Photosynth Res (2015) 124:217–229 DOI 10.1007/s11120-015-0122-0
REGULAR PAPER
Characterization of a Synechocystis sp. PCC 6803 double mutant lacking the CyanoP and Ycf48 proteins of Photosystem II Simon A. Jackson1 • Julian J. Eaton-Rye1
Received: 23 October 2014 / Accepted: 12 March 2015 / Published online: 24 March 2015 Ó Springer Science+Business Media Dordrecht 2015
Abstract Homologs of the Photosystem II (PS II) subunit PsbP are found in plants, algae, and cyanobacteria. In higher plants, PsbP is associated with mature PS II centers, but in cyanobacteria, the homologous CyanoP protein appears sub-stoichiometric to PS II. We have investigated the role of CyanoP by characterizing knockout mutants of the cyanobacterium Synechocystis sp. PCC 6803. Removal of CyanoP resulted in changes to phycobilisome coupling and energy transfer to PS II, but the function of PS II itself remained similar to wild type. We therefore investigated the hypothesis that CyanoP is involved in the biogenesis or repair of PS II by creating a double mutant lacking both CyanoP and the PS II assembly factor Ycf48. This strain exhibited an additive reduction in the amplitude of variable chlorophyll a fluorescence induction relative to either of the single mutants but displayed increased oxygen evolution, slight increases in PS II monomer and dimer levels, and a reduction in accumulation of an early PS II assembly complex containing CP47, compared to the DYcf48 strain. Keywords Assembly Biogenesis CyanoP Photosystem II PsbP Synechocystis sp. PCC 6803 Ycf48 Abbreviations Bis–Tris 2-[Bis(2-hydroxyethyl)amino]-2(hydroxymethyl)propane-1,3-diol BMF Blue measuring flashes
BN-PAGE CP43 CP47 D1 D2 DCMU FO FM J and I LC/MS/MS LHC O OD P PS I PS II QA QB
& Julian J. Eaton-Rye [email protected] 1
Department of Biochemistry, University of Otago, Dunedin 9016, New Zealand
RC47 RMF S-states
Blue-native polyacrylamide gel electrophoresis 43-kDa chlorophyll a-binding protein of the core antenna 47-kDa chlorophyll a-binding protein of the core antenna Photosystem II reaction center protein subunit Photosystem II reaction center protein subunit 3-(3,4-Dichlorophenyl)-1,1-dimethylurea Dark-adapted (minimum) fluorescence Maximum level of chlorophyll a fluorescence Inflection points between O (FO) and P in the chlorophyll a fluorescence induction curve Liquid chromatography tandem mass spectrometry Light-harvesting complex Origin (FO) level of the chlorophyll a fluorescence induction curve Optical density Peak level in the chlorophyll a fluorescence induction curve Photosystem I Photosystem II Primary plastoquinone electron acceptor of PS II Secondary plastoquinone electron acceptor of PS II Reaction center complex lacking the CP43 protein Red measuring flashes Oxidation states of the oxygen-evolving complex of PS II
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SDS-PAGE Tris
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Sodium dodecyl sulfate polyacrylamide gel electrophoresis 2-Amino-2-hydroxymethyl-propane-1,3-diol
Introduction Photosystem II (PS II) is found in all oxygenic photosynthetic organisms where it is required for the light-driven oxidation of water (Vinyard et al. 2013). The biogenesis of PS II is a multistep process which involves the coordinated assembly of several smaller intermediate complexes to form the mature photosystem (Komenda et al. 2012). The early assembly intermediates include a pre-complex containing the reaction center D1 and D2 proteins and two additional pre-complexes, each containing one of the chlorophyll a-binding core antenna proteins CP43 or CP47. All of these pre-complexes also contain additional protein subunits that are either membranespanning or soluble proteins associated with the exposed hydrophilic domains on both the lumenal and cytosolic faces of the core D1, D2, CP43, or CP47 subunits (Komenda et al. 2012). Many of the proteins found in these intermediate complexes are not found in the mature complex but function as assembly factors or chaperones that are transiently associated with PS II during de novo biogenesis or the PS II repair pathway that operates following light-induced photodamage (Nixon et al. 2010; Nickelsen and Rengstl 2013). In cyanobacteria, a group of three extrinsic PS II subunits: CyanoP, CyanoQ, and Psb27, all possessing an N-terminal lipid modification, are believed to associate with the lumenal face of PS II (Fagerlund and Eaton-Rye 2011). All three subunits are absent from the most recent high resolution crystal structures of PS II (Umena et al. 2011; Suga et al. 2015), but their structures have been determined independently (Cormann et al. 2009; Mabbitt et al. 2009; Jackson et al. 2010; Michoux et al. 2010; Umena et al. 2011; Jackson et al. 2012; Michoux et al. 2014). The CyanoQ and Psb27 subunits have been assigned roles in mature PS II centers and early PS II assembly intermediates, respectively (reviewed in (Bricker et al. (2012) and Mabbitt et al. (2014)). In contrast, any interaction or function of the CyanoP protein in PS II biogenesis, mature centers, or repair processes remains undetermined; however, the previously reported sub-stoichiometric relationship between the levels of intracellular CyanoP and PS II centers suggests that CyanoP might play a role in either de novo biogenesis or repair (Thornton et al. 2004). The Psb27 protein is known to be involved in both biogenesis and repair of PS II, and we recently reported the phenotypic characterization of Synechocystis sp. PCC 6803 (hereafter Synechocystis 6803) strains lacking Psb27 in the
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wild-type background and a mutant background which lacked the PS II assembly factor Ycf48 (Jackson et al. 2014). Ycf48 has been shown to be associated with early assembly complexes of PS II (Komenda et al. 2008) and the additive phenotypes we observed, for the DYcf48:DPsb27 strain compared to the corresponding single-deletion mutants, led to the conclusion that Psb27 and Ycf48 are associated with complexes that interact during the biogenesis or repair cycles (Jackson et al. 2014). In this communication, we test the hypothesis that if CyanoP functions during biogenesis or repair of PS II, then the function of this subunit might also be better revealed when PS II assembly is perturbed by removal of Ycf48.
Materials and methods Cyanobacteria strains and physiology experiments The Synechocystis 6803 strain referred to throughout as wild type is a sub-cultured derivative of the Williams glucose-tolerant strain, designated GT-O1 (Williams 1988; Morris et al. 2014). The strains lacking CyanoP (DCyanoP) and Ycf48 (DYcf48) have been described previously (Summerfield et al. 2005; Jackson et al. 2014). For the double knockout, the ycf48 ORF was inactivated in the DCyanoP background, producing the DCyanoP:DYcf48 mutant. Transformations were performed and strains were maintained as described in Eaton-Rye (2011). All physiological characterizations and experiments were performed as described in Jackson et al. (2014) with the alterations detailed hereafter. Whole-cell absorption spectra were collected using an Evolution 201 UV/Vis spectrophotometer (Thermo Scientific, Waltham, MA, USA). The flashinduced oscillation pattern of oxygen production following single-turnover actinic flashes was measured using a bare platinum Joliot-type electrode based on the designs of Joliot and Joliot (1968), Meunier and Popovic (1988), Messinger et al. (1993), and Renger and Hanssum (2009). The light source employed was a cluster of light-emitting diodes with an emission peak at 627 nm (Philips, Andover, MA, USA) and a custom built driver unit with variable flash-width capability. Two microliters of sample at a chlorophyll a concentration of 1 mg mL-1 in BG-11 (pH 7.5) was loaded onto the electrode, then dark adapted for 5 min, given a singleturnover flash, and dark adapted for a further 5 min before commencement of a train of 25 flashes (32 ls duration) given with a spacing between flashes of 250 ms. Analyses of thylakoid membrane proteins Thylakoid membrane samples were solubilized using bdodecyl maltoside as per Jackson et al. (2014). For Western
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blotting, 2 lg of chlorophyll a equivalent material was loaded per sample on NativePAGE 3–12 % Bis–Tris gradient gels (Life Technologies, Waltham, MA, USA). Electroblot transfer was performed at 50 V for 16 h in buffer containing 25 mM Tris, 192 mM glycine, and 20 % v/v methanol. The primary antibodies for the PsbA (D1), PsbC (CP43), and PsbD (D2) subunits of PS II and the PsaA subunit of PS I were obtained from Agrisera, Va¨nna¨s, Sweden. For 2D gel electrophoresis NativePAGE, 4–16 % Bis–Tris gradient gels (Life Technologies, Waltham, MA, USA) were run as the first dimension, followed by NuPAGE 12 % Bis-Tris gels (Life Technologies, Waltham, MA, USA) as the second (SDS-PAGE) dimension. The SDS-PAGE gels were stained with SYPRO Ruby (Life Technologies, Waltham, MA, USA), imaged with a 16-bit linear CCD detector system (Fuji imager PS3000, Fujifilm, Tokyo, Japan) operating in fluorescence mode (460 nm excitation, Y515 emission filter), de-stained and then restained with colloidal Coomassie Brilliant Blue G-250. For the false-colored overlay comparisons, all SYPRO Rubystained gels were imaged using identical exposure times, and the intensity data are displayed as two linear channels (red or green, for each strain as indicated) on a composite image with identical contrast and brightness settings applied to all channels. For mass spectrometry analyses, the gel spots of interest were excised and digested in-gel with trypsin. The resulting peptides were eluted from the gel, dried using a centrifugal concentrator, and reconstituted in an aqueous solution of 5 % v/v acetonitrile and 0.2 % v/v formic acid. The samples were then subject to liquid chromatography tandem mass spectrometry (LC/MS/MS) using a nanoflow uHPLCcoupled TripleTOF 5600? mass spectrometry system (Ab Sciex, Framingham, MA, USA). Samples were separated using a C-18 emitter-tip column operated with a 5–90 % v/v acetonitrile gradient in 0.2 % formic acid. For precursor ion measurement, a mass range between m/z 400–2000 was used, followed by 10 data-dependent collision-induced dissociation (CID) fragment ion measurements per cycle. PeakView (Ab Sciex, Framingham, MA, USA) was used to process the data and generate peak lists that were searched using an in-house Mascot server (Matrix Science, Boston, MA, USA).
