Cyanobacteria convert light energy into ATP and NADPH through photosynthesis and use it in metabolic reactions to fix CO2
. Various intracellular reactions, such as light harvesting, photosynthetic electron transfer, CO2
fixation, and metabolic reactions, are stringently regulated and optimized with regard to the surrounding environment. Photosystems and Rubisco are the most abundant proteins in cyanobacteria, and their gene expression levels fluctuate greatly in response to nutrient and light conditions. However, mRNA and protein levels of only 10–15% genes in the cyanobacterium Synechocystis
sp. PCC 6803 are correlated; specifically, the mRNA and protein levels of photosynthesis-related genes are rarely correlated, and their post-transcriptional regulation is complex [1
]. Furthermore, protein accumulation and activity are closely regulated by post-translational modifications (PTMs) [2
Post-translational modifications (PTMs), such as phosphorylation, acetylation, and methylation, are an important process that regulates protein activity [2
]. Among these, phosphorylation is a highly conserved PTM in several organisms from prokaryotes to eukaryotes, and it is closely associated with intracellular metabolism and signaling [3
]. Phosphorylation usually occurs on serine (Ser), threonine (Thr), and tyrosine (Tyr) residues of eukaryotic proteins. In addition, in prokaryotic proteins, phosphorylation also occurs on histidine, arginine, and lysine residues [4
]. Phosphorylation leads to the formation of a phosphate ester bond in the hydroxyl groups of amino acids, such as Ser, Thr, and Tyr, in proteins, and alters the protein structure by imparting a negative charge following the addition of a phosphate group. Citrate dehydrogenase in the tricarboxylic acid (TCA) cycle is a well-known enzyme that is inactivated by phosphorylation and activated by dephosphorylation [5
]. In cyanobacteria, a signal protein called PII is phosphorylated under nitrogen-limited conditions, and it then controls metabolism by facilitating the expression of genes encoding enzymes involved in nitrogen assimilation, the pentose phosphate pathway, and glycolysis [6
]. Regarding the photosynthetic apparatus, turn-over of the D1 protein in Arabidopsis
and remodeling of the photosystem II (PSII)–LHCII supercomplex in Chlamydomonas
have also been reported to be regulated by phosphorylation [8
]. Thus, phosphorylation is a key PTM involved in many cellular controls, such as enzyme activity, signal transduction, and photosynthetic apparatus restructuring, and elucidating the governing roles of phosphorylation is imperative to understand the regulatory mechanisms underlying metabolism and photosynthesis. The genome of Synechocystis
sp. PCC 6803 encodes 11 Ser/Thr kinase, 1 Tyr kinase, and 7 phosphatases [10
], and approximately 5% of the proteins in the cell are phosphorylated [11
]. A comprehensive shotgun phosphoproteome of Synechocystis
has been established [11
]. However, quantitative analysis of this cyanobacterium is limited by the low levels of phosphoproteins present in the cells and their susceptibility to ionization suppression during liquid chromatography-triple quadrupole mass spectrometry (LC-MS/MS) [16
In this study, target proteomics was used to quantitatively analyze the phosphorylation levels of Ser, Thr, and Tyr residues of specific proteins. To elucidate the regulation of intracellular metabolism and photosynthesis by protein expression and phosphorylation levels, we obtained data of the expression of proteins and their phosphorylation levels during photosynthesis and metabolism under various growth conditions (photoautotrophic, mixotrophic, heterotrophic, dark, and nitrogen-deprived conditions). Based on the obtained data, we demonstrated changes in protein expression and phosphorylation levels of the photosynthetic apparatus and metabolic enzymes under these growth conditions. Furthermore, we evaluated the association of regulation of protein expression and phosphorylation with adaptability to different environments.
Cyanobacteria have a photosynthetic apparatus and metabolic enzymes for carbon fixation, and their biosynthesis and metabolism require numerous amino acids and resources. The roles of PTMs, such as phosphorylation, are also pivotal because the environment can dramatically change in a short period. In this study, to elucidate their regulation and roles, the levels of proteins involved in photosynthesis and metabolism as well as their phosphorylation states under various growth conditions were analyzed using targeted proteomics.
