TY - JOUR
AU1 - Haniewicz, Patrycja
AU2 - Abram, Mateusz
AU3 - Nosek, Luká¡
AU4 - Kirkpatrick, Joanna
AU5 - El-Mohsnawy, Eithar
AU6 - Olmos, Julian D. Janna
AU7 - Kouřil, Roman
AU8 - Kargul, Joanna M.
AB - Abstract The monomeric photosystem I-light-harvesting antenna complex I (PSI-LHCI) supercomplex from the extremophilic red alga Cyanidioschyzon merolae represents an intermediate evolutionary link between the cyanobacterial PSI reaction center and its green algal/higher plant counterpart. We show that the C. merolae PSI-LHCI supercomplex is characterized by robustness in various extreme conditions. By a combination of biochemical, spectroscopic, mass spectrometry, and electron microscopy/single particle analyses, we dissected three molecular mechanisms underlying the inherent robustness of the C. merolae PSI-LHCI supercomplex: (1) the accumulation of photoprotective zeaxanthin in the LHCI antenna and the PSI reaction center; (2) structural remodeling of the LHCI antenna and adjustment of the effective absorption cross section; and (3) dynamic readjustment of the stoichiometry of the two PSI-LHCI isomers and changes in the oligomeric state of the PSI-LHCI supercomplex, accompanied by dissociation of the PsaK core subunit. We show that the largest low light-treated C. merolae PSI-LHCI supercomplex can bind up to eight Lhcr antenna subunits, which are organized as two rows on the PsaF/PsaJ side of the core complex. Under our experimental conditions, we found no evidence of functional coupling of the phycobilisomes with the PSI-LHCI supercomplex purified from various light conditions, suggesting that the putative association of this antenna with the PSI supercomplex is absent or may be lost during the purification procedure. Extremophiles have evolved the remarkable strategies that allow them to thrive beyond some daunting physical and chemical limits of life on Earth. It is important to understand the molecular mechanisms that define these limits of life under extreme conditions. Dissecting the molecular mechanisms of adaptation to these challenging environmental conditions is crucial for understanding how life may have evolved and survived in the early history of our planet. The emergence of oxygenic photosynthesis in cyanobacteria over 2.5 billion years ago is often dubbed the Big Bang of evolution (Barber, 2004), as it gave rise to an aerobic atmosphere and protective ozone layer and allowed efficient aerobic cellular respiration and the colonization of Earth’s surface by metazoan life. As such, it triggered fundamental biosphere changes on an unprecedented scale. In natural photosynthesis, the absorption of two quanta of light triggers the primary charge separation in the reaction centers (RCs) of PSII and PSI followed by the vectorial electron flow from PSII to PSI via the cytochrome b 6 f complex, with the concomitant release of protons and molecular oxygen. With the input of four photons, initially absorbed by chlorophyll (Chl) molecules, the catalytic metal center of PSII accumulates four oxidizing equivalents required to produce a dioxygen molecule from two substrate water molecules (Babcock et al., 1989; Kargul and Barber, 2011). Concomitantly, PSI upon absorption of the second photon provides energy-rich electrons to reduce the final acceptors ferredoxin and NADP+ to NADPH via ferredoxin-NADP reductase. The photosynthetic apparatus of extremophilic microalgae has gained considerable interest due to the exceptionally high enzymatic stability and activity of its photoelectroactive components, making them attractive for numerous applications in fields ranging from structural biology (Klukas et al., 1999; Jordan et al., 2001; Zouni et al., 2001; Kamiya and Shen, 2003; Ferreira et al., 2004; Loll et al., 2005; Adachi et al., 2009; Guskov et al., 2009; Amunts et al., 2010; Umena et al., 2011; Suga et al., 2015; Ago et al., 2016) to biotechnology (León-Bañares et al., 2004) and biophotovoltaics (Krassen et al., 2009; Iwuchukwu et al., 2010; Utschig et al., 2011; Kargul et al., 2012; Mershin et al., 2012; Gordiichuk et al., 2014; Ocakoglu et al., 2014; Janna Olmos and Kargul, 2015; Olmos et al., 2017; Szalkowski et al., 2017). These applications stemmed from the use of robust photosystems purified to homogeneity from thermophilic prokaryotic cyanobacteria or eukaryotic thermoacidophilic red microalgae. On the other hand, extremophilic red microalgae have been the favorite model organisms in which to study the evolution of fundamental processes of cell division and intracellular transport (Kuroiwa, 1998; Kuroiwa et al., 1998; Miyagishima et al., 2003; Nishida et al., 2003) as well as, very recently, the evolution and function of the photosynthetic apparatus through structural, genomic, biochemical, spectroscopic, and mass spectrometric approaches (Adachi et al., 2009; Vanselow et al., 2009; Busch et al., 2010; Krupnik et al., 2013; Nilsson et al., 2014; Ago et al., 2016). The photosynthetic apparatus of the red thermoacidophilic microalga Cyanidioschyzon merolae has gained considerable interest, due to the unique evolutionary positioning of this species near the root of the red algal lineage that forms a basal group within the eukaryotes and diverged ∼1.3 billion years ago within the most ancient algal order of Cyanidiales (Nozaki et al., 2003; Reeb and Bhattacharya, 2010). It is considered as an evolutionary intermediate link between the photosynthetic apparatus of prokaryotic cyanobacteria and that of the eukaryotic phototrophs of the green lineage (Ohta et al., 2003; Busch and Hippler, 2011; Kargul et al., 2012). As such, it combines several prokaryotic and eukaryotic structural traits. In particular, PSII displays predominantly prokaryotic and some eukaryotic features. The prokaryotic characteristics include the presence of cyanobacterial-like phycobilisomes (PBSs), functioning as the peripheral light-harvesting antenna, as well as the presence of cyanobacterial subunits stabilizing the catalytic center of PSII, PsbV, and PsbU in addition to the evolutionarily conserved PsbO subunit. The fourth subunit unique to red algae that is positioned within the oxygen-evolving complex is the PsbQ′ protein (Krupnik et al., 2013; Ago et al., 2016). The red algal PSI-LHCI supercomplex is reminiscent of its higher plant and green algal counterparts in that it comprises the monomeric RC core complex composed of 13 subunits (PsaA–PsaF and PsaI–PsaO; Jensen et al., 2007; Vanselow et al., 2009) and is associated with an asymmetrically located, crescent-like peripheral light-harvesting antenna complex (LHCI) composed of a variable number of Chla-binding Lhcr subunits, depending on the species (Tan et al., 1997; Busch et al., 2010; Thangaraj et al., 2011; Tian et al., 2017a). The analysis of the Galdieria sulphuraria plastid genome suggests that the red algal PSI may have evolved even earlier than the present-day cyanobacterial, green algal, and higher plant counterparts (Vanselow et al., 2009). The interesting features of the red algal PSI are the retention of the cyanobacterial PsaM subunit and the lack of higher plant and green algal PsaH and PsaG subunits implied in both the docking of the mobile LHCII antenna during state transitions and the formation of the LHCI belt, respectively (Kargul et al., 2012). Moreover, the chimeric nature of the two core subunits PsaF and PsaL, accommodating both cyanobacterial and higher plant-like structural domains, further supports the evolutionarily intermediate character of the red algal PSI-LHCI supercomplex (Busch and Hippler, 2011; Kargul et al., 2012). Recently, we provided, to our knowledge, the first direct evidence that the C. merolae PSII complex employs two distinct molecular mechanisms of photoprotection upon exposure to high light (HL). These are the accumulation of the carotenoid zeaxanthin (Zea) in thylakoids and dimeric PSII complexes together with a reversible RC-based nonphotochemical quenching that is triggered by the acidification of thylakoid lumen upon exposure to HL intensities (Krupnik et al., 2013). Both features are likely to provide the basis for the remarkable robustness and sustained high oxygen-evolving activity of the C. merolae PSII across a wide range of extreme light, temperature, and pH conditions (Krupnik et al., 2013). In this study, we extended the mechanistic and structural investigation of the extremophilic red algal photosynthetic apparatus to the second photosystem of C. merolae, the PSI-LHCI supercomplex, in order to gain an insight into the molecular basis of the exceptional robustness of this complex upon its exposure to various extreme conditions. We provide several lines of evidence that the high photochemical activity and stability of the C. merolae PSI-LHCI supercomplex are due to the combined protective effects of Zea accumulation within this complex, changes of the oligomeric state of this complex, as well as dynamic structural remodeling of the LHCI antenna upon exposure to changing light conditions. RESULTS AND DISCUSSION Biochemical and Proteomic Characterization of the C. merolae PSI-LHCI Supercomplex Isolated from Varying Light Regimes The main aim of our study was to dissect the molecular mechanisms of photoadaptation of the C. merolae PSI-LHCI supercomplex. To this end, we purified the highly homogenous PSI-LHCI preparations by detergent-based solubilization of thylakoids obtained from cells grown in four distinct light regimes: low (LL; 35 µE m−2 s−1), medium (ML; 90 µE m−2 s−1), high (HL; 150 µE m−2 s−1), and extreme high (EHL; 350 µE m−2 s−1) white light irradiation. We applied three-step anion-exchange chromatography (AEC) and size-exclusion chromatography (SEC) to obtain pure supercomplex preparations. Figure 1 shows the typical AEC chromatograms (identical for all four light regimes) and SDS-PAGE protein profiles, confirming that the all four PSI-LHCI samples were purified to homogeneity and contained the typical core (e.g. PsaA/PsaB heterodimer and smaller core subunits) and Lhcr antenna subunits resolved as two