Growth and hydrogen production of the cyanobacterium Synechocystis sp. PCC 6803 in a marine salt medium

The cyanobacterium Synechocystis sp. PCC 6803 is a promising candidate for sustainable hydrogen production due to its ability to generate hydrogen under fermentative conditions. This study investigates the impact of marine salt (35 g L⁻¹) supplementation in BG11 medium on the growth, biochemical composition, and hydrogen production of Synechocystis sp. PCC 6803. Cultures were subjected to a three-phase experimental design consisting of growth, nitrogen starvation, and dark fermentation. Marine salt supplementation did not influence growth rate during the initial phase and did not hinder biomass accumulation under nitrogen-deprived conditions. Biochemical analyses revealed that marine salt did not affect carbohydrate accumulation but decreased polyhydroxybutyrate accumulation, while protein content remained comparable between treatments. Notably, cultures grown in marine salt-supplemented media exhibited moderately enhanced hydrogen production, achieving up to 9.14 ± 0.62 mL g^-1 dry weight over four days—slightly higher than in control cultures. Our results indicated that carbohydrates accumulated during the nitrogen starvation phase are only partially utilized for hydrogen production during the subsequent phase of dark fermentation, and that more than 90% of the hydrogen produced occurs within the first 3 days. These findings suggest that marine salt not only supports acceptable growth of Synechocystis but also enhances its hydrogen production potential by improving the sustainability of the process.

1 Introduction

The continuous rise in carbon emissions and atmospheric carbon dioxide levels has led to an increase in global surface temperatures, contributing to the urgent issue of global warming (Lee et al., 2023). Addressing this challenge requires the development of alternative energy sources that are sustainable and environmentally friendly. Among the various options, sunlight stands out as an abundant and renewable energy source. However, its effective long-term storage and utilization present significant challenges. In this context, hydrogen emerges as a promising energy carrier for the future (Karmaker et al., 2023).

Photobiological hydrogen production can be achieved by cyanobacteriophyta and green microalgae (Dzulkarnain et al., 2022). Cyanobacteriophyta are increasingly recognized as promising platforms for sustainable hydrogen production, not only for their ability to harness solar energy and fix atmospheric CO₂, but also because they can indirectly utilize photosynthetic electrons for hydrogenase-mediated hydrogen evolution by using the stored carbohydrates accumulated under stress conditions such as nitrogen deprivation. Their metabolic flexibility, genetic tractability, and ability to thrive in saline environments make them ideal candidates for scalable, low-input biohydrogen production systems. Among these, halotolerant and marine strains offer additional advantages for large-scale cultivation in saline environments, reducing freshwater dependency and enhancing resilience to environmental stresses. One of the most extensively studied halotolerant cyanobacteriophyta is Aphanothece halophytica, which demonstrates robust hydrogen production under dark fermentative conditions. When cultivated in natural seawater with minimal nutrient supplementation, A. halophytica can sustain hydrogen production for over two weeks under optimal conditions (35°C, pH 6.0, supplemented with glucose, NaCl, and Fe³⁺). This strain utilizes glycogen accumulated during photoautotrophic growth as a substrate for hydrogenase-mediated H₂ evolution (Taikhao et al., 2015). Recent studies have also explored the role of abiotic stress in enhancing hydrogen production in freshwater cyanobacteriophyta. For instance, Synechococcus elongatus PCC 7942 and its genetically engineered PAMCOD strain, which expresses the choline oxidase gene (codA), accumulate significant amounts of sucrose under salt and heat stress. This sucrose serves as an osmoprotectant and a fermentable carbon source at the same time (Broussos et al., 2024a).

The cyanobacterium Synechocystis sp. PCC 6803 is particularly noteworthy due to its bidirectional NiFe-hydrogenase, HoxEFUYH. This enzyme complex consists of a diaphorase sub-complex (HoxEFU), which interacts with redox partners like NAD(P)(H), ferredoxin, and flavodoxin, and a hydrogenase sub-complex (HoxYH), which catalyzes the production and consumption of hydrogen (Burgstaller et al., 2022). Hydrogen production in Synechocystis sp. PCC 6803 occurs under fermentative conditions that are typically induced by nitrogen deprivation followed by dark incubation. In the light phase, nitrogen starvation halts protein synthesis and cell proliferation, leading to the accumulation of carbon-rich compounds such as glycogen in the presence of light and abundant carbon dioxide. These reserves serve as fermentable substrates during the dark phase, where photosynthesis ceases and anaerobic metabolism is activated. Under these conditions, electrons derived from carbohydrate catabolism are redirected to the bidirectional NiFe-hydrogenase (HoxEFUYH), facilitating hydrogen evolution. This two-step strategy is essential for establishing the redox imbalance that drives hydrogenase activity, and it reflects a natural stress response that can be harnessed for biohydrogen production.

Despite attempts to scale up the growth of Synechocystis sp. PCC 6803 to pilot levels (Torzillo et al., 2023), industrial applications remain limited due to high production costs and the risk of contamination by microalgae grazers, which can lead to culture loss (Toulopakis et al., 2016). One potential solution to these challenges is the addition of salt (e.g., sodium chloride) to the growth media or the use of marine water. This approach can reduce the viability of grazing microorganisms, thereby extending biomass utilization and lowering production costs (Bartley et al., 2013), given that Synechocystis sp. PCC 6803 is well-suited to thrive in salty environments (Zhu et al., 2025).

Using seawater for microalgae cultivation can significantly reduce the costs associated with medium preparation. Seawater can, in primis, replace fresh water and add key elements in traditional growth media, such as MgSO₄, CaCl₂, and Na₂CO₃, thereby reducing the need for expensive chemicals. Furthermore, seawater-supplemented media can promote electro-flocculation, simplifying the harvesting process and reducing the production costs (Rafa et al., 2021). This method also mitigates the environmental impact by utilizing a readily available natural resource. By leveraging seawater, the microalgae biofuel industry can overcome some of the economic barriers to large-scale production. This approach aligns with the goals of sustainable energy development and environmental conservation (Wang et al., 2024). Additionally, seawater use in microalgae cultivation can enhance hydrogen production efficiency by providing a stable ionic environment that supports optimal metabolic activities. The presence of naturally occurring salts in seawater can also help maintain osmotic balance, which is crucial for the sustained growth and hydrogen production of microalgae (Bayro-Kaiser and Nelson, 2017). Furthermore, seawater can facilitate the establishment of anaerobic conditions necessary for hydrogen production, particularly under nutrient-deprived conditions (Batyrova et al., 2015).

Synechocystis can adapt to saline environments by accumulating compatible solutes and altering its metabolic profile, including amino acid and glycogen metabolism (Iijima et al., 2015). Despite thorough studies of stress effects caused by salinity on Synechocystis physiology, our knowledge is limited about long-term cultivation in this environment, with special regard to long-term hydrogen production. The ability of Synechocystis to be grown in a saline environment also presents the opportunity to use seawater for cultivation instead of freshwater, which is a limited resource. In a commercial scale-up setting in which cultures are grown continuously in vast quantities of non-sterile water, these properties are potentially valuable (Chaves et al., 2015).

