Live-cell imaging of photosystem II antenna dissociation ... - PNAS

0 downloads 0 Views 1MB Size Report
Feb 2, 2010 - and spreading of, a 250-ps lifetime chlorophyll fluorescence compo- ... The 250-ps component was also the dominant compo- ..... 267–275.
Live-cell imaging of photosystem II antenna dissociation during state transitions Masakazu Iwaia,1, Makio Yokonoa,2, Noriko Inadab, and Jun Minagawaa,3 a Institute of Low Temperature Science, Hokkaido University, N19 W8, Sapporo, Hokkaido 060-0819, Japan; and bGraduate School of Biological Sciences, Nara Institute of Science and Technology, 8916-5 Takayama-cho, Ikoma, Nara 630-0192, Japan

Edited by Graham R. Fleming, University of California, Berkeley, CA, and approved December 1, 2009 (received for review August 4, 2009)

|

photosynthesis fluorescence lifetime imaging microscopy light-harvesting energy dissipation

|

I

| green algae |

n the thylakoid membranes of chloroplasts, photosystems I and II (PSI and PSII) work in concert to carry out photosynthetic electron transfer from water to NADP+. For efficient photosynthesis under changing light conditions, balancing absorbed light energy between the two photosystems, or state transition, is important (1–3). The plastoquinone (PQ) pool, an intersystem electron carrier, monitors changes in light quality and quantity and activates a protein kinase for light-harvesting complex (LHC) II (3–5). LHCII phosphorylation leads to a decrease in PSII light absorption (6, 7), which occurs on a timescale of minutes without any changes in gene expression. The decrease is therefore considered to be due to the reorganization of the PSII antenna system, such as dissociation of phospho-LHCII from PSII. The fate of dissociated phospho-LHCII has been investigated in vitro. Subfractionation of phosphorylated thylakoid membranes showed that the stroma lamella (non-appressed) fraction, where PSI was predominantly located, contained more phospho-LHCII than the grana (appressed) fraction, where PSII was predominantly located (8–10). Isolation of PSII under PQ pool-oxidizing conditions provided the evidence that unphospho-LHCII remained bound to PSII, whereas under PQ pool-reducing conditions phospho-LHCII was dissociated from PSII (11). Isolation of PSI under the same conditions revealed that the dissociated LHCII (including both major and minor LHCII) were reassociated with PSI (12). Thus, state transitions are accompanied by LHCII relocation between the two photosystems. However, because such LHCII migrations have been observed only in vitro, it still remains to be uncovered what happens during the LHCII migration in vivo. Does LHCII really dissociate from PSII in live cells? Does the undocked phosphoLHCII simply migrate from PSII to PSI? What if the unbound phospho-LHCII absorbs light energy during the migration? Here, to address these questions directly, we show live-cell imaging of LHCII dissociation during state transitions in the www.pnas.org/cgi/doi/10.1073/pnas.0908808107

green alga Chlamydomonas reinhardtii by using fluorescence lifetime imaging microscopy (FLIM). Upon preferential excitation of PSII, we watched a shift in a chlorophyll fluorescence lifetime (CFL) component in the C. reinhardtii cells which was ascribed to the phospho-LHCII dissociation from PSII. Surprisingly, the dissociated phospho-LHCII was found to form energy-dissipative aggregations, which is in fact advantageous when they are in transit between the two photosystems during state transitions. These results are important because such energy-dissipative unbound LHCII could also be involved in other photoacclimation modes including nonphotochemical quenching under high-light stress. The apparently distinct photoacclimation modes thus may share an underlying mechanism. Results Monitoring Chlorophyll Fluorescence Lifetime During State Transitions. State transitions can be readily induced in live plant

and algal cells by alternately providing PSI light (720 ± 10 nm), which preferentially excites PSI, and PSII light (467 ± 10 nm), which preferentially excites PSII. When the cells are exposed to PSI light for 15 min, most of the LHCII remains unphosphorylated and associated with PSII in the PSII-LHCII supercomplex (11). That physiological condition, which increases the excitation level at PSII, is called state 1. When cells are transferred to PSII light for 5 min, the PSI fluorescence at 77K (∼718 nm) relatively increases as compared with state 1; that physiological condition is called state 2 (Fig. S1). Using this induction method, we attempted to visualize phospho-LHCII dissociation from PSII during the transition from state 1 to state 2 (state 2 transition) in live C. reinhardtii cells. To spatiotemporally differentiate between the PSII-LHCII supercomplex (before phospho-LHCII dissociation) and dissociated phospho-LHCII, we measured the fluorescence lifetime of the chlorophyll molecules in both PSII and LHCII using FLIM. Given that phospho-LHCII dissociation modulates excitation energy transfer pathways among the chlorophylls in PSII and LHCII, the lifetimes of the fluorescence originating from these complexes are expected to change. Using wild-type (WT) cells in state 1 and state 2, we excited chlorophyll at 405 nm and counted the photons emitted at 665–685 nm, which originate predominantly from PSII and LHCII at room temperature (13, 14) (see Fig. S2A for the experimental scheme). The dominant chlorophyll