Results Photoautotrophic growth, oxygen evolution, and pigment composition Under the conditions used, removal of CyanoP did not significantly influence the photoautotrophic growth rate (doubling time of 11.0 h) compared to wild type (11.5 h)
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(Fig. 1a), whereas inactivation of the ycf48 ORF impaired photoautotrophic growth, increasing the doubling time to 21.4 h for the DYcf48 strain. However, removal of Ycf48 from the DCyanoP strain did not result in an additive impairment in growth beyond that already imposed by inactivation of the ycf48 ORF; the doubling time of the DCyanoP:DYcf48 strain was 19.7 h. The saturated rate of steady-state oxygen evolution was found to be comparable for wild type and the DCyanoP cells (555 and 556 lmol O2 per mg chlorophyll a per h, respectively) (Table 1). Consistent with the photoautotrophic growth data, removal of Ycf48 in the wild-type background reduced the rate of oxygen evolution to approximately 43 %, and removal of both CyanoP and Ycf48 resulted in 48 % activity compared to wild type. The level of photoinactivation observed during the time course of the assay for the DYcf48 strain (as evident by the curvature of the trace during the light exposure) was not influenced by the presence or absence of the CyanoP subunit, suggesting that CyanoP is not involved in photoprotection (Fig. 1b). The previously characterized DYcf48:DPsb27 strain (Jackson et al. 2014) exhibited changes in the cellular pigment composition, notably large increases in the carotenoid absorption region and suppression of the 680-nm peak that originates from absorption by chlorophyll a. Upon investigation of the DCyanoP and DCyanoP:DYcf48 strains, we identified a similar effect, with the 680-nm peak lower for the DCyanoP:DYcf48 double mutant than observed for the DYcf48 strain (Fig. 1c–d). Low temperature (77 K) chlorophyll a fluorescence emission Using an excitation beam that specifically targets chlorophyll a (wavelength of 440 nm), we measured low temperature (77 K) fluorescence emission spectra from each of the strains (Fig. 2a, b). A general overview of interpretation of the emission spectrum for wild type is that the 685 and 695 nm maxima originate from the CP43 and CP47 subunits of PS II, respectively, whilst the maximum PS I emission occurs at 725 nm; however, for PS II assembly mutants, the 685 nm peak might also contain contributions from other species (e.g., Boehm et al. (2011) and Jackson et al. (2014)). Inactivation of the ycf48 gene in the wildtype background resulted in an apparent reduction in the PS II to PS I ratio and an increase in emission at 685 relative to 695 nm, in agreement with our previous observations (Jackson et al. 2014). For the DCyanoP strain, the emission spectra showed a small reduction in the PS II to PS I ratio compared to wild type. Inactivation of the ycf48 gene in the DCyanoP mutant background did not result in a decrease in the PS II to PS I ratio (as observed for the DYcf48 mutant versus wild type), but a comparable increase in the
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b
3.2
Oxygen evolved (μmol per mg Chl a)
a
1.6
OD730
0.8 0.4
0.2 0.1 0.05
Light off (i) (iii)
25 20 15
(ii) (iv)
10 5 0 Light on
-5 0
24
48
72
96
120
0
60
Time (h)
120
180
240
300
Time (s)
d
c
Normalized absorbance
1.0 0.8 0.6 0.4 0.2
1.0 0.8 0.6 0.4 0.2
0.0
0.0 400
500
600
700
Wavelength (nm)
Table 1 Oxygen evolution rates (lmol O2 mg Chl a wild type
555 ± 29
DYcf48
236 ± 32
DCyanoP
556 ± 38
DCyanoP:DYcf48
264 ± 18
-1
-1
h )
Oxygen evolution was measured in the presence of 2,6-dichloro-1,4benzoquinone and K3Fe(CN)6 as described in Jackson et al. (2014)
amplitude of the 685 nm compared to 695 nm emission peak was observed. The slightly higher PS II to PS I ratio for the DCyanoP:DYcf48 strain, compared to DYcf48 cells, correlated with the *10 % higher rate of oxygen evolution observed for this strain (Table 1). Examination of 77 K fluorescence emission using an excitation wavelength targeting phycobilisome pigments (580 nm) can reveal differences in phycobilisome-coupled energy distribution (Fig. 2c, d). The DYcf48 strain
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0.025
Normalized absorbance
Fig. 1 Photoautotrophic growth, oxygen evolution, and whole-cell absorption spectra. a Photoautotrophic growth curve determined by light scattering at 730 nm: wild type (solid squares), DYcf48 (open squares), DCyanoP (solid circles), and DCyanoP:DYcf48 (open circles). Error bars represent the standard error from at least 3 independent experiments. b Oxygen evolution traces: wild type (i), DYcf48 (ii), DCyanoP (iii), and DCyanoP:DYcf48 (iv). c Whole-cell absorption spectra: wild type (solid line) and DYcf48 (dotted line). d Wholecell absorption spectra: DCyanoP (solid line) and DCyanoP:DYcf48 (dotted line). In c and d, spectra are the average of three independent experiments and are normalized to the maxima at 435 nm
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400
500
600
700
Wavelength (nm)
exhibited a characteristic increase in the emission peak at 680 nm compared to wild type. The 680 nm maximum originates from phycobilisome emission, but PS II-specific fluorophores give this peak a red-shifted shoulder, thus resolution of the underlying components is difficult. The DCyanoP strain showed a marked reduction in the 680 nm emission compared to wild type, but in the case of the DCyanoP:DYcf48 strain, removal of Ycf48 in the DCyanoP mutant background resulted in a phenotype whereby the 680 nm peak increased to a level similar to that observed for the DYcf48 strain. Variable chlorophyll a fluorescence measurements Measurements of room temperature variable chlorophyll a fluorescence were used to further investigate phycobilisome to PS II coupling and also examine electron transfer on the acceptor side of PS II. Our instrument is equipped to probe the state of PS II centers using either blue measuring flashes (BMF) for direct excitation of chlorophyll a or red
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a
1
Normalised fluorescence
1.0
Normalized fluorescence
Fig. 2 77 K fluorescence emission spectra. a 440 nm excitation for wild type (solid line) and DYcf48 (dotted line). b 440 nm excitation for DCyanoP (solid line) and DCyanoP:DYcf48 (dotted line). c 580 nm excitation for wild type (solid line) and DYcf48 (dotted line). d 580 nm excitation for DCyanoP (solid line) and DCyanoP:DYcf48 (dotted line). Spectra are the average of at least three independent experiments and are normalized to the PS I emission maxima
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0.8
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4.0 625
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0 775 625 4
675
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725
Wavelength (nm)
measuring flashes (RMF) for assessment of phycobilisomecoupled fluorescence emission. Comparison of data obtained using either BMF or RMF provides information on energy partitioning to complement the 77 K fluorescence emission spectra. The amplitudes of fluorescence presented herein are arbitrary and specific to the optical configuration of the fluorometer used. In addition, the relative amplitudes of data obtained with BMF versus RMF are not directly comparable because these are dependent on the measuring flash intensity, duration, and detector gain parameters. The amplitudes of fluorescence emission during the induction time course were significantly decreased for the DYcf48 strain relative to wild type when measured using either BMF or RMF (Fig. 3a–d) (Jackson et al. 2014). In contrast, inactivation of cyanoP (sll1418) in the wild-type background did not significantly affect the shape or amplitude of the fluorescence induction traces. Also shown in Fig. 3, the DCyanoP:DYcf48 double mutant exhibited a
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d Wavelength (nm)
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Wavelength (nm) Normalised fluorescence
Normalized fluorescence
c
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3
2
1
0 775 625
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Wavelength (nm)
reduction in the maximal amplitudes of fluorescence observed compared to the DYcf48 strain but retained the O, J, I, and P features of the fluorescence induction curve (Strasser et al. 1995). This is in contrast to the DYcf48:DPsb27 strain, where as well as a reduction in amplitude, we discovered loss of the J-P rise that is normally observed in the absence of 3-(3,4-dichlorophenyl)-1,1-dimethylurea) DCMU (Jackson et al. 2014). The amplitude of the fluorescence in the P region changes with addition of DCMU. This is most likely because DCMU blocks electron flow to the plastoquinone pool and influences the redox potential of the acceptor side. For wild type or the DCyanoP strain, the fluorescence yield was similar or reduced in the presence of DCMU, whilst in the DYcf48 and DCyanoP:DYcf48 strains, the yield was increased by addition of DCMU. A mechanistic interpretation for this observation has yet to be elucidated, but it might result from centers with a perturbed acceptor side formed in the absence of Ycf48 (Jackson et al. 2014).
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- DCMU
+ DCMU
1
1.0
0.6
0.4 0.2 0.0 1
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10-3
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0.1 Time (s)
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-5 -4 -3 -2 0.1 10 1.2 10 10 10 10 Time (s)
f
1.0
Fluorescence (F-F0)/(FM-F0)
Fluorescence (F-FO)/(FM-FO)
0.2
0 10-5
e 0.8 0.6 0.4 0.2 0.0
10-5 10-4 10-3 10-2 0.1 Time (s)
Four BMF or RMF were used to determine the background dark-state fluorescence (FO) (Table 2), prior to the start of the activation of actinic illumination of the fluorescence induction traces. No appreciable difference between the FO for all strains was observed using BMF, but when using RMF, the FO level was increased in DYcf48 cells and substantially decreased in the
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1
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0 -5 -4 -3 -2 0.1 10 0.8 10 10 10 10 Time (s) d Fluorescence (F-F0) a.u.
-5 -4 -3 -2 0.1 0.8 10 10 10 10 Time (s) c
0.8
Red measuring flashes (RMF)
0.8
Blue measuring flashes (BMF)
b Fluorescence (F-F0) a.u.
Fluorescence (F-FO) a.u.
a
Fluorescence (F-FO) a.u.
Fig. 3 Room temperature fluorescence induction and relaxation of fluorescence following single-turnover actinic flashes. a–d Room temperature fluorescence induction observed upon illumination of dark-adapted cells with a constant actinic light. Strains are wild type (solid), DYcf48 (dots), DCyanoP (dashes), and DCyanoP:DYcf48 (dash-dots) in the absence (a, c) and presence (b, d) of DCMU. In all instances, constant blue (455 nm) illumination was used with either blue (a, b) or red (625 nm) (c, d) measuring flashes. The time scale is relative to the commencement of illumination. e Relaxation of fluorescence following a singleturnover actinic flash in the absence of DCMU. f Relaxation of fluorescence following a single-turnover actinic flash in the presence of DCMU. In e and f, the strains are wild type (solid), DYcf48 (dots), DCyanoP (dashes), and DCyanoP:DYcf48 (dash-dots). Data displayed are for the average of at least three independent experiments
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1.0 0.8 0.6 0.4 0.2
0.0 10 10-5 10-4 10-3 10-2 0.1 Time (s)
DCyanoP strain. Additional removal of Ycf48 from DCyanoP cells reversed the reduction, resulting in a FO (RMF) similar to the DYcf48 mutant. This mirrors the trend observed in the 77 K emission spectra obtained using a 580 nm excitation beam where lack of Ycf48 resulted in a similar phenotype for both DYcf48 and DCyanoP:DYcf48 cells.
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The oxidation state of QA following a single-turnover saturating blue flash (455 nm), in the absence and presence of DCMU, was investigated by probing the relaxation of chlorophyll a fluorescence emission using BMF (Fig. 3e, f). The fluorescence traces were analyzed by applying a Joliot correction and modeling two exponential and one hyperbolic decays in the absence of DCMU and one exponential and one hyperbolic in the presence of DCMU (Joliot and Joliot 1964; Cser and Vass 2007). The QA oxidation kinetics derived from this analysis are summarized in Table 3. In the absence of DCMU, the transfer kinetics of the DCyanoP strain were similar to those observed in wild type, indicating that the absence of CyanoP does not influence electron transfer on the acceptor side of PS II. The Ycf48 knockout in DYcf48 cells, however, resulted in a decrease in the rate of electron transfer from QA to QB for centers where the QB site was likely empty at the time of the actinic flash (middle phase). An increase in the rate and amplitude for the slow phase was also observed, suggestive of increased recombination via back reaction with the oxygen-evolving complex (Jackson et al. 2014). In the DCyanoP:DYcf48 double mutant, these same effects were observed, consistent with CyanoP not influencing the acceptor-side electron transfer kinetics, and removal of Ycf48 eliciting the primary influence in the rates and amplitudes observed. In the presence of DCMU, oxidation of QA in wild-type cells was dominated by the back reaction (slow phase) with a rate constant of 1.2 s (95.7 %) and a minor fast phase component (4.3 %) (Table 3). The DCyanoP strain exhibited similar back reaction characteristics to wild type, indicating that neither the donor nor acceptor sides of PS II are destabilized by the absence of CyanoP. In the absence of Ycf48, the fast phase increased in rate (half time of 4.1 ms) and almost doubled in amplitude (to 7.4 %), observable in the normalized fluorescence decay trace (Fig. 3f) (Jackson et al. 2014). The DCyanoP:DYcf48 strain showed only minor changes in these kinetics compared to those observed in DYcf48 cells (Table 3), suggesting that the impairment of acceptor-side function in the absence of Ycf48 is not influenced by the presence or absence of CyanoP. Table 2 Dark-adapted background (FO) fluorescence data BMF (a.u.)
