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Journal of Bacteriology, June 2004, p. 3991-3999, Vol. 186, No. 12
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.12.3991-3999.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Involvement of the SppA1 Peptidase in Acclimation to Saturating Light Intensities in Synechocystis sp. Strain PCC 6803

E. Pojidaeva,¹ V. Zinchenko,² S. V. Shestakov,² and A. Sokolenko^1*

Department für Biologie I, Bereich Botanik, Ludwig-Maximilians-Universität, 80638 Munich, Germany,¹ Department of Genetics, Moscow State University, 119899 Moscow, Russia²

Received 29 October 2003/ Accepted 24 February 2004

Top Abstract Introduction Materials and Methods Results Discussion References

 
The sll1703 gene, encoding an Arabidopsis homologue of the thylakoid membrane-associated SppA peptidase, was inactivated by interposon mutagenesis in Synechocystis sp. strain PCC 6803. Upon acclimation from a light intensity of 50 to 150 µE m^–2 s^–1, the mutant preserved most of its phycobilisome content, whereas the wild-type strain developed a bleaching phenotype due to the loss of about 40% of its phycobiliproteins. Using in vivo and in vitro experiments, we demonstrate that the sppA1 strain does not undergo the cleavage of the L_R³³ and L_CM⁹⁹ linker proteins that develops in the wild type exposed to increasing light intensities. We conclude that a major contribution to light acclimation under a moderate light regime in cyanobacteria originates from an SppA1-mediated cleavage of phycobilisome linker proteins. Together with changes in gene expression of the major phycobiliproteins, it contributes an additional mechanism aimed at reducing the content in phycobilisome antennae upon acclimation to a higher light intensity.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Light is the principal energy source for all photosynthetic organisms. Pigment-protein complexes, which act as light-harvesting antennae, transfer absorbed light to photochemical reaction centers that convert it into chemical energy utilized by living cells. However, despite its high physiological importance, light energy can also become harmful for cell viability when absorbed in excess, due to the production of radical species that, combined with molecular oxygen, alter macromolecular structures by destroying chemical bonds. Thus, photosynthetic organisms have evolved specific protective and acclimative mechanisms in order to cope with unfavorable environmental conditions. In cyanobacteria the major targets for light access in the cell are the phycobilisome (PBS) antennae. PBS are multimeric peripheral membrane structures that comprise pigmented phycobiliproteins and nonpigmented linker polypeptides. In Synechocystis sp. strain PCC 6803, the major phycobiliproteins, allophycocyanin (APC) and phycocyanin (PC), are retained in a multimeric structure by several types of linker proteins (23). The core-membrane linker, L_CM subunit, is responsible for the energy transfer from PBS to photosystem II (PSII) (12, 25); the rod-core linkers, L_RC, attach the peripheral rods to the core of PBS. In addition, rod linkers, L_R, and core linkers, L_C, are involved in the assembly of rods and core domains of PBS, respectively (35). The major mechanisms underlying light acclimation involve the modulation in size and structure of the PBS, although other changes have also been reported either in the stoichiometry or in the composition of photochemical reaction centers (26, 27) with respect to other components of the electron transfer chain or in the concentration of enzymes for CO₂ fixation (1).

The synthesis, assembly, and membrane binding of PBS structures are controlled by a variety of regulatory mechanisms, operating under different environmental conditions. The amount and composition of PBS are modified with light conditions and nutrient availability (2, 6, 16, 17). Acclimation to higher light intensity occurs primarily through changes in gene expression (4, 18, 22) that result in a decreased number of PBS per cell and in a shortening of PBS rods (18, 30). Other well-studied examples of acclimation are the degradation of PBS during nitrogen, phosphor, and sulfur starvations (29, 41). Screening of cyanobacterial mutants that retained their PBS during sulfur starvation led to the identification of several genes that control PBS degradation in cyanobacteria (9, 34). Among these are nblS, encoding a sensor histidine protein kinase, and nblR, which encodes a response regulator. These represent a two-component regulatory system controlling the expression of a series of factors critical for the acclimation of the photosynthetic apparatus in response to both light and nutrient stresses (34, 39), including nblA, which encodes a protein triggering PBS degradation under nutrient deprivation (7, 21). These last two studies and others (6, 9, 21, 41) showed that changes in the rates of PBS degradation also contribute to a cyanobacterial response to changes in light intensity or nutrient availability. Other studies concurred with the conclusion that peptidases also participate in the posttranslational modification of the PBS antenna (40, 41). However, the identification of proteases involved in PBS degradation or remodeling has not been achieved, with the exception of two peptidase activities from Anabaena that could be involved in heterocyst formation during nitrogen limitation (11). However, the study of an Anabaena gene, encoding a Ca-dependent peptidase, could not demonstrate its actual involvement in PBS degradation (24).

The systematic inactivation of putative genes for peptidase components in Synechocystis sp. strain PCC 6803 revealed four enzymes that could be involved in light acclimation and one in response to nutrient deprivation (36). One of these components, the SppA1 peptidase, is an integral membrane endopeptidase that initiates the degradation of signal peptides in bacteria (5, 28). It was recently identified as a thylakoid membrane-associated protein in Arabidopsis that showed light induction at the transcriptional, translational, and possibly posttranslational levels (20). As do all other bacterial organisms, cyanobacteria express two SppA homologues, SppA1 and SppA2 (36), one of which is described in the present study. We provide evidence for involvement of the SppA1 peptidase in light acclimation of the PBS antenna. Although transfer from light close to saturation for growth (50 µE m^–2 s^–1) to saturating light (150 µE m^–2 s^–1) caused similar decreases in the rate of synthesis of PBS proteins in the wild-type and mutant strains, the latter retained much more phycobiliprotein than the former. We show that this difference results from a defect in cleavage of membrane and rod linkers in the PBS structure from the mutant. These observations support the view that light acclimation involves changes in PBS antennae that are not exclusively due to a decreased expression of those genes that encode PBS components. It also results from a truncation of antenna rods by cleavage of distinct linker polypeptides.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Strains and growth conditions. A wild-type, nonmotile Synechocystis sp. strain, PCC 6803, was obtained from the culture collection of the Department of Genetics at Moscow State University. Wild-type and mutant strains were cultivated in standard BG11 medium (32) under white low light (LL) of 50 µE m^–2 s^–1 at 30°C under constant agitation. For some experiments, cell cultures were bubbled with 3% CO₂. Cell concentrations were routinely measured by determining the optical densities of the cultures at 750 nm (OD₇₅₀) (6). There was no significant change in cell size between the wild type and mutant grown under LL and medium light (ML) conditions (data not shown). In all instances, an OD₇₅₀ of 1.0 corresponded to 1.6 x 10⁸ cells/ml. For light acclimation experiments, cells were grown up to an OD₇₅₀ of 1.0 and then diluted with BG11 to an OD₇₅₀ of 0.5 and transferred to ML (150 µE m^–2 s^–1) for 3 days, except when otherwise indicated. For nitrogen depletion experiments, cells were grown in BG11 medium and then washed and subcultured in nitrate-free medium. Growth media for sppA1 and pVZsppA1-complemented strains were supplemented with 40 µg of kanamycin/ml and 10 µg of chloramphenicol/ml, respectively.

