Obstacles in the quantification of the cyclic electron flux around ...

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Feb 4, 2016 - Sixty years ago Arnon and co-workers discovered photophosphorylation driven by a cyclic electron flux (CEF) around Photosystem I. Since ...
Photosynth Res DOI 10.1007/s11120-016-0223-4

REVIEW

Obstacles in the quantification of the cyclic electron flux around Photosystem I in leaves of C3 plants Da-Yong Fan1,5 • Duncan Fitzpatrick5 • Riichi Oguchi2,5 • Weimin Ma3,5 Jiancun Kou4,5 • Wah Soon Chow5



Received: 27 October 2015 / Accepted: 24 January 2016 Ó Springer Science+Business Media Dordrecht 2016

Abstract Sixty years ago Arnon and co-workers discovered photophosphorylation driven by a cyclic electron flux (CEF) around Photosystem I. Since then understanding the physiological roles and the regulation of CEF has progressed, mainly via genetic approaches. One basic problem remains, however: quantifying CEF in the absence of a net product. Quantification of CEF under physiological conditions is a crucial prerequisite for investigating the physiological roles of CEF. Here we summarize current progress in methods of CEF quantification in leaves and, in some cases, in isolated thylakoids, of C3 plants. Evidently, all present methods have their own shortcomings. We conclude that to quantify CEF in vivo, the best way currently is to measure the electron flux through PS I (ETR1) and that through PS II and PS I in series (ETR2) for the whole leaf tissue under identical conditions. The difference between ETR1 and ETR2 is an upper estimate of CEF, mainly consisting, in C3 plants, of a major PGR5–PGRL1dependent CEF component and a minor chloroplast NDHdependent component, where PGR5 stands for Proton Gradient Regulation 5 protein, PGRL1 for PGR5-like

& Wah Soon Chow [email protected]

photosynthesis phenotype 1, and NDH for Chloroplast NADH dehydrogenase-like complex. These two CEF components can be separated by the use of antimycin A to inhibit the former (major) component. Membrane inlet mass spectrometry utilizing stable oxygen isotopes provides a reliable estimation of ETR2, whilst ETR1 can be estimated from a method based on the photochemical yield of PS I, Y(I). However, some issues for the recommended method remain unresolved. Keywords Cyclic electron flux  Linear electron flux  Membrane inlet mass spectrometry  P700  Photosystem I  Photosystem II Abbreviations ATP CEF Chl Cyt DFlux DpH

Adenosine triphosphate Cyclic electron flux around PS I Chlorophyll Cytochrome Difference between ETR1 and ETR2 Trans-thylakoid membrane pH difference

1

Da-Yong Fan [email protected]

State Key Laboratory of Vegetation and Environmental Change, Institute of Botany, The Chinese Academy of Sciences, Beijing 100093, China

2

Duncan Fitzpatrick [email protected]

Graduate School of Life Sciences, Tohoku University, Sendai, Miyagi 980-8578, Japan

3

Riichi Oguchi [email protected]

College of Life & Environment Sciences, Shanghai Normal University, Guilin Road 100, Shanghai 200234, China

4

Weimin Ma [email protected]

College of Animal Science and Technology, Northwest A&F University, Yangling 712100, China

5

Division of Plant Sciences, Research School of Biology, The Australian National University, 46 Sullivans Creek Road, Acton, ACT 2601, Australia

Jiancun Kou [email protected]

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Dw DCMU ECS ES ETR1, ETR2 ETR2(O2) fI and fII Fs and Fm

FNR I LHCII MIMS MV NADPH NDH P700 Pc PGR5 PGRL1 Pmf PQ PS II, PS I PsaF PTOX QA, QB qE Rph Y(I) and Y(II)

Trans-thylakoid membrane electric potential difference 3-(3:4-dichlorophenyl)-1:1-dimethylurea Electrochromic shift Energy storage The electron flux through PS I, PS II, respectively The electron flux through PS II, measured by gross O2 evolution The fraction of absorbed light partitioned to PS I and PS II, respectively Steady-state and maximum Chl fluorescence yield during illumination, respectively Ferredoxin/NADP reductase Irradiance Light-harvesting chlorophyll-protein complex II Membrane inlet mass spectrometry Methyl viologen Reduced nicotinamide adenine dinucleotide phosphate Chloroplast NADH dehydrogenase-like complex Special chlorophyll pair in PS I Plastocyanin Proton gradient regulation 5 protein PGR5-like photosynthesis phenotype 1 Protonmotive force Plastoquinone Photosystem II, Photosystem I, respectively Plastocyanin-binding subunit in PS I Plastid terminal oxidase Primary and secondary quinine electron acceptor in PS II, respectively Energy-dependent quenching The sum of the photochemical rates The photochemical yield of PS I and PSII, respectively, during illumination

Introduction Arnon et al. (1955) demonstrated photophosphorylation on illumination of isolated thylakoids in the presence of vitamin K via a cyclic electron flux around PS I (CEF). Since CEF is essential for efficient photosynthesis (Munekage et al. 2002) and photoprotection (Takahashi et al. 2009; Kukuczka et al. 2014), there has been a sustained effort to elucidate the mechanisms and roles of this cyclic electron flow (for reviews, see Bendall and Manasse

