Advances and Challenges in Biohydrogen Production by Photosynthetic Microorganisms

Abstract

Hydrogen (H₂) production by photosynthetic microorganisms is a viable option for renewable energy due to its sustainability and potential for widespread application. Green algae, cyanobacteria, and purple non-sulfur bacteria have shown great promise in bio-H₂ production. However, problems such as low H₂ production rates and high H₂ production costs continue to hinder the commercial scalability of these systems. To overcome these obstacles, genetic engineering selection of robust strains capable of coping with variable environmental conditions, optimization of growth conditions, use of wastewater, and biotechnological approaches such as immobilization are carefully considered. The aim of this review is to provide a thorough overview of the methods and developments that can improve H₂ production and to highlight current difficulties and future directions for further studies.

1. Introduction

Producing hydrogen (H₂) with a sustainable approach remains a challenge. The use of photosynthetic microorganisms such as microalgae, cyanobacteria, and photosynthetic bacteria for H₂ production is crucial. Specifically, cyanobacteria can adapt their metabolism depending on growth conditions, making it possible to extend the photobiological H₂ production under a wide range of climatic conditions. Moreover, the numerous species of both microalgae and cyanobacteria are an important source of biodiversity that encourages researchers to explore a variety of new applications. Photosynthesis is one of the most attractive ways of splitting water and storing solar energy in the form of H₂, as it can operate under climatic conditions, at ambient temperature and pressure, and without artificial catalysts. The process is carried out by two natural catalysts, namely, photosystem II (PSII), which splits water, and hydrogenase, which combines the electrons and protons produced by PSII to form H₂. In fact, some microalgae and cyanobacteria can switch from oxygenic photosynthesis to H₂ evolution under anaerobic conditions. Since water is a by-product of the reaction, this technique is particularly environmentally friendly [1]. The aim of this review is to discuss the potential of H₂ production by photosynthetic microorganisms, including photosynthetic bacteria. A comparison between the costs of photobiological H₂ production and those of water electrolysis is discussed.

1.1. Microalgae and Cyanobacteria

Unicellular photosynthetic organisms such as microalgae and cyanobacteria have hydrogenase enzymes that enable them to produce H₂ through photosynthetic activities (Figure 1). Due to the physiology and metabolism of microalgae and cyanobacteria, artificial and semi-artificial in vitro techniques have been developed that combine photoexcited materials, enzymes, and hydrogenases or their mimics to generate H₂. These in vitro methods have partially mimicked the combination of cyanobacterial and microalgal hydrogenases with photosystem I (PSI) in vivo, which has recently succeeded as a proof of concept [2,3].

One of the key elements in the production of H₂ from microalgae is the sensitivity to oxygen (O₂) of hydrogenase [4]. The production and function of this enzyme are inhibited when O₂ is present. This is a crucial step in the photosynthetic biological production of H₂ by photosynthetic microorganisms, where O₂ is produced as a by-product. The structure of hydrogenase and the sites at which it interacts with O₂ have been the subject of several studies to determine how to reduce or at least attenuate its sensitivity to O₂ [5,6]. However, since these studies were carried out in vitro, it is quite difficult to achieve this goal as the process needs to be repeated in vivo.

The aim of biotechnological approaches to produce H₂ is to develop a system based on oxygenic photosynthesis, since the only components required for H₂ production are sunlight and water.

Various strains of microalgae and cyanobacteria are able to produce H₂ under certain conditions. Some of the most known strains are reported in Table 1 with their main physiological characteristics.

1.1.1. Microalgal Strains

Among microalgae, Chlamydomonas reinhardtii has been extensively studied for H₂ production and serves as a model organism for the study of physiological and metabolic processes due to its easy cultivation and handling. In this microalga, there are two separate pathways that transport electrons to the hydrogenase via ferredoxin: (i) a direct pathway involving the residual activity of PSII that occurs under anaerobic conditions; (ii) an indirect pathway that does not require light, involving the use of electrons from internal substrates like carbohydrates through glycolysis or from external organic sources. Although Chlamydomonas reinhardtii is one of the most studied microalgae, several other genera can also produce H₂ under specific circumstances. According to Duangjan et al., these include Scenedesmus, Acutodesmus, Coelastrum, Desmodesmus, and Pandorina [7]. However, because of their ease of cultivation, known metabolism, and cell structure, Chlamydomonas and Chlorella are the most studied species. Therefore, many studies have been conducted on mutant strains of these microalgal genera to improve H₂ production [8,9].

Nutritional Strategies for Production and Improvement of H₂ in Microalgal Strains

Electrons for hydrogenase are generated throughout the process of H₂ production by the fermentative metabolism of carbohydrates previously collected in the light phase. Usually, carbohydrate synthesis is increased by nutrient starvation. Subsequently, carbohydrates are used for H₂ production by a combination of the enzymatic activity of pyruvate–ferredoxin oxidoreductase and a dark process. Sulfur, nitrogen, or potassium limitation negatively affects the efficiency of the photosynthetic apparatus, leading to a decrease in photosynthetic activity and a reduction in oxygen concentration [8]. Under these conditions, hydrogenase activity is triggered, and the process of H₂ production is initiated. Maintaining these conditions promotes the increase in H₂ evolution yield.