Construction of sppA1 mutant strains. The gene encoding the SppA1 peptidase (sll1703) was amplified from genomic DNA of Synechocystis sp. strain PCC 6803 with primers PrF (5'-GGTTTCGGCTGAGGCAGATC-3') and PrR (5'-GCCTTCGAGGTAAACAATGGC-3'). The resulting PCR product of 890 bp was cloned into pGEM-TEasy (Promega, Mannheim, Germany). A kanamycin resistance cassette from plasmid pUC4K (Pharmacia) was inserted into the Eco811 site of sll1703. This plasmid construct was used for transformation of Synechocystis sp. strain PCC 6803 as described previously (15). Complete segregation was verified by PCR analysis, using primers PrF and PrR and Southern analysis.

The sppA1 gene is a member of a gene cluster, sll1702-sll1703-sll1704, where sll1702 and sll1703 contain four overlapping nucleotides in their termination and start codons. Sll1704 is located downstream of sll1703. We ruled out any pleiotropic or polar effect of sll1703 disruption on upstream sll1702 and downstream sll1704 gene expression by performing Northern analysis of RNA transcripts for sll1704, sll1703, and sll1702 in the wild-type and mutant strains (experiments not shown). Furthermore, in order to assess the specificity of our gene-targeted inactivation, we generated a pVZsppA1-complemented strain of the sppA1 mutant by introducing an autonomously replicating plasmid, pVZ321, carrying the sll1703 gene. The DNA fragment containing the entire coding region of sll1703 was amplified by PCR with primers PrAF (5'-GTTTGGGGATGATTTTGGGCTGG-3') and PrAR (5'-GAAGGCAGTAGTAAATCCCGACCACA-3') and cloned into the pGEM-TEasy vector. This fragment was excised with PvuII and recloned into the SmaI site of a cyanobacterial autonomous replication vector, pVZ321 (43). The resulting plasmid, pVZsppA1, was transferred from Escherichia coli into Synechocystis sp. strain PCC 6803 cells by triparental mating (43). Transconjugants were selected on BG11-containing plates with chloramphenicol. We could exclude the possibility of reverse recombination between a DNA fragment containing the wild-type sequence of sll1703 in the complementation plasmid and the sppA1 Synechocystis genome, based on the homogeneous size of DNA fragments amplified by PCR with external primers sll1807forw (5'-ACCATTGGGCGGCTTGCAAAA-3') and sll1703Rfull (5'-TTAAGGATTAAGAAAATGCCA-3'), which were designed from genome sequences downstream and upstream from PrAF and PrAR, respectively.

Isolation of RNA and Northern analysis. RNA was extracted from the wild type and the sppA1 mutant grown under LL and acclimated for 36 h to ML. RNA was extracted using a TRIZOL-based procedure (GibcoBRL Life technologies) and ToTALLY RNA kit (Ambion, Austin, Tex.) according to the manufacturer's instructions. For Northern blots, 15 µg of RNA was separated by denaturing agarose gel electrophoresis and transferred to a nylon membrane. A gene-specific probe for Northern analysis was obtained after PCR amplification of the coding region of sppA1.

Isolation of thylakoid membranes and PBS. For isolation of thylakoid membrane proteins, cyanobacterium cells were broken with glass beads and resuspended in buffer containing 0.5 M sucrose and 50 mM HEPES-NaOH (pH 7.0). Unbroken cells were removed by centrifugation for 10 min at 1,600 x g. Membrane fractions were separated from soluble fractions by centrifugation at 45,000 x g for 20 min at 4°C. PBS were extracted according to the method of Glazer (12). Cells were disrupted with glass beads and solubilized with 2% Triton X-100, and PBS were subsequently separated by sucrose density gradient ultracentrifugation by using 0.9 M phosphate buffer (pH 7.0). The fraction of intact PBS that formed the lower band in the gradient was diluted in 0.9 M phosphate buffer and centrifuged in a Beckman TL100.3 rotor at 160,000 x g, 4°C, for 5 h. The PBS pellet was dissolved in H₂O and used for in vitro analysis.

SDS-PAGE and immunological analysis. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was performed as described previously (19). Approximately 50 µg of membrane proteins or 3.5 µg of chlorophyll (Chl) per lane was resolved by 12% SDS-PAGE. For samples collected at different light intensities, proteins were normalized with respect to their -ATPase subunit content, which should not be affected by changes between LL and ML. For visualization of proteins, gels were stained with Coomassie blue or silver. For Western analysis, proteins were transferred onto nitrocellulose membranes (38). Immunodetection was performed using the enhanced chemiluminescence system. Antisera against PBS proteins (APC, PC, and linker polypeptides) were kindly provided by A. Grossman (Carnegie Institution, Stanford, Calif.).