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1995; Allen 2003; Bukhov and Carpentier 2004; Johnson 2005, 2011; Joliot and Joliot 2006; Shikanai 2007, 2014; Alric 2010, 2015; Miyake 2010; Kramer and Evans 2011; Suorsa 2015). It is supposed that CEF has two major functions: to supply the ATP shortfall for the Calvin– Benson cycle as required by a 3 ATP: 2 NADPH stoichiometry in C3 photosynthesis, and to supplement the proton gradient that underpins non-photochemical quenching for photoprotection of PS II. Crucially, CEF confines electrons to travel along a physiologically normal cyclic path of reducing ferredoxin (Fd) and NADP? instead of forming damaging reactive oxygen species on the acceptor side of PS I (Chow and Hope 1998). A prerequisite to understanding the role of CEF is a knowledge of the magnitude of CEF under physiological conditions. However, quantification of CEF in physiological conditions is notoriously difficult due to the absence of a net product of cyclic electron flow (Shikanai 2014). Additional electron fluxes through PS I, which are hard to be separated from CEF, increase the complexity of interpreting the data, whilst regulation of CEF has been found to be highly sensitive to the redox poising of electron carriers between the two photosystems, and to the proton gradient (DpH) across the thylakoid membrane (Heber et al. 1978; Allen 2003; Kramer and Evans, 2011). Accordingly, any experimental procedures that change the redox poise and DpH concomitantly alter CEF. It is worth noting that conditions used in measurements in vitro are unlikely to be optimized for electron transport, resulting in in vitro estimations of CEF that are considerably lower than estimations under natural conditions. For example, Strand et al. (2015b) found that CEF rate is very low when measured in vitro, but considerably higher when measured in vivo. One plausible explanation for this discrepancy is the alteration of the in vitro redox poise of CEF components from the optimum. For example, after isolation of thylakoids, the granal stacks may not be as well formed as they are in vivo, in which case PS II is closer to PS I on average. This may result in over-reduction of the PQ molecules near where PS I is located and where it performs cyclic electron transport. An over-reduction of PQ molecules is not conducive to cyclic electron flow, and one function of the lateral heterogeneity in the distribution of the two photosystems is to avoid an over-reduction of the PQ pool in the vicinity of PS I complexes performing CEF (Alric 2015). Some isolation and rupture procedures used for in vitro assays may perturb the highly conserved, functionally significant fine structure of chloroplasts (Kirchhoff et al. 2013), whilst structural integrity has been demonstrated to be associated with CEF (Joliot and Joliot 2002; Johnson 2011). The integrity of protein complexes involved in CEF within the thylakoid membrane may be altered during these

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procedures. For example, one postulation for the absence of PGR5 in an isolated supercomplex in C. reinhardtii is that it might have been lost during the biochemical manipulations (Iwai et al. 2010). The absence of PGR5 would affect CEF of the isolated complex, since a recent study showed that the functional attributes of the C. reinhardtii PGR5 mutant are consistent with those attributes observed for the analogous mutants in vascular plants, suggesting that PGR5 is essential for CEF activity in C. reinhardtii (Johnson et al. 2014). This is consistent with the conclusion that PGR5 is crucial for CEF function and hence performance in Arabidopsis, particularly in a fluctuating and high-light environment (Munekage et al. 2002; Dang et al. 2014; Kono et al. 2014; Suorsa 2015). Current methods for measuring/inferring CEF can be grouped into two categories. In Category 1, at least eight methods have been devised to monitor CEF directly. In Category 2, CEF can be estimated from the difference between linear electron transport through PS II (ETR2) and total flux through PS I (ETR1).

Category 1: Direct monitoring/measurement of CEF

to monitor redox changes of ferredoxin in a light-to-dark transition. It was possible to monitor a form of CEF, but only in the absence of ETR2 and in anaerobic conditions. Post-illumination increase in Chl fluorescence yield The transient increase in chlorophyll (Chl) fluorescence yield on cessation of actinic illumination reflects the presence of CEF (Endo et al. 1998). Munekage et al. (2002) used an in vitro assay, combined with the post-illumination increase in Chl fluorescence yield, to test for CEF in conditions in which PQ was reduced by NADPH, in the presence of Fd and under low measurement irradiance (\10 lmol photons m-2 s-1). However, this reaction in the in vitro assay would have been too slow to be compatible with a CEF measured in vivo by post-illumination P700? re-reduction and the onset kinetics of P700 oxidation (Nandha et al. 2007), and probably is unrelated to CEF activity but related to a subpopulation of PS II with low potential variants of QA which is directly reducible by ferredoxin (Fisher and Kramer 2014). Secondly, the original assay in the dark does not provide quantitative information for the leaves when illuminated with typical actinic light (Shikanai 2014). This way of monitoring CEF is basically qualitative (Gotoh et al. 2010).

