FRAP Analysis on Red Alga Reveals the Fluorescence Recovery Is Ascribed to Intrinsic Photoprocesses of Phycobilisomes than Large-Scale Diffusion

Abstract

Background

Phycobilisomes (PBsomes) are the extrinsic antenna complexes upon the photosynthetic membranes in red algae and most cyanobacteria. The PBsomes in the cyanobacteria has been proposed to present high lateral mobility on the thylakoid membrane surface. In contrast, direct measurement of PBsome motility in red algae has been lacking so far.

Methodology/Principal Findings

In this work, we investigated the dynamics of PBsomes in the unicellular red alga Porphyridium cruentum in vivo and in vitro, using fluorescence recovery after photobleaching (FRAP). We found that part of the fluorescence recovery could be detected in both partially- and wholly-bleached wild-type and mutant F11 (UTEX 637) cells. Such partial fluorescence recovery was also observed in glutaraldehyde-treated and betaine-treated cells in which PBsome diffusion should be restricted by cross-linking effect, as well as in isolated PBsomes immobilized on the glass slide.

Conclusions/Significance

On the basis of our previous structural results showing the PBsome crowding on the native photosynthetic membrane as well as the present FRAP data, we concluded that the fluorescence recovery observed during FRAP experiment in red algae is mainly ascribed to the intrinsic photoprocesses of the bleached PBsomes in situ, rather than the rapid diffusion of PBsomes on thylakoid membranes in vivo. Furthermore, direct observations of the fluorescence dynamics of phycoerythrins using FRAP demonstrated the energetic decoupling of phycoerythrins in PBsomes against strong excitation light in vivo, which is proposed as a photoprotective mechanism in red algae attributed by the PBsomes in response to excess light energy.

About the Authors:

Lu-Ning Liu

Affiliations State Key Lab of Microbial Technology, Marine Biotechnology Research Center, Shandong University, Jinan, People's Republic of China, Department of Biophysics, Huygens Laboratory, Leiden University, Leiden, The Netherlands

Thijs J. Aartsma

Affiliation: Department of Biophysics, Huygens Laboratory, Leiden University, Leiden, The Netherlands

Jean-Claude Thomas

Affiliation: UMR 8186 CNRS & Ecole Normale Supérieure, Biologie Moléculaire des Organismes Photosynthétiques, Paris, France

Bai-Cheng Zhou

Affiliation: State Key Lab of Microbial Technology, Marine Biotechnology Research Center, Shandong University, Jinan, People's Republic of China

Yu-Zhong Zhang

* E-mail: [email protected]

Affiliation: State Key Lab of Microbial Technology, Marine Biotechnology Research Center, Shandong University, Jinan, People's Republic of China

Introduction

Photosynthetic organisms are able to capture solar energy using light-harvesting antennae in the photosynthesis process [1], [2]. In the eukaryotic red algae and most of the prokaryotic cyanobacteria, the light-harvesting antennae are extrinsic supramolecular complexes, designated phycobilisomes (PBsomes). PBsomes are arranged on the surface of photosynthetic membranes [3]–[5], absorbing light and transferring it to the reaction center with high efficiency [6]–[8].

The major components of PBsomes are the phycobiliproteins (PBPs). According to the different properties, PBPs are commonly divided into four main classes: allophycocyanin (APC), phycocyanin (PC), phycoerythrin (PE) and phycoerythrocyanin (PEC) [1], [3], [9]. In the presence of linker polypeptides, PBPs are assembled into two subcomplexes: the core that combines with the membrane-bound photosynthetic reaction centers, and the peripheral rods that attach to the core [7], [10]–[12]. The former is mainly composed of APCs, and the latter contain PCs, PCs+PEs or PCs+PECs, depending on the species [1], [3], [4]. In the unicellular red alga Porphyridium (P.) cruentum, the main components of PBPs are PEs, which consist of B-PE and b-PE [13], [14]. These two types of PEs are both heterooligomers, but they differ by the presence/absence of the γ subunit [13], [15], [16].

Fluorescence recovery after photobleaching (FRAP) has been exploited to study the protein diffusion since 1970s [17]–[19]. It has been applied in exploring the mobility of PBsomes in cyanobacteria, suggesting that PBsomes could diffuse laterally on the surface of thylakoid membrane [20]–[22]. Such rapid mobility of PBsomes was further proposed to be a prerequisite for the light-state transition [23], a physiological mechanism that adapts to distribute the excitation light between photosystem I and photosystem II [24]–[26]. To date, the measurement of PBsome mobility in red algae has been lacking. Structural explorations have revealed larger size of the hemiellipsoidal PBsomes in P. cruentum compared to the hemidiscoidal PBsomes in cyanobacteria, and significant macromolecular crowding of PBsomes on the thylakoid surface in red algae [14], [27], [28]. These findings raise the question: the rapid and long-range diffusion of PBsomes in red algae may be highly restricted by a combination of macromolecular crowding of membrane surface and specific protein-protein interactions upon the thylakoid surface [29], [30].

In this paper, using FRAP we directly investigated the dynamics of PBsomes in P. cruentum wild-type (WT) and mutant F11 cells, as well as the isolated PBsomes in vitro. Our findings revealed that the fluorescence recovery observed is due to the intrinsic photoprocesses of PBsomes in situ, rather than large-scale PBsome diffusion in vivo. Furthermore, in situ light-induced energetic decoupling of PBsomes investigated by FRAP was proposed as an energy dissipation mechanism in vivo in response to excess light energy.