protein bands by SDS-PAGE and identified by mass spectrometry in all four PSI-LHCI preparations (Table I). Figure 1. Open in new tabDownload slide Purification and biochemical characterization of LL, ML, HL, and EHL supercomplexes. A, AEC chromatogram from the first step of the C. merolae PSI-LHCI supercomplex purification procedure on a DEAE TOYOPEARL 650M column. Insets I and II show RT absorption spectra of PSI and PSII fractions, respectively. B, AEC chromatogram from the second step of the C. merolae PSI-LHCI supercomplex purification procedure on a DEAE TOYOPEARL 650S column. Inset III shows a RT absorption spectrum of the PSI-LHCI supercomplex after the second AEC step. C, SDS-PAGE protein profiles of T. elongatus PSI trimer and C. merolae PSI-LHCI supercomplexes purified from four different light regimes (LL, ML, HL, and EHL). D, Photochemical activity of the C. merolae PSI-LHCI supercomplex purified from four different light regimes compared with the activity of the T. elongatus PSI trimer from ML. Figure 1. Open in new tabDownload slide Purification and biochemical characterization of LL, ML, HL, and EHL supercomplexes. A, AEC chromatogram from the first step of the C. merolae PSI-LHCI supercomplex purification procedure on a DEAE TOYOPEARL 650M column. Insets I and II show RT absorption spectra of PSI and PSII fractions, respectively. B, AEC chromatogram from the second step of the C. merolae PSI-LHCI supercomplex purification procedure on a DEAE TOYOPEARL 650S column. Inset III shows a RT absorption spectrum of the PSI-LHCI supercomplex after the second AEC step. C, SDS-PAGE protein profiles of T. elongatus PSI trimer and C. merolae PSI-LHCI supercomplexes purified from four different light regimes (LL, ML, HL, and EHL). D, Photochemical activity of the C. merolae PSI-LHCI supercomplex purified from four different light regimes compared with the activity of the T. elongatus PSI trimer from ML. Mass spectrometry analysis of the BN-PAGE protein bands obtained from LL, ML, HL, and EHL C. merolae PSI-LHCI samples Table I. Mass spectrometry analysis of the BN-PAGE protein bands obtained from LL, ML, HL, and EHL C. merolae PSI-LHCI samples Shown are the subunits identified in BN-PAGE bands in Fig. 3. The relative contributions are represented by crosses, where four crosses represent proteins with iBAQ values (the sums of intensities of all tryptic peptides for each protein divided by the number of theoretically observable peptides) ≥ 1.75e10, three crosses ≥ 2.5e9, two crosses ≥ 5e8, and one cross ≥ 2.5e6. The negligible contribution of a protein is indicated with a minus sign, where the iBAQ value was < 2.5e6. Open in new tab Table I. Mass spectrometry analysis of the BN-PAGE protein bands obtained from LL, ML, HL, and EHL C. merolae PSI-LHCI samples Shown are the subunits identified in BN-PAGE bands in Fig. 3. The relative contributions are represented by crosses, where four crosses represent proteins with iBAQ values (the sums of intensities of all tryptic peptides for each protein divided by the number of theoretically observable peptides) ≥ 1.75e10, three crosses ≥ 2.5e9, two crosses ≥ 5e8, and one cross ≥ 2.5e6. The negligible contribution of a protein is indicated with a minus sign, where the iBAQ value was < 2.5e6. Open in new tab The photochemical activity of all four PSI-LHCI preparations (Fig. 1D) was in the range of 538 to 1,331 µmol oxygen consumed mg−1 Chl h−1, with the highest value obtained for the EHL PSI-LHCI supercomplex. Overall, the activity of the red algal PSI-LHCI complex was 1.3- to 3.2-fold higher than the activity of the trimeric PSI complex purified from the thermophilic cyanobacterium Thermosynechococcus elongatus, using an analogous purification procedure and the same experimental conditions for the oxygen consumption measurement (Fig. 1D). To further probe the robustness of the C. merolae PSI-LHCI supercomplex, we subjected the sample obtained from ML to a wide range of extreme temperature, light, and pH conditions. The choice of the ML sample was to avoid any putative detrimental effects of stress conditions prior to investigation of the robustness of the C. merolae PSI-LHCI supercomplex. Figure 2 shows that the C. merolae PSI-LHCI supercomplex retains most of its photochemical activity when exposed to light intensities as high as 25,000 µE m−2 s−1, temperature up to 80°C, and a pH range from 4 to 12. To our knowledge, this is the most robust PSI-LHCI supercomplex reported to date, even when compared directly with other relatively stable PSI complexes such as the PSI trimer from the thermophilic cyanobacterium T. elongatus (Fig. 1D), from which a near atomic x-ray structure was obtained (Jordan et al., 2001). The wide temperature tolerance of isolated C. merolae PSII (Krupnik et al., 2013) and PSI-LHCI complexes (this study) is most likely the reason for the remarkable wide temperature range tolerance of this extremophilic alga, as shown in a recent study (Nikolova et al., 2017). Figure 2. Open in new tabDownload slide Robustness of C. merolae PSI-LHCI in extreme conditions. A, Photochemical activity of the PSI-LHCI ML supercomplex in different light regimes. B, Photochemical activity of the PSI-LHCI supercomplex exposed to various temperatures. C, Photochemical activity of the PSI-LHCI supercomplex in the pH range 4 to 12. Figure 2. Open in new tabDownload slide Robustness of C. merolae PSI-LHCI in extreme conditions. A, Photochemical activity of the PSI-LHCI ML supercomplex in different light regimes. B, Photochemical activity of the PSI-LHCI supercomplex exposed to various temperatures. C, Photochemical activity of the PSI-LHCI supercomplex in the pH range 4 to 12. The C. merolae PSI-LHCI supercomplex exists as a monomer, similar to its other eukaryotic counterparts (Busch et al., 2010; Drop et al., 2011; Mazor et al., 2015; Qin et al., 2015; Tian et al., 2017a). An interesting observation is the presence of two native bands from this complex that were identified by blue native (BN)-PAGE in LL/ML (50 µE m−2 s−1) C. merolae complexes, varying in the core and possibly LHCI subunit composition (Tian et al., 2017a). This prompted us to examine the putative changes in the relative abundance of both isomers in response to changing illumination conditions. Figure 3 shows that the overall average size of the PSI-LHCI supercomplex decreases gradually upon increasing light intensity (Fig. 3A). In agreement with this observation, the relative abundance of the smaller and larger PSI-LHCI isomer (identified as band 1 and band 2, respectively, by BN-PAGE; Fig. 3B) changes dynamically in response to varying light conditions, with the amount of the larger form of this complex diminished severely in HL and EHL (Fig. 3, B and C) compared with LL and ML conditions. We observed an additional band (band 3; Fig. 3B) composed of the oligomeric forms of the PSI-LHCI supercomplex of varying abundance depending on the light regime used, with the larger oligomers observed preferentially under LL conditions. Figure 3. Open in new tabDownload slide Oligomeric state of the LL, ML, HL, and EHL C. merolae PSI-LHCI supercomplexes. A, SEC analysis of PSI-LHCI samples purified from four different light regimes. Yellow, ML PSI trimer from T. elongatus; blue, C. merolae LL PSI-LHCI; green, ML C. merolae PSI-LHCI; gray, C. merolae HL PSI-LHCI; red, C. merolae EHL PSI-LHCI. B, BN-PAGE analysis of the C. merolae PSI-LHCI supercomplex purified from four different light regimes. C, Densitometric analysis of the bands from B. Figure 3. Open in new tabDownload slide Oligomeric state of the LL, ML, HL, and EHL C. merolae PSI-LHCI supercomplexes. A, SEC analysis of PSI-LHCI samples purified from four different light regimes. Yellow, ML PSI trimer from T. elongatus; blue, C. merolae LL PSI-LHCI; green, ML C. merolae PSI-LHCI; gray, C. merolae HL PSI-LHCI; red, C. merolae EHL PSI-LHCI. B, BN-PAGE analysis of the C. merolae PSI-LHCI supercomplex purified from four different light regimes. C, Densitometric analysis of the bands from B. To get an insight into the intriguing dynamic changes of abundance of the three native bands, we examined their precise subunit composition by mass spectrometry. Most of the core subunits (except for PsaI) and all the three Lhcr subunits were identified in all the C. merolae PSI-LHCI isomers and oligomers at all the light intensities applied (Table I). The EHL band 3 contained the same subunits as the EHL band 2, except for the PBS rod-core linker polypeptide, which was absent in the EHL oligomers (Table I). In contrast to the recent study of Tian et al. (2017a), the core subunits PsaF and PsaO were associated with both bands rather stably in all the light regimes. The PsaO subunit is absent in the recent x-ray structures of the higher plant PSI-LHCI supercomplex, and it has been postulated to be associated loosely with the PSI core complex (Mazor et al., 2015, 2017; Qin et al., 2015). The presence of PsaO in the C. merolae supercomplex, therefore, may reflect differences in its interaction with other core subunits in comparison with the higher plant complex. The PsaF subunit, apart from its well-established role as the docking site for the mobile electron carriers (cytochrome c 553 in C. merolae; for review, see Kargul et al., 2012), has been postulated as the putative binding site for one of the LHCI subunits in the C. merolae PSI-LHCI supercomplex (Tian et al., 2017a), although this remains to be directly confirmed by structural data. Of all the core subunits, the PsaK protein seems to exhibit the largest variability in its abundance in both PSI-LHCI isomers and oligomers (Table I). In fact, this subunit was missing in both LL bands as well as in a larger supercomplex isomer and oligomers isolated from EHL conditions (Table I), suggesting that it is associated rather loosely with the PSI core in C. merolae. In higher plants, the PsaK subunit interacts with PsaB on the opposite side from a homologous PsaG subunit (absent in C. merolae) that forms the docking site for the