Author contributions: M.I. and J.M. designed research; M.I. and N.I. performed research; M.I., N.I., and J.M. contributed new reagents/analytic tools; M.I., M.Y., and J.M. analyzed data; and M.I. and J.M. wrote the paper. The authors declare no conflict of interest. This article is a PNAS Direct Submission. Freely available online through the PNAS open access option. 1

Present address: Real-Time Bio-Imaging Research Team, Extreme Photonics Research Group, RIKEN Advanced Science Institute, 2-1 Hirosawa, Wako, Saitama 351-0198, Japan.

2

Present address: Molecular Photoscience Research Center, Kobe University, 1-1 Rokkodaicho, Nada, Kobe, Hyogo 657-8501, Japan.

3

To whom correspondence should be addressed. E-mail: [email protected].

This article contains supporting information online at www.pnas.org/cgi/content/full/ 0908808107/DCSupplemental.

PNAS | February 2, 2010 | vol. 107 | no. 5 | 2337–2342

PLANT BIOLOGY

Plants and green algae maintain efficient photosynthesis under changing light environments by adjusting their light-harvesting capacity. It has been suggested that energy redistribution is brought about by shuttling the light-harvesting antenna complex II (LHCII) between photosystem II (PSII) and photosystem I (PSI) (state transitions), but such molecular remodeling has never been demonstrated in vivo. Here, using chlorophyll fluorescence lifetime imaging microscopy, we visualized phospho-LHCII dissociation from PSII in live cells of the green alga Chlamydomonas reinhardtii. Induction of energy redistribution in wild-type cells led to an increase in, and spreading of, a 250-ps lifetime chlorophyll fluorescence component, which was not observed in the stt7 mutant incapable of state transitions. The 250-ps component was also the dominant component in a mutant containing the light-harvesting antenna complexes but no photosystems. The appearance of the 250-ps component was accompanied by activation of LHCII phosphorylation, supporting the visualization of phospho-LHCII dissociation. Possible implications of the unbound phospho-LHCII on energy dissipation are discussed.

fluorescence decay component in WT cells showed a lifetime of ∼170 ps (CFL170 ps) in state 1 (Fig. 1A) and ∼250 ps (CFL250 ps) in state 2 (Fig. 1B). The photon count ratio of CFL250 ps/CFL170 ps was about five times as high in state 2 as in state 1 (Fig. 1C). To record the CFL shift during a state 2 transition, we transferred the cells from PSI light to PSII light and measured their CFL every min for 5 min. We found that the initial CFL of ∼170 ps shifted to ∼250 ps over time (Fig. 1D), corresponding to the CFL observed in Fig. 1 A and B, respectively. Because we did not observe this shift in control cells kept under PSI light (Fig. S3 B and C), we attributed it to the state 2 transition. When we transferred the cells back to PSI light, the CFL170 ps became dominant again (Fig. S2 B and C), confirming that the tested cells were viable during the measurements and the underlying mechanism for the CFL shift was reversible. To avoid cell damage and “state” disturbance by laser scannings, we minimized the scanning time to 1 s (see Figs. S2 and S3 for more information). We then conducted single-cell FLIM to determine the distribution of centers where the CFL shift occurred during the state 2 transition. Fig. 2A shows the overall CFL distribution in the cell and the two major components—CFL170 ps, whose photon counts decreased during the state 2 transition, and CFL250 ps, whose photon counts increased. CFL components longer than 500 ps were negligible (less than 2% of the total counted photons) in our FLIM measurement. The spatiotemporal distribution for both CFL components showed that in 5 min the number of pixels for CFL170 ps decreased about 57% (from ∼6,800 to ∼2,900) and the number for CFL250 ps increased about 197% (from ∼3,700 to ∼11,000; Fig. 2B). The total photon counts decreased about 57% (from ∼4,200 to ∼1,800) for the CFL170 ps component and increased about 167% (from ∼2,100 to ∼5,600) for the CFL250 ps component (Fig. 2C). FLIM also revealed that

