Trends Sci. 2025; 22(10): 10645

Fossil fuels, the primary driver of global economic growth, have caused significant environmental damage and are projected to be depleted within the next 50 - 100 years [1]. As a result, the search for renewable, sustainable, and clean energy sources has become a top priority. Hydrogen (H₂) stands out as a promising alternative, offering a clean, carbon-free energy source with a high energy density (142 kJ g⁻¹) that can be efficiently converted into electricity and utilized for domestic and industrial applications [2]. H₂ can be generated through thermochemical, electrochemical, and biochemical processes. Among these, biological H₂ production using green microalgae has gained increasing attention due to its potential for sustainability and reliance on inexhaustible solar energy via photosynthesis.

Green algae are a highly diverse group of phototrophic eukaryotic organisms that mostly perform oxygenic photosynthesis. They can produce H₂ using electrons and protons derived from the oxidation of water molecules during photosynthesis. Under anaerobic conditions, driven by sunlight, green algae can redirect their endogenous photosynthetic electron flow in the thylakoid membranes toward H₂ production [3]. This terminal reaction is catalyzed by the nuclear-encoded, chloroplast-localized [FeFe] hydrogenase enzyme [4,5]. H₂ production by green algae depends on several factors, including cell age and cell optical density [6], medium composition [7,8], temperature [9], pH [10], and light intensity [11].

Despite their potential, the overall efficiency of H₂ production in green algae remains low and species dependent. Key limitations include the extreme O₂ sensitivity of hydrogenase enzymes, competition for electron flow during photosynthesis, and suboptimal cultivation conditions [12,13]. Furthermore, most studies to date have focused on model organisms such as Chlamydomonas reinhardtii, with limited exploration of other algal taxa. The lack of systematic screening and strain development presents a significant bottleneck to the advancement of algal-based H₂ production technologies. Therefore, there is a pressing need to explore alternative green algal species that combine high H₂ productivity, stress tolerance, and compatibility with large-scale cultivation systems.

Monoraphidium sp., a planktonic freshwater microalga from the Selenastraceae family (Chlorophyceae, Chlorophyta), exhibits several traits desirable for industrial applications. It is known for its high biomass productivity, substantial lipid content, and adaptability to a range of environmental conditions [14-16]. This genus has been widely studied for applications in biodiesel production [17-21], wastewater treatment [20], and nutritional supplementation due to its rich content of protein, methionine, and α-linolenic acid [22]. Despite its promising biochemical profile, the H₂ production potential of Monoraphidium sp. remains largely unexplored. Preliminary studies have hinted at the H₂-producing capability of Monoraphidium, but comprehensive investigations are lacking. This knowledge gap underscores the need for detailed studies to assess its viability as a biohydrogen producer. Accordingly, this study focuses on Monoraphidium sp. KMITL-1, aiming to evaluate and optimize its H₂ production potential under varying culture conditions. The findings could contribute significantly to expanding the diversity of algal species utilized in sustainable H₂ production systems.

Materials and methods

Materials

The unicellular green alga Monoraphidium sp. KMITL-1 was isolated from hydroponic water at the Plant Tissue Culture Laboratory, Department of Biology, School of Science, King Mongkut’s Institute of Technology Ladkrabang, Bangkok, Thailand. The key materials used in the experimental procedures are listed in Table 1, and the overall experimental workflow is illustrated in Figure 1.

Table 1 Materials used in this study.

Figure 1 Flowchart illustrating the process of strain identification and optimization of H₂ production by the unicellular green alga Monoraphidium sp. KMITL-1.