This study aims to investigate the impact of marine salt, dissolved in BG11 growth media at a concentration of 35 g L^-1, on the physiology and hydrogen production of wild-type salt-adapted Synechocystis sp. PCC 6803. The current study is the first and a simplified step in the row towards the final aim to utilize marine water combined with wastewater for long-term microalgae cultivation and hydrogen production on an industrial scale. Although the current study did not experimentally verify the use of wastewater, the integration of marine water with wastewater represents a promising strategy for sustainable hydrogen production. Wastewater is rich in nutrients such as nitrogen and phosphorus, which can support cyanobacterial growth while simultaneously contributing to bioremediation. On the other hand, by understanding the effects of marine salts on the physiology of this cyanobacterium with special attention to hydrogen evolution, we are aiming to establish a novel strategy to lower the environmental impact and improve the reliability and feasibility of outdoor long-term green hydrogen production.

2 Materials and methods

2.1 Organism and culture conditions

The experiments were performed with wild-type Synechocystis sp. PCC 6803 (Stanier et al., 1971). Cells were cultured in both BG11 medium (BG11) and BG11 medium supplemented with marine salt at a concentration of 35 g L^-1 (BG11 ms). Marine salt was obtained from Tropic Marin Ltd. Besides sodium chloride as a major compound, it contains calcium in 380–420 ppm, magnesium in 1210–1310 ppm and further 70 trace elements. Final concentration was reached via 3 steps in glass columns (400 mL working volume, 5 cm diameter) to avoid unnecessary stress. Unlike other studies investigating Synechocystis sp. PCC 6803 in salt salt-supplemented environment for the purpose of describing the physiological responses to sudden salt stress, here we placed great emphasis on avoiding the salt stress and its possible effect on hydrogen evolution to reserve its evolution rate compared to the salt-free media. The Synechocystis culture was grown in BG11 for two weeks, and thereafter it was supplemented with 20 g L^-1 marine salt and grown for 2 weeks again. Finally, the media was changed to BG11 supplemented with marine salt in 35 g L^-1. The cultures were mixed by bubbling a mixture of air and CO₂ (97/3 v/v). For every cultivation, the artificial light intensity was set to 100 μmol photons m^-2 s^-1 and was supplied from one side (Osram, Dulux L, 55W/840). The temperature was maintained at 32°C. For long-term maintenance, stock cultures were refreshed every week and kept in glass tubes.

2.2 Cultivation

For the experiments, Roux bottles were used (1 L working volume, 5.5 cm depth, 12 cm width, 1 main and 4 side inlets on the top of the bottle). Synechocystis sp. PCC 6803 cultures grown in BG11 medium or in a BG11 medium supplemented with marine salt at a concentration of 35 g L^-1 were simultaneously investigated. The media were changed to fresh ones, and the starting volume was set to 1 L in every case. The artificial light intensity was set to 100 μmol photons m^-2 s^-1 and was supplied from one side. The temperature was maintained at 32°C. The culture was mixed simultaneously by a magnetic bar and air bubbling, to guarantee adequate turbulence. Pure CO₂ was injected directly and was regulated by a gas valve based on the value of the measured pH. The upper limit of the pH was set to pH 7.5. Samples were taken for dry weight measurements every day and three times per week for analytical measurements. The samples were quickly washed twice with sterile distilled water to eliminate excess salts and lyophilized for further analysis. The cultures were grown for 7 days, then the whole medium was changed with BG11 supplemented with marine salt but deprived of nitrate, to rapidly induce the nitrogen starvation effect. The biomass was washed with sterile water before the inoculation into nitrogen-deprived media to take away all nitrate residues. The starving culture was kept for 7 days under the same conditions as the first step. For the hydrogen evolution, 60 mL culture was transferred directly from the Roux bottles, keeping the obtained cell density, to serum bottles (120 mL volume) sealed with a rubber stopper and aluminum cap in triplicate. The nitrogen-deprived BG11 medium with or without marine salt has not been modified for dark fermentation; an additional carbon source was not added. Hydrogen evolution relied solely on endogenous carbohydrate reserves. These cultures were incubated in a closed water bath in the dark, mixed by a horizontal shaker at 32°C. Hydrogen evolution was measured every 24 h for 7 days. After every measurement, the serum bottles were purged with nitrogen to avoid the uptake of the accumulated hydrogen. Samples were taken for analytical measurements only after 7 days.

2.3 Photosynthesis measurements

Chlorophyll fluorescence quenching measurements were performed by using a pulse-amplitude-modulation fluorescence instrument (PAM-2100, H. Walz, Effeltrich, Germany) operated by means of PamWin (version 2.00f) PC software. The ratio between variable and maximum fluorescence yields, F_v/F_m, was measured in order to assess the maximum photochemical yield of photosystem II (PSII). For this purpose, culture samples were incubated in the dark for 15 min in order to remove any energy-dependent quenching, and one far-red light (above 700 nm) pulse with a duration of 10 s (5 W m^-2), supplied by the instrument, was applied before every measurement (Schreiber et al., 1994).

2.4 Chlorophyll a fluorescence transients

A Handy-PEA (Hansatech Instruments) was used to record transients of chlorophyll a fluorescence in 2 ml samples during a period of 15 min. The dark-adapted samples were continuously lit by light-emitting diodes (LEDs) with a peak wavelength of 650 nm and a light intensity of 3500 µmol photons m^-2 s^-1. Using BiolyzerHP3 software, each chlorophyll a fluorescence induction curve was examined using the JIP-test (Strasser et al., 1995) (HPEA/LPA2, Hansatech Instruments).

In order to facilitate comparisons between the samples, the chlorophyll fluorescence data were normalized on F₀ and F_m (basal and maximum fluorescence value, respectively). The transient was calculated as relative variable fluorescence V_t = (F_t - F₀)/(F_m - F₀) at all times, with F_t fluorescence at each considered time (Giorio and Sellami, 2021).

The intensity of chlorophyll fluorescence rises from a low (the O level) to a maximum (the P level) in less than one second via intermediate phases designated J (2 ms) and I (30 ms). The rise in O-J correlates with a single turnover drop in quinone A (Q_A). The increase J-I corresponds to the reduction of the secondary quinone acceptor, quinone B (Q_B), the plastoquinone (PQ) pool, and the cytochrome b6f complex, while the I step leads to the partial reoxidation of Q_B, which happens when electrons are transported to the electron acceptor side of Photosystem I. Their decline continues in the next slowest thermal phase (I-P), in less than one second (Giorio and Sellami, 2021).

