 generation. Similar to the high light isoform of the D1 protein in cyanobacteria, a glutamate occupies the position D1-130 in all higher plants with known sequences. In Chlamydomonas, substitution of alanine at D1-251 of the QB binding pocket to cysteine improved tolerance to cosmic radiation in space-flight and cell survival after returning to earth. PSII can undergo post-translational modifications associated to stress protection and repair. For example, the phosphorylation of the PSII-associated chlorophyll-binding protein CP29 protects from cold-stress. Moreover, the D1 subunit is under a circadian-regulated phosphorylation pattern and requires dephosphorylation before it undergoes degradation. Phosphorylation is also key to thylakoid membrane folding enabling access of repair enzymes and the migration of photosystem antennas in state transitions, as discussed later. For recent detailed reviews on photosynthesis-related phosphorylation readers are directed towards and.

Beside 1O2 other ROS play an important role in light-induced damage of the photosystems. Superoxide is mainly generated at the acceptor side of PSI in the so-called Mehler reaction. In this reaction, O2 is reduced by ferredoxin or by the PSI iron-sulphur acceptor Fx. In algae and cyanobacteria reduction of O2 can be the dominant electron transport pathway, such as before the light-induced activation of the Calvin-Benson cycle, while in higher plants its importance as alternative electron sink is thought to be less important. It has been recently reported that gymnosperms have an increased capacity of the Mehler reaction (about 10 % of the maximum electron flow) compared to angiosperms, but only during dark to light transition before the Calvin cycle is active. In angiosperms it is thought that the importance of the Mehler reaction increases under stress conditions, such as drought, when the CO2 availability is limited by stomatal closure. In addition it has been reported that the photoperiod plays a role in the partition between linear electron flow to NADP+ and Mehler reaction. Further investigations are needed to elucidate the physiological importance of the Mehler reaction, if and how it is regulated in vivo.

Besides being generated at the acceptor side of PSI, O2•- can also be produced in vitro at the level of the cytochrome b6f complex (cytb6f) and at the acceptor side of PSII. In addition, the cytochrome b559, an intrinsic protein subunit of PSII can act, depending on its redox potential as an oxygen reductase, as a superoxide reductase or as a superoxide oxidase. However, the capacities of these pathways of O2 reduction seem to play only very minor roles in an intact, functional electron transport chain in thylakoid membranes.

A major source of photosynthesis-associated ROS is the H2O2 produced by photorespiration during the recycling of bi-products of the oxygenase activity of RubisCO. This occurs outside the chloroplast in the peroxisome and will not be discussed here. However, H2O2 is generated in isolated chloroplasts, thylakoids and PSI or PSII preparations by the dismutation of O2•-, either spontaneously or catalyzed by superoxide dismutase (SOD). Small amounts of H2O2 can also be generated directly by incomplete water splitting at the donor side of PSII as has been shown in vitro using isolated PSII membranes. Furthermore, it can be formed by the reduction of O2•- by plastoquinol. H2O2 itself is not that toxic, but in the presence of transition metals, such as Fe2+ or Cu+, it is converted to the highly reactive HO• radical (Fig. 2). Apart from the Fenton reaction, HO• may also be formed by the reduction of peroxide by metal centres coordinated to the proteins involved in electron transport.

In the intact chloroplasts, several enzymes are present that detoxify ROS. O2•- is dismutated to H2O2 by SOD containing Cu and Zn (Cu/Zn-SOD) or Fe (Fe-SOD) as a cofactor. H2O2 is mainly detoxified by ascorbate peroxidase (APX) and by chloroplast-located peroxiredoxins (PRX). While APX activity requires ascorbate, PRX activity is dependent on re-reduction by thiol or thioredoxins. The ascorbate (20-300 mM) and glutathione (GSH; 0.5 - 3.5 mM) content of the chloroplast is sufficiently high enough to enable a very efficient H2O2 detoxifying system. Furthermore, thiols and thioredoxins are substrates for glutathione peroxidases and glutathione-s-transferases, which are important in detoxifying reactive lipid species formed by 1O2. 1O2 can be scavenged by tocopherol, plastoquinone, carotenoids and by ascorbate. Despite ascorbate, these scavengers are present in isolated systems and will help to protect the photosystems in the light. However, they have limited capacity because regeneration cannot take place and after a given time they become exhausted. The addition of catalase, a non-chloroplast located enzyme, increased the efficiency and stability of thylakoid bio-electrodes confirming that H2O2 production can be an issue in artificial devices.

It is accepted by the majority of researchers that ROS directly damage photosystems with PSII being more vulnerable against oxidative damage than PSI. However, some researchers have suggested that the repair mechanism of the D1 protein is solely damaged by ROS, and not the PSII reaction centre itself (see Special Issue Physiologia Plantarum 2011 for the current debate). In vivo, the repair of PSII is so efficient that damage is only transitory. Addition of methylviologen to isolated thylakoids, which enhances O2•- at the acceptor side of PSI, still led to greater inhibition of PSII than PSI, showing that PSII is much more susceptible to ROS-induced damage than PSI, even when the site of production is located at PSI. It is intriguing that the D1 reaction centre at the heart of PSII photoinhibiton has remained highly susceptible to photo-damage in all oxygenic photosynthetic organisms and at all light intensities. Despite the apparent wastefulness of PSII photoinhibition, it can also be regarded as a regulatory mechanism of photosynthesis, since it lowers linear electron transport under excess light conditions and may thereby prevent photoinhibition of PSI. Photoinhibition of PSI has a higher impact on the performance of the photosynthetic apparatus since no efficient repair cycle exists.