Green algal cultivation and identification

Monoraphidium sp. KMITL-1 was cultivated in a 250-mL Erlenmeyer flask containing 100 mL of TAP medium (pH 7.2) [23]. The culture was incubated on a rotary shaker at 120 rpm under continuous white light at an intensity of 30 µmol photons m⁻² s⁻¹ at 30 °C for 3 days. Morphological characteristics were examined using a bright-field microscope (Nikon Eclipse Ci-L, Japan). For molecular characterization, genomic DNA was extracted using the Wizard^® SV DNA Purification System Kit (Promega, Madison, WI, USA). The 23S plastid rRNA gene was amplified by PCR Biometra^® T Professional Thermocycler (Biometra, Göttingen, Germany) using the following primers: p23SrV_f1 (5’-GGACAGAAAGACCCTATGAA-3’) and p23SrVr1 (5’-TCAGCCTGTTATCCCTAGAG-3’) [24]. The 50 L of PCR reaction contained 1X KAPA Taq Ready Mix (KAPA biosystems, South Africa), 25 pmol of each primer, and 50 ng of total genomic DNA. PCR was performed under the following conditions: An initial denaturation at 94 °C for 5 min, followed by 35 cycles of denaturation at 94 °C for 45 s, annealing at 52 °C for 1 min, and extension at 72 °C for 90 s, with a final extension at 72 °C for 10 min. The PCR product was purified using the QIAquick^® PCR purification System Kit (Qiagen, Hilden, Germany) before sequencing. The 23S plastid rRNA gene sequence was analyzed for homology using the NCBI BLASTn tool [25]. Sequence alignment was performed using the ClustalW program [26], and molecular evolutionary genetic analysis was conducted using the Maximum Likelihood method with 1,000 bootstrap replicates, implemented in Molecular Evolutionary Genetics Analysis (MEGA) software version 11 [27].

Growth measurement

Monoraphidium sp. KMITL-1 was cultivated in TAP medium under the previously described conditions for 5 days, with an initial optical density at 750 nm (OD₇₅₀) of approximately 0.1. The culture was sampled every 6 h to monitor growth. Growth was determined by measuring OD₇₅₀ using a spectrophotometer (Shimadzu, UV-601, Japan). Total chlorophyll content was determined according to the method described by [28]. Cell concentration was measured using a hemocytometer (Boeco, Germany).

Measurement of H₂ production

Monoraphidium sp. KMITL-1 was cultivated in TAP medium for 48 h, harvested by centrifugation at 8,000× g at 4 °C for 10 min, and subsequently washed 3 times with fresh TAP medium. The cell suspension was adjusted to an OD₇₅₀ of 0.5, and the cell pellet was resuspended in 5 mL of fresh TAP medium. The cell suspension was then transferred into a 12-mL glass vial, which was sealed with a rubber stopper and an aluminum crimp. To eliminate O₂ from the system, the vial was purged with argon gas for 10 min and subsequently shaken at 120 rpm under a light intensity of 30 µmol photons m⁻² s⁻¹. H₂ production was analyzed using a gas chromatograph with a thermal conductivity detector (GC-TCD) (Hewlett-Packard, HP5890A GC, Japan), equipped with a molecular sieve 5 Å 60/80 mesh packed column, using 99.999% (v/v) argon gas as the carrier gas and 4% (v/v) H₂ in argon as the standard H₂ gas (Praxair, Thailand). The injector and detector temperatures were set at 100 °C, while the column temperature was maintained at 50 °C.

Measurement of hydrogenase activity

Hydrogenase (H₂ase) activity was measured in the presence of methyl viologen and sodium dithionite. A 2-mL reaction mixture contained 1 mL of algal cell suspension, 1 mL of 50 mM phosphate buffer solution (PBS) (pH 6.8) containing 10 mM methyl viologen, 40 mM sodium dithionite, and 0.2% (v/v) Triton X-100 [7]. The reaction was carried out at 30 °C in darkness for 30 min under anaerobic conditions. Hydrogenase activity, indicated by the amount of H₂ produced, was measured using a gas chromatograph.