The fluorescence data were used to determine the following parameters (JIP-test parameters): The initial slope at the beginning of the variable fluorescence, M_o = 4 (F_{300 μs} - F₀)/(F_m - F₀), corresponds to the net rate of the reaction centers closure, which increases by trapping and decreases through electron transport. The variable fluorescence at phase J, V_J = (F_J - F₀)/(F_m - F₀), with F_J the fluorescence value at the J step, indicates the level of Q_A reduction. The following flux ratios and characteristics were computed as specified by Appenroth et al. (2001): F_v/F_m = (F_m - F₀)/F_m, and the highest quantum yield of Photosystem II (PSII) for primary photochemistry.

2.5 Oxygen evolution measurements

Oxygen evolution measurements were carried out in triplicate on 2-mL culture samples, using a Liquid-Phase Oxygen Electrode Chamber (Hansatech, DW3) thermostated at 28°C and equipped with an oxygen control electrode unit (Hansatech, Oxy-lab). Light was supplied via a red LED light source (Hansatech LH36/2R) at a wavelength of 637 nm, providing a 600-μmol photons m^-2 s^-1 PFD. The O₂ concentration dissolved in the sample was continuously monitored at an acquisition rate of 0.2 readings s^-1. Dark respiration rates were measured before the photosynthesis rates had been measured. The obtained values were normalized on the actual chlorophyll content measured on the day of sampling.

2.6 Analytical measurements

2.6.1 Dry weight determination

Determination of dry biomass weight (DW) was performed in triplicate using 2 mL samples and pre-weighted 47 mm diameter glass microfiber filter membranes (Whatman GF/F, Maidstone, England) in an electronic balance (Acculab USA model ATL-224-I) with a precision of ± 0.1 mg. The samples were washed with physiological saline solution before filtration to eliminate the effect of the salt concentration difference between the two cultures. After applying the samples on the filters, the cells on the filters were washed with 50 mL deionized water to get rid of the salt and oven-dried at 105°C. Before the measurement, the filters were transferred to a desiccator to equilibrate to laboratory temperature (Myers and Kratz, 1955). The specific growth rate was calculated as follows:

$$\text{μ}=(\text{ln }{\text{X}}_2–\text{ ln }{\text{X}}_1)/\text{Δt}[{\text{d}}^{-1}]$$

2.6.2 Nutrient analysis

Nitrate concentration was measured with a nitrate reagent (Hanna Instruments, HI93728-01) using the C99 Multiparameter Bench Photometer (Hanna, Lucca, Italy). The collected cultures were centrifuged to separate the dry matter from the media. The obtained supernatant was diluted ten times to fit into the detection range of the reagent. Every sample was measured three times to generate technical triplicates (Gawankar and Masten, 2022).

2.6.3 Analysis of carbohydrate

Total carbohydrate content was assessed by using the phenolsulphuric acid method (Dubois et al., 1956). 1–2 mg lyophilized samples were measured into 50 mL Falcon tubes and suspended in 10 mL of distilled water. For determining the total carbohydrate concentration for a 1 mL sample, 1 mL of phenol and 5 mL of sulfuric acid were added. The final sample was measured at 488 nm using a spectrophotometer (Agilent, Cary 3500 Multicell UV-Vis Spectrophotometer). Every biological sample was measured three times to generate technical triplicates. Results were evaluated by using D+ glucose as a standard (Dubois et al., 1956).

2.6.4 Analysis of protein

Protein measurement was performed in triplicate according to Lowry et al. (1951). 1–2 mg lyophilized samples were measured into 50 mL Falcon tubes and suspended in 10 mL of distilled water. For determination 1N NaOH, 5% Na₂CO₃, 0.5% CuSO₄ × 5H₂O, 1% K-Na tartrate and 1N Folin Ciocalteau reagent were used. Four solutions were prepared before the analysis:

● Solution A: 5% sodium carbonate in H₂O.

● Solution B: 0.5% copper sulphate (CuSO₄ × 5H₂O) in 1% potassium sodium tartarate.

● Solution C: Mix 50mL of reagent A and 1 mL of reagent B prior to use.

● Solution D: Dilute Folin-Ciocalteau reagent with an equal volume of 1 N acid.

0.5 mL sample + 0.5 mL NaOH 1N was incubated in boiling water for 5 min. After cooling to room temperature, 2.5 mL of solution C was added. The sample rested for 10 min, then 0.5 mL of solution D was added. Samples were read after 30 min of incubation at 750 nm using a spectrophotometer (Agilent, Cary 3500 Multicell UV-Vis Spectrophotometer). Results were evaluated by using a solution of bovine albumin as a standard (10–400 mg L^-1) (Lowry et al., 1951).

2.6.5 Analysis of polyhydroxybutyrate

PHB was determined in the form of crotonic acid by HPLC using the following procedure. 5 mL of the culture were centrifuged in a Sorvall Super T21 centrifuge at 5000 g for 10 min, and the pellet was used for acid digestion with 1 mL of pure sulfuric acid at 105°C to convert PHB to crotonic acid. Crotonic acid was eluted from a Synergy-Hydro-RP C-18 column (205 × 4.6 mm i.d.) and measured by ultraviolet detection at 214 nm. The mobile phase was 15% (v/v) acetonitrile in 0.1% (v/v) H₃PO₄ in aqueous solution at a flow rate of 1 mL min⁻¹. Every analysis was performed in triplicate (Law and Slepecky, 1961).

2.6.6 Analysis of pigments

Chlorophyll a and total carotenoid concentrations were determined by using 1.5 mL samples centrifuged for 8 min at 2650 ×g in Eppendorf tubes using an ALC-PK110 centrifuge. After discarding the supernatant and re-suspending the pellet in 1.5 mL of pure methanol, the tubes were kept for 3 min in a 70°C water bath, then centrifuged again for 8 min at 2650 g. The absorbance of the supernatant was measured at 470, 665 and 750 nm against pure methanol blind (Ritchie, 2006; Wellburn, 1994).

$$\text{Chlorophyll a}:{\text{C}}_{\text{a}}=12.9447\times ({\text{A}}_{665}–{\text{A}}_{750})$$

$$\text{Carotenoids}:{\text{C}}_{\text{car}}=[1,000({\text{A}}_{470}-{\text{A}}_{750})-2.86(\text{Ch}{\text{l}}_{\text{a}}[\mu \text{g m}{\text{L}}^{-1}])]/221$$

2.6.7 Hydrogen measurement

7At the beginning of the hydrogen production phase, the vials were sealed with rubber septum stoppers and aluminum rings and then purged with nitrogen gas for 5 min to remove oxygen. Purging was also repeated after every measurement, every day, to avoid the uptake of the accumulated hydrogen by the bidirectional hydrogenase. Hydrogen accumulation was calculated from the summary of the daily hydrogen evolutions. The cultures were incubated in darkness to trigger hydrogen production. Samples from the headspace were collected and analyzed with a gas chromatograph (model Clarus 500, Perkin Elmer, Waltham, Massachusetts), using a packed column (model Carbosieve SII Spherical Carbon, Supelco) and a thermal conductivity detector. The temperature was set to 35°C and the running time was 2 min. A hydrogen standard curve was used to calculate the volume of hydrogen from the reported peak areas (Kosourov et al., 2003).