ATP synthesis Post-illumination P7001 re-reduction Reliable direct measurements of ATP synthesis driven by CEF can only be carried out in vitro. These measurements have demonstrated that a cyclic electron flux does lead to ATP synthesis (Bendall and Manasse 1995; Allen 2003). Therefore, the ATP synthesis rate may indicate the size of CEF. Unfortunately, the proton conductivity of the thylakoid membrane (gH?, Kramer et al. 2004), changes under variable environmental conditions such as varying CO2 concentrations (Kanazawa and Kramer 2002), or in the Arabidopsis pgr5 mutant (Shikanai 2014), such that the coupling between ATP synthesis and electron flow may be variable. In addition, mitochondrial respiration interacts with chloroplast ATP production through the export of reducing equivalents from the chloroplast via the malateoxalate cycle (Johnson 2011). Both effects render such a method inappropriate for quantification of CEF in vivo. The Mehler reaction, as part of the water–water cycle particularly at high irradiance, also leads to ATP synthesis (Badger 1985), as does a linear electron flux. Redox changes of ferredoxin In an in vitro system, Cleland and Bendall (1992) preilluminated thylakoids in anaerobic conditions to set up an elaborate redox poising sequence, followed by the addition of DCMU to eliminate ETR2, finally applying actinic light

This method stems from a light-induced absorbance change in the range 800–850 nm that reflects the operation of PS I, specifically the photo-oxidation of P700, the primary electron donor in PS I (Harbinson and Woodward 1987; Schreiber et al. 1988). A quantitative assay of CEF via the post-illumination rereduction of P700? is based on the following rationale: when a steady-state level of P700? is attained in the light, the rate of electron flow through PS I is thought to equal the rate of P700? reduction measured immediately after cessation of illumination (Maxwell and Biggins 1976). The addition of DCMU inhibits the linear electron flux, ETR2, allowing the deduction of CEF (Joliot and Joliot 2002). As the redox poising of electron carriers involved in CEF is altered in the presence of DCMU, however, the CEF so obtained from spinach leaf discs during the photosynthetic induction period is much diminished (Fan et al. 2007). Allen (2003) pointed out that the optimal redox condition for CEF operation is attained when half of the PQ pool is reduced and half of Fd is reduced; the electrons then can cycle around P700 without the limitation of donor or acceptor sides of PSI. In the presence of DCMU, particularly when electron sinks at the acceptor side of PSI are active (for example, under aerobic conditions), on the one hand PSII cannot provide electrons to PQ pool, and on the

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other hand electrons on the acceptor side of PSI will be diverted to other pathways. Therefore, the re-injection of electrons into the oxidized PQ pool would gradually decrease, and consequently CEF would diminish. That is the reason for DCMU-treated leaves in aerobic conditions having a very slow post-illumination P700? re-reduction rate (Fan et al. 2007). A second pitfall, under certain circumstances such as anaerobicity or a highly reduced acceptor side of PS I (particularly for an in vitro system), results from the promotion of fast charge recombination of a light-induced radical pair. In such cases the P700? re-reduction is at least a combination of CEF and charge recombination, making the estimation unreliable (Takahashi et al. 2013). In another variation, CEF could be eliminated by methyl viologen (MV), and the difference of post-illumination P700? re-reduction rate with and without MV could be attributed to CEF. However, MV itself retards linear electron flow to some extent during continuous illumination (Jia et al. 2008; Fan et al. 2009), necessitating a correction of ETR2 in the presence of MV. The correction procedure using Chl fluorescence further suffers from the pitfall that a modulated fluorescence signal comes from a localized and uncertain depth in the leaf tissue (Oguchi et al. 2011); therefore, other ways of correction may need to be applied. Further, MV should be applied with caution, as MV can generate reactive oxygen species in aerobic conditions. A most serious objection to the use of the initial postillumination re-reduction of P700? as a quantitative measure of CEF is the magnitude of the initial slope, which is too small even when compared with the linear electron flux obtained from parallel oxygen evolution measurements (see the section ‘‘Measurement of ETR1 via post-illumination P700? re-reduction’’ below). The onset kinetics of P700 oxidation CEF could be monitored by the oxidation kinetics of P700 illuminated with far-red light (Joliot and Joliot 2002, 2005; Nandha et al. 2007; Vredenberg and Bulychev 2010). The extremely variable P700 oxidation (reflecting variation in the dark redox state of the chloroplast) could be removed by a short-time pulse of saturating green light (Joliot and Joliot 2005; Nandha et al. 2007). Although the use of farred actinic light is non-intrusive, it does not give the CEF in white light which normally drives photosynthesis in nature. The redox state of the cytochrome bf complex In a supercomplex composed of PS I, LHCII, Cyt b6f, FNR and PGRL1 from C. reinhardtii cells, Cyt b was photoreduced in