Effect of cell age and cell density on H₂ production

Monoraphidium sp. KMITL-1 was cultivated in TAP for 24, 48, 72, and 96 h. Cells were subsequently harvested by centrifugation, washed, and resuspended in fresh TAP before measuring H₂ production. To investigate the effect of cell density on H₂ production, the cell suspension was adjusted to OD₇₅₀ values of 0.2, 0.4, 0.6, 0.8, and 1.0 before measuring H₂ production by a gas chromatograph.

Effect of nutrient deprivation on H₂ production

Monoraphidium sp. KMITL-1 was cultivated in TAP for 24 h, harvested by centrifugation, washed, and resuspended in different types of TAP media, including potassium-deprived TAP (TAP-K), nitrogen-deprived TAP (TAP-N), phosphorus-deprived TAP (TAP-P), and sulfur-deprived TAP (TAP-S). Normal TAP served as the control. To remove sulfur in TAP-S, MgSO₄.6H₂O, FeSO₄.7H₂O, ZnSO₄.7H₂O, and CuSO₄.5H₂O were excluded from the medium, but the metal ions Mg²⁺, Fe²⁺, Zn²⁺, and Cu²⁺ were replaced with MgCl₂, FeCl₂, ZnCl₂ and CuCl₂, respectively, at the same ion concentrations. In TAP-N, NH₄Cl was removed from the medium. In TAP-P, KH₂PO₄ and K₂HPO₄ were removed, but K⁺ was replaced by adding KCl. In TAP-K, KH₂PO₄ and K₂HPO₄ were removed, but PO₄³⁻ was replaced by adding NaH₂PO₄ and Na₂HPO₄. The composition modifications for each TAP medium are summarized in Table 2. The cells were incubated under nutrient-deprived conditions for 48 h under light aerobic conditions. After incubation, the cells were harvested, resuspended in 5 mL of the respective media, and transferred into a 12 mL gas vial before measurement of H₂ production and hydrogenase activity using a gas chromatograph.

Table 2 Composition of standard TAP medium and single-nutrient-deprived TAP media, including TAP-S, TAP-N, TAP-P, and TAP-K. The presence (+) and absence (−) symbols indicate the availability and unavailability of each chemical compound in the culture medium, respectively. Some chemical compounds were replaced with the indicated alternatives.

Effect of carbon source and concentration on H₂ production

Monoraphidium sp. KMITL-1 cells were harvested, washed, and resuspended in optimal nutrient-deprived TAP medium supplemented with different carbon sources, including glucose, lactose, ethanol, butanol, propanol, citric acid, and glycerol. Acetic acid served as the control carbon source. The corresponding carbon source concentrations were 17.5, 35, 70, 175, 350, 700 and 1,750 mmol C-atom L⁻¹, with the control condition set at 35 mmol C-atom L⁻¹ (C-atom concentration in TAP). H₂ production was measured using a gas chromatograph.

Effect of pH, temperature, and light intensity on H₂ production

Monoraphidium sp. KMITL-1 cells were harvested, washed, and resuspended in optimized TAP medium under various conditions, including pH levels (6.0, 7.0, 7.2, 8.0, and 9.0), incubation temperatures (25, 30, 35, and 40 °C), and light intensities (0, 30, 60, 90, and 120 µmol photons m⁻² s⁻¹). The control conditions of pH, temperature, and light intensity were 7.2, 30 °C, and 30 µmol photons m⁻² s⁻¹, respectively. H₂ production was measured using a gas chromatograph.

Statistical analysis

All data are expressed as means of at least 3 independent experiments. Error bars represent the standard deviation. Significant differences were determined using one-way analysis of variance (ANOVA). The data were analyzed using Duncan’s multiple range test in IBM SPSS Statistics version 29.0, with a significance level of p < 0.05.