2.7 Statistical analysis

Each measurement was conducted using three biological and three technical replicates. Error bars represent the standard deviation of the measured values. For the significance analysis of JIP-test values in Table 1, a Two-way ANOVA test with Tukey’s post-hoc test was used. For each parameter, different letters indicate significant differences among the different periods. For the pairwise comparison of the related columns of hydrogen values in Figure 1, a Two-way ANOVA test was used. Means of the rows were compared, and the Šidák test was used for statistical hypothesis testing.

Table 1

Table 1. Changes of JIP-test parameters.

Figure 1

Figure 1. Changes in the biomass density in the Synechocystis sp. PCC 6803 cultures were grown in Roux bottles (growth and nitrogen starvation phase) and in serum vials (H₂ evolution phase). Graph (a) shows the volumetric biomass density of the culture cultivated in BG11 media (empty circle, continuous line) and in BG11 media supplemented with marine salt (filled square, continuous line). Graph (b) shows the sodium nitrate concentration changes in BG11 media (empty circle, continuous line) and in BG11 media supplemented with marine salt (filled square, continuous line). On day 7^th, both cultures were shifted to nitrate-deprived conditions by washing the cells with nitrate-deprived media. Each measurement was conducted using three biological and three technical replicates. Error bars represent the standard deviation of the measured values.

3 Results

3.1 Culture growth

Synechocystis sp. PCC 6803 cultures were cultivated in BG11 medium (BG11) and in a BG11 medium supplemented with marine salt at a concentration of 35 g L^-1 (BG11 ms). The experiment consisted of three distinct steps: growth, nitrogen starvation and hydrogen evolution. The sodium nitrate concentration in the media differed as a result of the reaction between the nitrate salt and the marine salt in the media, which caused a partial nitrate precipitation. Thus, their concentration started from 1.49 g L^-1 and 1.05 g L^-1 (Figure 1). Despite the different nitrate concentration, the actual nitrate content in BG11 medium supplemented with marine salt was sufficient to support the growth of Synechocystis culture, since nitrate concentration had not reached the level that suppresses growth rate. Both cultures were growing continuously during the growth phase, utilizing the nitrate sources, which finished at 0.39 g L^-1 and 0.43 g L^-1. For the following nitrogen starvation phase, both media were shifted to nitrate-free media; therefore, the measured sodium nitrate concentration was 0 g L^-1 for the entire starvation phase. The initial concentrations of the dry weight were 0.33 ± 0.08 g L^-1 and 0.39 ± 0.12 g L^-1 in both the cultures cultivated in BG11 and in BG11 supplemented with marine salt (Figure 1). In the growth phase, the dry weight reached 1.17 ± 0.17 g L^-1 and 1.11 ± 0.32 g L^-1. Their volumetric productivities were 0.119 ± 0.018 g L^-1 d ^-1 (μ = 0.18 ± 0.02 d^-1) and 0.102 ± 0.039 g L^-1 d ^-1 (μ = 0.15 ± 0.02 d^-1), respectively. In the nitrogen starvation phase, the dry weights reached 2.74 ± 0.27 g L^-1 and 2.75 ± 0.14 g L^-1,and their volumetric productivities were 0.225 ± 0.012 g L^-1 d^-1 (μ = 0.12 ± 0.01 d^-1) and 0.234 ± 0.059 g L^-1 d^-1 (μ = 0.14 ± 0.05 d^-1) (Figure 1). In the third phase, cultures were directly transferred from light to dark conditions to initiate hydrogen evolution via dark fermentation. At the end of this phase, the dry weight decreased to 2.12 ± 0.08 g L^-1 and 1.92 ± 0.26 g L^-1, and their volumetric productivities were -0.099 ± 0.023 g L^-1 d^-1 (μ = -0.04 ± 0.01 d^-1) and -0.143 ± 0.054 g L^-1 d^–1 (μ = -0.07 ± 0.03 d^-1).

3.2 Photosynthetic performance

Photosynthetic performance and electron transport dynamics were assessed using chlorophyll fluorescence quenching measurements, JIP-test parameters and gas exchange measurements. As shown in Table 1, the initial slope of fluorescence rise (M_o) increased during the starvation phase in both BG11 and BG11 ms cultures, indicating that the reduced rate of the primary quinone receptor Q_A increased, and that electron transfer to further Q_A was affected. This was accompanied by a moderate increase in the variable fluorescence at the J-step (V_J). As shown in Figure 2a, the maximum quantum yield of PSII (F_v/F_m) remained relatively stable; it increased slightly from 0.437 to 0.520 (BG11) and from 0.494 to 0.564 (BG11 ms).

Figure 2

Figure 2. Changes of PSII quantum yield, oxygen evolution and respiration in Synechocystis sp. PCC 6803 cultures during the growth and nitrogen starvation periods in Roux bottles (growth and nitrogen starvation phase): (a) changes of F_v/F_m values in in BG11 media (black column) and in BG11 media supplemented with marine salt (grey column), (b) oxygen evolution in BG11 media (black column) and in BG11 media supplemented with marine salt (white column) and respiration in BG11 media (dark grey column) and in BG11 media supplemented with marine salt (light grey column). Each measurement was conducted using three biological and three technical replicates. Error bars represent the standard deviation of the measured values.

Following nitrogen starvation, M_o values dropped sharply—by approximately 90%—in both media, reflecting a substantial decline in PSII activity. Simultaneously, V_J values increased more prominently in BG11 cultures (53%) than in BG11 ms (15%), suggesting a greater accumulation of reduced Q_A in the absence of marine salt. F_v/F_m values also declined to 0.218 in BG11 cultures compared to 0.392 in BG11 ms ones.

Complementary measurements of oxygen evolution and respiration (Figure 2b) aligned with these findings. During the growth phase, oxygen evolution was higher in BG11 cultures, while BG11 ms cultures exhibited slightly elevated respiration rates. Upon nitrogen starvation, both oxygen evolution and respiration declined, with a more pronounced drop in oxygen evolution in BG11 ms cultures. This shift suggests a reduction of electron flow due to the increasing stress and the damage to PSII.