Lhca1 antenna protein (Mazor et al., 2015, 2017; Qin et al., 2015). The PsaK subunit has been postulated to exist in two copies in another red microalga, G. sulphuraria (Vanselow et al., 2009). However, the precise positioning of the second copy remains to be established. Therefore, the absence of PsaK in the larger isoform of the C. merolae PSI-LHCI supercomplex in LL and EHL conditions may reflect its close interaction with one of the Lhcr subunits and their co-dissociation following significant structural changes in PsaK. Such conformational changes were postulated in the latest x-ray structure of the higher plant PSI-LHCI supercomplex during light-induced remodeling of the LHCI antenna (Mazor et al., 2017). Spectroscopic Investigation of the C. merolae Light-Harvesting Antenna Interaction with the PSI RC under Variable Light Conditions Spectroscopic analysis of the LL, ML, HL, and EHL PSI-LHCI supercomplex samples showed the typical absorbance red maxima at 678.5 nm and 77K red-shifted emission peaks at 726 to 729.5 nm following excitation of Chla at 435 nm (Fig. 4, A and B, respectively), due to the presence of red Chls postulated to serve as intermediate energy traps for excitations transferred to the P700 RC (Werst et al., 1992). Importantly, the EHL PSI-LHCI sample showed a 3.5-nm red shift of the 77K emission maximum compared with LL samples (Fig. 4B), most likely due to the accumulation of additional red Chls in the supercomplex upon exposure to extreme HL. Figure 4. Open in new tabDownload slide Spectroscopic characterization of the LL, ML, HL, and EHL C. merolae PSI-LHCI supercomplexes. A, RT absorption spectra of PSI-LHCI samples purified from four different light regimes. Yellow, ML PSI trimer from T. elongatus; blue, C. merolae LL PSI-LHCI; green, ML C. merolae PSI-LHCI; gray, C. merolae HL PSI-LHCI; red, C. merolae EHL PSI-LHCI. B, 77K emission spectra of PSI-LHCI complexes from four different light regimes, excited at 435 nm. C, 77K emission spectra of complexes from four different light regimes, excited at 600 nm. D, 77K excitation spectra of all the PSI-LHCI samples with emission recorded at 728 nm. Figure 4. Open in new tabDownload slide Spectroscopic characterization of the LL, ML, HL, and EHL C. merolae PSI-LHCI supercomplexes. A, RT absorption spectra of PSI-LHCI samples purified from four different light regimes. Yellow, ML PSI trimer from T. elongatus; blue, C. merolae LL PSI-LHCI; green, ML C. merolae PSI-LHCI; gray, C. merolae HL PSI-LHCI; red, C. merolae EHL PSI-LHCI. B, 77K emission spectra of PSI-LHCI complexes from four different light regimes, excited at 435 nm. C, 77K emission spectra of complexes from four different light regimes, excited at 600 nm. D, 77K excitation spectra of all the PSI-LHCI samples with emission recorded at 728 nm. The presence of the red-absorbing Chls is a characteristic feature of PSI (Morosinotto et al., 2005; Wientjes et al., 2012; Croce and van Amerongen, 2013). The red energy traps have been shown to slow down the rate of energy trapping in P700 through alteration of the kinetics of excitation energy transfer pathways from the peripheral and core antennae to the RC of the eukaryotic and cyanobacterial PSI complexes (Croce et al., 2000; Gobets and van Grondelle, 2001; Ihalainen et al., 2002; Jennings et al., 2003; Gibasiewicz et al., 2005; Melkozernov et al., 2005; Engelmann et al., 2006; Snellenburg et al., 2013). The slower fluorescence decay kinetics evoked by the red traps is due mainly to the uphill energy transfer from the low-energy forms to the bulk and/or inner core Chl molecules (Croce et al., 2000; Jennings et al., 2003). The physiological role of red traps is a matter of debate, although their plausible role may be to increase the PSI absorption cross section in a shaded environment or in the conditions favoring cyclic electron flow around PSI (Rivadossi et al., 1999). Red Chls have been localized mainly in the LHCI complex by thermal broadening spectroscopic analyses (Croce et al., 1996, 1998), which has been confirmed directly by the latest x-ray crystallography studies of the higher plant PSI-LHCI supercomplex (Mazor et al., 2015; Qin et al., 2015). However, it is not inconceivable that they also may accumulate in the C. merolae PSI core complex in response to light stress, where they may affect the kinetics and pathways of energy transfer to the P700 trap. Experiments to verify this hypothesis are currently under way. A common observation was that, in all but the last chromatographic step, a significant amount of PBSs, which serve as the peripheral antenna of PSII in C. merolae (Krupnik et al., 2013), was detected. To probe the putative functional coupling of the residual PBSs, identified by mass spectrometry in the final pure PSI-LHCI samples (Table I) with the PSI RC of C. merolae, we measured the emission spectra of all four final PSI-LHCI samples upon their excitation at 600 nm, a wavelength that selectively excites phycocyanin (a pigment present in the PBS antenna). Figure 4C shows a small 722-nm emission peak corresponding to the PSI-LHCI supercomplex. However, this peak was extremely small, due to the lack of energy transfer between PBSs and the PSI RC at all the light regimes applied in this study. To further probe the energy transfer between the peripheral antennae and the PSI RCs, we measured the excitation spectra of all four ultra-pure PSI-LHCI samples following excitation at 400 to 700 nm and recording emission at 728 nm. The excitation spectra (Fig. 4D) corresponded to the pure PSI-LHCI supercomplexes containing excitonically coupled LHCI antennae, albeit with no energy transfer detected between PBSs and the PSI RC for all the PSI-LHCI samples analyzed. Overall, we found no spectroscopic evidence of a functional association of PBSs with the C. merolae PSI RC in all four light regimes applied in this study. This observation is in contrast with the spectroscopic and proteomic results reported by Hippler and colleagues (Busch et al., 2010), who postulated the physical and functional association of a small fraction of PBSs with a subpool of the LL C. merolae PSI-LHCI supercomplex. This discrepancy is most likely due to the higher homogeneity of the supercomplex samples obtained in this study, because of the more stringent purification procedure, and the possible dissociation of the putative PBS fraction from the PSI-LHCI samples. However, we cannot exclude that such a functional association may occur in vivo and is lost during the purification procedure. In fact, during the purification of all four PSI-LHCI samples, we observed a high degree of heterogeneity of the photosystem complexes, with some PSI-LHCI fractions enriched significantly in PBSs (data not shown). Whether they may form functionally coupled PBS-PSI-LHCI assemblies remains to be established. Photoprotective Role of Zea Accumulating in the C. merolae PSI-LHCI Supercomplex upon Exposure to HL In this study, we examined the putative photoprotective roles of carotenoids accumulating in the C. merolae cells and PSI-LHCI supercomplex preparations upon exposure to various light conditions. An important issue was to determine the precise loci of the carotenoid accumulation within the PSI-LHCI supercomplex upon its exposure to various light regimes. To this end, we separated the LHCI antenna from the PSI core particles by the detergent treatment followed by Suc gradient fractionation of the solubilized complexes (Fig. 5, A and B). Figure 5. Open in new tabDownload slide Biochemical and spectroscopic analyses of LHCI-depleted PSI core complexes and LHCI antennae from the LL, ML, HL, and EHL C. merolae PSI-LHCI supercomplexes. A, Separation of LHCI-depleted PSI core complexes (F2) and LHCI antennae (F1) by Suc gradient ultracentrifugation. B, Coomassie Blue-stained SDS-PAGE protein profiles for the fractions from the Suc gradient from LL, ML, HL, and EHL samples. C, Representative 77K emission spectra of PSI-LHCI, LHCI, and LHCI-depleted PSI excited at 435 nm and emission at 728 nm following 400- to 700-nm excitation. For clarity, only ML emission spectra are shown. D, Table with the Qy band absorption and main peaks in 77K emission spectra of PSI-LHCI, LHCI, and LHCI-depleted PSI fractions excited at 435 nm and emission at 728 nm for all four light regimes. Figure 5. Open in new tabDownload slide Biochemical and spectroscopic analyses of LHCI-depleted PSI core complexes and LHCI antennae from the LL, ML, HL, and EHL C. merolae PSI-LHCI supercomplexes. A, Separation of LHCI-depleted PSI core complexes (F2) and LHCI antennae (F1) by Suc gradient ultracentrifugation. B, Coomassie Blue-stained SDS-PAGE protein profiles for the fractions from the Suc gradient from LL, ML, HL, and EHL samples. C, Representative 77K emission spectra of PSI-LHCI, LHCI, and LHCI-depleted PSI excited at 435 nm and emission at 728 nm following 400- to 700-nm excitation. For clarity, only ML emission spectra are shown. D, Table with the Qy band absorption and main peaks in 77K emission spectra of PSI-LHCI, LHCI, and LHCI-depleted PSI fractions excited at 435 nm and emission at 728 nm for all four light regimes. As expected, the isolated LHCI antenna displayed significant absorption in the 400-500 nm region and the 77K Chla emission peak at 682-684 nm (Fig. 5, C and D), arising from the enrichment of this fraction with carotenoids and the depletion of some red Chls (due to the detergent treatment), respectively. The LHCI-depleted PSI fraction was characterized by a 4.5-6-nm blue shift of the 77K Chla emission peak compared with the intact PSI-LHCI supercomplex (Fig. 5, C and D). This is most likely due to the loss of a fraction of the red Chls, which are associated predominantly with the LHCI antenna (Mazor et al., 2015, 2017; Qin et al., 2015). Two main peaks of 722 and 675 nm were present in the 77K emission spectra of the LHCI-depleted PSI fractions (for ML samples; Fig. 5C). We interpret this observation by the presence of tightly bound LHCI antenna whose energetic coupling with the