REGULATION OF LIGHT HARVESTING

What makes photosynthesis remarkable is that it efficiently functions under highly fluctuating photon flux densities, under environmental constraints and in accordance with the metabolic demands of the organism. Photoregulation is coordinated at multiple levels; At the pigment and protein levels via energy-transfer processes involving carotenoids and chlorophylls, at the membrane and cellular levels with supramolecular organization in the thylakoid membrane, at the cellular level by chloroplast relocation and at the organism level including heliotropism of plants and phototaxis of microorganisms. Within the thylakoid membrane, light harvesting complexes facilitate in capturing light energy and its transfer to the reaction centres for charge separation. This partitioning between light harvesting and reaction centres provides an opportunity in regulating how much and to which reaction centre energy is delivered to, thereby preventing excessive excitation, ROS production and the costs associated with photoinhibitory damage. Collectively, this plethora of regulatory mechanisms controlling light energy in intact organism is known as non-photochemical quenching (NPQ), due to their detection by measurements of chlorophyll fluorescence quenching that are distinct from photochemical quenching (i.e., use of the captured light energy in chemical reactions such as CO2 fixation). For a complete guide on chlorophyll measurements of photosynthetic efficiency readers are directed to. There are three components to NPQ; 1) dissipation of excess light energy in antenna to heat before it reaches the reaction centre (qE), 2) state transitions where light adsorption is balanced between the photosystems by movement of antenna (qT) and 3) ‘short term’ photoinhibition of PSII (qI), which recovers slower than qE or qT. As discussed below, qT and qE are governed by the plastoquinone (PQ) pool redox state and the thylakoid proton-motive force, respectively, which makes photosynthesis a highly efficient self-regulated process. For a comprehensive review on how photosynthetic organisms respond to excess light see.

THE QE COMPONENT OF NPQ

The qE component of NPQ, where light energy is dissipated before reaching the reaction centres, has been assigned to a synergistic action of the ∆pH since a low pH in the thylakoid lumen activates the xanthophylls cycle and leads to protonation of luminal residues of proteins such as PsbS and LhcSR3 that reduce energy transfer from the antenna to the PSII reaction centre (Fig. 3). In the xanthophyll cycle, the low pH activates a de-epoxidation of violaxanthin to zeaxanthin and requires minutes to hours to become fully activated, but zeaxanthin can temporarily remain active after the loss of the ΔpH. Lutein is another xanthophyll implicated in qE and protection from high light in both Arabidopsis and Chlamydomonas. Another ∆pH-dependent component of qE is protonation of light harvesting complexes (LHC) and an LHC-type protein, which rapidly induces NPQ within seconds. In higher plants this trans-membrane LHC-type protein is PsbS, whereas in green algae (e.g., Chlamydomonas) it is LhcSR3. Differences between the proteins include PsbS being constitutively expressed and not binding pigments, whereas LhcSR3 is highly inducible by excess light and binds chlorophyll and carotenoids. Moreover, the mechanism of LhcSR3 is less dynamic than PsbS and leads to quenching even in low light, which is perhaps why LhcSR3 is a high light-inducible protein. Evidence suggests the switch from LhcSR3 to PsbS happened when organisms colonised the land and was likely associated with the extra stresses associated with terrestrial life. Mosses can contain both proteins as has been shown for Physcomitrella patens and Rhytidium rugusum (data not shown). Regardless, mutants of Arabidopsis or Chlamydomonas deficient in either PsbS or LhcSR3, both referred to as npq4, have retarded abilities to dissipate excess energy and show sensitivity to high or naturally fluctuating light. The aggregation of LHC II trimers and detachment from PSII is also observed during qE induction. Due to the influence of ΔpH on qE, a regulation of ATP synthase activity, which uses the ΔpH to produce ATP also influences NPQ, possibly through thioredoxin-mediated redox control, but this has yet to be confirmed. Recommended introductory reviews to qE are for model plants and in other plants, but also see for further debate.

We have recently shown in vitro using Arabidopsis chloroplasts that 1O2 production is influenced by the presence or absence of PsbS-dependent NPQ capacity. Other markers of 1O2 production are the oxidation of β-carotene inside the reaction centre and of prenyllipids such as α-tocopherol and plastoquinone. It has also been reported that β-carotene is more oxidised in Arabidopsis npq4 than wild-type following excess light stress, confirming qE protects against reaction centre damage by 1O2. In addition to qE, other mechanisms of energy dissipation operate in specialised situations. For example, a light- and nigericin-insensitive activation of the xanthophyll cycle has been shown in a desiccation-tolerant fern and in lichens. Moreover, an induction of non-radiative charge recombinations in PSII (see above) occurs in cyanobacterial desert crusts in response to high light, avoiding the formation of 3Chl and associated 1O2. Although NPQ is essential to the contribution of desiccation tolerance, the contribution of the xanthophyll cycle to NPQ is not always supported, indicating that other mechanisms, such as chlorophyll cations that are very efficient quenchers, are responsible for the desiccation-induced NPQ. However, xanthophylls may play other roles in protection from excess light due to their efficient ability to scavenge ROS, such as zeaxanthin protecting from 1O2 and neoxanthin from O2•-. In cyanobacteria, a different type of qE has been described which relies on the light-induced conversion of the Orange-Carotenoid-Protein (OCP) to its active red form that quenches fluorescence (for review see).

THE QT COMPONENT OF NPQ

Another light-inducible response, the so-called state transitions, influences excitation delivery to the photosystems. Light Harvesting Complex II (LHCII) can migrate to change the energy deliverance to PSII or PSI. Moreover, the movement of LHCII away from PSII relieves excitation pressure. The migration of LHCII is under governance of the plastoquinone pool redox state, and therefore, the relative activities of PSII and PSI. Under high PSII activity and a reduced plastoquinone pool, plastoquinol binds to the Qo site of the cytb6f, which activates protein kinase STN7 to phosphorylate LHCII. This is a pre-requisite for its movement to PSI and the induction of ‘state 2’ (Fig. 4). A more oxidised plastoquinone pool leads to the dissociation of plastoquinol from the Qo site of cytb6f, thereby deactivating the kinase and activating a phosphatase (as reviewed by). Consequently LHCII migrates back to PSII, referred to as ‘state 1’. State transitions play a minor role in higher plants while they are highly important in green algae. In Arabidopsis, where only up to 15-20% of LHCII can be mobilised, state transitions are important in acclimating to changing light intensities. However, in Chlamydomonas 70- 80% of can disassociates from PSII, but only 20 % associates to PSI. It had been accepted in the literature for a long time that in Chlamydomonas the transition from state 1 to 2 is accompanied by a switch from linear to cyclic electron flow. However, it has recently been demonstrated that this is not necessarily the case. Although both state transitions and cyclic electron flow are induced by reducing conditions, the migration of LHCII to PSI is not a prerequisite for the induction of cyclic electron flow.

The activation of high light responses are upregulated at different times and at different intensities depending upon the organism, presumably to address the different demands and adjustments. The fasted responses are qE (first minutes upon exposure to high light, followed by state transitions (qT) and finally leading to photoinhibition of PSII (qI). Although each mechanism has unique functions they also possess overlapping protective roles.

