Abstract
Plants in natural environments must cope with diverse, highly dynamic, and unpredictable conditions. They have mechanisms to enhance the capture of light energy when light intensity is low, but they can also slow down photosynthetic electron transport to prevent the production of reactive oxygen species and consequent damage to the photosynthetic machinery under excess light. Plants need a highly responsive regulatory system to balance the photosynthetic light reactions with downstream metabolism. Various mechanisms of regulation of photosynthetic electron transport under stress have been proposed, however the data have been obtained mainly under environmentally stable and controlled conditions. Thus, our understanding of dynamic modulation of photosynthesis under dramatically fluctuating natural environments remains limited. In this review, first I describe the magnitude of environmental fluctuations under natural conditions. Next, I examine the effects of fluctuations in light intensity, CO2 concentration, leaf temperature, and relative humidity on dynamic photosynthesis. Finally, I summarize photoprotective strategies that allow plants to maintain the photosynthesis under stressful fluctuating environments. The present work clearly showed that fluctuation in various environmental factors resulted in reductions in photosynthetic rate in a stepwise manner at every environmental fluctuation, leading to the conclusion that fluctuating environments would have a large impact on photosynthesis.
Similar content being viewed by others
Avoid common mistakes on your manuscript.
Introduction
Researches on regulations of steady-state photosynthesis in response to variations in light intensity (Evans et al. 1993; Ögren and Evans 1993; Yamori et al. 2010a), CO2 concentration (Farquhar et al. 1980; Yamori et al. 2005; Yamori and von Caemmerer 2009), temperature (Yamori et al. 2005, 2006, 2010b), and humidity (Bunce 1997; Lu et al. 2015; Rawson et al. 1977) have been extensively examined under controlled laboratory conditions. Predicting the environmental responses of the steady-state photosynthetic rate is central to many models of changes in the future global carbon cycle and terrestrial biosphere (Bernacchi et al. 2013; Groenendijk et al. 2011; Zhu et al. 2004). However, plants in natural environments must cope with highly dynamic and unpredictable conditions during the day. Models of steady-state photosynthesis tend to overestimate photosynthesis under fluctuating light (Naumburg and Ellsworth 2002; Timm et al. 2004). To improve models of dynamic photosynthesis under fluctuating environmental conditions, a better understanding of the responses of photosynthesis to fluctuating environments is needed.
Light intensity is the most variable factor in natural environments; during the day, it changes on the order of seconds, minutes, or hours because of changes in leaf angle, cloud cover, and overshadowing canopy. Leaves could receive 50–300 sunflecks per day (some shorter than 10 s) in the understory (Pearcy 1988; Pfitsch and Pearcy 1989). Sunflecks longer than 120 s represent only 5 % of the total number but contribute more than 75 % of total daily light, and therefore such sunflecks in forest understories would be more important for photosynthesis than the short ones. Previous researches on photosynthetic responses to fluctuating light have focused on their mechanisms and interspecific variations (Pearcy 1990; Pearcy and Way 2012). Photosynthetic responses to sunflecks differ among and within species depending on sunfleck duration, frequency, and intensity (Chazdon and Pearcy 1986; Leakey et al. 2004; Sims and Pearcy 1993; Watling et al. 1997; Yin and Johnson 2000). A sudden increase in light intensity typically leads to a hyperbolic increase in the leaf photosynthetic rate (e.g., Bai et al. 2008; Han et al. 1999; Pearcy 1990; Valladares et al. 1997). Because of biochemical and stomatal limitations, there is a time lag from onset of light to achievement of the maximum rate of photosynthesis (Allen and Pearcy 2000; Bai et al. 2008; Chen et al. 2011; Han et al. 1999; Pearcy 1990; Rijkers et al. 2000; Yamori et al. 2012, 2016a).
In contrast, the effects of fluctuations in temperature, CO2 concentration, relative humidity, and leaf-to-air vapor pressure deficit (VPD, which combines temperature and relative humidity into a single value) on dynamic photosynthesis have not been examined (Kaiser et al. 2015; Way and Pearcy 2012). The leaves of many plants are thin and have only minimal heat capacity; when exposed to strong sunlight, they can warm up substantially above air temperature, especially when stomata are closed. Leaves with low transpiration rates (e.g., oak leaves) suffer frequent high-temperature episodes, during which leaf temperature can increase above air temperature by as much as 10–15 °C within 1 min (Hanson et al. 1999; Singsaas et al. 1999; Singsaas and Sharkey 1998). In cotton leaves, which have high transpirational cooling, the temperature repeatedly exceeded 40 °C and fluctuated by as much as 8–10 °C in a matter of seconds (Wise et al. 2004). Since leaf temperature and thus VPD often change in parallel with light intensity (Peak and Mott 2011; Schymanski et al. 2013), fluctuation of leaf temperature and VPD as well as light intensity would affect the photosynthetic rate.
CO2 concentration at the leaf surface also fluctuates, since active photosynthesis could significantly reduce the CO2 concentration in the air at the leaf surface. Subsequent air flow at the leaf surface could transiently restore it to near ambient. Thus, the frequency of fluctuations in CO2 at the leaf surface would be highly variable and determined by photosynthetic rate and wind conditions.
Plant growth and yield depend largely on photosynthesis (Long et al. 2006; Yamori et al. 2016b; Zhu et al. 2010). Understanding not only steady-state photosynthetic characteristics but also non-steady-state photosynthetic characteristics under fluctuating environments is needed for improving dynamic photosynthesis models and for using biotechnological strategies to improve photosynthetic performance under natural conditions (Yamori 2013). In this review article, after introducing the magnitude of environmental fluctuations under natural conditions, the effects of fluctuations in light intensity, CO2 concentration, leaf temperature, and relative humidity on dynamic photosynthesis has been analyzed in rice. Finally, photoprotective strategies that allow plants to maintain the photosynthesis under stressful fluctuating environments have been summarized. The present work clearly showed that various fluctuating environmental factors led to reductions in photosynthetic rate in a stepwise manner at every environmental fluctuation, resulting in photoinhibition. Thus, it is concluded that fluctuating environments have a large impact on photosynthetic performance and are stressful for plants.
Light-dependent reactions of photosynthesis
Light is absorbed by light-harvesting systems, which contain chlorophylls and carotenoids. The energy captured by the photosynthetic pigments is transferred to the reaction centers of photosystem I (PSI) and photosystem II (PSII) in the thylakoid membranes of chloroplasts (Fig. 1). Electrons derived from water splitting in PSII are ultimately transferred to NADP+ via PSI, resulting in NADPH production. This process is known as linear electron transport (Fig. 1). This linear electron transport, in which electrons pass through the cytochrome (Cyt) b6/f complex, generates a proton (H+) gradient across the thylakoid membrane (ΔpH) (Allen 2003). Together with protons generated by the water-splitting complex, ΔpH induces the qE component of non-photochemical quenching (NPQ) of excitation energy. The ΔpH and transmembrane electrical potential (Δψ) enable ATP production by ATP synthase (Fig. 1). In contrast to linear electron transport, cyclic electron transport around PSI depends solely on PSI; the electrons also pass through the Cyt b 6/f complex (Fig. 1). Cyclic electron transport around PSI can generate ΔpH and drives ATP synthesis without NADPH accumulation (Yamori and Shikanai 2016). Generated ATP and NADPH fuel the Calvin–Benson cycle and other metabolic pathways in the chloroplast stroma. Because of a concentration gradient, CO2 diffuses from the air through the stomata and then through intercellular airspaces into cells and ultimately chloroplasts, where it is fixed into carbohydrates (Fig. 1).
Photosynthesis is sensitive to various environmental changes, because it needs to balance absorbed light energy with the energy consumed by various metabolic pathways. The rate of photosynthesis rises with the increase in light intensity until saturation (Fig. 2). Low light limits photosynthesis and thus plant growth, whereas excessive light can cause photoinhibition of PSII, which results in reductions in photosynthetic rate (Aro et al. 1993; Takahashi and Badger 2011). PSII is very sensitive to light stress and is rapidly inactivated under strong light, a phenomenon that is referred to as PSII photoinhibition (Aro et al. 1993; Powles 1984). The light-induced photodamage to PSII and the repair of photodamaged PSII occur simultaneously. PSII photoinhibition becomes apparent when the photodamage rate exceeds the repair rate. Photosynthetic rate is also limited by other factors such as low or high temperature, low humidity, and low CO2 concentration, where light intensity would be frequently in excess of that required for CO2 assimilation, which can lead to photoinhibition (Fig. 2). Thus, photoinhibition is common even at light intensities that would be otherwise optimal (Murchie et al. 1999).
Dynamic photosynthetic responses to fluctuating environments
The rate of photosynthesis is determined by not only the rate of steady-state photosynthesis but also the speed of photosynthetic responses to fluctuating environments (Way and Pearcy 2012). In the present study, chlorophyll fluorescence, P700 redox state, and gas exchange were simultaneously measured in rice (Oryza sativa ‘Hitomebore’) plants exposed to fluctuating environments (for Materials and methods, see Fig. 3). Under constant conditions, the rates of CO2 assimilation and electron transport around PSI (ETR I) and PSII (ETR II) were constant for 5 h, and the reduction level of the plastoquinone pool (1 − qL) and NPQ (an indicator of thermal dissipation of excess energy) were constant. Under fluctuating light, ETR I, ETR II, and CO2 assimilation rate decreased and 1 − qL increased in a stepwise manner at every transition to low light, whereas NPQ was slightly suppressed during both the low-light and high-light phases (Fig. 3). After 5 h of fluctuating light, ETR I, ETR II, and CO2 assimilation rate during both low- and high-light phases were decreased. Under fluctuating temperature (moderate–low or moderate–high), relative humidity (moderate–low), and CO2 concentration (ambient–low or ambient–high), all the photosynthetic parameters (i.e., ETR I, ETR II, and CO2 assimilation rate) decreased in a stepwise manner at every transition, whereas 1 − qL and NPQ gradually increased over 5 h (Fig. 3).