RMF (a.u.)
wild type
0.504 ± 0.021
0.725 ± 0.025
DYcf48
0.536 ± 0.030
0.805 ± 0.085
DCyanoP
0.446 ± 0.025
0.549 ± 0.047
DCyanoP:DYcf48
0.544 ± 0.040
0.849 ± 0.136
The amplitudes between BMF and RMF are not directly comparable. Data shown were collected in the absence of DCMU. No significant difference was observed in its presence
Flash-induced oxygen evolution Efficient acceptor-side electron transfer and plastoquinone exchange are necessary to support full activity of the PS II complex, including maintaining function of the donor side and oxygen evolution. To investigate whether, in the absence of Ycf48, donor side function is impacted by the impairment of the acceptor side, we measured oxygen release from dark-adapted cells following single-turnover actinic flashes. The flash-induced oxygen patterns for all strains were normalized to the amplitude of the signal following the third flash and the amplitudes of oxygen released upon each flash plotted (Fig. 4). No difference in the pattern of oxygen release was observed, indicating that the dark-state resting distribution of S-states and function of the oxygen-evolving centers in all strains were similar. These results also suggest the impaired steady-state oxygen evolution and sensitivity to photoinactivation observed in the absence of Ycf48 stems from the perturbed acceptor side, not from any alteration to the donor side. Assembly of PS II complexes The thylakoid membrane-bound protein complexes of each strain were solubilized using b-dodecyl maltoside and then separated by blue-native polyacrylamide gel electrophoresis (BN-PAGE) (Fig. 5a). The subunit composition of each complex was subsequently probed by Western blotting (Fig. 5b–e). In wild type, the three main macromolecular PS II complexes were observed (i.e., dimers, monomers, and RC47 (CP43-less monomers)) along with several large supercomplexes with a size greater than that of a PS II dimer. In addition, complexes with masses between that of monomers and dimers and exhibiting reactivity toward the D1 and D2 but not CP43 antibodies were present. Because the CP43-containing precursor complex is the last major component of PS II to join the mature center and the first to dissociate during the repair cycle, we theorize that these intermediate bands are either precursor complexes prior to formation of mature PS II dimers or centers from which CP43 has dissociated—but with additional repair machinery such as proteases or chlorophyll biosynthesis enzymes and insertion apparatus bound. We have therefore labeled these as uncharacterized PS II intermediates. None of the mutant strains were notably impaired in their ability to assemble the large macromolecular PS II complexes observed in wild type. However, the mutants lacking Ycf48 accumulated a low-molecular-weight complex containing the CP43 subunit (Fig. 5d). It is possible that these low-molecular-weight complexes are responsible for the elevated emission at 685 nm in the 77 K fluorescence emission spectra, because isolated CP43 and CP47 complexes have been shown to display such emission (Boehm et al. 2011).
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Table 3 Kinetic analysis of QA oxidation No addition
Fast phase (t1/2 | amplitude)
wild type
626 ls (±21) | 60 % (±1.0)
6.4 ms (±0.5) | 27 % (±1.6)
7.7 s (±1.1) | 13 % (±0.7)
DYcf48
721 ls (±33) | 56 % (±1.2)
12.1 ms (±3.8) | 27 % (±2.8)
5.3 s (±1.0) | 17 % (±2.3)
DCyanoP
661 ls (±12) | 61 % (±1.8)
6.9 ms (±0.6) | 27 % (±2.4)
6.6 s (±0.7) | 12 % (±0.7)
DCyanoP:DYcf48
659 ls (±86) | 55 % (±2.9)
11.9 ms (±4.9) | 25 % (±4.3)
4.5 s (±1.3) | 20 % (±3.2)
With DCMU
Middle phase (t1/2 | amplitude)
Slow phase (t1/2 | amplitude)
Fast phase (t1/2 | amplitude)
Slow phase (t1/2 | amplitude)
wild type
5.7 ms (±1.1) | 4.3 % (± 0.2)
1.20 s (±0.04) | 95.7 % (±0.2)
DYcf48 DCyanoP
4.1 ms (±0.9) | 7.4 % (± 0.4) 5.0 ms (±0.5) | 4.2 % (± 0.4)
1.50 s (±0.06) | 92.6 % (±0.5) 1.18 s (±0.06) | 95.8 % (±0.4)
DCyanoP:DYcf48
3.8 ms (±0.3) | 8.8 % (± 1.0)
1.48 s (±0.08) | 91.2 % (±1.0)
Fig. 4 Flash-induced turnover of the oxygen-evolving complex. The oxygen production of dark-adapted cells following a train of flashes and normalized to the yield from the third flash for the wild type (a), DYcf48 (b), DCyanoP (c), and DCyanoP:DYcf48 (d) strains
Normalized oxygen yield
Kinetic analysis was performed on corrected fluorescence relaxation curves following a saturating single-turnover flash in the absence or presence of DCMU as described in Jackson et al. (2014)
a
b
c
d
1.0
0.5
Normalized oxygen yield
0.0
1.0
0.5
0.0 0
4
8
12
16
Flash number
To verify the findings obtained by western blot analysis, we performed 2D gel electrophoresis on detergent-solubilized thylakoid membrane samples from each of the strains (Fig. 6). Protein complexes were separated in the first
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dimension by BN-PAGE, and then the constituent proteins were separated in the second dimension by denaturing SDS-PAGE. The gels were stained with SYPRO Ruby then subsequently colloidal Coomassie. After the Coomassie
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a (BN-PAGE) 1
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Supercomplexes PS I trimer PS II intermediates PS II dimer PS II intermediates PS I monomer PS II monomer RC47 Pre-cursor complexes Unassembled subunits
Fig. 5 Analysis of photosynthetic complex assembly. Solubilized thylakoid membranes run on BN-PAGE (3–12 %) (a) followed by western blotting using antibodies for the D1 (b), D2 (c), CP43 (d),
and PsaA (e) proteins. Lanes are wild type (1), DYcf48 (2), DCyanoP (3), and DCyanoP:DYcf48 (4)
staining, multiple gel spots were excised, and LC/MS/MS was used to identify their major contributing proteins (Fig. 6a–d). SYPRO ruby is a fluorescent stain that, in conjunction with a linear fluorescence detector system, allows more accurate quantitative comparison of the protein separations between mutants than can be attained with Coomassie staining (Nishihara and Champion 2002). Using the linear fluorescence intensity data, we overlaid the gel images and used false coloring to visually illustrate the quantitative differences in spot intensity between strains (Fig. 6e–g). Comparison of wild type and the CyanoP knockout did not reveal any marked changes in protein abundance; the majority of spots appear yellow, indicative of similar fluorescence emission intensities (Fig. 6e). In contrast, the DYcf48 strain displayed reduced levels (spots that appear red) of PS II dimers and monomers and an increase (spots that appear green) in early assembly complexes, consistent with those observed in the western blot analyses (Fig. 6f). These precursor complexes displayed migration in the BN-PAGE dimension consistent with the previously identified CP47* and CP43a* complexes (Komenda et al. 2004), and the presence of the PsbH and PsbY subunits belonging to the CP47* complex (or CP47 pre-complex (Boehm et al. 2011)) were confirmed by mass spectrometry. Additionally, the DYcf48 strain showed an increase in the level of cytochrome b6f dimers, the PII nitrogen regulator GlnB, and the carbon concentrating proteins CcmK1 and CcmK2. Comparison of the DYcf48 strain with the DCyanoP:DYcf48 strain revealed a small increase in the levels of PS II dimers and monomers in the double knockout (green spots for the core D1 and D2 proteins) and a reduction in the accumulation of CP47* (Fig. 6g).
Discussion Removal of CyanoP does not impair function of the mature PS II complex Knockout of CyanoP from the wild-type background did not reduce the rates of photoautotrophic growth or oxygen evolution of the cells under the conditions tested. These findings are generally consistent with earlier studies of sll1418 knockout mutants of Synechocystis 6803 grown under nutrient-replete conditions, although these earlier studies do report conflicting levels of impairment in the oxygen-evolving capacity of PS II, ranging from a 0–40 % reduction compared to wild type. (Thornton et al. 2004; Ishikawa et al. 2005; Summerfield et al. 2005). The work by Sveshnikov et al. (2007) investigated CyanoP knockout mutants from several laboratories under comparable conditions, including flash-induced oxygen evolution measurements where differences were observed in the data obtained for the DCyanoP strains and wild type. In contrast, our measurements of flash-induced oxygen evolution did not reveal any difference between CyanoP and wild type. Furthermore, our fluorescence induction measurements and the relaxation of variable fluorescence following a single-turnover actinic flash, together with corresponding kinetic analyses, show no observable alteration of electron transfer from QA to QB, PQ exchange, or recombination with the donor side. In Thermosynechococcous elongatus, the CyanoP protein has recently been shown to bind inactive monomers of PS II in a mutually exclusive position to the PsbO protein, and it was further hypothesized that CyanoP might coordinate the free C-terminus of the D1 peptide (Cormann et al. 2014). This fits with our findings that in the absence of CyanoP, even though maturation or
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Photosynth Res (2015) 124:217–229 PS I trimer PS II dimer ATP synthase NADH dehydrogenase PS I monomer PS II monomer ATP synthase RC47 Cyt b6 f dimer Cyt b6 f monomer CP47* CP43a* CP43b*
PS I trimer PS II dimer ATP synthase NADH dehydrogenase PS I monomer PS II monomer ATP synthase RC47 Cyt b6 f dimer Cyt b6 f monomer CP47* CP43a* CP43b*
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b ΔCyanoP
wild type DnaK2 NdhH ATP synthase γ
CP47 CP43
62 49
factor Tu
38
Phycobillisome linkers
D2 D1
98
28
Cyt f
ATP synthase δ ATP synthase β ATP synthase β’
17 14
b559 α PsbH
Molecular weight (kDa)
a
6 3
c
d
ATP synthase α
ΔCyanoP:ΔYcf48
62 49
CP47 CP43
38
Cyt f
Cyt f
98
28 Cyt b6
β C-phycocyanin α C-phycocyanin
17 14
PII nitrogen regulator GlnB CcmK1, CcmK2 PsbH
b559 α
6
PsbY
e
wild type
ΔCyanoP
f
wild type
Molecular weight (kDa)
ΔYcf48
3 ΔYcf48
g
ΔYcf48
ΔCyanoP:ΔYcf48
62 49 38 28 17 14
6 3
Quenching of stain fluorescence by pigments from the PS I trimers and monomers or phycocyanin molecules
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Reduced level of PS II dimers and monomers
Increased PS II assembly precursor complexes
Increased level of PS II dimers and monomers
Reduced CP47*
Molecular weight (kDa)
98
Photosynth Res (2015) 124:217–229 b Fig. 6 2D gel electrophoresis analysis of thylakoid membrane
protein complexes. Solubilized thylakoid membrane proteins run on BN-PAGE (4–16 %) then SDS-PAGE (12 %). Coomassie-stained images for the wild type (a), DCyanoP (b), DYcf48 (c), and DCyanoP:DYcf48 (d) and false-colored comparison of SYPRO Ruby-stained fluorescence emission for the wild type (red) and DCyanoP (green) (e), wild type (red) and DYcf48 (green) (f), and DYcf48 (red) and DCyanoP:DYcf48 (green) (g)
repair of centers may be perturbed, once PS II centers have matured beyond the point of CyanoP involvement, they are functionally indistinct from centers formed in the presence of CyanoP. Removal of CyanoP influences antenna coupling In contrast to the above, we did observe a consistent difference between DCyanoP and wild-type cells when their 77 K fluorescence emission spectra were measured using 580 nm excitation. The 680 nm emission was reduced in the DCyanoP strain, suggesting a major difference in phycobilisome coupling to PS II. Support for this hypothesis is provided by the room temperature fluorescence data obtained using measuring flashes that specifically excite phycobilisome pigments (627 nm, RMF). A decrease in emission from the dark-adapted state FO (RMF) for the DCyanoP strain was observed, whereas the opposite effect was seen for DYcf48 cells where an increase in FO (RMF) was observed (Table 2), consistent with the increased 77 K peak at 680 nm (Fig. 2c) for this strain. We also verified this phenotype in a separate, independently created DCyanoP knockout mutant (data not shown). A link between proteins found in the thylakoid lumen influencing phycobilisome coupling is also supported by Veerman et al. (2005). Precedent for an involvement of the higher plant CyanoP homolog (PsbP) in regulation of light harvesting and energy distribution has been observed where the formation and stabilization of supercomplexes containing PS II and peripheral light-harvesting complexes (LHCs) were disrupted in the absence of PsbP (Boekema et al. 2000; Ido et al. 2009; Ifuku et al. 2011). Although cyanobacteria do not contain LHCs but instead utilize phycobilisomes, using BN-PAGE or subsequent western blotting, we did not observe analogous changes in supercomplex formation in either the DCyanoP or DCyanoP:DYcf48 strains. Nevertheless, phycobilisome to PS II coupling is linked to PS II assembly and repair, and we noted a similar decrease in the 680 nm emission for a DPsb27 strain (Hwang et al. 2008; Jackson et al. 2014). Given that CyanoP is suggested to be involved in the biogenesis or repair of PS II (Cormann et al. 2014), this difference possibly reflects altered coupling between incompletely assembled PS II centers and phycobilisomes.
