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First published online December 30, 2005; 10.1105/tpc.105.035808 The Plant Cell 18:457-464 (2006) © 2006 American Society of Plant Biologists Mobilization of Photosystem II Induced by Intense Red Light in the Cyanobacterium Synechococcus sp PCC7942School of Biological Sciences, Queen Mary, University of London, London E1 4NS, United Kingdom 1 To whom correspondence should be addressed. E-mail c.mullineaux{at}qmul.ac.uk; fax 44-20-8983-0973.
We use confocal fluorescence microscopy and fluorescence recovery after photobleaching to show that a specific light signal controls the diffusion of a protein complex in thylakoid membranes of the cyanobacterium Synechococcus sp PCC7942 in vivo. In low light, photosystem II appears completely immobile in the membrane. However, exposure to intense red light triggers rapid diffusion of up to  50% of photosystem II reaction centers. Particularly intense or prolonged red light exposure also leads to the redistribution of photosystem II to specific zones within the thylakoid membranes. The mobilization does not result from photodamage but is triggered by a specific red light signal. We show that mobilization of photosystem II is required for the rapid initiation of recovery from photoinhibition. Thus, intense red light triggers a switch from a static to a dynamic configuration of thylakoid membrane protein complexes, and this facilitates the rapid turnover and repair of the complexes. The localized concentrations of photosystem II seen after red light treatment may correspond to specific zones where the repair cycle is active.
The photosystem II (PSII) reaction centers of plants and cyanobacteria are subject to photodamage (Barber and Andersson, 1992  ). A complex repair cycle operates that involves the partial disassembly of damaged reaction centers, the proteolysis of damaged subunits, and their replacement by newly synthesized subunits (Andersson and Aro, 2001). This repair cycle is of crucial importance for photosynthesis. It is also a beautiful experimental model for the turnover of protein complexes, because damage and turnover can be initiated simply and specifically by exposure to light. For these reasons, the repair cycle has been intensively studied. A key enzyme involved in the early stages of the PSII repair cycle in plants and cyanobacteria was recently identified (Bailey et al., 2002; Silva et al., 2003). However, many questions remain concerning the ways in which the activity of the repair cycle is controlled and the possible localization of different stages of the repair cycle. In green plants, biochemical fractionation studies suggest that key stages of the repair cycle occur in the stroma lamellae and that the migration of photodamaged PSII complexes from the grana to the stroma lamellae plays an important role (Baena-Gonzalez et al., 1999; Andersson and Aro, 2001).
PSII core complexes are naturally fluorescent, raising the possibility that redistribution of PSII complexes during photoinhibition could be observed in vivo, in real time, using fluorescence microscopy. However, the intricate structure of green plant thylakoid membranes makes such studies difficult (Mullineaux, 2004 Here, we show that PSII in Synechococcus 7942 is not always immobile. After exposure of cells to intense red light, a proportion of PSII centers start to diffuse quite rapidly in the thylakoid membrane. Under some conditions, this leads to a redistribution of PSII within the thylakoid membrane system. We present data suggesting that this process facilitates the rapid initiation of the PSII repair cycle. Our results indicate a novel signal transduction process controlling the dynamics of the thylakoid membrane.
The diffusion of PSII complexes in cells of Synechococcus 7942 may be visualized using confocal FRAP with a blue excitation light that is mainly absorbed by chlorophyll a combined with detection of chlorophyll fluorescence in the red region of the spectrum (Sarcina and Mullineaux, 2004 ). We use the 457-nm band from an argon laser as the blue excitation light and detect emission wavelengths >665 nm (Sarcina and Mullineaux, 2004). Figure 1 shows a fluorescence emission spectrum for cells of Synechococcus 7942 excited at 457 nm. For comparison, a spectrum with excitation at 600 nm (predominantly absorbed by phycocyanin) is shown as well (Figure 1). Spectra were recorded in the presence of DCMU to close PSII reaction centers. This helps to replicate conditions during the confocal measurements, in which the very high intensity of illumination will cause the closure of PSII reaction centers, leading to increased PSII fluorescence. The 680- to 685-nm peak may come either from chlorophyll or from long-wavelength pigments in the phycobilisome core, whereas the 650-nm peak comes only from the phycobilisomes: it originates from phycocyanin in the phycobilisome rods (Sidler, 1994). We used a red glass filter to select longer-wavelength fluorescence (Figure 1). The presence of the 650-nm peak with 600-nm excitation (Figure 1) is indicative of strong excitation of the phycobilisomes. The absence of this peak with 457-nm excitation (Figure 1) indicates that there is little or no excitation of phycobilisomes at this wavelength; thus, the fluorescence we see must all come essentially from chlorophyll.