The maximum level of the P700 signal (P m; full oxidation of P700) in the dark and the maximum quantum yield of PSII (F v/F m) were measured before and after exposure to constant or fluctuating conditions for 5 h (Fig. 4). Fluctuating CO2 concentration, leaf temperature, and relative humidity significantly reduced F v/F m in comparison with constant conditions, indicating that PSII is susceptible to fluctuating environments (Figs. 4, 5). On the other hand, only fluctuating light significantly reduced P m, indicating that PSI is more susceptible to fluctuating light than to constant high light or other fluctuating conditions (Figs. 4, 5). In fluctuating environments, the electron transport system accumulates excess reducing power, which cannot be dissipated as heat and may cause a strong reducing burst (Fig. 3), eventually leading to photoinhibition of PSI or PSII (Fig. 4). As a result, plants decreased their CO2 assimilation rate via a reduction in ETR I and ETR II (Fig. 3). Thus, these experiments showed that fluctuating conditions are stressful for rice plants (Fig. 5).
Various components of photosynthesis respond to environmental fluctuations with very different time constants. For example, when light intensity is increased suddenly after a prolonged period of darkness or low light, photosynthetic rate increases gradually over the course of several minutes to tens of minutes and approaches a new steady-state level (e.g., Yamori et al. 2012). In this phase of photosynthetic induction, the initial reactions involved in electron transport respond to changes in light intensity immediately (for a review, see Pearcy 1990). During the first 1–2 min of light induction, the photosynthetic rate is limited by the build-up of metabolic pools, especially by RuBP regeneration (Sassenrath-Cole and Pearcy 1992; Way and Pearcy 2012). Light fluctuation within a few seconds would not have a drastic effect on the Calvin–Benson cycle, because the activation of enzymes under light and their inactivation in the dark take several minutes to tens of minutes (Buchanan 1980, 1991). Because light controls the activity of a number of Calvin–Benson cycle enzymes and ATP synthase via the ferredoxin/thioredoxin system, the changes in light intensity within minutes may be critical for CO2 assimilation in the Calvin–Benson cycle [which includes ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco), chloroplast fructose-1, 6-bisphosphatase (FBPase), sedoheptulose-1,7-bisphosphatase (SBPase), phosphoribulokinase (PRK)] (Kirschbaum and Pearcy 1988; Sassenrath-Cole and Pearcy 1992, 1994). Light fluctuation for several tens of minutes may affect stomatal conductance (Cardon and Berry 1992; Lawson et al. 2002; Morison 1998). Because these reactions affect photosynthesis as a whole, drastic fluctuations in light intensity can cause photoinhibition and reduce plant growth (Kono et al. 2014; Tikkanen and Aro 2012; Yamori et al. 2016a).
Temperature changes have a strong impact on photosynthetic reactions. The temperature response curve of photosynthetic rate is generally parabolic, often with an optimum at the growth temperature (for reviews, see Way and Yamori 2014; Yamori et al. 2014). A decrease in temperature by 10 °C (e.g., from 20 to 10 °C) will reduce the activity of Calvin–Benson cycle enzymes by 50 %. This usually reduces the demand for the reducing equivalent (NADPH) and causes subsequent accumulation of reductants. High temperatures reduce the activity of the Calvin–Benson cycle enzymes, especially Rubisco. The activation state of Rubisco is regulated by Rubisco activase (Portis 2003), which fails to maintain a high Rubisco activation state at high temperatures because of its thermolability (Crafts-Brandner and Salvucci 2000; Salvucci and Crafts-Brandner 2004). Thus, fluctuating temperature (moderate–low or moderate–high) in combination with high light could lead to over-reduction of the photosynthetic electron transport chain and eventually to photoinhibition.
Drought stress induces stomatal closure, which limits CO2 transfer into leaves and limits photosynthesis (Chaves et al. 2009; Cornic 2000; Flexas and Medrano 2002). This limitation can decrease the activity of the Calvin–Benson cycle enzymes and result in over-reduction of the photosynthetic electron transport chain, similar to the effect of temperature. High CO2 concentrations reduce stomatal conductance (Ainsworth et al. 2002; Ainsworth and Rogers 2007; Medlyn et al. 2001). In Phaseolus vulgaris, fluctuations in CO2 concentration decreased CO2 assimilation rate via a reduction in stomatal conductance (Cardon et al. 1994, 1995). Although changes in CO2 concentration, temperature, and humidity may be more gradual than changes in light intensity, fluctuating conditions causing stomatal closure under high light can lead to photoinhibition.
Photoinhibition of photosystems I and II
Photoinhibition may represent downregulation of the photosynthetic apparatus when harvested light energy exceeds energy that can be used by the chloroplast. In our experiments, fluctuating conditions other than light damaged PSII (Fig. 4). Many studies also have reported the sensitivity of PSII to photodamage, and PSII has an efficient and dynamic repair machinery (for reviews, see Aro et al. 1993, 2005). The damaged PSII proteins, primarily D1, are replaced with newly synthesized proteins after partial PSII disassembly (Takahashi and Badger 2011). ROS generated under various stresses inhibit PSII repair (Takahashi and Murata 2008). The dynamic control of active PSII reaction centers via a photoinhibition–repair cycle may rescue PSI from photodamage under high light (Tikkanen et al. 2014).
In contrast to PSII, PSI lacks efficient repair, and thus its recovery is extremely slow (Kudoh and Sonoike 2002). PSII photodamage increases linearly with light intensity, whereas PSI damage appears only when the electron flow from PSII exceeds the capacity of PSI electron acceptors. PSI has generally been regarded as more stable than PSII in vivo (Sonoike 2011). However, under natural conditions, PSI photodamage could be much more frequent than anticipated. PSI is potentially susceptible to fluctuating light even in wild-type Arabidopsis (Kono et al. 2014; Suorsa et al. 2012; Tikkanen et al. 2010) and rice (Figs. 4, 5; Yamori et al. 2016a). Selective photoinhibition of PSI by moderate light treatment of intact leaves at chilling temperatures occurs in several plant species, including cotton (Kornyeyev et al. 2003a, b), cucumber (Sonoike and Terashima, 1994; Terashima et al. 1994), and potato (Havaux and Davaud 1994). Thus, PSI is the primary site of photoinhibition in many plant species under certain conditions (e.g., fluctuating light or chilling temperature with moderate light intensity), and PSI photodamage could limit crop growth in temperate climates.
Photoprotective strategies to prevent photoinhibition
Rapid photosynthetic responses to fluctuating environments which operate on the proper timescale should determine their viability and survival under such stressful environment conditions. A number of photoprotective mechanisms over a wide range of time scales have been reported (Fig. 6); (1) photoprotection by avoiding exposure to light and (2) photoprotection by coping with excess light absorbed by photosynthetic pigments (Baena-Gonzalez and Aro 2002; Holt et al. 2004; Takahashi and Badger 2011). Below, I review various photoprotective strategies, with emphasis on photoprotection by coping with excess absorbed light.
Photoprotection by avoiding exposure to light at the leaf or chloroplast levels
Leaf movement, such as paraheliotropism, is an efficient strategy to reduce light interception and thus to avoid excess light energy absorption (Arena et al. 2008; Jiang et al. 2006; Ludlow and Bjorkman 1984; Pastenes et al. 2005), and also to reduce heat and transpirational water loss which confers the plant to protect against photoinhibition (Forseth and Ehleringer 1982; Gamon and Pearcy 1989). Paraheliotropism protects against photoinhibition in bean (Bielenberg et al. 2003; Pastenes et al. 2005), soybean (Jiang et al. 2006), and wild grape (Gamon and Pearcy 1989, 1990). Another type of leaf movement is leaf rolling, which protects leaves from dehydration because it reduces the effective leaf area and therefore transpiration. At the same time, leaf rolling protects leaves from photodamage (Corlett et al. 1994).
Chloroplasts alter their positions to optimize the use of light (Suetsugu and Wada 2007; Wada et al. 2003). Under strong light, they move from the cell surface to the side walls of cells. This so-called “avoidance response” protects against photoinhibition (Kasahara et al. 2002) because chloroplast movements decrease light interception by the photosynthetic apparatus and may reduce light absorptance by approximately 20 % (Brugnoli and Bjorkman 1992). Chloroplasts move at velocities above 1 µm min−1 (Kagawa and Wada 2004), and thus their avoidance response would be completed in approximately 20 min. Two photoreceptors for chloroplast movement, phototropin 1 (PHOT1) and phototropin 2 (PHOT2), have been identified in Arabidopsis thaliana. Only PHOT2 participates in the avoidance response, but both play a role in chloroplast accumulation in illuminated cell areas under low light to maximize light absorption for photosynthesis (Wada et al. 2003).
Photoprotection by coping with excess light absorbed by photosynthetic pigments
Several photoprotective processes involving in the photosynthetic electron transport have been proposed (Fig. 7).
Thermal energy dissipation of absorbed excess light energy
Plants possess a rapidly (on the time scale of seconds) inducible NPQ mechanism, termed ΔpH-dependent quenching (qE), for harmless thermal dissipation of excess light energy absorbed in the light-harvesting antenna of PSII (Fig. 7a). The mechanism responsible for qE component is associated with the conversion of violaxanthin via antheraxanthin to zeaxanthin by violaxanthin de-epoxidase and the protonation of the PSII subunit PsbS, which functions as a sensor of lumen pH in plants (Li et al. 2000; Niyogi et al. 2005). Both reactions are enhanced by low lumen pH, which is accompanied by ΔpH generation through electron transport at high light (Niyogi 1999). Thus, qE component of NPQ can be considered as feedback regulation of the light-dependent reactions of photosynthesis.
