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. 2024 Mar 5;65(5):790–797. doi: 10.1093/pcp/pcae021

High Myristic Acid in Glycerolipids Enhances the Repair of Photodamaged Photosystem II under Strong Light

Kazuki Kurima 1, Haruhiko Jimbo 2,*, Takashi Fujihara 3, Masakazu Saito 4,5, Toshiki Ishikawa 6, Hajime Wada 7
PMCID: PMC11138363  PMID: 38441322

Abstract

Cyanobacteria inhabit areas with a broad range of light, temperature and nutrient conditions. The robustness of cyanobacterial cells, which can survive under different conditions, may depend on the resilience of photosynthetic activity. Cyanothece sp. PCC 8801 (Cyanothece), a freshwater cyanobacterium isolated from a Taiwanese rice field, had a higher repair activity of photodamaged photosystem II (PSII) under intense light than Synechocystis sp. PCC 6803 (Synechocystis), another freshwater cyanobacterium. Cyanothece contains myristic acid (14:0) as the major fatty acid at the sn-2 position of the glycerolipids. To investigate the role of 14:0 in the repair of photodamaged PSII, we used a Synechocystis transformant expressing a T-1274 encoding a lysophosphatidic acid acyltransferase (LPAAT) from Cyanothece. The wild-type and transformant cells contained 0.2 and 20.1 mol% of 14:0 in glycerolipids, respectively. The higher content of 14:0 in the transformants increased the fluidity of the thylakoid membrane. In the transformants, PSII repair was accelerated due to an enhancement in the de novo synthesis of D1 protein, and the production of singlet oxygen (1O2), which inhibited protein synthesis, was suppressed. The high content of 14:0 increased transfer of light energy received by phycobilisomes to PSI and CP47 in PSII and the content of carotenoids. These results indicated that an increase in 14:0 reduced 1O2 formation and enhanced PSII repair. The higher content of 14:0 in the glycerolipids may be required as a survival strategy for Cyanothece inhabiting a rice field under direct sunlight.

Keywords: Cyanothece sp. PCC 8801, Membrane fluidity, Myristic acid, Photoinhibition, Synechocystis sp. PCC 6803

Introduction

Excess light damages photosynthesis and often becomes lethal for photosynthetic organisms. Cyanobacteria, oxygenic photosynthetic bacteria, have a broad habitat and some species can survive even under severe conditions such as hot springs, deserts, and polar regions. Cyanothece sp. PCC 8801 (hereafter Cyanothece), a nitrogen-fixing cyanobacterium, was found in a Taiwanese rice field (Huang and Chow 1988), where the natural light environment is harmful to photosynthetic microorganisms. The intensity of the sunlight on the surface of the field reached ∼2000 photons m−2 s−1. Under such intense light, photosystem II (PSII), an oxygen-evolving photosynthetic complex, is photodamaged, and photosynthesis is inhibited. Photodamaged PSII is repaired by proteolytic-based turnover and PSII activity is recovered. When the rate of photodamage exceeds that of repair, PSII activity decreases, referred to as photoinhibition. The action of reactive oxygen species (ROS), which are produced from the over-reduced photosynthetic electron transport under strong light (Asada 1999), is critical for the activity of PSII repair (Nishiyama et al. 2001). ROS attack the translation machinery required for protein synthesis and inhibit PSII repair (Jimbo et al. 2018, Murata and Nishiyama 2018). Therefore, the cyanobacterial species that inhabit niches exposed to strong light intensity, such as Cyanothece, must overcome ROS-related photoinhibition.

Cyanothece cells contain a high amount of myristic acid (14:0) at the sn-2 position of membrane glycerolipids (Saito et al. 2018). In other freshwater cyanobacteria, such as Synechocystis sp. PCC 6803 (hereafter Synechocystis), Synechococcus elongatus PCC 7942 and Anabaena sp. PCC 7120, palmitic acid (16:0) is a major saturated fatty acid at the sn-2 position of membrane glycerolipids. The occurrence of shorter chain lengths and more unsaturated fatty acids in membrane lipids increases the fluidity of biological membranes, which confers resistance to low-temperature-induced stress in Synechocystis cells (Wada et al. 1990, 1994, Murata et al. 1992). Therefore, 14:0, a shorter fatty acid present in Cyanothece than 16:0 in Synechocystis, might enhance the resilience of photosynthesis to environmental stress. However, the transformation techniques for Cyanothece have not been established, and the physiological role of 14:0 in Cyanothece has not been clarified.

In the present study, the sll1848 gene in Synechocystis encoding a major lysophosphatidic acid acyltransferase (LPAAT) was knocked out, as reported previously (Okazaki et al. 2006). Next, T-1274 of Cyanothece encoding an LPAAT responsible for the transfer of 14:0 to lysophosphatidic acids (LPAs) was expressed in the sll1848 knock-out mutant Synechocystis cells to elevate the contents of 14:0. In the transformants obtained, the 14:0 contents in membrane lipids increased, and the repair of the photodamaged PSII was accelerated because of the enhancement of the de novo synthesis of the D1 protein. The high content of 14:0 in the transformants also enhanced the fluidity of the thylakoid membrane, energy transfer from phycobilisomes to PSI and CP43 in PSII and the content of carotenoids, and decreased the production of singlet oxygen (1O2), which inhibited the repair of photodamaged PSII. These results suggest that 14:0 has an important role for the survival of Cyanothece under intense light.