3.3 Carbohydrate, protein and PHB

In each experimental phase, biomass was collected for the analysis of carbohydrate, protein and polyhydroxybutyrate contents (Figure 3). On the first day of the growth phase, the carbohydrate content was 0.2 ± 0.04 g g^-1 in the BG11 culture and 0.17 ± 0.01 g g^-1 in the BG11 culture supplemented with marine salt. Protein content was 0.56 ± 0.04 g g^-1 and 0.53 ± 0.04 g g^-1 in the same cultures. After one week of growth in full media, the carbohydrate content decreased to 0.12 ± 0.01 g g^-1 and 0.11 ± 0.01 g g^-1, respectively. Simultaneously, the protein content slightly increased to 0.56 ± 0.04 g g^-1 and 0.57 ± 0.04 g g^-1. In the next phase, both media were replaced by sodium nitrate-deprived media to force the physiological acclimation to the nitrogen-starved conditions. As a result, the carbohydrate content increased to 0.61 ± 0.02 g g^-1 and 0.59 ± 0.01 g g^-1. On the contrary, protein content fell to 0.23 ± 0.04 g g^-1 in both cultures. Polyhydroxybutyrate was not detectable during the growth phase, but at the end of the nitrogen starvation, it accumulated to 0.03 ± 0.01 g g^-1 and 0.02 ± 0.01 g g^-1. For the induction of hydrogen evolution, the cultures remained in the dark in the same medium for another week. During the dark fermentation, carbohydrate levels decreased to 0.59 ± 0.03 g g^-1 and 0.55 ± 0.07 g g^-1, meanwhile protein levels increased to 0.24 ± 0.05 g g^-1 in both cultures. Polyhydroxybutyrate increased to 0.1 ± 0.01 g g^-1 and 0.04 ± 0.01 g g^-1 till the last day of the cultivation.

Figure 3

Figure 3. Changes of carbohydrate, protein and PHB contents in Synechocystis sp. PCC 6803 cells were grown in Roux bottles under continuous light (growth and nitrogen starvation phase) and thereafter transferred to serum vials and incubated in the dark (H₂ evolution phase). The results of the cultivation in BG11 media are presented with an empty triangle and continuous line, the results of the cultivation in BG11 media supplemented with marine salt are depicted with a filled triangle and continuous line: (a) carbohydrate content; (b) protein content, (c) PHB content. Each measurement was conducted using three biological and three technical replicates. Error bars represent the standard deviation of the measured values.

3.4 Pigments

For pigment analysis, the concentration of chlorophyll a and carotenoids was monitored during all three phases of the experiment. The highest chlorophyll a and carotenoid content was obtained during the growth phase, 21.48 ± 3.51 mg g^-1/19.04 ± 1.72 mg g^-1 and 5.55 ± 0.33 mg g^-1/3.69 ± 0.59 mg g^-1 in BG11 and BG11 ms cultures (Figure 4). Chlorophyll a was very similar in both cultures. In contrast, carotenoids had significantly higher levels in BG11 cultures compared to BG11 ms cultures. These differences remained till the end of the cultivation. In the nitrogen deprivation phase, both the chlorophyll a and carotenoid concentrations decreased to 7.18 ± 0.76 mg g^-1/6.86 ± 0.69 mg g^-1 and 3.38 ± 0.44 mg g^-1/2.15 ± 0.42 mg g^-1 in BG11 and BG11 ms cultures. With the exception of the carotenoid content in BG11 cultures (4.05 ± 0.06 mg g^-1), the chlorophyll a and carotenoid contents further decreased to 6.29 ± 0.13 mg g^-1/6.56 ± 0.29 mg g^-1 and 1.41 ± 0.56 mg g^-1 till the end of the experiment.

Figure 4

Figure 4. Chlorophyll a and carotenoid contents in the Synechocystis sp. PCC 6803 cultures grown in Roux bottles (growth and nitrogen starvation phase) and serum vials (H₂ evolution phase): (a) chlorophyll a and (b) carotenoid contents changes in BG11 media (empty circle) and in BG11 media supplemented with marine salt (filled square). Each measurement was conducted using three biological and three technical replicates. Error bars represent the standard deviation of the measured values.

3.5 Hydrogen evolution

Hydrogen evolution measurements were carried out after the growth and starvation phase in 120 mL sealed serum vials and incubated in the dark. Hydrogen yields in the headspace were measured every day, then the accumulated hydrogen was discarded by purging the headspace of the vials to minimize the hydrogen uptake nature of the bidirectional NiFe-hydrogenase of Synechocystis and to obtain the actually produced hydrogen yields. The obtained gas yields were normalized on the dry weight of the 14^th day of cultivation for the purpose of a better comparison. The highest accumulated yields were recorded on the first day, 4.17 ± 1.53 mL g^-1 in BG11 culture and 4.26 ± 0.58 mL g^-1 in BG11 with marine salt culture (Figure 5a). Later, the daily yields stepwise decreased to 0.081 ± 0.013 mL g^-1 in BG11 and 0.332 ± 0.014 mL g^-1 in BG11 with marine salt culture on the fourth day. Hydrogen accumulation values were obtained by summing the daily yields to provide a comprehensive picture of the total hydrogen production (Figure 5b). The total hydrogen evolution yields at the end of the four-day-long monitoring period were 8.5 ± 1.81 mL g^-1 in BG11 culture and 9.14 ± 0.62 mL g^-1 in BG11 with marine salt culture.

Figure 5

Figure 5. Hydrogen evolution of Synechocystis sp. PCC 6803 during the first four days of the hydrogen production phase (a) daily hydrogen yields in the headspace of the culture cultivated in BG11 media (black column) and in BG11 media supplemented with marine salt (grey column) normalized on dry matter; (b) accumulated hydrogen yields in the headspace of the culture cultivated in BG11 media (black column) and in BG11 media supplemented with marine salt (grey column) normalized on dry matter. Each measurement was conducted using three biological and three technical replicates. Error bars represent the standard deviation of the measured values.

4 Discussion

This study explored the influence of marine salt (35 g L⁻¹) on the growth, physiological responses, and hydrogen production of Synechocystis sp. PCC 6803. The findings reveal that despite marine salt slightly reducing growth rates, it doesn’t significantly affect the hydrogen production under dark fermentative conditions.

The growth performance of Synechocystis in BG11 and BG11 supplemented with marine salt (BG11 ms) was comparable during the initial growth phase, with only a modest reduction in volumetric productivity in the salt-treated cultures. This is consistent with the known adaptation studies of Synechocystis sp. PCC 6803, which applied stepwise laboratory evolution to elevated salinity (Wang et al., 2014; Zhu et al., 2025). The salt-evolved Synechocystis strains exhibited full adaptation to high-salt environments, maintaining morphology and photosynthetic characteristics comparable to the wild-type strain while undergoing significant alterations in central metabolism under salt stress. It was demonstrated that salt exposure induces metabolic reprogramming, particularly in amino acid and fatty acid biosynthesis, supporting cellular adaptation. The continued biomass accumulation during nitrogen starvation in both conditions suggests that Synechocystis can redirect carbon flux toward storage compounds such as glycogen and polyhydroxybutyrate (PHB), even in the absence of nitrogen. This metabolic flexibility is a hallmark of cyanobacterial stress responses and has been exploited in biotechnological applications for biofuel and bioplastic production (Singhon et al., 2021).