Results

Fluorescence Recovery of Partially Photobleached WT Cell

FRAP has been performed to study the mobility of PBsomes on cyanobacterial thylakoid membrane [20], [31]–[33]. Here we applied the same strategy to individual cells of the red alga P. cruentum for analyzing the diffusion dynamics of PBsomes upon the thylakoid membranes. In the experiments, confocal fluorescence images were acquired of a particular plane in the cell. We excited the PBsomes in P. cruentum cells with a 568 nm laser which was mostly absorbed by PEs in the peripheral PBsome rods. The fluorescence emission of PBsome terminal emitters was detected in the range of 650–750 nm. For pre- and post-scanning 5% of the laser power was applied, and photobleaching was performed by zooming-in scan at 100% laser power (10 mW). Figure 1A shows a typical FRAP image sequence of PBsome emission in a single P. cruentum cell. Part of the cellular area was bleached to a depth of 65% and the fluorescence recovery of the bleached area was detected by a post-scan. The recovery of the fluorescence in bleached cellular region is depicted in Figure 1B (dashed line) and adequately fitted to an exponential function (Equation 1, see Materials and Methods). The fluorescence recovery levels off at 66% of the initial fluorescence intensity only (from 35% after bleaching). The recovery rate (R) was calculated to be 39.5 s−1 with Equation 1. Furthermore, we found that more intense illumination could induce less fluorescence recovery. When the photobleaching reaches 80–90% depth, the fluorescence recovery of PBsomes was much less, though still visible. This was corroborated with previous finding in cyanobacterium Thermosynechococcus elongates [33].

[Figure omitted. See PDF.]

Figure 1. Qualitative FRAP experiments of red alga P. cruentum WT cell.

A, selected fluorescence images from typical sequences recorded before bleaching, immediately after bleaching of PBsomes, and at various time lapses. Excitation is at 568 nm and detection range is from 650 to 750 nm. Scale bar: 5 µm; B, total fluorescence intensity of bleached cell region as a function of time. The recovery of the fluorescence is presented as square spots and fitted to an exponential function (solid line).

https://doi.org/10.1371/journal.pone.0005295.g001

Fluorescence Recovery of Wholly Photobleached WT Cell

To further survey the photodynamics of PBsome complexes, we explored a series of comparative experiments with the aforementioned FRAP procedure to whole cells. Figure 2 shows the time-course photobleaching of the whole P. cruentum cell. Three continuous cycles of photobleaching were carried out on the same cell. All the PBsomes were generally bleached by whole-cell scanning with intense laser power (100%). Generally, no fluorescence recovery by the lateral mobility of PBsomes was expected in this case. However, the partial recovery of fluorescent intensity from the whole cell was still observed. It was also found that stronger bleaching leads to less recovery, which has been observed in partially photobleached cell. There is one probability that such fluorescence recovery may be ascribed to the diffusion of PBsomes from neighboring lamellae surfaces. However, we found the fact that scanning with full power caused bleaching of PBsomes not only in the focusing plane, but also perpendicularly, in a few-micrometer flanking areas through the cell (data not shown).

[Figure omitted. See PDF.]

Figure 2. FRAP selected fluorescence images of the P. cruentum WT cells.

A, wholly bleaching on the cells. Excitation is at 568 nm and detection range is from 650 to 750 nm. Selected fluorescence images from sequences recorded before bleaching, immediately after bleaching of PBsomes, and at various time lapses. Cycle 1: 1-2-3; cycle 2: 3-4-5; cycle 3: 5-6-7; Scale bar: 5 µm; B, one-dimensional bleaching profiles derived from the sequences of fluorescence images of panel A.

https://doi.org/10.1371/journal.pone.0005295.g002

Fluorescence Recovery of Glutaraldehyde/Betaine-Treated WT Cell

Furthermore, FRAP experiments were performed on whole cells pre-treated with 1% glutaraldehyde or 0.5 M betaine, which have been documented to be able to chemically promote cross-linking between PBsomes and thylakoid membrane [27], [34]–[39]. Figure 3 shows that the fluorescence recovery was unexpectedly detected in all cases. The recovery rates in all conditions were calculated to be constantly 40.0±1.0 s−1, similar to the results of the native cells mentioned above.

[Figure omitted. See PDF.]

Figure 3. FRAP selected fluorescence images of P. cruentum WT cells pretreated with glutaraldehyde/betaine.

Excitation is at 568 nm and detection range is from 650 to 750 nm. A, partially-bleached cell pretreated with betaine; B, wholly-bleached cell pretreated with betaine; C, partially-bleached cell pretreated with glutaraldehyde; D, wholly-bleached cell pretreated with glutaraldehyde. Scale bar: 5 µm.

https://doi.org/10.1371/journal.pone.0005295.g003

Analysis of The Fluorescence Profiles

Investigations on one-dimensional fluorescence profiles obtained by summing pixel values across the edge of the bleached area (along the X direction in Figure 4) may allow us to analyze the PBsome mobility in real time. As the PBsome complexes diffuse upon the membrane surface, one should observe: (1) a fluorescence loss in the unbleached area if the PBsomes diffuse across the cells (Figure 4A–C); (2) successive flattening of the slope of the fluorescence intensity gradient (Figure 4A–C); (3) changes of the recovery rate as a function of distance from the edge of the bleached area (Figure 4D–F). However, as depicted in Figure 4, the above-mentioned effects were all not obvious, indicating that there was no remarkable difference of the fluorescence behaviors between all native cells, and glutaraldehyde- or betaine-treated cells.

[Figure omitted. See PDF.]

Figure 4. Selected analysis of the fluorescence profiles across the edge of the bleached area in partially-bleached cells, allowing the real-time exploration of the fluorescence dynamics of PBsomes correlated to their lateral diffusion.