Thermal dissipation might protect PSII from photoinhibition by decreasing the rate of PSII photodamage under strong light (Havaux and Niyogi 1999; Li et al. 2002). However, a recent study on Arabidopsis mutants defective in NPQ revealed that defects in thermal dissipation inhibited PSII repair without notable effects on PSII photodamage; this inhibition was attributed to suppression of de novo protein synthesis, especially that of D1 (Takahashi et al. 2009). ROS accelerated photoinhibition by inhibiting PSII repair rather than by being directly involved in photodamage (Nishiyama et al. 2006). Thus, it appears that thermal dissipation plays a role in preventing ROS generation and avoiding ROS-mediated inhibition of de novo D1 synthesis. There is still a possibility that the ROS could damage PSII depending on the light conditions, types of ROS (e.g., O2 −, H2O2 and OH·) and the ROS concentrations (Blot et al. 2011; Fan et al. 2016), since it has been also reported that ROS can photodamage PSII directly (Fan et al. 2016; Hideg et al. 2011; Kornyeyev et al. 2010).
Cyclic electron transport around PSI
Cyclic electron transport around PSI enhances generation of ΔpH across the thylakoid membrane by increasing electron transfer from PSI to the plastoquinone pool, and then back to PSI through the proton-pumping Cyt b 6/f complex (Fig. 7b). Cyclic electron transport around PSI is proposed to be essential for balancing the ATP/NADPH production ratio by increasing ∆pH to increase ATP synthesis (Yamori and Shikanai 2016). In addition, the additional generation of ΔpH induces qE to dissipate excess absorbed light energy (Müller et al. 2001; Niyogi 1999). The electron flow through the Cyt b 6/f complex slows as ∆pH increases. Thus, ∆pH plays a regulatory roles via the acidification of the thylakoid lumen by the down-regulation of electron transport through the Cyt b 6/f complex (Golding et al. 2004; Kramer et al. 2004; Suorsa et al. 2012; Tikkanen et al. 2014). Therefore, cyclic electron transport around PSI protects both photosystems from damage caused by stromal over-reduction (Miyake 2010; Takahashi and Badger 2011; Yamori and Shikanai 2016). It is worth noting that cyclic electron transport around PSII, which requires only PSII photochemical reactions, has been proposed to operate efficiently under excess light conditions (Miyake and Okamura 2003; Prasil et al. 1996), although its physiological roles and molecular mechanisms are still not clear.
In angiosperms, two alternative pathways of cyclic electron transport around PSI have been identified (for a review, see Yamori and Shikanai 2016). The major pathway in Arabidopsis depends on two proteins, PROTON GRADIENT REGULATION 5 (PGR5) (Munekage et al. 2002, 2004) and PGR5-Like PHOTOSYNTHETIC PHENOTYPE 1 (PGRL1) (DalCorso et al. 2008; Hertle et al. 2013), whereas the minor pathway is mediated by the chloroplast NADH dehydrogenase-like (NDH) complex (Burrows et al. 1998; Horváth et al. 2000; Kofer et al. 1998; Shikanai et al. 1998), which forms a supercomplex with PSI (Peng et al. 2011). Both the PGR5/PGRL1-dependent pathway (DalCorso et al. 2008; Hertle et al. 2013; Munekage et al. 2002, 2004) and the NDH-dependent pathway (Yamamoto et al. 2011; Yamamoto and Shikanai 2013) are involved in ferredoxin-dependent cyclic electron transport around PSI. In C3 plants, approximately ≤10–15 % of total electron transport is derived from cyclic electron transport around PSI in the steady state (e.g., Fan et al. 2007; Kramer et al. 2004; Kuvykin et al. 2011; Laisk et al. 2005, 2007; Miyake et al. 2005). However, when demand for ATP is higher than that for NADPH (e.g., during photosynthetic induction, at high or low temperature, at low CO2 concentration, or under drought), cyclic electron transport around PSI is likely to be activated (for a recent review, see Yamori and Shikanai 2016). Complete inhibition of both cyclic electron transports around PSI in the Arabidopsis crr2 pgr5 double mutant severely impairs photosynthesis and growth (Munekage et al. 2004), indicating that cyclic electron transport around PSI is essential for photosynthesis in C3 species.
Plastoquinol terminal oxidase (PTOX)
Plastoquinol terminal oxidase (PTOX) serves as an alternative electron sink, which oxidizes plastoquinol (PQH2) and reduces O2–H2O when the plastoquinone pool is over-reduced (McDonald et al. 2011; Nawrocki et al. 2015; Peltier and Cournac 2002). In principle, PTOX could contribute to ATP production, and it plays a role in the control of the redox poise in chloroplasts (Fig. 7c). It has been proposed that PTOX is capable of modulating the balance between linear electron transport and cyclic electron transport around PSI during the dark-to-light transition (Trouillard et al. 2012). The alpine plant species Ranunculus glacialis has high levels of PTOX, which may act in effective alternative electron transport (Streb et al. 2005). However, because PTOX is expressed at very low levels in most C3 plants (e.g., Arabidopsis or tomato), it is likely to act as a safety valve to keep the plastoquinone pool oxidized and to prevent photo-oxidative damage under stress conditions (Josse et al. 2000; McDonald et al. 2011).
It is likely that PTOX is involved in chloroplast development (McDonald et al. 2011). During carotenoid synthesis, desaturation requires plastoquinone and is driven by a redox chain in which PTOX is likely to re-oxidize the reduced plastoquinone (Carol and Kuntz 2001; Kuntz 2004). The lack of PTOX causes the variegated leaf phenotype of the Arabidopsis immutans mutant and the tomato ghost mutant, in which the bleached spots originate at early stage during chloroplast biogenesis as a result of shortage in carotenoid synthesis (Carol and Kuntz 2001). Therefore, PTOX appears to play a role in carotenoid synthesis. It is likely that the relative importance of PTOX to electron transport could vary depending on the developmental and physiological context.
ROS scavenging through the water–water cycle
The water–water cycle (Asada 1999, 2006), which includes the Mehler reaction of O2 reduction by PSI (Mehler 1951), occurs by the photoreduction of O2–H2O at the reducing side of PSI via electrons generated from H2O in PSII (H2O → PSII → PSI → O2 → H2O) (Fig. 7d). The superoxide (O2 −) formed in this reaction is scavenged by superoxide dismutase (SOD) and ascorbate peroxidase (APX) with consumption of NADPH (Asada 1999, 2006). The consumption of NADPH by the water–water cycle allows linear electron transport to continue, thereby producing ATP. Thus, the water–water cycle is eventually coupled to the generation of ΔpH, which drives ATP synthesis without NADPH accumulation. It should be noted that ROS production under excess light is accelerated not only at PSI but also at PSII, although each photosystem produces different ROS types; superoxide (O2 −) and hydrogen peroxide (H2O2) in PSI and singlet-state oxygen (1O2) in PSII (for a review, see Asada 2006). To avoid such oxidative stress, chloroplasts detoxify ROS effectively using multiple enzymes, including SOD, APX and peroxiredoxin (Prx).
In the leaves of most C3 species, the water–water cycle contributes <5 % of linear electron transport even when CO2 assimilation is inhibited (Clarke and Johnson 2001; Ruuska et al. 2000), but in rice leaves this cycle appears to operate at a substantial level during photosynthetic induction (Makino et al. 2002). Thus, the water–water cycle likely plays a photoprotective role by ROS detoxification and dissipation of excess energy, and could also balance the levels of ATP and reductants (for reviews, see Asada 1999; Miyake 2010; Ort and Baker 2002).
It should be noted that, to minimize the effects of oxidative stress, plants have also evolved a non-enzymatic antioxidant system, such as low-molecular weight antioxidants of plant cells (e.g., glutathione, ascorbate, tocopherol and carotenoids) (for a review, see Apel and Hirt 2004; DellaPenna and Pogson 2006; Foyer et al. 2006). Mutants with decreased contents of ascorbic acid are hypersensitive to stress (Conklin et al. 1996). Moreover, overexpression of β-carotene hydroxylase in Arabidopsis leads to increased amounts of xanthophyll in chloroplasts, resulting in enhanced tolerance to oxidative stress (Davison et al. 2002). Studies using various double knockout mutants in Arabidopsis showed compensatory effects of ROS scavengers, including tocopherol, ascorbate and glutathione (Kanwischer et al. 2005). The enhancement of chloroplast antioxidant defenses has proved to be one of the most effective ways of protecting plant cells from abiotic stress (Chang et al. 2009; Ishikawa and Shigeoka 2008).
Metabolic interactions between chloroplasts and mitochondria: the malate–oxaloacetate (OAA) shuttle
NADPH generated by photosynthetic electron transport under high light can be re-oxidized by the mitochondrial respiratory chain, in which NADPH is consumed by the reduction of OAA to malate and the malate is exported from chloroplasts to mitochondria (Fig. 7e). This pathway is called ‘malate–OAA shuttle’ (Noguchi and Yoshida 2008; Scheibe 2004).
Although metabolic interactions between chloroplasts and mitochondria have multiple physiological consequences, the contribution of the mitochondria as an electron sink for photosynthesis in vivo is still unclear (for a review, see Noctor et al. 2004). The malate–OAA shuttle appears to have low capacity to regulate the ATP/NADPH ratio (Scheibe et al. 2005). However, in transgenic potato with reduced levels of malate dehydrogenase (MDH), the malate–OAA shuttle can contribute up to 10 % of total electron flow from PSII (Laisk et al. 2007). Thus, mitochondrial respiration could neutralize excess photosynthetic reducing power and prevent oxidative damage of thylakoid membranes and other cellular components (Noguchi and Yoshida 2008; Raghavendra and Padmasree 2003).