Chlorophyll fluorescence could come either from PSII or from photosystem I (PSI). PSI fluorescence emission is generally low except at low temperatures, as a result of the very rapid decay of fluorescence in PSI (Turconi et al., 1993 ). However, PSI fluorescence emission is not completely insignificant in cyanobacteria, in which PSI has a larger chlorophyll light-harvesting antenna and the PSI/PSII ratio is relatively high, typically 3 in Synechococcus 7942 (Joshua and Mullineaux, 2005). Picosecond fluorescence decay measurements give accurate resolution of PSII and PSI fluorescence emission, allowing an estimate of the relative contribution of PSII and PSI fluorescence. Lifetime data with chlorophyll excitation and closed PSII reaction centers indicate that under these conditions, the ratio of PSII to PSI fluorescence emission is 4; thus, 80% of the fluorescence observed comes from PSII and 20% comes from PSI (Mullineaux and Holzwarth, 1993). Therefore, to a first approximation, FRAP measurements under our conditions report on the mobility of PSII, although a minor contribution from PSI is also likely.
For the FRAP measurement, cells are immobilized on an agar surface. The confocal laser spot is used to bleach a line across the center of the cell. Diffusion of fluorescent membrane components may then be visualized by repeatedly imaging the cell and observing the spread and recovery of the bleached line (Mullineaux and Sarcina, 2002
In the PSII diffusion measurements reported previously, the cells were exposed only to blue light at 457 nm (Sarcina and Mullineaux, 2004
The mobile fraction of chlorophyll fluorescence depends on the dose of red light (Figure 4). After a 3-s exposure to red light, 20% of chlorophyll fluorescence is mobile, and the effect saturates within 15 s. The maximum proportion of PSII that becomes mobile is 50 to 60%, and longer exposure does not lead to any further increase in the mobile fraction (Figure 4). After comparable pretreatments with blue light (457 nm) and green light (543 nm), with similar exposure times and photon flux densities, there is no detectable PSII diffusion. Figure 5 shows images from a typical FRAP sequence for a cell pretreated with 457-nm light (exposure time of 3 min, average photon flux density of 8.8 x 105 µE·m–2·s–1).
To determine whether the effect could be replicated under more physiological conditions, we exposed cells in bulk liquid culture to bright red light. In this case, cells were incubated in growth medium at 30°C and exposed to broad-band red light (>620 nm) at a photon flux density of 2000 µE·m–2·s–1. Aliquots were then immobilized on agar, and FRAP measurements were performed as before. After preillumination for 30 min, an average of 10% ± 3% of PSII complexes were mobile, although the proportion of mobile complexes is likely to have decreased during the delay required for the preparation of the microscope sample. The mobility of PSII ceased within 30 min of returning the cells to normal growth conditions after red light illumination. A comparable pretreatment with broad-band blue-green light (<560 nm) did not induce any PSII mobility.
In green plants, photodamage has been proposed as the trigger for a series of events leading to the migration of damaged PSII centers from the grana to the stroma lamellae (Baena-Gonzalez et al., 1999
Treatment with the 633-nm confocal laser spot not only mobilizes chlorophyll fluorescence but also causes it to rapidly redistribute within the cell (Figure 7). Chlorophyll fluorescence is initially distributed very evenly along the length of the cell (Figure 7A). However, after red light treatment, it becomes concentrated in localized zones within the thylakoid membranes (Figure 7B). In small cells, these are usually at the poles of the cell, whereas longer cells have additional concentrations throughout the length of the cell (Figure 7B). This effect could easily be induced with short, very intense red light treatments with the confocal microscope. However, when cells in liquid culture were exposed to red light (of much lower intensity), it could be induced only by very prolonged treatment. Exposure of a liquid culture to red light at 1200 µE·m–2·s–1 for 3 h was required to induce localized concentrations of chlorophyll fluorescence (data not shown). Localized concentrations of chlorophyll fluorescence could not be induced by 457-nm light treatment with the confocal microscope (data not shown).