Results and discussion

Microalgal identification

In this study, Monoraphidium sp. KMITL-1 was identified using both morphological and molecular analyses. Under light microscopy, it exhibited unicellular, green-coloured cells with a crescent or lunate shape, tapering at the ends. The cells measured 15 - 20 µm in length and 1 - 5 µm in width (Figure 2(A)). Monoraphidium is classified in the family Selenastraceae, and is characterized morphologically by cells containing a single nucleus, a parietal chloroplast without pyrenoids, and asexual reproduction via autospore formation [29]. The morphology of Monoraphidium sp. KMITL-1 closely resembled Monoraphidium sp. HDMA-11 [30] and showed similarities to various Monoraphidium strains previously reported [14,31]. Identification of Monoraphidium species is primarily based on key morphological traits, including the shape of the cell apices (round, tapered, finger-like, or needle-like), overall cell shape (cylindrical, straight, slightly curved, reniform, arched, sigmoid, or spiral), and cell length [31]. Additionally, species can be distinguished by the presence or absence of a starch envelope and pyrenoid-penetrating thylakoids [14]. However, morphological characteristics alone are insufficient for precise genus identification; therefore, molecular characterization was employed to confirm the morphological classification.

For molecular analysis, the 23S plastid rRNA gene of Monoraphidium sp. KMITL-1 was sequenced and submitted to the NCBI GenBank under accession number PQ469933.1. Phylogenetic analysis revealed that Monoraphidium sp. KMITL-1 was closely related with 97.12% similarity to Monoraphidium sp. P5-4 and clustered with other Monoraphidium strains and green algae within the family Selenastraceae, such as Ankistrodesmus, Selenastrum, and Raphidocelis (Figure 2(B)). These strains were phylogenetically distinct from other genera such as Scenedesmus, Tetradesmus, Chlorella, Chlamydomonas, and Chlorosarcinopsis (Figure 2(B)). The molecular data confirmed that Monoraphidium sp. KMITL-1 belongs to the genus Monoraphidium. A previous study reported that Monoraphidium sp. P5-4, isolated from tropical freshwater, showed potential as a biodiesel feedstock [32]. However, to date, H₂ production by Monoraphidium species has not been much investigated.

Figure 2 Observation using microscopy of Monoraphidium sp. KMITL-1 (A) and phylogenetic tree of Monoraphidium sp. KMITL-1 and other green microalgae constructed from the 23S plastid rRNA gene (B).

Growth of Monoraphidium sp. KMITL-1

Growth of Monoraphidium sp. KMITL-1 was determined by measuring OD₇₅₀, total chlorophyll content, and total cell concentration (Figure 3). The strain exhibited rapid growth, with a specific growth rate of 0.45 ± 0.02 day⁻¹ and a doubling time of approximately 1.5 days. The highest total chlorophyll content of 9.67 ± 0.99 g mL⁻¹ and a maximum cell density of 7.44 ± 0.01×10⁶ cells mL⁻¹ were observed after 3 days of cultivation (Figure 3). This growth rate was approximately twice as high as that of Monoraphidium neglectum [33] and Monoraphidium sp. DeK19 [34]. The differences in growth rates are likely due to variations in cultivation media and algal species. In this study, Monoraphidium sp. KMITL-1 was grown in TAP medium under photoheterotrophic conditions using acetic acid as a carbon source, whereas M. neglectum was cultivated photoautotrophically in BG11 medium without an added carbon source, and Monoraphidium sp. DeK19 was grown in wastewater effluent [33,34]. These results suggest that TAP is a suitable medium for cultivating Monoraphidium species, as it provides essential nutrients along with various metals and minerals.

Figure 3 Growth of Monoraphidium sp. KMITL-1 cultivated in TAP by OD₇₅₀ (▲), total chlorophyll concentration (●) and total cell concentration (■) measurements.