Proteolytic assay in vitro. For testing of the "self" proteolytic activity of isolated PBS fractions (in H₂O), the fractions were incubated in a water bath at 37°C under ML (150 µE m^–2 s^–1) for 3 h. As a control, PBS were incubated on ice in the dark. The reaction was stopped by addition of Laemmli loading dye, and samples were analyzed with SDS-PAGE.

Pigment analysis and optical and fluorescence spectroscopy. Chl and PC concentrations were determined as described previously (3). Absorption spectra of thylakoid membranes were measured at room temperature using a UV2401 UV/VISIBLE spectrophotometer (Shimadzu Biotech). Low-temperature fluorescence excitation spectra (77K) were recorded using an LS55 luminescence spectrometer (Perkin-Elmer). Cell suspensions were normalized at a Chl concentration of 2 µg/ml in BG11. After a 10-min incubation in the dark, samples were rapidly frozen in liquid nitrogen. Excitation spectra were collected for an emission wavelength set at 695 nm, i.e., in the PSII emission band (37).

Protein pulse labeling. To probe protein synthesis, intact cells were pulse labeled with L-[³⁵S]methionine (Amersham Biosciences, Freiburg, Germany). Cells were grown in liquid BG11 medium to an OD₇₅₀ of 1.5 and then diluted with BG11 to a Chl concentration of 2 µg/ml (OD₇₅₀ = 0.6). Cells were transferred to ML or kept under LL for the next 12, 24, or 36 h. After each time point, L-[³⁵S]methionine was added to the medium to a final concentration of 2 µCi/ml and cells were incubated for an additional 30 min under the chosen light regime. Translation was arrested with chloramphenicol (150 µg/ml). Samples were analyzed by SDS-PAGE.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Phenotypic and spectroscopic characterization of the sppA1 mutant. We have previously reported that the sppA1 mutant of Synechocystis displays a pattern of light sensitivity distinct from that of the wild-type strain (36). Figure 1A illustrates the phenotypical differences of the sppA1 mutant versus the wild type and the pVZsppA1-complemented strain grown under two light regimes, either 50 µE m^–2 s^–1, hereafter referred to as LL, or 150 µE m^–2 s^–1, referred to as ML. In these particular experiments, cell batches grown under LL conditions to the end of exponential phase were diluted to an OD₇₅₀ of 0.5 and either transferred to ML or kept under LL for another 3 days. As previously reported (34), the transfer of wild-type cells to ML caused a bleaching phenotype (Fig. 1A). In marked contrast, the sppA1 mutant showed no evidence for bleaching under ML. When the mutant strain was complemented with the sppA1 gene, yielding the pVZsppA1 strain (see Materials and Methods for details), the wild-type behavior was restored, i.e., bleaching was again observed under ML. Typical absorption spectra of wild-type cells grown under LL and ML are shown in Fig. 1B (bold line). The blue region (400 to 500 nm) displays a broad absorption peak, with several shoulders that correspond to the Soret region of the Chl a absorption spectrum (440 nm) and to the absorption of various carotenoid bands (above 450 nm). The red region shows two distinct peaks, one centered on 620 nm due to phycobilin-containing proteins, PC and APC, the other around 680 nm corresponding to Chl a. Consistent with the bleaching of the wild-type cultures in ML, their absorption spectrum changed from LL to ML with a significant decrease of both the Chl a and PC/APC absorption bands relative to that of the carotenoids (as can be seen in Fig. 1B, middle panel, by the higher contribution to the overall spectrum of the shoulder at 480 nm). However, the decrease in PC/APC was more pronounced than that of Chl a, leading to a marked change in the ratio of the two absorption peaks in the red region of the spectrum. The absorbance spectrum of the sppA1 mutant was similar to that of the wild type under LL but became markedly different from that of the wild type under ML (Fig. 1B, middle panel). Although a decrease in Chl a absorption relative to that of carotenoids was also observed, the PC/APC absorption peak at 620 nm remained higher for the sppA1 mutant. Consequently, the ratio between the Chl a and PC/APC absorption peaks in the mutant cells under ML remained similar to that under LL. These observations suggest that the wild type adapts to ML by losing more of its PC/APC-containing phycobiliproteins than does the sppA1 mutant. The loss of phycobiliproteins in the wild type of Synechocystis is well documented in another instance, when cells are deprived of nitrogen sources. To check whether SppA1 inactivation also resulted in the preservation of PBS under nitrogen starvation, the wild type and the sppA1 mutant were grown in nitrogen-depleted medium and the loss of PBS was visualized by the loss of the PC/APC absorbance peak in the 620-nm region of the spectrum (Fig. 1B, right panel). The same decrease in the PC/APC absorbance was observed with the two strains, showing that SppA1 plays no part in the loss of PBS under nitrogen starvation conditions.

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FIG. 1. Growth phenotype (A) and absorbance spectra (B) of the wild type (WT) and the sppA1 mutant during acclimation to the ML regime and nitrogen deprivation. Synechocystis cultures were grown under LL to an OD750 of 1.0. Cells were then transferred to ML or kept at LL for the next 3 days. For nitrogen limitations, cells were grown in BG11 medium and then transferred to nitrogen-free BG11 (-N) medium for 3 days as described in Materials and Methods. The spectra were measured on whole cyanobacterial cells. Bold line, wild type; dashed line, sppA1 mutant.

Figure 2 shows the growth rates and changes in pigment content during acclimation of the wild-type and mutant strains to ML for 72 h. The mutant grew faster than the wild type under ML and reached a higher cell density after 3 days of growth (Fig. 2A). The changes in pigment content are given on a per-cell basis (see Materials and Methods for relations between cell number and OD₇₅₀) relative to the initial concentration measured before acclimation (Fig. 2B and C). There was a parallel decrease in Chl content in the two strains over 3 days of growth in ML, whereas the PC content decreased twice as much in the wild type as in the sppA1 mutant. Taken together, the higher cell density and higher PC content explain the darker color of the mutant culture grown for 3 days in ML. Table 1 shows the generation time of the two strains at three light intensities, 20 µE m^–2 s^–1 (dim light), 50 µE m^–2 s^–1 (LL), and 150 µE m^–2 s^–1 (ML). Cell division was observed to be light limited in the 20- to 50-µE m^–2 s^–1 range. This intensity was close to saturation, since a further increase in light intensity by a factor of 3 produced only a moderate increase in growth rates for the two strains. That these growth conditions were not limited by CO₂ availability is demonstrated by the similar generation times observed whether cultures were or were not bubbled with 3% CO₂.