The photorespiration pathway
Since any efficient sink for electrons produced by water splitting in PSII may lower the risk of photoinhibition, it is relevant to include reactions of CO2 assimilation itself and photorespiration (Fig. 7f). In the Calvin–Benson cycle, Rubisco can fix both CO2 in photosynthesis and O2 in photorespiration (Bauwe et al. 2010). Photosynthetic carbon fixation produces two 3-phosphoglycerate (PGA) molecules for every carbon fixed, whereas photorespiration produces one PGA and one 2-phosphoglycolate, which is recycled to PGA with the loss of CO2 and ammonia. Thus, the photorespiratory pathway can be considered to consist of the photorespiratory carbon and nitrogen cycles. Photosynthesis results in net fixation of CO2, whereas the photorespiratory pathway requires ATP and releases previously fixed CO2. Photorespiration rate increases with a decrease in the CO2/O2 ratio in the chloroplast, and also under drought and high temperature (Ogren 1984; Sage et al. 2012). At current atmospheric CO2 concentrations and 30 °C, the rate of photorespiratory CO2 release from the mitochondria is approximately 25 % of the rate of net CO2 assimilation (Sage et al. 2012).
Although photorespiration is generally seen as a wasteful pathway, photorespiration is thought to act as a mechanism by which excess light energy can be used, reducing photodamage in the chloroplast (André 2011). The impairment of the photorespiratory pathway diminishes CO2 assimilation (because of the shortage of metabolites in the Calvin–Benson cycle and accumulation of intermediates of the photorespiratory pathway that can inhibit the Calvin–Benson cycle) and accelerates photoinhibition (Kozaki and Takeba 1996; Osmond 1981). In Arabidopsis mutants with impairments of ferredoxin-dependent glutamate synthase, serine hydroxymethyltransferase, glutamate/malate transporter, and glycerate kinase, photoinhibition caused by the impairment of the photorespiratory pathway is due to inhibition of the repair of photodamaged PSII, not acceleration of PSII photodamage (Takahashi et al. 2007). Therefore, the photorespiratory pathway could play a role in maintaining PSII repair by maintaining energy utilization in the Calvin–Benson cycle, which is important for decreasing ROS generation under stress.
Recent findings on the regulatory mechanisms of photosynthesis under fluctuating light
Analysis in several Arabidopsis mutants indicates that PGR5/PGRL1-dependent cyclic electron transport around PSI is crucial for photosynthesis regulation under fluctuating light (for a review, see Yamori and Shikanai 2016), especially at early developmental stages (Suorsa et al. 2012). In wild type, rapid induction of NPQ upon increase in light intensity prevents over-reduction of the plastoquinone pool. The pgr5 mutant and the NPQ mutants npq1 and npq4 induce little NPQ; fluctuating light hardly gives damage to the npq1 and npq4 mutants (Tikkanen et al. 2010), but it is lethal for the pgr5 mutant (Kono et al. 2014; Suorsa et al. 2012; Tikkanen et al. 2010). Unlike the npq4 mutant (Grieco et al. 2012; Tikkanen et al. 2015), the pgr5 mutant cannot oxidize P700 under high light, and this defect leads to PSI photodamage (Munekage et al. 2002). Thus, the absence of NPQ seems only to play an indirect role in response to fluctuating light. Similar to Arabidopsis pgr5 mutants, PGR5-knockdown rice (Yamori et al. 2016a) suffers from fluctuating light, with PSI as the primary target of photodamage and a stunted phenotype. The cumulative strong reduction of the entire electron transport system under fluctuating light for a couple of hours would cause a strong reducing burst at the acceptor side of PSI, leading to photodamage at PSI (Kono et al. 2014; Yamori et al. 2016a). Therefore, in higher plants the PGR5/PGRL1-dependent pathway is essential for effective responses to considerable fluctuations of light intensity and for avoiding photodamage (for a review see, Yamori and Shikanai 2016).
In rice, the impairment of NDH-dependent cyclic electron transport around PSI reduces photosynthetic rate under fluctuating light, leading to PSI photoinhibition, and consequently reduces biomass (Yamori et al. 2016a). During low-light phases, the NDH-dependent pathway maintains the electron transport chain on the acceptor side of PSI oxidized, which seems to be essential to prevent PSI over-reduction during subsequent high-light phases. This finding is supported by the important role of NDH-dependent PSI cyclic electron transport in regulation of the chloroplast redox state under constant low light, when light reactions limit photosynthesis (Yamori et al. 2015). In Arabidopsis, even the complete absence of the NDH complex does not reduce growth and photosynthesis under fluctuating light (Kono et al. 2014; Suorsa et al. 2012), indicating an interspecific difference in the physiological role of NDH-dependent cyclic electron transport around PSI under fluctuating light. This difference may depend on the activity of other alternative electron transport pathways, including the water–water cycle (Yamori and Shikanai 2016).
During acclimation to growth light environments, many plants change biochemical composition and morphology (e.g., Smith 1982; Terashima et al. 2005). Acclimation to low light enhances nitrogen allocation to components involved in light acquisition (e.g., chlorophylls and light-harvesting complexes), whereas acclimation to high light enhances components involved in light energy utilization (e.g., Rubisco and other Calvin–Benson cycle enzymes) and photoprotective potential. Photosynthetic acclimation to fluctuating light would require specific mechanisms, because plants acclimated to low light cannot cope with high light, and plants acclimated to high light cannot use low light efficiently. In rice, both PGR5/PGRL1-dependent and NDH-dependent cyclic electron transport around PSI sustain photosynthesis and plant growth under fluctuating light (Yamori et al. 2016a). The highly responsive regulatory system controlled by cyclic electron transport around PSI could optimize photosynthesis and plant growth under naturally fluctuating light (Yamori and Shikanai 2016).
Future perspectives
Despite extensive research on improving photosynthesis to increase crop yields (Yamori 2013; Yamori et al. 2016b), the photosynthetic responses to naturally fluctuating environments remains unclear. To optimize energy gain at low light and to protect the photosynthetic machinery at high light, plants have various photosynthetic regulation mechanisms that are either constitutively active or are activated when needed (Fig. 7; Demmig-Adams and Adams 1992; Niyogi 2000). It has not been clarified what would be the strategy to cope with fluctuations in each environmental factor (e.g., humidity, temperature, CO2). Thus, it would be needed to examine a role of each photoprotective strategy in avoidance of photoinhibition under various fluctuating environmental factors. Photoprotection is an essential adaptation to prevent severe photoinhibition and reduction of photosynthesis and thus plant growth. Therefore, understanding the physiological and molecular basis of photoprotection under fluctuating environments would help to develop selection markers for breeding aimed at enhancing stress tolerance in crops and ensuring food security. Recent advances in nuclear or chloroplast genome transformation facilitate manipulation of photosynthesis. Analysis of mutants or transformants could considerably improve our understanding of photosynthetic regulation under fluctuating conditions. Additionally, understanding of the responses of photosynthesis to fluctuating environments could lead to improvements of model predictions of dynamic photosynthesis under fluctuating environments in nature.
References
Ainsworth EA, Rogers A (2007) The response of photosynthesis and stomatal conductance to rising [CO2]: mechanisms and environmental interactions. Plant Cell Environ 30:258–270. doi:10.1111/j.1365-3040.2007.01641.x
Ainsworth EA, Davey PA, Bernacchi CJ et al (2002) A meta-analysis of elevated [CO2] effects on soybean (Glycine max) physiology, growth and yield. Glob Chang Biol 8:695–709. doi:10.1046/j.1365-2486.2002.00498.x
Allen JF (2003) Cyclic, pseudocyclic and noncyclic photophosphorylation: new links in the chain. Trends Plant Sci 8:15–19. doi:10.1016/S1360-1385(02)00006-7
Allen MT, Pearcy RW (2000) Stomatal versus biochemical limitations to dynamic photosynthetic performance in four tropical rainforest shrub species. Oecologia 122:479–486. doi:10.1007/s004420050969
André MJ (2011) Modelling 18O2 and 16O2 unidirectional fluxes in plants: II. Analysis of Rubisco evolution. Biosystems 103:252–264. doi:10.1016/j.biosystems.2010.10.003
Apel K, Hirt H (2004) Reactive oxygen species: metabolism, oxidative stress, and signal transduction. Ann Rev Plant Biol 55:373–399. doi:10.1146/annurev.arplant.55.031903.141701