Effect of cell age and cell density on H₂ Production

The growth phases of algal cells, or cell ages, are associated with different metabolic activities, including H₂ production. In this study, cells harvested at 24, 48, 72, and 96 h corresponded to the lag, mid-log, late-log, and stationary phases, respectively. The results showed that Monoraphidium sp. KMITL-1 cells cultivated in TAP for 48 h exhibited the highest H₂ production rate of 2.990 ± 0.021 µmol H₂ mg Chl⁻¹ h⁻¹ (Figure 4). Cells harvested at earlier or later time points displayed significantly lower H₂ production rates. Moreover, the 48-hour-old cells achieved the highest cumulative H₂ production, reaching 110.163 ± 0.406 µmol H₂ mg Chl⁻¹ after 96 h of anaerobic incubation (Figure 4). These findings indicate that the growth phase significantly affects H₂ production, with cells in the logarithmic phase yielding the highest levels. This may be attributed to the high metabolic activity and photosynthetic efficiency during the logarithmic phase, which generates a large number of electrons through water oxidation at Photosystem II (PSII). These electrons serve as substrates for the hydrogenase enzymes responsible for H₂ production. In contrast, Chlamydomonas reinhardtii UTEX 90 was reported to produce the highest H₂ levels during the late exponential phase [35].

Figure 4 H₂ production rate (A) and maximum cumulative H₂ production (B) by Monoraphidium sp. KMITL-1 under different cell ages. Statistical analysis was performed using one-way ANOVA, with significance considered at p < 0.05. Data are presented as mean ± standard deviation.

The effect of cell density on H₂ production was investigated using 48-hour-old cultures of Monoraphidium sp. KMITL-1 at varying OD₇₅₀ values. The results showed that increasing the OD₇₅₀ from 0.2 to 0.8 led to higher H₂ production rates, corresponding with increased cell densities. Cultures with an OD₇₅₀ of 0.8 exhibited the highest H₂ production rate of 4.803 ± 0.060 µmol H₂ mg Chl⁻¹ h⁻¹ and achieved the maximum cumulative H₂ production of 288.127 ± 1.197 µmol H₂ mg Chl⁻¹ after anaerobic incubation for 48 h (Figure 5). This enhanced production at OD₇₅₀ of 0.8 can be attributed to optimal PSII activity, resulting in greater electron availability for H₂ase-mediated H₂ production. At lower cell densities (OD₇₅₀ less than 0.8), reduced PSII activity likely led to insufficient electron generation. Conversely, at OD₇₅₀ values higher than 0.8, H₂ production decreased, probably due to limited light penetration within the culture vials, which hindered effective photosynthesis and electron generation. Similar findings have been reported in Chlamydomonas reinhardtii CC-125 and Scenedesmus sp. KMITL-OVG1 [8,36]. Based on these findings, Monoraphidium sp. KMITL-1 cultures at a cell age of 48 h and an OD₇₅₀ of 0.8 were selected for subsequent studies on H₂ production.

Figure 5 H₂ production rate (A) and maximum cumulative H₂ production (B) by Monoraphidium sp. KMITL-1 under different cell densities. Statistical analysis was performed using one-way ANOVA, with significance considered at p < 0.05. Data are presented as mean ± standard deviation.

Effect of nutrient deprivation on H₂ and O₂ production, and H₂ase activity

The H₂ and O₂ production rates, along with H₂ase activity, of Monoraphidium sp. KMITL-1 was investigated under various nutrient-deprived conditions. Monoraphidium sp. KMITL-1 cells exhibited the highest H₂ production rate of 11.388 ± 0.032 µmol H₂ mg Chl⁻¹ h⁻¹, the lowest O₂ production rate of 4.455 ± 0.016 µmol O₂ mg Chl⁻¹ h⁻¹ and the highest H₂ase activity at 3.540 ± 0.026 µmol H₂ mg Chl⁻¹ min⁻¹ in cells incubated in TAP-K (Table 3). This H₂ production rate was approximately 2- to 5-fold higher than those observed under other nutrient deprivation conditions. Similarly, the maximum cumulative H₂ production yield of 677.102 ± 0.574 µmol H₂ mg Chl⁻¹ was observed in cells incubated in TAP-K after 48 h of anaerobic incubation (Table 3). H₂ production by cells incubated in TAP-N, -P, and -S was significantly lower than that in normal TAP (Table 3). The lowest H₂ production rate of 2.426 ± 0.034 µmol H₂ g Chl⁻¹ h⁻¹, was recorded in cells incubated in TAP-S (Table 3). In this study, H₂ production was inversely related to O₂ production but corresponded with H₂ase activity (Table 3). Since O₂ is a strong inhibitor of H₂ase, a decrease in O₂ levels resulted in higher H₂ase activity, ultimately enhancing H₂ production.