Results

Higher PSII repair activity in Cyanothece

Synechocystis, a cyanobacterial strain used globally for photosynthesis research, was initially found in a shaded pond in Oakland, California (Stanier et al. 1971). Other ecotypes are also found in shaded fields. Cyanothece, located close to the Synechocystis branch in phylogenic analysis (Bandyopadhyay et al. 2011), was initially found in a Taiwanese rice field (Huang and Chow 1988). The sunlight intensity in rice fields reaches over 2000 μmol photons m−2 s−1. Therefore, the light sensitivity of PSII was examined by measuring the O2 evolution of cells exposed to strong light at 1500 μmol photons m−2 s−1. The maximum activities of PSII in Synechocystis and Cyanothece cells before exposure to strong light were similar (Supplementary Fig. S1). After 80 min of exposure, the PSII activity of Synechocystis cells declined by 60% (Fig. 1A). Under the same conditions, Cyanothece cells retained more than 90% of their original activity (Fig. 1A). In the presence of 200 μg ml−1 lincomycin that inhibits PSII repair, the PSII activity of Synechocystis and Cyanothece reduced at the same rate under strong light (Fig. 1B). Therefore, the PSII repair activity was higher in Cyanothece than that in Synechocystis.

Fig. 1.

Fig. 1

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Photoinhibition of PSII in Synechocystis sp. PCC 6803 (Synechocystis) and Cyanothece sp. PCC 8801 (Cyanothece) cells. WT cells of Synechocystis (circles) and Cyanothece (triangles) were incubated at 32°C under intense light at 1500 μmol photons m−2 s−1 with aeration by ambient air in the absence (A) or presence (B) of 200 μg ml−1 lincomycin. The PSII activity of Synechocystis and Cyanothece cells was measured in terms of O2 evolution in the presence of 1 mM 1,4-benzoquinone and 1 mM K3Fe(CN)6. The mean activities ± SD considered as 100% for Synechocystis and Cyanothece cells were 276.4 ± 15.7 and 254.6 ± 19.3 μmol O2 mg−1 Chl h−1, respectively. The values are the mean of the results from triplicates ± SD. *P < 0.01, calculated using ANOVA test.

Expression of the Cyanothece LPAAT encoding gene, T-1274, in a Δsll1848 mutant of Synechocystis increased the 14:0 content of glycerolipids and fluidity of the thylakoid membrane

In a previous study, we found that Cyanothece have a high level of myristic acid in their glycerolipids (Saito et al. 2018). Synechocystis or other freshwater cyanobacteria such as Synechococcus elongatus sp. PCC 7942 contain palmitic acid in their glycerolipids (Wada and Murata 1998). Thus, it was assumed that 14:0, a fatty acid with a shorter chain length than 16:0, may enhance the repair of PSII in Cyanothece. In a previous study, we identified T1274 in the Cyanothece genome as a gene encoding a LPAAT, which preferably transfers 14:0 to the sn-2 position of LPA (Saito et al. 2018) (Fig. 2A). Synechocystis has three LPAATs (Weier et al. 2005), with Sll1848 being the major LPAAT (Okazaki et al. 2006) (Fig. 2B). A Synechocystis sll1848 mutant was obtained by inserting a kanamycin-resistant gene cassette (KnR), as reported previously (Okazaki et al. 2006). T1274 of Cyanothece was expressed in the Synechocystis mutants to increase the 14:0 content of glycerolipids (Fig. 2A), and the strain obtained expressing was named T1274Δ1848. A previous study revealed that the expression of T1274 in the wild-type (WT) cells of Synechocystis enhanced the content of 14:0 in glycerolipids by up to 15.7 ± 1.5 mol% of total fatty acids (FAs) (Saito et al. 2018). In T1274Δ1848 cells, the content of 14:0 further increased and reached 20.1 ± 3.9 mol% of total FAs (Fig. 2C). However, 16:0 in the glycerolipids was not completely substituted with 14:0, probably due to the presence of other minor LPAATs in slr2060 and sll1752 or an insufficient level of 14:0 acyl-carrier protein (ACP), a substrate for LPAAT, in Synechocystis cells.

Fig. 2.

Fig. 2

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Construction of a transformant of Synechocystis, T1274Δ1848, expressing T1274 encoding a LPAAT from Cyanothece and its fatty acid content. (A) Schematic representation of the introduction of T1274 from Cyanothece into the neutral site between slr2030 and 2031, and disruption of sll1848 encoding a major LPAAT of Synechocystis by inserting a kanamycin-resistant gene cassette (KnR). (B) Schematic representation of lipid biosynthesis in Synechocystis and Cyanothece. GPAT, glycerol-3-phosphate acyltransferase; 14:0-ACP, ACP binding a myristic acid (14:0); 16:0-ACP, ACP binding a palmitic acid (16:0); 18:0/14:0 PA, PA binding stearic acid (18:0) and 14:0; 18:0/16:0 PA, PA binding 18:0 and 16:0. (C) Contents of fatty acids in glycerolipids of the WT Synechocystis (green), T1274Δ1848 (blue), and Cyanothece (pink) cells. Total lipids were extracted from the cells using the Bligh–Dyer method, and fatty acids bound to glycerolipids were methyl-esterified with KOH–methanol. Fatty acid methyl esters were analyzed by gas chromatography. 16:1, palmitoleic acid; 18:1, oleic acid; 18:2, linoleic acid; and 18:3, γ-linolenic acid. *P < 0.01, calculated using ANOVA test.