One of the most commonly used protocols, sulfur starvation, which is used in C. reinhardtii to reduce photosynthetic O₂ evolution, has several disadvantages. First, the culture must be centrifuged several times to remove all the sulfur traces from the medium. Second, the H₂ production rate is reduced due to the strong down-regulation of PSII. Given these limitations, scientists are searching for strains with a higher respiration to photosynthesis ratio than C. reinhardtii. In this regard, the microalga Chlorella appears to be very promising for H₂ production, as it is known to generate a significant amount of H₂ through carbohydrate catabolism. The shuttle system of this microalga for the uptake of carbohydrates is crucial because when cultivated in a medium containing sugars, the culture can achieve a very high biomass concentration. Moreover, this biomass could be rich in carbohydrates that could support the process of H₂ production. Chlorella can produce a significant amount of H₂ in both light and dark environments [10,11].

Table 1. Microalgal strains and their characteristics in H₂ production.

Microalgal Strain	Main Characteristics	H2 Production Methods	Advantages	Disadvantages	References
Chlamydomonas reinhardtii	Model organism for H2 production	Direct pathway (PSII) and indirect pathway (carbohydrate catabolism)	Easy to cultivate and manipulate	O2-sensitive; requires sulfur deprivation	[7,8,9]
Chlorella vulgaris	High ability to produce H2 from carbohydrates	Carbohydrate catabolism, production in both light and dark	High biomass and H2 production	Requires sugar-containing media. Augmented risk of contamination	[7,11]
Scenedesmus obliquus	Increased H2 production under potassium deprivation	PSII-independent pathway	Higher production without O₂ generation	Requires specific deprivation conditions	[7,12,13,14]
Synechocystis PCC 6803	Model cyanobacterium, genetically manipulable	Indirect light-driven and anaerobic metabolism production	Acclimatable to various environmental conditions	Limited production in some mutations. High risk of grazing by Poterioochromonas	[15]

Table 1. Microalgal strains and their characteristics in H₂ production.

Microalgal Strain	Main Characteristics	H2 Production Methods	Advantages	Disadvantages	References
Chlamydomonas reinhardtii	Model organism for H2 production	Direct pathway (PSII) and indirect pathway (carbohydrate catabolism)	Easy to cultivate and manipulate	O2-sensitive; requires sulfur deprivation	[7,8,9]
Chlorella vulgaris	High ability to produce H2 from carbohydrates	Carbohydrate catabolism, production in both light and dark	High biomass and H2 production	Requires sugar-containing media. Augmented risk of contamination	[7,11]
Scenedesmus obliquus	Increased H2 production under potassium deprivation	PSII-independent pathway	Higher production without O₂ generation	Requires specific deprivation conditions	[7,12,13,14]
Synechocystis PCC 6803	Model cyanobacterium, genetically manipulable	Indirect light-driven and anaerobic metabolism production	Acclimatable to various environmental conditions	Limited production in some mutations. High risk of grazing by Poterioochromonas	[15]

To avoid nutrient starvation, a strain of Chlorella vulgaris, BEIJ strain G-120, was studied for its ability to produce H₂ through both direct and indirect pathways [11]. Under light and dark conditions, a total H₂ production of 896 mL and 406 mL was recorded, respectively, and this strain was able to achieve a maximum H₂ production rate of 12 mL H₂/L/h [11]. By avoiding all the steps required for nutrient deprivation, the ability to induce hydrogenase activity throughout the medium allows the use of a simpler procedure in terms of imposing nutrient starvation. In this way, the process saves time and reduces the risk of cultural contamination during set-up.

In Scenedesmus obliquus, potassium deprivation also increases H₂ generation [12]. In most biological reactions, sodium can partially replace potassium so that the system can continue to function. This discovery considerably boosts the ability of the PSII-independent pathway to produce H₂. The breakdown of glucose resulted in a constant donation of electrons to the PQ pool of the photosynthetic electron chain, leading to the above-mentioned finding. Furthermore, the PSII-independent route leads to a sustained increase in H₂ production through PSII inactivation (which inhibits O₂ generation), PSI synthesis, and plastid hydrogenase overactivation [12].

The microalga Scenedesmus obliquus showed the ability to biodegrade dichlorophenols (dcps) under specific autotrophic and mixotrophic conditions [13]. In this microalga, an overactivated photosynthetic electron flow and H₂-productivity were promoted by the addition of 3,4-dichlorophenol in nitrogen-starved cultures, which was due to the large synthesis of Cytb6f that quenched the excess electrons generated by dichlorophenol biodegradation. This work provided a detailed analysis of the bioenergetic strategy of the green alga S. obliquus under nitrogen deficiency and the simultaneous presence of the hazardous phenolic chemical 3,4-dichlorophenol (dcp). This information is useful for finding new strategies to implement the H₂ production process. Using Scenedesmus sp. in PHM-S medium, the anaerobic and photosynthetic processes were used to study H₂ production [14]. The data show that the second experiment produced the most H₂. This was due to the extended incubation time of the second experiment, which prolonged the photosynthetic process and reduced the amount of H₂ gas that the algae could produce. It is interesting to note that H₂ is not produced in a specific period of time. We believe that this is related to the enzymatic reaction in algae, which is mainly caused by O₂. The enzyme hydrogenase was inhibited by O₂, which decreased during anaerobic adaptation, resulting in increased H₂ synthesis. Table 2 and Table 3 show the H₂ production rates for the most productive strains (microalgae and cyanobacteria) reported in the last five years.

1.1.2. H₂ Production in the Cyanobacterium Synechosyctis PCC 6803

Among photosynthetic microorganisms, the cyanobacterium Synechocystis PCC6803 was considered for H₂ production experiments because it thrives well under various environmental conditions, is easy to manipulate due to its small genome, and has been well characterized. For this reason, it is considered a cyanobacterial model strain [16]. Unfortunately, this organism is strongly grazed by the Crysophyta Poterioochromonas malhamensis, a globally distributed species that poses a serious threat to mass culture. However, strategies have been tested to successfully protect the culture from contamination. These include the use of high pH (approximately 11) and high salinity (similar to seawater) with negligible loss of productivity [17].