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Inactivation of Ycf48 dominates the phenotype of the double mutant In most of our experiments, the phenotype of the DCyanoP:DYcf48 double mutant was similar but not identical to the DYcf48 strain. Additive deleterious effects were present for the room temperature fluorescence measurements; however, the increased oxygen evolution, elevated PS II to PS I ratio, and increased levels of PS II monomer and dimer assembly in the double mutant suggest that removal of CyanoP partially alleviates an assembly block that is present in the absence of Ycf48. These findings are in contrast to the results obtained with the DYcf48:DPsb27 strain, where assembly of mature PS II centers was severely perturbed (Jackson et al. 2014). On balance, our current findings do not indicate an additive restriction for the DCyanoP:DYcf48 double mutant relative to the parental DYcf48 strain. Combined with the recent finding from Cormann et al. (2014), that CyanoP binds inactive monomers which also have Psb27 present, this suggests that Ycf48 dissociates before CyanoP binds. This hypothesis is supported by the findings of Komenda et al. (2008), who showed Ycf48 to be involved in early biogenesis but not the RC47 complex and beyond. Additionally, from our 2D gel data comparison, accumulation of a PS II precursor complex containing CP47, in the absence of Ycf48, appears reduced in the absence of both Ycf48 and CyanoP (Fig. 6c, d, g). These observations might indicate either a direct role for CyanoP earlier in biogenesis than the inactive monomer stage (but not associated with the RC core) or that biosynthesis of the CP47 pre-complex is related to the presence of CyanoP. We previously observed the DYcf48 strain to accumulate a precursor complex containing CP43 which likely gives rise to the increase in 77 K fluorescence emission at a peak of 685 nm (Jackson et al. 2014). This putative pre-complex is also evident in the DCyanoP:DYcf48 strain, demonstrating that its accumulation is not reliant on the presence of CyanoP (Figs. 5d, 6d).
Concluding remarks Our findings for the inactivation of CyanoP in the wildtype background support current models where the role of CyanoP is neither in the mature complex nor is it required for function under normal conditions. Moreover, our results are consistent with there being an altered interaction between incompletely assembled PS II centers and phycobilisomes in strains lacking CyanoP, and that this gives rise to fluorescence quenching at 680 nm in the 77 K fluorescence emission spectra when the phycobilisomes are directly excited by 580 nm light. Investigation of the role of
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CyanoP in early biogenesis in conjunction with the Ycf48 assembly factor indicates that it is unlikely there is a direct interaction between complexes containing CyanoP or Ycf48 during the biogenesis or repair of PS II. Compared to DYcf48 cells, a reduction in the PsbH- and PsbY-containing CP47 pre-complex was observed in the DCyanoP:DYcf48 strain. Furthermore, a small increase in the accumulation of PS II dimers and monomers, supported by 77 K fluorescence emission spectra and quantitative 2D gel analyses, along with an increase in oxygen evolution capacity, compared to the DYcf48 single mutant, suggests that the CyanoP protein impedes PS II assembly in the absence of Ycf48. Acknowledgments S.A.J. was supported by a University of Otago Division of Health Sciences Career Development Postdoctoral Fellowship. Other funding for this project was provided by an Otago University Research grant to J.E.-R. We thank Asher J. Dale for assistance with preliminary experiments.
References Boehm M, Romero E, Reisinger V, Yu J, Komenda J, Eichacker LA, Dekker JP, Nixon PJ (2011) Investigating the early stages of Photosystem II assembly in Synechocystis sp. PCC 6803: isolation of CP47 and CP43 complexes. J Biol Chem 286:14812–14819 Boekema EJ, Van Breemen JF, Van Roon H, Dekker JP (2000) Conformational changes in Photosystem II supercomplexes upon removal of extrinsic subunits. Biochemistry 39:12907–12915 Bricker TM, Roose JL, Fagerlund RD, Frankel LK, Eaton-Rye JJ (2012) The extrinsic proteins of Photosystem II. Biochim Biophys Acta 1817:121–142 Cormann KU, Bangert JA, Ikeuchi M, Ro¨gner M, Stoll R, Nowaczyk MM (2009) Structure of Psb27 in solution: implications for transient binding to Photosystem II during biogenesis and repair. Biochemistry 48:8768–8770 Cormann KU, Bartsch M, Ro¨gner M, Nowaczyk MM (2014) Localization of the CyanoP binding site on Photosystem II by surface plasmon resonance spectroscopy. Front Plant Sci. doi:10. 3389/fpls.2014.00595 Cser K, Vass I (2007) Radiative and non-radiative charge recombination pathways in Photosystem II studied by thermoluminescence and chlorophyll fluorescence in the cyanobacterium Synechocystis 6803. Biochim Biophys Acta 1767:233–243 Eaton-Rye JJ (2011) Construction of gene interruptions and gene deletions in the cyanobacterium Synechocystis sp. strain PCC 6803. Methods Mol Biol 684:295–312 Fagerlund RD, Eaton-Rye JJ (2011) The lipoproteins of cyanobacterial Photosystem II. J Photochem Photobiol, B 104:191–203 Hwang HJ, Nagarajan A, McLain A, Burnap RL (2008) Assembly and disassembly of the Photosystem II manganese cluster reversibly alters the coupling of the reaction center with the light-harvesting phycobilisome. Biochemistry 47:9747–9755 Ido K, Ifuku K, Yamamoto Y, Ishihara S, Murakami A, Takabe K, Miyake C, Sato F (2009) Knockdown of the PsbP protein does not prevent assembly of the dimeric PSII core complex but impairs accumulation of Photosystem II supercomplexes in tobacco. Biochim Biophys Acta 1787:873–881
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Photosynth Res (2015) 124:217–229 Ifuku K, Ido K, Sato F (2011) Molecular functions of PsbP and PsbQ proteins in the Photosystem II supercomplex. J Photochem Photobiol, B 104:158–164 Ishikawa Y, Schro¨der WP, Funk C (2005) Functional analysis of the PsbP-like protein (sll1418) in Synechocystis sp. PCC 6803. Photosynth Res 84:257–262 Jackson SA, Fagerlund RD, Wilbanks SM, Eaton-Rye JJ (2010) Crystal structure of PsbQ from Synechocystis sp. PCC 6803 at ˚ : implications for binding and function in cyanobacterial 1.8 A Photosystem II. Biochemistry 49:2765–2767 Jackson SA, Hinds MG, Eaton-Rye JJ (2012) Solution structure of CyanoP from Synechocystis sp. PCC 6803: new insights on the structural basis for functional specialization amongst PsbP family proteins. Biochim Biophys Acta 1817:1331– 1338 Jackson SA, Hervey JR, Dale AJ, Eaton-Rye JJ (2014) Removal of both Ycf48 and Psb27 in Synechocystis sp. PCC 6803 disrupts Photosystem II assembly and alters QA oxidation in the mature complex. FEBS Lett 588:3751–3760 Joliot A, Joliot P (1964) Etude cine´tique de la re´action photochimique libe´rant l’oxyge`ne au cours de la photosynthe`se. C R Acad Sci Paris 258:4622–4635 Joliot P, Joliot A (1968) A polarographic method for detection of oxygen production and reduction of Hill reagent by isolated chloroplasts. Biochim Biophys Acta 153:625–634 Komenda J, Reisinger V, Mu¨ller BC, Doba´kova´ M, Granvogl B, Eichacker LA (2004) Accumulation of the D2 protein is a key regulatory step for assembly of the Photosystem II reaction center complex in Synechocystis PCC 6803. J Biol Chem 279:48620–48629 Komenda J, Nickelsen J, Tichy´ M, Pra´sˇil O, Eichacker LA, Nixon PJ (2008) The cyanobacterial homologue of HCF136/YCF48 is a component of an early Photosystem II assembly complex and is important for both the efficient assembly and repair of Photosystem II in Synechocystis sp. PCC 6803. J Biol Chem 283:22390–22399 Komenda J, Sobotka R, Nixon PJ (2012) Assembling and maintaining the Photosystem II complex in chloroplasts and cyanobacteria. Curr Opin Plant Biol 15:245–251 Mabbitt PD, Rautureau GJ, Day CL, Wilbanks SM, Eaton-Rye JJ, Hinds MG (2009) Solution structure of Psb27 from cyanobacterial Photosystem II. Biochemistry 48:8771–8773 Mabbitt PD, Wilbanks SM, Eaton-Rye JJ (2014) Structure and function of the hydrophilic Photosystem II assembly proteins: Psb27, Psb28 and Ycf48. Plant Physiol Biochem 81:96–107 Messinger J, Schro¨der WP, Renger G (1993) Structure-function relations in Photosystem II. Effects of temperature and chaotropic agents on the period four oscillation of flash-induced oxygen evolution. Biochemistry 32:7658–7668 Meunier PC, Popovic R (1988) High-accuracy oxygen polarograph for photosynthetic systems. Rev Sci Instrum 59:486–491 Michoux F, Takasaka K, Boehm M, Nixon PJ, Murray JW (2010) ˚ : implications for the evolution and Structure of CyanoP at 2.8 A function of the PsbP subunit of Photosystem II. Biochemistry 49:7411–7413 Michoux F, Boehm M, Bialek W, Takasaka K, Maghlaoui K, Barber J, Murray JW, Nixon PJ (2014) Crystal structure of CyanoQ from the thermophilic cyanobacterium Thermosynechococcus elongatus and detection in isolated Photosystem II complexes. Photosynth Res 122:57–67 Morris JN, Crawford TS, Jeffs A, Stockwell PA, Eaton-Rye JJ, Summerfield TC (2014) Whole genome re-sequencing of two ‘wild-type’ strains of the model cyanobacterium Synechocystis sp. PCC 6803. N Z J Bot 52:36–47 Nickelsen J, Rengstl B (2013) Photosystem II assembly: from cyanobacteria to plants. Annu Rev Plant Biol 64:609–635
Photosynth Res (2015) 124:217–229 Nishihara JC, Champion KM (2002) Quantitative evaluation of proteins in one- and two-dimensional polyacrylamide gels using a fluorescent stain. Electrophoresis 23:2203–2215 Nixon PJ, Michoux F, Yu J, Boehm M, Komenda J (2010) Recent advances in understanding the assembly and repair of Photosystem II. Ann Bot 106:1–16 Renger G, Hanssum B (2009) Oxygen detection in biological systems. Photosynth Res 102:487–498 Strasser RJ, Srivastava A, Govindjee (1995) Polyphasic chlorophylla fluorescence transient in plants and cyanobacteria. Photochem Photobiol 61:32–42 Suga M, Akita F, Hirata K, Ueno G, Murakami H, Nakajima Y, Shimizu T, Yamashita K, Yamamoto M, Ago H, Shen J-R ˚ resolution (2015) Native structure of Photosystem II at 1.94 A viewed by femotosecond X-ray pulses. Nature 517:99–103 Summerfield TC, Winter RT, Eaton-Rye JJ (2005) Investigation of a requirement for the PsbP-like protein in Synechocystis sp. PCC 6803. Photosynth Res 84:263–268 Sveshnikov D, Funk C, Schro¨der WP (2007) The PsbP-like protein (sll1418) of Synechocystis sp. PCC 6803 stabilises the donor side of Photosystem II. Photosynth Res 93:101–109
229 Thornton LE, Ohkawa H, Roose JL, Kashino Y, Keren N, Pakrasi HB (2004) Homologs of plant PsbP and PsbQ proteins are necessary for regulation of Photosystem II activity in the cyanobacterium Synechocystis 6803. Plant Cell 16:2164–2175 Umena Y, Kawakami K, Shen J-R, Kamiya N (2011) Crystal structure of oxygen-evolving Photosystem II at a resolution of ˚ . Nature 473:55–60 1.9 A Veerman J, Bentley FK, Eaton-Rye JJ, Mullineaux CW, Vasil’ev S, Bruce D (2005) The PsbU subunit of Photosystem II stabilizes energy transfer and primary photochemistry in the phycobilisome-photosystem II assembly of Synechocystis sp. PCC 6803. Biochemistry 44:16939–16948 Vinyard DJ, Ananyev GM, Dismukes GC (2013) Photosystem II: the reaction center of oxygenic photosynthesis. Annu Rev Biochem 82:577–606 Williams JGK (1988) Construction of specific mutations in Photosystem-II photosynthetic reaction center by genetic-engineering methods in Synechocystis-6803. Methods Enzymol 167:766–778