PSI RC is destabilized due to the detergent and freeze/thaw treatment during separation of the LHCI antenna. These spectroscopic results indicate the heterogeneity of Lhcrs present in the C. merolae PSI-LHCI supercomplex in terms of varying amounts of associated red Chls. The red peak of 722 nm in the LHCI-depleted core fraction points to the presence of red Chls also in the core complex, similar to the cyanobacterial counterpart (Jordan et al., 2001). The Lhcr subunits were detected as single bands of ∼20 kD by BN-PAGE (data not shown), suggesting that they exist as monomers in contrast to the higher plant Lhca dimers (Mazor et al., 2017). We then examined the putative changes in the accumulation of carotenoids in whole cells exposed to all four light regimes as well as purified intact PSI-LHCI supercomplexes, LHCI-depleted PSI core particles, and the outer peripheral LHCI antenna complexes. To this end, we performed quantitative HPLC analysis of the pigments present in all of the above samples to determine their molar ratios with respect to the total number of Chla molecules estimated in each fraction (Tables II and III). Three main carotenoids were identified in all the samples obtained from four different light regimes (Tables II and III; Supplemental Figs. S1 and S2): Zea, β-carotene (β-car), and β-cryptoxanthin (β-crypto [an intermediate of Zea biosynthesis from β-car in C. merolae]; Cunningham et al., 2007). Of all the carotenoids identified in our study, Zea accumulated in considerable amounts in both intact cells as well as the isolated PSI-LHCI, PSI core, and LHCI antenna complexes in response to an increasing light intensity (Tables II and III; Supplemental Figs. S1 and S2). Our results confirm the observations of Gantt and colleagues (Cunningham et al., 1989), who showed that, in the mesophilic unicellular red alga Porphyridium cruentum, the cellular content of Zea increased with growth irradiance, confirming a pivotal role for this carotenoid in long-term light adaptation. Nevertheless, the molar ratios of Zea/Chla seem to be up to 3.3-fold higher in C. merolae cells (Table II) compared with the cells of P. cruentum, indicating that this carotenoid plays an important role in acclimation of the extremophilic red microalgae exposed to low pH and high temperatures under variable light. Under such challenging conditions, the additional energetic pressure may be exerted on the C. merolae photosynthetic apparatus to cope with an increased demand on ATP to actively extrude protons from the cytoplasm. Therefore, the accumulation of photoprotective Zea may be one of the crucial strategies to maintain the functional and structural integrity of the photosynthetic apparatus and thylakoid membranes in this thermoacidophilic microalga. Interestingly, the β-car/Chl ratio showed only a minor increase, up to 20% for the EHL cells compared with LL cells, with the exception of HL cell samples, for which a rather low β-car/Chl ratio was observed (Table II). The significance of the latter observation is unknown at present, although it can be stipulated that the cells exposed to HL conditions decrease the overall content of β-car, perhaps due to the enhanced Zea synthesis from β-car that is triggered by light stress. Quantification of molar ratios of pigments in LL, ML, HL, and EHL C. merolae cells Table II. Quantification of molar ratios of pigments in LL, ML, HL, and EHL C. merolae cells Preparation . Pigment and Light Regime . . LL (35 µE) . ML (90 µE) . HL (150 µE) . EHL (350 µE) . Zea/Chla 0.856 ± 0.001 1.061 ± 0.002 1.169 ± 0.049 2.878 ± 0.040 β-Car/Chla 0.323 ± 0.002 0.362 ± 0.001 0.080 ± 0.002 0.397 ± 0.002 β-Crypto/Chla 0.061 ± 0.002 0.071 ± 0.001 0.056 ± 0.001 0.151 ± 0.003 Preparation . Pigment and Light Regime . . LL (35 µE) . ML (90 µE) . HL (150 µE) . EHL (350 µE) . Zea/Chla 0.856 ± 0.001 1.061 ± 0.002 1.169 ± 0.049 2.878 ± 0.040 β-Car/Chla 0.323 ± 0.002 0.362 ± 0.001 0.080 ± 0.002 0.397 ± 0.002 β-Crypto/Chla 0.061 ± 0.002 0.071 ± 0.001 0.056 ± 0.001 0.151 ± 0.003 Molar ratios of pigments were calculated by integration of an area underneath the relevant peak and using extinction coefficients, as described in “Materials and Methods.” sd values were calculated from one injection from two independent preparations (n = 2). Open in new tab Table II. Quantification of molar ratios of pigments in LL, ML, HL, and EHL C. merolae cells Preparation . Pigment and Light Regime . . LL (35 µE) . ML (90 µE) . HL (150 µE) . EHL (350 µE) . Zea/Chla 0.856 ± 0.001 1.061 ± 0.002 1.169 ± 0.049 2.878 ± 0.040 β-Car/Chla 0.323 ± 0.002 0.362 ± 0.001 0.080 ± 0.002 0.397 ± 0.002 β-Crypto/Chla 0.061 ± 0.002 0.071 ± 0.001 0.056 ± 0.001 0.151 ± 0.003 Preparation . Pigment and Light Regime . . LL (35 µE) . ML (90 µE) . HL (150 µE) . EHL (350 µE) . Zea/Chla 0.856 ± 0.001 1.061 ± 0.002 1.169 ± 0.049 2.878 ± 0.040 β-Car/Chla 0.323 ± 0.002 0.362 ± 0.001 0.080 ± 0.002 0.397 ± 0.002 β-Crypto/Chla 0.061 ± 0.002 0.071 ± 0.001 0.056 ± 0.001 0.151 ± 0.003 Molar ratios of pigments were calculated by integration of an area underneath the relevant peak and using extinction coefficients, as described in “Materials and Methods.” sd values were calculated from one injection from two independent preparations (n = 2). Open in new tab Quantification of molar ratios of pigments in LL, ML, HL, and EHL C. merolae PSI-LHCI supercomplexes, LHCI antennae, and LHCI-depleted PSI core particles Table III. Quantification of molar ratios of pigments in LL, ML, HL, and EHL C. merolae PSI-LHCI supercomplexes, LHCI antennae, and LHCI-depleted PSI core particles Molar ratios of pigments were calculated by integration of an area underneath the relevant peak and using extinction coefficients, as described in “Materials and Methods.” Values in parentheses show the number of pigment molecules normalized to 210, 182, 168, and 154 Chls for the LL, ML, HL, and EHL PSI-LHCI supercomplexes, respectively. The calculations were performed assuming 14 Chls per Lhcr; eight, six, five, and four Lhcrs per LL, ML, HL, and EHL supercomplexes, respectively (see Table IV); 98 Chls per PSI core complex; as well as 112, 84, 70, and 56 Chls (Mazor et al., 2017) per each respective LHCI complex. sd values were calculated from one to two injections from two independent preparations (n = 2). Open in new tab Table III. Quantification of molar ratios of pigments in LL, ML, HL, and EHL C. merolae PSI-LHCI supercomplexes, LHCI antennae, and LHCI-depleted PSI core particles Molar ratios of pigments were calculated by integration of an area underneath the relevant peak and using extinction coefficients, as described in “Materials and Methods.” Values in parentheses show the number of pigment molecules normalized to 210, 182, 168, and 154 Chls for the LL, ML, HL, and EHL PSI-LHCI supercomplexes, respectively. The calculations were performed assuming 14 Chls per Lhcr; eight, six, five, and four Lhcrs per LL, ML, HL, and EHL supercomplexes, respectively (see Table IV); 98 Chls per PSI core complex; as well as 112, 84, 70, and 56 Chls (Mazor et al., 2017) per each respective LHCI complex. sd values were calculated from one to two injections from two independent preparations (n = 2). Open in new tab The relative amounts of total carotenoids estimated in the intact PSI-LHCI supercomplex were 52, 39, 51, and 54 molecules (normalized to 14 Chls per Lhc subunit, as well as 8, 6, 5, and 4 Lhcrs per LL, ML, HL, and EHL PSI RC, and 98 Chls per PSI RC) for the LL, ML, HL, and EHL samples, respectively, indicating that the overall amount of total carotenoids remained similar in the PSI-LHCI supercomplexes pre-adapted to various light regimes (Table III). However, the relative amount of total carotenoids in the LHCI antenna complexes showed a steady increase during adaptation of the cells to excessive light. Thus, we identified 4, 6, 8, and 12 carotenoids per Lhcr subunit from LL, ML, HL, and EHL LHCI fractions, respectively (Table III). The total numbers of carotenoids in the C. merolae HL and EHL LHCI antenna complexes are significantly higher than for the higher plant counterpart, where an average of 3-4 carotenoids were identified per Lhca subunit by x-ray crystallography (Qin et al., 2015; Mazor et al., 2017). Similarly, an average of 8 carotenoids per Lhcr was reported in the LL/ML LHCI fraction from the same alga (Tian et al., 2017a). The quantification of the Zea/Chla molar ratios allowed us to estimate the relative numbers of Zea molecules as 28, 33, 36, and 45 per LL, ML, HL, and EHL PSI-LHCI supercomplex, respectively, following the same normalization as above. The analogous estimation gives a total number of β-car molecules of 19, 4, 13, and 6 per LL, ML, HL, and EHL PSI-LHCI supercomplex, respectively (Table III), indicating a 73% decrease in the content of this carotenoid in the EHL C. merolae PSI-LHCI sample compared with the x-ray structure of the pea (Pisum sativum) PSI-LHCI supercomplex, for which 22 β-car molecules were identified (Qin et al., 2015; Mazor et al., 2017). All the above data point to the conclusion that there may be significant structural differences between the red algal PSI-LHCI supercomplex and the higher plant counterpart with respect to the intrinsic pigment organization during adaptation to excessive light, whereby Zea accumulation is accompanied by a steady decrease in the β-car content. Significant differences in the pigment organisation and interactions were also observed for the C. merolae PSI-LHCI complex and its cyanobacterial counterpart, as evidenced by the circular dichroism (CD) spectroscopy (Supplemental Fig. S3). The comparison of pigment quantification data between various samples prompted us to the conclusion that the main locus for the accumulation of Zea in response to increasing light intensities is the LHCI antenna (Table III), in agreement with a previous study (Tian et al., 2017a). In the higher plant PSI-LHCI supercomplex, 2% of the total amount of Zea was detected in the core complex, in addition to the majority of this pigment being present