Table 3 H₂ production, O₂ production and H₂ase activity of Monoraphidium sp. KMITL-1 under different nutrient-deprived conditions. Data are presented as mean ± standard deviation.

Essential macronutrients, including S, N, P, and K, play critical roles in regulating the growth and metabolism of phototrophic microalgae. Deficiencies in these nutrients impose cellular stress, ultimately leading to the cessation of cell division and growth [37]. In this study, K deprivation was found to significantly enhance H₂ production. The lack of K impaired protein synthesis and decreased PSII activity, primarily by disrupting the replacement of the D1 protein [38]. As a result, the rate of O₂ evolution through PSII fell below the rate of respiratory O₂ consumption, creating anoxic conditions in closed systems. These anoxic conditions, in turn, induced H₂ase activity and promoted H₂ production [38]. In addition, the increased H₂ production might be due to the reduction of starch accumulation and the acceleration of starch degradation, leading to the provision of many electrons, the substrate for H₂ase [36]. Previous reports showed that K deprivation promoted H₂ production only in Scenedesmus strains [36,38]. This study shows that Monoraphidium is another green algal strain whose H₂ production is enhanced by K deprivation.

The deprivation of N, S, and P imposes severe stress on green algal growth, as these nutrients are essential components of proteins, nucleic acids, and other critical biomolecules. In Monoraphidium sp. KMITL-1, deprivation of N, P, or S did not enhance H₂ production. Among these nutrient deprivations, N deprivation appeared to have the least impact on H₂ production (Table 3). N deprivation has been reported to stimulate H₂ production in other algal species, such as Chlamydomonas reinhardtii [39], Chlorella sp. ChiW4 [40], and Chlorella sp. LSD-W2 [28], primarily through increased electron availability from photosynthesis and the degradation of accumulated starch. P deprivation has been shown to enhance H₂ production in marine green microalgae Chlorella sp., by promoting starch accumulation and facilitating anaerobic conditions [41].

In the present study, S deprivation led to the most significant reduction in both H₂ase activity and H₂ production, likely due to the lack of S required for H₂ase synthesis. Nevertheless, previous studies have demonstrated prolonged H₂ photoproduction under S deprivation in C. reinhardtii [42]. Under S-deprived conditions, the inability to resynthesize the D1 protein results in the gradual inhibition of PSII-dependent O₂-evolving activity, subsequently lowering O₂ levels, which can induce H₂ase activity and promote H₂ production. The decrease in H₂ production by Monoraphidium sp. KMITL-1 under N, P, and S deprivation requires further investigation. These findings suggest that the pattern of H₂ production differs depending on the algal strains. H₂ production observed in Monoraphidium sp. KMITL-1 closely resembles that of Scenedesmus species but differs from that of other green algal strains.

Effect of carbon source and concentration on H₂ production

H₂ production by Monoraphidium sp. KMITL-1 was evaluated in TAP-K supplemented with various carbon sources, including acetic acid, butanol, citric acid, glucose, glycerol, ethanol, lactose, propanol, and sucrose, each at an equivalent carbon atom concentration of 35 mmol C-atom L⁻¹ (matching the carbon content of standard TAP medium). Among the tested carbon sources, glucose supplementation resulted in the highest H₂ production rate of 28.555 ± 0.093 µmol H₂ mg Chl⁻¹ h⁻¹, with a cumulative H₂ yield of 1,446.683 ± 0.585 µmol H₂ mg Chl⁻¹ after anaerobic incubation for 48 h (Figure 6). This yield was approximately 2.5-fold higher than that observed with acetic acid supplementation. Other carbon sources led to comparatively lower H₂ production rates and yields. The enhanced H₂ production with glucose is likely attributable to increased intracellular reducing equivalents generated via glycolysis, which facilitate electron transfer to H₂ase [43]. Therefore, glucose was selected for further investigation into the effects of carbon source concentration on H₂ production.