The fluidity of a membrane is affected by the composition of FAs bound to membrane lipids. To clarify the effects of 14:0 on the biophysical characteristics of the thylakoid membrane, the fluidity of the thylakoid membranes isolated from Synechocystis cells, T1274Δ1848, and Cyanothece cells was measured using the electron paramagnetic resonance (EPR) spectroscopy with a spin probe, 16-DOXYL stearic acid (16SAL) (Sigma-Aldrich, MA, USA), at room temperature. The moment parameters calculated from the EPR spectra of the thylakoid membranes isolated from the T1274Δ1848 and Cyanothece cells were significantly lower than that from WT cells (Table 1). These results indicated that the fluidity of the thylakoid membranes from Cyanothece and T1274Δ1848 cells was higher than those of the wild-type Synechocystis cells because of the high 14:0 levels in their glycerolipids.

Table 1.

Parameters were obtained from EPR spectrum to measure the fluidity of thylakoid membranes

Strains EPRt0 (×10−11 s)
S. WT 5.60 ± 0.35
S. T1274Δ1848 *4.22 ± 0.47
Cyanothece *4.78 ± 0.20
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The moment parameter, τ0, was calculated from the EPR spectrum using an equation described in the “Materials and Methods” section.

*

P < 0.01, calculated using ANOVA test.

An increase in the 14:0 content in Synechocystis-enhanced PSII repair and de novo D1 synthesis

Photosynthetic activities of the WT, T1274Δ1848, and Cyanothece cells in the presence of NaHCO3 (net photosynthesis activity) or 1, 4-benzoquinone (PSII activity) were measured using a Clark-type oxygen electrode (Supplementary Fig. S2). Cyanothece cells showed a lower net photosynthesis activity than the WT and T1274Δ1848 cells (Supplementary Fig. S2A). The net activity of T1274Δ1848 cells was similar to that of the WT cells. Thus, the net activity was not related to the content of 14:0 glycerolipids. The PSII activities of the WT, T1274Δ1848 and Cyanothece cells were similar (Supplementary Fig. S2B).

To test whether the tolerance of photosynthesis to intense light was related to the high content of 14:0, the WT and T1274Δ1848 cells were exposed to 1500 μmol photons m−2 s−1 at 32°C and changes in PSII activity were measured (Fig. 3). The PSII activity in both cells declined under strong light, which was less conspicuous in the T1274Δ1848 than the WT cells (Fig. 3A). The PSII photodamage rates measured in the presence of lincomycin were the same in the WT and T1274Δ1848 cells (Fig. 3B). The photoinhibition in the Δsll1848 mutant cells of Synechocystis (Δ1848) was also measured to determine whether the sll1848 disruption affected photoinhibition. As shown in Supplementary Fig. S3, there was no difference in the photoinhibition of PSII between the WT and Δ1848 cells; thus, there was no effect of sll1848 on the photoinhibition of PSII. The rate constants for photoinactivation (Kpi) in WT and T1274Δ1848 cells were similar, 0.0180 ± 0.0038 and 0.0262 ± 0.002 min−1, respectively. However, the rate constants for PSII repair (Krec) in WT and T1274Δ1848 cells were significantly different, 0.0273 ± 0.0048 and 0.072 ± 0.0336 min−1, respectively. These results indicated that T1274Δ1848 cells had a higher PSII repair activity than the WT cells.

Fig. 3.

Fig. 3

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Photoinhibition of PSII in the WT and T1274Δ1848 cells. WT (green circles) and T1274Δ1848 cells (blue squares) were incubated at 32°C under strong light at 1500 μmol photons m−2 s−1 with aeration by ambient air in the absence (A) or presence (B) of 200 μg ml−1 lincomycin. The PSII activity of the WT and T1274Δ1848 cells was measured in terms of O2 evolution in the presence of 1 mM 1,4-benzoquinone and 1 mM K3Fe(CN)6. The mean activities ± SD considered as 100% for the WT and T1274Δ1848 cells were 309.5 ± 10.0 and 325.7 ± 14.2 μmol O2 mg−1 Chl h−1, respectively. The values are the mean of the results from triplicates ± SD. *P < 0.01, calculated using ANOVA test.

The de novo synthesis of D1, a reaction center protein of PSII, plays a central role in PSII repair (Murata and Nishiyama 2018). Thus, the translational activity was measured under the same conditions as those used for the analysis of the photoinhibition of PSII in the WT and T1274Δ1848 cells using 35S-labeled methionine/cysteine (35S-Met/Cys). In both the WT and T1274Δ1848 cells, the newly synthesized proteins in the thylakoid membrane, especially D1, were labeled with 35S-Met/Cys for 20 min (Fig. 4A). Densitometric analysis of the labeled D1 indicated that D1 was synthesized linearly in both the WT and T1274Δ1848 cells, and the rate of de novo synthesis of D1 was 1.65-fold faster in the T1274Δ1848 than the WT cells (Fig. 4B). These results indicated that an increase of 14:0 in the glycerolipids enhanced PSII repair by accelerating the de novo synthesis of D1.

Fig. 4.

Fig. 4

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De novo synthesis of proteins in thylakoid membranes of the WT and T1274Δ1848 cells under strong light. Proteins of the WT and T1274Δ1848 cells were pulse-labeled by incubating cells with 35S-labeled Met and Cys for the indicated times under strong light at 1500 μmol photons m−2 s−1. Thylakoid membranes were isolated and membranes corresponding to 4 μg Chl a were analyzed by SDS–PAGE. (A) Representative radiogram of pulse-labeled proteins from thylakoid membranes. (B) Quantification of the relative levels of the labeled D1 protein in the WT (green circles) and T1274Δ1848 cells (blue squares). The value for the WT cells after pulse-labeling for 20 min was designated as 1.0. *P < 0.01, calculated using ANOVA test.