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FIG. 2. Cell growth and pigment analysis of the wild type and the sppA1 mutant strain under the ML regime. (A) Synechocystis cells were grown first under LL till exponential phase, diluted with BG11 medium, and transferred to ML for 72 h. The cell growth (A) and Chl (B) and PC (C) concentrations were measured as percentages per OD750. Bold line, wild type; dashed line, sppA1 mutant.

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TABLE 1. Doubling time of the wild type and the sppA1 mutant strain of Synechocystis sp. strain PCC 6083 at different light intensitiesa

In an attempt to correlate the changes in pigment content and absorption properties with changes in the functional organization of the antennae, we determined 77K excitation spectra of the 695-nm emission peak that originates from PSII cores (13), using cyanobacterial cells adapted for 36 h to either LL or ML conditions. As shown in Fig. 3A, the wild-type excitation spectrum of PSII emission from cells grown under LL displayed a small excitation component in the blue region that corresponds to Chl a from the PSII cores and a major 624-nm component that corresponds to PBS antennae functionally connected to the Chl a-containing PSII cores. These features are seen as well in the sppA1 strain grown under LL. The wild type grown under ML showed a marked decrease in the contribution of PBS sensitization relative to Chl sensitization of the PSII emission (Fig. 3B) that was not observed with the sppA1 mutant grown under ML. Taken together, the absorbance and 77K fluorescence data show that light acclimation, viewed as a decreased PBS-to-Chl ratio in thylakoid membranes of the wild type grown under ML, is prevented in the absence of SppA1.

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FIG. 3. 77K excitation spectra of the wild type and the sppA1 strain with LL and ML regimes. Synechocystis cells of the wild type and the sppA1 mutant were grown in LL and adapted to ML for 36 h. The excitation spectra were recorded for PSII emission at 695 nm. Bold line, wild type; dashed line, sppA1 mutant.

Characterization of supramolecular photosynthetic assemblies. The changes in intracellular pigment content in the wild type and the sppA1 mutant under various light regimes could reflect changes in the amount of thylakoid membranes per cell or changes in the relative content of pigment-binding protein per thylakoid membrane or both. Thylakoid membrane proteins were therefore isolated from wild-type and mutant strains after growth for 3 days with LL or ML, and their protein patterns were compared after SDS-PAGE (data not shown) and immunological analysis (Fig. 4). Thylakoid proteins were immunodetected with antisera raised against the ß subunit of the ATP synthase, the PsaA/B proteins of the PSI reaction center, the D1 protein of PSII, the Rieske FeS protein of the cytochrome b/f complex, and major PBS rod proteins (Fig. 4, PC). No significant differences in the content of ATP synthase and that of the cytochrome b/f complex were observed with various light regimes between the two strains. In contrast, the content in Chl a-containing reaction center proteins of PSI and PSII decreased with exposure to higher light intensities. However, no differences in the relative contents of these protein complexes were detected between the sppA1 mutant and the wild type. In contrast, the content in phycobiliproteins in thylakoid membranes (Fig. 4) and whole-cell extracts (Fig. 5) was markedly different between the wild type and the sppA1 mutant when grown under ML. The amount of PC decreased extensively with increasing light regimes in the wild type (Fig. 4), whereas the sppA1 mutant retained most of its PC/APC content under ML. The preservation of most of the antenna proteins in the mutant strain under the ML regime is consistent with the spectral data. Thus, we conclude that the sppA1 mutant is mainly altered in the acclimation of PBS structures to an increase in light intensity.

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FIG. 4. Biochemical analysis of major photosynthetic complexes. Thylakoid membrane proteins isolated from cyanobacterial cells adapted for 3 days to LL and ML were separated by 12% PAGE. The major photosynthetic proteins were visualized by immunodetection with antisera raised against the ß subunit of ATP synthase, PsaA/B reaction center proteins of PSI, the D1 protein of PSII, the Rieske FeS protein of cytochrome b/f complex, and major bilin-containing proteins of PBS antennae. The proteins were normalized to the content in the ß subunit of ATP synthase. WT, wild type.

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FIG. 5. Evolution of whole-cell content in PC in vivo. Wild-type (WT) and sppA1 strains were grown in LL till mid-log phase and then diluted with BG11 to an OD750 of 0.5 (point 0) and transferred to ML or kept in LL for 36 h. Cells were taken after 12, 24, or 36 h in ML and after 36 h in LL. Total proteins were extracted from cells at the same OD750 and separated by 12% SDS-PAGE. Proteins were visualized by Coomassie blue staining. Coomassie-stained cell proteins from the wild type (A) and visualization of PC and PCß bands during acclimation to ML (B) are shown. The upper part of the panel with - and ß-ATPase subunits was used as a loading control.

Figure 5 shows a kinetic analysis of the changes in cellular content in phycobiliproteins for the wild type and the sppA1 strain grown under ML over a 36-h time period. The whole-protein content from cell cultures acclimated to ML after 12, 24, or 36 h was separated on a 12% SDS-polyacrylamide gel and stained with Coomassie blue. The prominent 16-kDa band in this gel corresponds to PC, with a faint APC band migrating immediately below PC (Fig. 5A). We focused on the changes in these bands upon light acclimation (Fig. 5B). As a loading control, the content in and ß subunits of ATP synthase is also presented. The comigration of these proteins with Coomassie-stained bands was tested immunologically by using anti- and -ß ATP synthase antibodies. No significant changes were detected in the wild type during the first 12 h of acclimation to ML. The decrease in total cell content in PC became visible only after 24 h and further extended up to 36 h, a time period that corresponds to the bleaching phenotype readily visible by the mere observation of the cultures. In contrast, sppA1 cells retained most of the PC subunits after 36 h of exposure to ML. These data proved that the loss of phycobiliproteins under higher light regimes is a delayed and slow process in cyanobacterial cells that requires about 24 h before it can be detected.

