TY - JOUR
AU - Schreiber, Ulrich
AB - Abstract Non-photochemical chlorophyll fluorescence quenching (NPQ) plays a major role in the protection of the photosynthetic apparatus against damage by excess light, which is closely linked to the production of reactive oxygen species (ROS). The effect of a short heat treatment on NPQ and ROS production was studied with detached tobacco leaves by fluorescence imaging of chlorophyll and of the ROS sensor dye HO-1889NH. NPQ was stimulated >3-fold by 3 min pre-treatment at 44°C, in parallel with suppression of CO2 uptake, while no ROS formation could be detected. In contrast, after 3 min pre-treatment at 46°C, NPQ was suppressed and ROS formation was indicated by quenching of HO-1889NH fluorescence. After 3 min pre-treatment at 46°C and above, partial inactivation of ascorbate peroxidase and light-driven accumulation of H2O2 was also observed. These data are discussed as evidence for a decisive role of the Mehler ascorbate peroxidase or water–water cycle in the formation of the NPQ that reflects down-regulation of PSII. Introduction Chlorophyll fluorescence allows deep insights into the complex process of photosynthesis in vivo (see review in Papageorgiou and Govindjee 2004). In particular, pulse amplitude modulation (PAM) fluorometry and fluorescence quenching analysis by the saturation pulse method have proven useful for non-invasive assessment of the efficiency of photosynthetic energy conversion (Schreiber 2004). Measurements of non-photochemical quenching, mostly expressed by the parameter NPQ (Bilger and Björkman 1990), have allowed the mechanisms that protect plants from damage by excess light to be studied (for reviews, see, for example, Demmig-Adams and Adams 1992, Osmond 1994, Horton et al. 1996, Müller et al. 2001). There is general consensus that NPQ reflects the dissipation of excess excitation energy in the form of harmless heat (down-regulation of PSII), thus protecting the plant from the damaging effects of reactive oxygen species (ROS). The majority of studies on NPQ have been focused on the underlying molecular mechanisms, emphasizing the pivotal roles of the trans-thylakoid proton gradient (ΔpH) and zeaxanthin. Less attention has been paid to the question of what kind of electron flow is responsible for the formation of the ΔpH that drives violaxanthin de-epoxidation and renders zeaxanthin an efficient quencher, when CO2-dependent electron flow is limiting. This happens during the first minute of illumination after dark adaptation, when Calvin–Benson cycle enzymes are not yet light activated or in stressed leaves when these enzymes are damaged (Schreiber and Bilger 1987). Whenever CO2 is not available, molecular oxygen may substitute as electron acceptor, either in photorespiration (Osmond 1981) or in the Mehler ascorbate peroxidase (MAP) cycle (Nakano and Asada 1981, Asada and Badger 1984, Asada and Takahashi 1987). Electron flow associated with photorespiration is characterized by a higher ATP/NADPH ratio than CO2 fixation and, hence, does not qualify for NPQ generation. On the other hand, no ATP is consumed in the MAP cycle, so that in the intact system even low rates of this cycle can build up an appreciable ΔpH (Hormann et al. 1993, Hormann et al. 1994). Hence, it has been proposed that O2-dependent electron flow in the MAP cycle is mainly responsible for generation of the NPQ associated with the down-regulation of PSII (Schreiber and Neubauer 1990, Neubauer and Yamamoto 1992, Schreiber et al. 1995). One turnover of this cycle (transient formation of one H2O2) consumes 8 quanta and leads to the translocation of at least 16 H+ from the stroma to the lumen. As all H2O2 that is formed via O2 reduction is eventually re-reduced to H2O and as the electrons for the reduction of O2–· and H2O2 originate from the splitting of a stoichiometric amount of H2O in PSII, the name ‘water–water cycle’ was coined for this sequence of reactions (Asada 1999). A central role for the water–water cycle in generating NPQ in intact leaves has been questioned by Heber and co-workers (Wu et al. 1991, Kobayashi and Heber 1994, Heber 2002), who favor cyclic PSI electron flow instead, with the water–water cycle only serving a ‘poising’ function to relieve the intersystem electron transport chain from ‘over-reduction’. This view has been supported