REGULATION OF THE ELECTRON TRANSPORT CHAIN

Beside the regulation of the amount and distribution of light energy to the reaction centres, the activity of the electron transport chain can be down regulated at different sites. The size of the ΔpH is the most important component that not only controls qE (see above), but also regulates electron transport at the level of the cytb6f and at the donor side of PSII. A decreased luminal pH limits electron transport by slowing the activity of the cytb6f, a regulation mechanism called “photosynthetic control”. The lumen pH in Arabidopsis leaves under ambient CO2 was estimated to range from approximately pH 7.5 to 6.5 under weak and saturating light, respectively. These moderate pH values in the lumen allow regulation at the antenna level via qE and via electron transport through the cytb6f, as well as preventing acid-induced damages. The pH value for zeaxanthin accumulation and PsbS protonation was estimated to be about 6.8. When net ATP synthesis is zero, the pH in the lumen can decrease as low as pH 5.2 (for review see, a pH at which the water-splitting activity and the reduction kinetics of P680+ start to be slowed down. Below pH 5.5, Ca2+, an obligatory co-factor of the water-splitting complex is reversibly removed, evoking a shift of QA to the high potential form and protecting PSII against 1O2 generation (see above).

Beside processes that are regulated by the luminal pH, alternative pathways of photosynthetic electron transport can release the pressure from the electron transport chain and prevent photoinhibtion. Under conditions of limiting light and no limitation on the electron acceptor side (i.e., sufficient CO2), linear electron transport is dominating and electrons released from splitting H2O in PSII are used by PSI to reduce electron acceptors such as NADP+. As electron acceptors become limited (i.e., low CO2 that limits NADPH oxidation for carbon assimilation), pseudocyclic electron flow / Mehler reaction (where O2 is the electron acceptor) and cyclic electron flow are increasingly able to compete for reducing power (Fig. 5). In cyclic electron flow the reducing power is not from PSII, but instead recycled back from PSI into the plastoquinone pool and via cytb6f. This occurs directly via ferredoxin and other proteins, such as PGR5 and PGRL1, or via NAD(P)H dehydrogenases (NDH). As the reinvested reducing power of cyclic electron flow passes through the Q-cycle of cytb6f, the accompanied proton transport from stroma to lumen facilitates the formation of ΔpH, and hence, ATP production. For photosynthetic organisms switching between cyclic and non-cyclic pathways provides a degree of flexibility in the ratio of ATP and NAPDH production to meet metabolic needs. This is particularly important in ATP-expensive photosynthesis, such as CAM plants and in the bundle sheath cells of C4 plants, which both require a higher ATP:NADPH ratio for CO2 fixation than C3 photosynthesis. Furthermore, cyclic flow is enhanced when CO2 becomes limiting in both higher plants and Chlamydomonas, the latter which has a high ATP demand under CO2 limitation because CO2-concentrating mechanisms operate at the expense of ATP. Cyclic electron flow is clearly linked to stress, but the exact regulatory mechanism that switches it on is still unknown. A joint PSI-cytb6f supercomplex for cyclic electron flow has been demonstrated biochemically to be formed in Chlamydomonas, but not yet in higher plants. The NAD(P)H dehydrogenase (NDH) route in Chlamydomonas is achieved by a monomeric protein called Nda2, while in higher plants it is achieved by the NDH complex. Both can operate in the dark with non-photosynthetic supplies of NAD(P)H. In Arabidopsis, at least, both the PGR5/PGRL1-dependent and the NDH-dependent cyclic pathways seem to be under redox regulation by thioredoxin m4, while the PSI-cytb6f supercomplex is Ca2+-dependent in Chlamydomonas.

As discussed above, cyclic electron transport is enhanced to support the production of a ΔpH under conditions that limit carbon assimilation, including environmental stress. Thus it allows ATP generation without the net formation of reductants, such as reduced Ferredoxin or NADPH. The exclusion of linear electron flow also limits the Mehler reaction and O2•- generation. Moreover, cyclic electron flow protects against photodamage of PSI since it keeps the acceptor side oxidized.

In addition to partitioning between linear electron flow, cyclic flow and Mehler reaction (Fig. 5), the plastid terminal oxidase (PTOX), which directly oxidises plastoquinol while reducing O2 to H2O, has been proposed to act as a safety valve and to avoid photo-oxidative damage. PTOX activity may also enhance the formation of a ΔpH, as demonstrated in marine organisms that survive in iron-depleted waters. Here, the cost of building iron-rich PSI is very high, but organisms can operate with PSII activity alone. In alpine plants the level of PTOX protein is elevated, which may be linked to their ability to tolerate harsh conditions like very high irradiation at low temperatures. In agreement with this, PTOX levels are increased in plants exposed to extreme temperatures or to high salinity. In Chlamydomas two isoforms of PTOX exist, PTOX and PTOX2. PTOX2 has been shown to keep the PQ pool oxidized in the dark. However, in other model plants such as Arabidopsis thaliana, Nicotiana tabacum and Solanum lycopersicum grown under standard conditions, the role of PTOX in mature leaves is less clear. Overexpression of PTOX did not protect plants from photoinhibition but actually enhanced it in some circumstances. Recent evidence indicates that PTOX rather modulates the balance between linear and cyclic flow than acting as a safety valve since the capacity of electron flow via PTOX has been measured to be very limited in S. lycopersicum leaves. Further investigations are needed to show the importance of PTOX as a safety valve under stress conditions.

In summary, the competition for absorbed quanta by alternative electron flows becomes important when the electron acceptor NADP+ is limited. These electron flows include cyclic flow around PSI, the Mehler reaction and PTOX activity. Not only does this enable metabolic adjustment through regulating ATP:NADPH ratios, but also releases reducing pressure off the electron transport chain, lowering incidences of charge recombination and preventing 1O2 production (see above). Hence, the incorporation of a regulatory mechanism to prevent over-reduction of charge carriers could also be attractive to artificial systems that may suffer damage under high loads.