It has been previously proposed that the decrease in the size of the PBS which occurs with increasing light intensity was partially due to a down-regulation of the genes coding for the major phycobiliproteins (4, 18). Therefore, we wondered whether the difference in PC/APC content between the sppA1 mutant and the wild type could be attributed to a lack of down-regulation in PC/APC gene expression in the mutant grown under ML. The rates of synthesis of the major bilin-containing proteins were monitored by a pulse-labeling study using L-[³⁵S]methionine under LL and ML conditions (Fig. 6; for details, see Materials and Methods). Most importantly, the wild-type and sppA1 strains displayed the same behavior with respect to changes in light regimes: as previously reported (18, 22), there was a drop in the rate of synthesis of the major PC and APC polypeptides under ML versus LL for the two strains. This change occurred within the first 12 h of transfer to ML conditions, with no further decrease over the next 36 h of acclimation. These data account for the decreased content in PC that remains in the sppA1 mutant (Fig. 2). They led us to exclude that the larger decrease in phycobiliproteins in the wild type when placed under ML conditions is due to a mere translational regulation: the sppA1 mutant undergoes a similar down-regulation of translation, although it preserves a higher phycobiliprotein content under ML.

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FIG. 6. Analysis of protein translation rate in the sppA1 mutant and the wild type under various light regimes by pulse labeling with L-[35S]methionine. Cells were adapted to LL and ML for 12, 24, or 36 h. Cells were labeled with L-[35S]methionine as described in Materials and Methods, after immediate transfer to various light regimes for 30 min (lane 0.5) and after each time point of incubation under ML. The whole-cell proteins were separated by 12% SDS-PAGE, and the gel was fluorographed in a Fuji phosphorimager.

SppA controls a light-dependent cleavage of rod linkers. The amount of linker proteins in PBS bound to the thylakoid membranes in the wild type and mutant adapted to various light conditions for 3 days was then analyzed by an immunological assay (Fig. 7A). The amount of L_R³⁵ was stable in the wild type under two light conditions, while those of L_CM⁹⁹ and L_R³³ strongly decreased upon acclimation to ML. In marked contrast, these two linker proteins remained stable in the mutant strain under ML. These observations are consistent with the loss of membrane-bound APC and PC observed in the wild type but not in the sppA1 mutant and reflect an SppA1-driven loss in phycobiliproteins in Synechocystis during light acclimation to ML intensities.

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FIG. 7. Degradation of linker polypeptides of PBS antennae. (A) Wild-type and sppA1 mutant cells were grown under LL and then transferred to the ML regime for the next 72 h. Thylakoid membrane proteins were separated, transferred onto nitrocellulose membranes, and immunodetected with antisera against the various linker proteins: membrane linker LCM99 and rod linkers LR35 and LR33. Protein loading was normalized to the ß-ATPase subunit as shown in Fig. 4. (B) Gene expression of sppA1 under LL and ML. RNA was extracted from the wild-type and mutant strains grown under LL and then acclimated to ML for 36 h. SppA1 transcripts were identified by hybridization analysis with a gene-specific probe. (C) PBS were isolated from wild-type, pVZsppA1-complemented, and sppA1 cells grown in LL. Isolated PBS were incubated at 4°C in the dark (lanes 1, 3, and 5) and at 37°C under ML (lanes 2, 4, and 6) for 3 h. The reaction was stopped by placing the samples on ice. Proteins were separated by 12% SDS-PAGE. For protein visualization, the gel was silver stained.

The fact that SppA1 controls acclimation of antennae to increased light intensity could be due to a light-dependent expression of the sppA1 gene. This prompted us to probe the transcript pattern for sppA1 in wild-type cells grown under LL before being acclimated (or not) for 36 h to ML (Fig. 7B). We found that the sppA1 transcript was equally expressed under LL and ML regimes. These data indicated that SppA1, if light regulated, is controlled at a (post)translational level.

Early studies with isolated PBS have mentioned that they were subjected to rapid degradation in vitro (33). In an attempt to link these early observations with the present findings, PBS from the wild-type, pVZsppA1-complemented, and sppA1 cells adapted to LL conditions were isolated and subsequently incubated at 4°C in the dark and at 37°C for 3 h under ML conditions (Fig. 7C). A degradation of the membrane (L_CM⁹⁹) and rod (L_R³³) linkers was observed exclusively with PBS preparations from the wild-type and pVZsppA1 strains. L_CM⁹⁹ and L_R³³ linker proteins were stable in the sppA1 mutant. This experiment demonstrates that at least two types of linker peptides, L_CM⁹⁹ and L_R³³, are degraded in isolated PBS fractions. In addition, it shows that PBS isolated from the sppA1 mutant lack the peptidase activity responsible for linker degradation in the wild type. Taken together, the in vitro and in vivo experimental data demonstrate the proteolytic resistance of two major PBS linker polypeptides, L_CM⁹⁹ and L_R³³, in the absence of the SppA1 peptidase.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Under our experimental conditions, the wild-type strain of Synechocystis has a doubling time of about 11 h when grown at a light intensity of 50 µE m^–2 s^–1 under white light. This light intensity is close to saturation, since a switch to 150 µE m^–2 s^–1 produces only a modest decrease in generation time, by about 10%. This generation time is not limited by CO₂ availability, since bubbling CO₂ in the culture does not change the rate of cell division under LL and ML regimes. However, this transition from close-to-saturating light to saturating light was accompanied by a marked bleaching of the cultures that became visible after 24 h of exposure to the new light regime and slowly developed over the next 48 h. This delay in bleaching was observed for initial cell OD₇₅₀s of either 0.1 or 0.5 as well as in culture conditions that were kept at constant cell density by a daily dilution up to an OD₇₅₀ of 1.0. The 40% decrease in phycobilins and Chl after 3 days under 150 µE m^–2 s^–1 was accounted for by the loss of a significant proportion of the PC and APC phycobiliproteins and of the PSI and PSII Chl-binding proteins. The former change reflects an acclimation of Synechocystis to increasing light intensities that has been previously attributed to changes in the expression of genes encoding phycobiliproteins (4, 8).