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the presence of Fd, under anaerobic conditions and after a short illumination (indicated by an absorbance change in the range of 540–580 nm, Lam and Malkin 1982). This probably represents electron donation to haem bH from reduced ferredoxin via CEF (Iwai et al. 2010). However, Takahashi et al. (2013) argued that under anaerobic conditions, serious acceptor-side limitation promotes charge recombination within PS I in a short illumination; therefore, the photoreduction of Cyt b only reflects a different degree of charge recombination instead of CEF. Photoacoustic spectroscopy This technique measures the conversion of light energy to heat in an absorbing sample and hence the storage of light energy as chemical energy (energy storage, ES), under conditions in which the photobaric component is depressed, as described by Malkin and Canaani (1994). There are some pitfalls in this method. First, ES can only be measured provided the photobaric component is depressed, particularly for a leaf sample, by (1) a high-modulation frequency or (2) by the use of water infiltration (Malkin et al. 1992) or plastic wrap film without specific requirement of modulation frequency (Joe¨t et al. 2002). In the former case, ES depends on the modulation frequency because ES reflects energy stored in photochemical products that decay with a time constant larger than the modulation frequency of excitation (Malkin and Canaani 1994). Therefore, ES measured with a high-modulation frequency cannot detect fluxes at rates lower than the modulation frequency, which is likely the case when CEF occurs under steady-state illumination (Joe¨t et al. 2002). In the second case, water infiltration or plastic wrap film changes the CO2 and O2 concentrations inside leaf tissue, altering the redox poise and eventually affecting CEF. This method is further limited to the use of far-red actinic light, unless inhibitors are used to prevent the linear electron flux in white light (Canaani et al. 1989; Veeranjaneyulu et al. 1998). In this case the magnitude of CEF is altered due to inhibition of ETR2 and consequent changes in the redox poise, as previously discussed. The electrochromic signal The protonmotive force (pmf) that drives a proton efflux across the thylakoid membrane includes two components: DpH and Dw (the membrane potential difference). Quenching of 9-aminoacridine fluorescence can reflect the amplitude of DpH, but is not practical for in vivo measurements (Baker et al. 2007). The electrochromic shift (ECS), due to the presence of an electric field and resulting in a difference in light absorption of membrane pigments at a wavelength of 515–520 nm, can be used to monitor the

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formation of the protonmotive force (pmf) in leaves. Joliot and Joliot (2002) used post-illumination ECS kinetics to estimate cyclic electron flow through PS I upon inhibition of PS II by DCMU, but the CEF so obtained is different from that in the absence of DCMU because of altered redox poising. The amplitude of DpH can be applied to quantify the rate of electron transport indirectly. An inconvenience is that since the proton current obtained by the ECS method is in unit of s-1, calculating the CEF on a leaf area basis requires further information, such as the content of PS I reaction centres per unit leaf area. Further, the ECS signal overlaps with the 535 nm signal that depends on energydependent quenching (qE), and careful evaluation is needed for the application of the ECS (Johnson and Ruban 2014).

Category 2: Indirect methods of estimating CEF Some researchers have estimated CEF when linear electron flow occurs concurrently, using relative flux assays, taking advantage of the fact that ETR2, the rate of linear electron flux through PS II, should equal the linear electron flux through PS I (Klughammer and Schreiber 1994) or the cytochrome bf complex (Sacksteder and Kramer 2000). Therefore, a reliable measurement of both ETR2 and the total electron flux through PS I (ETR1) is a prerequisite for CEF estimation in this category of methods. Some studies took an increased ratio of the photochemical yields of PS I and PS II (Harbinson et al. 1989), of ETR1 and ETR2 (Kono et al. 2014), or of light-driven proton flux and ETR2 (Strand et al. 2015a), as evidence of CEF occurring. A quantitative measure of CEF could in principle be obtained from the difference between ETR1 and ETR2 and as such all of these methods fall into this category. ETR1 measurements Measurement of the electron flux through PS I (ETR1) via a Y(I)-based electron flux The photochemical yield of PS I, Y(I), in actinic light at a given irradiance can be obtained by the percentage of P700 that can be photo-oxidized with a saturating pulse superimposed on the actinic light; the percentage of photo-oxidizable P700 is calculated relative to the maximum P700? signal that can be obtained by a saturating single-turnover flash (or a saturating pulse) superimposed on weak continuous far-red light (Klughammer and Schreiber 2008). It is worth noting that a proper determination of the photochemical yield of PS I in actinic light requires strong farred light immediately before and during the application of a saturating light pulse (Siebke et al. 1997). ETR1, based

on Y(I), is then calculated as ETR1 = Y(I) 9 I 9 0.85 9 fI, where I is the irradiance, 0.85 is the assumed leaf absorptance and fI is the fraction (the estimation of which is described later) of the absorbed white light partitioned to PS I. This method rests on three key assumptions. First, the absolute quantum efficiency of reduced P700 is not known, although it is generally expected to be in the order of 0.95 (Lavergne and Trissl, 1995). It is therefore considered that in most cases taking the relative yield as the absolute yield is reasonable (Baker et al. 2007). Second, three PS I-related components (Pc?, P700? and Fd-) can contribute disproportionately to the 810 nm signal, depending on the reducing condition (Harbinson et al. 1989; Klughammer and Schreiber 1991; Kobayashi and Heber 1994; Schansker et al. 2003). Although current commercial instruments apply a dual-wavelength (810–870 nm) subtraction method to minimize the Pc contribution, it may not be sufficient, and additional deconvolution methods (such as ‘‘oxidative titration by far-red light’’, Oja et al. 2003) may be necessary. Third, although leaf absorptance could be measured with the aid of an integrating sphere, or estimated from the Chl content per leaf area (Evans and Poorter 2001), it is not easy to determine fI experimentally under all conditions. In some special cases such as low irradiance and/or in the presence of antimycin A, where CEF is assumed to be zero, ETR2 is supposed to be approximately equal to ETR1 and hence fI could be estimated (fI values are in the range 0.4–0.5, Kou et al. 2013, 2015). A variation of fI estimation is to measure Y(II) by Chl fluorescence and the oxygen evolution rate RO2 simultaneously, at low irradiance and high CO2 concentration. Assuming RO2 = IA 9 fII 9 Y(II)/4, where IA is the absorbed irradiance, the fraction of absorbed light partitioned to PS II fII can be obtained (Kono et al. 2014), and fI = 1 - fII. The validity of these two methods needs further evaluation; for example fI may vary according to experimental conditions. Furthermore, antimycin A is supposed to have multiple effects on photosynthesis, such as diminishing qE through either LHCII disaggregation or protonophoric activity (Horton et al. 1991). An alternative for the estimation of fI, through the use of photoacoustic spectrometry (Malkin and Canaani, 1994), is subject to the limitations relating to modulation frequency outlined earlier. Measurement of ETR1 via post-illumination P700? re-reduction Based on the rationale proposed by Maxwell and Biggins (1976), the initial rate of re-reduction of P700? should equal the electron flux to P700? immediately before cessation of illumination. The post-illumination re-reduction of P700? can be described as the sum of three sub-fluxes; the initial

