JXB Advance Access originally published online on March 21, 2006
Journal of Experimental Botany 2006 57(6):1211-1223; doi:10.1093/jxb/erj104
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REVIEW ARTICLE |
Conservation and dissipation of light energy as complementary processes: homoiohydric and poikilohydric autotrophs
1Julius von Sachs Institute of Biosciences, University of Würzburg, D-97082 Würzburg, Germany
2Institute of Basic Biological Problems, Russian Academy of Sciences, Pushchino-na-Oke, Moscow Region, and Laboratory of Biophysics, Belozersky Institute of Chemical and Physical Biology, Moscow State University, Moscow 119992, Russia
* To whom correspondence should be addressed. E-mail: heber{at}botanik.uni-wuerzburg.de
Received 26 July 2005; Accepted 21 December 2005
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Abstract |
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Top
Abstract
Introduction: photosynthesis...The relationship between photosynthetic energy conservation and thermal dissipation of light energy is considered, with emphasis on organisms which tolerate full desiccation without suffering photo-oxidative damage in strong light. As soon as water becomes available to dry poikilohydric organisms, they resume photosynthetic water oxidation. Only excess light is then thermally dissipated in mosses and chlorolichens by a mechanism depending on the protonation of a thylakoid protein and availability of zeaxanthin. Upon desiccation, another mechanism is activated which requires neither protonation nor zeaxanthin although the zeaxanthin-dependent mechanism of energy dissipation remains active, provided desiccation occurs in the light. Increased thermal energy dissipation under desiccation finds expression in the loss of variable, and in the quenching of, basal chlorophyll fluorescence. Spectroscopical analysis revealed the activity of photosystem II reaction centres in the absence of water. Oxidized ß-carotene (Car+) and reduced chlorophyll (Chl–), perhaps ChlD1 next to P680 within the D1 subunit, accumulates reversibly under very strong illumination. Although recombination between Car+ and Chl– is too slow to contribute significantly to thermal energy dissipation, a much faster reaction such as the recombination between P680+ and the neighbouring Chl– is suggested to form the molecular basis of desiccation-induced energy dissipation in photosystem II reaction centres. Thermal dissipation of absorbed light energy within a picosecond time domain deactivates excited singlet chlorophyll, thereby preventing triplet accumulation and the consequent photo-oxidative damage by singlet oxygen.
Key words: Chlorophyll fluorescence, energy dissipation, lichens, mosses, photoprotection, photosystem II, reaction centre, zeaxanthin
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Introduction: photosynthesis versus photoprotection |
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The central requirements for autotrophic plant life are the availability of light, water, and CO2. Whereas light absorption by photosynthetic pigments is proportional to the incident light intensity, photosynthesis is not. Essentially, all absorbed light is used for photosynthetic reactions by unstressed leaves only as long as photon flux is very low. By contrast, light is invariably far in excess when plants are fully exposed to sunlight. CO2 then becomes rate-limiting for photosynthesis. Light absorbed by the photosynthetic apparatus may activate atmospheric oxygen which, as singlet oxygen (1O2), is either indiscriminately oxidative (Krieger-Liszkay, 2005
) or, in its univalently reduced form 
, gives rise to reactive oxygen radicals (Siefermann-Harms, 1987; Owens, 1996; Asada, 1999). The question of the nature of the mechanisms which prevent excess excitation energy from damaging the photosynthetic apparatus in hydrated plants by unspecific oxidation has been and still is a subject of intensive research (Demmig-Adams, 1990; Schreiber and Neubauer, 1990; Krieger and Weiss, 1993; Björkman and Demmig-Adams, 1995; Bruce et al., 1997; Gilmore and Govindjee, 1999; Niyogi, 1999; Heber et al., 2001; Finazzi et al., 2004; Holt et al., 2004, 2005; Krause and Jahns, 2004; Horton et al., 2005; Pascal et al., 2005; and others).
Whereas the survival of the majority of higher plants depends on the maintenance of hydration, many mosses, lichens, and aerophilic algae tolerate full dehydration (Green and Lange, 1995; Alpert, 2000). For metabolic activity, they depend on moisture they take from their environment. Even under temperate conditions, epilithic or terrestrial lichens are hydrated for only 35–65% of the annual time period, depending on lichen type and habitat. Desert lichens experience less than 10% of their life in conditions of water potential that allow for photosynthetic and/or respiratory activities (Evans and Lange, 2003). For the remainder of the time they are desiccated. Survival under high irradiance depends on mechanisms which permit rapid thermal dissipation of perceived radiation. The capability of poikilohydric autotrophs to cope with strong irradiance is species-specific. Many lichens are less easily damaged by strong light in the desiccated state than under hydrated conditions (Demmig-Adams et al., 1990). Tolerance of exposure has a strong adaptive component and depends on habitat conditions or pretreatment (Gauslaa and Solhaug, 2004). Lichens from shade habitats are damaged when they are exposed to high light even when dehydrated (Gauslaa and Solhaug, 1999; Gauslaa et al., 2001).
In photosynthesis, only a few specialized chlorophyll molecules are involved in converting light energy into redox energy. A much larger number serves, together with several carotenoids, to harvest light energy. Absorbed excitation energy migrates to and is trapped by a limited number of chlorophyll-containing reaction centres (RCs) where energy conservation is initiated by charge separation within about 3 ps (Zinth and Kaiser, 1993). The RC of photosystem II (PSII) where water is oxidized is composed of two proteins, D1 and D2, and contains six chlorophylls, two pheophytins, two ß-carotenes, two quinones, and one or two haems of cytochrome b559 (Fig. 1). Endergonic oxidation of a chlorophyll dimer termed P680, or P, creates an oxidant of a redox potential high enough to oxidize water in several steps. The liberated electrons are transfered first to one of the bound quinones of the RC, a plastoquinone (PQ) termed QA, then to QB and from there, through a number of further steps involving also the reaction centre of photosystem I, PSI, to a physiological electron acceptor such as CO2.
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As long as water is present as a donor molecule, its oxidation reduces the oxidized primary electron donor, P680+, which is created by charge separation in the RC of PSII. This prevents P680+ from unspecific oxidative side reactions which can destroy components of the photosynthetic apparatus. In the absence of effective thermal energy dissipation, excessive excitation of light-harvesting chlorophylls under strong irradiance leads to increased long-lived triplet formation, not only within the pigment bed but also within PSII RCs (Krieger-Liszkay 2005
). If triplet detoxification by carotenoids (Siefermann-Harms, 1987) cannot keep pace with triplet accumulation, highly oxidative singlet oxygen will be created by
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will, finally, result in cell death. Obviously, the problem of over-excitation is exacerbated when sun-exposed photosynthetic organisms desiccate. In the present communication, conservation of light energy in photosynthesis will only be considered briefly, but attention will be directed to problems of regulated thermal energy dissipation in hydrated and desiccated photosynthetic organisms as the main means of photoprotection.