The sppA1 mutant grows at rates similar to those for the wild type—even a little faster—under the various conditions used in this study. In that respect it is similar to other genetically modified cyanobacterial strains that were previously reported to have higher growth rates than wild-type strains (10, 21). When cultures of the sppA1 mutant were exposed to a 50/150-µE m^–2 s^–1 light transition, we observed the same changes in Chl and photosystem contents and the same drop in synthesis of the major phycobiliproteins as in the wild type. However, the mere comparison of the cultures showed that the pronounced bleaching observed with the wild type was no longer seen with the mutant. Although a minor part of this difference can be ascribed to the higher pigment/cell content and higher cell density reached by the mutant cultures, the major contribution came from the better preservation of phycobiliproteins in sppA1 grown at 150 µE m^–2 s^–1. The higher content in peripheral antenna in the mutant was documented both at the protein level, relative to other photosynthetic proteins, and by a higher PBS sensitization of PSII fluorescence emission at 77K. Because the two strains showed a similar down-regulation of PBS expression at 150 µE m^–2 s^–1, we are bound to conclude that the greater loss of phycobiliproteins in the wild type is due to a degradation process that is hampered in the sppA1 mutant.

The participation of SppA1 in the degradation of phycobiliproteins when reaching light intensities immediately above saturation is specific to this acclimation process. The loss in phycobiliproteins and PBS due to other changes in environmental conditions, such as nitrogen deprivation (this study) or deprivation in S, P, Fe, and Cu (36), remained unaltered in the sppA1 mutant. PBS degradation responses are thus controlled by different proteases depending on the environmental stimulus. They differ in both amplitude and kinetics, with nitrogen depletion leading to a rather rapid decrease of up to 90% of PBS (21), while the drop in intracellular phycobiliproteins reported here, upon acclimation to saturating light, is a delayed process and does not exceed 40%. During nitrogen deprivation, cell division and synthesis of phycobiliproteins stop, while preexisting phycobiliproteins are degraded. Therefore, there is a net loss in phycobiliproteins. During light acclimation, the synthesis of new phycobiliproteins is decreased, and the content in preexisting phycobiliproteins also declines due to a limited and specific degradation process. However, since the cell cultures continue to grow, there still is a relative increase in phycobiliproteins per unit volume although the content per cell decreases. We note that the degradation of LHCII antenna upon acclimation of higher plants to high light conditions is also a slow process, with 30% degradation only, observed after 3 days of acclimation to high light intensities (42).

It was previously demonstrated (31, 33) that linker proteins could be degraded in vitro within isolated PBS by some coisolated proteolytic enzyme(s). Here we confirmed this observation: there was a selective loss of L_R³³ and L_CM⁹⁹ linker proteins in PBS isolated from the wild type when incubated at 37°C for 3 h. However, this linker degradation was no longer observed when using PBS isolated from sppA1. This points to an acclimation process with saturating light regimes that is caused by a linker-targeted and SppA1-mediated degradation process. That linker proteins were degraded in vitro under ML, whereas they remained stable in the dark, demonstrated that the protease can be activated by light, most likely by some conformational change. Indeed, at variance with the SppA1 homologue from Arabidopsis, which is light induced at the transcriptional level (20), we found that sppA1 from Synechocystis is constitutively transcribed. Therefore, the light-dependent regulation of SppA1 proteolytic activity should rather involve some posttranslational modifications of the protease or/and some conformational changes of its protein substrates. We note that an extensive degradation of rod linker L_R³³, but not of the membrane linker, was also reported during nitrogen starvation (21), a process that does not require SppA1. This means that PBS may undergo similar modifications of their structure during degradation through different regulatory mechanisms.

Proteolysis of PBS encompasses a diversity of phenomena from extensive degradation of all PBS subunits, as observed during nitrogen starvation, to some limited modifications in their supramolecular structure due to the selective action of endopeptidases. Our in vitro experiments suggest that such a fine regulation in antenna organization should start with the cleavage of the distal linker protein L_R³³ and the membrane linker L_CM⁹⁹. Decreased energy transfer from PBS to PSII at saturating light intensities can occur through shortening of the PBS rods via a detachment of the external rod segments or of the whole rods from PBS cores and/or through a decreased ratio of PBS to PSII per photosynthetic membrane due to their detachment from the membrane, leading ultimately to their degradation in the cytoplasm. Degradation of L_R³³ and L_CM⁹⁹ can account for both a shortening and a release of PBS from the membranes. The former process should be driven by the loss of L_R³³-PC, which represents the distal chains of the rods. We have shown that this loss is controlled by SppA1, an observation which is consistent with previous reports showing that the regulation of L_R³³ accumulation is not primarily due to transcriptional changes but rather is due to a control at the translational or posttranslational level (8). The release of PBS from the membranes probably involves the other linker, L_CM, which participates in assembling the PBS structure in an energy transfer-competent position towards PSII (12, 25). L_CM linker represents a chimeric protein with a heterogeneous domain structure. The C-terminal part of this protein contains three repeat domains (REP1-3) which show high homology to conserved domains of the rod and rod-core linker polypeptides and provides the binding domains that interact with the APC trimer. Sequencing of a 23-kDa peptide that was associated with an APC (^APß^AP) subcomplex showed that it originated from the C-terminal part of this membrane linker (14), which carried only the last REP domain. This crucial experiment demonstrates that there is a peptidase able to cleave the C-terminal sequence of the L_CM linker, which is tightly interacting with APC. Since there are two copies of L_CM per PBS structure, each APC trimeric cylinder could potentially be detached, leading to a complete dissociation of PBS from the thylakoid membrane. Thus, one would expect that degradation of L_CM should lead to a decrease in the whole-cell content in assembled PBS, an observation which was indeed reported for some cyanobacteria during light acclimation (30).