by Munekage et al. (2002) who explained the suppression of NPQ in a ‘proton gradient-deficient’ Arabidopsis mutant (PGR5) by a defect in the ferredoxin-dependent cyclic PSI pathway. These authors, however, explicitly do not rule out the possibility ‘that O2 reduction is mediated by the PGR5-dependent pathway’. The main reason why this important question is not clarified yet is the fact that to date no methods are available to quantify reliably the fluxes of the water–water cycle and cyclic PSI in illuminated intact leaves. Both reactions can be identified only indirectly, via the NPQ and the ‘scattering change’ associated with ΔpH formation. Recently we introduced a new PAM fluorescence imaging system, which allows parallel imaging of NPQ and ROS formation via fluorescence quenching of the ROS sensor dye HO-1889NH (Hideg and Schreiber 2007). In this previous study, the performance of the new system was demonstrated after leaf infiltration with methyl viologen in order to induce ROS formation upon illumination. In the present study, a mild, short heat treatment is applied for the same purpose, as a stress treatment that may also occur in the natural environment. Schreiber and Klughammer (2008) very recently showed that with increasing treatment temperatures NPQ was first substantially stimulated (at 44–46°C in outdoor-grown rose leaves) but then was strongly suppressed at only 2°C higher temperature. In this study, the heat-induced suppression of NPQ was assumed to be caused by an inhibition of the water–water cycle, while the heat-induced stimulation of NPQ was explained by inhibition of CO2 fixation (Bilger et al. 1987). If indeed both the Calvin–Benson cycle and the water–water cycle would be inactivated, the suppression of NPQ should be correlated with ROS formation. It is the aim of the present study to check on this hypothesis by parallel imaging of NPQ and fluorescence quenching of a ROS sensor. Results Figure 1 shows that while at up to 42°C there was only little inhibition, 3 min exposure to 44°C caused about 75% inhibition of CO2 uptake capacity. In leaves exposed to 46°C, CO2 uptake was completely blocked. Fig. 1 View largeDownload slide Photosynthetic carbon dioxide uptake in tobacco leaves pre-treated for 3 min at the indicated temperatures (n = 4). Fig. 1 View largeDownload slide Photosynthetic carbon dioxide uptake in tobacco leaves pre-treated for 3 min at the indicated temperatures (n = 4). With the same leaf material, chlorophyll fluorescence parameters were imaged after 3 min heat treatment (Figs. 2, 3). By exposing only half the areas of the leaves to the heat, any heat-induced change in photosynthetic parameters can be readily distinguished by comparison with the untreated half. Fig. 2 shows that, in agreement with the CO2 uptake data in Fig. 1, the 42°C pre-treatment did not have any effect on Fv/Fm and the photochemical yield [Y(II)], both of which displayed a very homogenous distribution over the imaged area. The non-photochemical quenching parameter, NPQ, showed some heterogeneity along the leaf veins, which was more pronounced in the untreated half. Distinct changes were induced by treatment at 44°C, which were particularly pronounced in Y(II) (large decrease) and NPQ (large increase), whereas Fv/Fm was just marginally decreased. It may be noted that heating not only affected the heated half of the leaf, but also spread to some extent into the untreated half. Exposure to 46°C resulted in further decreases of Fv/Fm and Y(II), and almost complete suppression of NPQ. In contrast to the leaf treated at 44°C, there is a sharp border between the untreated and treated halves of the leaf, which is characterized by very high NPQ (deep blue, equivalent to 1.8–1.9). As temperatures in this border area are uncertain, these areas were not included in data averaging. Fig. 2 View largeDownload slide Images of various chlorophyll fluorescence parameters of 32 × 24 mm center parts of detached tobacco leaves. The lower halves of the imaged areas were exposed to 3 min heat treatment at the indicated temperatures, while the upper parts were untreated. Images are color coded according to the patterns shown next to the images. The following parameters derived by the saturation pulse method are shown: A, Fv/Fm; B, Y(II) at 75 μmol m–2 s–1 PAR; C, NPQ at 75 μmol m–2 s–1 PAR. Fig. 