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REGULAR PAPER
Characterization of a Synechocystis sp. PCC 6803 double mutant lacking the CyanoP and Ycf48 proteins of Photosystem II Simon A. Jackson1 • Julian J. Eaton-Rye1
Received: 23 October 2014 / Accepted: 12 March 2015 / Published online: 24 March 2015 Ó Springer Science+Business Media Dordrecht 2015
Abstract Homologs of the Photosystem II (PS II) subunit PsbP are found in plants, algae, and cyanobacteria. In higher plants, PsbP is associated with mature PS II centers, but in cyanobacteria, the homologous CyanoP protein appears sub-stoichiometric to PS II. We have investigated the role of CyanoP by characterizing knockout mutants of the cyanobacterium Synechocystis sp. PCC 6803. Removal of CyanoP resulted in changes to phycobilisome coupling and energy transfer to PS II, but the function of PS II itself remained similar to wild type. We therefore investigated the hypothesis that CyanoP is involved in the biogenesis or repair of PS II by creating a double mutant lacking both CyanoP and the PS II assembly factor Ycf48. This strain exhibited an additive reduction in the amplitude of variable chlorophyll a fluorescence induction relative to either of the single mutants but displayed increased oxygen evolution, slight increases in PS II monomer and dimer levels, and a reduction in accumulation of an early PS II assembly complex containing CP47, compared to the DYcf48 strain. Keywords Assembly Biogenesis CyanoP Photosystem II PsbP Synechocystis sp. PCC 6803 Ycf48 Abbreviations Bis–Tris 2-[Bis(2-hydroxyethyl)amino]-2(hydroxymethyl)propane-1,3-diol BMF Blue measuring flashes
BN-PAGE CP43 CP47 D1 D2 DCMU FO FM J and I LC/MS/MS LHC O OD P PS I PS II QA QB
& Julian J. Eaton-Rye [email protected] 1
Department of Biochemistry, University of Otago, Dunedin 9016, New Zealand
RC47 RMF S-states
Blue-native polyacrylamide gel electrophoresis 43-kDa chlorophyll a-binding protein of the core antenna 47-kDa chlorophyll a-binding protein of the core antenna Photosystem II reaction center protein subunit Photosystem II reaction center protein subunit 3-(3,4-Dichlorophenyl)-1,1-dimethylurea Dark-adapted (minimum) fluorescence Maximum level of chlorophyll a fluorescence Inflection points between O (FO) and P in the chlorophyll a fluorescence induction curve Liquid chromatography tandem mass spectrometry Light-harvesting complex Origin (FO) level of the chlorophyll a fluorescence induction curve Optical density Peak level in the chlorophyll a fluorescence induction curve Photosystem I Photosystem II Primary plastoquinone electron acceptor of PS II Secondary plastoquinone electron acceptor of PS II Reaction center complex lacking the CP43 protein Red measuring flashes Oxidation states of the oxygen-evolving complex of PS II
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SDS-PAGE Tris
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Sodium dodecyl sulfate polyacrylamide gel electrophoresis 2-Amino-2-hydroxymethyl-propane-1,3-diol
Introduction Photosystem II (PS II) is found in all oxygenic photosynthetic organisms where it is required for the light-driven oxidation of water (Vinyard et al. 2013). The biogenesis of PS II is a multistep process which involves the coordinated assembly of several smaller intermediate complexes to form the mature photosystem (Komenda et al. 2012). The early assembly intermediates include a pre-complex containing the reaction center D1 and D2 proteins and two additional pre-complexes, each containing one of the chlorophyll a-binding core antenna proteins CP43 or CP47. All of these pre-complexes also contain additional protein subunits that are either membranespanning or soluble proteins associated with the exposed hydrophilic domains on both the lumenal and cytosolic faces of the core D1, D2, CP43, or CP47 subunits (Komenda et al. 2012). Many of the proteins found in these intermediate complexes are not found in the mature complex but function as assembly factors or chaperones that are transiently associated with PS II during de novo biogenesis or the PS II repair pathway that operates following light-induced photodamage (Nixon et al. 2010; Nickelsen and Rengstl 2013). In cyanobacteria, a group of three extrinsic PS II subunits: CyanoP, CyanoQ, and Psb27, all possessing an N-terminal lipid modification, are believed to associate with the lumenal face of PS II (Fagerlund and Eaton-Rye 2011). All three subunits are absent from the most recent high resolution crystal structures of PS II (Umena et al. 2011; Suga et al. 2015), but their structures have been determined independently (Cormann et al. 2009; Mabbitt et al. 2009; Jackson et al. 2010; Michoux et al. 2010; Umena et al. 2011; Jackson et al. 2012; Michoux et al. 2014). The CyanoQ and Psb27 subunits have been assigned roles in mature PS II centers and early PS II assembly intermediates, respectively (reviewed in (Bricker et al. (2012) and Mabbitt et al. (2014)). In contrast, any interaction or function of the CyanoP protein in PS II biogenesis, mature centers, or repair processes remains undetermined; however, the previously reported sub-stoichiometric relationship between the levels of intracellular CyanoP and PS II centers suggests that CyanoP might play a role in either de novo biogenesis or repair (Thornton et al. 2004). The Psb27 protein is known to be involved in both biogenesis and repair of PS II, and we recently reported the phenotypic characterization of Synechocystis sp. PCC 6803 (hereafter Synechocystis 6803) strains lacking Psb27 in the
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wild-type background and a mutant background which lacked the PS II assembly factor Ycf48 (Jackson et al. 2014). Ycf48 has been shown to be associated with early assembly complexes of PS II (Komenda et al. 2008) and the additive phenotypes we observed, for the DYcf48:DPsb27 strain compared to the corresponding single-deletion mutants, led to the conclusion that Psb27 and Ycf48 are associated with complexes that interact during the biogenesis or repair cycles (Jackson et al. 2014). In this communication, we test the hypothesis that if CyanoP functions during biogenesis or repair of PS II, then the function of this subunit might also be better revealed when PS II assembly is perturbed by removal of Ycf48.
Materials and methods Cyanobacteria strains and physiology experiments The Synechocystis 6803 strain referred to throughout as wild type is a sub-cultured derivative of the Williams glucose-tolerant strain, designated GT-O1 (Williams 1988; Morris et al. 2014). The strains lacking CyanoP (DCyanoP) and Ycf48 (DYcf48) have been described previously (Summerfield et al. 2005; Jackson et al. 2014). For the double knockout, the ycf48 ORF was inactivated in the DCyanoP background, producing the DCyanoP:DYcf48 mutant. Transformations were performed and strains were maintained as described in Eaton-Rye (2011). All physiological characterizations and experiments were performed as described in Jackson et al. (2014) with the alterations detailed hereafter. Whole-cell absorption spectra were collected using an Evolution 201 UV/Vis spectrophotometer (Thermo Scientific, Waltham, MA, USA). The flashinduced oscillation pattern of oxygen production following single-turnover actinic flashes was measured using a bare platinum Joliot-type electrode based on the designs of Joliot and Joliot (1968), Meunier and Popovic (1988), Messinger et al. (1993), and Renger and Hanssum (2009). The light source employed was a cluster of light-emitting diodes with an emission peak at 627 nm (Philips, Andover, MA, USA) and a custom built driver unit with variable flash-width capability. Two microliters of sample at a chlorophyll a concentration of 1 mg mL-1 in BG-11 (pH 7.5) was loaded onto the electrode, then dark adapted for 5 min, given a singleturnover flash, and dark adapted for a further 5 min before commencement of a train of 25 flashes (32 ls duration) given with a spacing between flashes of 250 ms. Analyses of thylakoid membrane proteins Thylakoid membrane samples were solubilized using bdodecyl maltoside as per Jackson et al. (2014). For Western
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blotting, 2 lg of chlorophyll a equivalent material was loaded per sample on NativePAGE 3–12 % Bis–Tris gradient gels (Life Technologies, Waltham, MA, USA). Electroblot transfer was performed at 50 V for 16 h in buffer containing 25 mM Tris, 192 mM glycine, and 20 % v/v methanol. The primary antibodies for the PsbA (D1), PsbC (CP43), and PsbD (D2) subunits of PS II and the PsaA subunit of PS I were obtained from Agrisera, Va¨nna¨s, Sweden. For 2D gel electrophoresis NativePAGE, 4–16 % Bis–Tris gradient gels (Life Technologies, Waltham, MA, USA) were run as the first dimension, followed by NuPAGE 12 % Bis-Tris gels (Life Technologies, Waltham, MA, USA) as the second (SDS-PAGE) dimension. The SDS-PAGE gels were stained with SYPRO Ruby (Life Technologies, Waltham, MA, USA), imaged with a 16-bit linear CCD detector system (Fuji imager PS3000, Fujifilm, Tokyo, Japan) operating in fluorescence mode (460 nm excitation, Y515 emission filter), de-stained and then restained with colloidal Coomassie Brilliant Blue G-250. For the false-colored overlay comparisons, all SYPRO Rubystained gels were imaged using identical exposure times, and the intensity data are displayed as two linear channels (red or green, for each strain as indicated) on a composite image with identical contrast and brightness settings applied to all channels. For mass spectrometry analyses, the gel spots of interest were excised and digested in-gel with trypsin. The resulting peptides were eluted from the gel, dried using a centrifugal concentrator, and reconstituted in an aqueous solution of 5 % v/v acetonitrile and 0.2 % v/v formic acid. The samples were then subject to liquid chromatography tandem mass spectrometry (LC/MS/MS) using a nanoflow uHPLCcoupled TripleTOF 5600? mass spectrometry system (Ab Sciex, Framingham, MA, USA). Samples were separated using a C-18 emitter-tip column operated with a 5–90 % v/v acetonitrile gradient in 0.2 % formic acid. For precursor ion measurement, a mass range between m/z 400–2000 was used, followed by 10 data-dependent collision-induced dissociation (CID) fragment ion measurements per cycle. PeakView (Ab Sciex, Framingham, MA, USA) was used to process the data and generate peak lists that were searched using an in-house Mascot server (Matrix Science, Boston, MA, USA).