14:0 reduced 1O2 production

Excess activity of the photosynthetic electron transport under strong light induces energy transfer to molecular oxygen (O2) and produces ROS such as 1O2. 1O2 inhibited translational elongation involved in the repair of the photodamaged PSII (Kusama et al. 2015, Jimbo et al. 2018). Thus, the 1O2 production rates were measured in the WT, T1274Δ1848, and Cyanothece cells under 2000 μmol photons m−2 s−1 at 32°C using a histidine-trapping method (Rehman et al. 2013). The rate of 1O2 production in T1274Δ1848 and Cyanothece cells were slower than that in WT Synechocystis cells (Fig. 5).

Fig. 5.

Fig. 5

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Rates of 1O2 production in the S. WT, S. T1274Δ1848 and Cyanothece cells. The production rates of 1O2 in the S. WT (green), S. T1274Δ1848 (blue) and Cyanothece cells (pink) were monitored in terms of the difference between the uptake of O2 in the presence and absence of 5 mM histidine under intense light at 2000 μmol photons m−2 s−1. *P < 0.01, calculated using ANOVA test.

Production of ROS is associated with the content of carotenoids and the distribution of energy absorbed by the antennae to the PSII and PSI (Chen et al. 2015, Izuhara et al. 2020). First, the fluorescence emission spectra of the WT, T1274Δ1848 and Cyanothece cells were measured at 77 K. When chlorophylls (Chls) were excited by monochromatic light at 435 nm, the fluorescence spectra of the WT and T1274Δ1848 cells were similar, and the F725/F695 ratios were 2.42 ± 0.63 and 2.47 ± 1.1, respectively (Supplementary Fig. S4). These results indicated that PSI/PSII ratios were similar in the WT and T1274Δ1848 cells and were unaffected by the 14:0 content. In contrast, when phycobilins were excited by monochromatic light at 580 nm in the WT and T1274Δ1848 cells, significant changes in energy transfer from the phycobilisomes to PSII (F695) or PSI (F725) between the WT and T1274Δ1848 cells were observed (Fig. 6). The F725/F695 ratios calculated using the spectra were 0.75 ± 0.13, 0.82 ± 0.14 and 1.06 ± 0.13 in the WT, T1274Δ1848 and Cyanothece cells, respectively. These results indicated that the higher level of 14:0 in glycerolipids modified energy transfer from the phycobilisomes to the photosystems. We also check the content of carotenoids by measuring the absorption spectrum of the cell extract. The absorption at 470 nm that corresponds to the absorption by carotenoids in the cell extract from T1274Δ1848 cells was higher than that from WT cells (Fig. 7). The content of carotenoids determined spectroscopically in T1274Δ1848 cells (0.331 ± 0.164 μg μg−1 Chl a) was significantly higher than that in the WT (0.167 ± 0.035 μg μg−1 Chl a). This result suggests that higher level of 14:0 in glycerolipids increased carotenoid content as well.

Fig. 6.

Fig. 6

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Fluorescence spectra of the S. WT, S. T1274Δ1848 and Cyanothece cells. (A) Fluorescence spectra of the S. WT (green), S. T1274Δ1848 (blue) and Cyanothece (pink) cells with excitation at 580 nm and 77 K were normalized with the fluorescence peak of CP43. (B) Differential spectra were calculated by subtracting the spectra of the S. WT cells from those of the S. T1274Δ1848 (blue) and Cyanothece cells (pink). Peaks were annotated according to Ogawa et al. (2013).

Fig. 7.

Fig. 7

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Absorption spectra of the extracts from WT and T1274Δ1848 cells. Absorption spectra (400–700 nm) of the extracts from WT (green), T1274Δ1848 (blue) cells were normalized with the absorption at 663 nm. The values are the mean of the results from triplicates ± SD.

Discussion

Role of 14:0-containing glycerolipids in PSII repair

The present study demonstrated that an increase in the 14:0 content of glycerolipids in the thylakoid membrane of Synechocystis cells enhanced PSII repair by suppressing ROS production, which inhibited the de novo synthesis of D1 required for the recovery of the photodamaged PSII. Most thylakoid membranes in cyanobacteria and chloroplasts contain glycerolipids that bind to highly polyunsaturated fatty acids (PUFAs) (Wada and Murata 1998), which make the membranes more fluid than other membranes that do not contain PUFAs. The fluidity of thylakoid membranes is critical for the proper functioning and maintenance of photosynthesis under environmental stresses (Wada et al. 1990, 1994, Murata et al. 1992, Wada and Murata 2004). However, the mechanism by which the fluidity of the thylakoid membranes affected photosynthesis is not fully understood. Here, we revealed that an increase in the fluidity of the thylakoid membranes by the modification of glycerolipids with 14:0 enhanced PSII repair under strong light. The 14:0-containing glycerolipids increased membrane fluidity and energy transfer from phycobilisomes to PSI. This transfer of light energy absorbed by the antennae complexes to photosystems by relocating the antennae complexes to PSII (state I) and PSI (state II) is referred to as state transition, which is a part of the nonphotochemical quenching mechanisms that reduces the transfer of excess light energy to the photosystems to suppress ROS production (Minagawa 2011). Although the structures of antenna complexes are different between plants and cyanobacteria, both regulatory mechanisms rely on the redox state of the plastoquinone (PQ) pool (Minagawa 2011, Calzadilla and Kirilovsky 2020). Membrane fluidity affects the mobility of protein complexes, including antennae complexes, in the thylakoid membranes (Pali et al. 2003); thus, the states of antennae complexes are altered by membrane fluidity and the redox state of the PQ pool. Notably, the increased fluidity of the thylakoid membranes may also enhance PSII repair by accelerating the disassembly of PSII dimers to PSII monomers and then to CP43-less PSII monomers (Nickelsen and Rengstl 2013, Jimbo and Wada 2023). However, the molecular mechanisms by which the increase in the fluidity of thylakoid membranes enhances energy transfer from phycobilisomes to PSI (state II) remain to be elucidated. Furthermore, 1O2 can be quenched by carotenoids in thylakoid membranes. The content of carotenoids in T1274Δ1848 cells was higher than that in WT cells (Fig. 7). Therefore, the increase in 14:0 content altered the carotenoid content and affected the 1O2 production in T1274Δ1848 cells. The molecular mechanism underlying the observed increase in carotenoid contents in the mutant cells also remains to be investigated.