Photosynthetic performance, assessed via JIP-test parameters, showed a marked decline in PSII efficiency (F_v/F_m) during both nitrogen starvation and following the dark fermentation, with slightly better retention in BG11 ms cultures. F_v/F_m values declined to 0.218 in BG11 cultures and to a lesser extent (0.392) in BG11 ms cultures. These findings align with complementary gas exchange data: oxygen evolution was higher in BG11 cultures during growth, while BG11 ms cultures exhibited slightly elevated respiration. Similar trends were reported by Touloupakis et al. (2016), who found that Synechocystis maintained higher photosynthetic activity under modified media conditions (Touloupakis et al., 2016). The observed reduction in M_o and increase in V_J values during hydrogen production indicate a shift in electron transport dynamics, likely due to the redirection of reducing equivalents toward hydrogenase activity. Burgstaller et al. (2022) described the role of the bidirectional NiFe-hydrogenase in balancing redox poise under fermentative conditions, a mechanism that appears to be enhanced in salt-treated cultures (Burgstaller et al., 2022).

Carbohydrate content increased significantly during nitrogen starvation, reaching similar levels in both BG11 and BG11 ms cultures. This is consistent with the known response of cyanobacteriophyta to nitrogen limitation, where carbon is stored as glycogen (Singhon et al., 2021). The slight reduction in carbohydrate content during dark fermentation suggests its partial utilization as a substrate for hydrogen production (Navarro et al., 2009). Protein content decreased during nitrogen starvation, reflecting halted protein synthesis and degradation of existing proteins to recycle nitrogen (Elmorjani and Herdman, 1987; Huang et al., 2013). PHB accumulation was more pronounced in BG11 ms cultures, particularly during dark fermentation. This suggests that marine salt may enhance carbon storage under stress, potentially by modulating the activity of key enzymes in the PHB biosynthetic pathway (Meixner et al., 2022).

Chlorophyll a and carotenoid contents declined during nitrogen starvation and dark fermentation, reflecting the downregulation of photosynthetic activity. However, carotenoid levels were consistently lower in BG11 ms cultures, possibly due to altered pigment biosynthesis under saline conditions. In other studies, the highest carotenoid contents were found at 400 mM (23.38 g L^-1) and 342 mM (20 g L^-1) NaCl concentrations in Synechocystis sp. CCNM 2501 and in Synechocystis sp. PCC 6803, respectively (Schubert et al., 1993; Paliwal et al., 2015). From these concentrations, the carotenoid levels generally declined as the salt concentration stepwise increased.

In this study, the hydrogen production in marine salt-enriched BG11 cultures was moderately enhanced. Although the total hydrogen yield was not dramatically higher, cultures grown in BG11 supplemented with marine salt consistently produced slightly more hydrogen than the control. This suggests that marine salt may support fermentative hydrogen production by stabilizing metabolic activity under stress. Several former studies may support this improvement. Salt stress may increase the availability of reducing equivalents by promoting carbohydrate catabolism, as suggested by the higher initial carbohydrate content and PHB accumulation (Meixner et al., 2022). Similar findings have been described with Synechococcus elongatus PCC7942, PAMCOD and Synechocystis sp. PCC6714, where salt-induced sucrose accumulation enhanced hydrogen production under anaerobic dark fermentation (Broussos et al., 2024a, 2024b). Additionally, marine salt may facilitate the establishment of anaerobic conditions by reducing oxygen solubility, thereby favoring hydrogenase activity (Oren, 2013). While the enhancement observed in this study was modest, it supports the feasibility of using saline media to optimize hydrogen yields in cyanobacterial systems.

A mass balance between carbohydrate consumption and H₂ production demonstrates that during the fermentation phase, there was in both cultures a slight decrease in the amount of carbohydrates of 0.03 g g^-1 DW, which were likely used for hydrogen production. Fermentation of carbohydrates and hydrogen and acetic acid production occur according to the following reaction (Das and Veziroǧlu, 2001);

$${\text{C}}_6{\text{H}}_{12}{\text{O}}_6+2{\text{H}}_2\text{O}\to 2{\text{C}}_2{\text{H}}_4{\text{O}}_2+2\text{C}{\text{O}}_2+4{\text{H}}_2\text{ΔG}{0}^{\prime }=-206\text{ kJ mol}$$

Hence, with 30 mg g^-1 of carbohydrates, consumed during the fermentation phase (Figure 4), a total theoretical amount of 1.33 mg of H₂ g^-1 DW should be produced, corresponding to 14.8 mL of H₂ (assuming 89.8 mg mL^-1 the density of H₂). The amount of hydrogen actually produced was 8.5 mL g^-1 DW (BG11) and 9.14 mL g^-1 DW (BG11 ms), which were 57% and 62% of the theoretical yield, respectively. However, these yields are significantly higher than those reported by Touloupakis et al. (2016). Nevertheless, the amount of carbohydrates involved in dark fermentation represented only a minimal part of that synthesized during the nitrogen starvation (Figure 4), indicating that only some classes of carbohydrates are involved in the production of hydrogen by dark fermentation (Broussos et al., 2024a). Notwithstanding the scarce utilization of carbohydrates during the dark fermentation phase, and the stable amount of protein, the cell dry weight strongly decreased in both the cultures (by -0.62 g L^-1 in BG11) and (by -0.83 g L^-1 in BG11 ms). This significant reduction could be only partially explained by the synthesis of PHB, characterized by a higher energy content (24–30 KJ g^-1) than that of the biomass (close to 20 KJ g^-1) and by an increased lipid fraction of the biomass (biochemical component not assessed). Indeed, lipid accumulation can be induced by stress conditions (Neag et al., 2019). Therefore, a relevant cell lysis during the too-long fermentation phase cannot be ruled out. Our results clearly indicated that more than 90% of total hydrogen produced is attained within 3 days (Figure 5); therefore, the length of the fermentation phase can be reduced by more than 50% of the total time, thus reducing the length of stress due to long anaerobiosis conditions. Therefore, stopping the fermentation phase on day 3, the drop of dry weight could have been much less relevant (Figure 1).

It is worth pointing out that the use of marine salt or seawater in microalga cultivation offers several advantages: it reduces freshwater demand, lowers contamination risk, and enhances hydrogen production. This aligns with the goals of sustainable energy development and supports the feasibility of large-scale hydrogen production using brackish or marine water sources. Moreover, the compatibility of Synechocystis with saline environments makes it a promising candidate for outdoor cultivation in coastal regions. The findings of this study provide a strong rationale for further optimization of saline cultivation systems, including the use of natural seawater and the development of salt-tolerant strains through genetic engineering.

Data availability statement

The original contributions presented in the study are included in the article/supplementary material. Further inquiries can be directed to the corresponding author.

Author contributions

GL: Writing – original draft, Investigation, Formal Analysis, Conceptualization, Methodology, Writing – review & editing. BC: Writing – original draft, Investigation, Visualization, Methodology, Data curation. FB: Writing – review & editing, Investigation, Methodology. AP: Writing – review & editing, Methodology. ET: Investigation, Writing – review & editing, Methodology. GC: Writing – review & editing. GT: Writing – review & editing, Writing – original draft. CF: Writing – review & editing, Writing – original draft, Methodology, Supervision, Investigation.

Funding

The author(s) declared that financial support was received for this work and/or its publication. This research was supported by the European Innovation Council (EIC) under the Horizon Europe program through the EIC Pathfinder grant (No. 101070948).