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Chlorophyll fluorescence as a tool to assess the effectiveness of thermal energy dissipation as a sink for excess excitation energy |
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The energy of light absorbed by the photosynthetic apparatus may either be used for photochemical charge separation in the RCs, or it may be re-emitted as fluorescence, or it may be dissipated harmlessly in the form of heat. As these processes are mutually exclusive, i.e. they compete with one another, measurements of fluorescence can give important information, not only on photosynthetic electron flow but also on energy dissipation by the photosynthetic apparatus (Bradbury and Baker, 1981
; Schreiber et al., 1986; Krause and Weis, 1991; Krause and Jahns, 2004). Fluorescence is decreased, or quenched, when light energy is used for photosynthetic electron transport and when it is thermally dissipated. Both modes of fluorescence quenching can be experimentally distinguished when modulated light is used for measuring fluorescence (Schreiber and Bilger, 1993; Krause and Jahns, 2003).
Modulated fluorescence emitted by hydrated plants is at its minimum level (termed Fo) under strictly rate-limiting light. Electron carriers of the photosynthetic electron transport chain such as QA are largely oxidized under very low light. The potential for charge separation in PSII RCs is maximal. However, when actinic light is strong enough to reduce QA fully, modulated fluorescence is at its maximum level (termed Fm) because photochemical charge separation between excited P, P*, and QA is no longer possible. In this situation, fluorescence intensity is determined by the state of energy dissipation. Fluorescence is at the Fm level, when mechanisms of thermal energy dissipation have not yet been activated. It is at the lower F'm level when thermal energy dissipation competes effectively with fluorescence emission. Non-photochemical fluorescence quenching 
describes the extent of energy dissipation. However, under desiccation, F'm is often not distinguishable from F'o, which signifies a quenched Fo level (see Figs. 4 and 5). In this case, 
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Thus, regulation of energy dissipation can be monitored by following the changes in fluorescence, provided proper precautions are taken to avoid interference by photochemical fluorescence quenching.
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Use of light for photosynthesis |
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In rate-limiting light, almost all absorbed quanta are used for photosynthesis by hydrated plants. As a result, the initial slope of photosynthesis versus photon flux is practically identical in many different plants. As the light intensity is increased, the slope decreases. In fully sun-exposed leaves, light is usually far in excess. Practically all absorbed light energy needs to be converted into heat in desiccated poikilohydric autotrophs if oxidative damage is to be avoided. This requires highly effective mechanisms of thermal energy dissipation. In fact, the quantum yield of fluorescence is very low in desiccated poikilohydric plants. Fluorescence is strongly quenched. It increases upon hydration indicating rapid deactivation of the mechanisms which are responsible for fluorescence quenching in the desiccated state. Sensitive regulation of energy dissipation ascertains the dominance of photosynthetic energy conservation after water has become available. Protons play a key role both in photosynthesis and in the opposing process of energy dissipation. When two molecules of water are oxidized inside the closed thylakoid membrane system of chloroplasts, four protons remain inside the thylakoids. Eight more are pumped into the thylakoids as the four electrons liberated during water oxidation are on their path to reduce one molecule of CO2 (Fig. 2). Efflux of these protons lowers the acidification of the intrathylakoid space and is sufficient, or almost sufficient (Allen, 2003a
), to power the endergonic synthesis of three molecules of ATP which are needed for the reduction of one CO2 in the so-called C3 pathway of carbon assimilation. Thus, influx of protons into and efflux from the thylakoids are largely balanced during carbon reduction. Thylakoid acidification is low because a relatively small proton gradient satisfies the ATP requirements of carbon assimilation. The alternative electron transport pathways of cyclic electron transport and electron flow to oxygen support stronger thylakoid acidification (Allen, 2003a; Heber, 2003). They are regulated by the redox state of electron carriers. By permitting protonation of the PsbS protein, they activate thermal energy dissipation as will be shown below.
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Thermal energy dissipation: role of light-harvesting chlorophyll–protein complexes |
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Light in excess of that used for photosynthesis is either re-emitted as fluorescence or thermally dissipated so that fluorescence decreases. When the main light-harvesting complex of the photosynthetic membrane, LHCII is isolated in a detergent solution, it is strongly fluorescent. Upon aggregation, fluorescence is quenched. Fluorescence life-times decrease accordingly from about 4 ns to values ranging between 100 ps and 1.5 ns (Mullineaux et al., 1993
; Moya et al., 2001). Fluorescence is also quenched in LHCII crystals (Pascal et al., 2005). The fluorescence life-time was 890 ps compared to 4 ns for LHCII trimers. In attempts to find a molecular basis for regulated energy dissipation, aggregation phenomena are proposed to lead to observed fluorescence quenching in intact organisms (Horton et al., 1991, 2005). Still, the energy dissipation indicated by a decrease of fluorescence life-times in isolated chlorophyll–protein complexes from 4 ns to about 100 ps or more is insufficient to drain excitation energy sufficiently fast from open PSII RCs to afford photoprotection of PSII RCs. More effective mechanisms of energy dissipation are observed in intact organisms. |
Role of zeaxanthin in thermal energy dissipation by hydrated plants |
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Although it has long been known that the concentration of the xanthophyll zeaxanthin in photosynthetic membranes of higher plants reversibly increases in the light at the expense of violaxanthin (Yamamoto et al., 1962
; Sapozhnikov et al., 1966), a role of zeaxanthin in thermal energy dissipation has been proposed by Barbara Demmig at the University of Würzburg only in 1987 (Demmig-Adams, 1990). By now, an impressive amount of documentation has become available not only on the wide distribution of zeaxanthin-dependent thermal energy dissipation within green plants, but also on its regulation (Gilmore and Govindjee, 1999; Niyogi, 1999; Ruban et al., 2002; Ma et al., 2003; Holt et al., 2004, 2005; Horton et al., 2005). In summary, a special protein in the antenna of PSII, the gene product of the PsbS gene, is thought to change its conformation on protonation. Once protonated, it is capable of forming a complex with zeaxanthin which is active as an effective sink for excitation energy. Sensitive regulation (Li et al., 2004) ensures that assimilation has precedence over thermal energy dissipation. Whereas linear light-dependent electron transport from water to CO2 as depicted in Fig. 2 is accompanied by balanced deposition of protons inside thylakoids and proton efflux from the thylakoids, additional thylakoid acidification by proton-coupled cyclic and pseudocyclic electron transport (Allen, 2003a; Heber, 2003) provides protons for protonation of the PsbS-protein when CO2 becomes rate-limiting for carbon reduction. Also, a low intrathylakoid pH is known to cause the de-epoxidation of the xanthophyll violaxanthin to zeaxanthin (Pfündel and Dilley, 1993). Availability of zeaxanthin and protonation of the PsbS-protein make it possible to form energy-dissipating centres which drain excitation energy from PSII RCs.