COST-BENEFIT RATIO OF REGULATORY MECHANISMS

An obvious cost of photoregulatory processes is the loss in efficiency in photosynthetic yield. A delay in recovery of NPQ from the residual presence of zeaxanthin after high light treatment may have high costs in carbon assimilation of agricultural plants. In the non-native setting of agriculture that strives for maximum growth rates, the protection afforded by NPQ may well be, at times, too conservative. Therefore, opportunities for genetically manipulating light harvesting mechanisms for optimising yields may exist. However, photoregulation prevents photoinhibition, which itself is costly in energetics and resources for repairing damaged reaction centres as well as in the photosynthesis forgone during repair (reviewed by). Other protective expenses to photosynthetic organisms are the investment in antioxidants, such as ascorbate, that require complex biosynthetic pathways and reductants with other enzymes to recycle their activity. Therefore, the cost-benefit ratio of photoregulation versus photoinhibition is extremely complex, especially considering an ecological setting with unpredictable resource availability. As mentioned above, photoinhibition itself is a regulatory process of biological photosystems that can rapidly repair themselves. This brings into question the perhaps impossible task of increasing the longevity of isolated reaction centres in biomimetic systems, but the incorporation of exogenous antioxidants can help.

USE OF REGULATORY MECHANISMS FOR TECHNICAL EXPLOITATION

The question arises as to what we can learn from the regulatory mechanisms of natural photosynthesis for using photosynthetic machinery in technical applications. First, one has to distinguish between technical applications that are either based on exploiting whole organisms as biosensors or using isolated complexes enriched with photosynthetic complexes. Inhibition of the photosynthetic electron transport in whole organisms is easily detectable by monitoring chlorophyll fluorescence and is used to detect heavy metal or herbicide pollution in water. Recent exploits using laser printing have achieved greater contact of electrodes with whole cells or thylakoids, increasing efficiency of charge transfer and sensitivity of biosensors to the nM range. In an intact organism all natural regulatory mechanism are present and can be activated depending on the environmental conditions. To allow a long-term use of a biosensor based on intact organisms it has to be ensured that enough nutrients and CO2 are available and that waste is removed when intact organisms are embedded into a matrix for the technical application.

Regarding isolated systems, PSII complexes or PSII-enriched membrane fragments have already been tested as sensors to detect herbicides. Instead of using active PSII, with the highly vulnerable water-splitting complex, it may be interesting to use PSII with an inactivated donor side. Inactivation of the donor side leads to the shift of the midpoint potential of QA to the high potential form as described above. PSII with high potential QA is protected against damage by 1O2. Herbicides bind efficiently to these modified PSII and herbicide binding could be detected by either measuring thermoluminescence or herbicide-induced changes in the decay kinetics of chlorophyll fluorescence. Isolated photosynthetic systems may well be devoid of metabolic control required in whole organisms in responding to changes in the environments like fluctuating light, but in an outside environment they will still be subjected to highly variable conditions of temperature and light intensity, affecting efficiency and performance. Therefore, when isolated thylakoid membranes or isolated PSI preparations will be used in a technical device, it may be useful to add a safety valve in analogy to the PTOX or the Mehler reaction present in the natural system. This safety valve should only operate when PSI (or both photosystems in case of thylakoids) becomes saturated. To design such a safety valve, a better understanding of the regulation of the Mehler reaction and of PTOX are first needed before reasonable suggestion can be made based on physiologically relevant protection mechanisms. As the field of bio-sensors and transducers is only in its infancy, we can expect great advances when technological advances are coupled with an enhanced understanding of photosynthesis and its control.

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    Faunce TA, Lubitz W, Rutherford AW, MacFarlane D, Moore GF, Yang P, Nocera DG, Moore TA, Gregory DH, Fukuzumi S, Yoon KB, Armstrong FA, Wasielewski MR, Styring S. Energy and environment policy case for a global project on artificial photosynthesis. Energ. Environ. Sci. 2013;6:695-698.

    Not in our library

  5. 5.

    Giardi MT, Pace E. Photosynthetic proteins for technological applications. Trends Biotechnol. 2005;23:257-263.

    Not in our library10.1016/j.tibtech.2005.03.003 (opens in a new tab)

  6. 6.

    Cha Y, Mauzerall D. Energy storage of linear and cyclic electron flows in photosynthesis. Plant Physiol. 1991;100:1869-1877.

    Not in our library10.1104/pp.100.4.1869 (opens in a new tab)

  7. 7.

    Bhardwaj R, Pan RL, Gross EL. Solar energy conversion by chloroplast photoelectrochemical cells. Nature. 1981;289:396-398.

    Not in our library

  8. 8.

    Sjoholm KH, Rasmussen M, Minteer SD. Bio-Solar Cells Incorporating Catalase for Stabilization of Thylakoid Bioelectrodes during Direct Photoelectrocatalysis. ECS Electrochemistry Letters. 2012;1:G7-G9.

    Not in our library

  9. 9.

    Rasmussen M, Minteer SD. Investigating the mechanism of thylakoid direct electron transfer for photocurrent generation. Electrochemical Society 224th Meeting. 2013;2464.

    Not in our library

  10. 10.

    Rasmussen M, Shrier A, Minteer SD. High performance thylakoid bio-solar cell using laccase enzymatic biocathodes. Phys. Chem. Chem. Phys. 2013;15:9062.

    Not in our library10.1039/c3cp51813b (opens in a new tab)

  11. 11.

    Vittadello M, Gorbunov MY, Mastrogiovanni DT, Wielunski LS, Garfunkel EL, Guerrero F, Kirilovsky D, Sugiura M, Rutherford AW, Safari A, Falkowski PG. Photoelectron generation by photosystem II core complexes tethered to gold surfaces. ChemSusChem. 2010;3:471-475.

    Not in our library10.1002/cssc.200900255 (opens in a new tab)

  12. 12.

    Kato M, Cardona T, Rutherford AW, Reisner E. Photoelectrochemical water oxidation with photosystem II integrated in a mesoporous indium-tin oxide electrode. J. Am. Chem. Soc. 2012;134:8332-8335.

    Not in our library10.1021/ja301488d (opens in a new tab)

  13. 13.

    Yehezkeli O, Tel-Vered R, Michaeli D, Nechushtai R, Willner I. Photosystem I (PSI)/Photosystem II (PSII)-Based Photo-Bioelectrochemical Cells Revealing Directional Generation of Photocurrents. Small. 2013;9(17):2970-2978.

    Not in our library10.1002/smll.201300051 (opens in a new tab)

  14. 14.