Conflict of interest

The author(s) declared that this work was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Generative AI statement

The author(s) declared that generative AI was not used in the creation of this manuscript.

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References

Appenroth K. J., Stöckel J., Srivastava A., and Strasser R. J. (2001). Multiple effects of chromate on the photosynthetic apparatus of Spirodela polyrhiza as probed by OJIP chlorophyll a fluorescence measurements. Environ. pollut. 115, 49–64. doi: 10.1016/S0269-7491(01)00116-5

PubMed Abstract | Crossref Full Text | Google Scholar

Bartley M. L., Boeing W. J., Corcoran A. A., Holguin F. O., and Schaub T. (2013). Effects of salinity on growth and lipid accumulation of biofuel microalga Nannochloropsis salina and invading organisms. Biomass Bioenergy 54, 83–88. doi: 10.1016/j.biombioe.2013.03.015

Crossref Full Text | Google Scholar

Batyrova K., Gavrisheva A., Ivanova E., Liu J., and Tsygankov A. (2015). Sustainable hydrogen photoproduction by phosphorus-deprived marine green microalgae Chlorella sp. Int. J. Mol. Sci. 16, 2705–2716. doi: 10.3390/ijms16022705

PubMed Abstract | Crossref Full Text | Google Scholar

Bayro-Kaiser V. and Nelson N. (2017). Microalgal hydrogen production: prospects of an essential technology for a clean and sustainable energy economy. Photosynth. Res. 133, 49–62. doi: 10.1007/s11120-017-0350-6

PubMed Abstract | Crossref Full Text | Google Scholar

Broussos P. I., Romanos G. E., and Stamatakis K. (2024a). Salt and heat stress enhance hydrogen production in cyanobacteria. Photosynth. Res. 161, 117–125. doi: 10.1007/s11120-024-01071-2

PubMed Abstract | Crossref Full Text | Google Scholar

Broussos P. I., Romanos G. E., and Stamatakis K. (2024b). H₂ production by the unicellular freshwater cyanobacterium Synechococcus elongatus PCC 7942 PAMCOD strain. Int. J. Hydrogen Energy 52, 1298–1303. doi: 10.1016/j.ijhydene.2023.10.123

Crossref Full Text | Google Scholar

Burgstaller H., Wang Y., Caliebe J., Hueren V., Appel J., Boehm M., et al. (2022). Synechocystis sp. PCC 6803 requires the bidirectional hydrogenase to metabolize glucose and arginine under oxic conditions. Front. Microbiol. 13. doi: 10.3389/fmicb.2022.896190

PubMed Abstract | Crossref Full Text | Google Scholar

Chaves J. E., Kirst H., and Melis A. (2015). Isoprene production in Synechocystis under alkaline and saline growth conditions. J. Appl. Phycol. 27, 1089–1097. doi: 10.1007/s10811-014-0395-2

Crossref Full Text | Google Scholar

Das D. and Veziroǧlu T. N. (2001). Hydrogen production by biological processes: a survey of literature. Int. J. Hydrogen Energy 26, 13–28. doi: 10.1016/S0360-3199(00)00058-2

Crossref Full Text | Google Scholar

Dubois M., Gilles K. A., Hamilton J. K., Rebers P. A., and Smith F. (1956). Colourimetric method for the determination of sugars and related substances. Anal. Chem. 28, 350–356. doi: 10.1021/ac60111a017

Crossref Full Text | Google Scholar

Dzulkarnain E. L. N., Audu J. O., Wan Dagang W. R. Z., and Abdul-Wahab M. F. (2022). Microbiomes of biohydrogen production from dark fermentation of industrial wastes: current trends, advanced tools and future outlook. Bioresour. Bioprocess. 9, 16. doi: 10.1186/s40643-022-00501-x

PubMed Abstract | Crossref Full Text | Google Scholar

Elmorjani K. and Herdman M. (1987). Metabolic control of phycocyanin degradation in the cyanobacterium Synechocystis PCC 6803: a glucose effect. J. Gen. Microbiol. 133, 1685–1694. doi: 10.1099/00221287-133-7-1685

Crossref Full Text | Google Scholar

Gawankar S. and Masten S. J. (2022). Development of an inexpensive, rapid method to measure nitrates in freshwater to enhance student learning. J. Chem. Educ. 99, 1234–1240. doi: 10.1021/acs.jchemed.2c00381

Crossref Full Text | Google Scholar

Giorio P. and Sellami M. H. (2021). Polyphasic OKJIP Chlorophyll a Fluorescence Transient in a Landrace and a Commercial Cultivar of Sweet Pepper (Capsicum annuum, L.) under Long-Term Salt Stress. Plants 10, 887–901. doi: 10.3390/plants10050887

PubMed Abstract | Crossref Full Text | Google Scholar

Huang S., Chen L., Te R., Qiao J., Wang J., and Zhang W. (2013). Complementary iTRAQ proteomics and RNA-seq transcriptomics reveal multiple levels of regulation in response to nitrogen starvation in Synechocystis sp. PCC 6803. Mol. Biosyst. 9, 2565–2574. doi: 10.1039/c3mb70109c

PubMed Abstract | Crossref Full Text | Google Scholar

Iijima H., Nakaya Y., Kuwahara A., Hirai M. Y., and Osanai T. (2015). Seawater cultivation of freshwater cyanobacterium Synechocystis sp. PCC 6803 drastically alters amino acid composition and glycogen metabolism. Front. Microbiol. 6. doi: 10.3389/fmicb.2015.00326

PubMed Abstract | Crossref Full Text | Google Scholar

Karmaker S. C., Chapman A., Sen K. K., Hosan S., and Saha B. B. (2023). Renewable Energy Pathways toward Accelerating Hydrogen Fuel Production: Evidence from Global Hydrogen Modeling. Sustainability 15 (1), 588. doi: 10.3390/su15010588

Crossref Full Text | Google Scholar

Kosourov S., Seibert M., and Ghirardi M. L. (2003). Effects of extracellular pH on the metabolic pathways in sulfur-deprived, H₂-producing Chlamydomonas reinhardtii cultures. Plant Cell Physiol. 44, 146–155. doi: 10.1093/pcp/pcg013

PubMed Abstract | Crossref Full Text | Google Scholar

Law J. H. and Slepecky R. A. (1961). Assay of poly-β-hydroxybutyric acid. J. Bacteriol. 82, 33–36. doi: 10.1128/jb.82.1.33-36.1961

PubMed Abstract | Crossref Full Text | Google Scholar

Lee H., Calvin K., Dasgupta D., Krinner G., Mukherji A., Thorne P., et al. (2023). IPCC 2023: Climate Change 2023: Synthesis Report, Summary for Policymakers. Contribution of Working Groups I, II and III to the Sixth Assessment Report of the Intergovernmental Panel on Climate Change. Eds. Lee H. and Romero J. (Geneva, Switzerland: IPCC).