Under dark and anaerobic conditions, Synechocystis PCC 6803 produces organic acids such as acetate, lactate, and four-carbon dicarboxylic acids (succinate, fumarate, and malate) via the glycogen catabolism pathway, as well as H₂. Both pathways that lead to the production of organic acids and the hydrogenase enzyme that leads to H₂ production use NADH. Altering the fermentation pathway for organic acid formation may lead to a change in NADH utilization that affects the increase or decrease in H₂ production under dark, anaerobic conditions.

Some mutant strains were studied to investigate the role of the balance of intracellular redox potential (involved in the production of organic compounds) in the production of H₂, under anaerobic conditions [18]. In fact, overexpression of malate dehydrogenase, inhibition of d-lactate dehydrogenase, or inhibition of acetate kinase altered the ratios of organic acids and H₂ in these mutants under dark anaerobiosis. In particular, mutant strains showed variations from the original wild-type strain in terms of the NADH/NAD⁺ ratio and the oxidation of reducing powers. When organic acid accumulation was increased in the mutant strains, the NADH/NAD+ ratio was lower and the H₂ production decreased compared to a glucose-tolerant strain. These studies provided important information on the role of the intracellular redox balance of Synechocystis PCC 6803, particularly the NADH/NAD+ ratio, in the regulation of organic acid and H₂ production under dark anaerobic conditions.

The growth of Synechocystis PCC 6803 was studied outdoors in a 50 L tubular photobioreactor (PBR) at different mean initial biomass concentrations (1.10, 0.60, and 0.32 g/L) to test the ability of this strain to acclimate to solar light [15]. These studies were interesting to investigate the possibility of producing H₂ with this strain using solar light. To this end, the ability of Synechocystis PCC 6803 to produce H₂ by an indirect light-driven mechanism was investigated both in the laboratory and in a 50 L outdoor PBR. The rate of H₂ production was 0.05 mL H₂/L/h, while 3.07 mL H₂/L/h was achieved in the laboratory. The fermentation of carbohydrates that accumulated in the light during nitrogen starvation of the cells maintained H₂ production in the dark.

Table 2. H₂ production rate by various microalgae.

Microalgae	Growth Conditions	H2 Production Conditions	H2 Production	References
Closterium moniliferum AARL G041	Jaworski’s medium (JM), autotrophically, 30.8 μE/m2/s, 25 °C	Sulfur-free JM medium (JM-S), 54 μE/m2/s 25 °C, under Ar, 60 mL vial serum bottles	0.38 mmol/h/mg (Chl)	[7]
Chlorella vulgaris	Artificial wastewater medium, immobilized, 140 μE/m2/s, 25 ± 1 °C	Wastewater medium 10 g/L glucose, sulfur deprivation, 140 μE/m2/s, purple light, 25 ± 1 °C, under N2 atmosphere, pH 8	1.63 mL/L/h (or 39.18 mL/L/day)	[19]
Scenedesmus obliquus	Artificial wastewater medium, immobilized, 140 μE/m2/s, 25 ± 1 °C	Wastewater medium 10 g/L glucose, sulfur deprivation, 140 μE/m2/s, purple light, 25 ± 1 °C, under N2 atmosphere, pH 8	8.53 mL/L/h (or 204.8 mL/L/day)	[19]
Chlorella pyrenoidosa	Tris-acetate-phosphate (TAP) medium þ 10 mM NaHCO3 (TCP medium), immobilized, pH 7, 180 ± 10 μE/m2/s, 28 °C, a 3.925 L airlift PBR	TCP medium þ injection of 10 mM DCMU (3-(3,4-dichlorophenyl)-1,1-dimethylurea), after 9 h injection, under N2, under 24 h darkness then 180 ± 10 μE/m2/s, 28 °C, anaerobic bottles with 75 mL TCP medium	93.86 mL/L	[20]
Tetraspora sp. CU2551	Tris-acetate-phosphate (TAP) medium, immobilized, pH 7.2, 29 μE/m2/s, 36 °C, 125 mL Erlenmeyer flasks	Sulfur-free TAP medium (TAP-S), under air, pH 7.2, 29 μE/m2/s, 36 °C, 100 mL gas-tight vials	0.182 ± 0.020 mmol/h/mg dry wt	[21]
Chlorella lewinii KU201	Tris-acetate-phosphate (TAP) medium 0.7 mM NH4Cl, 25 °C, 5 μE/m2/s, 14:10 h light/dark cycle, pH 7.3	Sulfur-free TAP medium (TAP-S) 0.7 mM NH4Cl, 25 °C, 35 μE/m2/s, pH 7.3, under Ar, 650 mL bioreactors	13.03 mL/L	[22]
Chlorella sp. KU209	Tris-acetate-phosphate (TAP) medium þ 0.7 mM NH4Cl, 25 °C, 35 μE/m2/s, 14:10 h light/dark cycle, pH 7.3	Sulfur-free TAP medium (TAP-S) þ 0.7 mM NH4Cl, 25 °C, 35 μE/m2/s, pH 7.3, under Ar, 650 mL bioreactors	12.67 mL/L	[22]
Chlorella sorokiniana	KU204 tris-acetate-phosphate (TAP) medium 0.7 mM NH4Cl, 25 °C, 35 μE/m2/s, 14:10 h light/dark cycle, pH 7.3	Sulfur-free TAP medium (TAP-S) 0.7 mM NH4Cl, 25 °C, 35 μE/m2/s, pH 7.3, under Ar, 650 mL bioreactors	Maximum H2 1.30 mL/L/h, total H2 89.64 mL/L	[22]

Table 2. H₂ production rate by various microalgae.