Our results showed that the expression levels of only certain proteins changed in response to growth conditions. For example, under the Hetero condition, although photosynthesis was terminated, the levels of only RbcL, RbcS, and CO2
-concentrating mechanism proteins among the metabolic enzymes were drastically reduced. However, the levels of other enzymes in the Calvin cycle did not change significantly (Figure 2
a). As RbcL and RbcS are the most abundant proteins in the cell, reducing the excess RbcL and RbcS levels under the Hetero condition, where photosynthesis was inhibited, would greatly reduce the amount of amino acids required for protein biosynthesis.
Under nitrogen deficiency, cells lose phycobilisomes, a huge protein complex, and redistribute nitrogen to other highly important metabolites [24
]. In fact, the results of absorption spectra, presented in Figure 1
c, showed that the absorption peak of phycobilisome at 630 nm was drastically reduced, indicating phycobilisome degradation. However, not all subunits of the phycobilisome complex were uniformly degraded; levels of only CpcC, a rod-linker protein of the phycobilisome complex, were drastically reduced. This result indicates that the phycobilisome complex possibly dissociates under nitrogen deficiency, but the individual subunits remain intact, allowing resumption of photosynthesis as soon as the growth environment improves.
In contrast, phosphorylation levels varied greatly than protein levels under different growth conditions (Figure 3
a, Figure S2
). Regulation through PTMs, such as phosphorylation, may play an important role. In particular, little changes at the proteome level were observed under Dark condition (Figure 2
a). A previous study has reported that mRNA expression levels of photosynthesis-related genes decreased between 1 and 9 h after transition from light to dark [25
]. However, the effects of protein synthesis and degradation were still considered to be negligible because the change in protein levels was small under Dark condition after 6 h (Figure 2
a, Figure 3
b). In contrast, the phosphorylation levels of proteins involved in metabolism, such as Pgm, CcmM, RbcL, RbcS, anti-sigB1, and anti-sigB2, were evidently altered. These results suggest that the photosynthetic apparatus and metabolic enzymes are maintained at the proteome level in dark conditions and are regulated via phosphorylation such that cellular activity can be resumed immediately under light conditions.
Furthermore, the phosphorylation levels of CpcB, ApcA (Y17 or S19), OCP, and PsbV were elevated under nitrogen deficiency (Figure 4
). CpcB and ApcA construct the rod and core of the phycobilisome complex, respectively. Under nitrogen deficiency, the phycobilisome complex is degraded by Clp proteases and a small protein called NblA [26
]. During this process, NblA binds to the N
-terminal side of CpcB and ApcA and then dissociates the phycobilisome complex [27
]. The phosphorylation sites of CpcB and ApcA detected in this study are located on the N
-terminal side, and CpcB and ApcA phosphorylation may be involved in their binding with NblA. In addition, the phosphorylation levels of S118 or T121 in ApcA were reduced. This amino acid is located in the trimeric interaction domain region of ApcA [31
]. Therefore, phosphorylation of this amino acid is involved in ApcA trimerization, and its dephosphorylation under nitrogen deficiency may be involved in dissociation of the ApcA trimer. OCP, a water-soluble carotenoid-binding protein, has been suggested to be involved in NPQ, which dissipates excess light as heat [22
]. PsbV is a small subunit of PSII and contributes to the stabilization of the manganese cluster of PSII. Moreover, in Synechocystis
sp. PCC 6803, PsbV suppresses PSII activity under nitrogen deficiency [1
]. The psbV
-disrupted strains maintain PSII activity even under nitrogen deficiency, while cell proliferation is reduced, indicating that the suppression of PSII activity by PsbV plays an important role in growth under nitrogen deficiency [1
]. Our results, together with the previous reports, suggest that PsbV phosphorylation can suppress PSII activity under nitrogen deficiency.