    Terasaki N, Yamamoto N, Tamada K, Hattori M, Hiraga T, Tohri A, Sato I, Iwai M, Iwai M, Taguchi S, Enami I, Inoue Y, Yamanoi Y, Yonezawa T, Mizuno K, Murata M, Nishihara H, Yoneyama S, Minakata M, Ohmori T, Sakai M, Fujii M. Bio-photosensor: Cyanobacterial photosystem I coupled with transistor via molecular wire. Biochim Biophys Acta. 2007;1767:653-659.

    Not in our library10.1016/j.bbabio.2006.11.008 (opens in a new tab)

  15. 15.

    Mershin A, Matsumoto K, Kaiser L, Yu D, Vaughn M, Nazeeruddin MK, Bruce BD, Graetzel M, Zhang S. Self-assembled photosystem-I biophotovoltaics on nanostructured TiO2 and ZnO. Scientific Reports. 2012;2:234.

    Not in our library10.1038/srep00234 (opens in a new tab)

  16. 16.

    Carmeli I, Frolov L, Carmeli C, Richter S. Photovoltaic activity of photosystem I-based self-assembled monolayer. J. Am. Chem. Soc. 2007;129:12352-12353.

    Not in our library10.1021/ja073040c (opens in a new tab)

  17. 17.

    Lukashev EP, Nadtochenko VA, Permenova EP, Sarkisov OM, Rubin AB. Electron phototransfer between photosynthetic reaction centers of the bacteria Rhodobacter sphaeroides and semiconductor mesoporous tiO2 films. Dokl. Biochem. Biophys. 2007;415:211-216.

    Not in our library10.1134/s1607672907040138 (opens in a new tab)

  18. 18.

    Das R, Kiley PJ, Segal M, Norville J, Yu AA, Wang L, Trammell SA, Reddick LE, Kumar R, Stellacci F, Lebedev N, Schnur J, Bruce BD, Zhang S, Baldo M. Integration of photosynthetic protein molecular complexes in solid-state electronic devices. Nano Letters. 2004;4:1079-1083.

    Not in our library

  19. 19.

    Trammell SA, Spano A, Price R, Lebedev N. Effect of protein orientation on electron transfer between photosynthetic reaction centers and carbon electrodes. Biosens Bioelectron. 2006;21:1023-1028.

    Not in our library10.1016/j.bios.2005.03.015 (opens in a new tab)

  20. 20.

    Bhalla V, Zazubovich V. Self-assembly and sensor response of photosynthetic reaction centers on screen-printed electrodes. Analytica Chimica Acta. 2011;707:184-190.

    Not in our library10.1016/j.aca.2011.09.020 (opens in a new tab)

  21. 21.

    Ventrella A, Catucci L, Placido T, Longobardi F, Agostiano A. Biomaterials based on photosynthetic membranes as potential sensors for herbicides. Biosens. Bioelectron. 2011;26:4747-4752.

    Not in our library10.1016/j.bios.2011.05.043 (opens in a new tab)

  22. 22.

    Rasmussen M, Minteer SD. Self-powered herbicide biosensor utilizing thylakoid membranes. Anal. Methods. 2013;5:1140.

    Not in our library

  23. 23.

    Breton F, Euzet P, Piletsky SA, Giardi MT, Rouillon R. Integration of photosynthetic biosensor with molecularly imprinted polymer-based solid phase extraction cartridge. Analytica Chimica Acta. 2006;569:50-57.

    Not in our library

  24. 24.

    Touloupakis E, Boutopoulos C, Buonasera K, Zergioti I, Giardi MT. A photosynthetic biosensor with enhanced electron transfer generation realized by laser printing technology. Anal. Bioanal. Chem. 2012;402:3237-3244.

    Not in our library10.1007/s00216-012-5771-7 (opens in a new tab)

  25. 25.

    Ferro Y, Perullini M, Jobbagy M, Bilmes SA, Durrieu C. Development of a biosensor for environmental monitoring based on microalgae immobilized in silica hydrogels. Sensors. 2012;12:16879-16891.

    Not in our library10.3390/s121216879 (opens in a new tab)

  26. 26.

    Lambreva MD, Giardi MT, Rambaldi I, Antonacci A, Pastorelli S, Bertalan I, Husu I, Johanningmeier U, Rea G. A powerful molecular engineering tool provided efficient Chlamydomonas mutants as bio-sensing elements for herbicides detection. PloS One. 2013;8:e61851.

    Not in our library10.1371/journal.pone.0061851 (opens in a new tab)

  27. 27.

    Husu I, Rodio G, Touloupakis E, Lambreva MD, Buonasera K, Litescu SC, Giardi MT, Rea G. Insights into photo-electrochemical sensing of herbicides driven by Chlamydomonas reinhardtii cells. Sensor. Actuat. B-Chem. 2013;185:321-330.

    Not in our library

  28. 28.

    Zamaleeva AI, Sharipova IR, Shamagsumova RV, Ivanov AN, Evtugyn GA, Ishmuchametova DG, Fakhrullin RF. A whole-cell amperometric herbicide biosensor based on magnetically functionalised microalgae and screen-printed electrodes. Anal Methods. 2011;3:509.

    Not in our library10.1039/c0ay00627k (opens in a new tab)

  29. 29.

    Shitanda I, Takamatsu S, Watanabe K, Itagaki M. Amperometric screen-printed algal biosensor with flow injection analysis system for detection of environmental toxic compounds. Electrochimica Acta. 2009;54:4933-4936.

    Not in our library

  30. 30.

    Buonasera K, Lambreva M, Rea G, Touloupakis E, Giardi MT. Technological applications of chlorophyll a fluorescence for the assessment of environmental pollutants. Anal. Bioanal. Chem. 2011;401:1139-1151.

    Not in our library10.1007/s00216-011-5166-1 (opens in a new tab)

  31. 31.

    Doval DA, Areephong J, Bang EK, Bertone L, Charbonnaz P, Fin A, Lin NT, Lista M, Matile S, Montenegro J, Orentas E, Sakai N, Tran DH, Jentzsch AV. Recent progress with functional biosupramolecular systems. Langmuir. 2011;27:9696-9705.

    Not in our library10.1021/la200593p (opens in a new tab)

  32. 32.