Google Scholar

Lowry O. H., Rosebrough N. J., Farr A. L., and Randall R. J. (1951). Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265–275. doi: 10.1016/S0021-9258(19)52451-6

Crossref Full Text | Google Scholar

Meixner K., Daffert C., Dalnodar D., Mrázová K., Hrubanová K. V., Krzyzanek V., et al. (2022). Glycogen, poly(3-hydroxybutyrate) and pigment accumulation in three Synechocystis strains when exposed to a stepwise increasing salt stress. J. Appl. Phycol. 34, 1227–1241. doi: 10.1007/s10811-022-02693-3

PubMed Abstract | Crossref Full Text | Google Scholar

Myers J. and Kratz W. A. (1955). Relations between pigment content and photosynthetic characteristics in a blue-green alga. J. Gen. Physiol. 39, 11–22. doi: 10.1085/jgp.39.1.11

PubMed Abstract | Crossref Full Text | Google Scholar

Navarro E., Montagud A., de Córdoba P. F., and Urchueguía J. F. (2009). Metabolic flux analysis of the hydrogen production potential in Synechocystis sp. PCC6803. Int. J. Hydrogen Energy 34, 8828–8838. doi: 10.1016/j.ijhydene.2009.08.018

Crossref Full Text | Google Scholar

Neag E., Torök A. I., Cadar O., Băbălău V., and Roman F. C. (2019). Enhancing lipid production of Synechocystis PCC 6803 for biofuels production, through environmental stress exposure. Renew. Energy 143, 243–251. doi: 10.1016/j.renene.2019.04.148

Crossref Full Text | Google Scholar

Oren A. (2013). “Life at high salt concentrations,” in The Prokaryotes. Eds. Rosenberg E., DeLong E. F., Lory S., Stackebrandt E., and Thompson F. (Springer, Berlin, Heidelberg). doi: 10.1007/978-3-642-30123-0_57

Crossref Full Text | Google Scholar

Paliwal C., Pancha I., Ghosh T., Maurya R., Chokshi K., Bharadwaj S. V., et al. (2015). Selective carotenoid accumulation by varying nutrient media and salinity in Synechocystis sp. CCNM 2501. Bioresour. Technol. 197, 363–368. doi: 10.1016/j.biortech.2015.08.110

PubMed Abstract | Crossref Full Text | Google Scholar

Rafa N., Ahmed S. F., Badruddin I. A., Mofijur R. M., and Kamangar S. (2021). Strategies to produce cost-effective third-generation biofuel from microalgae. Front. Energy Res. 9. doi: 10.3389/fenrg.2021.749968

Crossref Full Text | Google Scholar

Ritchie R. J. (2006). Consistent sets of spectrophotometric chlorophyll equations for acetone, methanol and ethanol solvents. Photosynth. Res. 89, 27–41. doi: 10.1007/s11120-006-9065-9

PubMed Abstract | Crossref Full Text | Google Scholar

Schreiber U., Bilger W., and Neubauer C. (1994). Chlorophyll fluorescence as a nonintrusive indicator for rapid assessment of in vivo photosynthesis. Ecol. Stud. 100, 49–70. doi: 10.1007/978-3-642-78910-7_5

Crossref Full Text | Google Scholar

Schubert H., Fulda S., and Hagemann M. (1993). Effects of adaptation to different salt concentrations on photosynthesis and pigmentation of the cyanobacterium Synechocystis sp. PCC 6803. J. Plant Physiol. 142, 291–295. doi: 10.1016/S0176-1617(11)81147-1

Crossref Full Text | Google Scholar

Singhon P., Phoraksa O., Incharoensakdi A., and Monshupanee T. (2021). Increased bioproduction of glycogen, lipids, and poly(3-hydroxybutyrate) under partial supply of nitrogen and phosphorus by photoautotrophic cyanobacterium Synechocystis sp. PCC 6803. J. Appl. Phycol. 33, 2833–2843. doi: 10.1007/s10811-021-02494-0

Crossref Full Text | Google Scholar

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. doi: 10.1128/br.35.2.171-205.1971

PubMed Abstract | Crossref Full Text | Google Scholar

Strasser R., Srivastava A., and Govindjee (1995). Polyphasic chlorophyll a fluorescence transient in plants and cyanobacteria. Photochem. Photobiol. 61, 32–42. doi: 10.1111/j.1751-1097.1995.tb09239.x

PubMed Abstract | Crossref Full Text | Google Scholar

Taikhao S., Incharoensakdi A., and Phunpruch S. (2015). Dark fermentative hydrogen production by the unicellular halotolerant cyanobacterium Aphanothece halophytica grown in seawater. J. Appl. Phycol. 27, 187–196. doi: 10.1007/s10811-014-0292-8

Crossref Full Text | Google Scholar

Torzillo G., Zittelli G. C., Cicchi B., Diano M., Parente M., Silva Benavides A. M., et al. (2023). Effect of plate distance on light conversion efficiency of a Synechocystis culture grown outdoors in a multiplate photobioreactor. Sci. Total Environ. 842, 156840. doi: 10.1016/j.scitotenv.2022.156840

PubMed Abstract | Crossref Full Text | Google Scholar

Touloupakis E., Cicchi B., Benavides A. M. S., and Torzillo G. (2016). Effect of high pH on growth of Synechocystis sp. PCC 6803 cultures and their contamination by golden algae (Poterioochromonas sp.). Appl. Microbiol. Biotechnol. 100, 1333–1341. doi: 10.1007/s00253-015-7094-3

PubMed Abstract | Crossref Full Text | Google Scholar

Wang M., Ye X., Bi H., and Shen Z. (2024). Microalgae biofuels: illuminating the path to a sustainable future amidst challenges and opportunities. Front. Energy Res. 17, 10. doi: 10.1186/s13068-024-02461-0

PubMed Abstract | Crossref Full Text | Google Scholar

Wang J., Zhang X., Shi M., Gao L., Niu X., Te R., et al. (2014). Metabolomic analysis of the salt-sensitive mutants reveals changes in amino acid and fatty acid composition important to long-term salt stress in Synechocystis sp. PCC 6803. Funct. Integr. Genomics 14, 431–440. doi: 10.1007/s10142-014-0370-7

PubMed Abstract | Crossref Full Text | Google Scholar

Wellburn A. R. (1994). The spectral determination of chlorophylls a and b, as well as total carotenoids, using various solvents with spectrophotometers of different resolution. J. Plant Physiol. 144, 307–313. doi: 10.1016/S0176-1617(11)81192-2

Crossref Full Text | Google Scholar

Zhu X., Zou R., Liu D., Liu J., Wu X., Jiang J., et al. (2025). Enhanced salt tolerance in Synechocystis sp. PCC 6803 through adaptive evolution: Mechanisms and applications for environmental bioremediation. Microbiol. Res. 296, 128140. doi: 10.1016/j.micres.2025.128140

PubMed Abstract | Crossref Full Text | Google Scholar