Although analysis of the deficient strains and other factors are required to obtain direct evidence, based on the results of our proteomic and phosphoproteomic analyses as well as PAM measurements, a mechanism for adaptation to nitrogen deficiency was proposed, as illustrated in Figure 5
. First, degradation of phycobilisome via phosphorylation and suppression of PSII photosynthetic activity by PsbV phosphorylation may occur. Although our results showed no difference in Fv
values between Auto and −N conditions, Ogawa et al. [23
] have reported that Fv
under nitrogen deficiency is about half of the value under Auto condition. Moreover, Ogawa et al. [23
] have reported that the estimation of photosynthesis by PAM measurement of cyanobacteria is always problematic owing to the interference from respiratory electron transfer and phycocyanin fluorescence; however, subtracting basal phycobilisome and PSI fluorescence allows to accurately estimate the maximum quantum yield (ΦII) of PSII, and difference between the “apparent” and “true” maximum quantum yield of PSII becomes larger under high phycocyanin conditions. Our PAM measurement system could not measure the “true” maximum quantum yield of PSII and measured the “apparent” maximum quantum yield. However, the results reported by Ogawa et al. [23
] support our findings at the proteome and phosphoproteome levels. In other words, the uptake of light may be reduced at the entrance, such as phycobilisome, under nitrogen deficiency. ΦII and qP under nitrogen deficiency were lower than those under Auto condition, indicating that the rates of photosynthetic electron transfer and CO2
fixation were suppressed. In nitrogen-deficient cells, carbonate fixation was reduced owing to the blockade of metabolism caused by the C/N balance. In other words, the amount of energy required for the output seemed to have decreased.
Under −N (N24 and N48) conditions, qN and NPQ were reduced (Table 1
). This result suggests that the rate of dissipation of light energy as heat was reduced and the efficiency of input was increased. Under −N (N24 and N48) conditions, the PQ pool is drastically reduced compared with that under Auto condition, even at the same light intensity because of delayed metabolism. This notion is supported by decreased qP under nitrogen deficiency. Therefore, under nitrogen deficiency, cells are stressed, similar to that under high light conditions, and become acclimatized to higher light intensities. It seems that NPQ under nitrogen deficiency was lower than that under Auto condition, because PAM measurements were performed at an actinic light intensity of 80 μm−2
and cultivation was performed under a light intensity of 50 μm−2
. The results of phosphoproteomic analysis showed that OCP phosphorylation was accelerated under nitrogen deficiency, suggesting that heat dissipation by OCP is negatively regulated by phosphorylation.
Overall, our results indicate that, in addition to the regulation of protein expression, the regulation of phosphorylation levels of cyanobacterial photosynthetic apparatus and metabolic enzymes is crucial for adapting to changing environmental conditions. In the future, in addition to further detailed analyses of phosphoproteome, elucidation of the phosphorylation control of individual proteins is warranted to clarify the regulatory mechanism of overall photosynthesis, which is expected to allow for the artificial optimization of photosynthesis.
4. Materials and Methods
4.1. Culture Conditions
sp. PCC 6803 GT strain, isolated by Williams [35
], was used in this study. The cyanobacteria were grown in modified BG11 medium [36
] containing 5 mM NH4
Cl as a nitrogen source. Cells were grown in 200 mL of medium in 500 mL Erlenmeyer flasks for batch culture under photoautotrophic conditions with continuous light (~50 μmol μm−2
) at preculture. Synechocystis
sp. PCC 6803 (GT) was cultured under the following five conditions (Figure 1
a): (1) photoautotrophic (Auto; photosynthesis only), (2) mixotrophic (Mixo; photosynthesis and glucose utilization), (3) heterotrophic (Hetero; glucose utilization only), (4) dark (Dark), and (5) nitrogen-deprived (−N). Cultivation was performed by rotary shaking under five different nutrient conditions at 34°C with continuous illumination by LED lights at 50 μmol μm−2