Results Photoautotrophic growth, oxygen evolution, and pigment composition Under the conditions used, removal of CyanoP did not significantly influence the photoautotrophic growth rate (doubling time of 11.0 h) compared to wild type (11.5 h)
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(Fig. 1a), whereas inactivation of the ycf48 ORF impaired photoautotrophic growth, increasing the doubling time to 21.4 h for the DYcf48 strain. However, removal of Ycf48 from the DCyanoP strain did not result in an additive impairment in growth beyond that already imposed by inactivation of the ycf48 ORF; the doubling time of the DCyanoP:DYcf48 strain was 19.7 h. The saturated rate of steady-state oxygen evolution was found to be comparable for wild type and the DCyanoP cells (555 and 556 lmol O2 per mg chlorophyll a per h, respectively) (Table 1). Consistent with the photoautotrophic growth data, removal of Ycf48 in the wild-type background reduced the rate of oxygen evolution to approximately 43 %, and removal of both CyanoP and Ycf48 resulted in 48 % activity compared to wild type. The level of photoinactivation observed during the time course of the assay for the DYcf48 strain (as evident by the curvature of the trace during the light exposure) was not influenced by the presence or absence of the CyanoP subunit, suggesting that CyanoP is not involved in photoprotection (Fig. 1b). The previously characterized DYcf48:DPsb27 strain (Jackson et al. 2014) exhibited changes in the cellular pigment composition, notably large increases in the carotenoid absorption region and suppression of the 680-nm peak that originates from absorption by chlorophyll a. Upon investigation of the DCyanoP and DCyanoP:DYcf48 strains, we identified a similar effect, with the 680-nm peak lower for the DCyanoP:DYcf48 double mutant than observed for the DYcf48 strain (Fig. 1c–d). Low temperature (77 K) chlorophyll a fluorescence emission Using an excitation beam that specifically targets chlorophyll a (wavelength of 440 nm), we measured low temperature (77 K) fluorescence emission spectra from each of the strains (Fig. 2a, b). A general overview of interpretation of the emission spectrum for wild type is that the 685 and 695 nm maxima originate from the CP43 and CP47 subunits of PS II, respectively, whilst the maximum PS I emission occurs at 725 nm; however, for PS II assembly mutants, the 685 nm peak might also contain contributions from other species (e.g., Boehm et al. (2011) and Jackson et al. (2014)). Inactivation of the ycf48 gene in the wildtype background resulted in an apparent reduction in the PS II to PS I ratio and an increase in emission at 685 relative to 695 nm, in agreement with our previous observations (Jackson et al. 2014). For the DCyanoP strain, the emission spectra showed a small reduction in the PS II to PS I ratio compared to wild type. Inactivation of the ycf48 gene in the DCyanoP mutant background did not result in a decrease in the PS II to PS I ratio (as observed for the DYcf48 mutant versus wild type), but a comparable increase in the
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b
3.2
Oxygen evolved (μmol per mg Chl a)
a
1.6
OD730
0.8 0.4
0.2 0.1 0.05
Light off (i) (iii)
25 20 15
(ii) (iv)
10 5 0 Light on
-5 0
24
48
72
96
120
0
60
Time (h)
120
180
240
300
Time (s)
d
c
Normalized absorbance
1.0 0.8 0.6 0.4 0.2
1.0 0.8 0.6 0.4 0.2
0.0
0.0 400
500
600
700
Wavelength (nm)
Table 1 Oxygen evolution rates (lmol O2 mg Chl a wild type
555 ± 29
DYcf48
236 ± 32
DCyanoP
556 ± 38
DCyanoP:DYcf48
264 ± 18
-1
-1
h )
Oxygen evolution was measured in the presence of 2,6-dichloro-1,4benzoquinone and K3Fe(CN)6 as described in Jackson et al. (2014)
amplitude of the 685 nm compared to 695 nm emission peak was observed. The slightly higher PS II to PS I ratio for the DCyanoP:DYcf48 strain, compared to DYcf48 cells, correlated with the *10 % higher rate of oxygen evolution observed for this strain (Table 1). Examination of 77 K fluorescence emission using an excitation wavelength targeting phycobilisome pigments (580 nm) can reveal differences in phycobilisome-coupled energy distribution (Fig. 2c, d). The DYcf48 strain
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0.025
Normalized absorbance
Fig. 1 Photoautotrophic growth, oxygen evolution, and whole-cell absorption spectra. a Photoautotrophic growth curve determined by light scattering at 730 nm: wild type (solid squares), DYcf48 (open squares), DCyanoP (solid circles), and DCyanoP:DYcf48 (open circles). Error bars represent the standard error from at least 3 independent experiments. b Oxygen evolution traces: wild type (i), DYcf48 (ii), DCyanoP (iii), and DCyanoP:DYcf48 (iv). c Whole-cell absorption spectra: wild type (solid line) and DYcf48 (dotted line). d Wholecell absorption spectra: DCyanoP (solid line) and DCyanoP:DYcf48 (dotted line). In c and d, spectra are the average of three independent experiments and are normalized to the maxima at 435 nm
Photosynth Res (2015) 124:217–229
400
500
600
700
Wavelength (nm)
exhibited a characteristic increase in the emission peak at 680 nm compared to wild type. The 680 nm maximum originates from phycobilisome emission, but PS II-specific fluorophores give this peak a red-shifted shoulder, thus resolution of the underlying components is difficult. The DCyanoP strain showed a marked reduction in the 680 nm emission compared to wild type, but in the case of the DCyanoP:DYcf48 strain, removal of Ycf48 in the DCyanoP mutant background resulted in a phenotype whereby the 680 nm peak increased to a level similar to that observed for the DYcf48 strain. Variable chlorophyll a fluorescence measurements Measurements of room temperature variable chlorophyll a fluorescence were used to further investigate phycobilisome to PS II coupling and also examine electron transfer on the acceptor side of PS II. Our instrument is equipped to probe the state of PS II centers using either blue measuring flashes (BMF) for direct excitation of chlorophyll a or red
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a
1
Normalised fluorescence
1.0
Normalized fluorescence
Fig. 2 77 K fluorescence emission spectra. a 440 nm excitation for wild type (solid line) and DYcf48 (dotted line). b 440 nm excitation for DCyanoP (solid line) and DCyanoP:DYcf48 (dotted line). c 580 nm excitation for wild type (solid line) and DYcf48 (dotted line). d 580 nm excitation for DCyanoP (solid line) and DCyanoP:DYcf48 (dotted line). Spectra are the average of at least three independent experiments and are normalized to the PS I emission maxima
221
0.8
0.6
0.4
0.2
0.0
4.0 625
650
700
725
750
3.0
2.0
1.0
650
0.6
0.4
0.2
0 775 625 4
675
700
725
Wavelength (nm)
measuring flashes (RMF) for assessment of phycobilisomecoupled fluorescence emission. Comparison of data obtained using either BMF or RMF provides information on energy partitioning to complement the 77 K fluorescence emission spectra. The amplitudes of fluorescence presented herein are arbitrary and specific to the optical configuration of the fluorometer used. In addition, the relative amplitudes of data obtained with BMF versus RMF are not directly comparable because these are dependent on the measuring flash intensity, duration, and detector gain parameters. The amplitudes of fluorescence emission during the induction time course were significantly decreased for the DYcf48 strain relative to wild type when measured using either BMF or RMF (Fig. 3a–d) (Jackson et al. 2014). In contrast, inactivation of cyanoP (sll1418) in the wild-type background did not significantly affect the shape or amplitude of the fluorescence induction traces. Also shown in Fig. 3, the DCyanoP:DYcf48 double mutant exhibited a
650
675
700
725
750
775
750
775
d Wavelength (nm)
0.0
625
0.8
Wavelength (nm) Normalised fluorescence
Normalized fluorescence
c
675
b
750
3
2
1
0 775 625
650
675
700
725
Wavelength (nm)
reduction in the maximal amplitudes of fluorescence observed compared to the DYcf48 strain but retained the O, J, I, and P features of the fluorescence induction curve (Strasser et al. 1995). This is in contrast to the DYcf48:DPsb27 strain, where as well as a reduction in amplitude, we discovered loss of the J-P rise that is normally observed in the absence of 3-(3,4-dichlorophenyl)-1,1-dimethylurea) DCMU (Jackson et al. 2014). The amplitude of the fluorescence in the P region changes with addition of DCMU. This is most likely because DCMU blocks electron flow to the plastoquinone pool and influences the redox potential of the acceptor side. For wild type or the DCyanoP strain, the fluorescence yield was similar or reduced in the presence of DCMU, whilst in the DYcf48 and DCyanoP:DYcf48 strains, the yield was increased by addition of DCMU. A mechanistic interpretation for this observation has yet to be elucidated, but it might result from centers with a perturbed acceptor side formed in the absence of Ycf48 (Jackson et al. 2014).
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- DCMU
+ DCMU
1
1.0
0.6
0.4 0.2 0.0 1
0.6
0.4
0.2
0.0 10-4
10-3
10-2
0.1 Time (s)
1
1
10
1
10
1
10
0.6
0.4
0.2
-5 -4 -3 -2 0.1 10 1.2 10 10 10 10 Time (s)
f
1.0
Fluorescence (F-F0)/(FM-F0)
Fluorescence (F-FO)/(FM-FO)
0.2
0 10-5
e 0.8 0.6 0.4 0.2 0.0
10-5 10-4 10-3 10-2 0.1 Time (s)
Four BMF or RMF were used to determine the background dark-state fluorescence (FO) (Table 2), prior to the start of the activation of actinic illumination of the fluorescence induction traces. No appreciable difference between the FO for all strains was observed using BMF, but when using RMF, the FO level was increased in DYcf48 cells and substantially decreased in the
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Blue measuring flashes (BMF)
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0.6
0 -5 -4 -3 -2 0.1 10 0.8 10 10 10 10 Time (s) d Fluorescence (F-F0) a.u.
-5 -4 -3 -2 0.1 0.8 10 10 10 10 Time (s) c
0.8
Red measuring flashes (RMF)
0.8
Blue measuring flashes (BMF)
b Fluorescence (F-F0) a.u.
Fluorescence (F-FO) a.u.
a
Fluorescence (F-FO) a.u.
Fig. 3 Room temperature fluorescence induction and relaxation of fluorescence following single-turnover actinic flashes. a–d Room temperature fluorescence induction observed upon illumination of dark-adapted cells with a constant actinic light. Strains are wild type (solid), DYcf48 (dots), DCyanoP (dashes), and DCyanoP:DYcf48 (dash-dots) in the absence (a, c) and presence (b, d) of DCMU. In all instances, constant blue (455 nm) illumination was used with either blue (a, b) or red (625 nm) (c, d) measuring flashes. The time scale is relative to the commencement of illumination. e Relaxation of fluorescence following a singleturnover actinic flash in the absence of DCMU. f Relaxation of fluorescence following a single-turnover actinic flash in the presence of DCMU. In e and f, the strains are wild type (solid), DYcf48 (dots), DCyanoP (dashes), and DCyanoP:DYcf48 (dash-dots). Data displayed are for the average of at least three independent experiments
Photosynth Res (2015) 124:217–229
1.0 0.8 0.6 0.4 0.2
0.0 10 10-5 10-4 10-3 10-2 0.1 Time (s)
DCyanoP strain. Additional removal of Ycf48 from DCyanoP cells reversed the reduction, resulting in a FO (RMF) similar to the DYcf48 mutant. This mirrors the trend observed in the 77 K emission spectra obtained using a 580 nm excitation beam where lack of Ycf48 resulted in a similar phenotype for both DYcf48 and DCyanoP:DYcf48 cells.
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The oxidation state of QA following a single-turnover saturating blue flash (455 nm), in the absence and presence of DCMU, was investigated by probing the relaxation of chlorophyll a fluorescence emission using BMF (Fig. 3e, f). The fluorescence traces were analyzed by applying a Joliot correction and modeling two exponential and one hyperbolic decays in the absence of DCMU and one exponential and one hyperbolic in the presence of DCMU (Joliot and Joliot 1964; Cser and Vass 2007). The QA oxidation kinetics derived from this analysis are summarized in Table 3. In the absence of DCMU, the transfer kinetics of the DCyanoP strain were similar to those observed in wild type, indicating that the absence of CyanoP does not influence electron transfer on the acceptor side of PS II. The Ycf48 knockout in DYcf48 cells, however, resulted in a decrease in the rate of electron transfer from QA to QB for centers where the QB site was likely empty at the time of the actinic flash (middle phase). An increase in the rate and amplitude for the slow phase was also observed, suggestive of increased recombination via back reaction with the oxygen-evolving complex (Jackson et al. 2014). In the DCyanoP:DYcf48 double mutant, these same effects were observed, consistent with CyanoP not influencing the acceptor-side electron transfer kinetics, and removal of Ycf48 eliciting the primary influence in the rates and amplitudes observed. In the presence of DCMU, oxidation of QA in wild-type cells was dominated by the back reaction (slow phase) with a rate constant of 1.2 s (95.7 %) and a minor fast phase component (4.3 %) (Table 3). The DCyanoP strain exhibited similar back reaction characteristics to wild type, indicating that neither the donor nor acceptor sides of PS II are destabilized by the absence of CyanoP. In the absence of Ycf48, the fast phase increased in rate (half time of 4.1 ms) and almost doubled in amplitude (to 7.4 %), observable in the normalized fluorescence decay trace (Fig. 3f) (Jackson et al. 2014). The DCyanoP:DYcf48 strain showed only minor changes in these kinetics compared to those observed in DYcf48 cells (Table 3), suggesting that the impairment of acceptor-side function in the absence of Ycf48 is not influenced by the presence or absence of CyanoP. Table 2 Dark-adapted background (FO) fluorescence data BMF (a.u.)