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magnitude of each subflux equals the product of the corresponding rate coefficient and normalized amplitude, in unit of turnovers of each P700 per second (Jia et al. 2008). A potential error in measuring the rate of electron transport to P700? based on post-illumination P700 re-reduction is that the sub-millisecond kinetics of electron transport from the reduced Pc and Cyt b6f to P700? are ignored by current commercial instruments due to their poor time resolution (Kramer et al. 2004; Baker et al. 2007). It has been demonstrated that the initial rate of postillumination re-reduction of P700? is a non-linear indicator of flux through P700, if the equilibrium constant for sharing of electrons between the Pc/Cyt f pool and P700 is larger than one (Sacksteder and Kramer 2000). In consideration of this, Golding et al. (2005) suggested provided that P700 is more than about 20 % oxidized in the light, it could be possible to use the post-illumination kinetics of P700? to estimate the electron flux through PS I. However, we found that the Y(I)-based electron flux exceeded the initial post-illumination flux by a factor of about three, as shown in Fig. 1. This discrepancy, which existed regardless of a fast (closed circles) or a relatively slow (open circles) cessation of illumination, seems to be intrinsic rather than a result of limitation with data acquisition. How does the initial post-illumination rate of P700? re-reduction vastly underestimate ETR1? A plausible mechanism concerns a possible sudden change in the electric potential difference across the thylakoid membrane on cessation of illumination. In steady-state photosynthesis, a proton efflux from the lumen through the ATP synthase is balanced by (1) proton deposition in the lumen after water oxidation and (2) proton influx into the lumen at the secondary quinone acceptor (QB) in PS II and the Qi site (near the stromal surface) in the Cyt b6f complex (Hauska et al. 1996). At the instant the light is turned off, the proton deposition in and influx into the lumen may be stopped abruptly, while protons tend to continue to exit through the ATP synthase. The proton efflux must be rapidly compensated by the movement of counterions to avoid the lumen reaching a more negative electric potential (Vredenberg and Tonk 1975), but this charge compensation may not occur sufficiently rapid if the thylakoid membrane is not as permeable to the counterions as the activated ATP synthase is to protons. The sudden establishment of a more negative potential in the lumen may slow electron donation from Pc to P700? by impairing the docking of Pc with PS I. The N-terminal of the PsaF subunit of PS I contains a series of basic amino acids (Lys-12, Lys-16, Lys-19, Lys-23) that interact electrostatically with an acidic area (the ‘Southern patch’) on Pc (Hippler et al. 1996, 1997). Electron transfer from Pc to P700? takes place in a complex formed between Pc and the PsaF, involving both electrostatic and hydrophobic interactions. It is believed that formation of this complex and the subsequent conformational changes may be rate

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Fig. 1 A comparison of two ways of estimating the total electron flux through PS I (ETR1) in actinic light of irradiance 1300 lmol m-2 s-1. The Y(I)-based electron flux was obtained as described by Kou et al. (2013). The post-illumination initial flux was obtained by fitting the time course as the sum of three exponential decays and the initial slope was calculated by differentiation; the flux was converted into a value on a leaf area basis by multiplying it by 1.7 mmol P700 (mol Chl)-1 (Chow and Hope 1987) and by the Chl content, mol m-2. Cessation of illumination occurred fast (50 % decrease in irradiance in t‘ & 0.5 ls) or relatively slow (50 % decrease in irradiance in t‘ & 100 ls), using a different batch of plants in each case. The 1:1 correspondence of the two electron fluxes would follow the dashed line shown. Variation of the electron fluxes was effected by infiltration with various concentrations of DCMU

limiting for electron transfer (Sigfridsson et al. 1997). When the lumen electric potential becomes more negative in the vicinity of the basic amino acids of the PsaF subunit, the approach of net negatively charged Pc towards PS I may be hampered, impairing the formation of a Pc-PS I complex, as illustrated diagrammatically in Fig. 2. Therefore, it is likely that electron transfer from Pc to PS I is slowed considerably by a more negative electric potential in the lumen, that develops immediately at the cessation of illumination, hampering the use of the post-illumination kinetics to correctly quantify the on-going electron flux during illumination. We found that vacuum infiltration of leaf discs with 0.5 lM valinomycin hastened the initial post-illumination flux by 18 % in spite of a decrease in the Y(I)-based flux of 21 %. Higher concentrations of valinomycin decreased the Y(I)-based flux further, making it difficult to see any further increase in initial post-illumination fluxes. It is clear, however, that the half-time for relaxation from the minimum post-illumination electrochromic signal (ECS, partly indicating a more negative electric potential in the lumen)