in the LHCI antenna (Ballottari et al., 2014). Indeed, our LHCI-depleted PSI complexes obtained from various light regimes contained 1-5 Zea molecules (Table III), suggesting that this pigment also may evoke a light-harvesting role in the RC of PSI-LHCI supercomplex under light-limiting conditions. However, at this stage, it is impossible to determine unequivocally whether these Zea molecules were associated exclusively with the pure PSI core complex, as this preparation still contained the residual amount of tightly bound, if excitonically uncoupled, Lhcr antenna subunits (Fig. 5C). Our pigment quantification differs from the values obtained in the study by Tian et al. (2017a), probably due to different light illumination and cell culture growth conditions used for obtaining the PSI-LHCI samples. In the previous study, 18 molecules of Zea were reported for the LL/ML PSI-LHCI supercomplex of C. merolae, in contrast to 28-45 Zea molecules identified in this work. One important difference between our study and the above-mentioned results was that fact that we applied the specific light treatment to mid-log phase cultures to ensure the complete penetration of light, in contrast to the previous study, in which stationary cultures were used for all the biochemical and spectroscopic analyses. The direct photoprotective role of Zea was proposed for the higher plant PSI-LHCI supercomplex, where this carotenoid, bound within the LHCI domain, was implicated in a novel type of Zea-dependent nonphotochemical quenching, involving the formation of the carotenoid radical cations, that was coupled to an improved photostability of the PSI-LHCI supercomplex (Ballottari et al., 2014). It is important to emphasize that this phenomenon occurred in the npq2 mutant of Arabidopsis thaliana that constitutively accumulated Zea and lacked violaxanthin, whereas HL-adapted wild-type plants showed no Zea in the PSI core complex (Ballottari et al., 2014). The postulate of a direct role of Zea in photoprotection of the higher plant PSI-LHCI supercomplex was recently challenged due to the similar time-resolved fluorescence kinetics of the PSI-LHCI complex in dark-adapted and HL conditions (Tian et al., 2017b), suggesting that this carotenoid may play a different role in photoprotection of the photosynthetic apparatus than initially proposed (Ballottari et al., 2014). A feasible physiological photoprotective role for Zea may be to act as a direct antioxidant in scavenging free radicals and singlet oxygen molecules produced in HL in the lipid phase of the thylakoid membranes, in the vicinity of PSII and PSI-LHCI macrodomains (Havaux et al., 2007; Johnson et al., 2007). Indeed, we observed a large amount of Zea also associated with pure ML dimeric PSII samples from C. merolae (Krupnik et al., 2013), in addition to the presence of this carotenoid in purified PSI-LHCI supercomplex identified in this study. Free Zea molecules bound within the C. merolae PSI-LHCI and PSII complex-lipid interface also may protect the intrinsic lipid molecules, identified in the x-ray structures of both red algal PSII and higher plant PSI (Mazor et al., 2015, 2017; Qin et al., 2015; Ago et al., 2016), from singlet oxygen-mediated peroxidation (Havaux et al., 2007). An additional structural role for Zea may be to act as the rigidifying molecule (Gruszecki and Strzałka, 2005), which could lead to better stability of C. merolae photosynthetic complexes at high temperatures and in HL conditions. Antenna Remodeling of the C. merolae PSI-LHCI Supercomplex under Varying Light Conditions In Rhodophyta, the peripheral LHCI antenna system is formed by Chla-binding Lhcr proteins whose number per supercomplex is species dependent (Wolfe et al., 1994a, 1994b). Biochemical analyses showed the presence of six Lhcr proteins in the red alga P. cruentum (Tan et al., 1997) and five Lhcr polypeptides in G. sulphuraria (Marquardt et al., 2001). The nuclear genome of C. merolae encodes three Lhcr polypeptides (Matsuzaki et al., 2004), which are present at various stoichiometries in the PSI-LHCI complex to form the asymmetrically located LHCI antenna belt, as confirmed by proteomic and electron microscopy coupled to single-particle analyses (Busch et al., 2010). In this work, we tested the putative remodeling of the C. merolae LHCI antenna upon exposure to various light intensities. Such remodeling of the LHCI antenna was observed for LL-grown cells of two extremophilic red microalgae, G. sulphuraria and Cyanidium caldarium; however, the precise number of the Lhcr subunits could not be determined (Gardian et al., 2007; Thangaraj et al., 2011). To gain an insight into the putative rearrangement of the peripheral light-harvesting antenna subunits as the adaptation mechanism to varying light conditions, we visualized the negatively stained PSI-LHCI particles (purified from cells grown in LL, ML, and EHL conditions) by electron microscopy (Fig. 6) followed by single-particle analysis of their 2D projections. Single-particle averaging of a large set of particles showed that they were mostly present in a top-view position from the stromal side (Fig. 7). Single-particle analysis revealed two to four specific forms of PSI-LHCI supercomplexes whose presence and relative abundance depend on the growth light conditions. Under LL conditions, only two large and medium forms are present (Fig. 7, A and B), with dimensions of 19 × 17 nm and 19 × 15 nm, respectively. Under EHL conditions, the largest form of the PSI-LHCI supercomplex is completely absent and two smaller forms can be identified, in addition to the medium size particles obtained from LL and ML conditions (Fig. 7, H and I), with dimensions of 19 × 14 nm and 17 × 15 nm, respectively. In ML conditions, all forms of the PSI-LHCI supercomplex were present, indicating the largest structural heterogeneity of this complex preparation. Figure 6. Open in new tabDownload slide Sections of the electron micrographs of negatively stained PSI-LHCI supercomplex particles purified from C. merolae cells grown under three different light conditions (LL, ML, and EHL). Figure 6. Open in new tabDownload slide Sections of the electron micrographs of negatively stained PSI-LHCI supercomplex particles purified from C. merolae cells grown under three different light conditions (LL, ML, and EHL). Figure 7. Open in new tabDownload slide Structural characterization of PSI-LHCI supercomplexes from the red alga C. merolae grown under different light conditions. Single-particle image analysis revealed two forms of PSI-LHCI supercomplexes under LL conditions (A and B), four forms of PSI-LHCI supercomplexes under ML conditions (C–F), and three forms of PSI-LHCI supercomplexes under EHL conditions (G–I). Figure 7. Open in new tabDownload slide Structural characterization of PSI-LHCI supercomplexes from the red alga C. merolae grown under different light conditions. Single-particle image analysis revealed two forms of PSI-LHCI supercomplexes under LL conditions (A and B), four forms of PSI-LHCI supercomplexes under ML conditions (C–F), and three forms of PSI-LHCI supercomplexes under EHL conditions (G–I). To determine the location and estimate the number of the antenna subunits, we overlaid the x-ray structure of the higher plant PSI-LHCI supercomplex (Mazor et al., 2017) onto the C. merolae PSI-LHCI supercomplex projections. Figure 8 shows four structural models (A–D) of the C. merolae supercomplex purified from three light regimes (LL, ML, and EHL), which consider the varying number of Lhcr subunits, as well as the absence of some core subunits, as determined by mass spectrometry in this study (Table I) and in the other published data (Busch et al., 2010; Tian et al., 2017a). Figure 8. Open in new tabDownload slide Structural modeling of the PSI-LHCI supercomplex isolated from the red alga C. merolae grown under different light conditions. The largest form of the PSI-LHCI supercomplex consists of the PSI core complex and eight antenna proteins (A); smaller forms represent the association between the PSI core complex and either seven (B and D) or six (C) Lhcr antenna subunits. Magenta contours and black asterisks indicate strong densities and stain-accumulated areas, respectively, in the projection map of the largest PSI-LHCI supercomplex (A). Overlay of the contour model with the projection maps of smaller forms of the PSI-LHCI supercomplexes (B–D) indicates their structural differences compared with the largest form. Structural assignment of the PSI-LHCI supercomplexes was based on overlay of the x-ray structure of the plant PSI-LHCI supercomplex (Mazor et al., 2017). The PsaG and PsaH subunits were removed from the structure, as they are absent in C. merolae (Matsuzaki et al., 2004). PsaK is missing in the PSI core structure in the structural model of the smaller PSI-LHCI supercomplex (C). The structure of the Lhca4 protein was used to fit the second row of Lhcr antenna subunits. Figure 8. Open in new tabDownload slide Structural modeling of the PSI-LHCI supercomplex isolated from the red alga C. merolae grown under different light conditions. The largest form of the PSI-LHCI supercomplex consists of the PSI core complex and eight antenna proteins (A); smaller forms represent the association between the PSI core complex and either seven (B and D) or six (C) Lhcr antenna subunits. Magenta contours and black asterisks indicate strong densities and stain-accumulated areas, respectively, in the projection map of the largest PSI-LHCI supercomplex (A). Overlay of the contour model with the projection maps of smaller forms of the PSI-LHCI supercomplexes (B–D) indicates their structural differences compared with the largest form. Structural assignment of the PSI-LHCI supercomplexes was based on overlay of the x-ray structure of the plant PSI-LHCI supercomplex (Mazor et al., 2017). The PsaG and PsaH subunits were removed from the structure, as they are absent in C. merolae (Matsuzaki et al., 2004). PsaK is missing in the PSI core structure in the structural model of the