However, activation of energy dissipation by protonation is not necessarily based on light-dependent electron transport. It has recently been shown that CO2 which, at a pK of 6.31, acts as a very weak protonating agent, is capable of promoting non-photochemical quenching, NPQ, in a moss, provided some zeaxanthin is present (Bukhov et al., 2001a). Similar observations were made with chlorolichens (Heber et al., 2000; Kopecky et al., 2005). This is surprising because protonation of the amino acids E122 and E226 of the PsbS-protein (Li et al., 2002) is expected to require a much lower pH than can be provided by CO2. NPQ formation increased and the quantum efficiency of charge separation in PSII RCs decreased with increasing CO2 concentration (Bukhov et al., 2001a). Actinic light was not necessary. Removal of the CO2 reversed the NPQ and increased the quantum efficiency of charge separation in PSII RCs. Dithiothreitol, an inhibitor of zeaxanthin formation (Yamamoto and Kamite, 1972) inhibited NPQ formation by CO2. All this showed that protonation governed energy dissipation. The presence of zeaxanthin was essential, but a few molecules of zeaxanthin per PSII RC were sufficient for effective competition with open RCs for excitation energy (Bukhov et al., 2001a).
Importantly, leaves which possess the zeaxanthin cycle were very slow to respond to high concentrations of CO2 by increasing NPQ or decreasing charge separation in PSII RCS when actinic light was absent. By contrast, responses of the moss Rhytidiadelphus to CO2 were fast and dramatic (Bukhov et al., 2001b). Figure 3 compares the extent of the reversible suppression of charge separation by CO2 in the moss Rhytidiadelphus with that in a spinach leaf in the absence of actinic light. Apparently, regulation of zeaxanthin-dependent energy dissipation by protonation was different in the moss (or in lichens, Kopecky et al., 2005) on one side and in higher plants such as spinach on the other side. Also, protonation by CO2 suppressed Fo fluorescence in mosses and lichens, but not in spinach or other higher plants. Maximum rates of photosynthesis in the different groups of photosynthetic organisms are known to be different (Green and Lange, 1995; Lange, 2003; Larcher, 2003). Maximum charge separation in PSII RCs as expressed by 
F/Fm is also different. It is consistently lower in lichens and mosses than in higher plants (Jensen, 2002). Apparently, the balance between energy conservation and energy dissipation is tilted towards dissipation in many poikilohydric autotrophs, whereas, in higher plants, energy conservation assumes dominance over energy dissipation. It thus appears that sensitivity to excess light is higher in the mosses and the lichens than in higher plants. More sensitive regulation of thermal energy dissipation ensures protection against oxidative damage in these organisms.
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Thermal energy dissipation in desiccated poikilohydric autotrophs: simple consequence of drying or the result of the activation of a special mechanism? |
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After hydrated lichens or phototolerant poikilohydric mosses have been slowly desiccated, variable fluorescence
F=(Fm–Fo) is lost and even Fo fluorescence is more or less dramatically decreased (see F'o in Figs 4 and 5), depending on species and season. Whereas full quenching of F increases NPQ in hydrated plants from zero to less than 5 if Fo remains unchanged, NPQ may increase to values of more than 50 during the desiccation of some lichens. The increase in thermal energy dissipation revealed by the loss of fluorescence yield is readily reversed by hydration.
Importantly, drying of leaves, or of poikilohydric mosses, when these are in a photosensitive state, does not result in much fluorescence quenching (Kopecky et al., 2005; Heber and Shuvalov, 2005). This shows that the large loss of fluorescence during desiccation of phototolerant poikilohydric organisms is not a simple result of drying a photosynthetic membrane. Rather, it indicates the activation of a specific mechanism, or of specific mechanisms, of energy dissipation.
Whereas many lichens are associated with a green alga, others harbour a cyanobacterium as the photobiont. Only those with a green alga possess a zeaxanthin cycle (Demmig-Adams et al., 1990). Nevertheless, both types of lichens exhibit strong desiccation-induced fluorescence quenching. Figure 4 compares modulated chlorophyll fluorescence of the desiccated chlorolichen Hypogymnia physodes and the cyanolichen Peltigera neckeri before and after hydration. A modulated measuring beam of a very low average intensity (less than 0.005% of sunlight) increased chlorophyll fluorescence to the level F'o. After the initial fluorescence rise two short light pulses of extremely strong actinic light (PPFD=13 000 µmol m–2 s–1, i.e. five times sunlight) were given while the lichens were still dry. Although they were strong enough to excite all chlorophylls several times, including those located within the RCs, they caused no appreciable fluorescence responses. Apparently, stable charge separation was suppressed in the desiccated lichens. Hydration increased fluorescence strongly. Reversible pulse-induced fluorescence changes demonstrated that PSII RCs became rapidly functional after hydration. After the Fo level had risen during hydration, fluorescence was transiently quenched below the elevated Fo level immediately after the 1 s light pulses only in the Hypogymnia experiment of Fig. 4A, but not in the Peltigera experiment of Fig. 4B. Transient pulse-induced Fo quenching is typical for the presence of active zeaxanthin-dependent energy dissipation (Katona et al., 1992). The cyanolichen does not possess a zeaxanthin cycle.