Arena C, Vitale L, De Santo AV (2008) Paraheliotropism in Robinia pseudoacacia L.: an efficient strategy to optimise photosynthetic performance under natural environmental conditions. Plant Biol 10:194–201. doi:10.1111/j.1438-8677.2008.00032.x
Aro EM, Virgin I, Andersson B (1993) Photoinhibition of photosystem II. Inactivation, protein damage and turnover. Biochim Biophys Acta 1143:113–134. doi:10.1016/0005-2728(93)90134-2
Aro E-M, Suorsa M, Rokka A et al (2005) Dynamics of photosystem II: a proteomic approach to thylakoid protein complexes. J Exp Bot 56:347–356. doi:10.1093/jxb/eri041
Asada K (1999) The water-water cycle in chloroplasts: scavenging of active oxygens and dissipation of excess photons. Annu Rev Plant Physiol Plant Mol Biol 50:601–639. doi:10.1146/annurev.arplant.50.1.601
Asada K (2006) Production and scavenging of reactive oxygen species in chloroplasts and their functions. Plant Physiol 141:391–396. doi:10.1104/pp.106.082040
Baena-Gonzalez E, Aro EM (2002) Biogenesis, assembly and turnover of photosystem II units. Philos Trans R Soc Lond B Biol Sci 357:1451–1459. doi:10.1098/rstb.2002.1141
Bai KD, Liao DB, Jiang DB, Cao KF (2008) Photosynthetic induction in leaves of co-occurring Fagus lucida and Castanopsis lamontii saplings grown in contrasting light environments. Trees 22:449–462. doi:10.1007/s00468-007-0205-4
Bauwe H, Hagemann M, Fernie AR (2010) Photorespiration: players, partners and origin. Trends Plant Sci 15:330–336. doi:10.1016/j.tplants.2010.03.006
Bernacchi CJ, Bagley JE, Serbin SP et al (2013) Modelling C3 photosynthesis from the chloroplast to the ecosystem. Plant Cell Environ 36:1641–1657. doi:10.1111/pce.12118
Bielenberg DG, Miller JD, Berg VS (2003) Paraheliotropism in two Phaseolus species: combined effects of photon flux density and pulvinus temperature, and consequences for leaf gas exchange. Environ Exp Bot 49:95–105. doi:10.1016/S0098-8472(02)00062-X
Blot N, Mella-Flores D, Six C et al (2011) Light history influences the response of the marine cyanobacterium Synechococcus sp. WH7803 to oxidative stress. Plant Physiol 156:1934–1954. doi:10.1104/pp.111.174714
Brugnoli E, Bjorkman O (1992) Chloroplast movements in leaves: influence on chlorophyll fluorescence and measurements of light-induced absorbance changes related to pH and zeaxanthin formation. Photosynth Res 32:23–35. doi:10.1007/BF00028795
Buchanan BB (1980) Role of light in the regulation of chloroplast enzymes. Annu Rev Plant Physiol 31:341–374. doi:10.1146/annurev.pp.31.060180.002013
Buchanan BB (1991) Regulation of CO2 assimilation in oxygenic photosynthesis: the ferredoxin/thioredoxin system. Perspective on its discovery, present status, and future development. Arch Biochem Biophys 288:1–9. doi:10.1016/0003-9861(91)90157-E
Bunce JA (1997) Does transpiration control stomatal responses to water vapour pressure deficit? Plant Cell Environ 20:131–135
Burrows PA, Sazanov LA, Svab Z, Maliga P, Nixon PJ (1998) Identification of a functional respiratory complex in chloroplasts through analysis of tobacco mutants containing disrupted plastid ndh genes. EMBO J 17:868–876. doi:10.1093/emboj/17.4.868
Cardon ZG, Berry J (1992) Effects of O2 and CO2 concentration on the steady-state fluorescence yield of single guard cell pairs in intact leaf discs of Tradescantia albiflora. Plant Physiol 99:1238–1244. doi:10.1104/pp.99.3.1238
Cardon ZG, Berry JA, Woodward IE (1994) Dependence of the extent and direction of average stomatal responses in Zea mays L. and Phaseolus vulgaris L. on the frequency of fluctuations in environmental stimuli. Plant Physiol 105:1007–1013. doi:10.1104/pp.105.3.1007
Cardon ZG, Berry JA, Woodward IE (1995) Fluctuating [CO2] drives species specific changes in water use efficiency. J Biogeogr 22:203–208. doi:10.2307/2845911
Carol P, Kuntz M (2001) A plastid terminal oxidase comes to light: implications for carotenoid biosynthesis and chlororespiration. Trend Plant Sci 6:31–36. doi:10.1016/S1360-1385(00)01811-2
Chang CC, Slesak I, Jordá L et al (2009) Arabidopsis chloroplastic glutathione peroxidases play a role in cross talk between photooxidative stress and immune responses. Plant Physiol 150:670–683. doi:10.1104/pp.109.135566
Chaves MM, Flexas J, Pinheiro C (2009) Photosynthesis under drought and salt stress: regulation mechanisms from whole plant to cell. Ann Bot 103:551–560. doi:10.1093/aob/mcn125
Chazdon RL, Pearcy RW (1986) Photosynthetic responses to light variation in rainforest species. II. Carbon gain and photosynthetic efficiency during lightflecks. Oecologia 69:524–531. doi:10.1007/BF00410358
Chen JW, Zhang Q, Li XS, Cao KF (2011) Steady and dynamic photosynthetic responses of seedlings from contrasting successional groups under low-light growth conditions. Physiol Plant 141:84–95. doi:10.1111/j.1399-3054.2010.01414.x
Clarke JE, Johnson GN (2001) In vivo temperature dependence of cyclic and pseudocyclic electron transport in barley. Planta 212:808–816. doi:10.1007/s004250000432
Conklin PL, Williams EH, Last RL (1996) Environmental stress sensitivity of an ascorbic acid-deficient Arabidopsis mutant. Proc Natl Acad Sci USA 93:9970–9974
Corlett JE, Jones HG, Massacci A, Masojidek J (1994) Water deficit, leaf rolling and susceptibility to photoinhibition in field grown sorghum. Physiol Plant 92:423–430. doi:10.1111/j.1399-3054.1994.tb08831.x
Cornic G (2000) Drought stress inhibits photosynthesis by decreasing stomatal aperture—not by affecting ATP synthesis. Trends Plant Sci 5:187–188. doi:10.1016/S1360-1385(00)01625-3
Crafts-Brandner SJ, Salvucci ME (2000) Rubisco activase constrains the photosynthetic potential of leaves at high temperature and CO2. Proc Natl Acad Sci USA 97:13430–13435. doi:10.1073/pnas.230451497
DalCorso G, Pesaresi P, Masiero S et al (2008) A complex containing PGRL1 and PGR5 is involved in the switch between linear and cyclic electron flow in Arabidopsis. Cell 132:273–285. doi:10.1016/j.cell.2007.12.028
Davison PA, Hunter CN, Horton P (2002) Overexpression of β-carotene hydroxylase enhances stress tolerance in Arabidopsis. Nature 418:203–206. doi:10.1038/nature00861
DellaPenna D, Pogson BJ (2006) Vitamin synthesis in plants: tocopherols and carotenoids. Annu Rev Plant Biol 57:711–738. doi:10.1146/annurev.arplant.56.032604.144301
Demmig-Adams B, Adams WW III (1992) Photoprotection and other responses of plants to high light stress. Annu Rev Plant Physiol Plant Mol Biol 43:599–626. doi:10.1146/annurev.pp.43.060192.003123
Eberhard S, Finazzi G, Wollman FA (2008) The dynamics of photosynthesis. Annu Rev Genet 42:463–515. doi:10.1146/annurev.genet.42.110807.091452
Evans JR, Jakobsen I, Ogren E (1993) Photosynthetic light-response curves 2. Gradients of light absorption and photosynthetic capacity. Planta 189:191–200. doi:10.1007/BF00195076
Fan DY, Nie Q, Hope AB, Hillier W, Pogson BJ, Chow WS (2007) Quantification of cyclic electron flow around photosystem I in spinach leaves during photosynthetic induction. Photosynth Res 94:347–357. doi:10.1007/s11120-006-9127-z
Fan D, Ye Z, Wang S, Chow WS (2016) Multiple roles of oxygen in the photoinactivation and dynamic repair of photosystem II in spinach leaves. Photosynth Res 127:307–319. doi:10.1007/s11120-015-0185-y
Farquhar GD, Caemmerer S, Berry JA (1980) A biochemical model of photosynthetic CO2 assimilation in leaves of C3 species. Planta 149:78–90. doi:10.1007/BF00386231
Flexas J, Medrano H (2002) Drought-inhibition of photosynthesis in C3 plants: stomatal and non-stomatal limitations revisited. Ann Bot 89:183–189. doi:10.1093/aob/mcf027
Forseth IN, Ehleringer JR (1982) Ecophysiology of two solar-tracking desert winter annuals. II. Leaf movements, water relations and microclimate. Oecologia 54:41–49. doi:10.1007/BF00541105
Foyer CH, Trebst A, Noctor G (2006) Signaling and integration of defense functions of tocopherol, ascorbate and glutathione. In: Demmig-Adams B, Adams WW III, Mattoo AK (eds) Photoprotection, photoinhibition, gene regulation and environment. Springer, Dordrecht, pp 241–268. doi:10.1007/1-4020-3579-9_16
Gamon JA, Pearcy RW (1989) Leaf movement, stress avoidance and photosynthesis in Vitis californica. Oecologia 79:475–481. doi:10.1007/BF00378664
Gamon JA, Pearcy RW (1990) Photoinhibition in Vitis californica: interactive effects of sunlight, temperature and water status. Plant Cell Environ 13:267–275. doi:10.1111/j.1365-3040.1990.tb01311.x
Golding AJ, Finazzi G, Johnson GN (2004) Reduction of the thylakoid electron transport chain by stromal reductants: evidence for activation of cyclic electron transport upon dark adaptation or under drought. Planta 220:356–363. doi:10.1007/s00425-004-1345-z
Grieco M, Tikkanen M, Paakkarinen V, Kangasjarvi S, Aro EM (2012) Steady-state phosphorylation of light-harvesting complex II proteins preserves photosystem I under fluctuating white light. Plant Physiol 160:1896–1910. doi:10.1104/pp.112.206466