2 View largeDownload slide Images of various chlorophyll fluorescence parameters of 32 × 24 mm center parts of detached tobacco leaves. The lower halves of the imaged areas were exposed to 3 min heat treatment at the indicated temperatures, while the upper parts were untreated. Images are color coded according to the patterns shown next to the images. The following parameters derived by the saturation pulse method are shown: A, Fv/Fm; B, Y(II) at 75 μmol m–2 s–1 PAR; C, NPQ at 75 μmol m–2 s–1 PAR. Fig. 3 View largeDownload slide Effect of heat treatment on dark–light induction kinetics of chlorophyll fluorescence parameters in tobacco leaves. (A) Effective photochemical quantum yield Y(II); (B) non-photochemical quenching NPQ. Illumination at 75 μmol m–2 s–1 PAR. Treatments and measurements are as described for Fig. 2 (n = 3). Fig. 3 View largeDownload slide Effect of heat treatment on dark–light induction kinetics of chlorophyll fluorescence parameters in tobacco leaves. (A) Effective photochemical quantum yield Y(II); (B) non-photochemical quenching NPQ. Illumination at 75 μmol m–2 s–1 PAR. Treatments and measurements are as described for Fig. 2 (n = 3). For a more detailed analysis, the dark–light induction kinetics of Y(II) and NPQ in terms of dependence on the pre-treatment temperature are presented in Fig. 3A and B, respectively. At the applied moderate light intensity of 75 μmol m–2 s–1 photosynthetically active radiation (PAR), in untreated control samples the kinetics reflect the activation of CO2 fixation in the Calvin–Benson cycle (Schreiber et al. 1986): during the first minute after onset of illumination, Y(II) decreases, whereas NPQ increases. Subsequently, Y(II) rises to a high level in the steady state, whereas NPQ drops to a low steady-state level. These transients have been interpreted as the ‘fingerprint’ of an energizing, non-assimilatory electron flow, which after dark adaptation activates Calvin–Benson cycle enzymes and thus primes CO2 fixation (Schreiber and Bilger 1987). This interpretation is supported by the present data. Both measurements of CO2 uptake and chlorophyll fluorescence parameters show a clearly defined critical threshold between 42 and 44°C treatment temperatures, below which CO2 uptake and fluorescence induction kinetics essentially are not affected and above which CO2 uptake is inhibited, while the characteristic increase of Y(II) and decrease of NPQ are suppressed. The physiological state after 3 min pre-treatment at 44°C is of special interest, because it is characterized by high NPQ in the light, which reflects down-regulation of PSII and thus protection of the plant from photoinhibitory damage (for reviews, see Demmig-Adams and Adams 1992, Horton et al. 1996, Müller et al. 2001). The question is, what kind of proton-coupled electron transport is responsible for this high NPQ (see Introduction and Discussion). In this context, it is important that treatment at just 2°C higher temperature (3 min at 46°C) causes complete suppression of NPQ. Hence, there is another clearly defined critical threshold between 44 and 46°C, which reflects the inactivation of the protective NPQ-generating reaction and thus should help to elucidate the identity of these reactions. If, as has been suggested before, the water–water cycle is responsible for generation of the protective NPQ (Schreiber and Neubauer 1990, Schreiber et al. 1995, Asada 1999), the abrupt suppression of NPQ by 3 min treatment at temperatures above 44°C should be due to the inactivation of one or several of the enzymes involved in this cycle, namely superoxide dismutase (SOD), ascorbate peroxidase (APX) and monodehydroascorbate reductase (MDAR), so that ROS can accumulate upon illumination. The following experiments were designed to check the validity of this hypothesis. ROS were detected on the basis of fluorescence quenching of a sensor probe (HO-1889NH), a solution of which was fed via pinholes into untreated and heat-pre-treated leaf halves, as illustrated in Fig. 4. For optimal differentiation between heated and untreated leaf halves, a relatively long illumination time (30 min) at relatively low intensity (25 μmol m–2 s–1) proved optimal. Fig. 4A and C shows fluorescence images before, and Fig. 4B and D after illumination. While generally no light-induced quenching was observed in