PBS linker proteins were protected from degradation in the sppA1 strain under ML. This is in favor of SppA1 being the peptidase coisolated with PBS that cleaves the linkers. However, we cannot exclude an indirect role of SppA. For instance, it could control the susceptibility of the PBS structure to another peptidase or regulate the expression of this PBS-targeted peptidase. The study of other intracellular targets for SppA1 should provide a better view of the regulatory function of thylakoid-bound peptidases in cyanobacteria.

 
The work was supported by the Deutsche Forschungsgemeinschaft Grant DFG to A.S. and E.P. (DFG SO448/2), the Russian Foundation for Basic Research (01-04-48081) to V.Z., and the Russian Program of Leading Scientific Institutions to S.V.S. and V.Z.

We are grateful to A. Grossman for the kind gift of antisera against phycobiliproteins. We thank V. M. Glaser for technical help in the preparation of the mutant strains. We also thank R. Bassi, F.-A. Wollman, and R. G. Herrmann for critical reading and discussion of the manuscript.

 
* Corresponding author. Mailing address: Department für Biologie I, Bereich Botanik, Ludwig-Maximilians-Universität, Menzingerstr. 67, 80638 München, Germany. Phone: 49-89-17861242. Fax: 49-89-1782274. E-mail: Anna.Sokolenko{at}lrz.uni-muenchen.de.

Top Abstract Introduction Materials and Methods Results Discussion References

 