Fig. 1. Measurement of CFL in live C. reinhardtii WT cells in states 1 and 2. CFL images in state 1 (A) and state 2 (B) cells as indicated by the color scale from 100 to 300 ps. (Scale bars, 10 μm.) (C) The photon count ratio of CFL250 ps /CFL170 ps (±SD, n = 11 cells) in state 1 (S1) and state 2 (S2) cells. (D) Timelapse CFL images of WT cells during state 2 transitions as indicated by the color scale from 100 to 300 ps. (Scale bar, 10 μm.)

2338 | www.pnas.org/cgi/doi/10.1073/pnas.0908808107

several large spotted areas of CFL250 ps became apparent in state 2 (Fig. 2B, 4–5 min). Because phosphorylation of LHCII causes it to dissociate from PSII (6, 7, 11), we carried out immunoblot analysis using an antiphosphothreonine antibody and WT cells grown in liquid media that were sampled during the same state 2 induction. We found that activation of LHCII phosphorylation was accompanied by an increase in the CFL250 ps photon count during state 2 transition (Fig. 2C). The phosphorylation signal was about 8.4 times as high in state 2 as in state 1. To corroborate that the increase in the CFL250 ps reflected the dissociated phospho-LHCII, we tested the stt7 mutant, which is deficient in a protein kinase for LHCII (15). Because of the lack of LHCII phosphorylation, LHCII should remain associated with PSII in this mutant even under state 2-inducing conditions. As expected, FLIM showed that the CFL170 ps was the dominant component in the stt7 mutant all of the time (Fig. 2 D–F), and no LHCII phosphorylation was observed during the PSII-light illumination (Fig. 2F, Lower). It is therefore likely that the CFL170 ps signal in state 1 represented the PSII-LHCII supercomplex, and the increase in CFL250 ps signal in state 2 reflected the dissociated phospho-LHCII; however, we could not exclude the possibility that CFL250 ps originated from the PSII core or the PSI supercomplex associated with LHCII. Chlorophyll Fluorescence Lifetime of Dissociated Phospho-LHCII In Vivo. To determine the CFL of dissociated LHCII in vivo, we

used ΔPSI/II, a C. reinhardtii mutant that lacks both PSI and PSII but accumulates a normal amount of all of the other thylakoid proteins, including PSI light-harvesting antenna complex (LHCI) and LHCII, when grown heterotrophically (Fig. 3A). The FLIM measurement, performed exactly as for the WT cells, revealed that the major chlorophyll fluorescence had a lifetime of ∼265 ps (CFL265 ps; Fig. 2G) in ΔPSI/II, and we observed no clear evidence of CFL170 ps. Because CFL170 ps is close to the values obtained for PSII membranes (16), the absence of CFL170 ps was likely due to the lack of PSII in the mutant. We also observed no clear CFL shift in ΔPSI/II under the state 2-inducing conditions (Fig. 2 H and I), which was likely due to the lack of state transitions in this mutant. The spatiotemporal distribution for CFL265 ps indicated that the large spotted areas were observed throughout state 2 induction (Fig. 2H). The absence of PSI in ΔPSI/II could cause the emergence of fluorescence from LHCI, but that would be minimal because LHCII is much more abundant than LHCI (Fig. 3 B and C) and overwhelms the spectral contribution at 665–685 nm at room temperature. The CFL265 ps from ΔPSI/II thus most likely originated from the dissociated LHCII, with no effect from PSI and PSII. Immunoblot analysis of WT cells showed that the increase in CFL250 ps occurred simultaneously with the activation of LHCII phosphorylation (Figs. 2C, Lower), so we also examined the phosphorylation state of LHCII in the ΔPSI/II mutant by immunoblot analysis. The results indicated that LHCII phosphorylation was already activated under PSI light and was sustained during the illumination with PSII light (Fig. 2I, Lower). This could account for the preshifted CFL in this mutant (Fig. 2 G–I). The cause of the activation of LHCII kinase in the mutant is unknown, but it is possible that reducing equivalents derived from chlororespiration (17) keep the PQ pool reduced and thereby cause constitutive LHCII phosphorylation. Because the phospho-LHCII in ΔPSI/II cells is not associated with a photosystem, it could reflect the properties of the dissociated form of phospho-LHCII during state 2 transition in WT cells. The slight difference between the CFL250 ps in WT cells and the CFL265 ps in ΔPSI/II mutant cells could be a result of the differences in antenna size and/or could be within the margin of error of our measurements because the time resolution of our FLIM system was 40 ps. Another possibility is that some CFL250 ps in WT Iwai et al.