In this study, the introduction of 14:0 into glycerolipids accelerated PSII repair. In our previous study, supplementation of the BG-11 medium with 14:0 slightly inhibited PSII repair in Synechocystis cells under intense light (Jimbo et al. 2020). Under strong light, the supplemented free fatty acids were specifically incorporated into the sn-2 position of phosphatidylglycerol (PG), which is the major phospholipid in cyanobacterial cells, through the turnover of PG (Jimbo et al. 2021b, Kojima et al. 2022). The FAs incorporated from the medium reached 20 mol% of the total FAs in PG (Jimbo et al. 2021b), which was as little as 2 mol% of the cellular FAs. Therefore, the incorporation of 14:0 into PG may affect some biological processes related to PSII repair rather than the fluidity of thylakoid membranes. The chain length of FAs bound to the sn-2 position of PG played crucial roles in PSII activity and the accumulation of Chl a (Endo et al. 2022), suggesting a specific role of FAs bound to the PG molecules. T1274Δ1848 cells contained more than 20 mol% of the total FAs. Together with the previous findings, particular modification of fatty acids in PG molecules would inhibit PSII repair. However, modifying FAs in all glycerolipids can mask the inhibitory effect of modified PG molecules and even enhance PSII repair.

Ecophysiological significance of 14:0 in glycerolipids

Most cyanobacterial strains contain 16:0 as the major FA in the sn-2 position of the glycerolipids (Murata et al. 1992). The present study demonstrated that introducing 14:0 into the sn-2 position of glycerolipids in Synechocystis cells enhanced PSII repair under intense light, which appeared advantageous for photosynthetic organisms. The question thus arises regarding the reason behind most cyanobacterial strains containing 16:0 but not 14:0 in the sn-2 position of glycerolipids. The higher fluidity of thylakoid membranes induced by the elevated levels of 14:0 enhanced the energy transfer from phycobilisomes to PSI (state II). State II accelerates the excitation of the acceptor side, which is suitable for driving photosynthetic electron transfer under strong light. However, the frequent excitation of the acceptor side of the PSI causes its photoinhibition, which is toxic for photosynthetic organisms because the inactivated PSI is not repaired as rapidly as PSII (Sonoike 2011, Allahverdiyeva et al. 2015). Some cyanobacterial strains isolated from marine/brackish water, such as Spirulina sp. 6313, Synechococcus WH7803 and Prochlorococcus marinus, have high 14:0 content in their glycerolipids (Kenyon et al. 1972, Pittera et al. 2018). On the ocean surface, the intensity of sunlight reaches more than 3000 μmol photons m−2 s−1, which is adverse for photosynthetic organisms. Therefore, these strains may evolutionarily adjust the allocation of photosynthetic antennae to PSI (state II) to adapt to the intense light conditions, and the high content of 14:0 may help in the adaptation. If applied in these 14:0-containing cyanobacterial strains, future gene-editing techniques can provide us with a better ecophysiological understanding of the role of 14:0 in environmental adaptation.

Materials and Methods

Strains used and culture conditions

The WT and transformant Synechocystis and Cyanothece cells were grown photoautotrophically in liquid BG-11 medium at 32°C under 10 μmol photons m−2 s−1 and aeration by sterile air. Cultures with an OD730 of 1.0 ± 0.2, which corresponded to a Chl concentration of ∼ 3.6 mg ml−1, were used for the assays. A sll1848 Synechocystis mutant [Δ1848] (Okazaki et al. 2006) was gifted by Prof. Ikuo Nishida (Saitama University, Japan). T1274Δ1848 cells were generated by transforming Δ1848 cells with a DNA construct containing PpsbA-T1274:CmR as described previously (Saito et al. 2018). Transformants were isolated on a BG-11 plate containing 40 μg ml−1 chloramphenicol. The complete segregation of all the copies of the Synechocystis genome was confirmed by PCR.