1
1. Anderson, J. M., W. S. Chow, and Y. I. Park. 1995. The grand design of photosynthesis: acclimation of the photosynthetic apparatus to environmental cues. Photosynth. Res. 46:129-139.[CrossRef]
2
2. Anderson, L. K., M. C. Rayner, R. M. Sweet, and F. A. Eiserling. 1983. Regulation of Nostoc phycobilisome structure by light and temperature. J. Bacteriol. 155:1407-1416.[Abstract/Free Full Text]
3
3. Arnon, D. I., B. D. McSwain, H. Y. Tsujimoto, and K. Wad. 1974. Photochemical activity and components of membrane preparations from blue-green algae. I. Coexistence of two photosystems in relation to chlorophyll a and removal of phycocyanin. Biochim. Biophys. Acta 357:231-245.[Medline]
4
4. Belknap, W. R., and R. Haselkorn. 1987. Cloning and light regulation of expression of the phycocyanin operon of the cyanobacterium Anabaena. EMBO J. 6:871-884.[Medline]
5
5. Bolhuis, A., A. Matzen, H. L. Hyyrylainen, V. P. Kontinen, R. Meima, J. Chapuis, G. Venema, S. Bron, R. Freudl, and J. M. van Dijl. 1999. Signal peptide peptidase- and ClpP-like proteins of Bacillus subtilis required for efficient translocation and processing of secretory proteins. J. Biol. Chem. 274:24585-24592.[Abstract/Free Full Text]
6
6. Collier, J. L., and A. R. Grossman. 1992. Chlorosis induced by nutrient deprivation in Synechococcus sp. strain PCC 7942: not all bleaching is the same. J. Bacteriol. 174:4718-4726.[Abstract/Free Full Text]
7
7. Collier, J. L., and A. R. Grossman. 1994. A small polypeptide triggers complete degradation of light-harvesting phycobiliproteins in nutrient-deprived cyanobacteria. EMBO J. 13:1039-1047.[Medline]
8
8. de Lorimier, R. M., R. L. Smith, and S. E. Stevens, Jr. 1991. Regulation of phycobilisomes structure and gene expression by light intensity. Plant Physiol. 98:1003-1010.
9
9. Dolganov, N., and A. R. Grossman. 1999. A polypeptide with similarity to phycocyanin alpha-subunit phycocyanobilin lyase involved in degradation of phycobilisomes. J. Bacteriol. 181:610-617.[Abstract/Free Full Text]
10
10. Emlyn-Jones, D., M. K. Ashby, and C. W. Mullineaux. 1999. A gene required for the regulation of photosynthetic light harvesting in the cyanobacterium Synechocystis 6803. Mol. Microbiol. 33:1050-1058.[CrossRef][Medline]
11
11. Foulds, I. J., and N. G. Carr. 1977. A proteolytic enzyme degrading phycocyanin. FEMS Microbiol. Lett. 2:117-119.
12
12. Glazer, A. N. 1988. Phycobiliproteins. Methods Enzymol. 167:291-303.[Medline]
13
13. Goedheer, J. C. 1964. Fluorescence bands and chlorophyll a forms. Biochim. Biophys. Acta 88:304-317.[Medline]
14
14. Gottschalk, L., F. Lottspeich, and H. Scheer. 1994. Reconstitution of an allophycocyanin trimer complex containing the C-terminal 21-23 kDa domain of the core-membrane linker polypeptide L_CM. Z. Naturforsch. 49c:331-336.
15
15. Grigorieva, G. A., and S. V. Shestakov. 1982. Transformation in the cyanobacterium Synechocystis PCC 6803. FEMS Microbiol. Lett. 13:367-370.[CrossRef]
16
16. Grossman, A. R., M. R. Schaefer, G. G. Chiang, and J. L. Collier. 1993. The phycobilisome, a light-harvesting complex responsive to environmental conditions. Microbiol. Rev. 57:725-749.[Abstract/Free Full Text]
17
17. Grossman A. R., M. R. Schaefer, G. G. Chiang, and J. L. Collier. 1994. The responses of cyanobacteria to environmental conditions: light and nutrients, p. 641-675. In D. A. Bryant (ed.), The molecular biology of cyanobacteria. Kluwer Academic Publishing, Dordrecht, The Netherlands.
18
18. Kalla, R., R. P. Bhalerao, and P. Gustaffson. 1993. Regulation of phycobilisome rod proteins and mRNA at different light intensities in the cyanobacterium Synechococcus 6301. Gene 126:77-83.[CrossRef][Medline]
19
19. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685.[CrossRef][Medline]
20
20. Lensch, M., R. G. Herrmann, and A. Sokolenko. 2001. Identification and characterization of SppA, a novel light-inducible chloroplast protease complex associated with thylakoid membranes. J. Biol. Chem. 276:33645-33651.[Abstract/Free Full Text]
21
21. Li, H., and L. A. Sherman. 2002. Characterization of Synechocystis sp. strain PCC 6803 and nbl mutants under nitrogen-deficient conditions. Arch. Microbiol. 178:256-266.[CrossRef][Medline]
22
22. Lomax, T. L., P. B. Conley, J. Schilling, and A. R. Grossman. 1987. Isolation and characterization of light-regulated phycobilisome linker polypeptide genes and their transcription as a polycistronic mRNA. J. Bacteriol. 169:2675-2684.[Abstract/Free Full Text]
23
23. Lundell, D. J., R. C. Williams, and A. N. Glazer. 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.[Abstract/Free Full Text]
24
24. Maldener, I., W. Lockau, Y. P. Cai, and C. P. Wolk. 1991. Calcium-dependent protease of the cyanobacterium Anabaena: molecular cloning and expression of the gene in Escherichia coli, sequencing and site-directed mutagenesis. Mol. Gen. Genet. 225:113-120.[Medline]
25
25. Manodori, A., M. Alhadeff, A. N. Glazer, and A. Melis. 1985. Phycobilisome-photosystem II association in Synechococcus 6301 (Cyanophycea). FEBS Lett. 181:79-82.[CrossRef]
26
26. Melis, A., C. W. Mullineaux, and J. F. Allen. 1989. Acclimation of the photosynthetic apparatus to photosystem I or photosystem II light: evidence from quantum yield measurements and fluorescence spectroscopy of cyanobacterial cells. Z. Naturforsch. 44c:109-118.
27
27. Murakami, A., and Y. Fujita. 1991. Regulation of photosystem stoichiometry in the photosynthetic system of the cyanophyte Synechocystis PCC 6714 in response to light intensity. Plant Cell Physiol. 32:223-230.[Abstract/Free Full Text]
28
28. Novak, P., and I. K. Dev. 1988. Degradation of a signal peptide by protease IV and oligopeptidase A. J. Bacteriol. 170:5067-5075.[Abstract/Free Full Text]
29
29. Ownby, J. D., M. Shannahan, and E. Hood. 1979. Protein synthesis and degradation in Anabaena during nitrogen starvation. J. Gen. Microbiol. 110:255-261.
30
30. Raps, S., J. H. Kycia, M. C. Ledbetter, and H. W. Siegelman. 1985. Light intensity adaptation and phycobilisome composition of Mycrocystis aeruginosa. Plant Physiol. 79:983-987.[Abstract/Free Full Text]
31
31. Reuter, W., and C. Nickel-Reuter. 1993. Molecular assembly of the phycobilisomes from the cyanobacterium Mastigocladus laminosus. J. Photochem. Photobiol. B Biol. 18:51-66.[CrossRef]
32
32. Rippka, R. 1988. Isolation and purification of cyanobacteria. Methods Enzymol. 167:3-27.[Medline]
33
33. Rusckowski, M., and B. A. Zilinskas. 1982. Allophycocyanin I and the 95 kilodalton polypeptide. The bridge between phycobilisomes and membranes. Plant Physiol. 70:1055-1059.[Abstract/Free Full Text]
34
34. Schwarz, R., and A. R. Grossman. 1998. A response regulator of cyanobacteria integrates diverse environmental signals and is critical for survival under extreme conditions. Proc. Natl. Acad. Sci. USA 95:11008-11013.[Abstract/Free Full Text]
35
35. Sidler, W. A. 1994. Phycobilisome and phycobilisome structures, p. 139-216. In D. A. Bryant (ed.), The molecular biology of cyanobacteria. Kluwer Academic Publishers, Dordrecht, The Netherlands.
36
36. Sokolenko, A., E. Pojidaeva, V. Zinchenko, V. Panichkin, V. M. Glaser, R. G. Herrmann, and S. V. Shestakov. 2002. The gene complement for proteolysis in the cyanobacterium Synechocystis sp. PCC 6803 and Arabidopsis thaliana chloroplasts. Curr. Genet. 41:291-310.[CrossRef][Medline]
37
37. Su, X., P. G. Fraenkel, and L. Bogorad. 1992. Excitation energy transfer from phycocyanin to chlorophyll in an apcA-defective mutant of Synechocystis sp. PCC 6803. J. Biol. Chem. 267:22944-22950.[Abstract/Free Full Text]
38
38. Towbin, H., T. Staehelin, and J. Gordon. 1979. Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. Proc. Natl. Acad. Sci. USA 76:4350-4354.[Abstract/Free Full Text]
39
39. van Waasbergen, L. G., N. Dolganov, and A. R. Grossman. 2002. nblS, a gene involved in controlling photosynthesis-related gene expression during high light and nutrient stress in Synechococcus elongatus PCC 7942. J. Bacteriol. 184:2481-2490.[Abstract/Free Full Text]
40
40. Wood, N. B., and T. Haselkorn. 1980. Control of phycobiliprotein proteolysis and heterocyst differentiation in Anabaena. J. Bacteriol. 141:1375-1385.[Abstract/Free Full Text]
41
41. Yamanaka, G., and A. N. Glazer. 1980. Dynamic aspects of phycobilisome structure. Phycobilisome turnover during nitrogen starvation in Synechococ-cus sp. Arch. Microbiol. 124:39-47.[CrossRef]
42
42. Yang, D.-H., J. Webster, Z. Adam, M. Lindahl, and B. Andersson. 1998. Induction of acclimative proteolysis of the light-harvesting chlorophyll a/b protein of photosystem II in response to elevated light intensities. Plant Physiol. 118:827-834.[Abstract/Free Full Text]
43
43. Zinchenko, V. V., I. V. Piven, V. A. Melnik, and S. V. Shestakov. 1999. Vectors for the complementation analysis of cyanobacterial mutants. Mol. Genet. 35:228-232.

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Journal of Bacteriology, June 2004, p. 3991-3999, Vol. 186, No. 12
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.12.3991-3999.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

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