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dark-interval relaxation kinetics (DIRK, Sacksteder and Kramer 2000) or the ‘dark pulse’ kinetics (Joliot and Joliot 2002) of the proton current immediately after cessation of illumination. In steady state, the sum of the photochemical rates (Rph) contributed by PS I (both cyclic and linear) and PS II (linear) could be deduced from the difference between the slopes of the ECS signal measured immediately before and after the light is switched off. If the quantum yield of PS II photochemistry is estimated from fluorescence, the CEF can be deduced from Rph (Joliot and Joliot 2002). However, fluorescence-based estimation of PS II photochemistry suffer from some drawbacks as discussed below. ETR2 measurements

Fig. 2 A schematic showing the concentration of plastocyanin (Pcn-) and the electric potential (w) as a function of distance from the lumenal surface of the thylakoid membrane. a. In steady-state illumination, w is less negative, and [Pcn-] is higher near the surface, favouring the docking of Pc with PS I. b. On abrupt cessation of illumination the H? diffusion potential, due to the proton gradient that has been established in the light, makes w much more negative, lowering [Pcn-] near the surface and impairing the docking of Pc with PS I

to a quasi-steady state in darkness was shortened by valinomycin. The half-time decreased from 5 s for the control to 2.3 s for leaf discs treated with 0.5 lM valinomcyin and to 1.2 s for 2 lM valinomycin, showing that dissipation of the H? diffusion potential was accelerated by the K?ionophore (Fan and Chow, unpublished data). The ECS signal A method to estimate ETR1 is to use the electrochromic signal in continuous actinic light of a particular spectral quality (reviewed by Bailleul et al. 2010) by monitoring the

Because the quantum yield of PS II photochemistry is directly related to the rate ETR2, it is possible in principle to estimate ETR2 through quantum yield of PS II photochemistry, using the following equation: ETR2 = Y(II) 9 I 9 0.85 9 0.5, where Y(II) is the quantum yield of PS II photochemistry, measured by Chl fluorescence as (1 Fs/Fm0 ), 0.85 is the assumed absorptance, I is the irradiance and 0.5 is the assumed fraction of absorbed light partitioned to PS II (fII). Fs and Fm0 are the steady-state and maximum Chl fluorescence yields, respectively, during illumination. Despite empirical tests (Genty et al. 1989; Siebke et al. 1997) that have demonstrated the robustness of the use of fluorescence to investigate ETR2 in the absence of photorespiration, there are some uncertainties for this method. First, it is difficult to determine fII experimentally, as discussed above. Second, the contribution of PS I fluorescence emission to Y(II) cannot be ignored (Pfundel 1998; Baker et al. 2007; Lazar 2013). Third, an inherent problem with using Chl fluorescence is that the detected signal originates from an unspecified depth in the leaf tissue and that depth may vary during the course of an experiment, for example, due to chloroplast movement. For another example, as functional PS II complexes at a shallow depth are rendered less fluorescent upon photoinactivation, the contribution to the Chl fluorescence yield from deeper tissue becomes more prominent (Oguchi et al. 2011). In fact, ETR2 based on chlorophyll fluorescence was only 72 % of that based on gross oxygen evolution when a spinach leaf was illuminated at irradiance 1460 lmol m-2 s-1 (Kou et al. 2013). A better way to obtain ETR2 based on whole-tissue measurement is the gross rate of oxygen evolution recorded by a gas-phase oxygen electrode. The ETR2 thus obtained can be validly compared with ETR1 obtained from Y(I). This is because the P700? signal is also a whole-tissue measurement, by virtue of the fact that the measuring beams at 820 and 870 nm are only weakly absorbed by the leaf tissue

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and are, therefore, multiply scattered in the tissue until they are finally absorbed; subtraction of ETR2 from ETR1 is then valid, as both refer to the same leaf issue. Since four electrons are needed for each oxygen molecule evolved, ETR2 equals four times the gross rate of oxygen evolution. A leaf disc in a sealed chamber, however, would exhaust the ambient CO2 quickly when illuminated. Thus, steady-state ETR2 can only be obtained in CO2-enriched air, which has the added advantage of inhibiting photorespiration that could otherwise complicate the measurement of oxygen evolution. However, gross rates of O2 evolution measured with a gas-phase oxygen electrode can only be estimated with this method by adding the post-illumination slope algebraically to the net rate in the light. The post-illumination slope is a little hard to estimate because its magnitude decreases over several minutes after cessation of illumination, particularly if there is a cooling effect after strong light is turned off (unpublished observation). Membrane inlet mass spectrometry, utilizing the stable 18O2 isotope to differentially and simultaneously measure rates of O2 uptake and evolution, provides a more precise method to accurately quantify ETR2 under nearnatural conditions (Beckmann et al. 2009). It also allows the [CO2] to be monitored in the cuvette to ensure photorespiration does not significantly contribute to the O2 uptake signal. The difference between ETR1 and ETR2 Scrutiny of the difference between ETR1 [measurement based on Y(I)] and ETR2 (measured by a gas-phase oxygen electrode) shows that it represents an overestimation of CEF, since contributions to ETR1 include not only CEF, but also electron fluxes associated with the Mehler reaction, electron donation from stromal reductants and any direct charge recombination in PS I. The Mehler reaction pathway is the electron flow from water via PS II, Cyt b6f and PS I to molecular oxygen. The role of the Mehler reaction as a major alternative electron sink under variable conditions remains controversial. Depending on species phylogeny, materials, experimental conditions and instruments used, the rate of the Mehler reaction can vary from a few percentage of ETR1 in some angiosperm plants (Ruuska et al. 2000; Vredenberg and Bulychev 2010; Driever and Baker 2011) to 50–100 % in algae and cyanobacteria (Badger et al. 2000; Asada 2000; Roberty et al. 2014). Shirao et al. (2013) confirmed that the magnitude of the Mehler reaction in vascular plants is small (\10 % of ETR2); therefore, the Mehler reaction may regulate CEF by adjusting redox poising instead of competing for electrons (Miyake 2010). A second role of the Mehler reaction may be as a partial compensation pathway when CEF is dysfunctional (Kono et al. 2014).