Measurements of photosynthetic activities and detection of 1O2

Cells were exposed to 1500 μmol photons m−2 s−1 at 32°C for specific times to induce the photoinhibition of PSII as described previously (Jimbo and Wada 2023). For photodamage assays, lincomycin was added to the cell suspension at a final concentration of 200 μg ml−1 just before illumination. PSII activity was measured in terms of O2 evolution in the presence of 1 mM 1,4-benzoquinone and 1 mM K3Fe(CN)6 with a Clark-type oxygen electrode (Hansatech Instruments, Pentney, UK) under 2000 μmol photons m−2 s−1 at 32°C. The rate constants of photoinactivation (Kpi) and PSII repair (Krec) were determined according to Campbell and Tyystjärvi (2012). The net photosynthesis activity was also measured in terms of O2 evolution in the presence of 1 mM NaHCO3. For the detection of 1O2, the protocol, as described previously, was followed with a slight change (Kusama et al. 2015, Izuhara et al. 2020). Cells were exposed to 2000 μmol photons m−2 s−1 at 32°C in the presence and absence of 5 mM histidine, and O2 evolution was measured in the absence of electron acceptors. The generation of 1O2 was quantitated by subtracting the rate of O2 evolution in the absence of His from that observed in its presence.

Lipid analysis

Lipids were extracted from cells as described previously (Jimbo et al. 2021a). Fatty acids bound to glycerolipids were methyl-esterified by the KOH–methanol as described previously (Jimbo et al. 2021b). The fatty acid methyl esters obtained were analyzed by gas chromatography.

Measurement of the fluorescence spectra

Cells corresponding to 5 µg Chl a ml−1 were frozen in a cryocell using liquid N2 and transferred to the sample holder of RF-5300PC for fluorescence spectroscopy (SHIMADZU, Kyoto, Japan). The fluorescence emitted at 430–650 nm at excitation wavelengths of 435 or 580 nm was recorded. The fluorescence peaks were annotated according to a previous study (Ogawa et al. 2013).

Measurement of absorption spectra

Chl a and carotenoids in isolated thylakoid membrane were extracted in the 80% acetone. Absorption spectra of the cell extract were measured with spectrophotometer (V-730BIO, JASCO, Japan). The concentrations of Chl a and carotenoids were determined as described previously (Lichtenthaler and Wellburn 1983).

Labeling of proteins in vivo

Labeling of proteins in vivo with 35S Met/Cys was performed and the labeled proteins were analyzed according to Jimbo et al. (2019).

Assay of fluidity of the thylakoid membrane

For assaying the fluidity of the thylakoid membranes by EPR, the thylakoid membranes were isolated from the cells by disrupting with glass beads in KD buffer (50 mmol l−1 MES-NaOH [pH 6.0], 10 mmol l−1 MgCl2, 5 mmol l−1 CaCl2 and 25% glycerol). The fluidity of thylakoid membranes was analyzed using the EPR spectrum as described previously (Murata et al. 1975, Wada et al. 1984). Thylakoid membranes isolated from the WT, T1274Δ1848 and Cyanothece cells were adjusted to 0.6 mg Chl a ml−1 in 50 mM MES-NaOH buffer (pH 6.0) containing 10 mM MgCl2, 5 mM CaCl2, 25% (w/v) glycerol, 0.5 mM K3[Fe (CN)6], 0.5 mM K4[Fe (CN)6] and 100 µM of 16SAL. EPR spectra were measured using Magnettech ESR5000 (Bruker, Billerica, MA, USA) with a microwave at a power of 5 mW, modulation width of 2.0 Gauss and modulation frequency of 100 kHz. Samples were placed in a glass capillary tube and sealed with cray at one end before insertion into the EPR cavity. The rotational correlation times of 16SAL in the samples were calculated using the formula mentioned below.

graphic file with name UM0001-Latex.gif
graphic file with name UM0002-Latex.gif

Supplementary Material

pcae021_Supp
pcae021_supp.zip (271.1KB, zip)

Acknowledgments

The authors would like to thank Mr. Tsutomu Takizawa (the University of Tokyo) for technical assistance in the radioisotopic analysis of proteins.

Contributor Information

Kazuki Kurima, Department of Life Sciences, Graduate School of Arts and Sciences, The University of Tokyo, 3-8-1 Komaba, Meguro-ku, Tokyo 153-8902, Japan.

Haruhiko Jimbo, Department of Life Sciences, Graduate School of Arts and Sciences, The University of Tokyo, 3-8-1 Komaba, Meguro-ku, Tokyo 153-8902, Japan.

Takashi Fujihara, Comprehensive Analysis Center for Science, Saitama University, 255 Shimo-Okubo, Sakura-ku, Saitama 338-8570, Japan.

Masakazu Saito, Department of Life Sciences, Graduate School of Arts and Sciences, The University of Tokyo, 3-8-1 Komaba, Meguro-ku, Tokyo 153-8902, Japan; Department of Bioscience, Nagahama Institute of Bio-Science and Technology, 1266, Tamura, Nagahama, Shiga 526-0829, Japan.

Toshiki Ishikawa, Department of Biochemistry and Molecular Biology, Graduate School of Science and Engineering, Saitama University, 255 Shimo-Okubo, Sakura-ku, Saitama 338-8570, Japan.

Hajime Wada, Department of Life Sciences, Graduate School of Arts and Sciences, The University of Tokyo, 3-8-1 Komaba, Meguro-ku, Tokyo 153-8902, Japan.

Supplementary Data

Supplementary Data are available at PCP online.

Data Avaiavility

The data underlying this article are available in the article and in its online supplementary material.

Funding

Grants-in-Aid for Scientific Research from the Japan Society for the Promotion of Science (No. 22K14795 to H.J. and No. 20K06701 to H.W. I part); a research fund for young researchers from the Institute for Fermentation, Osaka (IFO), Japan (No. Y-2021-2-012 to H.J.).