the untreated leaf halves or in leaves treated at lower temperatures (Fig. 4A, B), HO-1889NH fluorescence decreased markedly in the leaf half exposed to 46°C (Fig. 4D), which clearly reflects formation of ROS. A quantitative analysis based on such images is presented in Fig. 5, which shows that the threshold temperature for ROS detection was between 44 and 46°C. Because HO-1889NH is not reactive to H2O2, this species was detected on the basis of observing brown coloring in leaves treated with DAB (diamino benzidine tetrahydrochloride). This was most pronounced in leaves pre-exposed to 48°C and only slightly noticeable in response to 46°C (Fig. 6). Fig. 4 View largeDownload slide Fluorescence images of 18 × 24 mm tobacco leaf segments infiltrated through two pinholes with a 2 mM solution of HO-1889NH after 3 min heat exposure to 38 and 46°C, as indicated in the figure. Images were taken before (A, C) and after (B, D) exposure to 25 μmol m–2 s–1 PAR for 30 min. Fluorescence intensities are color coded according to the pattern shown next to the images. Fig. 4 View largeDownload slide Fluorescence images of 18 × 24 mm tobacco leaf segments infiltrated through two pinholes with a 2 mM solution of HO-1889NH after 3 min heat exposure to 38 and 46°C, as indicated in the figure. Images were taken before (A, C) and after (B, D) exposure to 25 μmol m–2 s–1 PAR for 30 min. Fluorescence intensities are color coded according to the pattern shown next to the images. Fig. 7 demonstrates a strong temperature sensitivity of APX. Enzyme activity was decreased by approximately 30% even by the relatively short, 3 min heat treatments, if these were performed at 46°C or higher temperatures. This was due to a loss of function and not to any loss of APX protein, as illustrated by the protein gel immunoblot in Fig. 8. This experiment was carried out using heat-treated pea leaves, because commercially available antibodies were not reactive with tobacco APX. Discussion NPQ is induced whenever ΔpH formation exceeds the consumption of ΔpH in ATP synthesis for CO2 fixation (and photorespiration). In Fig. 1 it was demonstrated that CO2 uptake in tobacco leaves can be inhibited by a 3 min heat treatment at ≥44°C. The same treatment resulted in >3-fold stimulation of NPQ by 3 min heat treatment at 44°C and almost complete suppression of NPQ by 3 min heat treatment at 46°C (Figs. 2, 3). The same critical treatment temperature was also observed for the occurrence of light-induced fluorescence quenching of the ROS sensor dye HO-1889NH (Figs. 4, 5), for DAB coloration (Fig. 6) and for inactivation of APX (Fig. 7). This is the first direct demonstration of ROS formation in leaves under in vivo conditions. It shows that ROS are formed and can cause damage as soon as the NPQ-generating process is inhibited, which can be induced by heat inactivation of the APX. These findings suggest that the ROS-scavenging and NPQ-generating processes are closely linked. Hence, the presented data support the hypothesis that the MAP or water–water cycle is responsible for formation of the ΔpH that induces NPQ in leaves and thus provides 2-fold protection of the plant against damage by ROS and excess radiation (Asada 1999). Fig. 5 View largeDownload slide Effect of 3 min heat treatment at 38–48°C on relative intensity of HO-1889NH fluorescence in infiltrated tobacco leaves. Fluorescence quenching indicates light-induced ROS production. Fluorescence intensities were averaged from approximatelty 1 cm–2 areas of the HO-1889NH-infiltrated leaf segments (from four different images like the ones shown in Fig. 4) before and after exposure to 25 μmol m–2 s–1 PAR for 30 min. Changes in average intensities are shown as a percentage of mean fluorescence intensity before illumination. Error bars represent standard deviations. Values significantly (P < 0.05, in a paired t-test) different from the activity measured in untreated samples are marked with an asterisk. Fig. 5 View largeDownload slide Effect of 3 min heat treatment at 38–48°C on relative intensity of HO-1889NH fluorescence in infiltrated tobacco leaves. Fluorescence quenching indicates light-induced ROS production. Fluorescence intensities were averaged from approximatelty 1 cm–2 areas of the HO-1889NH-infiltrated leaf segments (from four different images like the ones shown in Fig. 4) before