Groenendijk M, Dolman AJ, van der Molen MK et al (2011) Assessing parameter variability in a photosynthesis model within and between plant functional types using global Fluxnet eddy covariance data. Agric For Meteorol 151:22–38. doi:10.1016/j.agrformet.2010.08.013
Han Q, Yamaguchi E, Odaka N, Kakubari Y (1999) Photosynthetic induction responses to variable light under field conditions in three species grown in the gap and understory of a Fagus crenata forest. Tree Physiol 19:625–634. doi:10.1093/treephys/19.10.625
Hanson DT, Swanson S, Graham LE, Sharkey TD (1999) Evolutionary significance of isoprene emission from mosses. Am J Bot 86:634–639. doi:10.2307/2656571
Havaux M, Davaud A (1994) Photoinhibition of photosynthesis in chilled potato leaves is not correlated with a loss of photosystem-II activity. Photosynth Res 40:75–92. doi:10.1007/BF00019047
Havaux M, Niyogi KK (1999) The violaxanthin cycle protects plants from photooxidative damage by more than one mechanism. Proc Natl Acad Sci USA 96:8762–8767. doi:10.1073/pnas.96.15.8762
Hertle AP, Blunder T, Wunder T et al (2013) PGRL1 is the elusive ferredoxin-plastoquinone reductase in photosynthetic cyclic electron flow. Mol Cell 49:511–523. doi:10.1016/j.molcel.2012.11.030
Hideg É, Deák Z, Hakala-Yatkin M et al (2011) Pure forms of the singlet oxygen sensors TEMP and TEMPD do not inhibit photosystem II. Biochim Biophys Acta 1807:1658–1661. doi:10.1016/j.bbabio.2011.09.009
Holt NE, Fleming GR, Niyogi KK (2004) Toward an understanding of the mechanism of non-photochemical quenching in green plants. Biochemistry 43:8281–8289. doi:10.1021/bi0494020
Horváth EM, Peter SO, Joet T et al (2000) Targeted inactivation of the plastid ndhB gene in tobacco results in an enhanced sensitivity of photosynthesis to moderate stomatal closure. Plant Physiol 123:1337–1350. doi:10.1104/pp.123.4.1337
Ishikawa T, Shigeoka S (2008) Recent advances in ascorbate biosynthesis and the physiological significance of ascorbate peroxidase in photosynthesizing organisms. Biosci Biotechnol Biochem 72:1143–1154. doi:10.1271/bbb.80062
Jiang CD, Gao HY, Zou Q, Jiang GM, Li LH (2006) Leaf orientation, photorespiration and xanthophyll cycle protect young soybean leaves against high irradiance in the field. Environ Exp Bot 55:87–96. doi:10.1016/j.envexpbot.2004.10.003
Josse EM, Simkin AJ, Gaffé J, Labouré AM, Kuntz M, Carol P (2000) A plastid terminal oxidase associated with carotenoid desaturation during chromoplast differentiation. Plant Physiol 123:1427–1436. doi:10.1104/pp.123.4.1427
Kagawa T, Wada M (2004) Velocity of chloroplast avoidance movement is fluence rate dependent. Photochem Photobiol Sci 3:592–595. doi:10.1039/B316285K
Kaiser E, Morales A, Harbinson J, Kromdijk J, Heuvelink E, Marcelis LFM (2015) Dynamic photosynthesis in different environmental conditions. J Exp Bot 66:2415–2426. doi:10.1093/jxb/eru406
Kanwischer M, Porfirova S, Bergmüller E, Dörmann P (2005) Alterations in tocopherol cyclase activity in transgenic and mutant plants of Arabidopsis affect tocopherol content, tocopherol composition, and oxidative stress. Plant Physiol 137:713–723. doi:10.1104/pp.104.054908
Kasahara M, Kagawa T, Oikawa K et al (2002) Chloroplast avoidance movement reduces photodamage in plants. Nature 420:829–832. doi:10.1038/nature01213
Kirschbaum MUF, Pearcy RW (1988) Gas exchange analysis of the fast phase of photosynthetic induction in Alocasia macrorrhiza. Plant Physiol 87:818–821. doi:10.1104/pp.87.4.818
Kofer W, Koop HU, Wanner G, Steinmüller K (1998) Mutagenesis of the genes encoding subunits A, C, H, I, J and K of the plastid NAD(P)H-plastoquinone-oxidoreductase in tobacco by polyethylene glycol-mediated plastome transformation. Mol Gen Genet 258:166–173. doi:10.1007/s004380050719
Kono M, Noguchi K, Terashima I (2014) Roles of cyclic electron flow around PSI (CEF-PSI) and O2-dependent alternative pathways in regulation of the photosynthetic electron flow in short-term fluctuating light in Arabidopsis thaliana. Plant Cell Physiol 55:990–1004. doi:10.1093/pcp/pcu033
Kornyeyev D, Logan BA, Allen RD, Holaday AS (2003a) Effect of chloroplastic overexpression of ascorbate peroxidase on photosynthesis and photoprotection in cotton leaves subjected to low temperature photoinhibition. Plant Sci 165:1033–1041. doi:10.1016/S0168-9452(03)00294-2
Kornyeyev D, Logan BA, Payton PR, Allen RD, Holaday AS (2003b) Elevated chloroplastic glutathione reductase activities decrease chilling-induced photoinhibition by increasing rates of photochemistry, but not thermal energy dissipation, in transgenic cotton. Funct Plant Biol 30:101–110. doi:10.1071/FP02144
Kornyeyev D, Logan BA, Holaday AS (2010) Excitation pressure as a measure of the sensitivity of photosystem II to photoinactivation. Funct Plant Biol 37:943–951. doi:10.1071/fp09276
Kozaki A, Takeba G (1996) Photorespiration protects C3 plants from photooxidation. Nature 384:557–560. doi:10.1038/384557a0
Kramer DM, Avenson TJ, Edwards GE (2004) Dynamic flexibility in the light reactions of photosynthesis governed by both electron and proton transfer reactions. Trends Plant Sci 9:349–357. doi:10.1016/j.tplants.2004.05.001
Kudoh H, Sonoike K (2002) Irreversible damage to photosystem I by chilling in the light: cause of the degradation of chlorophyll after returning to normal growth temperature. Planta 215:541–548. doi:10.1007/s00425-002-0790-9
Kuntz M (2004) Plastid terminal oxidase and its biological significance. Planta 218:896–899. doi:10.1007/s00425-004-1217-6
Kuvykin IV, Ptushenko VV, Vershubskii AV, Tikhonov AN (2011) Regulation of electron transport in C3 plant chloroplasts in situ and in silico. Short-term effects of atmospheric CO2 and O2. Biochim Biophys Acta 1807:336–347. doi:10.1016/j.bbabio.2010.12.012
Laisk A, Eichelmann H, Oja V, Peterson RB (2005) Control of cytochrome b6f at low and high light intensity and cyclic electron transport in leaves. Biochim Biophys Acta 1708:79–90. doi:10.1016/j.bbabio.2005.01.007
Laisk A, Eichelmann H, Oja V, Talts E, Scheibe R (2007) Rates and roles of cyclic and alternative electron flow in potato leaves. Plant Cell Physiol 48:1575–1588. doi:10.1093/pcp/pcm129
Lawson T, Oxborough K, Morison JIL, Baker NR (2002) Responses of photosynthetic electron transport in stomatal guard cells and mesophyll cells in intact leaves to light, CO2, and humidity. Plant Physiol 128:52–62. doi:10.1104/pp.010317
Leakey ADB, Scholes JD, Press MC (2004) Physiological and ecological significance of sunflecks for dipterocarp seedlings. J Exp Bot 56:469–482. doi:10.1093/jxb/eri055
Li XP, Björkman O, Shih C, Grossman AR, Rosenquist M, Jansson S, Niyogi KK (2000) A pigment-binding protein essential for regulation of photosynthetic light harvesting. Nature 403:391–395. doi:10.1038/35000131
Li XP, Muller-Moule P, Gilmore AM, Niyogi KK (2002) PsbS-dependent enhancement of feedback de-excitation protects photosystem II from photoinhibition. Proc Natl Acad Sci USA 99:15222–15227. doi:10.1073/pnas.232447699
Long SP, Zhu XG, Naidu SL, Ort DR (2006) Can improvement in photosynthesis increase crop yields? Plant Cell Environ 29:315–330
Lu N, Nukaya T, Kamimura T et al (2015) Control of vapor pressure deficit (VPD) in greenhouse enhanced tomato growth and productivity during the winter season. Sci Hortic 197:17–23. doi:10.1016/j.scienta.2015.11.001
Ludlow MM, Bjorkman O (1984) Paraheliotropic leaf movement in Siratro as a protective mechanism against drought-induce damage to primary photosynthetic reactions: damage by excessive light and heat. Planta 161:505–518. doi:10.1007/BF00407082
Makino A, Miyake C, Yokota A (2002) Physiological functions of the water-water cycle (Mehler reaction) and the cyclic electron flow around PSI in rice leaves. Plant Cell Physiol 43:1017–1026. doi:10.1093/pcp/pcf124
McDonald AE, Ivanov AG, Bode R, Maxwell DP, Rodermel SR, Huner NPA (2011) Flexibility in photosynthetic electron transport: the physiological role of plastoquinol terminal oxidase (PTOX). Biochim Biophys Acta 1807:954–967. doi:10.1016/j.bbabio.2010.10.024
Medlyn BE, Barton CVM, Broadmeadow MSJ et al (2001) Stomatal conductance of forest species after long-term exposure to elevated CO2 concentration: a synthesis. New Phytol 149:247–264. doi:10.1046/j.1469-8137.2001.00028.x
Mehler AH (1951) Studies on reactions of illuminated chloroplasts. I. Mechanisms of the reduction of oxygen and other Hill reagents. Arch Biochem Biophys 33:65–77. doi:10.1016/0003-9861(51)90082-3
Miyake C (2010) Alternative electron flows (water-water cycle and cyclic electron flow around PSI) in photosynthesis: molecular mechanisms and physiological functions. Plant Cell Physiol 51:1951–1963. doi:10.1093/pcp/pcq173