Author Contributions

K.K. performed most of the experiments and wrote the first draft of the manuscript; T.F. and T.I. contributed to the measurement and analysis of EPR spectra; H.J. designed the research, wrote the manuscript and edited the final version of the manuscript; H.W. supervised the project, wrote the manuscript and edited the final version of the manuscript.

Disclosures

The authors have no conflicts of interest to declare.

References

  1. Allahverdiyeva  Y., Suorsa  M., Tikkanen  M. and Aro  E.M. (2015) Photoprotection of photosystems in fluctuating light intensities. J. Exp. Bot.  66: 2427–2436. [DOI] [PubMed] [Google Scholar]
  2. Asada  K. (1999) The water-water cycle in chloroplasts: scavenging of active oxygens and dissipation of excess photons. Ann. Rev. Plant Physiol. Plant Mol. Biol.  50: 601–639. [DOI] [PubMed] [Google Scholar]
  3. Bandyopadhyay  A., Elvitigala  T., Welsh  E., Stockel  J., Liberton  M., Min  H., et al. (2011) Novel metabolic attributes of the genus cyanothece, comprising a group of unicellular nitrogen-fixing Cyanothece. mBio  2: e00214–11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Calzadilla  P.I. and Kirilovsky  D. (2020) Revisiting cyanobacterial state transitions. Photochem. Photobiol. Sci.  19: 585–603. [DOI] [PubMed] [Google Scholar]
  5. Campbell  D.A. and Tyystjärvi  E. (2012) Parameterization of photosystem II photoinactivation and repair. Biochim. Biophys. Acta  1817: 258–265. [DOI] [PubMed] [Google Scholar]
  6. Chen  Z., Zhan  J., Chen  Y., Yang  M., He  C., Ge  F., et al. (2015) Effects of phosphorylation of β subunits of phycocyanins on state transition in the model cyanobacterium Synechocystis sp. PCC 6803. Plant Cell Physiol.  56: 1997–2013. [DOI] [PubMed] [Google Scholar]
  7. Endo  K., Abe  M., Kawanishi  N., Jimbo  H., Kobayashi  K., Suzuki  T., et al. (2022) Crucial importance of length of fatty-acyl chains bound to the sn-2 position of phosphatidylglycerol for growth and photosynthesis of Synechocystis sp. PCC 6803. Biochim. Biophys. Acta  1867: 159158–159164. [DOI] [PubMed] [Google Scholar]
  8. Huang  T.C. and Chow  T.J. (1988) Comparative studies of some nitrogen-fixing unicellular cyanobacteria isolated from rice fields. Microbiology  134: 3089–3097. [Google Scholar]
  9. Izuhara  T., Kaihatsu  I., Jimbo  H., Takaichi  S. and Nishiyama  Y. (2020) Elevated levels of specific carotenoids during acclimation to strong light protect the repair of photosystem II in Synechocystis sp. PCC 6803. Front. Plant Sci.  11: 1030–1039. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Jimbo  H., Izuhara  T., Hirashima  T., Endo  K., Nakamura  Y. and Wada  H. (2021a) Membrane lipid remodeling is required for photosystem II function under low CO2. Plant J.  105: 245–253. [DOI] [PubMed] [Google Scholar]
  11. Jimbo  H., Izuhara  T., Hihara  Y., Hisabori  T. and Nishiyama  Y. (2019) Light-inducible expression of translation factor EF-Tu during acclimation to strong light enhances the repair of photosystem II. Proceedings of the National Academy of Sciences.  116: 21268–21273. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Jimbo  H., Yuasa  K., Takagi  K., Hirashima  T., Keta  S., Aichi  M., et al. (2021b) Specific incorporation of polyunsaturated fatty acids into the sn-2 position of phosphatidylglycerol accelerates photodamage to photosystem II under strong light. Int. J. Mol. Sci.  22: 10432–10444. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Jimbo  H., Takagi  K., Hirashima  T., Nishiyama  Y. and Wada  H. (2020) Long-chain saturated fatty acids, palmitic and stearic acids, enhance the repair of photosystem II. Int. J. Mol. Sci.  21: 7509–7518. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Jimbo  H. and Wada  H. (2023) Deacylation of galactolipids decomposes photosystem II dimers to enhance degradation of damaged D1 protein. Plant Physiol.  191: 87–95. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Jimbo  H., Yutthanasirikul  R., Nagano  T., Hisabori  T., Hihara  Y. and Nishiyama  Y. (2018) Oxidation of translation factor EF-Tu inhibits the repair of photosystem II. Plant Physiol.  176: 2691–2699. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Kenyon  C.N., Rippka  R. and Stanier  R.Y. (1972) Fatty acid composition and physiological properties of some filamentous blue-green algae. Arch. Mikrobiol.  83: 216–236. [DOI] [PubMed] [Google Scholar]
  17. Kojima  K., Matsumoto  U., Keta  S., Nakahigashi  K., Ikeda  K., Takatani  N., et al. (2022) High-light-induced stress activates lipid deacylation at the sn-2 position in the cyanobacterium Synechocystis sp. PCC 6803. Plant Cell Physiol.  63: 82–91. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Kusama  Y., Inoue  S., Jimbo  H., Takaichi  S., Sonoike  K., Hihara  Y., et al. (2015) Zeaxanthin and echinenone protect the repair of photosystem II from inhibition by singlet oxygen in Synechocystis sp. PCC 6803. Plant Cell Physiol.  56: 906–916. [DOI] [PubMed] [Google Scholar]