Fig. 2. Quantitative measurements of the CFL shift observed during state 2 transitions in single WT (A–C), stt7 (D–F), and ΔPSI/II (G–I) cells. The overall CFL distribution measured in single WT (A), stt7 (D), and ΔPSI/II (G) cells during the 5-min state 2 transition. The spatiotemporal distribution of the CFL shift observed in single WT (B), stt7 (E), and ΔPSI/II (H) cells during the same time course shown in A, D, and G, respectively. Colors indicate pixels whose strongest CFL was 170 ± 10 ps (blue) and 250 ± 10 ps (red). These colors are overlaid on the total fluorescence images indicated in gray scale. (Scale bars, 5 μm.) The change in the photon count for CFL170 ps (blue) and CFL250 ps (red) observed in single WT (C), stt7 (F), and ΔPSI/II (I) cells during the same time course shown in A, D, and G, respectively. Phosphorylation of major LHCII detected by immunoblot analysis using an anti-phosphothreonine antibody is shown under the photon count data.

Biochemical Analysis of Phospho-LHCII. Although phospho-LHCII dissociation is fundamental to state transitions, we know little about the fate of dissociated LHCII in vivo. As shown in Fig. 2, and also in our previous study (11), LHCII dissociates only when it is phosphorylated. When we subjected isolated LHCII to sucrose density gradient ultracentrifugation, the density was higher for that from state 2 cells than that from state 1 cells (Fig. 4A). Immunoblot analysis confirmed that higher-density LHCII was more phosphorylated than lower-density LHCII (Fig. 4B). Moreover, negative-staining electron microscopy revealed that phospho-LHCII formed a significantly larger (possibly aggregated) structure (Fig. 4 Iwai et al.

C and D). We estimate that the large aggregated structure consists of 4–7 LHCII trimers. A similar structure was previously observed as a native form of LHCII in thylakoid membranes (21). The structural alteration conferred by LHCII phosphorylation (11, 22– 24) could cause these aggregations. Furthermore, FLIM and spectroscopic measurements indicated that the phospho-LHCII was energy-dissipative as compared with unphosphorylated LHCII (Fig. 4 E and F and Fig. S4). Discussion It is widely accepted that the light energy balance between PSII and PSI is maintained by redistributing LHCII, which dissociates from PSII upon LHCII phosphorylation under conditions where PSII is more excited than PSI (25). Advanced technology for live-cell imaging has long been awaited to visualize phosphoLHCII dissociation during state transitions in vivo. In this study using FLIM to differentiate chlorophyll autofluorescence by its lifetime, we revealed a CFL shift in live C. reinhardtii cells in response to changing light conditions (Fig. 1). Our observations strongly suggest that the CFL shift originates from phosphoLHCII dissociation from PSII in vivo (Fig. 2). Single-cell FLIM further indicates that the dissociated phospho-LHCII spreads PNAS | February 2, 2010 | vol. 107 | no. 5 | 2339

PLANT BIOLOGY

cells originates from the PSI associated with LHCII formed in state 2. This possibility can be excluded, however, because less than 2% of the fluorescence at 665–685 nm is derived from PSI at room temperature (13), and the CFL of PSI fluorescence at those wavelengths is shorter than ∼70 ps (18–20), a time range that our FLIM system cannot resolve. Taken together, these observations suggest that the CFL250 ps observed in WT cells indicates phosphoLHCII dissociation, and that FLIM enabled visualization of the spatiotemporal spread of dissociated phospho-LHCII during state 2 transitions in live cells.

Fig. 3. Protein composition in the ΔPSI/II mutant. (A) Immunoblot analysis of thylakoid membranes isolated from WT and ΔPSI/II cells using antibodies specific for PSI (PsaA, PsaF), PSII (CP47, D1), cytochrome b6f complex (Cyt b6, Cyt f), minor LHCII (CP26, CP29), and major LHCII (LhcbM6). Protein samples were normalized to the amount of major LHCII. (B) Sucrose density gradient centrifugation to separate chlorophyll-protein complexes of thylakoid membranes isolated from ΔPSI/II. (C) SDS/PAGE analysis of the chlorophyllprotein complexes separated in the sucrose density gradient in B. The gel was stained with Coomassie brilliant blue R-250. Arrows and arrowheads indicate the protein bands for LHCII and LHCI, respectively.