and after exposure to 25 μmol m–2 s–1 PAR for 30 min. Changes in average intensities are shown as a percentage of mean fluorescence intensity before illumination. Error bars represent standard deviations. Values significantly (P < 0.05, in a paired t-test) different from the activity measured in untreated samples are marked with an asterisk. Fig. 6 View largeDownload slide DAB staining for hydrogen peroxide in cuttings from tobacco leaves pre-exposed to heat for 3 min. Leaves were kept under 25 μmol m–2 s–1 PAR for 2 h in the DAB solution before chlorophyll was removed. Fig. 6 View largeDownload slide DAB staining for hydrogen peroxide in cuttings from tobacco leaves pre-exposed to heat for 3 min. Leaves were kept under 25 μmol m–2 s–1 PAR for 2 h in the DAB solution before chlorophyll was removed. Fig. 7 View largeDownload slide Activity of the ascorbate peroxide enzyme in extracts made from tobacco leaves after a 3 min exposure to heat as detailed in Materials and Methods. Symbols correspond to mean values, and error bars show standard deviations (n = 4). Values significantly (P < 0.05, in a paired t-test) different from the activity measured in untreated samples are marked with an asterisk. Fig. 7 View largeDownload slide Activity of the ascorbate peroxide enzyme in extracts made from tobacco leaves after a 3 min exposure to heat as detailed in Materials and Methods. Symbols correspond to mean values, and error bars show standard deviations (n = 4). Values significantly (P < 0.05, in a paired t-test) different from the activity measured in untreated samples are marked with an asterisk. Fig. 8 View largeDownload slide Protein gel blot analysis of ascorbate peroxidase content of heat-pre-treated pea leaves (see Materials and Methods for details). Fig. 8 View largeDownload slide Protein gel blot analysis of ascorbate peroxidase content of heat-pre-treated pea leaves (see Materials and Methods for details). A close link between NPQ formation and ROS scavenging has already been established. Previous work with intact and class D spinach chloroplasts has shown that for ΔpH and NPQ formation the presence of molecular oxygen (apparent Km of 60 μM) and ascorbate (apparent Km of 7 mM) is essential (Schreiber and Neubauer 1990, Schreiber et al. 1995). Furthermore, it was previously shown that it is the reduction of the H2O2 formed via the Mehler reaction and superoxide dismutation, involving enzymatic activity of APX and MDAR (Asada and Takahashi 1987, Asada 1999), which is mainly responsible for energy-dependent NPQ (Schreiber et al. 1991). In agreement with these findings, we now show that the suppression of room temperature NPQ by heat pre-treatment indeed occurs together with accumulation of H2O2 (Fig. 6) and a suppression of APX activity (Figs. 7, 8). Therefore, it appears likely that it is the heat inactivation of the chloroplast APX which causes the suppression of NPQ depicted in Figs. 2 and 3, the formation of ROS demonstrated in Figs. 4 and 5, and the accumulation of H2O2 shown in Fig. 6. Upon heating of leaves, electron donation to the intersystem electron transport chain in the dark is reportedly stimulated (Schreiber et al. 1976, Havaux 1996). The resulting acceleration of P700+ re-reduction following oxidation by far-red light and the NPQ generated by far-red illumination have been discussed as evidence for a role for cyclic PSI in the down-regulation of PSII reflected in NPQ (Bukhov et al. 1999). Hence, it could be argued that the stimulation of NPQ after 3 min pre-treatment at 44°C demonstrated in Figs. 2 and 3 is due to stimulation of cyclic PSI. Such a role for cyclic PSI electron flow in NPQ formation was also suggested by Heber and co-workers (Wu et al. 1991, Kobayashi and Heber 1994, Heber 2002). The following points, however, argue against this interpretation. (i) Far-red light-induced electron flow does not necessarily correspond to cyclic PSI, particularly if a sufficiently large electron pool at the donor side and molecular oxygen at the acceptor side are available. (ii) PSI is known to display relatively high heat stability and heat-induced acceleration of P700 re-reduction was still observed after 15 min at 51°C (Havaux 1996), whereas in our experiments the light-induced NPQ was almost completely suppressed after 3 min treatment at 46°C (Figs. 2, 3). (iii) Reising (1994) showed by simultaneous measurements of pulse-modulated fluorescence and photoacoustics of tobacco leaves that almost 30% of linear electron flow persisted after inactivation of CO2 fixation at 44°C. As this linear flux was not reflected by a net photobaric signal, the underlying O2 evolution must have been balanced by an equivalent O2 uptake. The latter was also reflected by strong stroma alkalization (Reising and Schreiber 1992). In agreement with the data of the present study, both linear flux and stroma alkalization were suppressed above 45°C. (iv) A recent domain model of Albertsson (2001) suggests that cyclic PSI is restricted to the stroma lamellae, whereas the PSI located in the margin region is mainly responsible for linear electron transport. To our knowledge, there is no evidence for a control of PSII, most of which is located in the stacked grana region, via the ΔpH formed in the stroma lamellae. The observed suppression of NPQ in heat pre-treated tobacco leaves strongly resembles that in the PRG5 mutant of Arabidopsis (Munekage et al. 2002). While Munekage et al. favored a role for PGR5 in cyclic electron flow around PSI, they did not exclude the possibility that the activity of the water–water cycle was also affected by the mutation and suggested that the PGR5 mutation may lead to pleiotropic effects on both cyclic PSI and the water–water cycle. They also showed that removal of O2 leads to strong limitation at the PSI acceptor side, which argues for a substantial O2-dependent electron flow, i.e. the reduction of O2 and monodehydroascorbate (MDA). Such electron flow can protect the photosynthetic apparatus in two ways: first, by relieving the PSI acceptor side, thus preventing PSI photoinhibition (Sonoike and Terashima 1994); and, secondly, by ΔpH/NPQ formation and the resulting down-regulation of PSII, with a direct photoprotective effect on PSII as well as an indirect photoprotective effect on PSI due to decreased ‘electron pressure’. In the pro/contra discussion of cyclic PSI vs. the water–water cycle it is often overlooked that cyclic PSI cannot relieve the PSI acceptor side as much as a reduction of O2 and MDA, because during the cyclic flow every electron taken up at the acceptor side is fed into the donor side, from where it is returned to the acceptor side again. The indirect effect of cyclic PSI via ΔpH/NPQ formation, however, is well known to depend on ‘poising’ by O2-dependent electron flow. This suggests that in any case cyclic PSI is functionally linked to the water–water cycle. The decisive question remains what are the relative fluxes of these two processes. For a regulatory function, the fluxes do not need to be high. Analogous heat treatment studies with Arabidopsis wild type and the PRG5 mutant may contribute to a better understanding of these processes. In conclusion, the presented data emphasize the 2-fold protective function of the water–water cycle, as previously outlined by Asada (1999): first, whenever O2 is reduced by PSI, the generated ROS is scavenged and in this way enzyme damage is prevented. Secondly, in the course of this scavenging process an exceptionally efficient electron acceptor, MDA, is formed, reduction of which by linear electron transport generates the NPQ that leads to down-regulation of PSII and thus to protection against damage by excess radiation. In the heat-pre-treated leaves, even partial inactivation of APX, a key component of the water–water cycle, leads to both ROS production and suppressed NPQ. The source of the detected ROS is probably heterogeneous, partly originating in the inefficient water–water cycle and partly in PSII, which is not down-regulated in the absence of NPQ. Materials and Methods Tobacco (Nicotiana tabaccum, L.) and garden pea (Pisum sativum) plants were grown in the greenhouse at 22–24°C with a natural photoperiod and daytime irradiation maxima around 220–250 μmol m–2 s–1 PAR. With the exception of protein immunoblots, all experiments were carried out using tobacco leaves. The youngest fully expanded leaves of 3-week-old tobacco plants were detached and one half of the leaf was immersed in water heated up to one of the following temperatures for 3 min: 38, 40, 42, 44, 46 or 48°C in dim (2–5 μmol m–2 s–1 PAR) green light. After this relatively mild heat treatment and drying of the