Two different pathways of thermal energy dissipation in poikilohydric autotrophs which possess a zeaxanthin cycle
As long as sufficient light is available and hydration still supports proton-coupled electron transport in poikilohydric autotrophs which possess the zeaxanthin cycle, zeaxanthin-dependent energy dissipation is active while water is slowly lost. Dissipating centres not only appear to survive desiccation but also to remain functional in the fully desiccated state (Eickmeier et al., 1993; Deltoro et al., 1998; Kopecky et al., 2005). Nevertheless, desiccation-induced thermal energy dissipation does not depend on the presence of zeaxanthin even in poikilohydric organisms which possess the zeaxanthin cycle. This is shown in the experiment of Fig. 5 for the sun-tolerant moss Rhytidium rugosum, but can also be demonstrated in chlorolichens. The moss was taken sun-dried from a sun-exposed habitat and used for the experiment in Fig. 5A. A parallel sample was hydrated and kept for 2 d in dim light in order to decrease or eliminate zeaxanthin. It was then slowly dried in darkness and taken for the experiment in Fig. 5B. A low-intensity measuring beam increased chlorophyll fluorescence of the desiccated mosses less in Fig. 5A than in Fig. 5B, indicating that chlorophyll fluorescence was more strongly quenched in the sun-dried moss than in the moss which had been dried in darkness. Two very strong 1 s light pulses were as ineffective in causing appreciable fluorescence responses in the desiccated mosses as they were in the lichen experiments of Fig. 4. Apparently, stable charge separation was suppressed, not only in the lichens but also in the desiccated mosses.
Hydration by a drop of water changed the fluorescence responses. A small rapid initial lowering of fluorescence appeared to have optical reasons, but the following increase is based on the recovery of function. Quenching was reversed by hydration. Pulse-induced fluorescence now indicated charge separation in the PSII RCs and transient reduction of oxidized QA both in the sun-dried and the dark-dried moss. However, the extent of charge separation as indicated by the size of fluorescence responses was smaller in the sun-dried moss than in the dark-dried moss (F/Fm=0.14 and 0.42, respectively, in Fig. 5A and B shortly after hydration). Appreciable transient Fo quenching following the strong light pulses occurred only in Fig. 5A suggesting the presence of active zeaxanthin-dependent energy dissipation (Katona et al., 1992; Kopecky et al., 2005). It was very small or absent in Fig. 5B. Apparently, in the sun-dried moss, zeaxanthin had been present. Lowered Fo fluorescence in Fig. 5A compared with Fo fluorescence in Fig. 5B suggests that it had contributed to photoprotection during desiccation. Nevertheless, the large reversal of quenching by hydration in the dark-dried moss also reveals considerable desiccation-induced thermal energy dissipation which does not depend on the presence of zeaxanthin.
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Acquisition and loss of phototolerance in poikilohydric mosses |
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When collected in rainy weather during the autumn or winter seasons and then desiccated in the dark, the shade-adapted moss Rhytidiadelphus squarrosus was found to be photosensitive. Desiccation did not result in appreciable fluorescence quenching (Heber and Shuvalov, 2005
). Remoistening failed to increase Fo fluorescence. When the moss was collected in the spring or summer, fluorescence decreased strongly whether or not light was present during desiccation. Hydration reversed the fluorescence quenching (see also Fig. 5 for Rhytidium rugosum). Laboratory experiments with different shade-adapted mosses species established interchangeability between the different fluorescence responses (U Heber, VA Shuvalov, unpublished experiments). Exposing a phototolerant ‘summer’ moss in the hydrated state for prolonged periods of time to dim light before drying it in darkness made it photosensitive, as shown by increased sensitivity to strong irradiation in the desiccated state. Damage became manifest by the irreversible loss of fluorescence characteristics not only after hydration but also in the desiccated state. Conversely, drying a photosensitive ‘winter’ moss under illumination made it more phototolerant, as shown by decreased sensitivity to strong irradiation, whether or not zeaxanthin was present. |
Photosystem II reaction centres as possible sites of desiccation-induced energy dissipation |
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Activation of desiccation-induced thermal energy dissipation during drying of shade-adapted mosses requires the interaction of light and desiccation. A main factor involved in zeaxanthin-dependent energy dissipation is protonation of the PsbS protein. However, protonation does not appear to be involved in desiccation-induced energy dissipation because the protonophore nigericin did not prevent fluorescence quenching during drying. Another possibility to explain the light requirement of the acquisition of phototolerance is redox regulation which is known to be common in plant metabolism (Allen, 2003b
). Illumination changes the redox state of several components of the photosynthetic apparatus. In order to restrict reduction to a specific site, the electron transport inhibitor DCMU was used. It is known to block electron transport selectively between QA and QB in the RC of PSII (Trebst, 1986). This restricts reduction to a component of the RC such as QA. Importantly, desiccation of a DCMU-inhibited photosensitive moss in the presence of light, which was sufficient to reduce QA in the RC of PSII, but insufficient to induce zeaxanthin synthesis, increased phototolerance in Rhytidiadelphus squarrosus (U Heber and VA Shuvalov, unpublished data). The experiment suggested the RC of PSII as the site of desiccation-induced energy dissipation, but does not rule out the possibility of a light requirement for conformational changes of chlorophyll–protein complexes which may be involved in desiccation-induced energy dissipation. |
Spectral changes in fluorescence emission during desiccation |
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Spectral changes in fluorescence excitation and emission caused by desiccation of a chlorolichen and a cyanobacterium have been investigated by Bilger et al. (1989)
. The results of low temperature fluorescence spectroscopy were interpreted to indicate interruption of functional energy transfer from the antenna of photosystem II or from accessory pigments to PSII RCs which were considered inactive. These conclusions now appear to be in question in view of more recently observed RC activity in dried leaves (Shuvalov and Heber, 2003; Kopecky et al., 2005) and desiccated mosses (Heber and Shuvalov, 2005; see also below). Nevertheless, fluorescence spectroscopy still provides valuable information on energy dissipation. Figure 6 shows fluorescence emission spectra of two desiccated mosses. The excitation wavelength was 440 nm. In the lower spectrum of Fig. 6A for sun-dried Hypnum, fluorescence emission peaked at about 720 nm. The 685 nm band of fluorescence emission shown in the upper spectrum was largely absent. By contrast, when zeaxanthin had been converted to violaxanthin by exposure to dim light before drying Hypnum in the dark, a large maximum peaking at 685 nm dominated the upper spectrum in Fig. 6A. The 720 nm emission was still present. In sun-dried Rhytidium (Fig. 6B, lower spectrum), the 685 nm emission was less strongly suppressed than in sun-dried Hypnum. When Rhytidium was dried in the dark after previous exposure to dim light to decrease zeaxanthin (upper spectrum of Fig. 6B), the 685 nm peak was as large as in the Hypnum experiment of Fig. 6A). The 720 nm emission was also present. It is important to note that the spectra were taken at room temperature where photosystem I is known to be largely non-fluorescent. Therefore, emission at 720 nm is thought to originate from PSII. In fact, excitation with monochromatic far-red light up to about 740 nm was shown to result in fluorescence emission from PSII (Heber and Shuvalov, 2005). The observations shown in Fig. 6 suggest that, in desiccated mosses, zeaxanthin-dependent energy dissipation drains excitation energy preferably from centres which emit 685 nm fluorescence.