through the cell during state 2 transitions and forms several large spotted areas (Fig. 2B). Our biochemical analyses indicated phospho-LHCII formed a large aggregated structure whereas unphosphorylated LHCII did not (Fig. 4 C and D). The underlying mechanism for the aggregation is unclear. We know, however, that aggregated phosphoLHCII were in an energy-dissipative state (Fig. 4 E and F and Fig. S4; see also refs. 26–28). Thus, the free phospho-LHCII aggregates appearing during state 2 transitions could be energy-dissipative, as has recently been suggested (29, 30). Because our experimental procedure did not give high light illumination (>500 μmol photons m−2 s−1, which is required to induce qE quenching in C. reinhardtii; see ref. 31), the energy-dissipative phospho-LHCII aggregates observed in this study were not likely exhibiting qE quenching, but exhibiting qT (state transition) quenching. However, a recent report suggested that qE quenching also involved LHCII dissociation (32). It is thus tempting to speculate that both qE and qT quenching are causally related by the energy-dissipative free LHCII aggregates, even though the trigger, high light or reduced PQ pool, and the detaching mechanism, PsbS or LHCII phosphorylation, are different. Although the fluorescence lifetime increases from CFL170 ps to CFL250 ps during the formation of phospho-LHCII aggregates in state 2, an 80-ps difference would only make a marginal fluorescence yield difference. However, there is a substantial difference between the CFL of phospho-LHCII in vivo (∼250 ps; Fig. 2) and that in vitro (∼1300 ps; Fig. 4), which suggests that an additional factor(s) must be involved in the shorter CFL (energy dissipation) in phospho-LHCII in vivo. Recently, it was proposed that dissociated LHCII from the PSII supercomplex have contact with PsbS, thereby forming a quenching center under high light conditions (32). An unidentified component might be similarly associated with unbound phospho-LHCII during state 2 in C. reinhardtii cells. 2340 | www.pnas.org/cgi/doi/10.1073/pnas.0908808107

Fig. 4. Biochemical differences between LHCII and phospho-LHCII. (A) Thylakoid membranes isolated from WT cells in state 1 (S1) and state 2 (S2) were subjected to sucrose density gradient ultracentrifugation, which separated LHCII, PSII, and PSI-LHCI protein complexes. (B) Phosphorylated proteins in the LHCII fractions obtained in A are shown by immunoblot analysis using an anti-phosphothreonine antibody. Proteins were normalized with the amount of LHCII protein (α-LHCII). Electron micrographs of the negativestained LHCII fractions in S1 (C) and S2 (D) obtained in A. (Scale bars, 50 nm.) Energy-dissipative state of phospho-LHCII are shown by mean CFL of LHCII fractions in state 1 (E) and state 2 (F) which were obtained in A. Color scale indicates mean CFL. (Scale bars, 5 μm.)

On the basis of these observations, we propose a model for LHCII migration between the two photosystems during state transitions in C. reinhardtii (Fig. 5). As previously shown (8–11), LHCII phosphorylation induces the dissociation of LHCII (both minor and major LHCII, as shown in refs. 11, 12, 29) from PSII, leading to a decrease in the excitation level of PSII (Fig. 5A). Several dissociated phospho-LHCII are then incorporated into aggregated structures (Fig. 5B), which might represent the substantial amounts of free LHCII at the margin of grana and the appressed region of thylakoid membranes (22, 33). Aggregated phospho-LHCII is more energy-dissipative than the monomeric form, thereby suppressing the deleterious effects of the excess energy. This is advantageous because PSI-localized stroma lamellae and the non-appressed region of the thylakoid membranes could be too distant for LHCII to travel to from PSIIlocalized grana (the appressed region). Subsequently, to increase the excitation level in PSI, a certain number of LHCII (including minor LHCII) need to migrate from the aggregated structure (Fig. 5C) to the stroma lamellae (non-appressed region) and reassociate with PSI (12, 34, 35), thereby completing the state 2 transition (Fig. 5D). During the transition back to state 1, LHCII dissociates from PSI (Fig. 5D′) and associates with the aggregated structure (Fig. 5C′). Several LHCII will then move back to PSII (Fig. 5 A′ and B′). Above all, FLIM technology allows us to watch the dissociation of phospho-LHCII from PSII during state transitions. Surprisingly, the undocked phospho-LHCII formed an energydissipative aggregation. Many questions are then raised: What is the significance of a state transition—to decrease the absorption cross-section of PSII, to increase that of PSI, or both? Does a state transition really increase the PSI absorption cross-section? Is there a pool of unbound LHCII all of the time? How much Iwai et al.