RMF (a.u.)
wild type
0.504 ± 0.021
0.725 ± 0.025
DYcf48
0.536 ± 0.030
0.805 ± 0.085
DCyanoP
0.446 ± 0.025
0.549 ± 0.047
DCyanoP:DYcf48
0.544 ± 0.040
0.849 ± 0.136
The amplitudes between BMF and RMF are not directly comparable. Data shown were collected in the absence of DCMU. No significant difference was observed in its presence
Flash-induced oxygen evolution Efficient acceptor-side electron transfer and plastoquinone exchange are necessary to support full activity of the PS II complex, including maintaining function of the donor side and oxygen evolution. To investigate whether, in the absence of Ycf48, donor side function is impacted by the impairment of the acceptor side, we measured oxygen release from dark-adapted cells following single-turnover actinic flashes. The flash-induced oxygen patterns for all strains were normalized to the amplitude of the signal following the third flash and the amplitudes of oxygen released upon each flash plotted (Fig. 4). No difference in the pattern of oxygen release was observed, indicating that the dark-state resting distribution of S-states and function of the oxygen-evolving centers in all strains were similar. These results also suggest the impaired steady-state oxygen evolution and sensitivity to photoinactivation observed in the absence of Ycf48 stems from the perturbed acceptor side, not from any alteration to the donor side. Assembly of PS II complexes The thylakoid membrane-bound protein complexes of each strain were solubilized using b-dodecyl maltoside and then separated by blue-native polyacrylamide gel electrophoresis (BN-PAGE) (Fig. 5a). The subunit composition of each complex was subsequently probed by Western blotting (Fig. 5b–e). In wild type, the three main macromolecular PS II complexes were observed (i.e., dimers, monomers, and RC47 (CP43-less monomers)) along with several large supercomplexes with a size greater than that of a PS II dimer. In addition, complexes with masses between that of monomers and dimers and exhibiting reactivity toward the D1 and D2 but not CP43 antibodies were present. Because the CP43-containing precursor complex is the last major component of PS II to join the mature center and the first to dissociate during the repair cycle, we theorize that these intermediate bands are either precursor complexes prior to formation of mature PS II dimers or centers from which CP43 has dissociated—but with additional repair machinery such as proteases or chlorophyll biosynthesis enzymes and insertion apparatus bound. We have therefore labeled these as uncharacterized PS II intermediates. None of the mutant strains were notably impaired in their ability to assemble the large macromolecular PS II complexes observed in wild type. However, the mutants lacking Ycf48 accumulated a low-molecular-weight complex containing the CP43 subunit (Fig. 5d). It is possible that these low-molecular-weight complexes are responsible for the elevated emission at 685 nm in the 77 K fluorescence emission spectra, because isolated CP43 and CP47 complexes have been shown to display such emission (Boehm et al. 2011).
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Table 3 Kinetic analysis of QA oxidation No addition
Fast phase (t1/2 | amplitude)
wild type
626 ls (±21) | 60 % (±1.0)
6.4 ms (±0.5) | 27 % (±1.6)
7.7 s (±1.1) | 13 % (±0.7)
DYcf48
721 ls (±33) | 56 % (±1.2)
12.1 ms (±3.8) | 27 % (±2.8)
5.3 s (±1.0) | 17 % (±2.3)
DCyanoP
661 ls (±12) | 61 % (±1.8)
6.9 ms (±0.6) | 27 % (±2.4)
6.6 s (±0.7) | 12 % (±0.7)
DCyanoP:DYcf48
659 ls (±86) | 55 % (±2.9)
11.9 ms (±4.9) | 25 % (±4.3)
4.5 s (±1.3) | 20 % (±3.2)
With DCMU
Middle phase (t1/2 | amplitude)
Slow phase (t1/2 | amplitude)
Fast phase (t1/2 | amplitude)
Slow phase (t1/2 | amplitude)
wild type
5.7 ms (±1.1) | 4.3 % (± 0.2)
1.20 s (±0.04) | 95.7 % (±0.2)
DYcf48 DCyanoP
4.1 ms (±0.9) | 7.4 % (± 0.4) 5.0 ms (±0.5) | 4.2 % (± 0.4)
1.50 s (±0.06) | 92.6 % (±0.5) 1.18 s (±0.06) | 95.8 % (±0.4)
DCyanoP:DYcf48
3.8 ms (±0.3) | 8.8 % (± 1.0)
1.48 s (±0.08) | 91.2 % (±1.0)
Fig. 4 Flash-induced turnover of the oxygen-evolving complex. The oxygen production of dark-adapted cells following a train of flashes and normalized to the yield from the third flash for the wild type (a), DYcf48 (b), DCyanoP (c), and DCyanoP:DYcf48 (d) strains
Normalized oxygen yield
Kinetic analysis was performed on corrected fluorescence relaxation curves following a saturating single-turnover flash in the absence or presence of DCMU as described in Jackson et al. (2014)
a
b
c
d
1.0
0.5
Normalized oxygen yield
0.0
1.0
0.5
0.0 0
4
8
12
16
Flash number
To verify the findings obtained by western blot analysis, we performed 2D gel electrophoresis on detergent-solubilized thylakoid membrane samples from each of the strains (Fig. 6). Protein complexes were separated in the first
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0
4
8
12
16
20
24
Flash number
dimension by BN-PAGE, and then the constituent proteins were separated in the second dimension by denaturing SDS-PAGE. The gels were stained with SYPRO Ruby then subsequently colloidal Coomassie. After the Coomassie
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a (BN-PAGE) 1
2
3
4
b
c
(D1) 1
2
3
4
d
(D2) 1
2
3
4
e
(CP43) 1
2
3
4
(PsaA) 1
2
3
4
Supercomplexes PS I trimer PS II intermediates PS II dimer PS II intermediates PS I monomer PS II monomer RC47 Pre-cursor complexes Unassembled subunits
Fig. 5 Analysis of photosynthetic complex assembly. Solubilized thylakoid membranes run on BN-PAGE (3–12 %) (a) followed by western blotting using antibodies for the D1 (b), D2 (c), CP43 (d),
and PsaA (e) proteins. Lanes are wild type (1), DYcf48 (2), DCyanoP (3), and DCyanoP:DYcf48 (4)
staining, multiple gel spots were excised, and LC/MS/MS was used to identify their major contributing proteins (Fig. 6a–d). SYPRO ruby is a fluorescent stain that, in conjunction with a linear fluorescence detector system, allows more accurate quantitative comparison of the protein separations between mutants than can be attained with Coomassie staining (Nishihara and Champion 2002). Using the linear fluorescence intensity data, we overlaid the gel images and used false coloring to visually illustrate the quantitative differences in spot intensity between strains (Fig. 6e–g). Comparison of wild type and the CyanoP knockout did not reveal any marked changes in protein abundance; the majority of spots appear yellow, indicative of similar fluorescence emission intensities (Fig. 6e). In contrast, the DYcf48 strain displayed reduced levels (spots that appear red) of PS II dimers and monomers and an increase (spots that appear green) in early assembly complexes, consistent with those observed in the western blot analyses (Fig. 6f). These precursor complexes displayed migration in the BN-PAGE dimension consistent with the previously identified CP47* and CP43a* complexes (Komenda et al. 2004), and the presence of the PsbH and PsbY subunits belonging to the CP47* complex (or CP47 pre-complex (Boehm et al. 2011)) were confirmed by mass spectrometry. Additionally, the DYcf48 strain showed an increase in the level of cytochrome b6f dimers, the PII nitrogen regulator GlnB, and the carbon concentrating proteins CcmK1 and CcmK2. Comparison of the DYcf48 strain with the DCyanoP:DYcf48 strain revealed a small increase in the levels of PS II dimers and monomers in the double knockout (green spots for the core D1 and D2 proteins) and a reduction in the accumulation of CP47* (Fig. 6g).
Discussion Removal of CyanoP does not impair function of the mature PS II complex Knockout of CyanoP from the wild-type background did not reduce the rates of photoautotrophic growth or oxygen evolution of the cells under the conditions tested. These findings are generally consistent with earlier studies of sll1418 knockout mutants of Synechocystis 6803 grown under nutrient-replete conditions, although these earlier studies do report conflicting levels of impairment in the oxygen-evolving capacity of PS II, ranging from a 0–40 % reduction compared to wild type. (Thornton et al. 2004; Ishikawa et al. 2005; Summerfield et al. 2005). The work by Sveshnikov et al. (2007) investigated CyanoP knockout mutants from several laboratories under comparable conditions, including flash-induced oxygen evolution measurements where differences were observed in the data obtained for the DCyanoP strains and wild type. In contrast, our measurements of flash-induced oxygen evolution did not reveal any difference between CyanoP and wild type. Furthermore, our fluorescence induction measurements and the relaxation of variable fluorescence following a single-turnover actinic flash, together with corresponding kinetic analyses, show no observable alteration of electron transfer from QA to QB, PQ exchange, or recombination with the donor side. In Thermosynechococcous elongatus, the CyanoP protein has recently been shown to bind inactive monomers of PS II in a mutually exclusive position to the PsbO protein, and it was further hypothesized that CyanoP might coordinate the free C-terminus of the D1 peptide (Cormann et al. 2014). This fits with our findings that in the absence of CyanoP, even though maturation or
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Photosynth Res (2015) 124:217–229 PS I trimer PS II dimer ATP synthase NADH dehydrogenase PS I monomer PS II monomer ATP synthase RC47 Cyt b6 f dimer Cyt b6 f monomer CP47* CP43a* CP43b*
PS I trimer PS II dimer ATP synthase NADH dehydrogenase PS I monomer PS II monomer ATP synthase RC47 Cyt b6 f dimer Cyt b6 f monomer CP47* CP43a* CP43b*
226
b ΔCyanoP
wild type DnaK2 NdhH ATP synthase γ
CP47 CP43
62 49
factor Tu
38
Phycobillisome linkers
D2 D1
98
28
Cyt f
ATP synthase δ ATP synthase β ATP synthase β’
17 14
b559 α PsbH
Molecular weight (kDa)
a
6 3
c
d
ATP synthase α
ΔCyanoP:ΔYcf48
62 49
CP47 CP43
38
Cyt f
Cyt f
98
28 Cyt b6
β C-phycocyanin α C-phycocyanin
17 14
PII nitrogen regulator GlnB CcmK1, CcmK2 PsbH
b559 α
6
PsbY
e
wild type
ΔCyanoP
f
wild type
Molecular weight (kDa)
ΔYcf48
3 ΔYcf48
g
ΔYcf48
ΔCyanoP:ΔYcf48
62 49 38 28 17 14
6 3
Quenching of stain fluorescence by pigments from the PS I trimers and monomers or phycocyanin molecules
123
Reduced level of PS II dimers and monomers
Increased PS II assembly precursor complexes
Increased level of PS II dimers and monomers
Reduced CP47*
Molecular weight (kDa)
98
Photosynth Res (2015) 124:217–229 b Fig. 6 2D gel electrophoresis analysis of thylakoid membrane
protein complexes. Solubilized thylakoid membrane proteins run on BN-PAGE (4–16 %) then SDS-PAGE (12 %). Coomassie-stained images for the wild type (a), DCyanoP (b), DYcf48 (c), and DCyanoP:DYcf48 (d) and false-colored comparison of SYPRO Ruby-stained fluorescence emission for the wild type (red) and DCyanoP (green) (e), wild type (red) and DYcf48 (green) (f), and DYcf48 (red) and DCyanoP:DYcf48 (green) (g)
repair of centers may be perturbed, once PS II centers have matured beyond the point of CyanoP involvement, they are functionally indistinct from centers formed in the presence of CyanoP. Removal of CyanoP influences antenna coupling In contrast to the above, we did observe a consistent difference between DCyanoP and wild-type cells when their 77 K fluorescence emission spectra were measured using 580 nm excitation. The 680 nm emission was reduced in the DCyanoP strain, suggesting a major difference in phycobilisome coupling to PS II. Support for this hypothesis is provided by the room temperature fluorescence data obtained using measuring flashes that specifically excite phycobilisome pigments (627 nm, RMF). A decrease in emission from the dark-adapted state FO (RMF) for the DCyanoP strain was observed, whereas the opposite effect was seen for DYcf48 cells where an increase in FO (RMF) was observed (Table 2), consistent with the increased 77 K peak at 680 nm (Fig. 2c) for this strain. We also verified this phenotype in a separate, independently created DCyanoP knockout mutant (data not shown). A link between proteins found in the thylakoid lumen influencing phycobilisome coupling is also supported by Veerman et al. (2005). Precedent for an involvement of the higher plant CyanoP homolog (PsbP) in regulation of light harvesting and energy distribution has been observed where the formation and stabilization of supercomplexes containing PS II and peripheral light-harvesting complexes (LHCs) were disrupted in the absence of PsbP (Boekema et al. 2000; Ido et al. 2009; Ifuku et al. 2011). Although cyanobacteria do not contain LHCs but instead utilize phycobilisomes, using BN-PAGE or subsequent western blotting, we did not observe analogous changes in supercomplex formation in either the DCyanoP or DCyanoP:DYcf48 strains. Nevertheless, phycobilisome to PS II coupling is linked to PS II assembly and repair, and we noted a similar decrease in the 680 nm emission for a DPsb27 strain (Hwang et al. 2008; Jackson et al. 2014). Given that CyanoP is suggested to be involved in the biogenesis or repair of PS II (Cormann et al. 2014), this difference possibly reflects altered coupling between incompletely assembled PS II centers and phycobilisomes.