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On heating desiccated mosses and lichens, fluorescence increased reversibly. Charge separation in PSII RCs returned indicating temperature-induced reversal of thermal energy dissipation (Heber and Shuvalov, 2005
). From a comparison of the shift of fluorescence emission towards 720 nm in desiccated mosses (Fig. 6) and lichens with the activation energy of the reversible part of the temperature-dependent fluorescence increase, it was suggested that energy is dissipated by dissipation centres which emit 720 nm fluorescence with a very low quantum yield. |
Electron transport in PSII RCs in the absence of water |
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Even very low light raised the fluorescence of dry leaves (Kopecky et al., 2005
) or desiccated photosensitive mosses above the Fo level indicating efficient energy migration to PSII RCs and reduction of QA. Once reduced, QA remained reduced in the dark or reoxidized only slowly. It could be permanently kept in the fully reduced state by low intensity background light. This was used to separate other photoreactions of PSII from electron transport to QA (Shuvalov and Heber, 2003). Light-dependent bleaching around 500 nm showed that ß-carotene (Car), a constituent of PSII RCs, was photo-oxidized with a low quantum yield, not only in dried spinach leaves (Fig. 7), but also in dried spinach chloroplasts or subchloroplast particles. Oxidation was almost proportional to light intensity up to PPFDs close to sunlight. While ß-carotene was photo-oxidized, kinetically related formation of bands at 450 and 810 nm and bleaching at 436 and 680 nm suggested the accumulation of a reduced chlorophyll. Sieve effects, which are stronger in the blue than in the green region of the spectrum, made quantitative analysis of the relationship between chlorophyll reduction and carotene oxidation difficult in dry leaves. However, spectra obtained with subchloroplast particles suggested a ratio Chl–/Car+ close to 1. No indication was obtained for the photoaccumulation of reduced pheophytin. Also, redox changes around 559 nm were notably absent suggesting that cytochrome b559 in the RC of PSII remained in the oxidized state while reversible redox reactions of ß-carotene and chlorophyll were observed. On darkening, Car+ was reduced and Chl– was oxidized with half-times below 1 s (Shuvalov and Heber, 2003).
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The data were interpreted as showing that a chlorophyll next to P680 is the primary electron acceptor in the RC of PSII rather than pheophytin, up to now considered the primary electron acceptor in higher plants. Charge separation in PSII resulted in the initial formation of the radical pair P680+Chl–. After Car was oxidized in the dry leaves by P680+, reduction of Car+ by Chl– completed an electron cycle within the RC of PSII (Fig. 8). By decreasing the concentration of P680+, both the fast recombination of the closely spaced radical pair P680+Chl– and the slower recombination between the more widely spaced Car+ and Chl– within PSII RCs are photoprotective reactions. However, the slower recombination reaction is not expected to contribute significantly to energy dissipation. Details on spatial relations in the PSII RC have been published by Kamiya and Shen (2003)
. Edge to edge distances between P680 and ChlD1 are about 5 Å, between P680 and PheoD1 about 9 Å, and between ChlD1 and Car about 23 Å (Christian Fufezan, University of Freiburg, Germany, personal communication). The recombination P680+Chl–
P680 Chl is thought to compete with and actually eliminate, in desiccated organisms, the recombination of the pair P680+Pheo– which is known to be accompanied by fluorescence emission and by the formation of highly reactive singlet oxygen (Krieger-Liszkay, 2005).
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Transmittance spectra very similar to the spectrum shown in Fig. 7 for a dried spinach leaf were also obtained with the poikilohydric fern Polypodium vulgare (Shuvalov and Heber, 2003
) and with the desiccated moss Rhytidiadelphus squarrosus, but only after the moss had been slowly dried in the dark. Owing to strong sieve effects, the fern and moss spectra were well resolved only in the 500 nm region. Attempts to obtain transmission spectra from sun-dried mosses were unsuccessful because stable charge separation in the RCs was suppressed as a consequence of increased desiccation-induced energy dissipation. Desiccated lichens do not transmit weak measuring light. However, reflectance could be used to obtain information on photoreactions. Strong desiccation-dependent quenching made it necessary to use extremely high photon flux densities for analysis. In this situation, light-dependent band formation around 800 nm was observed. Dark relaxation of the band had several phases. Since both chlorophyll cations and anions absorb in this region, no reliable assignments could be made, although a contribution of P700+ could be excluded by doing measurements in the presence of a background of far-red light.
Formation of an absorption band was also observed close to 950 nm where carotenes are known to absorb (Tracewell et al., 2001). Absorption increased in mosses and lichens in proportion to light intensity as carotene oxidation had done in dried spinach leaves (Shuvalov and Heber, 2003) showing that the quantum efficiency of oxidation was low. The reaction was observed whether or not attempts had been made to eliminate zeaxanthin. Dark relaxation was too fast to be resolved by the available instrumentation. The reflectance data suggest that the electron cycle within the RC of PSII, which has been observed in dry spinach leaves, is faster in desiccated mosses and lichens.
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Significance of Fo suppression: time scale of thermal energy dissipation |
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As has been mentioned above, Fo fluorescence elicited by very low PPFDs of preferably less than 0.1 µmol m–2 s–1 describes a situation in which essentially all PSII RCs are open in hydrated photosynthetic organisms while QA is oxidized. Minimum or Fo fluorescence is in equilibrium with energy capture by the reaction centres, which is known to result in charge separation within about 3 ps (Zinth and Kaiser, 1993
). Nevertheless, protonation by high concentrations of CO2 quenched Fo fluorescence in hydrated mosses and lichens by activating zeaxanthin-dependent energy dissipation (Heber et al., 2000; Bukhov et al., 2001a). This proves that the dissipating centres formed as a result of protonation were capable of draining excitation energy even from open PSII reaction centres. It means simultaneously that energy dissipation was faster than energy capture by PSII RCs. Fo fluorescence is quenched not only by activating zeaxanthin-dependent energy dissipation in hydrated mosses and lichens, but more strongly in desiccated phototolerant mosses, chlorolichens and cyanolichens, whether or not zeaxanthin is present, and whether or not drying is done in near darkness to keep PSII RCs in the open state. It was concluded that desiccation of poikilohydric autotrophs results in energy loss from dissipating centres which is much faster than energy trapping by normally functioning PSII RCs.