Fluorescence Lifetime Imaging Microscopy. For FLIM measurements, we used a Leica TCS SP5 confocal laser scanning microscope equipped with a timecorrelated single-photon counting module (SPC-830; Becker & Hickl). A diode pulsed laser (LDH-P-C-405B/PDL 800-B; PicoQuant; 405 nm, 40 MHz, 480 nW) was used to excite chlorophyll, and the emission from 665 to 685 nm was collected through an HCX PL APO lbd.BL 63 × 1.4 NA oil objective (Leica Microsystems) in 256 × 256 pixel mode. Live cells grown on TAP agar media were placed directly on a slide glass and observed with a coverslip under continuous illumination with weak PSI or PSII light. We observed isolated LHCII in a coverglass chamber in the dark. The lifetime data for live cells and isolated LHCII were collected according to the manufacturer’s recommendations (Leica 1-ch D-FLIM; http://www.becker-hickl.de/leicaman.htm) with modified TAC gain and offset to 5% and 30%, respectively. The collection time was 1 s for live cells and 30 s for isolated LHCII.

N

undocked LHCII actually reassociates with PSI? And, is there a relationship between state transitions and qE quenching? Further investigation of these issues is required if we are to fully understand how plants adapt to changing light environments. Materials and Methods Strains and Growth Conditions. Wild-type C. reinhardtii strain 137c and the stt7 mutant (a kind gift from Jean-David Rochaix, Geneva, Switzerland) were grown in Tris-acetate-phosphate media (36) under low light (∼20 μmol photons m−2 s−1). The ΔPSI/II mutant was generated biolistically with tungsten particles coated with pAA400 plasmid in which the psaA gene was deleted, and a host ΔPSII mutant FuD7 (37) that we obtained from the Chlamy Center (http://www.chlamy.org). The ΔPSI/II mutant was grown in Tris-acetate-phosphate media in the dark.

τm ¼ ∑ ai τi i¼1



N

∑ ai : i¼1

Biochemical Analysis. Isolation of thylakoid membranes, solubilization of the isolated thylakoid membranes with tridecyl-β-D-maltoside, fluorescence spectroscopy, pigment analysis by HPLC, and SDS/PAGE were performed as described previously (11). Sucrose density gradient centrifugation and immunoblotting were performed essentially as described (12), except here we used a P40ST rotor (Hitachi) at 91,500 × g for 24 h and antibodies against cytochrome b6 and cytochrome f (Agrisera) and LhcbM6 polypeptide (a kind gift from Michael Hippler, Münster, Germany). Electron Microscopy. Single-particle isolated LHCII were observed on carboncoated grids under an H-7650 Zero A electron microscope (Hitachi) and captured digitally at ×50,000 as described previously (11).

Induction of State 2 Transitions by Illumination with Two Different Lights. As shown previously (11), C. reinhardtii cells grown on Tris-acetate-phosphate agar in liquid media under low light (WT and stt7 mutant) or in the dark (ΔPSI/II mutant) were placed in the dark for 15 min. They were illuminated with weak PSI light through a band-pass filter (720 ± 10 nm; ∼10 μmol photons m−2 s−1) for 15 min to induce state 1. After that, to induce a state 2 transition, we illuminated them with weak PSII light through a band-pass filter (467 ± 10 nm; ∼10 μmol photons m−2 s−1), sampling them every min for 5 min for FLIM measurements and immunoblot analysis. To recover state 1, the state 2 cells at 5 min were further illuminated by the PSI light for 15 min. The experimental scheme is depicted in Fig. S2A.

ACKNOWLEDGMENTS. We thank Profs. Jean-David Rochaix and Michael Hippler for providing us with the stt7 mutant and LhcbM6 antibody, respectively. This study was supported by the Research Fellowship for Young Scientists from the Japan Society for the Promotion of Science (M.I.), Grants-inAid for Scientific Research for Plant Graduate Students from Nara Institute of Science and Technology, the Ministry of Education, Culture, Sports, Science, and Technology (N.I.), a Research Grant in the Natural Sciences by the Mitsubishi Foundation, the Strategic International Cooperative Program by the Japan Science and Technology Agency, and Grants-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science, and Technology (J.M.).