A, untreated cell; B, betaine-treated cell; C, glutaraldehyde-treated cell. Area (1), the unbleached cellular area; Area (2), the slope of the fluorescence intensity gradient; Area (3), the fluorescence recovery as a function of distance from the edge of the bleached area. The mean fluorescence intensities of three different regions (R1, R2 and R3 in Figure 4A–C) at distinct recovery time were analyzed: D, from untreated cell; E, from betaine-treated cell; F, from glutaraldehyde-treated cell. Fluorescence intensities are normalized at zero second. As shown, no remarkable difference of the fluorescence intensities in these three cellular areas could be observed, indicating the fluorescence recovery of PBsomes does not differ as a function of distance from the edge of the bleached area of theses cells.

https://doi.org/10.1371/journal.pone.0005295.g004

Fluorescence Recovery of F11 Cells

In vitro single-molecule result has elaborated that two types of PEs in the PBsomes of P. cruentum play different roles in the energy transfer [40]. Unlike B-PEs, b-PEs are not involved in the energy decoupling of PBsomes. We measured the cells of P. cruentum mutant F11 strain (UTEX 637) which is depleted of B-PEs using FRAP. In Figure 5, the fluorescence recovery of PBsomes in all partially-bleached, wholly-bleached, as well as chemical-fixed cells by glutaraldehyde or betaine was obviously observed, similar to the results of WT cells.

[Figure omitted. See PDF.]

Figure 5. FRAP selected fluorescence images of P. cruentum F11 cells.

Excitation is at 568 nm and detection range is from 650 to 750 nm. Open squares indicate bleached areas by zooming-in illumination. Scale bar: 5 µm. A, partially-bleached cell; B, wholly-bleached cell; C, partially-bleached cell pretreated with glutaraldehyde; D, partially-bleached cell pretreated with betaine.

https://doi.org/10.1371/journal.pone.0005295.g005

Moreover, PBsomes from the mutant P. cruentum F11 cell have a declined dimension due to lower level of PE content [40]–[42]. PBsomes with a smaller size were expected previously to enhance their mobility [31]. However, by examining the FRAP profiles of mutant F11 cell, we found that the fluorescence recovery has similar recovery rates as that of the native cell, suggesting that the fluorescence recovery is not affected by the dimension of PBsomes in red algae.

Fluorescence Recovery of Ensemble PBsomes in vitro

FRAP experiment was also exploited on isolated PBsomes from P. cruentum. Intact PBsomes functionally isolated from the thylakoid membrane were spread on the clean surface of glass cover slide. Photobleaching was performed on the pool of immobilized PBsome complexes by zooming-in on a small area of the sample (Figure 6, square). The fluorescence recovery of bleached PBsome aggregations were unexpectedly observed in vitro in the first 150 s timespan of our measurements. The fluorescence feature, more specifically the shape of the aggregate, remained fairly stable in the course of the experiment. It revealed that the fluorescence recovery could be detected even in the immobile PBsomes. Taken together, our experimental data strongly demonstrated that the fluorescence recovery observed is most likely ascribed to the intrinsic photoprocesses of the bleached PBsomes in situ, rather than the diffusion of PBsomes.

[Figure omitted. See PDF.]

Figure 6. Fluorescence images of isolated PBsomes from P. cruentum WT cells.

Excitation is at 568 nm and detection range is from 650 to 750 nm. Selected fluorescence images from sequences recorded before bleaching, immediately after bleaching of PBsomes, and at various time lapses. The positions of PBsome aggregations imaged out of the bleaching area did not change, indicating the immobilization of PBsomes above the substrate. Scale bar: 10 µm.

https://doi.org/10.1371/journal.pone.0005295.g006

Fluorescence Properties of PE Emission in vivo

In the meanwhile, we studied the fluorescence properties of PE during photobleaching by altering the detecting region from 650–750 nm into 550–600 nm. A lower power of scanning laser (10%) was applied in the prebleaching and postbleaching in order to record continuous fluorescence images, and maximum laser intensity (100%) was introduced to bleach the samples of interest by zooming-in scanning.

Figure 7 illustrates selected images of PE fluorescence in the cells in response to intense illumination. Interestingly, wholly (Figure 7A) and partially bleaching (Figure 7B) on the WT cells of P. cruentum both resulted in the increase of PE fluorescence intensity. Such increase was also observed in the adjacent non-bleached area near the edges of bleaching region in the partially bleached cells, probably due to the scattered bleaching light. We further studied the PE fluorescent behavior by using glutaraldehyde-treated cell as a control. As a result, the fluorescence increase was not detected (Figure 7C), demonstrating that the increase of PE fluorescence imaged is due to the energetic decoupling of PE molecules within the PBsome rods. Furthermore, experimental results of P. cruentum mutant F11 strain, which only contain b-PE assemble with neighboring R-PC [40], did not show the fluorescence increase either (Figure 7D), implying the different roles of B-PE and b-PE in the energetic decoupling. These are corroborated with the previous conclusion obtained from single-molecule experiment [40]. Furthermore, it provides the direct evidence of the energetic decoupling of PBsomes against intense exciting energy in vivo. Similar fluorescence increase of PE has also been observed in the FRAP bleach in cryptophyte [43].

[Figure omitted. See PDF.]

Figure 7. Selected PE fluorescence images in cells imaged with confocal microscopy by detecting fluorescence at 550–600 nm. A, B, native WT cells; C, WT cell pretreated with glutaraldehyde; D, F11 cell. Scale bar: 5 µm.

https://doi.org/10.1371/journal.pone.0005295.g007

Figure 8 shows quantitatively the fluorescence intensities of PE in cells as a function of time. It is seen that the bleaching on WT natural cell induced the increase of PE emission (curves 1 and 2), and longer irradiation time of photobleaching was capable to trigger more remarkable fluorescence reduction (curves 2, 3, 4 and 5). The PE fluorescence intensity of the bleached areas in WT cells pretreated with glutaraldehyde and F11 cells is relatively constant, lower than the initial fluorescence intensities before photobleaching (curves 6 and 7). The plots revealed that there is no significant recovery of PE, either from increased or declined fluorescence levels. This suggested, on the other hand, that there is no significant mobility of PEs that are the major components of PBsomes.

[Figure omitted. See PDF.]

Figure 8. Time-lapse PE fluorescence intensity in cells in the process of photobleaching.