Figure 6 H₂ production rate (A) and maximum cumulative H₂ production (B) by Monoraphidium sp. KMITL-1 incubated in TAP-K medium supplemented with different carbon sources. Statistical analysis was performed using one-way ANOVA, with significance considered at p < 0.05. Data are presented as mean ± standard deviation.

By varying the glucose concentration from 17.5 to 1,750 mmol C-atom L⁻¹, the highest H₂ production rate of 45.211 ± 0.156 µmol H₂ mg Chl⁻¹ h⁻¹ was achieved at a glucose concentration of 350 mmol C-atom L⁻¹ (Table 4). This rate represented approximately a fourfold increase compared to the K-deprived medium supplemented with 35 mmol C-atom L⁻¹ glucose. Increasing glucose concentration likely enhanced the availability of electrons, the substrates for H₂ase activity, as glucose can be metabolized through glycolysis and subsequent fermentative pathways to generate reducing equivalents [44]. However, further increases in glucose concentration beyond 350 mmol C-atom L⁻¹ led to a decline in both H₂ production rate and cumulative yield (Table 4). This reduction may be attributed to the toxic effects of excess glucose, which can impose metabolic stress on cells by necessitating additional energy for glucose export [45,46]. Moreover, the low pH of the medium (pH 3.4) (Table 4) at higher glucose concentrations may have created an acidic environment that inhibited key enzymatic activities. Based on these findings, glucose supplementation at 350 mmol C-atom L⁻¹ was selected for subsequent experiments.

Table 4 H₂ production rate and maximum cumulative H₂ production by Monoraphidium sp. KMITL-1 cultured in TAP-K medium supplemented with varying glucose concentrations. Data are presented as mean ± standard deviation.

Effect of pH, temperature and light intensity on H₂ production

H₂ production by Monoraphidium sp. KMITL-1 incubated in TAP-K supplemented with 350 mmol C-atom L⁻¹ glucose was measured across a pH range of 6.0 to 9.0. A pH of 7.2, corresponding to the standard TAP medium, was used as the control. The maximum H₂ production rate of 45.506 ± 0.331 µmol H₂ mg Chl⁻¹ h⁻¹ was observed at pH 7.2 (Figure 7(A)). This rate was significantly higher than those recorded at other pH values. These results suggest that the H₂ase of Monoraphidium sp. KMITL-1 exhibits its highest catalytic activity for H₂ production at pH 7.2. Although lower pH conditions provide a higher concentration of protons, the substrate for hydrogenase, the enzyme activity was reduced under more acidic conditions, likely due to pH-induced enzyme inactivation. This result is consistent with previous studies found that Chlorella sp. KLSc59 and C. reinhardtii exhibited maximum H₂ production at pH 7.0 [6,47].

Figure 7 H₂ production rate of Monoraphidium sp. KMITL-1 under different initial pH values (A), temperatures (B), and light intensities (C). Statistical analysis was performed using one-way ANOVA, with significance considered at p < 0.05. Data are presented as mean ± standard deviation.