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Besides the Mehler reaction which produces reactive oxygen species (ROS), there is another route of O2 photoreduction by photosynthetic electron transfer, one that involves A-type flavodiiron proteins that reduce O2 directly to water (Helman et al. 2003). Since the activity of flavodiiron proteins is very high in cyanobateria and probably algae, whereas angiosperms do not conserve them (Peltier et al. 2010; Cardol et al. 2011), it is possible to explain why the rate of the Mehler reaction/O2 photoreduction can vary from a few percentage of ETR1 in some angiosperm plants (Ruuska et al. 2000; Vredenberg and Bulychev 2010; Driever and Baker 2011) to 50–100 % in algae and cyanobacteria (Badger et al. 2000; Asada 2000; Roberty et al. 2014). If the Mehler reaction occurs, the electron flux is counted as a contribution to ETR1. However, it does not affect ETR2 measurement when assayed by gross oxygen evolution in a gas-phase oxygen electrode because the product, superoxide, dismutates to produce H2O2, which, on further reaction, gives back oxygen. Similarly, activity of flavodiiron proteins, in producing water using electrons from water oxidation in PS II, will not affect the (NADPHforming) oxygen evolution measured with a Clarke-type electrode, but the electrons do pass through PS I and are, therefore, included in ERT1. Therefore, in either the Mehler reaction or the flavodiiron protein-catalyzed reaction, DFlux obtained using an oxygen electrode tends to overestimate CEF, even if electron donation from stromal reductants and direct charge recombination in PS I were negligible. On the other hand, some electrons may leak out of the electron transport chain before reaching PSI, via the plastid terminal oxidase (PTOX) (Bailey et al. 2008; Cardol et al. 2008; Laureau et al. 2013). This means that ETR1 does not include electrons from PS II that have been ‘‘hijacked’’, whereas ETR2 (assayed by gross oxygen evolution using an oxygen electrode) is unaffected because any superoxide formed via PTOX dismutates to form O2 and H2O2, thereby turning into O2 again. Therefore, DFlux tends to underestimate CEF due to the PTOX effect, which counteracts the overestimation associated with the Mehler reaction to some extent. Because all the fluxes including CEF are intertwined with one another and are hard to be quantified separately using an oxygen electrode, how other electron fluxes affect the quantification of CEF remains uncertain. Other pathways involving oxygen uptake include mitochondrial respiration and, in limiting CO2, photorespiration. These may affect the quantification of ETR2 by a traditional gas-phase oxygen electrode under physiological conditions. Through the utilization of MIMS and separation of 18O2 uptake fluxes from the 16O2 evolution flux from PS II, a more reliable quantification of ETR2 by gross oxygen evolution ETR2(O2) can be obtained. Results of

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simultaneous measurements of ETR2(O2) by MIMS and the Y(I)-based ETR1 in spinach discs are shown in Fig. 3. First, fI was estimated (Fig. 3a) utilizing 200 lM antimycin A to vacuum infiltrate spinach leaf discs. At irradiance \1000 lmol m-2 s-1, fI was close to 0.50, similar to the value of *0.47 obtained previously using an oxygen electrode and the P700? signal (Kou et al. 2013). There was an apparent decrease in fI to 0.40 at 2000 lmol m-2 s-1, but the reason for the apparent decrease in fI is not clear. In estimating fI, the assumption is that the linear electron flux (LEFO2) equals the total electron flux through PS I (=ETR1 = Y(I) 9 I 9 0.85 9 fI). This is true if antimycin A has inhibited CEF, the Mehler reaction is negligible and direct charge recombination in PS I is also negligible. Perhaps at high irradiance (2000 lmol m-2 s-1), there was some direct charge recombination, which keeps P700 more reduced, in which case Y(I) would be greater than in the absence of direct charge recombination. Therefore, the ratio fI = LEF(O2)/ [Y(I) 9 I 9 0.85] would be smaller. Second, using a value of fI = 0.50, we calculated ETR1 from Y(I) at each irradiance. At irradi-2 -1 ance \300 lmol m s , ETR1 &ETR2(O2), as found previously using an oxygen electrode and the P700? signal (Kou et al. 2013); that is, there seemed to be little or no CEF in glasshouse-grown spinach below this irradiance. DFlux increased steadily with irradiance above 300 lmol m-2 s-1, not saturating even at the highest irradiance used, as also found previously using an oxygen electrode and the P700? signal (Kou et al. 2013). The measurements conducted with MIMS appear to confirm previous measurements conducted with an oxygen electrode. Direct charge recombination in PS I due to both donorand acceptor-side limitations has been regarded as a futile photochemical loop internal to the photosystem I. In the absence of environmental stress, as electron donation to P700? and removal from the terminal acceptors are very efficient, charge recombination in PS I is expected to be small, particularly for plants that are acclimated to high light. However, if both donor and acceptor sides are severely limited, the fast charge recombination components (for example, 10–250 ls for an electron to return from A1 to P700?, Vassiliev et al. 1997) could occur and efficiently compete with other fluxes through PS I. Unfortunately, due to poor time resolution of current commercial instruments, the contribution of charge recombination in PS I can only be indirectly inferred from the extents of donor and acceptor limitation. The CEF estimated as the difference between ETR1 and ETR2 (DFlux) in the leaves of glasshouse-grown spinach,