Powerful irradiation can cause initial increase of PE and prolonged illumination time may induce bleaching of PE. (1), whole bleaching of native WT cell; (2), 5 times scan; (3), 10 times scan; (4), 20 times scan; (5), 30 times scan; (6), WT cell pretreated with glutaraldehyde; (7), F11 mutant cell.

https://doi.org/10.1371/journal.pone.0005295.g008

Fluorescence Recovery of PEs in Ensemble PBsomes in vitro

Figure 9 shows the fluorescence dynamics of PE in isolated PBsomes upon intense illumination at 514 nm. Immediately after photobleaching, continuous scan recorded a dramatic fluorescence increase in the wholly-bleached area of WT PBsomes (Figure 9A), whereas PE fluorescence was bleached in both WT PBsomes treated with glutaraldehyde (Figure 9B) and B-PE-lacking PBsomes from the F11 mutant (Figure 9C). This further confirmed the energetic decoupling of PBsomes in response to intense exciting light.

[Figure omitted. See PDF.]

Figure 9. Selected PE fluorescence images of ensemble PBsome isolated from P. cruentum WT cells (A), WT cell pretreated with glutaraldehyde (B) and F11 mutant (C) obtained with confocal microscopy by detecting 550–600 nm.

Scale bar: 5 µm.

https://doi.org/10.1371/journal.pone.0005295.g009

Discussion

Thin-section electron microscopic images of P. cruentum cell present a globular shape with a typical diameter of 5–8 µm [44], [45]. The large chloroplasts consist of parallel-arranged and unstacked lamellae that occupy most of the intra-cellular space in vivo. Such a regular thylakoid organization is supposedly suitable for FRAP measurements [21], [22].

In cyanobacteria, PBsomes have been suggested to diffuse rapidly on the surface of thylakoid membrane [20]. However, recent investigations have shown that the thylakoid membrane in the unicellular red alga P. cruentum exhibits dense crowding of PBsome complexes [27], [28]. In addition to determining the arrangements of PBsomes on the thylakoid membrane, the supramolecular crowding has also profound implications for the dynamic behavior of PBsomes in P. cruentum: the rapid and long-range movement of PBsomes may be inhibited by the dense packing of membrane surface, and the limited free vertical spacing between opposite thylakoid layers, as well as remarkable steric hindrance, taking into account the large size of individual PBsomes. Furthermore, PBsomes attach quite strongly to the membrane surfaces [46], whereas the rapid diffusion of PBsomes requires looser association of PBsomes to the photosynthetic reactions centers.

This contradiction was explored in the present work by studying the fluorescence dynamics of PBsomes on thylakoid membranes from P. cruentum using FRAP. First, FRAP results of WT cells present partial fluorescence recovery. Assuming the fluorescence recovery is attributed to the diffusion of PBsome complexes [20], it then indicated that some PBsomes are presumably immobile on the thylakoid membranes of red algae. However, this probability was not supported by the finding that the fluorescence recovery of PBsomes declines as the illumination strength increases. The partial fluorescence recovery was also observed in partially-bleached F11 cells, wholly-bleached WT and F11 cells, and even in immobilized PBsome complexes in vitro. We also detected, similar fluorescence recovery behavior of P. cruentum cells pretreated with protein cross-linking agents glutaraldehyde or betaine with the fixation effect. By comparative analysis of the fluorescence feature and recovery rate, no significant difference of the fluorescence behavior between those cells was observed. All these data strongly indicate the observed recovery is not likely ascribed to the lateral diffusion of PBsome complexes on the thylakoid membranes.

Another evidence of the PBsome diffusion in red algae was provided by recording the PE fluorescent signal instead of the fluorescence emission of PBsome terminal emitters. After photobleaching, rapid diffusion of PBsomes was expected to cause the recovery of PE fluorescence to the initial fluorescence intensity. Alternatively, we could not observe such tendency in either the area of WT cells having increased PE fluorescence intensity or the area of glutaraldehyde/betaine-pretreated WT cells and F11 cells showing decreased PE fluorescence intensity (Figures 7 and 8).

Based on all the observations we conclude that, lateral diffusion of red algal PBsomes is not the predominant mechanism which could potentially explain the fluorescence recovery that we observed by means of FRAP. Instead, the fluorescence recovery is probably ascribed to the intrinsic photoprocesses of the bleached PBsomes in situ, rather than the rapid diffusion of PBsomes on thylakoid in vivo, although the mechanism of the intrinsic photoprocesses still remains to be determined.

The mechanism for light-state transition in red algae remains controversial. Unlike the state transition in higher plant, light-dependent phosphorylation was absent in cyanobacteria and rhodophytes [47]. Rapid mobility of PBsomes in cyanobacteria has been proposed to be required for the PBsome-dependent state transition [31]. In red alga P. cruentum, the long-distance diffusion of PBsome complexes may be highly restricted, however potential diffusion of PBsomes confined to small domains at sub-optical scales would not be excluded. For instance the local conformational changes and movement of PBsomes between adjacent photosystem II and photosystem I complexes [48] can probably be involved in the energy redistribution in red algae.

By monitoring the fluorescence of PEs in the PBsomes, we studied the fluorescence dynamics of ensemble PBsomes in situ during photobleaching. We demonstrated directly the energetic decoupling of PBsomes in vivo in response to the bleaching laser. The PEs (specifically B-PEs), rather than PC/APC, are mainly involved in the energetic decoupling. This is in agreement with single-molecule observations [40] and ensemble results of isolated PBsomes (data not shown). Such an energetic decoupling of PBsomes is proposed to have important implication on the photoprotective role of PBsomes: It may allow excess photon energy from PE to photosynthetic reaction centers to be modulated to minimize the risk of chlorophyll photooxidation. Therefore, like the light-state transition, this mechanism might contribute to prevent PE-containing algae from excessive photoinhibition.