Miyake C, Okamura M (2003) Cyclic electron flow within PSII protects PSII from its photoinhibition in thylakoid membranes from spinach chloroplasts. Plant Cell Physiol 44:457–462. doi:10.1093/pcp/pcg053
Miyake C, Miyata M, Shinzaki Y, Tomizawa K (2005) CO2 response of cyclic electron flow around PSI (CEF-PSI) in tobacco leaves—relative electron fluxes through PSI and PSII determine the magnitude of non-photochemical quenching (NPQ) of Chl fluorescence. Plant Cell Physiol 46:629–637. doi:10.1093/pcp/pci067
Morison JIL (1998) Stomatal response to increased CO2 concentration. J Exp Bot 49:443–453. doi:10.1093/jxb/49.Special_Issue.443
Müller P, Li XP, Niyogi KK (2001) Non-photochemical quenching. A response to excess light energy. Plant Physiol 125:1558–1566. doi:10.1104/pp.125.4.1558
Munekage Y, Hojo M, Meurer J et al (2002) PGR5 is involved in cyclic electron flow around photosystem I and is essential for photoprotection in Arabidopsis. Cell 110:361–371. doi:10.1016/S0092-8674(02)00867-X
Munekage Y, Hashimoto M, Miyake C et al (2004) Cyclic electron flow around photosystem I is essential for photosynthesis. Nature 429:579–582. doi:10.1038/nature02598
Murchie EH, Chen YZ, Hubbart S, Peng SB, Horton P (1999) Interactions between senescence and leaf orientation determine in situ patterns of photosynthesis and photoinhibition in field-grown rice. Plant Physiol 119:553–563. doi:10.1104/pp.119.2.553
Naumburg E, Ellsworth DS (2002) Short-term light and leaf photosynthetic dynamics affect estimates of daily understory photosynthesis in four tree species. Tree Physiol 22:393–401. doi:10.1093/treephys/22.6.393
Nawrocki WJ, Tourasse NJ, Taly A, Rappaport F, Wollman FA (2015) The plastid terminal oxidase: its elusive function points to multiple contributions to plastid physiology. Ann Rev Plant Biol 66:49–74. doi:10.1146/annurev-arplant-043014-114744
Nishiyama Y, Allakhverdiev SI, Murata N (2006) A new paradigm for the action of reactive oxygen species in the photoinhibition of photosystem II. Biochim Biophys Acta 1757:742–749. doi:10.1016/j.bbabio.2006.05.013
Niyogi KK (1999) Photoprotection revisited: genetic and molecular approaches. Annu Rev Plant Physiol Plant Mol Biol 50:333–359. doi:10.1146/annurev.arplant.50.1.333
Niyogi KK (2000) Safety valves for photosynthesis. Curr Opin Plant Biol 3:455–460. doi:10.1016/S1369-5266(00)00113-8
Niyogi KK, Li XP, Rosenberg V, Jung HS (2005) Is PsbS the site of non-photochemical quenching in photosynthesis? J Exp Bot 56:375–382. doi:10.1093/jxb/eri056
Noctor G, Dutilleul C, De Paepe R, Foyer CH (2004) Use of mitochondrial electron transport mutants to evaluate the effects of redox state on photosynthesis, stress tolerance and the integration of carbon/nitrogen metabolism. J Exp Bot 55:49–57. doi:10.1093/jxb/erh021
Noguchi K, Yoshida K (2008) Interaction between photosynthesis and respiration in illuminated leaves. Mitochondrion 8:87–99. doi:10.1016/j.mito.2007.09.003
Ogren WL (1984) Photorespiration: pathways, regulation, and modification. Ann Rev Plant Physiol 35:415–442. doi:10.1146/annurev.pp.35.060184.002215
Ögren E, Evans JR (1993) Photosynthetic light response curves. 1. The influence of CO2 partial pressure and leaf inversion. Planta 189:182–190. doi:10.1007/BF00195075
Ort DR, Baker NR (2002) A photoprotective role for O2 as an alternative electron sink in photosynthesis? Curr Opp Plant Biol 5:193–197. doi:10.1016/S1369-5266(02)00259-5
Osmond CB (1981) Photorespiration and photoinhibition. Some implications for the energetics of photosynthesis. Biochim Biophys Acta 639:77–98. doi:10.1016/0304-4173(81)90006-9
Pastenes C, Pimentel P, Lillo J (2005) Leaf movements and photoinhibition in relation to water stress in field-grown beans. J Exp Bot 56:425–433. doi:10.1093/jxb/eri061
Peak D, Mott KA (2011) A new, vapour-phase mechanism for stomatal responses to humidity and temperature. Plant Cell Environ 34:162–178. doi:10.1111/j.1365-3040.2010.02234.x
Pearcy RW (1988) Photosynthetic utilization of lightflecks by understory plants. Aust J Plant Physiol 15:223–238. doi:10.1071/PP9880223
Pearcy RW (1990) Sunflecks and photosynthesis in plant canopies. Annu Rev Plant Physiol Plant Mol Biol 41:421–453. doi:10.1146/annurev.pp.41.060190.002225
Pearcy RW, Way DA (2012) Two decades of sunfleck research: looking back to move forward. Tree Physiol 32:1059–1061. doi:10.1093/treephys/tps084
Peltier G, Cournac L (2002) Chlororespiration. Annu Rev Plant Biol 53:523–550. doi:10.1146/annurev.arplant.53.100301.135242
Peng L, Yamamoto H, Shikanai T (2011) Structure and biogenesis of the chloroplast NAD(P)H dehydrogenase complex. Biochim Biophys Acta 1807:945–953. doi:10.1016/j.bbabio.2010.10.015
Pfitsch WA, Pearcy RW (1989) Steady-state and dynamic photosynthetic response of Adenocaulon bicolor (Asteraceae) in its redwood forest habitat. Oecologia 80:471–476. doi:10.1007/BF00380068
Portis AR Jr (2003) Rubisco activase: Rubisco’s catalytic chaperone. Photosynth Res 75:11–27. doi:10.1023/A:1022458108678
Powles SB (1984) Photoinhibition of photosynthesis induced by visible light. Annu Rev Plant Physiol 35:15–44. doi:10.1146/annurev.pp.35.060184.000311
Prasil O, Kolber Z, Berry JA, Falkowski PG (1996) Cyclic electron flow around photosystem II in vivo. Photosynth Res 48:395–410. doi:10.1007/BF00029472
Raghavendra AS, Padmasree K (2003) Beneficial interactions of mitochondrial metabolism with photosynthetic carbon assimilation. Trends Plant Sci 8:546–553. doi:10.1016/j.tplants.2003.09.015
Rawson HM, Begg JE, Woodward RG (1977) The effect of atmospheric humidity on photosynthesis, transpiration and water use efficiency of leaves of several plant species. Planta 134:5–10. doi:10.1007/BF00390086
Rijkers T, de Vries PJ, Pons TL, Bongers F (2000) Photosynthetic induction in saplings of three shade-tolerant tree species: comparing understorey and gap habitats in a French Guiana rain forest. Oecologia 125:331–340. doi:10.1007/s004420000459
Ruuska SA, Badger MR, Andrews TJ, von Caemmerer S (2000) Photosynthetic electron sinks in transgenic tobacco with reduced amounts of Rubisco: little evidence for significant Mehler reaction. J Exp Bot 51:357–368. doi:10.1093/jexbot/51.suppl_1.357
Sage RF, Sage TL, Kocacinar F (2012) Photorespiration and the evolution of C4 photosynthesis. Ann Rev Plant Biol 63:19–47. doi:10.1146/annurev-arplant-042811-105511
Salvucci ME, Crafts-Brandner SJ (2004) Relationship between the heat tolerance of photosynthesis and the thermal stability of Rubisco activase in plants from contrasting thermal environments. Plant Physiol 134:1460–1470. doi:10.1104/pp.103.038323
Sassenrath-Cole GF, Pearcy RW (1992) The role of ribulose-1,5-bisphosphate regeneration in the induction requirement of photosynthetic CO2 exchange under transient light conditions. Plant Physiol 99:227–234
Sassenrath-Cole GF, Pearcy RW (1994) Regulation of photosynthetic induction state by the magnitude and duration of low light exposure. Plant Physiol 105:1115–1123. doi:10.1104/pp.105.4.1115
Scheibe R (2004) Malate valves to balance cellular energy supply. Physiol Plant 120:21–26. doi:10.1111/j.0031-9317.2004.0222.x
Scheibe R, Backhausen JE, Emmerlich V, Holtgrefe S (2005) Strategies to maintain redox homeostasis during photosynthesis under changing conditions. J Exp Bot 56:1481–1489. doi:10.1093/jxb/eri181
Schymanski SJ, Or D, Zwieniecki M (2013) Stomatal control and leaf thermal and hydraulic capacitances under rapid environmental fluctuations. PLoS One 8:e54231. doi:10.1371/journal.pone.0054231
Shikanai T, Endo T, Hashimoto T et al (1998) Directed disruption of the tobacco ndhB gene impairs cyclic electron flow around photosystem I. Proc Natl Acad Sci USA 95:9705–9709
Sims DA, Pearcy RW (1993) Sunfleck frequency and duration affect growth-rate of the understorey plant, Alocasia macrorrhiza. Funct Ecol 7:683–689. doi:10.2307/2390189
Singsaas EL, Sharkey TD (1998) The regulation of isoprene emission responses to rapid leaf temperature fluctuations. Plant Cell Environ 21:1181–1188. doi:10.1046/j.1365-3040.1998.00380.x
Singsaas EL, Laporte MM, Shi JZ et al (1999) Leaf temperature fluctuation affects isoprene emission from red oak (Quercus rubra) leaves. Tree Physiol 19:917–924. doi:10.1093/treephys/19.14.917
Smith H (1982) Light quality, photoperception, and plant strategy. Ann Rev Plant Physiol 33:481–518. doi:10.1146/annurev.pp.33.060182.002405
Sonoike K (2011) Photoinhibition of photosystem I. Physiol Plant 142:56–64. doi:10.1111/j.1399-3054.2010.01437.x
Sonoike K, Terashima I (1994) Mechanism of photosystem-I photoinhibition in leaves of Cucumis sativus L. Planta 194:287–293. doi:10.1007/BF00196400
Streb P, Josse EM, Gallouet E, Baptist F, Kuntz M, Cornic G (2005) Evidence for alternative electron sinks to photosynthetic carbon assimilation in the high mountain plant species Ranunculus glacialis. Plant Cell Environ 28:1123–1135. doi:10.1111/j.1365-3040.2005.01350.x