The wavelength specificity of the red light effect allows us to test the possibility that the mobilization of chlorophyll complexes plays a role in the response of the cells to photoinhibitory conditions. Figure 8 compares oxygen evolution during photoinhibitory treatments with red or blue-green light. No electron acceptors were added; therefore, oxygen evolution reports on the activity of the entire photosynthetic electron transport chain. Lincomycin inhibits de novo protein synthesis and therefore blocks the PSII repair cycle. The response of the cells to photoinhibitory treatment may be gauged by comparing oxygen evolution during photoinhibitory light treatments in the presence and absence of lincomycin. Higher oxygen evolution in the absence of lincomycin indicates an active PSII repair cycle (Silva et al., 2003 ). In the presence of lincomycin, oxygen evolution declines to zero in 50 min in both blue-green light (Figure 8A) and red light (Figure 8B). Thus, both light treatments are effective at inducing photodamage. In blue light in the absence of lincomycin, oxygen evolution initially declines almost as fast as when lincomycin is present. Thus, there is initially little activity of the repair cycle, although it becomes significant after 20 to 30 min. In red light in the absence of lincomycin, there is initially a significant increase in oxygen evolution. This is not seen when lincomycin is present, indicating an adaptation process requiring protein synthesis. Oxygen evolution eventually declines during the photoinhibitory treatment, but it always remains significantly higher than in the presence of lincomycin, indicating that repair and adaptation processes operate efficiently from the start under these conditions (Figure 8B).
The difference between the response of the cells to photoinhibition under red and blue light is confirmed by t tests performed for the data shown in Figure 8 at 15 min after the onset of photoinhibition. In the presence of lincomycin, oxygen evolution declines on average to 80% ± 4% in blue light and to 84% ± 3% in red light. This difference is not significant (P = 0.53), showing that blue and red light are equally effective at inducing photoinhibitory damage. In the absence of lincomycin, mean oxygen evolution is 86% ± 3% in blue light and 108% ± 2% in red light. Oxygen evolution under red light is significantly higher (P = 3 x 10–6). The presence of lincomycin makes no significant difference under blue light (P = 0.27), but oxygen evolution is significantly lower when lincomycin is present under red light (P = 10–6).
A proportion of chlorophyll fluorescence in the cyanobacterium Synechococcus 7942 begins to diffuse after exposure of the cells to intense red light (Figure 2). Depending on the dose of red light, up to 50 to 60% of chlorophyll fluorescence becomes mobile under these conditions (Figure 4). The majority of chlorophyll fluorescence ( 80%) comes from PSII. Therefore, PSII reaction centers must be diffusing under these conditions, although it is possible that a part of the diffusion that we see comes from PSI. Prolonged exposure to intense red light also leads to a very clear redistribution of chlorophyll complexes within the thylakoid membrane system. Under normal conditions, chlorophyll fluorescence is rather evenly distributed within the thylakoid membranes (Figure 7A), but after exposure to intense red light, there are obvious concentrations at the poles of the cell and at other points in long cells (Figure 7B).
The diffusion of chlorophyll complexes is triggered by red light (633 nm) but not by comparable doses of blue light (457 nm) (Figure 5) or green light (543 nm) (data not shown). The wavelength specificity of the effect suggests that it is caused by light perception and signal transduction rather than being a simple consequence of photodamage. This is confirmed by an experiment showing that a red light signal at one end of the cell triggers diffusion at the other end (Figure 6), indicating that the reaction centers that become mobile are not simply those that were photodamaged by the light pretreatment. There are two possible explanations for this. (1) The red light may induce a redox signal, or a specific kind of photodamage, as a result of its preferential absorption by the phycobilisomes. This could then trigger a signal transduction pathway leading to the mobilization of reaction centers. (2) The red light may be perceived by a specific photoreceptor, initiating a signal transduction pathway leading to the mobilization of PSII throughout the cell. The nature of the possible photoreceptor is unknown, but it could plausibly be a phytochrome. Phytochromes are generally considered to be low-irradiance sensors, but there are indications that plant phytochromes can also act as high-irradiance sensors of red light (Shinomura et al., 2000 Mobilization of chlorophyll complexes is induced only by light at intensities considerably greater than those required to saturate photosynthesis. This suggests that the physiological role of the effect is somehow to minimize photoinhibition. There are two obvious possibilities for this. (1) Mobilization of chlorophyll complexes allows redistribution into localized concentrations within the cell (Figure 7). When this occurs, overall light absorption will be decreased and thus photodamage could be reduced. (2) Mobilization of PSII could facilitate the PSII repair cycle, allowing the rapid turnover and repair of photodamaged PSII. Other adaptation and repair processes could also be facilitated by switching the thylakoid membrane to a more dynamic state, allowing the diffusion of protein complexes. Possibility 1 seems unlikely because concentrations of PSII are formed rapidly only after treatment with photon flux densities of red light far greater than anything likely to be encountered in nature. At more physiologically realistic light intensities, they are seen only after very prolonged high-light treatment, which has already caused major photodamage. Therefore, this cannot be an efficient mechanism for minimizing photodamage by reducing light absorption. Thus, we favor possibility 2, that the mobilization of PSII facilitates the PSII repair cycle. The wavelength specificity of the effect allows us to test this possibility. PSII becomes mobile when cells are exposed to photoinhibitory doses of red light but not blue or green light. Therefore, we predict that the response of the cells to photoinhibition should be more effective during exposure to red light than to blue light. We find that this is the case (Figure 8). As a result, we suggest that the mobilization of PSII is required to allow rapid initiation of the repair cycle and possibly other responses involving de novo protein synthesis. However, it is clear that the PSII repair cycle does eventually become active under blue light when no PSII diffusion is induced (Figure 8). Further studies will be required to determine whether this represents a slower induction of essentially the same repair process or perhaps an alternative mechanism requiring the de novo synthesis of repair enzymes. The data shown in Figure 8 indicate that the rapid initiation of repair significantly enhances photosynthetic performance during short-term high-light treatments.