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Inactivation of Ycf48 dominates the phenotype of the double mutant In most of our experiments, the phenotype of the DCyanoP:DYcf48 double mutant was similar but not identical to the DYcf48 strain. Additive deleterious effects were present for the room temperature fluorescence measurements; however, the increased oxygen evolution, elevated PS II to PS I ratio, and increased levels of PS II monomer and dimer assembly in the double mutant suggest that removal of CyanoP partially alleviates an assembly block that is present in the absence of Ycf48. These findings are in contrast to the results obtained with the DYcf48:DPsb27 strain, where assembly of mature PS II centers was severely perturbed (Jackson et al. 2014). On balance, our current findings do not indicate an additive restriction for the DCyanoP:DYcf48 double mutant relative to the parental DYcf48 strain. Combined with the recent finding from Cormann et al. (2014), that CyanoP binds inactive monomers which also have Psb27 present, this suggests that Ycf48 dissociates before CyanoP binds. This hypothesis is supported by the findings of Komenda et al. (2008), who showed Ycf48 to be involved in early biogenesis but not the RC47 complex and beyond. Additionally, from our 2D gel data comparison, accumulation of a PS II precursor complex containing CP47, in the absence of Ycf48, appears reduced in the absence of both Ycf48 and CyanoP (Fig. 6c, d, g). These observations might indicate either a direct role for CyanoP earlier in biogenesis than the inactive monomer stage (but not associated with the RC core) or that biosynthesis of the CP47 pre-complex is related to the presence of CyanoP. We previously observed the DYcf48 strain to accumulate a precursor complex containing CP43 which likely gives rise to the increase in 77 K fluorescence emission at a peak of 685 nm (Jackson et al. 2014). This putative pre-complex is also evident in the DCyanoP:DYcf48 strain, demonstrating that its accumulation is not reliant on the presence of CyanoP (Figs. 5d, 6d).
Concluding remarks Our findings for the inactivation of CyanoP in the wildtype background support current models where the role of CyanoP is neither in the mature complex nor is it required for function under normal conditions. Moreover, our results are consistent with there being an altered interaction between incompletely assembled PS II centers and phycobilisomes in strains lacking CyanoP, and that this gives rise to fluorescence quenching at 680 nm in the 77 K fluorescence emission spectra when the phycobilisomes are directly excited by 580 nm light. Investigation of the role of
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CyanoP in early biogenesis in conjunction with the Ycf48 assembly factor indicates that it is unlikely there is a direct interaction between complexes containing CyanoP or Ycf48 during the biogenesis or repair of PS II. Compared to DYcf48 cells, a reduction in the PsbH- and PsbY-containing CP47 pre-complex was observed in the DCyanoP:DYcf48 strain. Furthermore, a small increase in the accumulation of PS II dimers and monomers, supported by 77 K fluorescence emission spectra and quantitative 2D gel analyses, along with an increase in oxygen evolution capacity, compared to the DYcf48 single mutant, suggests that the CyanoP protein impedes PS II assembly in the absence of Ycf48. Acknowledgments S.A.J. was supported by a University of Otago Division of Health Sciences Career Development Postdoctoral Fellowship. Other funding for this project was provided by an Otago University Research grant to J.E.-R. We thank Asher J. Dale for assistance with preliminary experiments.
References Boehm M, Romero E, Reisinger V, Yu J, Komenda J, Eichacker LA, Dekker JP, Nixon PJ (2011) Investigating the early stages of Photosystem II assembly in Synechocystis sp. PCC 6803: isolation of CP47 and CP43 complexes. J Biol Chem 286:14812–14819 Boekema EJ, Van Breemen JF, Van Roon H, Dekker JP (2000) Conformational changes in Photosystem II supercomplexes upon removal of extrinsic subunits. Biochemistry 39:12907–12915 Bricker TM, Roose JL, Fagerlund RD, Frankel LK, Eaton-Rye JJ (2012) The extrinsic proteins of Photosystem II. Biochim Biophys Acta 1817:121–142 Cormann KU, Bangert JA, Ikeuchi M, Ro¨gner M, Stoll R, Nowaczyk MM (2009) Structure of Psb27 in solution: implications for transient binding to Photosystem II during biogenesis and repair. Biochemistry 48:8768–8770 Cormann KU, Bartsch M, Ro¨gner M, Nowaczyk MM (2014) Localization of the CyanoP binding site on Photosystem II by surface plasmon resonance spectroscopy. Front Plant Sci. doi:10. 3389/fpls.2014.00595 Cser K, Vass I (2007) Radiative and non-radiative charge recombination pathways in Photosystem II studied by thermoluminescence and chlorophyll fluorescence in the cyanobacterium Synechocystis 6803. Biochim Biophys Acta 1767:233–243 Eaton-Rye JJ (2011) Construction of gene interruptions and gene deletions in the cyanobacterium Synechocystis sp. strain PCC 6803. Methods Mol Biol 684:295–312 Fagerlund RD, Eaton-Rye JJ (2011) The lipoproteins of cyanobacterial Photosystem II. J Photochem Photobiol, B 104:191–203 Hwang HJ, Nagarajan A, McLain A, Burnap RL (2008) Assembly and disassembly of the Photosystem II manganese cluster reversibly alters the coupling of the reaction center with the light-harvesting phycobilisome. Biochemistry 47:9747–9755 Ido K, Ifuku K, Yamamoto Y, Ishihara S, Murakami A, Takabe K, Miyake C, Sato F (2009) Knockdown of the PsbP protein does not prevent assembly of the dimeric PSII core complex but impairs accumulation of Photosystem II supercomplexes in tobacco. Biochim Biophys Acta 1787:873–881
123
Photosynth Res (2015) 124:217–229 Ifuku K, Ido K, Sato F (2011) Molecular functions of PsbP and PsbQ proteins in the Photosystem II supercomplex. J Photochem Photobiol, B 104:158–164 Ishikawa Y, Schro¨der WP, Funk C (2005) Functional analysis of the PsbP-like protein (sll1418) in Synechocystis sp. PCC 6803. Photosynth Res 84:257–262 Jackson SA, Fagerlund RD, Wilbanks SM, Eaton-Rye JJ (2010) Crystal structure of PsbQ from Synechocystis sp. PCC 6803 at ˚ : implications for binding and function in cyanobacterial 1.8 A Photosystem II. Biochemistry 49:2765–2767 Jackson SA, Hinds MG, Eaton-Rye JJ (2012) Solution structure of CyanoP from Synechocystis sp. PCC 6803: new insights on the structural basis for functional specialization amongst PsbP family proteins. Biochim Biophys Acta 1817:1331– 1338 Jackson SA, Hervey JR, Dale AJ, Eaton-Rye JJ (2014) Removal of both Ycf48 and Psb27 in Synechocystis sp. PCC 6803 disrupts Photosystem II assembly and alters QA oxidation in the mature complex. FEBS Lett 588:3751–3760 Joliot A, Joliot P (1964) Etude cine´tique de la re´action photochimique libe´rant l’oxyge`ne au cours de la photosynthe`se. C R Acad Sci Paris 258:4622–4635 Joliot P, Joliot A (1968) A polarographic method for detection of oxygen production and reduction of Hill reagent by isolated chloroplasts. Biochim Biophys Acta 153:625–634 Komenda J, Reisinger V, Mu¨ller BC, Doba´kova´ M, Granvogl B, Eichacker LA (2004) Accumulation of the D2 protein is a key regulatory step for assembly of the Photosystem II reaction center complex in Synechocystis PCC 6803. J Biol Chem 279:48620–48629 Komenda J, Nickelsen J, Tichy´ M, Pra´sˇil O, Eichacker LA, Nixon PJ (2008) The cyanobacterial homologue of HCF136/YCF48 is a component of an early Photosystem II assembly complex and is important for both the efficient assembly and repair of Photosystem II in Synechocystis sp. PCC 6803. J Biol Chem 283:22390–22399 Komenda J, Sobotka R, Nixon PJ (2012) Assembling and maintaining the Photosystem II complex in chloroplasts and cyanobacteria. Curr Opin Plant Biol 15:245–251 Mabbitt PD, Rautureau GJ, Day CL, Wilbanks SM, Eaton-Rye JJ, Hinds MG (2009) Solution structure of Psb27 from cyanobacterial Photosystem II. Biochemistry 48:8771–8773 Mabbitt PD, Wilbanks SM, Eaton-Rye JJ (2014) Structure and function of the hydrophilic Photosystem II assembly proteins: Psb27, Psb28 and Ycf48. Plant Physiol Biochem 81:96–107 Messinger J, Schro¨der WP, Renger G (1993) Structure-function relations in Photosystem II. Effects of temperature and chaotropic agents on the period four oscillation of flash-induced oxygen evolution. Biochemistry 32:7658–7668 Meunier PC, Popovic R (1988) High-accuracy oxygen polarograph for photosynthetic systems. Rev Sci Instrum 59:486–491 Michoux F, Takasaka K, Boehm M, Nixon PJ, Murray JW (2010) ˚ : implications for the evolution and Structure of CyanoP at 2.8 A function of the PsbP subunit of Photosystem II. Biochemistry 49:7411–7413 Michoux F, Boehm M, Bialek W, Takasaka K, Maghlaoui K, Barber J, Murray JW, Nixon PJ (2014) Crystal structure of CyanoQ from the thermophilic cyanobacterium Thermosynechococcus elongatus and detection in isolated Photosystem II complexes. Photosynth Res 122:57–67 Morris JN, Crawford TS, Jeffs A, Stockwell PA, Eaton-Rye JJ, Summerfield TC (2014) Whole genome re-sequencing of two ‘wild-type’ strains of the model cyanobacterium Synechocystis sp. PCC 6803. N Z J Bot 52:36–47 Nickelsen J, Rengstl B (2013) Photosystem II assembly: from cyanobacteria to plants. Annu Rev Plant Biol 64:609–635
Photosynth Res (2015) 124:217–229 Nishihara JC, Champion KM (2002) Quantitative evaluation of proteins in one- and two-dimensional polyacrylamide gels using a fluorescent stain. Electrophoresis 23:2203–2215 Nixon PJ, Michoux F, Yu J, Boehm M, Komenda J (2010) Recent advances in understanding the assembly and repair of Photosystem II. Ann Bot 106:1–16 Renger G, Hanssum B (2009) Oxygen detection in biological systems. Photosynth Res 102:487–498 Strasser RJ, Srivastava A, Govindjee (1995) Polyphasic chlorophylla fluorescence transient in plants and cyanobacteria. Photochem Photobiol 61:32–42 Suga M, Akita F, Hirata K, Ueno G, Murakami H, Nakajima Y, Shimizu T, Yamashita K, Yamamoto M, Ago H, Shen J-R ˚ resolution (2015) Native structure of Photosystem II at 1.94 A viewed by femotosecond X-ray pulses. Nature 517:99–103 Summerfield TC, Winter RT, Eaton-Rye JJ (2005) Investigation of a requirement for the PsbP-like protein in Synechocystis sp. PCC 6803. Photosynth Res 84:263–268 Sveshnikov D, Funk C, Schro¨der WP (2007) The PsbP-like protein (sll1418) of Synechocystis sp. PCC 6803 stabilises the donor side of Photosystem II. Photosynth Res 93:101–109
229 Thornton LE, Ohkawa H, Roose JL, Kashino Y, Keren N, Pakrasi HB (2004) Homologs of plant PsbP and PsbQ proteins are necessary for regulation of Photosystem II activity in the cyanobacterium Synechocystis 6803. Plant Cell 16:2164–2175 Umena Y, Kawakami K, Shen J-R, Kamiya N (2011) Crystal structure of oxygen-evolving Photosystem II at a resolution of ˚ . Nature 473:55–60 1.9 A Veerman J, Bentley FK, Eaton-Rye JJ, Mullineaux CW, Vasil’ev S, Bruce D (2005) The PsbU subunit of Photosystem II stabilizes energy transfer and primary photochemistry in the phycobilisome-photosystem II assembly of Synechocystis sp. PCC 6803. Biochemistry 44:16939–16948 Vinyard DJ, Ananyev GM, Dismukes GC (2013) Photosystem II: the reaction center of oxygenic photosynthesis. Annu Rev Biochem 82:577–606 Williams JGK (1988) Construction of specific mutations in Photosystem-II photosynthetic reaction center by genetic-engineering methods in Synechocystis-6803. Methods Enzymol 167:766–778
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