. Under Mixo and Hetero conditions, 5 mM of glucose was added as a carbon source, and under Hetero condition, 10 μM of 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), a photosynthetic inhibitor, was added. Under −N and Dark conditions, cells were incubated under Auto until confluence (OD ~ 1.0) and then transferred to the respective conditions. Under −N, the cells were collected by centrifugation (4000× g
, 5 min, room temperature (RT)), and washed with a nitrogen-free medium (BG110
) and then re-cultured in the BG110
medium. Samples for proteomic analysis were collected 24 and 48 h after nitrogen deprivation (hereafter referred to as N24 and N48, respectively). Under Dark condition, upon reaching confluence (OD ~ 1.0), the culture was covered with an aluminum foil and then re-cultured in the dark for 6 h. Cells grown under Auto, Hetero, and Mixo conditions were collected when they reached an OD730
of ~1.0 (at 96 h, 48 h, and 48 h after inoculation, respectively). Cells grown under Dark condition was collected 6 h after culture in darkness. Cells grown under N24 and N48 were collected 24 and 48 h after being inoculated in BG110
4.2. Sample Preparation for Proteomic Analysis
Total proteins were extracted as described by Picotti et al. [37
]. The suspension containing cells in the logarithmic growth phase (200 mL, OD730
= 1.0) was collected by centrifugation (5000× g
, 4 °C, 5 min). Pellets were resuspended in 1 mL of lysis buffer (50 mM HEPES, 15% glycerol, 15 mM dithiothreitol (DTT), 100 mM KCl, 5 mM ethylenediamine-N,N,N′,N′
-tetraacetic acid disodium salt dihydrate (EDTA), one cOmplete protease inhibitors cocktail per 10 mL (Roche, Mannheim, Germany), and one PhosSTOP (Roche) per 10 mL). The suspension was transferred to an Eppendorf tube containing zirconia beads (diameter, 0.6 and 6 mm) and disrupted with Beads Crusher μT-12 (TAITEC, Saitama, Japan) (3000 min−1
, 6 min). The resulting solution was centrifuged (15,000× g
, 4 °C, 5 min), and the supernatant was transferred to a proteomics Eppendorf tube (protein low-adsorption tube) to obtain protein extraction samples. Protein concentration in the extracted sample was measured by the Bradford method, and the total protein content was adjusted to 3 mg. Denatured buffer (500 mM Tris–HCl, 10 mM EDTA, and 7 M guanidine HCl) was added to the adjusted samples to make up a total volume of 2.2 mL.
4.3. Reduction and Alkylation/Methanol Chloroform Precipitation
Dithiothreitol (DTT) 1 (50 mg mL−1
, 10 μL) was added to the protein samples and shaken at RT for 1 h using a tube mixer (CM-1000 Cute Mixer, EYELA, Tokyo, Japan). Next, 25 μL of 50 mg mL−1
iodoacetamide was added and shaken for 1 h to reduce/alkylate the proteins. Next, the proteins were purified by methanol/chloroform precipitation, as described by Wessel and Flügge [38
]. Cold methanol (6 mL) was added to the sample solution and mixed by inversion. Then, 1.5 mL of cold chloroform was added and mixed by inversion. Cold Milli-Q water (4.5 mL) was added and mixed by inversion, followed by centrifugation (4000× g
, 4 °C, 5 min). The upper layer was removed, and 450 μL of cold methanol was added and mixed gently by inversion. After centrifugation (4000× g
, 4 °C, 5 min) using a swing rotor, the supernatant was removed, and an additional cycle of centrifugation (4000× g
, 4 °C, 1 min) was performed to completely remove the supernatant.
4.4. Trypsin/LysC Digestion
Proteins were digested by trypsin and LysC (Promega). Trypsin hydrolyzes the ester bonds on the carboxyl side of Arg, while trypsin and LysC hydrolyze the ester bonds on the carboxyl side of Lys. The combination of trypsin and LysC enhances the efficiency of digestion. Therefore, trypsin/LysC digestion was carried out as described previously [27
]. To the supernatant obtained from the above steps, 90 μL of 6 M urea was added and shaken for about 10 min at RT using a tube mixer. Then, 360 μL of 0.1 M Tris-HCl (pH 8.5) was added, and ultrasonic treatment and standing on ice were repeated twice for 30 s using an ultrasonic washer (Branson 2510, Danbury, CT, USA) to resuspend the protein precipitate. Next, 10 μL of 0.5 mg mL−1
LysC solution and 25 μL of 1% Protease Max solution (Promega, Nacka, Sweden) were added and mixed by tapping, and the samples were incubated at 25 °C for 3 h. Finally, 10 μL of 0.5 mg mL−1
trypsin solution was added and mixed by tapping, and the samples were incubated at 37 °C for 16 h.