Among the tested incubation temperatures ranging from 25 to 40 °C, Monoraphidium sp. KMITL-1 showed the highest H₂ production rate of 45.585 ± 0.237 µmol H₂ mg Chl⁻¹ h⁻¹ at 30 °C (Figure 7(B)). Incubation at temperatures lower or higher than 30 °C led to a reduction in H₂ production. Similar to pH, the incubation temperature plays a critical role in regulating H₂ase activity, with 30 °C identified as the optimal temperature for maximal H₂ production by Monoraphidium sp. KMITL-1. However, the optimal incubation temperature for H₂ production varies depending on the species of green algae. Under light intensities ranging from 0 - 120 µmol photon m⁻² s⁻¹, Monoraphidium sp. KMITL-1 showed the highest H₂ production rate of 67.976 ± 1.096 µmol H₂ mg Chl⁻¹ h⁻¹ at 60 µmol photon m⁻² s⁻¹ (Figure 7(C)). This rate was approximately 10-fold and 1.5-fold higher than that observed under dark conditions and under a light intensity of 30 µmol photons m⁻² s⁻¹, respectively. Furthermore, Monoraphidium sp. KMITL-1 achieved the maximum cumulative H₂ production of 3,190.436 ± 2.219 µmol H₂ mg Chl⁻¹ after 72 h of incubation at 60 µmol photons m⁻² s⁻¹. Light serves as an essential energy source for H₂ production through photosynthesis, which generates the electrons required by H₂ase. However, photosynthesis also produces oxygen, a strong inhibitor of hydrogenase activity. Therefore, an optimal light intensity is necessary to balance sufficient electron supply with minimal O₂ accumulation. These findings are consistent with a previous study in which a light intensity of 60 µmol photons m⁻² s⁻¹ resulted in the highest H₂ production in Chlorella sp. LSD-W2 [10]. In contrast, Chlamydomonas reinhardtii exhibited maximum H₂ production at a higher light intensity of 200 µmol photons m⁻² s⁻¹ [48].

Conclusions

This study demonstrated that physiological and environmental factors significantly influence H₂ production by the unicellular green alga Monoraphidium sp. KMITL-1. Key factors examined included cell age, cell density, nutrient deprivation, carbon source and concentration, pH, temperature, and light intensity. The optimal conditions for H₂ production were identified as a 48-hour-old culture with OD₇₅₀ of 0.8. Potassium deprivation enhanced H₂ production by approximately 2.5-fold compared to non-deprived conditions. Among the carbon sources and concentrations tested, glucose at a concentration of 350 mmol C-atom L⁻¹ significantly stimulated H₂ production. These findings confirm that Monoraphidium sp. KMITL-1 is a promising candidate for sustainable and efficient biohydrogen production. Its ability to produce H₂ under defined and controllable conditions highlights its potential for integration into renewable energy systems. Future studies should focus on optimization of large-scale cultivation systems, and investigation of long-term H₂ production stability under outdoor conditions. Moreover, co-cultivation with bacteria or use of waste-derived substrates may further improve the feasibility and economic viability of algal-based H₂ production systems.

Acknowledgements

This research is a result of the paper entitled “Hydrogen Production by Green Algae by Co-cultivation with Bacteria Gant No. RE-KRIS/FF67/034” by King Mongkut’s Institute of Technology Ladkrabang (KMITL), which has been received funding support from the NSRF. The authors would like to thank the Department of Biology, School of Science, KMITL, for providing laboratory facilities and technical support throughout this study. Special thanks to the research staff and colleagues who contributed to the experimental work and data analysis.

Declaration of generative AI in scientific writing

The authors acknowledge the use of the generative AI tool ChatGPT in the preparation of this manuscript, specifically for language editing and grammar correction. No content generation or data interpretation was performed by the AI. The authors take full responsibility for the content and conclusions of this work.

CRediT author statement

Varanya Krutpan: Methodology, Investigation, Data curation, Formal analysis, Validation, and Writing–original draft preparation.

Thaninthorn Supakriangkrai: Data curation, Formal analysis, and Validation.

Pana Lohasupthawee: Resources, and Validation.

Saranya Phunpruch: Conceptualization, Methodology, Funding acquisition, Project administration, Resources, Supervision, Validation, and Writing–review and editing

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