smaller PSI-LHCI supercomplex (C). The structure of the Lhca4 protein was used to fit the second row of Lhcr antenna subunits. Structural model A corresponds to the largest form of the PSI-LHCI supercomplex, which consists of the PSI core complex and up to eight Lhcr antenna proteins. The inner belt of four antenna proteins is likely to be organized within the same architecture as the higher plant LHCI structure (Mazor et al., 2017). The second row of antenna proteins can contain up to four additional Lhcr subunits over and above the plant structure. In structural model B, a smaller form of the PSI-LHCI supercomplex lacks one Lhcr antenna protein in the second row of LHCI. Its absence can lead to different binding of the antenna proteins present in the inner belt of LHCI at the position where Lhca1 binds to the core complex in higher plants (Qin et al., 2015; Mazor et al., 2017). Structural model C corresponds to the smaller form of the PSI-LHCI supercomplex, which, in addition, lacks (compared with model B) the PsaK subunit (Table I) and another closely associated antenna protein. Thus, the PSI core complex associates with only six antenna proteins in this type of class average. Structural model D represents the smallest form of the C. merolae PSI-LHCI supercomplex. Although the PSI core complex can associate with up to seven antenna proteins in the same way as in model B, the protein density close to the PsaL subunit, which is essential for trimerization of the cyanobacterial PSI (Chitnis and Chitnis, 1993; Schluchter et al., 1996; Jordan et al., 2001), is absent in this projection. We compared our electron microscopy data by quantification of Lhcr subunits by measuring the P700/Chla ratios for all four PSI-LHCI complex preparations using chemical oxidation and reduction of the P700 RC and comparing the same measurements conducted on the isolated PSI complex of T. elongatus, which is known to bind 98 Chla molecules (Jordan et al., 2001). The results of this analysis are summarized in Table IV. In brief, we estimated the total number of Chla molecules per PSI-LHCI complex as 214, 186, 169, and 159. These values correspond to eight, six, five, and four Lhcr subunits per LL, ML, HL, and EHL PSI-LHCI supercomplex, respectively, taking into account an average of 14 Chl molecules per antenna subunit, as determined in the latest 2.6 Å x-ray structure of the higher plant PSI-LHCI supercomplex (Mazor et al., 2017). The total number of Lhcr subunits estimated for the smallest EHL complex using the redox spectroscopy approach was somewhat smaller than in our structural model C (four versus six Lhcrs, respectively), possibly due to some deactivation of the PSI RCs exposed to EHL conditions or the experimental error of both approaches. Quantification of Chla/P700 ratios in LL, ML, and HL C. merolae PSI-LHCI supercomplexes Table IV. Quantification of Chla/P700 ratios in LL, ML, and HL C. merolae PSI-LHCI supercomplexes PSI-LHCI Sample . Total No. of Chls . No. of Chls in the LHCI Antenna . Estimated No. of Lhcr Subunits . LL (35 µE m−2 s−1) 214.0 ± 2.34 116.0 ± 2.34 8 ML (90 µE m−2 s−1) 185.6 ± 4.56 87.6 ± 4.56 6 HL (150 µE m−2 s−1) 169.0 ± 5.29 71.0 ± 5.29 5 EHL (350 µE m−2 s−1) 159.0 ± 2.51 61.5 ± 2.51 4 PSI-LHCI Sample . Total No. of Chls . No. of Chls in the LHCI Antenna . Estimated No. of Lhcr Subunits . LL (35 µE m−2 s−1) 214.0 ± 2.34 116.0 ± 2.34 8 ML (90 µE m−2 s−1) 185.6 ± 4.56 87.6 ± 4.56 6 HL (150 µE m−2 s−1) 169.0 ± 5.29 71.0 ± 5.29 5 EHL (350 µE m−2 s−1) 159.0 ± 2.51 61.5 ± 2.51 4 Each value represents an average of at least three independent measurements from two to three independent samples. The number of Lhcr subunits was estimated taking into account an average of 14 Chl molecules per antenna subunit, as determined in the latest 2.6 Å x-ray structure of the higher plant PSI-LHCI supercomplex (Mazor et al., 2017). Open in new tab Table IV. Quantification of Chla/P700 ratios in LL, ML, and HL C. merolae PSI-LHCI supercomplexes PSI-LHCI Sample . Total No. of Chls . No. of Chls in the LHCI Antenna . Estimated No. of Lhcr Subunits . LL (35 µE m−2 s−1) 214.0 ± 2.34 116.0 ± 2.34 8 ML (90 µE m−2 s−1) 185.6 ± 4.56 87.6 ± 4.56 6 HL (150 µE m−2 s−1) 169.0 ± 5.29 71.0 ± 5.29 5 EHL (350 µE m−2 s−1) 159.0 ± 2.51 61.5 ± 2.51 4 PSI-LHCI Sample . Total No. of Chls . No. of Chls in the LHCI Antenna . Estimated No. of Lhcr Subunits . LL (35 µE m−2 s−1) 214.0 ± 2.34 116.0 ± 2.34 8 ML (90 µE m−2 s−1) 185.6 ± 4.56 87.6 ± 4.56 6 HL (150 µE m−2 s−1) 169.0 ± 5.29 71.0 ± 5.29 5 EHL (350 µE m−2 s−1) 159.0 ± 2.51 61.5 ± 2.51 4 Each value represents an average of at least three independent measurements from two to three independent samples. The number of Lhcr subunits was estimated taking into account an average of 14 Chl molecules per antenna subunit, as determined in the latest 2.6 Å x-ray structure of the higher plant PSI-LHCI supercomplex (Mazor et al., 2017). Open in new tab Overall, our combined P700/Chla quantification and modeling of the x-ray structure of the plant PSI-LHCI supercomplex onto 2D projections of the C. merolae PSI-LHCI supercomplex revealed that the smallest particle, which is predominant in EHL conditions, comprises the PSI core complex, with two rows of asymmetrically bound four to six Lhcr subunits located on the PsaF/PsaJ side of the core complex. On the other hand, the LL PSI-LHCI particle contained up to four additional Lhcr antenna subunits over and above the basic unit of the PSI-LHCI complex, displaying the additional loosely bound protein density extending the two rows of the proteins forming the belt-shaped LHCI complex. Our observation of such significant remodeling of the C. merolae PSI-LHCI complex is in stark contrast to the results of Busch et al. (2010) and Tian et al. (2017a), who reported only three to four Lhcr subunits forming the C. merolae LHCI antenna domain in LL and LL/ML conditions. Nevertheless, in agreement with our study, such significant structural readjustment of the LHCI antenna size has been observed for other microalgal species exposed to LL conditions, including the red algae G. sulphuraria (Thangaraj et al., 2011) and C. caldarium (Gardian et al., 2007) as well as the green alga Chlamydomonas reinhardtii (Drop et al., 2011). Therefore, we propose that significant structural remodeling of the LHCI antenna provides the molecular basis of photoadaptation in the extremophilic red alga C. merolae, whereby, upon exposure to EHL, the LHCI antenna has the smallest size to prevent overexcitation of the PSI RC. On the other hand, LL illumination induces a 2-fold increase of the effective antenna size (compared with the smallest PSI-LHCI complex predominant in EHL and HL conditions) to maximize the solar energy capture for efficient photochemistry to occur in the PSI RC. CONCLUSIONS In this work, we provided several lines of evidence that the ultra-robust C. merolae PSI-LHCI supercomplex evokes three distinct molecular mechanisms underlying its inherent robustness during adaptation to varying light conditions: (1) the accumulation of a photoprotective carotenoid Zea mainly in the LHCI antenna and, possibly, the PSI RC; (2) structural remodeling of the LHCI antenna and adjustment of the effective absorption cross section; and (3) dynamic readjustment of the stoichiometry of the two PSI-LHCI isomers identified in the C. merolae photosynthetic apparatus accompanied by the dissociation of the PsaK core subunit in the larger isomer or both isomers upon exposure to EHL and LL conditions, respectively. Our combined redox difference spectroscopy and single-particle analysis suggest that the largest C. merolae PSI-LHCI supercomplex can bind up to eight Lhcr antenna subunits, which are organized as two rows on the PsaF/PsaJ side of the core complex. In contrast to previous work (Busch et al., 2010), we found no evidence of functional coupling of the PBSs with the purified C. merolae PSI-LHCI supercomplex in all four light regimes studied, suggesting that the putative association of PBSs with PSI is absent (Yokono et al., 2011; Ueno et al., 2017) or it is transient and may be lost during the purification procedure. Future work will address this issue. MATERIALS AND METHODS Cell Culturing and Isolation of Thylakoids Cells of Cyanidioschyzon merolae strain NIES-1332 (obtained from the Microbial Culture Collection of the National Institute for Environmental Studies in Japan) were cultivated in a modified Allen medium at 42°C, pH 2.5 (Minoda et al., 2004), with continuous white light of 90 µE m−2 s−1 (ML) and bubbling with 5% CO2 in air, as described in detail by Krupnik et al. (2013). Cultures were inoculated to the start OD680 of 0.05-0.07 and grown under light intensities of 35, 90, 150, and 350 µE m−2 s−1 (Panasonic FL40SS-ENW/37 lamps) for 6, 5, 4, and 3 d, respectively, to reach the target OD680 of 0.5-0.7 prior to cell harvesting and thylakoid isolation. The procedure of thylakoid isolation was performed as described by Krupnik et al. (2013). Final thylakoid pellets were resuspended in buffer A (40 mm MES-KOH, pH 6.1, 10 mm CaCl2, 5 mm MgCl2, and 25% [w/v] glycerol) at a Chla concentration of 2 to 5 mg mL−1, snap frozen in liquid N2, and stored at −80°C prior to use. Cells of Thermosynechococcus elongatus BP-1 NIES-2133 wild-type strain (a gift from M. Nowaczyk, University of Bochum) were grown in BG-11 medium at 45°C, pH 8 (Castenholz, 1988), in continuous white light of 90 µmol photons m−2 s−1 (ML) with gentle bubbling with 5% CO2 in air. Cultures were grown for approximately 7 to 13 d to OD680 of 0.8 to 1 with continuous white light illumination. The procedure of thylakoid isolation was performed at 4°C in dim green light as described by Kuhl et al. (2000) with several modifications. Briefly, cells were harvested by centrifuging at 4,000g for 10 min, then resuspended in a buffer containing 20 mm MES-NaOH, pH 6.5, 10 mm MgCl2, and 10 mm CaCl2 supplemented with a protease inhibitor cocktail (Roche), DNase I (5 mg per 50 mL; Roche), and RNase I (10 μL from stock per 50 mL; Sigma-Aldrich). The