4.5. Samples Desalination
Sample desalination was performed as described previously [39
]. Milli-Q water (75 μL) and of 50% aqueous formic acid solution (30 μL) were added to the trypsin digestion product, and the mixture was stirred with a vortex mixer and centrifuged (15,000 rpm, 4 °C, 5 min). As a result, 580 μL of the supernatant was obtained. To prepare samples for relative quantitative analysis, 15
N samples (obtained by culturing Synechocystis
sp. PCC 6803 in BG11 medium with 15
Cl as the nitrogen source) and 14
N samples were mixed such that the protein contents were at a ratio of 1:1. The sample volumes were prepared such that the total peptide content could be 300 μg. Tryptic peptide concentrations were measured by the BCA method (Pierce BCA Protein Assay kit, Thermo, Rockford, USA).
The samples were diluted five times with Reagent A (5% acetonitrile, 0.1% formic acid). The diluted samples were desalted by MonoSpinC18 (GL Science, Tokyo, Japan). The column was equilibrated with the same bed volume of Reagent B (80% acetonitrile, 0.1% formic acid) and washed with Reagent A by centrifugation (200× g, 10 min, RT). The samples corresponding to 300 μg of proteins was loaded onto the column and centrifuged (200× g, 10 min, RT). The column was washed by a bed volume of Reagent A by centrifugation (200× g, 10 min, RT) twice. The same volume of Reagent B was added to the samples and centrifuged (200× g, 10 min, RT). The prepared samples were dried and solidified using a centrifugal concentrator and frozen until phosphopeptide enrichment.
4.6. Phosphopeptide Enrichment
Phosphopeptide enrichment was performed with a TiO2 column (GL science) as per the following protocol. First, the TiO2 column was equilibrated with 200 μL of Solution A (0.5% trifluoro acid and 80% acetonitrile), followed by 200 μL of Solution B (0.5% trifluoro acid, 80% acetonitrile, and 300 mg mL−1 lactic acid), and then centrifuged (100× g, 3 min, RT). Second, the desalted samples diluted with 2 mL of Solution B were loaded onto the TiO2 column and absorbed on the column by centrifugation (100× g, 3 min, RT). The column was washed first with 2 mL of Solution A once and then with Solution B twice by centrifugation (100× g, 3 min, RT). Finally, the absorbed phosphopeptides were eluted in 200 μL of 5% ammonium solution, followed by 200 μL of 5% pyrrolidine. The eluted solution was immediately acidified by 400 μL of 5% formic acid (pH, 2~3). The eluted acidified samples were desalted by MonoSpinC18 (GL Science) in the same manner mentioned before. Then, the desalted samples were dried and solidified with a centrifugal concentrator and frozen until nanoLC–MS/MS.
4.7. Design of MRM Assays
For quantitative proteomic analysis, 221 proteins related to the central metabolic pathways and photosynthetic apparatus were selected from the Kyoto Encyclopedia of Genes and Genomes (KEGG) database [42
]. The amino acid sequences of the target proteins were obtained from Cyanobase [43
]. MRM assays used to quantify these 221 proteins designed using the open-source software Skyline 2.6 [44
]. Each protein was subjected to a tryptic peptide filter of 8 to 25 residues, and 5 y-fragments (y1 to y5) were selected for each peptide. Samples of Synechocystis
sp. PCC 6803 were analyzed once by nanoLC–MS/MS (LCMS-8060, Shimadzu, Kyoto, Japan) using the provisional MRM assay. From the results of this analysis, peaks were selected based on the shape, coelution, and intensity, and the best transitions up to 5 were selected. For proteins with no suitable tryptic peptides and transitions, transitions were quantified for all y- and b-fragments, from which the tryptic peptides suitable for quantitation were selected again to design the final MRM assay.