Photosynthetic membrane has been considered as an ideal model to study the protein mobility in view of their naturally fluorescent properties [21]. However, the complexity of photosynthetic membrane should be taken into account in FRAP, as numbers of fluorescent protein are located densely in the membrane with a close functional association benefiting for efficient energy transfer. Not only does the spectral overlap make it hard to distinguish the individual contribution of each complex [20], but also the photoreactions among distinct cooperated protein complexes can be triggered by exposure laser, for instance the energetic decoupling in the PBsomes as we observed and the photoblinking behaviors revealed in single-molecule studies on phycobiliproteins [49]–[52]. In addition, it was proposed that all the fluorescent molecules can undergo reversible photobleaching [53]. Therefore, investigations on each functional protein complex involved may be prerequisite for understanding the fluorescence dynamics of photosynthetic membrane.

Materials and Methods

Growth of Cells and Separation of Intact PBsomes

P. cruentum WT and mutant F11 strain UTEX 637 were grown in an artificial sea water medium [54]. Flasks were supplied with 3% CO2 in air through a plug of sterile cotton at a constant temperature of 20°C. Cultures were illuminated continuously with light provided by daylight fluorescent lamps at 6 W·m−2. Intact PBsomes were separated following the previous protocol [27].

FRAP Measurements

Growing cells or cells with corresponding pre-treatments by 1% (v/v) glutaraldehyde or 0.5 M betaine buffer were immobilized on a slide covered with a glass slip and located under objective lens of laser scanning confocal fluorescence microscopy (LSCM).

FRAP experiments were carried out with a Leica TCS NT confocal microscope. Samples were excited with a 568 nm Yellow Krypton laser (10 mW) or a 514 nm Argon laser (10 mW). PBsome emission was detected at 650–750 nm or 550–600 nm for detecting PBsome emission and PE emission, respectively. After recording a prebleaching image, the sample was bleached by zooming in and scanning with maximum laser power for three seconds. Subsequently, images were captured with the same power as that of pre-bleaching image. No detectable photobleaching during the recording of successive image scans was observed with optimized laser power.

The fluorescence recovery curve is fitted by single exponential function, given by(1)where F(t) is the intensity as time t; A and B are the amplitudes of the time-dependent and time-independent terms, respectively; τ is the lifetime of the exponential term (time constant), and the recovery rate is given by R = 1/τ.

Author Contributions

Conceived and designed the experiments: LNL TA BCZ YZZ. Performed the experiments: LNL. Analyzed the data: LNL TA BCZ YZZ. Contributed reagents/materials/analysis tools: JCT. Wrote the paper: LNL TA.

Citation: Liu L-N, Aartsma TJ, Thomas J-C, Zhou B-C, Zhang Y-Z (2009) FRAP Analysis on Red Alga Reveals the Fluorescence Recovery Is Ascribed to Intrinsic Photoprocesses of Phycobilisomes than Large-Scale Diffusion. PLoS ONE4(4): e5295. https://doi.org/10.1371/journal.pone.0005295

References

1. Glazer AN (1989) Light guides directional energy transfer in a photosynthetic antenna. J Biol Chem 264: 1–4.AN Glazer1989Light guides directional energy transfer in a photosynthetic antenna.J Biol Chem26414

2. Mullineaux CW (2005) Function and evolution of grana. Trends Plant Sci 10: 521–525.CW Mullineaux2005Function and evolution of grana.Trends Plant Sci10521525

3. MacColl R (1998) Cyanobacterial phycobilisomes. J Struct Biol 124: 311–334.R. MacColl1998Cyanobacterial phycobilisomes.J Struct Biol124311334

4. Bald D, Kruip J, Rögner M (1996) Supramolecular architecture of cyanobacterial thylakoid membranes: How is the phycobilisome connected with the photosystems? Photosynth Res 49: 103–118.D. BaldJ. KruipM. Rögner1996Supramolecular architecture of cyanobacterial thylakoid membranes: How is the phycobilisome connected with the photosystems?Photosynth Res49103118

5. Zilinskas BA, Greenwald LS (1986) Phycobilisome structure and function. Photosynth Res 10: 7–35.BA ZilinskasLS Greenwald1986Phycobilisome structure and function.Photosynth Res10735

6. Adir N (2005) Elucidation of the molecular structures of components of the phycobilisome: reconstructing a giant. Photosynth Res 85: 15–32.N. Adir2005Elucidation of the molecular structures of components of the phycobilisome: reconstructing a giant.Photosynth Res851532

7. Grossman AR, Schaefer MR, Chiang GG, Collier JL (1993) The phycobilisome, a light-harvesting complex responsive to environmental conditions. Microbiol Rev 57: 725–749.AR GrossmanMR SchaeferGG ChiangJL Collier1993The phycobilisome, a light-harvesting complex responsive to environmental conditions.Microbiol Rev57725749

8. Biggins J, Bruce D (1989) Regulation of excitation energy transfer in organisms containing phycobilins. Photosynth Res 20: 1–34.J. BigginsD. Bruce1989Regulation of excitation energy transfer in organisms containing phycobilins.Photosynth Res20134

9. Colyer CL, Kinkade CS, Viskari PJ, Landers JP (2005) Analysis of cyanobacterial pigments and proteins by electrophoretic and chromatographic methods. Anal Bioanal Chem 382: 559–569.CL ColyerCS KinkadePJ ViskariJP Landers2005Analysis of cyanobacterial pigments and proteins by electrophoretic and chromatographic methods.Anal Bioanal Chem382559569

10. Lundell DJ, Williams RC, Glazer AN (1981) Molecular architecture of a light-harvesting antenna: in vitro assembly of the rod substructures of Synechococcus 6301 phycobilisomes. J Biol Chem 256: 3580–3592.DJ LundellRC WilliamsAN Glazer1981Molecular architecture of a light-harvesting antenna: in vitro assembly of the rod substructures of Synechococcus 6301 phycobilisomes.J Biol Chem25635803592