cells were centrifuged as above and resuspended in the same buffer supplemented with 500 mm d-mannitol and the other supplements as above. Cells were disrupted by vigorous agitation with 0.1-mm glass beads (as described by Krupnik et al. [2013]) for 13 cycles, each of 10 s of beating and 4 min of rest. The cell homogenate was recovered from the beads by filtering through Whatman paper and washing the beads with the buffer as above supplemented with 500 mm d-mannitol. The homogenate was centrifuged for 1 min at 1,000g to remove the unbroken cells, then ultracentrifuged at 180,000g for 25 min to harvest the thylakoid membranes. The thylakoid pellet was resuspended in the same buffer devoid of d-mannitol. The operation was repeated two to three times depending on the amount of PBSs present. The final thylakoid pellet was resuspended in the buffer as above supplemented with 500 mm d-mannitol, adjusted to a Chla concentration of 2 to 5 mg mL−1, snap frozen in liquid N2, and stored at −80°C prior to use. Purification of C. merolae PSI-LHCI and T. elongatus PSI Complexes Solubilization of C. merolae thylakoids and separation of the PSII from crude PSI-LHCI samples was performed according to the protocol described in Krupnik et al. (2013). The crude C. merolae PSI-LHCI fraction, eluted from the DEAE TOYOPEARL 650M column with 0.09 m NaCl, as described by Krupnik et al. (2013), was applied onto the DEAE TOYOPEARL 650S column, and pure PSI-LHCI supercomplex was eluted with a continuous 0 to 0.2 m NaCl gradient in the carrier buffer. The PSI-LHCI pool obtained after the DEAE 650S column was concentrated to 1 mg mL−1 Chla and further purified to remove any residual PBSs by additional chromatography purification steps performed, first, on the desalting Superdex G-25 column in buffer B (40 mm HEPES-NaOH, pH 8, 3 mm CaCl2, 25% [w/v] glycerol, and 0.03% [w/v] dodecyl-β-D-maltoside, DDM), followed by an anion-exchange chromatography (AEC) step using a UNOQ12 column. The ultrapure PSI-LHCI supercomplex fractions were collected with a 0.05 m NaCl gradient that was separated from the PBSs, which displayed a very strong affinity to the UNOQ12 resin. The fractions containing the pure PSI-LHCI supercomplex were collected and concentrated to 2 to 5 mg mL−1 Chla, snap frozen in liquid N2, and stored at −80°C prior to use. For the purification of T. elongatus PSI trimer, thylakoids were thawed 5 h on ice. Normally, a total of 66.3 mg of Chla was used for solubilization with a 10% [w/v] stock of DDM (Roth). The Chla concentration was adjusted to 1.61 mg mL−1 (1.8 mm) before solubilization with 0.5% [w/v] DDM (10 mm) at a detergent-to-Chla molar ratio of 5:1, in the presence of the protease inhibitor cocktail (Roche), for 20 min at room temperature (RT) in the dark. The solubilized thylakoids were ultracentrifuged at 80,000g for 30 min (T-865 rotor; Thermo Scientific), and the supernatant was collected for the subsequent purification steps. Trimeric PSI complex of T. elongatus was purified from solubilized thylakoids by eluting from the DEAE TOYOPEARL 650M column with 0.09 m NaCl, as described previously (Krupnik et al., 2013). The crude PSI trimer was then applied to a DEAE TOYOPEARL 650S column that was subsequently washed extensively with the carrier buffer to remove excess PBSs and carotenoids. PSI trimer was eluted with a linear gradient of 0 to 0.5 m NaCl, then buffer exchanged on the desalting Sephadex G-25 column into buffer B as above, concentrated to 2 to 5 mg mL−1 Chla, snap frozen in liquid N2, and stored at −80°C prior to use. The monomeric PSI complex of T. elongatus were purified according to El-Mohsnawy et al. (2010). Briefly, thylakoid membranes were resuspended with 0.8 m ammonium sulfate and stirred at 50°C for 20 min in the dark. The suspension was cooled down to RT before the isolation of monomeric PSI complex by solubilization of the membranes with 0.6% to 1% [w/v] DDM. After ultracentrifugation at 80,000g for 1 h at 4°C, the filtrate was subjected to two purification steps hydrophobic interaction chromatography (HIC) and AEC to purify the monomeric PSI complex. The Chla concentration was measured spectroscopically according to Porra et al. (1989) using an extinction coefficient of 86.3 μg μL−1 cm−1. Purity of the samples was confirmed spectroscopically, by size exclusion chromatography, and by SDS-PAGE, as described by Krupnik et al. (2013). Isolation and Biochemical Characterization of C. merolae LHCI Antenna and LHCI-Depleted PSI RC Particles The LHCI antenna and PSI core particles were isolated by solubilization of the PSI-LHCI supercomplex (0.3 mg Chla mL−1) with 1.5% [w/v] DDM and 0.6% [w/v] Zwittergent 3-16 and with five freeze/thaw cycles followed by Suc gradient centrifugation of the detergent-treated complexes, as described by Melkozernov et al. (2004). Fractions were collected following 17 h of ultracentrifugation at 140,000g (Surespin 630 rotor; Thermo Scientific), concentrated to 1 to 3 mg mL−1 Chla, and characterized by SDS-PAGE, RT absorption, and 77K fluorescence spectroscopy, as described below. HPLC Pigment Analysis Analytical HPLC of pigments was performed according to the method described by Krupnik et al. (2013) using a Nucleosil-100 C18 column (Teknokoma) and a linear gradient of 10% to 60% [v/v] ethyl acetate. The content of each pigment was expressed as a ratio of the area under the pigment-corresponding peak to the area under the Chla peak. For the pigment molar ratio calculations, extinction coefficients of 83.2, 91.7, and 125.3 mm −1 cm−1 for Zea, Chla, and β-car were used, respectively (Oren et al., 1996). Activity of PSI Photochemical activity of the purified PSI-LHCI (2.5 μg of Chla) was measured by the oxygen consumption assay (Vernon and Cardon, 1982; Allakhverdiev et al., 2000) using an oxygen Clark-type electrode (Hansatech). Standard measurements were performed at 30°C in the reaction buffer (40 mm HEPES-NaOH, pH 8, 3 mm CaCl2, 25% [w/v] glycerol, and 0.05% [w/v] DDM) in the presence of 0.2 mm methyl viologen as an exogenous electron acceptor with 10 mm sodium azide as an efficient physical quencher of singlet oxygen and 0.2 mm dichlorophenolindophenol as a mediator. For a standard measurement, the samples were incubated in the dark for 2 min, followed by the addition of 6 mm sodium ascorbate as the sacrificial electron donor and illumination with a white light intensity of 5,000 µE m−2 s−1, using a KL 2500 LCD white light source (Schott). For measurements of the activity of PSI in the wide pH range, pH was regulated by adding HCl or NaOH into the buffer containing 40 mm HEPES-NaOH, 3 mm CaCl2, 25% [w/v] glycerol, and 0.05% [w/v] DDM. Samples were preincubated at various pH levels in the dark for 30 min with dilution of at least 50 times. The oxygen consumption was then immediately measured in the reaction buffer: 40 mm HEPES-NaOH, pH 8, 3 mm CaCl2, 25% [w/v] glycerol, and 0.05% [w/v] DDM. Activity was assayed with at least three independent measurements, and the values were expressed as means ± sd. RT Absorption and Circular Dichroism Spectroscopy Optical absorption spectra were recorded at 5 µg Chla mL−1 at RT in the range 800 to 350 nm using a UV-1800 Shimadzu spectrophotometer with the TCC-100 temperature-controlled cell holder, using a quartz cuvette with an optical path length of 10 mm. Determination of Chla/P700 ratios was done by differential absorption spectroscopy, as described (Kargul et al., 2003). The samples were diluted to 20 µg mL−1 in 50 mm MES-KOH, pH 6.5, 10 mm MgCl2, 10 mm CaCl2, and 0.05% [w/v] DDM. Chla/P700 ratios were quantified by measuring absorbance changes at 700 nm in the presence of 2 mm sodium ascorbate, 0.2 mm dichlorophenolindophenol as a reductant, and 0.5 mm ferricyanide as an oxidant. Circular dichroism (CD) absorption spectra of PSI complexes were measured according to Schlodder et al. (2007). PSI complexes were diluted to 10 µm Chla in 20 mm MES-KOH, pH 6.5, 10 mm CaCl2, 30 mm MgCl2, and 0.02% [w/v] DDM. A JASCO J-715 spectropolarimeter was adjusted to a speed of 20 nm min−1, bandwidth of 2 nm, and pitch of 0.2 nm. After five cycles, the spectra were averaged using JASCO software. 77K Fluorescence Spectroscopy Steady-state fluorescence emission spectra at 77K were acquired in an LS55 Fluorescence Spectrometer (Perkin Elmer), as described (Busch et al., 2010; Krupnik et al., 2013). For spectroscopic measurements, PSI samples (3 µg of Chla) were diluted in buffer B, then diluted 2-fold with 80% [w/v] glycerol and frozen in liquid N2. Fluorescence emission spectra were recorded in the range 600 to 800 nm using excitation wavelengths of 435 and 600 nm for Chla and phycocyanin, respectively. The emission spectra were normalized to the Chl peak of PSI at 728 nm. Action spectra were generated by exciting the samples in the range 400 to 700 nm and recording emission at 728 nm. The spectra obtained were normalized to the Chl peak at 674.5 nm. All spectra were obtained from two to three independent preparations in three replicates. BN-PAGE BN gel electrophoresis experiments were performed with 3% to 12% nondenaturing continuous gradient polyacrylamide gels according to Schägger and von Jagow (1991). Samples of PSI at 0.5 mg Chl mL−1 were mixed with 0.25 volumes of Coomassie Blue solution (5% [w/v] Serva Blue G, 750 mm aminocaproic acid, and 35% [w/v] Suc), and after incubation, they were loaded onto the gel. Commercially available markers from Invitrogen were used for protein mass identification. Electrophoresis was conducted in the running buffer at 90 V for 15 h at 4°C in the dark according to Farci et al. (2017). Densitometry analysis of the intensity of the bands on the BN gel was performed using Image Lab software (Bio-Rad Molecular Imager GelDoc XR). Sample Preparation for Mass Spectrometry The bands from the BN gels were excised, reduced in DTT (10 mm, 56°C, and 30 min), and subsequently alkylated with iodoacetamide (55 mm, 25°C, and 20 min in the dark). Following dehydration with acetonitrile, trypsin (1 ng μL−1 solution