1. Murata N (1969) Control of excitation transfer in photosynthesis. I. Light-induced

8. Andersson B, Åkerlund H-E, Jergil B, Larsson C (1982) Differential phosphorylation of

change of chlorophyll a fluorescence in Porphyridium cruentum. Biochim Biophys Acta 172:242–251. Bonaventura C, Myers J (1969) Fluorescence and oxygen evolution from Chlorella pyrenoidosa. Biochim Biophys Acta 189:366–383. Allen JF, Bennett J, Steinback KE, Arntzen CJ (1981) Chloroplast protein phosphorylation couples plastoquinone redox state to distribution of excitation energy between photosystems. Nature 291:25–29. Wollman F-A, Lemaire C (1988) Studies on kinase-controlled state transitions in photosystem II and b6 f mutants from Chlamydomonas reinhardtii which lack quinone-binding proteins. Biochim Biophys Acta 933:85–94. Bellafiore S, Barneche F, Peltier G, Rochaix J-D (2005) State transitions and light adaptation require chloroplast thylakoid protein kinase STN7. Nature 433:892–895. Bennett J, Steinback KE, Arntzen CJ (1980) Chloroplast phosphoproteins: Regulation of excitation energy transfer by phosphorylation of thylakoid membrane polypeptides. Proc Natl Acad Sci USA 77:5253–5257. Kyle DJ, Haworth P, Arntzen CJ (1982) Thylakoid membrane protein phosphorylation

the light-harvesting chlorophyll-protein complex in appressed and non-appressed regions of the thylakoid membrane. FEBS Lett 149:181–185. 9. Kyle DJ, Staehelin LA, Arntzen CJ (1983) Lateral mobility of the light-harvesting complex in chloroplast membranes controls excitation energy distribution in higher

2. 3.

4.

5. 6.

7.

leads to a decrease in connectivity between photosystem II reaction centers. Biochim Biophys Acta 680:336–342.

Iwai et al.

plants. Arch Biochem Biophys 222:527–541. 10. Bassi R, Giacometti GM, Simpson DJ (1988) Changes in the organization of stroma membranes induced by in vivo state 1–state 2 transition. Biochim Biophys Acta 935: 152–165. 11. Iwai M, Takahashi Y, Minagawa J (2008) Molecular remodeling of photosystem II during state transitions in Chlamydomonas reinhardtii. Plant Cell 20:2177–2189. 12. Takahashi H, Iwai M, Takahashi Y, Minagawa J (2006) Identification of the mobile light-harvesting complex II polypeptides for state transitions in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 103:477–482. 13. Wendler J, Holzwarth AR (1987) State transitions in the green alga Scenedesmus obliquus probed by time-resolved chlorophyll fluorescence spectroscopy and global data analysis. Biophys J 52:717–728. 14. Govindjee (1995) Sixty-three years since Kautsky: Chlorophyll a fluorescence. Aust J Plant Physiol 22:131–160.

PNAS | February 2, 2010 | vol. 107 | no. 5 | 2341

PLANT BIOLOGY

Fig. 5. A model for the dissociation of phospho-LHCII from PSII during state 2 transitions. (A) Dissociation of phospho-LHCII (both major and minor LHCII) from PSII. (B) Aggregation of phospho-LHCII. (C) Migration of LHCII from aggregates to PSI-LHCI supercomplex. (D) Association of LHCII (both major and minor LHCII) with PSI-LHCI supercomplex. The reverse process occurs during the transition from state 2 to state 1 (D’–A’). The crystal coordinates were obtained from the Protein Data Bank: PSII core dimer, 2AXT; LHCII, 2NHW; and PSI-LHCI supercomplex, 2O01.

FLIM Data Analysis. We analyzed the lifetime data measured by FLIM using SPCImage software (Becker & Hickl) with the default settings (scatter, shift, and decay components unfixed). A triple exponential decay model was employed for curve fitting to calculate lifetime components. We set the binning of pixels to 8–10 for improved lifetime accuracy and used only images with χ2 values of 0.95–1.30. The raw data containing each CFL component per pixel were exported to Mathematica (Wolfram Research) to obtain the CFL distribution against the photon count observed in measured cells. Mean lifetime (τm) for Fig. 4 E and F, which we calculated using SPCImage software, was defined by the equation