  19. Lichtenthaler  H.K. and Wellburn  A.R. (1983) Determinations of total carotenoids and chlorophylls a and b of leaf extracts in different solvents. Biochem. Soc. Trans.  11: 591–592. [Google Scholar]
  20. Minagawa  J. (2011) State transitions-the molecular remodeling of photosynthetic supercomplexes that controls energy flow in the chloroplast. Biochim. Biophys. Acta  1807: 897–905. [DOI] [PubMed] [Google Scholar]
  21. Murata  N. and Nishiyama  Y. (2018) ATP is a driving force in the repair of photosystem II during photoinhibition. Plant Cell Environ.  41: 285–299. [DOI] [PubMed] [Google Scholar]
  22. Murata  N., Troughton  J.H. and Fork  D.C. (1975) Relationships between the transition of the physical phase of membrane lipids and photosynthetic parameters in anacystis nidulans and lettuce and spinach chloroplasts. Plant Physiol.  56: 508–517. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Murata  N., Wada  H. and Gombos  Z. (1992) Modes of fatty-acid desaturation in cyanobacteria. Plant Cell Physiol.  33: 933–941. [Google Scholar]
  24. Nickelsen  J. and Rengstl  B. (2013) Photosystem II assembly: from cyanobacteria to plants. Annu. Rev. Plant Biol.  64: 609–635. [DOI] [PubMed] [Google Scholar]
  25. Nishiyama  Y., Yamamoto  H., Allakhverdiev  S.I., Inaba  M., Yokota  A. and Murata  N. (2001) Oxidative stress inhibits the repair of photodamage to the photosynthetic machinery. Embo J.  20: 5587–5594. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Ogawa  T., Harada  T., Ozaki  H. and Sonoike  K. (2013) Disruption of the ndhF1 gene affects Chl fluorescence through state transition in the cyanobacterium Synechocystis sp. PCC 6803, resulting in apparent high efficiency of photosynthesis. Plant Cell Physiol.  54: 1164–1171. [DOI] [PubMed] [Google Scholar]
  27. Okazaki  K., Sato  N., Tsuji  N., Tsuzuki  M. and Nishida  I. (2006) The significance of C16 fatty acids in the sn-2 positions of glycerolipids in the photosynthetic growth of Synechocystis sp. PCC 6803. Plant Physiol.  141: 546–556. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Pali  T., Garab  G., Horvath  L.I. and Kota  Z. (2003) Functional significance of the lipid-protein interface in photosynthetic membranes. Cell Mol. Life Sci.  60: 1591–1606. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Pittera  J., Jouhet  J., Breton  S., Garczarek  L., Partensky  F., Marechal  E., et al. (2018) Thermoacclimation and genome adaptation of the membrane lipidome in marine Synechococcus. Environ. Microbiol.  20: 612–631. [DOI] [PubMed] [Google Scholar]
  30. Rehman  A.U., Cser  K., Sass  L. and Vass  I. (2013) Characterization of singlet oxygen production and its involvement in photodamage of photosystem II in the cyanobacterium Synechocystis PCC 6803 by histidine-mediated chemical trapping. Biochim. Biophys. Acta  1827: 689–698. [DOI] [PubMed] [Google Scholar]
  31. Saito  M., Endo  K., Kobayashi  K., Watanabe  M., Ikeuchi  M., Murakami  A., et al. (2018) High myristic acid content in the cyanobacterium Cyanothece sp. PCC 8801 results from substrate specificity of lysophosphatidic acid acyltransferase. Biochim. Biophys. Acta  1863: 939–947. [DOI] [PubMed] [Google Scholar]
  32. Sonoike  K. (2011) Photoinhibition of photosystem I. Physiol. Planta  142: 56–64. [DOI] [PubMed] [Google Scholar]
  33. Stanier  R.Y., Kunisawa  R., Mandel  M. and Cohen-Bazire  G. (1971) Purification and properties of unicellular blue-green algae (order Chroococcales). Bacteriol. Rev  35: 171–205. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Wada  H., Gombos  Z. and Murata  N. (1990) Enhancement of chilling tolerance of a cyanobacterium by genetic manipulation of fatty acid desaturation. Nature  347: 200–203. [DOI] [PubMed] [Google Scholar]
  35. Wada  H., Gombos  Z. and Murata  N. (1994) Contribution of membrane lipids to the ability of the photosynthetic machinery to tolerate temperature stress. Proc. Natl. Acad. Sci. USA  91: 4273–4277. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Wada  H., Hirasawa  R., Omata  T. and Murata  N. (1984) The lipid phase of thylakoid and cytoplasmic membranes from the blue-green algae (cyanobacteria), Anacystis nidulans and Anabaena variabilis. Plant Cell Physiol.  25: 907–911. [Google Scholar]
  37. Wada  H. and Murata  N. (1998) Lipids in Photosynthesis: Structure, Function and Genetics. Kluwer Academic Publishers, Dourdrecht, the Netherlands. [Google Scholar]
  38. Wada  H. and Murata  N. (2004) Membrane Lipids in Cyanobacteria. In  Lipids in Photosynthesis: Structure, Function and Genetics. pp. 65–81. Springer: Dordrecht. [Google Scholar]
  39. Weier  D., Muller  C., Gaspers  C. and Frentzen  M. (2005) Characterisation of acyltransferases from Synechocystis sp. PCC 6803. Biochem. Biophys. Res. Commun.  334: 1127–1134. [DOI] [PubMed] [Google Scholar]

Associated Data

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Supplementary Materials

pcae021_Supp
pcae021_supp.zip (271.1KB, zip)

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