For quantitative phosphoproteomic analysis, the list of targeted phosphoproteins was obtained from previous studies [12
]. Thus, 101 phosphoproteins mainly involved in photosynthesis, central metabolism, and transcriptional regulation were selected. For designing MRM assays, y- and b-fragments (5 with m/z > precursor) were selected for each phosphopeptide.
Finally, 101 targeted phosphoproteins in Synechocystis sp. PCC 6803 were analyzed once by nanoLC–MS/MS (LCMS-8060, Shimadzu) using the provisional MRM assay. From the results of this analysis, peaks were selected based on shape, coelution, and intensity, and the best 5 fragments were selected. The phosphopeptides with fewer than 3 fragments were excluded following analysis. Finally, the MRM assays for 32 phosphoproteins were successfully designed.
4.8. NanoLC–MS/MS Using MRM Assays
The obtained phosphopeptides were analyzed on a quadrupole mass spectrometer (LCMS-8060, Shimadzu) as described previously [45
]. Electrospray ionization (ESI) was performed, and the sample were separated by nanoLC (LC-20ADnano, Shimadzu). The analytical conditions were as follows: high-performance liquid chromatography column, L-column ODS (pore size: 5 μm, 0.1 × 150 mm; CERI, Tokyo, Japan); trap column, L-column ODS (pore size: 5 μm, 0.3 × 5 mm; CERI); solvent system, water (0.1% formic acid):acetonitrile (0.1% formic acid); gradient program, 10:90, v/v
at 0 min, 10:90 at 10 min, 40:60 at 45 min, 95:5 at 55 min, and 90:10 at 65 min; and flow rate, 400 nL·min−1
. MS was performed in the MRM mode, ESI was 1.6 kV, capillary temperature was 150 °C, collision gas was 270 kPa, resolution of Q 1 and Q 3 was low, dwell time was 1.0 ms, pause time was 1.0 ms, and retention time window was 2 min. Each peptide/phosphopeptide was quantified by the peak area ratio of 14
N samples to 15
N samples using Skyline 2.6.
4.9. Measurement of Ultraviolet–Visible (UV-VIS) Spectra
DU 800 was used to measure the UV-VIS spectra. Absorption spectra of cell suspensions were measured according to the “opal glass method,” with a translucent cuvette placed in front of the detector to minimize the effect of light scattering [46
]. The results obtained were normalized to absorbance at 730 nm (1.0).
4.10. Proteomic Data Analysis
The results of nanoLC–MS/MS were loaded into Skyline and quantified as the peak ratio of 14N samples to 15N internal standard samples.
For protein level, each peptide ratio was standardized by the mean of ratios under Auto conditions (n = 3). For more than two peptides of a single protein, the mean of all peptides was calculated. For a single peptide of a single protein, the obtained ratio was directly used for quantification. Finally, these parameters were expressed as fold change (FC).
For phosphoprotein level, each phosphopeptide ratio was standardized by the mean of ratios under Auto conditions (n = 3). The final FC of each phosphopeptide was calculated as the ratio of FCphosphopeptide to FCprotein to subtract the effect of background protein expression. The KEGG database and Cyanobase were referred to assess the function of each protein.
4.11. Chlorophyll Fluorescence Measurement
The PAM fluorescence measurements were performed using a fluorescence monitoring system (model FMS1; Hansatech, Norfolk, UK). Aliquots of 2.0 mL of suspension were transferred to the measurement chamber. Cells were maintained in the dark for 15 min. Modulated light was provided with a setting of 1 and an amplifier gain of 70. Actinic light was provided with a power setting of 15 (corresponding to a light intensity of 80 μmol m−2·s−1) to drive photosynthesis. Pulses of saturating light of 0.2 s duration were applied under power setting of 100 (light intensity 10,000 μmol m−2·s−1) at 30 s intervals to measure the maximum quantum yield. At the end of each measurement, DCMU (10 μM) was added in the presence of actinic light at a setting 30 to obtain Fm.