Microalgae	Growth Conditions	H2 Production Conditions	H2 Production	References
Closterium moniliferum AARL G041	Jaworski’s medium (JM), autotrophically, 30.8 μE/m2/s, 25 °C	Sulfur-free JM medium (JM-S), 54 μE/m2/s 25 °C, under Ar, 60 mL vial serum bottles	0.38 mmol/h/mg (Chl)	[7]
Chlorella vulgaris	Artificial wastewater medium, immobilized, 140 μE/m2/s, 25 ± 1 °C	Wastewater medium 10 g/L glucose, sulfur deprivation, 140 μE/m2/s, purple light, 25 ± 1 °C, under N2 atmosphere, pH 8	1.63 mL/L/h (or 39.18 mL/L/day)	[19]
Scenedesmus obliquus	Artificial wastewater medium, immobilized, 140 μE/m2/s, 25 ± 1 °C	Wastewater medium 10 g/L glucose, sulfur deprivation, 140 μE/m2/s, purple light, 25 ± 1 °C, under N2 atmosphere, pH 8	8.53 mL/L/h (or 204.8 mL/L/day)	[19]
Chlorella pyrenoidosa	Tris-acetate-phosphate (TAP) medium þ 10 mM NaHCO3 (TCP medium), immobilized, pH 7, 180 ± 10 μE/m2/s, 28 °C, a 3.925 L airlift PBR	TCP medium þ injection of 10 mM DCMU (3-(3,4-dichlorophenyl)-1,1-dimethylurea), after 9 h injection, under N2, under 24 h darkness then 180 ± 10 μE/m2/s, 28 °C, anaerobic bottles with 75 mL TCP medium	93.86 mL/L	[20]
Tetraspora sp. CU2551	Tris-acetate-phosphate (TAP) medium, immobilized, pH 7.2, 29 μE/m2/s, 36 °C, 125 mL Erlenmeyer flasks	Sulfur-free TAP medium (TAP-S), under air, pH 7.2, 29 μE/m2/s, 36 °C, 100 mL gas-tight vials	0.182 ± 0.020 mmol/h/mg dry wt	[21]
Chlorella lewinii KU201	Tris-acetate-phosphate (TAP) medium 0.7 mM NH4Cl, 25 °C, 5 μE/m2/s, 14:10 h light/dark cycle, pH 7.3	Sulfur-free TAP medium (TAP-S) 0.7 mM NH4Cl, 25 °C, 35 μE/m2/s, pH 7.3, under Ar, 650 mL bioreactors	13.03 mL/L	[22]
Chlorella sp. KU209	Tris-acetate-phosphate (TAP) medium þ 0.7 mM NH4Cl, 25 °C, 35 μE/m2/s, 14:10 h light/dark cycle, pH 7.3	Sulfur-free TAP medium (TAP-S) þ 0.7 mM NH4Cl, 25 °C, 35 μE/m2/s, pH 7.3, under Ar, 650 mL bioreactors	12.67 mL/L	[22]
Chlorella sorokiniana	KU204 tris-acetate-phosphate (TAP) medium 0.7 mM NH4Cl, 25 °C, 35 μE/m2/s, 14:10 h light/dark cycle, pH 7.3	Sulfur-free TAP medium (TAP-S) 0.7 mM NH4Cl, 25 °C, 35 μE/m2/s, pH 7.3, under Ar, 650 mL bioreactors	Maximum H2 1.30 mL/L/h, total H2 89.64 mL/L	[22]

Table 3. H₂ production rate by various cyanobacteria.

Cyanobacteria	Growth Conditions	H2 Production Conditions	H2 Production	References
Immobilized Synechocystis PCC 6803	BG11 medium, 28 °C, 70 μE/m2/s, 97% air 3% CO2, Glass tube (400 mL)	Nitrogen-free medium BG110-Tris, 60 μE/m2/s, 28 °C, under N2 atmosphere, serum vials (60 mL)	Total H2 5.80 ± 0.14 mL or maximum H2 5.73 ± 0.69 mL/mg cells	[23]
Synechocystis sp. PCC 6803	BG11 medium, 28 °C, 70 μE/m2/s, 97% air 3% CO2, Glass tube (400 mL)	BG11 medium, 28 °C, 70 μE/m2/s, use of oxygen absorber to create anaerobiosis	3–5 mL/g/day (direct biophotolysis)	Faraloni et al. (unpublished)
Fischerella muscicola	BG11 medium, pH 7.5, bubbling with air, 50 μE/m2/s, 30 °C	Sulfur- and nitrogen-free BG11 medium (BG110-s medium) 0.1% glucose, under Ar, optimal pH 7.5, 25 °C, 250 μE/m2/s, 20 mL vial	0.35 ± 0.08 mmol/g(Chl)/h	[24]
Nostoc calcicola	BG11 medium, pH 7.5, bubbling with air, 50 μE/m2/s, 30 °C	Nitrate-free BG11 medium (BG110 medium) 0.1% glucose, under Ar, optimal pH 7.5, 25 °C, 250 μE/m2/s 20 mL vial	0.09 ± 0.01 mmol/g(Chl)/h	[24]
Scytonema bohneri	BG11 medium, pH 7.5, bubbling with air, 50 μE/m2/s, 30 °C	Nitrate-free BG11 medium (BG110 medium) 0.3% glucose, under Ar, optimal pH 7.5, 25 °C, 250 μE/m2/s, 20 mL vial	0.09 ± 0.01 mmol/mg(Chl)/h	[24]
Tolypothrix distorta	BG11 medium, pH 7.5, bubbling with air, 50 μE/m2/s, 30 °C	Nitrate-free BG11 medium (BG110 medium) 0.1% glucose, under Ar, optimal pH 7.5, 25 °C, 250 μE/m2/s, 20 mL vial	0.21 ± 0.05 mmol/mg(Chl)/h	[24]
Geitlerinema sp. RMK-SH10	ASN III medium, 30 μE/m2/s, 30 °C Nitrogen-free ASN IIIeN medium 0.2 M NaCl, 18.9 mmol C-atom/L glucose 0.1 mM Ni2, 30 °C	Under dark and Ar atmosphere, a 10 mL gas-tight vial	0.271 mmol/h/mg dry wt	[25]
Anabaena sp. PCC 7120	DHup 8-times diluted Allen and Arnon medium (AA/8), 30 μE/m2/s, 27 °C, 99% air 1% CO2	A total of 8-times diluted Allen and Arnon medium without nitrogen (AA/8-N), 290 and 340 μE/m2/s, Ar with 5% CO2, and 3.3% N2, pH 8.2, 22 °C, plastic bags (1 L)	0.86 mL/h/L or 20.6 mL/day/L or 33.2 mL/L/during 5 days	[26]