in 50 mm ammonium bicarbonate) was added, and the gel pieces were allowed to swell on ice for 30 min. They were then digested overnight at 37°C with shaking. After digestion, the peptide content was extracted twice with sonication (using a solution of 50:50 water:acetonitrile and 1% [v/v] formic acid). The pooled extracts were placed in a clean tube and dried with a speed vacuum centrifuge. The dried pool was finally redissolved in 50 μL of Oasis Solvent A (water and 0.05% [v/v] formic acid). The digests were then desalted with the Waters Oasis HLB µElution Plate 30 µm in the presence of a slow vacuum. In this process, the columns were conditioned with 3 × 100 µL of Oasis Solvent B (80% acetonitrile [v/v] and 0.05% [v/v] formic acid) and equilibrated with 3 × 100 µL of Oasis Solvent A. The samples were loaded, washed three times with 100 µL of Oasis Solvent A, and then eluted into PCR tubes with 50 µL of Oasis Solvent B. The eluates were dried down with the speed vacuum centrifuge and dissolved in 20 µL of 5% [v/v] acetonitrile, 95% MilliQ water, and 0.1% [v/v] formic acid prior to analysis by liquid chromatography-tandem mass spectrometry (LC-MS/MS). Liquid Chromatography-MS/MS Analysis Peptides were separated using the nanoAcquity ultra-performance liquid chromatography system (Waters) fitted with a trapping column (nanoAcquity Symmetry C18, 5 µm, 180 µm × 20 mm) and an analytical column (nanoAcquity BEH C18, 1.7 µm, 75 µm × 250 mm). The outlet of the analytical column was coupled directly to an Orbitrap Fusion Lumos (Thermo Fisher Scientific) using the Proxeon nanospray source. Solvent A was water and 0.1% [v/v] formic acid and solvent B was acetonitrile and 0.1% [v/v] formic acid. The samples (5 µL) were loaded with a constant flow of solvent A (5 µL min−1) onto the trapping column. Trapping time was 6 min. Peptides were eluted via the analytical column with constant flow (0.3 µL min−1). During the elution step, the percentage of solvent B increased in a linear fashion from 3% to 25% in 30 min, then increased to 32% in 5 min, and finally to 50% in a further 0.1 min. Total run time was 60 min. The peptides were introduced into the mass spectrometer via a Pico-Tip Emitter 360-µm o.d. × 20-µm i.d., 10-µm tip (New Objective), and a spray voltage of 2.2 kV was applied. The capillary temperature was set at 300°C. The ion funnel radio frequency (RF) lens was set to 30%. Full-scan mass spectra with mass range 375 to 1,500 mass-to-charge ratio (m/z) were acquired in profile mode in the Orbitrap with resolution of 120,000. The filling time was set at a maximum of 50 ms with limitation of 2 × 105 ions. The top speed method was employed to take the maximum number of precursor ions (with an intensity threshold of 5 × 103) from the full-scan mass spectra for fragmentation (using higher-energy collisional dissociation or HCD of 30%) and quadrupole isolation (1.4-D window) and measurement in the ion trap, with a cycle time of 3 s. The monoisotopic precursor selection peptide algorithm was employed, but with relaxed restrictions when too few precursors meeting the criteria were found. The fragmentation was performed after the accumulation of 2 × 103 ions or after filling time of 300 ms for each precursor ion (whichever occurred first). MS/MS data were acquired in centroid mode, with the rapid scan rate and a fixed first mass of 120 m/z. Only multiply charged (2+ to 7+) precursor ions were selected for MS/MS. Dynamic exclusion was employed with a maximum retention period of 60 s and relative mass window of 10 ppm. Isotopes were excluded. Additionally, only one data-dependent scan was performed per precursor (only the most intense charge state was selected). Ions were injected for all available parallelizable time. In order to improve the mass accuracy, a lock mass correction using a background ion (m/z 445.12003) was applied. For data acquisition and processing of the raw data, Xcalibur 4.0 (Thermo Scientific) was employed. Proteomic Data Analysis Raw data from the mass spectra were searched using MaxQuant (version 1.5.3.30; Cox and Mann, 2008). Data were searched against a species-specific (C. merolae; http://merolae.biol.s.u-tokyo.ac.jp/download/cds.fasta) database, with a list of common contaminants appended using the Andromeda search engine (Cox et al., 2011). The search criteria were set as follows: full tryptic specificity was required (cleavage after Lys or Arg residues, unless followed by Pro); two missed cleavages were allowed; oxidation (M) and acetylation (protein N-term) were applied as variable modifications, carbamidomethyl Cys was applied as a fixed modification, and a mass tolerance of 20 ppm (precursor) and 0.5 D (fragments) was set. The reversed sequences of the target database were used as a decoy database. Peptide and protein hits were filtered at a false discovery rate of 1% using a target-decoy strategy (Elias and Gygi, 2007). Data from entries that are relevant to the photosynthetic apparatus are represented in Table I. They are depicted according to their iBAQ intensities (extracted from the MaxQuant protein groups output) for each of the proteins corresponding to each of the BN complexes from the BN-PAGE gels (Fig. 3). Here, the relative contributions are represented by icons, where four crosses represent proteins with iBAQ values ≥ 1.75e10, three crosses ≥ 2.5e9, two crosses ≥ 5e8, one cross ≥ 2.5e6, and a minus sign < 2.5e6. For a full list of proteins identified in each band (including those not considered to be part of the photosynthetic apparatus), with absolute iBAQ values shown, see Supplemental Table S1. Data were filtered according to iBAQ value; any protein with an iBAQ value less than 1e6 in any band was filtered from the list. One peptide per protein hits were retained where the iBAQ value was above this threshold in at least one condition, as a number of the proteins with this peptide number belong to the lower molecular weight proteins in the list, where not so many peptides can be expected upon digestion. Electron Microscopy and Image Processing Specimens for electron microscopy were prepared on glow-discharged carbon-coated copper grids and negatively stained with 2% [w/v] uranyl acetate. Electron microscopy was performed on a Tecnai TF20 microscope (FEI) equipped with a field emission gun operated at 200 kV. Images were recorded with an Eagle 4K CCD camera (FEI) at 83,000× magnification with a pixel size of 0.18 nm. Automated data acquisition software for single-particle analysis (EPU; FEI) was used for the acquisition of about 800, 1,100, and 700 micrographs of LL, ML, and HL samples, respectively. Data sets of ?60,000, 150,000, and 90,000 single-particle projections of PSI-LHCI supercomplexes were selected for LL, ML, and HL samples, respectively. Single-particle image analysis (Boekema et al., 2009) was performed using RELION software (Scheres, 2012). Image analysis revealed that about 75% to 80% of the projections from each data set could be assigned to one of the specific classes of the PSI-LHCI supercomplexes. Pseudo-atomic models of the PSI-LHCI supercomplexes were created using PYMOL (DeLano, 2002). Accession Numbers Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers. CMV135C, CMV136C, CMV059C, CMV144C, CMV128C, CMV201C, CMV202C, CMV055C, CMV236C, CMP086C, CMN234C, CMN235C, CMQ142C, CMV063C, CMP166C, CMV158C, CMV064C, CMV159C, and CMV051C. Supplemental Data The following supplemental materials are available. Supplemental Figure S1. HPLC pigment analysis of the C. merolae PSI-LHCI supercomplex. Supplemental Figure S2. HPLC pigment analysis of the C. merolae cells. Supplemental Figure S3. CD-absorption spectra of PSI monomers and trimers of T. elongatus and the ML PSI-LHCI supercomplex of C. merolae. Supplemental Table S1. 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Nature 409 : 739 – 743 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 J.M.K., P.H., and M.A. acknowledge support from the Polish National Science Centre (OPUS grant no. UMO-2014/15/B/NZ1/00975 to J.M.K.). Part of this work was also supported by the Polish Ministry of Science and Higher Education and the European Science Foundation within the Eurocores/EuroSolarFuels/Solarfueltandem programme (grant no. 844/N-ESF EuroSolarFuels/10/2011/0 to J.M.K.). The work of L.N. and R.K. was supported by grant LO1204 (Sustainable Development of Research in the Centre of the Region Haná) from the National Program of Sustainability I from the Ministry of Education, Youth, and Sports, Czech Republic. CIISB research infrastructure project LM2015043 funded by MEYS CR is gratefully acknowledged for the financial support of the measurements at the CF Cryo-Electron Microscopy and Tomography. E.E.-M. acknowledges support from the Ministry of Higher Education Egypt. 2 Address correspondence to j.kargul@uw.edu.pl. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Joanna M. Kargul (j.kargul@uw.edu.pl). P.H., M.A., L.N., J.K., J.M.K., E.E.-M., and R.K. generated and processed the data and prepared the figures; J.D.J.O. contributed to purification and biochemical characterization of PSI and studies on its interaction with PBSs; J.K., J.M.K., P.H., and R.K. designed the experiments and analyzed and interpreted the data; J.M.K., R.K., and P.H. cowrote the article; J.M.K. conceived and coordinated the study. [OPEN] Articles can be viewed without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.17.01022 © 2018 American Society of Plant Biologists. All Rights Reserved. © The Author(s) 2018. Published by Oxford University Press on behalf of American Society of Plant Biologists. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.
TI - Molecular Mechanisms of Photoadaptation of Photosystem I Supercomplex from an Evolutionary Cyanobacterial/Algal Intermediate  
JF - Plant Physiology
DO - 10.1104/pp.17.01022
DA - 2018-02-06
UR - https://www.deepdyve.com/lp/oxford-university-press/molecular-mechanisms-of-photoadaptation-of-photosystem-i-supercomplex-LCcvfOpdpY
SP - 1433
EP - 1451
VL - 176
IS - 2
DP - DeepDyve
ER -