surfaces, the leaves were used in photosynthesis or ROS measurements. Non-immersed halves of the leaves served as controls. Leaf stems were kept wet through all treatments and measurements. ROS were measured with a newly developed imaging apparatus capable of detecting changes in the fluorescence of the ROS sensor 3-(N-dansyl)aminomethyl-2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrole (HO-1889NH; Kálai et al 1998) as described earlier (Hideg and Schreiber 2007). Briefly, the infiltrating solution containing 2 mM HO-1889NH in a water : ethanol (99 : 1, v/v) solution was forced into the middle layer of leaf tissue using a plastic syringe without a needle, through a pinhole made with a sharp pin, as described earlier (Hideg et al 2002). Images of HO-1889NH fluorescence were taken before and after exposing the leaf to low intensity (25 μmol m–2 s–1) PAR for 30 min. PAR was provided from a KL-1500 lamp (DMP, Geneva, Switzerland) via fiberoptics. ROS production was assessed as lowering of HO-1889NH fluorescence during this period. Results are shown as color-coded images of the ROS sensor fluorescence. Fluorescence intensities were also averaged from areas of approximately 1–1.5 cm–2 where infiltration was relatively uniform. HO-1889NH trapping experiments were repeated three times. Hydrogen peroxide production was visualized with DAB staining (Thordal-Christensen et al. 1997). In these experiments, segments were cut from the leaves after the indicated heat pre-treatments, then floated on 1 mM DAB solution for 2 h under 25 μmol m–2 s–1 PAR. This was followed by removing chlorophyll from the leaf cuttings by ethanol (65°C, 2 h) to visualize brown staining characteristic of the presence of H2O2. These experiments served as illustrations only; H2O2 production was not quantified. Rates of CO2 assimilation in the leaves were determined by a portable infrared gas analyser (LI-6400, LI-COR Biosciences, Lincoln, NE, USA) using reference air with 400–450 μmol mol–1 CO2, corresponding to the growth conditions. To record light response curves, 6 cm2 areas of untreated or heat-pre-treated leaves were enclosed in the analyzer's leaf chamber and irradiated through a transparent window with a KL-1500 lamp (DMP). Incident PAR was measured using the gas analyzer's built-in sensor. PAR was increased from 0 to 1,000 μmol m–2 s–1, stepwise. After keeping the leaf at the chosen PAR level for 5 min, net photosynthesis rates were measured and expressed as rates of CO2 uptake (μmol CO2 m–2 s–1). Experiments were repeated four times, using four different leaves undergoing the same temperature exposure before measuring CO2 uptake. Values represent means of these repetitions with standard deviations. Chlorophyll fluorescence was imaged at room temperature, using the MINI-version of the Imaging-PAM (Heinz Walz GmbH, Effeltrich, Germany), with which areas up to 24 × 32 mm can be assessed. This instrument employs the same blue power light-emitting diodes (LEDs) for pulse modulated measuring light, continuous actinic illumination and saturation pulses. Fo, the minimal fluorescence yield of dark-acclimated samples, is imaged at low frequency of pulse-modulated measuring light, while images of the maximal fluorescence yield, Fm, are obtained with the help of a saturation pulse. Based on Fo and Fm, the images of maximal, dark-acclimated PSII quantum yield, Fv/Fm, are derived. With illuminated samples, the maximal fluorescence yield, Fm′, is non-photochemically quenched with respect to Fm (Schreiber et al. 1986). The effective PSII quantum yield of illuminated samples is calculated from the expression Y(II) = (Fm′ – F)/Fm′ (Genty et al. 1989). Non-photochemical quenching is quantified by the parameter NPQ = (Fm – Fm′)/Fm′ (Bilger and Björkman 1990) For assessment of heat-induced changes in photosynthesis, dark–light induction curves were recorded: first Fo and Fm images were measured with dark-acclimated samples, from which an Fv/Fm image was derived. This was followed by 3 min exposure at 75 μmol m–2 s–1 PAR, with repetitive measurements of Fo and Fm′ images every 20 s, from which automatically images of Y(II) and NPQ were calculated by the ImagingWin software. Kinetics of Y(II) decrease and NPQ build-up were constructed by averaging data from corresponding areas of interest in the images. The data points of these