Table 3. H₂ production rate by various cyanobacteria.

Cyanobacteria	Growth Conditions	H2 Production Conditions	H2 Production	References
Immobilized Synechocystis PCC 6803	BG11 medium, 28 °C, 70 μE/m2/s, 97% air 3% CO2, Glass tube (400 mL)	Nitrogen-free medium BG110-Tris, 60 μE/m2/s, 28 °C, under N2 atmosphere, serum vials (60 mL)	Total H2 5.80 ± 0.14 mL or maximum H2 5.73 ± 0.69 mL/mg cells	[23]
Synechocystis sp. PCC 6803	BG11 medium, 28 °C, 70 μE/m2/s, 97% air 3% CO2, Glass tube (400 mL)	BG11 medium, 28 °C, 70 μE/m2/s, use of oxygen absorber to create anaerobiosis	3–5 mL/g/day (direct biophotolysis)	Faraloni et al. (unpublished)
Fischerella muscicola	BG11 medium, pH 7.5, bubbling with air, 50 μE/m2/s, 30 °C	Sulfur- and nitrogen-free BG11 medium (BG110-s medium) 0.1% glucose, under Ar, optimal pH 7.5, 25 °C, 250 μE/m2/s, 20 mL vial	0.35 ± 0.08 mmol/g(Chl)/h	[24]
Nostoc calcicola	BG11 medium, pH 7.5, bubbling with air, 50 μE/m2/s, 30 °C	Nitrate-free BG11 medium (BG110 medium) 0.1% glucose, under Ar, optimal pH 7.5, 25 °C, 250 μE/m2/s 20 mL vial	0.09 ± 0.01 mmol/g(Chl)/h	[24]
Scytonema bohneri	BG11 medium, pH 7.5, bubbling with air, 50 μE/m2/s, 30 °C	Nitrate-free BG11 medium (BG110 medium) 0.3% glucose, under Ar, optimal pH 7.5, 25 °C, 250 μE/m2/s, 20 mL vial	0.09 ± 0.01 mmol/mg(Chl)/h	[24]
Tolypothrix distorta	BG11 medium, pH 7.5, bubbling with air, 50 μE/m2/s, 30 °C	Nitrate-free BG11 medium (BG110 medium) 0.1% glucose, under Ar, optimal pH 7.5, 25 °C, 250 μE/m2/s, 20 mL vial	0.21 ± 0.05 mmol/mg(Chl)/h	[24]
Geitlerinema sp. RMK-SH10	ASN III medium, 30 μE/m2/s, 30 °C Nitrogen-free ASN IIIeN medium 0.2 M NaCl, 18.9 mmol C-atom/L glucose 0.1 mM Ni2, 30 °C	Under dark and Ar atmosphere, a 10 mL gas-tight vial	0.271 mmol/h/mg dry wt	[25]
Anabaena sp. PCC 7120	DHup 8-times diluted Allen and Arnon medium (AA/8), 30 μE/m2/s, 27 °C, 99% air 1% CO2	A total of 8-times diluted Allen and Arnon medium without nitrogen (AA/8-N), 290 and 340 μE/m2/s, Ar with 5% CO2, and 3.3% N2, pH 8.2, 22 °C, plastic bags (1 L)	0.86 mL/h/L or 20.6 mL/day/L or 33.2 mL/L/during 5 days	[26]

1.2. Enzymes

Enzyme hydrogenase allows H₂ photoproduction by catalyzing a reversible conversion between H₂ and H⁺. The active site of this enzyme contains one or two metal ions (Fe or Ni) that work together to produce and consume H₂ during the catalytic cycle.

Hydrogenase enzymes are divided into three different types according to the following metal groups: [FeFe]-hydrogenase (in algae and other eukaryotes), [NiFe]-, and [Fe]-hydrogenase (in bacteria and cyanobacteria).