11. Redlinger T, Gantt E (1981) Phycobilisome structure of Porphyridium cruentum: polypeptide composition. Plant Physiol 68: 1375–1379.T. RedlingerE. Gantt1981Phycobilisome structure of Porphyridium cruentum: polypeptide composition.Plant Physiol6813751379

12. Liu L-N, Chen X-L, Zhang Y-Z, Zhou B-C (2005) Characterization, structure and function of linker polypeptides in phycobilisomes of cyanobacteria and red algae: An overview. Biochim Biophys Acta 1708: 133–142.L-N LiuX-L ChenY-Z ZhangB-C Zhou2005Characterization, structure and function of linker polypeptides in phycobilisomes of cyanobacteria and red algae: An overview.Biochim Biophys Acta1708133142

13. Gantt E, Lipschultz CA (1974) Phycobilisomes of Porphyridium cruentum: pigment analysis. Biochemistry 13: 2960–2966.E. GanttCA Lipschultz1974Phycobilisomes of Porphyridium cruentum: pigment analysis.Biochemistry1329602966

14. Gantt E, Lipschultz CA, Zilinskas B (1976) Further evidence for a phycobilisome model from selective dissociation, fluorescence emission, immunoprecipitation, and electron microscopy. Biochim Biophys Acta 430: 375–388.E. GanttCA LipschultzB. Zilinskas1976Further evidence for a phycobilisome model from selective dissociation, fluorescence emission, immunoprecipitation, and electron microscopy.Biochim Biophys Acta430375388

15. Ficner R, Huber R (1993) Refined crystal structure of phycoerythrin from Porphyridium cruentum at 0.23-nm resolution and localization of the gamma subunit. Eur J Biochem 218: 103–106.R. FicnerR. Huber1993Refined crystal structure of phycoerythrin from Porphyridium cruentum at 0.23-nm resolution and localization of the gamma subunit.Eur J Biochem218103106

16. Lundell DJ, Glazer AN, DeLange RJ, Brown DM (1984) Bilin attachment sites in the alpha and beta subunits of B-phycoerythrin. Amino acid sequence studies. J Biol Chem 259: 5472–5480.DJ LundellAN GlazerRJ DeLangeDM Brown1984Bilin attachment sites in the alpha and beta subunits of B-phycoerythrin. Amino acid sequence studies.J Biol Chem25954725480

17. Axelrod D, Koppel DE, Schlessinger J, Elson E, Webb WW (1976) Mobility measurement by analysis of fluorescence photobleaching recovery kinetics. Biophys J 16: 1055–1069.D. AxelrodDE KoppelJ. SchlessingerE. ElsonWW Webb1976Mobility measurement by analysis of fluorescence photobleaching recovery kinetics.Biophys J1610551069

18. Zhang F, Lee GM, Jacobson K (1993) Protein lateral mobility as a reflection of membrane microstructure. Bioessays 15: 579–588.F. ZhangGM LeeK. Jacobson1993Protein lateral mobility as a reflection of membrane microstructure.Bioessays15579588

19. Lippincott-Schwartz J, Altan-Bonnet N, Patterson GH (2003) Photobleaching and photoactivation: following protein dynamics in living cells. Nat Cell Biol: Suppl S7–14.J. Lippincott-SchwartzN. Altan-BonnetGH Patterson2003Photobleaching and photoactivation: following protein dynamics in living cells.Nat Cell Biol: SupplS714

20. Mullineaux CW, Tobin MJ, Jones GR (1997) Mobility of photosynthetic complexes in thylakoid membranes. Nature 390: 421–424.CW MullineauxMJ TobinGR Jones1997Mobility of photosynthetic complexes in thylakoid membranes.Nature390421424

21. Mullineaux CW (2004) FRAP analysis of photosynthetic membranes. J Exp Bot 55: 1207–1211.CW Mullineaux2004FRAP analysis of photosynthetic membranes.J Exp Bot5512071211

22. Mullineaux CW, Sarcina M (2002) Probing the dynamics of photosynthetic membranes with fluorescence recovery after photobleaching. Trends Plant Sci 7: 237–240.CW MullineauxM. Sarcina2002Probing the dynamics of photosynthetic membranes with fluorescence recovery after photobleaching.Trends Plant Sci7237240

23. Joshua S, Mullineaux CW (2004) Phycobilisome diffusion is required for light-state transitions in cyanobacteria. Plant Physiology 135: 2112–2119.S. JoshuaCW Mullineaux2004Phycobilisome diffusion is required for light-state transitions in cyanobacteria.Plant Physiology13521122119

24. Mullineaux CW, Emlyn-Jones D (2005) State transitions: an example of acclimation to low-light stress. J Exp Bot 56: 389–393.CW MullineauxD. Emlyn-Jones2005State transitions: an example of acclimation to low-light stress.J Exp Bot56389393

25. Schluchter WM, Shen G, Zhao J, Bryant DA (1996) Characterization of psaI and psaL mutants of Synechococcus sp. strain PCC 7002: a new model for state transitions in cyanobacteria. Photochem Photobiol 64: 53–66.WM SchluchterG. ShenJ. ZhaoDA Bryant1996Characterization of psaI and psaL mutants of Synechococcus sp. strain PCC 7002: a new model for state transitions in cyanobacteria.Photochem Photobiol645366

26. Bruce D, Brimble S, Bryant DA (1989) State transitions in a phycobilisome-less mutant of the cyanobacterium Synechococcus sp. PCC 7002. Biochim Biophys Acta 974: 66–73.D. BruceS. BrimbleDA Bryant1989State transitions in a phycobilisome-less mutant of the cyanobacterium Synechococcus sp. PCC 7002.Biochim Biophys Acta9746673