In green plants, the phosphorylation of PSII core proteins may facilitate the diffusion of photodamaged PSII complexes from the grana to the stroma lamellae (Baena-Gonzalez et al., 1999 A requirement of PSII mobility for rapid PSII repair suggests that photodamaged PSII complexes are not readily repaired in situ but must instead migrate to specialized repair zones, where the necessary enzymes are concentrated. The localized concentrations of PSII seen after brief, very intense red light treatment (Figure 7) may correspond to such repair zones. We propose that the red light signal leads to the mobilization of a proportion of PSII centers and that this allows photodamaged centers to diffuse to repair zones. When photodamage is faster than repair, PSII will accumulate in the repair zones. Concentrations of PSII are not observed during less intense red light photoinhibitory treatments, such as those used in Figure 8B, although PSII is mobilized under these conditions. However, under these conditions, there is little net photoinhibition, indicating that PSII repair keeps pace with PSII photodamage (Figure 8B). Therefore, we would not expect PSII to accumulate in the repair zones. We observe the concentrations of PSII only after extremely intense red light treatments or after very prolonged treatments at more physiologically realistic light intensities. In the first case, the very intense red light treatment will cause the rapid production of photodamaged PSII, overwhelming the cell's capacity for repair. In the second case, the prolonged high-light treatment compromises the cell's capacity for PSII repair, so that we observe very sustained photoinhibition under these conditions (data not shown). Under both of these extreme conditions, damaged PSII centers may accumulate in the repair zones faster than they can be turned over, allowing us to visualize the repair zones as concentrations of PSII.
Cell fractionation studies suggest that the early stages of PSII biosynthesis in cyanobacteria may occur in the plasma membrane rather than in the thylakoids (Zak et al., 2001
Intense red light treatment leads to the mobilization of up to
Cell Growth and Sample Preparation The cyanobacterium Synechococcus sp PCC7942 was obtained from the Pasteur Culture Collection and grown in liquid cultures in an illuminated orbital shaking incubator in BG11 medium (Castenholz, 1988 ) supplemented with 10 mM NaHCO3. Growth was at 30°C under white light at 10 µE·m–2·s–1. Before FRAP measurements, cells were grown for 16 h in the presence of 0.5% DMSO to induce cell elongation (Aspinwall et al., 2004). For measurements with the confocal microscope, drops of cell culture were adsorbed onto an agar surface (1.5% Difco Bacto-agar containing BG11 medium). Blocks of agar were placed in the well of a laboratory-built, temperature-controlled sample holder and covered with glass cover slips. All measurements were performed at 30°C.
Fluorescence Spectroscopy
Confocal Microscopy and FRAP
Photoinhibition and Oxygen Evolution
We thank P.J. Nixon and E. Lopez-Juez for discussion and A. Casal and S. Garcia for technical support. This work was supported by Biotechnology and Biological Science Research Council and Wellcome Trust grants to C.W.M.
The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) is: Conrad W. Mullineaux (c.mullineaux{at}qmul.ac.uk). Article, publication date, and citation information can be found at www.plantcell.org/cgi/doi/10.1105/tpc.105.035808. Received July 8, 2005; Revision received November 7, 2005. accepted December 13, 2005.
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