    Mazor Y, Toporik H, Nelson N. Temperature-sensitive PSII and promiscuous PSI as a possible solution for sustainable photosynthetic hydrogen production. Biochim. Biophys. Acta. 2012;1817:1122-1126.

    Not in our library10.1016/j.bbabio.2012.01.005 (opens in a new tab)

  33. 33.

    Lubner CE, Grimme R, Bryant DA, Golbeck JH. Wiring photosystem I for direct solar hydrogen production. Biochemistry. 2010;49:404-414.

    Not in our library10.1021/bi901704v (opens in a new tab)

  34. 34.

    Kargul J, Janna Olmos JD, Krupnik T. Structure and function of photosystem I and its application in biomimetic solar-to-fuel systems. J. Plant Physiol. 2012;169:1639-1653.

    Not in our library10.1016/j.jplph.2012.05.018 (opens in a new tab)

  35. 35.

    Seibert M, Picorel R, Rubin AB, Connolly JS. Spectral photophysical and stability properties of isolated photosystem II reaction center. Plant Physiol. 1988;87:303-306.

    Not in our library10.1104/pp.87.2.303 (opens in a new tab)

  36. 36.

    Asada K. Production and scavenging of reactive oxygen species in chloroplasts and their functions. Plant Physiol. 2006;141:391-396.

    Not in our library10.1104/pp.106.082040 (opens in a new tab)

  37. 37.

    Fischer BB, Hideg É, Krieger-Liszkay A. Production detection and signaling of singlet oxygen in photosynthetic organisms. Antioxid. Redox Signal. 2013;18:2145-2162.

    Not in our library10.1089/ars.2012.5124 (opens in a new tab)

  38. 38.

    Foyer CH, Noctor G. Redox regulation in photosynthetic organisms signaling acclimation and practical implications. Antioxid Redox Signal. 2009;11:861-905.

    Not in our library10.1089/ars.2008.2177 (opens in a new tab)

  39. 39.

    Keren N, Krieger-Liszkay A. Photoinhibition molecular mechanisms and physiological significance. Physiol. Plant. 2011;142:1-5.

    Not in our library10.1111/j.1399-3054.2011.01467.x (opens in a new tab)

  40. 40.

    Krieger-Liszkay A, Fufezan C, Trebst A. Singlet oxygen production in photosystem II and related protection mechanism. Photosynth. Res. 2008;98:551-564.

    Not in our library10.1007/s11120-008-9349-3 (opens in a new tab)

  41. 41.

    Förster B, Osmond CB, Pogson BJ. Improved survival of very high light and oxidative stress is conferred by spontaneous gain-of-function mutations in Chlamydomonas. Biochim. Biophys. Acta. 2005;1709:45-57.

    Not in our library10.1016/j.bbabio.2005.05.012 (opens in a new tab)

  42. 42.

    Krieger-Liszkay A, Rutherford AW. Influence of herbicide binding on the redox potential of the quinone acceptor in photosystem II relevance to photodamage and phytotoxicity. Biochemistry. 1998;37:17339-17344.

    Not in our library10.1021/bi9822628 (opens in a new tab)

  43. 43.

    Vass I. Molecular mechanisms of photodamage in the Photosystem II complex. Biochim. Biophys. Acta. 2012;1817:209-217.

    Not in our library10.1016/j.bbabio.2011.04.014 (opens in a new tab)

  44. 44.

    Rutherford AW, Krieger-Liszkay A. Herbicide-induced oxidative stress in photosystem II. Trends Biochem. Sci. 2001;26:648-653.

    Not in our library10.1016/s0968-0004(01)01953-3 (opens in a new tab)

  45. 45.

    Fufezan C, Drepper F, Juhnke HD, Lancaster CRD, Un S, Rutherford AW, Krieger-Liszkay A. Herbicide-induced changes in charge recombination and redox potential of Q(A) in the T4 mutant of Blastochloris viridis. Biochemistry. 2005;44:5931-5939.

    Not in our library10.1021/bi050055j (opens in a new tab)

  46. 46.

    Johnson GN, Rutherford AW, Krieger A. A change in the midpoint potential of the quinone QA in Photosystem II associated with photoactivation of oxygen evolution. Biochim Biophys Acta. 1995;1229:202-207.

    Not in our library

  47. 47.

    Fufezan C, Gross CM, Sjödin M, Rutherford AW, Krieger-Liszkay A, Kirilovsky D. Influence of the redox potential of the primary quinone electron acceptor on photoinhibition in photosystem II. J. Biol. Chem. 2007;282:12492-12502.

    Not in our library10.1074/jbc.m610951200 (opens in a new tab)

  48. 48.

    Fischer BB, Krieger-Liszkay A, Hideg E, Snyrychová I, Wiesendanger M, Eggen RIL. Role of singlet oxygen in chloroplast to nucleus retrograde signaling in Chlamydomonas reinhardtii. FEBS Lett. 2007;581:5555-5560.

    Not in our library10.1016/j.febslet.2007.11.003 (opens in a new tab)

  49. 49.

    Roach T, Sedoud A, Krieger-Liszkay A. Acetate in mixotrophic growth medium affects photosystem II in Chlamydomonas reinhardtii and protects against photoinhibition. Biochim. Biophys. Acta. 2013;1827:1183-1190.

    Not in our library10.1016/j.bbabio.2013.06.004 (opens in a new tab)

  50. 50.

    Hideg E, Spetea C, Vass I. Singlet oxygen production in thylakoid membranes during photoinhibition as detected by EPR spectroscopy. Photosynth. Res. 1994;39:191-199.

    Not in our library10.1007/bf00029386 (opens in a new tab)

  51. 51.

    Fischer BB, Eggen RIL, Trebst A, Krieger-Liszkay A. The glutathione peroxidase homologous gene Gpxh in Chlamydomonas reinhardtii is upregulated by singlet oxygen produced in photosystem II. Planta. 2006;223:583-590.

    Not in our library10.1007/s00425-005-0108-9 (opens in a new tab)

  52. 52.

    Jahns P, Holzwarth AR. The role of the xanthophyll cycle and of lutein in photoprotection of photosystem II. Biochim. Biophys. Acta. 2012;1817:182-193.

    Not in our library10.1016/j.bbabio.2011.04.012 (opens in a new tab)

  53. 53.

    Böhm F, Edge R, Truscott TG. Interactions of dietary carotenoids with singlet oxygen 1O2 and free radicals: potential effects for human health. Acta Biochimica Polonica. 2012;59:27-30.