[FeFe]- and [NiFe]-hydrogenases are found in numerous microorganisms and are divided into multiple subclasses, whereas [Fe]-hydrogenases are limited to methanogens [27]. Although all [NiFe]-hydrogenases can be reactivated without O₂, and some, referred to as O₂-tolerant enzymes, can operate at normal O₂ levels, all identified [FeFe]-enzymes are O₂-sensitive and require resynthesis of the enzyme or at least the metal cluster of the active site to regain activity [28,29].

Microalgal [FeFe] hydrogenases generate H₂ in vitro at rates close to 10⁴ H₂/s, which is more than ten times higher than the typical rates observed for cyanobacterial [NiFe] hydrogenases (1000–2000 H₂/s) [30]. Due to the limitations of the electron input, the in vivo rates are reduced. However, the [FeFe]-hydrogenases are more efficient in H₂ production as their turnover rate is higher than that of the cyanobacterial [NiFe]-hydrogenases. These two classes of enzymes are bidirectional, but H₂ oxidation is more active in the [NiFe]-hydrogenase, and these factors contribute to the highest H₂ production efficiency of [FeFe]-hydrogenases [31].

The cyanobacterial enzyme [Ni-Fe] hydrogenase can reach 640 μmol H₂/mg Chl/h when exposed to a light intensity of about 800 μE for about 10 s. In general, instantaneous rates of 100–200 μmol H₂/mg Chl/h are observed in algal strains, although in some cases, rates up to 450 μmol H₂/mg Chl/h have been measured [31]. Additionally, this peak rate is sustained for roughly ten seconds. It is also important to note that algal cells contain a higher concentration of chlorophyll.

Cyanobacteria possess an additional enzyme, nitrogenase, which operates as a hydrogenase. Nitrogenases convert dinitrogen (N₂) into ammonia and produce an H₂ molecule during each catalytic cycle [32]. When the nitrogen and O₂ sources are removed from the cultures, or when the uptake of hydrogenase, which eliminates the reducing power of H₂ for the cells, is inhibited, some cyanobacterial strains can produce H₂ for prolonged periods of time. The nitrogenase process consumes two molecules of ATP. To decrease the potential to the point where N₂ is diminished for each electron moved, a significant amount of energy is required to generate H₂ (specifically, 16 ATP for each H₂ in the standard case or potentially only 4 ATP if an inert gas is used instead of N₂). In vivo generation of photosystem–hydrogenase fusions could enhance these strategies, and since semi-artificial systems can easily differentiate between H₂-producing PSI and O₂-generating PSII in separate half-cells, both approaches could benefit from each other. A summary of the characteristics, mechanism of functioning, advantages, and disadvantages can be found in

Table 4. Enzymes involved in H₂ production.

Enzyme Type	Characteristics	H2 Production Mechanism	Advantages	Disadvantages	References
[FeFe]-Hydrogenase	Found in microalgae; highly active but O2-sensitive	Catalyzes H2 production at rates of ~104 H2/s in vitro	High efficiency in H2 production	Highly sensitive to O2, requiring anaerobic conditions	[30]
[NiFe]-Hydrogenase	Found in cyanobacteria; more resistant to O2 exposure	Lower H2 production rates (~1000–2000 H2/s)	Some variants can function in oxygenated environments	Lower catalytic efficiency compared to [FeFe]-hydrogenase	[28]
Nitrogenase	Converts N₂ into ammonia, producing H2 as a by-product	Requires ATP (16 ATP per H2 with N2, 4 ATP without N2)	Can produce H2 for longer periods in anaerobic conditions	High energy cost due to ATP consumption	[32]

.

2. Strategies to Optimize H₂ Production

Different strategies can be considered to optimize the sustainability of the entire H₂ production process (

Table 5. Optimization strategies for H₂ production.

Optimization Strategy	Description	Advantages	Challenges	References
Genetic Engineering	Modifying microalgae/cyanobacteria for improved H2 output	Can enhance enzyme efficiency and O2 tolerance	Ethical concerns, regulatory restrictions	[33,34]
Co-Cultivation with Bacteria	Using bacteria to assist H2 production (e.g., dark fermentation)	Converts biomass into additional H2	Requires controlled symbiotic relationships	[35,36]
Nutrient Starvation	Limiting sulfur, nitrogen, or potassium to enhance H2 yield	Enhances hydrogenase activity, reduces O2 production	Can stress cells, lowering overall biomass production	[19]
Wastewater Utilization	Using wastewater as a nutrient source for microalgae	Reduces costs, reuses waste, increases biomass	Variability in wastewater composition	[37,38]
Light Optimization	Using specific wavelengths (e.g., purple light)	Increases photosynthetic efficiency and H2 production	Requires precise control of light exposure	[39,40]

).

2.1. Genetic Engineering

Two main strategies to increase H₂ production are metabolic engineering and genetic engineering [34]. Metabolic engineering can improve H₂ production by optimizing microbial metabolic pathways and introducing new functionalities. Metabolic engineering includes gene overexpression and knockout, heterologous gene expression, regulatory protein manipulation, carbon source optimization, and in vitro pathway construction. Studies have been carried out to investigate the possible effects of genetic engineering on particular metabolic pathways to enhance the H₂ production process. These studies mainly focus on the efficiency of electron transport from ferredoxin to hydrogenase enzymes. The development of commercially viable systems for H₂ production from microalgae is significantly hampered by the sensitivity to O₂ of the enzymes. Yang et al. genetically modified the hydrogenase diaphorase A (HYDA) of Chlorella sp. to alter the activity of the enzyme [33]. These changes were made primarily to prevent O₂ from entering the active site, thereby increasing the amount of H₂ produced by biophotocatalysis. The results showed a significant increase in H₂ production, which can be up to thirty times higher than that of the wild type when exposed to O₂.