Suetsugu N, Wada M (2007) Chloroplast photorelocation movement mediated by phototropin family proteins in green plants. Biol Chem 388:927–935. doi:10.1515/BC.2007.118
Suorsa M, Järvi S, Grieco M et al (2012) PROTON GRADIENT REGULATION5 is essential for proper acclimation of Arabidopsis photosystem I to naturally and artificially fluctuating light conditions. Plant Cell 24:2934–2948. doi:10.1105/tpc.112.097162
Takahashi S, Badger MR (2011) Photoprotection in plants: a new light on photosystem II damage. Trends Plant Sci 16:53–60. doi:10.1016/j.tplants.2010.10.001
Takahashi S, Murata N (2008) How do environmental stresses accelerate photoinhibition? Trends Plant Sci 13:178–182. doi:10.1016/j.tplants.2008.01.005
Takahashi S, Bauwe H, Badger M (2007) Impairment of the photorespiratory pathway accelerates photoinhibition of photosystem II by suppression of repair process and not acceleration of damage process in Arabidopsis thaliana. Plant Physiol 144:487–494. doi:10.1104/pp.107.097253
Takahashi S, Milward SE, Fan DY, Chow WS, Badger MR (2009) How does cyclic electron flow alleviate photoinhibition in Arabidopsis? Plant Physiol 149:1560–1567. doi:10.1104/pp.108.134122
Terashima I, Funayama S, Sonoike K (1994) The site of photoinhibition in leaves of Cucumis sativus L. at low temperatures is photosystem I, not photosystem II. Planta 193:300–306. doi:10.1007/BF00192544
Terashima I, Araya T, Miyazawa SI, Sone K, Yano S (2005) Construction and maintenance of the optimal photosynthetic systems of the leaf, herbaceous plant and tree: an eco-developmental treatise. Ann Bot 95:507–519. doi:10.1093/aob/mci049
Tikkanen M, Aro EM (2012) Thylakoid protein phosphorylation in dynamic regulation of photosystem II in higher plants. Biochim Biophys Acta 1817:232–238. doi:10.1016/j.bbabio.2011.05.005
Tikkanen M, Grieco M, Kangasjärvi S, Aro EM (2010) Thylakoid protein phosphorylation in higher plant chloroplasts optimizes electron transfer under fluctuating light. Plant Physiol 152:723–735. doi:10.1104/pp.109.150250
Tikkanen M, Mekala NR, Aro EM (2014) Photosystem II photoinhibition-repair cycle protects photosystem I from irreversible damage. Biochim Biophys Acta 1837:210–215. doi:10.1016/j.bbabio.2013.10.001
Tikkanen M, Rantala S, Aro EM (2015) Electron flow from PSII to PSI under high light is controlled by PGR5 but not by PSBS. Front Plant Sci 6:521. doi:10.3389/fpls.2015.00521
Timm HC, Küppers M, Stegemann J (2004) Non-destructive analysis of architectural expansion and assimilate allocation in different tropical tree saplings: consequences of using steady-state and dynamic photosynthesis models. Ecotropica 10:101–121
Trouillard M, Shahbazi M, Moyet L, Rappaport F, Joliot P, Kuntz M, Finazzi G (2012) Kinetic properties and physiological role of the plastoquinone terminal oxidase (PTOX) in a vascular plant. Biochim Biophys Acta 1817:2140–2148. doi:10.1016/j.bbabio.2012.08.006
Valladares F, Allen MT, Pearcy RW (1997) Photosynthetic responses to dynamic light under field conditions in six tropical rainforest shrubs occurring along a light gradient. Oecologia 111:505–514. doi:10.1007/s004420050264
Wada M, Kagawa T, Sato Y (2003) Chloroplast movement. Annu Rev Plant Biol 54:455–468. doi:10.1146/annurev.arplant.54.031902.135023
Watling JR, Ball MC, Woodrow IE (1997) The utilization of lightflecks for growth in four Australian rain-forest species. Funct Ecol 11:231–239. doi:10.1046/j.1365-2435.1997.00073.x
Way DA, Pearcy RW (2012) Sunflecks in trees and forests: from photosynthetic physiology to global change biology. Tree Physiol 32:1066–1081. doi:10.1093/treephys/tps064
Way DA, Yamori W (2014) Thermal acclimation of photosynthesis: on the importance of definitions and accounting for thermal acclimation of respiration. Photosynth Res 119:89–100. doi:10.1007/s11120-013-9873-7
Wise RR, Olson AJ, Schrader SM, Sharkey TD (2004) Electron transport is the functional limitation of photosynthesis in field-grown pima cotton plants at high temperature. Plant Cell Environ 27:717–724. doi:10.1111/j.1365-3040.2004.01171.x
Yamamoto H, Shikanai T (2013) In planta mutagenesis of Src homology 3 domain-like fold of NdhS, a ferredoxin-binding subunit of the chloroplast NADH dehydrogenase-like complex in Arabidopsis. A conserved Arg-193 plays a critical role in ferredoxin binding. J Biol Chem 288:36328–36337. doi:10.1074/jbc.M113.511584
Yamamoto H, Peng L, Fukao Y, Shikanai T (2011) An Src homology 3 domain-like fold protein forms a ferredoxin-binding site for the chloroplast NADH dehydrogenase-like complex in Arabidopsis. Plant Cell 23:1480–1493. doi:10.1105/tpc.110.080291
Yamori W (2013) Improving photosynthesis to increase food and fuel production by biotechnological strategies in crops. J Plant Biochem Physiol 1:113. doi:10.4172/2329-9029.1000113
Yamori W, Shikanai T (2016) Physiological functions of cyclic electron transport around photosystem I in sustaining photosynthesis and plant growth. Annu Rev Plant Biol. doi:10.1146/annurev-arplant-043015-112002
Yamori W, von Caemmerer S (2009) Effect of Rubisco activase deficiency on the temperature response of CO2 assimilation rate and Rubisco activation state: insights from transgenic tobacco with reduced amounts of Rubisco activase. Plant Physiol 151:2073–2082. doi:10.1104/pp.109.146514
Yamori W, Noguchi K, Terashima I (2005) Temperature acclimation of photosynthesis in spinach leaves: analyses of photosynthetic components and temperature dependencies of photosynthetic partial reactions. Plant Cell Environ 28:536–547. doi:10.1111/j.1365-3040.2004.01299.x
Yamori W, Suzuki K, Noguchi K, Nakai M, Terashima I (2006) Effects of Rubisco kinetics and Rubisco activation state on the temperature dependence of the photosynthetic rate in spinach leaves from contrasting growth temperatures. Plant Cell Environ 29:1659–1670. doi:10.1111/j.1365-3040.2006.01550.x
Yamori W, Evans JR, von Caemmerer S (2010a) Effects of growth and measurement light intensities on temperature dependence of CO2 assimilation rate in tobacco leaves. Plant Cell Environ 33:332–343. doi:10.1111/j.1365-3040.2009.02067.x
Yamori W, Noguchi K, Hikosaka K, Terashima I (2010b) Phenotypic plasticity in photosynthetic temperature acclimation among crop species with different cold tolerances. Plant Physiol 152:388–399. doi:10.1104/pp.109.145862
Yamori W, Sakata N, Suzuki Y, Shikanai T, Makino A (2011) Cyclic electron flow around photosystem I via chloroplast NAD(P)H dehydrogenase (NDH) complex performs a significant physiological role during photosynthesis and plant growth at low temperature in rice. Plant J 68:966–976. doi:10.1111/j.1365-313X.2011.04747.x
Yamori W, Masumoto C, Fukayama H, Makino A (2012) Rubisco activase is a key regulator of non steady-state photosynthesis at any leaf temperature and to a lesser extent, of steady-state photosynthesis at high temperature. Plant J 71:871–880. doi:10.1111/j.1365-313X.2012.05041.x
Yamori W, Hikosaka K, Way DA (2014) Temperature response of photosynthesis in C3, C4 and CAM plants: temperature acclimation and temperature adaptation. Photosynth Res 119:101–117. doi:10.1007/s11120-013-9874-6
Yamori W, Shikanai T, Makino A (2015) Photosystem I cyclic electron flow via chloroplast NADH dehydrogenease-like complex performs a physiological role for photosynthesis at low light. Sci Rep 5:13908. doi:10.1038/srep13908
Yamori W, Makino A, Shikanai T (2016a) A physiological role of cyclic electron transport around photosystem I in sustaining photosynthesis under fluctuating light in rice. Sci Rep 6:20147. doi:10.1038/srep20147
Yamori W, Kondo E, Sugiura D et al (2016b) Enhanced leaf photosynthesis as a target to increase grain yield: insights from transgenic rice lines with variable Rieske FeS protein content in the cytochrome b6/f complex. Plant Cell Environ 39:80–87. doi:10.1111/pce.12594
Yin ZH, Johnson GN (2000) Photosynthetic acclimation of higher plants to growth in fluctuating light environments. Photosynth Res 63:97–107. doi:10.1023/A:1006303611365
Zhu XG, Portis AR, Long SP (2004) Would transformation of C3 crop plants with foreign Rubisco increase productivity? A computational analysis extrapolating from kinetic properties to canopy photosynthesis. Plant Cell Environ 27:155–165. doi:10.1046/j.1365-3040.2004.01142.x
Zhu XG, Long SP, Ort DR (2010) Improving photosynthetic efficiency for greater yield. Annu Rev Plant Biol 61:235–261. doi:10.1146/annurev-arplant-042809-112206
Acknowledgments
This work was supported by the Japan Science and Technology Agency, PRESTO (to W.Y.).
Author information
Authors and Affiliations
Corresponding author
Rights and permissions
About this article
Cite this article
Yamori, W. Photosynthetic response to fluctuating environments and photoprotective strategies under abiotic stress. J Plant Res 129, 379–395 (2016). https://doi.org/10.1007/s10265-016-0816-1
Received:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1007/s10265-016-0816-1