Using leaves of transgenic Arabidopsis thaliana, Holt et al. (2005) concluded from measurements of transient absorption changes in the picosecond range, that charge separation in an activated dissipation centre consisting of the protonated PsbS-protein, zeaxanthin, and chlorophyll results, within about 1 ps, in zeaxanthin oxidation and chlorophyll reduction:
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Zeaxanthin-dependent energy dissipation is thought to be localized in the antenna of PSII (Horton et al., 2005; but see Holt et al., 2004), but Finazzi et al. (2004) have recently demonstrated a zeaxanthin-independent energy dissipation pathway in hydrated leaves which is localized in or near PSII RCs. Nothing definite is known yet about the site of desiccation-induced energy dissipation in desiccated poikilohydric autotrophs, but there are arguments to suggest that it also proceeds in PSII RCs where charge separation results in the oxidation of P680 and, initially, in the reduction of a chlorophyll (Shuvalov and Heber, 2003):
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forms the molecular basis of zeaxanthin-dependent energy dissipation. The simultaneous occurrence of two photoprotective pathways in desiccated chlorolichens (Kopecky et al., 2005) and mosses and the strong suppression of Fo fluorescence in these organisms suggest that RC-based energy dissipation operates in a picosecond time domain not very different from that reported by Holt et al. (2005) for zeaxanthin-dependent energy dissipation. |
Concluding remarks |
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During the first decennium of modern photosynthesis research, the main emphasis was on the elucidation of the principal pathways of electron transport and of the reduction of CO2 and other physiological electron acceptors. Later, much attention was focused on the mechanisms involved in regulating the pathways. Always, the use of light energy for biosynthetic purposes was in the foreground of interest although it was soon recognized that light can also damage the photosynthetic apparatus. No mutants devoid of carotenoids have ever been observed which can survive illumination in an oxygen-containing atmosphere. Also, agents interfering with carotenoid biosynthesis are effective herbicides. Although this emphasized the role of carotenoids in providing protection against light-sensitized oxidation, the existence of physiological mechanisms able to regulate not only photosynthesis but also the thermal degradation of excess light energy was overlooked until fluorescence became a widely applied tool in photosynthesis research. Then it became obvious that changes in chlorophyll fluorescence were not only related to photochemical light use but also gave information on hitherto unknown mechanisms involved in protecting the photosynthetic apparatus against radical damage (Krause and Weis, 1991
; Krause and Jahns, 2004). As long as water is available and photon flux is low, light energy absorbed by the light-harvesting pigment system of PSII is trapped primarily by PSII RCs where charge separation takes place, initiating energy conservation. Water is oxidized and oxygen is evolved (Fig. 1). Zeaxanthin-dependent thermal energy dissipation is activated in the antenna of PSII only at higher photon flux densities by protonation of the PsbS protein. This occurs when alternative electron transport processes decrease the intrathylakoid pH so as to permit protein protonation. This governs thermal energy dissipation under conditions of hydration. Once they have been activated, zeaxanthin-containing dissipation centres remain functional even after desiccation. Loss of water during desiccation of poikilohydric autotrophs activates another mechanism of thermal energy dissipation which, like zeaxanthin-dependent energy dissipation, finds expression in reduced chlorophyll fluorescence. It is proposed to be localized in the RCs of PSII. Loss of water during desiccation, in combination with the reduction of an RC component such as QA, appears to change the conformation of PSII RCs so as to transform them from energy-conserving to energy-dissipating centres. Figure 8 compares the established view of linear electron transport in PSII RCs of hydrated photosynthetic organisms with electron transport in the RCs of desiccated poikilohydric autotrophs. The latter is thought to form the basis for desiccation-induced thermal energy dissipation. Together with zeaxanthin-dependent energy dissipation, energy dissipation in PSII RCs protects poikilohydric mosses and chlorolichens from photo-oxidation when water is absent. Both mechanisms are important for their ability to survive full sunlight for prolonged periods of time in the desiccated state. Cyanolichens appear to be protected primarily by the desiccation-activated mechanism.
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Acknowledgements |
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Our research was supported by Deutsche Forschungsgemeinschaft, Fonds der Chemischen Industrie and Russian Fund for Basic Research. Professor Hedrich, Würzburg, provided the working facilities. We also acknowledge co-operation with Drs Pfündel, Würzburg, and Bukhov and Azarkovich, Moscow. Dr Zellner, Würzburg, kindly helped in identifying several species of mosses and lichens.
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References |
|---|
Allen JF. 2003a. Cyclic, pseudocyclic and noncyclic photophosphorylation: new links in the chain. Trends in Plant Science 8, 15–19.[CrossRef][Web of Science][Medline]
Allen JF. 2003b. The function of genomes in bioenergetic organelles. Philosophical Transactions of the Royal Society: Biological Sciences 358, 19–38.
Alpert P. 2000. The discovery, scope, and puzzle of desiccation tolerance in plants. Plant Ecology 151, 5–17.[CrossRef]
Asada K. 1999. The water–water cycle in chloroplasts: scavenging of active oxygens and dissipation of excess photons. Annual Review of Plant Physiology and Plant Molecular Biology 50, 601–639.[CrossRef][Web of Science]
Bilger W, Rimke S, Schreiber U, Lange OL. 1989. Inhibition of energy transfer to photosystem II in lichens by dehydration: different properties of reversibility with green and blue-green photobionts. Journal of Plant Physiology 134, 261–268.
Björkman O, Demmig-Adams B. 1995. Regulation of photosynthetic light energy capture, conversion, and dissipation in leaves of higher plants. In: Schulze E-D, Caldwell MM, eds. Ecophysiology of photosynthesis. Ecological studies, 100. Berlin: Springer, 7–47.