In recent years, significant progress has been made in both the cyanobacterium Synechocystis PCC 6803 and the microalga Chlamydomonas reinhardtii, enabling biological H₂ production from sunlight and water while avoiding nutrient starvation. In these organisms, hydrogenases were genetically fused with PSI to capture electrons for H₂ production. H₂ production proved to be light-driven in both the engineered cyanobacterial strain (PsaD–HoxYH) and the engineered algal strain (ΨH1). The cyanobacterial NiFe-hydrogenase HoxYH was fused to the PSI subunit PsaD, while the algal FeFe-hydrogenase HydA2 was inserted into the PSI subunit PsaC [41]. The fusion of hydrogenase with PSI allows hydrogenase to compete with the Calvin–Benson–Bassham cycle, the main competitor of hydrogenase that oxidizes NADPH provided via ferredoxin–NADP+ reductase (FNR). FNR likely has a competitive advantage over other downstream electron acceptors because it binds to the PSI acceptor site [42]. Both engineered strains produce H₂ for several hours under anaerobic conditions (PSI–HoxYH strain: 0.6 μmol H₂/mg Chl/h; ΨH1: 21 μmol H₂/mg Chl/h). In WT green algae and cyanobacteria, the onset of CO₂ fixation terminates H₂ production within several minutes. However, ΨH1 continues to produce photobiological H₂ due to the diversion of most electrons to H₂ production away from CO₂ fixation. In both cases, the PSI–hydrogenase chimera that replaces normal PSI is still capable of photoautotrophic growth, albeit at low rates [41]. Recently, it was proposed that diverting 50% of the electron transfer rate (ETR) in Synechocystis to carbohydrate synthesis to support respiration for the creation of anaerobic conditions would result in a reduction in light conversion efficiency (LCE) from the theoretical maximum of 13.4% to 8.7% (light to H₂) [43]. This value remains more than eight times higher than what can be achieved with conventional sulfur starvation methods [44]. This hypothesis is based on the assumption that four electrons are needed to produce one molecule of H₂, while six electrons per carbon atom are required to synthesize one molecule of glucose (C₆H₁₂O₆). To achieve this balance, the respiration rate must be high enough to metabolize the increased amount of carbohydrates and maintain anaerobiosis. Our investigations indicate that the respiration rate is strongly dependent on light irradiance and salinity (Torzillo, unpublished). For example, cells of Synechocystis PCC 6803 grown under 100 to 200 µmol photons/m²/s show respiration rates of 35 to 80 µmol O₂/mg chlorophyll a (Chla)/h.

2.2. Co-Cultivation with Bacteria

Microalgae–bacteria consortia offer potential advantages for H₂ production. Co-cultures can increase H₂ synthesis and produce changes in yield, rate, and duration compared to monocultures. Co-cultivation of C. reinhardtii with isolated bacteria can increase the production of H₂. For example, compared to monocultures, the co-culture of Rhodopseudomonas palustris and C. vulgaris enhanced H₂ production by 125% [36]. With C. reinhardtii, a 5.3-fold increase was observed, and when Azotobacter chroococcum and Bradyrhizobium japonicum were used, a 14-fold increase was reached [45,46]. Co-cultivation of C. reinhardtii with Escherichia coli, Pseudomonas stutzeri, and Pseudomonas putida increased H₂ production by up to 24%, 46%, and 32%, respectively [47]. In addition to Chlamydomonas, photobiological H₂ production was also promoted in other microalgae–bacteria co-cultures, namely, Chlorella protothecoides, Chlorella pyrenoidosa, and Scenedesmus obliquus, all of which were individually paired with the bacterial species Pseudomonas sp. [48]. However, it must be pointed out that the co-cultivation of microalgae with bacteria is only possible with mineral media, in which the growth of bacteria is limited by the lack of organic substrate. Instead, in the presence of supplemented organic substrates, it can be risky for the bacteria to accelerate the process toward anaerobic fermentation with a consequent rapid drop in the pH and a predominant production of volatile acids such as lactic and acetic acid.

2.3. Use of Wastewater

The large amount of organic compounds contained in wastewater (WW) from various sources increases the potential to use this WW as a substrate for the growth of microalgae. It is interesting to note that the absorption of sugars from WW can lead to a more carbohydrate-rich microalgal biomass, as has been observed in Chlorella [37]. This microalga has shown a variety of applications and a high degree of adaptability. In some studies, C. vulgaris var. TISTR 8261 was cultured with WW from the frozen food sector. This strain removed phosphate, sulfate, nitrate, and nitrite from WW, and it generated more H₂ in WW with sodium acetate than in the control medium [37]. In the sector of H₂ production, numerous sources for the use of WW have been examined [49]. The growth and H₂ production of immobilized C. vulgaris and Scenedesmus obliquus in urban WW were investigated [19]. Sulfur deprivation and different light conditions were combined before anaerobiosis. Purple light generated the highest H₂ output (128 mL H₂/L for S. obliquus and 60.4 mL H₂/L for C. vulgaris, respectively), while blue light generated the highest growth. It was also found that a considerable amount of organic carbon was eliminated, leading to more reliable results for S. obliquus [19]. These two species have also been used together in the past to treat urban WW. The naturally occurring strains of C. vulgaris and Scenedesmus obliquus coexisted in urban WW. To induce the accumulation of carbohydrates, which ranged from 22% to 43%, they were able to eliminate the organic compounds present in the WW and were maintained on a restricted diet for a longer period of time. The biomass was then converted to H₂ by the bacterium Enterobacter aerogenes through dark fermentation. The amount of biogas produced was comparable to the amount produced with biomass grown in a conventional growth medium. This is an interesting and novel way to utilize microalgae biomass for H₂ production, even though it is an indirect method for H₂ production from microalgae. The effect of olive mill wastewater (OMW) on biomass production and composition was investigated using C. reinhardtii and a traditional growth medium [38]. OMW was only used in the initial stages of microalgal development, which produced biomass with a high carbohydrate content. These sulfur-starved cells produced a larger volume of biogas than the control. This was associated with a higher initial production rate, which is often associated with high H₂ evolution [50].