    Not in our library

  54. 54.

    Edelman M, Mattoo AK. D1-protein dynamics in photosystem II: the lingering enigma. Photosynth. Res. 2008;98:609-620.

    Not in our library10.1007/s11120-008-9342-x (opens in a new tab)

  55. 55.

    Keren N, Gong H, Ohad I. Oscillations of reaction center II-D1 protein degradation in vivo induced by repetitive light flashes Correlation between the level of RCII-QB- and protein degradation in low light. J. Biol. Chem. 1995;270:806-814.

    Not in our library10.1074/jbc.270.2.806 (opens in a new tab)

  56. 56.

    Szilárd A, Sass L, Hideg E, Vass I. Photoinactivation of photosystem II by flashing light. Photosynth. Res. 2005;84:15-20.

    Not in our library10.1007/s11120-004-7161-2 (opens in a new tab)

  57. 57.
    Mechanism of photosystem II photoinactivation and D1 protein degradation at low light: The role of back electron flow · 1997
    Not checked
  58. 58.

    Cser K, Vass I. Radiative and non-radiative charge recombination pathways in Photosystem II studied by thermoluminescence and chlorophyll fluorescence in the cyanobacterium Synechocystis 6803. Biochim. Biophys Acta. 2007;1767:233-243.

    Not in our library10.1016/j.bbabio.2007.01.022 (opens in a new tab)

  59. 59.

    Kós PB, Deák Z, Cheregi O, Vass I. Differential regulation of psbA and psbD gene expression and the role of the different D1 protein copies in the cyanobacterium Thermosynechococcus elongatus BP-1. Biochim. Biophys. Acta. 2008;1777:74-83.

    Not in our library10.1016/j.bbabio.2007.10.015 (opens in a new tab)

  60. 60.

    Giardi MT, Rea G, Lambreva MD, Antonacci A, Pastorelli S, Bertalan I, Johanningmeier U, Mattoo AK. Mutations of photosystem II D1 protein that empower efficient phenotypes of Chlamydomonas reinhardtii under extreme environment in space. PloS One. 2013;8:e64352.

    Not in our library10.1371/journal.pone.0064352 (opens in a new tab)

  61. 61.

    Bergantino E, Dainese P, Cerovic Z, Sechi S, Bassi R. A Post-translational Modification of the Photosystem II Subunit CP29 Protects Maize from Cold Stress. J. Biol. Chem. 1995;270:8474-8481.

    Not in our library10.1074/jbc.270.15.8474 (opens in a new tab)

  62. 62.

    Booij-James IS, Swegle WM, Edelman M, Mattoo AK. Phosphorylation of the D1 photosystem II reaction center protein is controlled by an endogenous circadian rhythm. Plant Physiol. 2002;130:2069-2075.

    Not in our library10.1104/pp.013441 (opens in a new tab)

  63. 63.

    Rintamäki E, Kettunen R, Aro EM. Differential D1 dephosphorylation in functional and photodamaged photosystem II centers Dephosphorylation is a prerequisite for degradation of damaged D1. J. Biol. Chem. 1996;271:14870-14875.

    Not in our library10.1074/jbc.271.25.14870 (opens in a new tab)

  64. 64.

    Fristedt R, Willig A, Granath P, Crèvecoeur M, Rochaix JD, Vener AV. Phosphorylation of photosystem II controls functional macroscopic folding of photosynthetic membranes in Arabidopsis. Plant Cell. 2009;21:3950-3964.

    Not in our library10.1105/tpc.109.069435 (opens in a new tab)

  65. 65.

    Rochaix JD, Lemeille S, Shapiguzov A, Samol I, Fucile G, Willig A, Goldschmidt-Clermont M. Protein kinases and phosphatases involved in the acclimation of the photosynthetic apparatus to a changing light environment. Philos. Trans. R. Soc. Lond. B Biol. Sci. 2012;367:3466-3474.

    Not in our library10.1098/rstb.2012.0064 (opens in a new tab)

  66. 66.

    Tikkanen M, Aro EM. Integrative regulatory network of plant thylakoid energy transduction. Trends Plant Sci. 2014;19(1):10-17.

    Not in our library10.1016/j.tplants.2013.09.003 (opens in a new tab)

  67. 67.

    Mehler A. Studies on the reactions of illuminated chloroplasts 11 Stimulation on inhibition of the reaction with molecular oxygen. Arch Biochem Biophys. 1951;34:339-351.

    Not in our library10.1016/0003-9861(51)90012-4 (opens in a new tab)

  68. 68.

    Takahashi M, Asada K. Superoxide production in aprotic interior of chloroplast thylakoids. Arch. Biochem. Biophys. 1988;267:714-722.

    Not in our library10.1016/0003-9861(88)90080-x (opens in a new tab)

  69. 69.

    Asada K. The water-water cycle as alternative photon and electron sinks. Philos. Trans. R. Soc. Lond. B Biol. Sci. 2000;355:1419-1431.

    Not in our library10.1098/rstb.2000.0703 (opens in a new tab)

  70. 70.

    Allen JF, Gantt E, Golbeck JH, Osmond B, Frank F, Houyoux PA. The Mehler Reaction in Chlamydomonas During Photosynthetic Induction and Steady-State Photosynthesis in Wild-Type and in a Mitochondrial Mutant. In Photosynthesis Energy from the Sun 14th International Congress on Photosynthesis. Springer Netherlands Dordrecht. 2008;581-584.

    Not in our library

  71. 71.

    Ort DR, Baker NR. A photoprotective role for O2 as an alternative electron sink in photosynthesis?. Curr. Opin. Plant Biol. 2002;5:193-198.

    Not in our library10.1016/s1369-5266(02)00259-5 (opens in a new tab)

  72. 72.

    Driever SM, Baker NR. The water-water cycle in leaves is not a major alternative electron sink for dissipation of excess excitation energy when CO2 assimilation is restricted. Plant Cell Environ. 2011;34:837-846.

    Not in our library10.1111/j.1365-3040.2011.02288.x (opens in a new tab)

  73. 73.

    Shirao M, Kuroki S, Kaneko K, Kinjo Y, Tsuyama M, Förster B, Takahashi S, Badger MR. Gymnosperms Have Increased Capacity for Electron Leakage to Oxygen (Mehler and PTOX reactions) in Photosynthesis Compared with Angiosperms. Plant Cell Physiol. 2013;54:1152-1163.