15. Depège N, Bellafiore S, Rochaix J-D (2003) Role of chloroplast protein kinase Stt7 in LHCII phosphorylation and state transition in Chlamydomonas. Science 299:1572–1575. 16. Broess K, et al. (2006) Excitation energy transfer and charge separation in photosystem II membranes revisited. Biophys J 91:3776–3786. 17. Bennoun P (1982) Evidence for a respiratory chain in the chloroplast. Proc Natl Acad Sci USA 79:4352–4356. 18. Ihalainen JA, et al. (2005) Kinetics of excitation trapping in intact photosystem I of Chlamydomonas reinhardtii and Arabidopsis thaliana. Biochim Biophys Acta 1706: 267–275. 19. van der Weij-de Wit CD, Ihalainen JA, van Grondelle R, Dekker JP (2007) Excitation energy transfer in native and unstacked thylakoid membranes studied by low temperature and ultrafast fluorescence spectroscopy. Photosynth Res 93:173–182. 20. Yokono M, Iwai M, Akimoto S, Minagawa J (2008) Simulation of excitation energy transfer within the PSI-LHCI/II supercomplex from Chlamydomonas reinhardtii. Energy from the Sun, eds Allen J, Gantt E, Golbeck J, Osmond B (Springer, Dordrecht), pp 1027–1030. 21. Dekker JP, van Roon H, Boekem EJ (1999) Heptameric association of light-harvesting complex II trimers in partially solubilized photosystem II membranes. FEBS Lett 449: 211–214. 22. Peter GF, Thornber JP (1991) Biochemical composition and organization of higher plant photosystem II light-harvesting pigment-proteins. J Biol Chem 266:16745–16754. 23. Zer H, et al. (1999) Regulation of thylakoid protein phosphorylation at the substrate level: Reversible light-induced conformational changes expose the phosphorylation site of the light-harvesting complex II. Proc Natl Acad Sci USA 96:8277–8282. 24. Várkonyi Z, et al. (2009) Effect of phosphorylation on the thermal and light stability of the thylakoid membranes. Photosynth Res 99:161–171. 25. Rochaix J-D (2007) Role of thylakoid protein kinases in photosynthetic acclimation. FEBS Lett 581:2768–2775.

2342 | www.pnas.org/cgi/doi/10.1073/pnas.0908808107

26. Mullineaux CW, Pascal AA, Horton P, Holzwarth AR (1993) Excitation-energy quenching in aggregates of the LHC II chlorophyll-protein complex: A time-resolved fluorescence study. Biochim Biophys Acta 1141:23–28. 27. Ruban AV, Horton P (1992) Mechanism of ΔpH-dependent dissipation of absorbed excitation energy by photosynthetic membranes. I. Spectroscopic analysis of isolated light-harvesting complexes. Biochim Biophys Acta 1102:30–38. 28. Lambrev PH, et al. (2007) Importance of trimer-trimer interactions for the native state of the plant light-harvesting complex II. Biochim Biophys Acta 1767:847–853. 29. Tokutsu R, Iwai M, Minagawa J (2009) CP29, a monomeric light-harvesting complex II protein, is essential for state transitions in Chlamydomonas reinhardtii. J Biol Chem 284:7777–7782. 30. Ruban AV, Johnson MP (2009) Dynamics of higher plant photosystem cross-section associated with state transitions. Photosynth Res 99:173–183. 31. Baroli I, et al. (2004) Photo-oxidative stress in a xanthophyll-deficient mutant of Chlamydomonas. J Biol Chem 279:6337–6344. 32. Betterle N, et al. (2009) Light-induced dissociation of an antenna hetero-oligomer is needed for non-photochemical quenching induction. J Biol Chem 284:15255–15266. 33. Danielsson R, Albertsson PA, Mamedov F, Styring S (2004) Quantification of photosystem I and II in different parts of the thylakoid membrane from spinach. Biochim Biophys Acta 1608:53–61. 34. Kouřil R, et al. (2005) Structural characterization of a complex of photosystem I and light-harvesting complex II of Arabidopsis thaliana. Biochemistry 44:10935–10940. 35. Kargul J, et al. (2005) Light-harvesting complex II protein CP29 binds to photosystem I of Chlamydomonas reinhardtii under state 2 conditions. FEBS J 272:4797–4806. 36. Gorman DS, Levine RP (1965) Cytochrome f and plastocyanin: Their sequence in the photosynthetic electron transport chain of Chlamydomonas reinhardi. Proc Natl Acad Sci USA 54:1665–1669. 37. Bennoun P, et al. (1986) Characterization of photosystem II mutants of Chlamydomonas reinhardtii lacking the psbA gene. Plant Mol Biol 6:151–160.

Iwai et al.