induction curves are averages of three repetitions using new heat-pre-treated leaves, with error bars corresponding to standard deviations. Results are also shown as color-coded images of Fv/Fm, Y(II) and NPQ after the 3 min illumination. Due to the non-invasive nature of PAM fluorescence measurements, APX could be extracted afterwards from the same leaves. Following heat treatment and fluorescence measurements, soluble proteins, including enzymes, were extracted from the tobacco leaves in ice-cold buffer (10 ml g–1 leaf FW) containing 50 mM potassium phosphate (pH 7.0), 1 mM EDTA, 50 mM NaCl, 1% polyvinylpyrrolidone and 1 mM ascorbate. Extracts were centrifuged at 10 000×g for 20 min at 4°C and supernatants were stored at −80°C until use. APX activity was measured spectrophotometrically, according to Nakano and Asada (1981), in 50 mM potassium phosphate buffer (pH 7.0) in the presence of 0.5 mM ascorbate and 10 mM H2O2. APX activity was calculated from the decrease in the 290 nm absorption of ascorbate, using 2.8 mM–1 cm–1 as the molar extinction coefficient. Measurements were corrected for the direct oxidation of ascorbate by H2O2, which was <10% of the enzyme-related absorbance change. APX activity data are given as mM H2O2 consumed per mg total protein per min; values are averages of four repetitions using new leaves, and error bars represent standard deviations. APX protein contents were determined using Western blots. Due to the reactivity of commercially available antibodies, this measurement was carried out using garden pea leaves. After applying the same heat treatments to whole pea leaves as to the halves of tobacco leaves, the former were weighed and homogenized in 1.5 ml reaction tubes with an equal amount of extraction buffer [50 mM Tris–HCl, pH 7.2, containing 1 mM phenylmethylsulfonyl fluoride (PMSF)]. The samples were centrifuged at 4°C for 45 min at 18,000×g to remove the insoluble fraction. The protein content of the supernatant was determined by Bradford assay using Bradford Reagent (Sigma-Aldrich Gmbh, Germany) and bovine serum albumin (BSA) as standard. From each sample, 15 μg of total soluble protein was separated by 12% SDS–PAGE, then transferred to a nitrocellulose membrane (G. Kisker GbR, Szeged, Hungary) and the APX content was probed with anti-APX antibody (Agrisera, Vännäs, Sweden). Funding The Hungarian Research Fund (grant Nos. OTKA T49438, OTKA-NKTH K67597). ACKNOWLEDGMENTS We thank Professors Tamás Kálai and Kálmán Hideg (Pécs University, Department of Organic and Medicinal Chemistry, Hungary) for the fluorescent ROS sensor HO-1889NH. REFERENCES Albertsson P.-A.  A quantitative model of the domain structure of the photosynthetic membrane,  Trends Plant Sci. ,  2001, vol.  6 (pg.  349- 354) Google Scholar CrossRef Search ADS PubMed  Asada K.  The water–water cycle in chloroplasts: scavenging of active oxygens and dissipation of excess photons,  Annu. Rev. Plant Physiol. Plant Mol. 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Acta ,  1991, vol.  104 (pg.  283- 291) Google Scholar CrossRef Search ADS   Abbreviations: Abbreviations: APX ascorbate peroxidase (EC 1.11.1.11) DAB diamino benzidine tetrahydrochloride Fm maximum fluorescence yield in the dark-acclimated state Fo minimum fluorescence yield in the dark-acclimated state Fm′ maximum fluorescence yield in the light-acclimated state Fv/Fm maximum photochemical yield of PSII in the dark-acclimated state HO-1889NH 3-(N-dansyl)aminomethyl-2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrole LED light-emitting diode MAP cycle Mehler ascorbate peroxidase cycle MDA monodehydroascorbate MDAR monodehydroascorbate reductase NPQ non-photochemical quenching PAM pulse amplitude modulation PAR photosynthetically active radiation ROS reactive oxygen species Y(II) photochemical yield © The Author 2008. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. 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TI - Imaging of NPQ and ROS Formation in Tobacco Leaves: Heat Inactivation of the Water–Water Cycle Prevents Down-Regulation of PSII
JF - Plant and Cell Physiology
DO - 10.1093/pcp/pcn170
DA - 2008-12-03
UR - https://www.deepdyve.com/lp/oxford-university-press/imaging-of-npq-and-ros-formation-in-tobacco-leaves-heat-inactivation-ynClRTecdu
SP - 1879
EP - 1886
VL - 49
IS - 12
DP - DeepDyve
ER -