(A)

(B)

Fig. 3 Simultaneously collected values for ETR1 utilizing Y(I) obtained by the P700? signal and ETR2(O2) based on gross oxygen fluxes measured with membrane inlet mass spectrometry. a The fraction of absorbed light partitioned to PS I (fI), calculated from measurements of spinach leaf discs (each 0.56 cm2) vacuum infiltrated with 200 lM antimycin A in water diluted from a stock solution in ethanol, and allowed to evaporate off excess inter-cellular water in darkness. Prior to measurement, each leaf disc was illuminated for 5 min in white light at an irradiance (I) of 250 lmol m-2 s-1. Y(I) was obtained as described by Kou et al. (2013). ETR1 was expressed as Y(I) 9 I 9 0.85 9 fI. The gross oxygen evolution rate, ETR2(O2), was obtained by adding the magnitude of the net 16O2 evolution to that of 18O2 uptake, corrected for changing concentrations of each gas in accordance with method described by Beckmann et al. (2009) at each irradiance, measured with a mass spectrometer. It was multiplied by 4 to convert the oxygen rate into a gross electron flux, ETR2(O2). By equating ETR2 with ETR1 in the presence of antimycin A, fI could be estimated. Each point is a mean of 3 replicates ± SE. b Steady-state ETR2 and ETR1, obtained by MIMS and the P700? signal, respectively, are plotted as a function of irradiance. ETR1 was calculated by assuming fI = 0.50 at all irradiances. The difference between them (DFlux) is shown as a function of irradiance. Each point is a mean of 5 replicates ± SE

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even when the above factors may be present, mainly consists of PGR5–PGRL1 protein-dependent CEF; that is, about 90 % of DFlux could be inhibited by antimycin A (Kou et al. 2013), showing that the chloroplast NADH dehydrogenase-like complex-dependent pathway is only minor in spinach. Indeed, in the pgr5 Arabidopsis mutant, the role of the chloroplast NDH-dependent pathway seems to be in compensation for the loss of the important PGR5 pathway to some extent (Munekage et al. 2004; Shikanai 2014). Antimycin A can be applied to separate the two CEF pathways (Shikanai 2014), although other electron fluxes should be borne in mind. For example, in the presence of antimycin A, low-light-grown Arabidopsis that lacks NDH still exhibited a substantial DFlux at high irradiance, attributable to charge recombination in PS I and/or the Mehler reaction (Kou et al. 2015).

Future prospects Here we have summarized current progress in methods of CEF quantification in chloroplasts of C3 leaves, both in vitro and in vivo, and have described the apparent advantages and disadvantages of current methods. We conclude that the quantification of CEF in C3 leaves is currently best approximated through measurements of ETR1 and ETR2 under identical conditions. In high-lightacclimated C3 plants the difference between ETR1 and ETR2 can be an upper estimate of CEF mainly consisting of PGR5–PGRL1-dependent CEF and a chloroplast NDHdependent CEF. These two CEF components can be separated by antimycin A. A more reliable estimation of ETR2 can be based on MIMS utilizing the stable 18O2 isotope, whereas ETR1 can be estimated from the Y(I)-based method. However, a number of unresolved issues remain for the recommended method: how to preclude the contribution of other components to the light-induced absorbance change in the range 800–850 nm besides that of P700?; how to obtain the fraction of absorbed light partitioned to photosystems; how to quantify charge recombination in PS I when both acceptor and donor sides are severely limiting; and finally, how to take account of the inclusion of the PTOX flux in ETR2 but not in ETR1. Methodological efforts to solve these questions should contribute greatly to CEF studies. A further challenge is to quantitatively estimate CEF in leaves of the three subtypes of C4 plants, which have increased requirements, to different extents, of ATP relative to NADPH, and hence of CEF. For example, the accumulation of NDH is [10-fold and that of PGR5/ PGRL1 is 3-fold as large in the NADP-malic enzyme subtype of Flaveria as in C3 Flaveria plants (Nakamura et al. 2013). Separating the CEF components in mesophyll

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cells and bundle sheath cells of C4 leaves will require great ingenuity. Acknowledgments The support of this work by an Australian Research Council Grant (DP1093827) awarded to W.S.C. and a Knowledge Innovation Program of the Chinese Academy of Sciences grant (KZCX2-XB3-09-02) and a grant of NNSF of China (No. 31370424) to D.Y. F. is gratefully acknowledged.

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