Bradbury M, Baker NR. 1981. Analysis of the slow phase of the in vivo fluorescence induction curve. Changes in the redox state of photosystem II electron acceptors and fluorescence emission from photosystems I and II. Biochimica et Biophysica Acta 635, 542–551.[Medline]
Bruce D, Samson G, Carpenter C. 1997. The origins of nonphotochemical quenching of chlorophyll fluorescence in photosynthesis. Direct quenching by P680+ in photosystem II enriched membranes at low pH. Biochemistry 36, 749–755.[Medline]
Bukhov NG, Kopecky J, Pfündel EE, Klughammer C, Heber U. 2001a. One molecule of zeaxanthin per reaction centre of photosystem II permits effective thermal dissipation of light energy in a poikilohydric moss. Planta 212, 739–748.[CrossRef][Web of Science][Medline]
Bukhov NG, Heber U, Wiese C, Shuvalov VA. 2001b. Energy dissipation in photosynthesis: quenching of chlorophyll fluorescence in reaction centers and antenna complexes. Planta 212, 749–758.[CrossRef][Web of Science][Medline]
Deltoro VL, Calatayud A, Gimeno C, Abadia A, Barreno E. 1998. Changes in chlorophyll a fluorescence, photosynthetic CO2 assimilation and xanthophyll cycle interconversions during dehydration in desiccation-tolerant and intolerant liverworths. Planta 207, 224–228.[CrossRef]
Demmig-Adams B. 1990. Carotenoids and photoprotection of plants: a role for the xanthophyll zeaxanthin. Biochimica et Biophysica Acta 1020, 1–24.[CrossRef]
Demmig-Adams B, Máguas C, Adams III WW, Meyer A, Kilian E, Lange OL. 1990. Effect of high light on the efficiency of photochemical energy conversion in a variety of lichen species with green and blue-green phycobionts. Planta 180, 400–409.[CrossRef]
Eickmeier WC, Casper C, Osmond CB. 1993. Chlorophyll fluorescence in the resurrection plant Selaginella lepidophylla (Hook. & Grev.) during high light and desiccation stress, and evidence for zeaxanthin-associated photoprotection. Planta 189, 30–38.
Evans DE, Lange OL. 2003. Biological soil crusts and ecosystem nitrogen and carbon dynamics. In: Belnap J, Lange OL, eds. Biological soil crusts: structure, function, and management. Ecological studies, 150. Berlin, Heidelberg, New York: Springer, 263–279.
Finazzi G, Johnson NG, Dallosto L, Joliot P, Wollman F-A, Bassi R. 2004. A zeaxanthin-independent nonphotochemical quenching mechanism localized in the photosystem II core complex. Proceedings of the National Academy of Sciences, USA 101, 12375–12380.
Gauslaa Y, Solhaug KA. 1999. High-light damage in air-dry thalli of the old forest lichen Lobaria pulmonaria: interactions of irradiance, exposure duration and high temperature. Journal of Experimental Botany 50, 697–705.
Gauslaa Y, Solhaug KA. 2004. Photoinhibition in lichens depends on cortical characteristics and hydration. Lichenology 36, 133–143.[CrossRef]
Gauslaa Y, Ohlson M, Solhaug KA, Bilger W, Nybakken L. 2001. Aspect-dependent high-irradiance damage in two transplanted foliose forest lichens, Lobaria pulmonaria and Parmelia sulcata. Canadian Journal of Forest Research 31, 1639–1649.[CrossRef]
Gilmore AM, Govindjee. 1999. How higher plants respond to excess light: energy dissipation in photosystem II. In: Singhal GS, Renger G, Sopory SK, Irrgang K-D, Govindjee, eds. Concepts in photobiology: photosynthesis and photomorphogenesis. New Delhi: Narosa Publishing House, 513–548.
Green TGA, Lange OL. 1995. Photosynthesis in poikilohydric plants: a comparison of lichens and bryophytes. In Schulze E-D, Caldwell MM, eds. Ecophysiology of photosynthesis. Ecological studies, 100. Berlin: Springer, 319–341.
Heber U. 2003. Irrungen, Wirrungen? The Mehler reaction in relation to cyclic electron transport in C3 plants. Photosynthesis Research 73, 223–231.
Heber U, Bilger W, Bligny R, Lange OL. 2000. Photoreactions in two lichens, a poikilohydric moss and higher plants in relation to phototolerance of alpine plants: a comparison. Planta 211, 770–780.[CrossRef][Web of Science][Medline]
Heber U, Bukhov NG, Shuvalov VA, Kobayashi Y, Lange OL. 2001. Protection of the photosynthetic apparatus against damage by excessive illumination in homoiohydric leaves and poikilohydric mosses and lichens. Journal of Experimental Botany 52, 1999–2006.
Heber U, Shuvalov VA. 2005. Photochemical reactions of chlorophyll in dehydrated photosystem II: two chlorophyll forms (680 and 700 nm). Photosynthesis Research 84, 85–91.[CrossRef][Web of Science][Medline]
Holt NE, Fleming GR, Niyogi KK. 2004. Toward an understanding of the mechanism of nonphotochemical quenching in green plants. Biochemistry 43, 8281–8289.[CrossRef][Medline]
Holt NE, Tigmantas D, Valkunas L, Li X-P, Niyogi KK, Fleming GR. 2005. Carotenoid cation formation and the regulation of photosynthetic light harvesting. Science 307, 433–436.
Horton P, Ruban AV, Rees D, Noctor G, Pascal AA, Young A. 1991. Control of the light-harvesting function of chloroplast membranes by aggregation of the LHCII chlorophyll protein complex. FEBS Letters 292, 1–4.[CrossRef][Web of Science][Medline]
Horton P, Wentworth M, Ruban A. 2005. Control of the light-harvesting function of chloroplast membranes: the LHCII-aggregation model for non-photochemical quenching. FEBS Leters 579, 4201–4206.
Jensen M. 2002. Measurement of chlorophyll fluorescence in lichens. In: Kranner I, Beckett R, Varma A, eds. Protocols in lichenology. Berlin, Heidelberg, New York: Springer Verlag, 135–151.
Kamiya N, Shen J-R. 2003. Crystal structure of oxygen-evolving photosystem II from Thermosynechococcus vulcanus at 3.7 Å resolution. Proceedings of the National Academy of Sciences, USA 100, 98–103.
Katona E, Neimanis S, Schönknecht G, Heber U. 1992. Photosystem I-dependent cyclic electron transport is important in controlling photosystem II activity in leaves under conditions of water stress. Photosynthesis Research 34, 449–464.[CrossRef]
Kopecky J, Azarkovich M, Pfündel EE, Shuvalov VA, Heber U. 2005. Thermal dissipation of light energy is regulated differently and by different mechanisms in lichens and higher plants. Plant Biology 7, 156–167.[CrossRef][Medline]
Krause GH, Jahns P. 2003. Pulse amplitude modulated chlorophyll fluorometry and its application in plant science. In: Green BR, Parson WW, eds. Light-harvesting antennae in photosynthesis. Dordrecht: Kluwer Academic Publishers, 373–399.