27. Arteni AA, Liu L-N, Aartsma TJ, Zhang Y-Z, Zhou B-C, et al. (2008) Structure and organization of phycobilisomes on membranes of the red alga Porphyridium cruentum. Photosynth Res 95: 169–174.AA ArteniL-N LiuTJ AartsmaY-Z ZhangB-C Zhou2008Structure and organization of phycobilisomes on membranes of the red alga Porphyridium cruentum.Photosynth Res95169174

28. Liu L-N, Aartsma TJ, Thomas J-C, Lamers GEM, Zhou B-C, et al. (2008) Watching the native supramolecular architecture of photosynthetic membrane in red algae: Topography of phycobilisomes, and their crowding, diverse distribution patterns. J Biol Chem 283: 34946–34953.L-N LiuTJ AartsmaJ-C ThomasGEM LamersB-C Zhou2008Watching the native supramolecular architecture of photosynthetic membrane in red algae: Topography of phycobilisomes, and their crowding, diverse distribution patterns.J Biol Chem2833494634953

29. Kirchhoff H (2008) Molecular crowding and order in photosynthetic membranes. Trends Plant Sci 13: 201–207.H. Kirchhoff2008Molecular crowding and order in photosynthetic membranes.Trends Plant Sci13201207

30. Kirchhoff H, Haferkamp S, Allen JF, Epstein D, Mullineaux CW (2008) Protein diffusion and macromolecular crowding in thylakoid membranes. Plant Physiol 146: 1571–1578.H. KirchhoffS. HaferkampJF AllenD. EpsteinCW Mullineaux2008Protein diffusion and macromolecular crowding in thylakoid membranes.Plant Physiol14615711578

31. Sarcina M, Tobin MJ, Mullineaux CW (2001) Diffusion of phycobilisomes on the thylakoid membranes of the cyanobacterium Synechococcus 7942. Effects of phycobilisome size, temperature, and membrane lipid composition. J Biol Chem 276: 46830–46834.M. SarcinaMJ TobinCW Mullineaux2001Diffusion of phycobilisomes on the thylakoid membranes of the cyanobacterium Synechococcus 7942. Effects of phycobilisome size, temperature, and membrane lipid composition.J Biol Chem2764683046834

32. Kondo K, Mullineaux CW, Ikeuchi M (2009) Distinct roles of CpcG1-phycobilisome and CpcG2-phycobilisome in state transitions in a cyanobacterium Synechocystis sp. PCC 6803. Photosynth Res 99: 217–225.K. KondoCW MullineauxM. Ikeuchi2009Distinct roles of CpcG1-phycobilisome and CpcG2-phycobilisome in state transitions in a cyanobacterium Synechocystis sp. PCC 6803.Photosynth Res99217225

33. Yang S, Su Z, Li H, Feng J, Xie J, et al. (2007) Demonstration of phycobilisome mobility by the time- and space-correlated fluorescence imaging of a cyanobacterial cell. Biochim Biophys Acta 1767: 15–21.S. YangZ. SuH. LiJ. FengJ. Xie2007Demonstration of phycobilisome mobility by the time- and space-correlated fluorescence imaging of a cyanobacterial cell.Biochim Biophys Acta17671521

34. Brimble S, Bruce D (1989) Pigment orientation and excitation energy transfer in Porphyridium cruentum and Synechococcus sp. PCC 6301 cross-linked in light state 1 and light state 2 with glutaraldehyde. Biochim Biophys Acta 973: 315–323.S. BrimbleD. Bruce1989Pigment orientation and excitation energy transfer in Porphyridium cruentum and Synechococcus sp. PCC 6301 cross-linked in light state 1 and light state 2 with glutaraldehyde.Biochim Biophys Acta973315323

35. Li D, Xie J, Zhao J, Xia A, Li D, et al. (2004) Light-induced excitation energy redistribution in Spirulina platensis cells: “spillover” or “mobile PBSs”? Biochim Biophys Acta 1608: 114–121.D. LiJ. XieJ. ZhaoA. XiaD. Li2004Light-induced excitation energy redistribution in Spirulina platensis cells: “spillover” or “mobile PBSs”?Biochim Biophys Acta1608114121

36. Biggins J (1983) Mechanism of the light state transition in photosynthesis. I. Analysis of the kinetics of cytochrome f oxidation in state 1 and state 2 in the red alga, Porphyridium cruentum. Biochim Biophys Acta 724: 111–117.J. Biggins1983Mechanism of the light state transition in photosynthesis. I. Analysis of the kinetics of cytochrome f oxidation in state 1 and state 2 in the red alga, Porphyridium cruentum.Biochim Biophys Acta724111117

37. Bruce D, Biggins J, Steiner T, Thewalt M (1985) Mechanism of the light state transition in photosynthesis. IV. Picosecond fluorescence spectroscopy of Anacystis nidulans and Porphyridium cruentum in state 1 and state 2 at 77 K. Biochim Biophys Acta 806: 237–246.D. BruceJ. BigginsT. SteinerM. Thewalt1985Mechanism of the light state transition in photosynthesis. IV.Picosecond fluorescence spectroscopy of Anacystis nidulans and Porphyridium cruentum in state 1 and state 2 at 77 K. Biochim Biophys Acta806237246

38. Dilworth MF, Gantt E (1981) Phycobilisome-thylakoid topography on photosynthetically active vesicles of Porphyridium cruentum. Plant Physiol 67: 608–612.MF DilworthE. Gantt1981Phycobilisome-thylakoid topography on photosynthetically active vesicles of Porphyridium cruentum.Plant Physiol67608612

39. Mustardy L, Cunningham FX Jr, Gantt E (1992) Photosynthetic membrane topography: quantitative in situ localization of photosystems I and II. Proc Natl Acad Sci USA 89: 10021–10025.L. MustardyFX Cunningham JrE. Gantt1992Photosynthetic membrane topography: quantitative in situ localization of photosystems I and II.Proc Natl Acad Sci USA891002110025