    Not in our library10.1093/pcp/pct066 (opens in a new tab)

  74. 74.

    Michelet L, Krieger-Liszkay A. Reactive oxygen intermediates produced by photosynthetic electron transport are enhanced in short-day grown plants. Biochim Biophys Acta. 2012;1817:1306-1313.

    Not in our library10.1016/j.bbabio.2011.11.014 (opens in a new tab)

  75. 75.

    Sang M, Xie J, Qin XC, Wang WD, Chen XB, Wang KB, Zhang JP, Li LB, Kuang TY. High-light induced superoxide radical formation in cytochrome b6f complex from Bryopsis corticulans as detected by EPR spectroscopy. J. Photochem. Photobiol. B. 2011;102:177-181.

    Not in our library10.1016/j.jphotobiol.2010.11.002 (opens in a new tab)

  76. 76.

    Pospíšil P. Molecular mechanisms of production and scavenging of reactive oxygen species by photosystem II. Biochim. Biophys. Acta. 2012;1817:218-231.

    Not in our library10.1016/j.bbabio.2011.05.017 (opens in a new tab)

  77. 77.

    Foyer CH, Bloom AJ, Queval G, Noctor G. Photorespiratory metabolism genes mutants energetics and redox signaling. Ann. Rev. Plant Biol. 2009;60:455-484.

    Not in our library10.1146/annurev.arplant.043008.091948 (opens in a new tab)

  78. 78.

    Arató A, Bondarava N, Krieger-Liszkay A. Production of reactive oxygen species in chloride- and calcium-depleted photosystem II and their involvement in photoinhibition. Biochim. Biophys. Acta. 2004;1608:171-180.

    Not in our library10.1016/j.bbabio.2003.12.003 (opens in a new tab)

  79. 79.

    Mubarakshina MM, Ivanov BN. The production and scavenging of reactive oxygen species in the plastoquinone pool of chloroplast thylakoid membranes. Physiol Plant. 2010;140:103-110.

    Not in our library10.1111/j.1399-3054.2010.01391.x (opens in a new tab)

  80. 80.

    Miyake C. Alternative electron flows (water-water cycle and cyclic electron flow around PSI) in photosynthesis molecular mechanisms and physiological functions. Plant Cell Physiol. 2010;51:1951-1963.

    Not in our library10.1093/pcp/pcq173 (opens in a new tab)

  81. 81.

    Dietz KJ. Peroxiredoxins in plants and cyanobacteria. Antioxid. Redox Signal. 2011;15:1129-1159.

    Not in our library10.1089/ars.2010.3657 (opens in a new tab)

  82. 82.

    Noctor G, Gomez L, Vanacker H, Foyer CH. Interactions between biosynthesis compartmentation and transport in the control of glutathione homeostasis and signalling. J. Exp. Bot. 2002;53:1283-1304.

    Not in our library10.1093/jexbot/53.372.1283 (opens in a new tab)

  83. 83.

    Smirnoff N, Conklin PL, Loewus FA. Biosynthesis of ascorbic acid in plants A Renaissance. Ann. Rev. Plant Physiol. Plant Mol. Biol. 2001;52:437-467.

    Not in our library10.1146/annurev.arplant.52.1.437 (opens in a new tab)

  84. 84.

    Ledford HK, Chin BL, Niyogi KK. Acclimation to singlet oxygen stress in Chlamydomonas reinhardtii. Eukaryotic Cell. 2007;6:919-930.

    Not in our library10.1128/ec.00207-06 (opens in a new tab)

  85. 85.

    Fischer BB, Ledford HK, Wakao S, Huang SG, Casero D, Pellegrini M, Merchant SS, Koller A, Eggen RIL, Niyogi KK. SINGLET OXYGEN RESISTANT 1 links reactive electrophile signaling to singlet oxygen acclimation in Chlamydomonas reinhardtii. Proc. Natl Acad. Sci. U.S.A. 2012;109:E1302-E1311.

    Not in our library10.1073/pnas.1116843109 (opens in a new tab)

  86. 86.

    Kruk J, Trebst A. Plastoquinol as a singlet oxygen scavenger in photosystem II. Biochim. Biophys Acta. 2008;1777:154-162.

    Not in our library10.1016/j.bbabio.2007.10.008 (opens in a new tab)

  87. 87.

    Bisby RH, Morgan CG, Hamblett I, Gorman AA. Quenching of singlet oxygen by trolox C ascorbate and amino acids Effects of pH and temperature. J. Phys Chem A. 1999;103:7454-7459.

    Not in our library

  88. 88.

    Krieger-Liszkay A, Trebst A. Tocopherol is the scavenger of singlet oxygen produced by the triplet states of chlorophyll in the PSII reaction centre. J. Exp Bot. 2006;57:1677-1684.

    Not in our library10.1093/jxb/erl002 (opens in a new tab)

  89. 89.

    Krieger-Liszkay A, Kós PB, Hideg E. Superoxide anion radicals generated by methylviologen in photosystem I damage photosystem II. Physiol. Plant. 2011;142:17-25.

    Not in our library10.1111/j.1399-3054.2010.01416.x (opens in a new tab)

  90. 90.

    Sonoike K. Photoinhibition of photosystem I. Physiol Plant. 2011;142:56-64.

    Not in our library10.1111/j.1399-3054.2010.01437.x (opens in a new tab)

  91. 91.

    Maxwell K, Johnson GN. Chlorophyll fluorescence a practical guide. J. Exp Bot. 2000;51:659-68.

    Not in our library10.1093/jxb/51.345.659 (opens in a new tab)

  92. 92.

    Quick WP, Stitt M. An examination of factors contributing to non-photochemical quenching of chlorophyll fluorescence in barley leaves. Biochim Biophys Acta. 1989;977:287-296.

    Not in our library

  93. 93.

    Nilkens M, Kress E, Lambrev P, Miloslavina Y, Müller M, Holzwarth AR, Jahns P. Identification of a slowly inducible zeaxanthin-dependent component of non-photochemical quenching of chlorophyll fluorescence generated under steady-state conditions in Arabidopsis. Biochim. Biophys. Acta. 2010;1797:466-475.

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  94. 94.

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