Krause GH, Weis E. 1991. Chlorophyll fluorescence and photosynthesis: the basics. Annual Review of Plant Physiology and Plant Molecular Biology 42, 313–349.[CrossRef][Web of Science]
Krause GH, Jahns P. 2004. Non-photochemical energy dissipation determined by chlorophyll fluorescence quenching: characterization and function. In: Papageorgiou GC, Govindjee, eds. Chlorophyll a fluorescence: a signature of photosynthesis. Springer, 463–495.
Krieger A, Weiss E. 1993. The role of calcium in the pH-dependent control of photosystem II. Photosynthesis Research 37, 117–130.[CrossRef]
Krieger-Liszkay A. 2005. Singlet oxygen production in photosynthesis. Journal of Experimental Botany 56, 337–346.
Lange OL. 2003. Photosynthesis of soil-crust biota as dependent on environmental factors. In: Belnap J, Lange OL, eds. Biological soil crusts: structure, function and management. Ecological studies, 150. Berlin: Springer, 217–240.
Larcher W. 2003. Physiological plant ecology. Ecophysiology and stress physiology of functional groups. Berlin: Springer.
Li X-P, Phippard A, Pasari J, Niyogi KK. 2002. Structure–function analysis of photosystem II subunit S (PsbS) in vivo. Functional Plant Biology 29, 1131–1139.[CrossRef]
Li X-P, Gilmore AM, Caffari S, Bassi R, Golan T, Kramer D, Niyogi KK. 2004. Regulation of photosynthetic light harvesting involves intrathylakoid lumen pH sensing by the PsbS protein. Journal of Biological Chemistry 279, 22866–22874.
Ma Y-Z, Holt NE, Li X-P, Niyogi KK, Fleming GR. 2003. Evidence for direct carotenoid involvement in the regulation of photosynthetic light harvesting. Proceedings of the National Academy of Sciences of the USA 100, 4377–4382.
Moya I, Silvestri M, Vallon O, Cinque G, Bassi R. 2001. Time-resolved fluorescence analysis of the photosystem II antenna proteins in detergent micelles and liposomes. Biochemistry 40, 12552–12561.[CrossRef][Medline]
Mullineaux CW, Pascal AA, Horton P, Holzwarth AR. 1993. Excitation-energy quenching in aggregates of the LHCII chlorophyll–protein complex: a time-resolved fluorescence study. Biochimica et Biophysica Acta 1141, 23–28.
Niyogi KK. 1999. Photoprotection revisited: genetic and molecular approaches. Annual Review of Plant Physiology and Plant Molecular Biology 50, 333–359.[CrossRef][Web of Science]
Owens TG. 1996. Processing of excitation energy by antenna pigments. In: Baker NR, ed. Photosynthesis and the environment. Dordrecht: Kluwer, 1–23.
Pascal AA, Liu Z, Broess K, van Oort B, van Amerongen H, Wang C, Horton P, Robert B, Chang W, Ruban A. 2005. Molecular basis of photoprotection and control of photosynthetic light-harvesting. Nature 436, 134–137.[CrossRef][Medline]
Pfündel EE, Dilley RA. 1993. The pH-dependence of violaxanthin deepoxidation in isolated pea chloroplasts. Plant Physiology 101, 65–71.[Abstract]
Ruban AV, Pascal AA, Robert B, Horton P. 2002. Activation of zeaxanthin is an obligatory event in the regulation of photosynthetic light harvesting. Journal of Biological Chemistry 277, 7785–7789.
Sapozhnikov DJ, Kolotova LR, Giller YE. 1966. Action spectrum of de-epoxidation of violaxanthin. Doklady Akademia Nauk SSSR 171, 740–741.
Schreiber U, Schliwa U, Bilger W. 1986. Continuous recording of photochemical and non-photochemical chlorophyll fluorescence quenching with a new type of modulation fluorometer. Photosynthesis Research 10, 51–62.
Schreiber U, Bilger W. 1993. Progress in chlorophyll fluorescence research: major developments during the past years in retrospect. Progress in Botany 54, 151–153.
Schreiber U, Neubauer C. 1990. O2-dependent electron flow, membrane energization and the mechanism of nonphotochemical quenching. Photosynthesis Research 25, 279–293.[CrossRef]
Shuvalov VA, Heber U. 2003. Photochemical reactions in dehydrated photosynthetic organisms, leaves, chloroplasts, and photosystem II particles: Reversible reduction of pheophytin and chlorophyll and oxidation of ß-carotene. Chemical Physics 294, 227–237.[CrossRef]
Siefermann-Harms D. 1987. The light harvesting and protective functions of carotenoids in photosynthetic membranes. Physiologia Plantarum 69, 561–568.
Trebst A. 1986. The topology of the plastoquinone and herbicide binding peptides of photosystem II in the thylakoid membrane. Zeitschrift für Naturforschung 41c, 240–245.
Tracewell CA, Vrettos JS, Bautista JA, Frank HA, Brudwig GW. 2001. Carotenoid photo-oxidation in photosystem II. Archives of Biochemistry and Biophysics 385, 61–69.[CrossRef][Web of Science][Medline]
Vermaas W. 1993. Molecular-biological approaches to analyze photosystem II structures and functions. Annual Review of Plant Physiology and Plant Molecular Biology 44, 457–481.[CrossRef][Web of Science]
Yamamoto HY, Nakayama TOM, Chichester CO. 1962. Studies on the light and dark interconversions of leaf xanthophylls. Archives of Biochemistry 97, 168–173.[CrossRef][Web of Science][Medline]
Yamamoto HY, Kamite L. 1972. The effects of dithiothreitol on violaxanthin de-epoxidation and absorbance changes in the 500 nm region. Biochimica et Biophysica Acta 267, 538–543.[Medline]
Zinth W, Kaiser W. 1993. Time-resolved spectroscopy of the primary electron transfer in reaction centers of Rhodopacter sphaeroides and Rhodopseudomonas viridis. In: Deisenhofer J, Norris JR, eds. The photosynthetic reaction center. San Diego: Academic Press, 71–88.
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