40. Liu L-N, Elmalk AT, Aartsma TJ, Thomas J-C, Lamers GEM, et al. (2008) Light-induced energetic decoupling as a mechanism for phycobilisome-related energy dissipation in red algae: a single molecule study. PLoS ONE 3: e3134.L-N LiuAT ElmalkTJ AartsmaJ-C ThomasGEM Lamers2008Light-induced energetic decoupling as a mechanism for phycobilisome-related energy dissipation in red algae: a single molecule study.PLoS ONE3e3134

41. Sivan A, Arad SM (1993) Induction and characterization of pigment mutants in the red microalga Porphyridium sp (Rhodophyceae). Phycologia 32: 68–72.A. SivanSM Arad1993Induction and characterization of pigment mutants in the red microalga Porphyridium sp (Rhodophyceae).Phycologia326872

42. Sivan A, Thomas JC, Dubacq JP, Moppes D, Arad S (1995) Protoplast fusion and genetic complementation of pigment mutations in the red microalga Porphyridium sp. J Phycol 31: 167–172.A. SivanJC ThomasJP DubacqD. MoppesS. Arad1995Protoplast fusion and genetic complementation of pigment mutations in the red microalga Porphyridium sp.J Phycol31167172

43. Kaňa R, Prášil O, Mullineaux CW (2009) Immobility of phycobilins in the thylakoid lumen of a cryptophyte suggests that protein diffusion in the lumen is very restricted. FEBS Lett 583: 670–674.R. KaňaO. PrášilCW Mullineaux2009Immobility of phycobilins in the thylakoid lumen of a cryptophyte suggests that protein diffusion in the lumen is very restricted.FEBS Lett583670674

44. Gantt E, Conti SF (1965) The ultrastructure of Porphyridium cruentum. J Cell Biol 26: 365–381.E. GanttSF Conti1965The ultrastructure of Porphyridium cruentum.J Cell Biol26365381

45. Gantt E, Conti SF (1966) Granules associated with the chloroplast lamellae of Porphyridium cruentum. J Cell Biol 29: 423–434.E. GanttSF Conti1966Granules associated with the chloroplast lamellae of Porphyridium cruentum.J Cell Biol29423434

46. Mullineaux CW (2008) Phycobilisome-reaction centre interaction in cyanobacteria. Photosynth Res 95: 175–182.CW Mullineaux2008Phycobilisome-reaction centre interaction in cyanobacteria.Photosynth Res95175182

47. Gantt E, Grabowski B, Cunningham FX Jr, Green BR, Parson WW (2003) Antenna systems of red algae: phycobilisomes with photosystem II and chlorophyll complexes with photosystem I. In: Green BR, Parson WW, editors. Light-Harvesting Antennas in Photosynthesis. Dordrecht: Kluwer Academic Publishers. pp. 307–322.E. GanttB. GrabowskiFX Cunningham JrBR GreenWW Parson2003Antenna systems of red algae: phycobilisomes with photosystem II and chlorophyll complexes with photosystem I.BR GreenWW ParsonLight-Harvesting Antennas in PhotosynthesisDordrechtKluwer Academic Publishers307322

48. McConnell MD, Koop R, Vasil'ev S, Bruce D (2002) Regulation of the distribution of chlorophyll and phycobilin-absorbed excitation energy in cyanobacteria. A structure-based model for the light state transition. Plant Physiol 130: 1201–1212.MD McConnellR. KoopS. Vasil'evD. Bruce2002Regulation of the distribution of chlorophyll and phycobilin-absorbed excitation energy in cyanobacteria. A structure-based model for the light state transition.Plant Physiol13012011212

49. Ray K, Chowdhury MH, Lakowicz JR (2008) Single-molecule spectroscopic study of enhanced intrinsic phycoerythrin fluorescence on silver nanostructured surfaces. Anal Chem 80: 6942–6948.K. RayMH ChowdhuryJR Lakowicz2008Single-molecule spectroscopic study of enhanced intrinsic phycoerythrin fluorescence on silver nanostructured surfaces.Anal Chem8069426948

50. Zehetmayer P, Hellerer T, Parbel A, Scheer H, Zumbusch A (2002) Spectroscopy of single phycoerythrocyanin monomers: dark state identification and observation of energy transfer heterogeneities. Biophys J 83: 407–415.P. ZehetmayerT. HellererA. ParbelH. ScheerA. Zumbusch2002Spectroscopy of single phycoerythrocyanin monomers: dark state identification and observation of energy transfer heterogeneities.Biophys J83407415

51. Loos D, Cotlet M, De SF, Habuchi S, Hofkens J (2004) Single-molecule spectroscopy selectively probes donor and acceptor chromophores in the phycobiliprotein allophycocyanin. Biophys J 87: 2598–2608.D. LoosM. CotletSF DeS. HabuchiJ. Hofkens2004Single-molecule spectroscopy selectively probes donor and acceptor chromophores in the phycobiliprotein allophycocyanin.Biophys J8725982608

52. Zehetmayer P, Kupka M, Scheer H, Zumbusch A (2004) Energy transfer in monomeric phycoerythrocyanin. Biochim Biophys Acta 1608: 35–44.P. ZehetmayerM. KupkaH. ScheerA. Zumbusch2004Energy transfer in monomeric phycoerythrocyanin.Biochim Biophys Acta16083544

53. Verkman AS (2002) Solute and macromolecule diffusion in cellular aqueous compartments. Trends Biochem Sci 27: 27–33.AS Verkman2002Solute and macromolecule diffusion in cellular aqueous compartments.Trends Biochem Sci272733

54. Jones RF, Speer HL, Kury W (1963) Studies on the growth of the red alga Porphyridium cruentum. Physiol Plant 16: 636–643.RF JonesHL SpeerW. Kury1963Studies on the growth of the red alga Porphyridium cruentum.Physiol Plant16636643