2.4. Light Utilization

Several studies have shown that sudden light can generate H₂, which increases H₂ production [39]. The effects of alternating photoperiods on the development of C. reinhardtii under light-driven, sulfur-deficient conditions were investigated [40]. The authors showed that the highest H₂ production was achieved during the first 7–10 days, with the highest production in the first 5 days.

2.5. Challenges in Scaling Up of Photobiological H₂ Production

A very important part of a successful solar H₂ production process relies on the development of an economically feasible PBR [51]. Although there is not much consensus in the literature on the cost of PBR, it is believed that the materials for the construction of the bioreactor account for about 35% of the production cost, which increases up to 63% when the nutrients for the preparation of the medium and the CO₂ supply are included [52]. The most important factors for successful direct photobiological H₂ production in large-scale outdoor PBRs include a PBR design in which it is possible to achieve efficient LCE, the use of an efficient mixing system, and adequate monitoring and control of culture parameters [44]. In the case of PBR for H₂ production, the design becomes more complex as the PBR must be perfectly sealed to avoid H₂ losses to the atmosphere. Another problem with H₂ production by direct biophotolysis (DBP) is the risk of explosion caused by the simultaneous release of H₂ and O₂ from the PSII water splitting system. This problem is substantially avoided in PBRs operated with microalgae exposed to nutrient starvation, by using the Melis protocol, which is based on the down-regulation of PSII activity to balance O₂ release by photosynthesis and O₂ by respiration, to create anaerobic conditions required for hydrogenase enzyme activation [53]. This protocol has the advantage of producing nearly pure H₂, which reduces the cost of its purification. Unfortunately, the use of nutrient starvation is far from economical, as the efficiency of solar to hydrogen (STH) barely reaches 1%. This is 1/13 of the efficiency that can theoretically be achieved by using DBP, assuming that a microalgal/cyanobacterium strain with a hydrogenase resistant to O₂ is achieved through genetic modification [43,44].

To fully exploit the LCE of the organism to be cultivated for outdoor H₂ production, the problem of the “light saturation” of photosynthesis, which is a strain-dependent property, must be avoided. In other words, the photosynthetic activity of the cells must remain in the linear part of the light vs. photosynthesis curve. This objective is very difficult to achieve outdoors, as light irradiance can reach values as much as 10 times higher than that required for photosynthesis saturation. To reduce this problem, PBRs with an increased surface area in relation to their footprint are preferred. This can be achieved by greatly increasing the cross-section of the reactor (i.e., the illuminated part of the reactor) with respect to the ground area it occupies. In this way, it is possible to reduce the “saturation effect” of photosynthesis by taking advantage of the so-called “light dilution effect”. Recently, a multiplate PBR of 1.3 m³ was tested outdoors for the growth of Synechocystis. PCC 6803. The authors concluded that the LCE calculated based on the ground area occupied by the reactor was significantly higher in the configuration with 0.5 m plate spacing, reaching 5.43% based on photosynthetically active radiation and 2.44% based on solar radiation, giving a 3.7-fold higher value than with 1.0 m spacing between plates [44,54].

3. Potential Use of Microalgae for H₂ Production as By-Products

Exhausted biomass from H₂-producing microalgal cultures can be a valuable source of other products. For instance, Chlamydomonas biomass can be used for pyrolysis, which can generate additional energy but is not very efficient when recovered after H₂ production. Chlamydomonas biomass can also be processed into charcoal and used as fertilizer [55]. Furthermore, these cells can be helpful as a natural antioxidant supply after sulfur deprivation and after H₂ synthesis is complete, as they are rich in carotenoids such as lutein, antheraxanthin, zeaxanthin, and β-carotene [8]. Chlorella biomass can be recycled for various purposes, including hydrolysis of algal biomass to produce fermentable sugars. A common natural microalga used as a dietary supplement is C. vulgaris, which has a high biomass concentration of proteins and chlorophyll. A significant amount of lutein, β-carotene, and zeaxanthin (5.3 mg/g, 0.31 mg/g, and 0.24 mg/g, respectively) with high bioavailability has also been demonstrated in this microalga [56,57]. Although microalgae have a balanced profile of essential amino acids, their digestibility may not be as good as traditional protein sources [58]. In this context, in vitro tests on the digestibility of various microalgae species were performed. The findings showed a broad range of protein digestibility rates, ranging from 50 to 82% depending on the species studied [59,60]. In the case of C. vulgaris, the digestibility of crude protein was in the high range of 76%. According to Niccolai et al., this microalga has a higher digestibility than some common protein sources, such as wheat, beans, and oats (78%, 72%, and 77%, respectively) [59]. Along with nutraceuticals, proteins derived from microalgae could also have technical applications in agriculture and the food sector. For instance, C. vulgaris would have the ability to gel, thicken, foam, or emulsify [58]. Because of their attractive biological properties, protein extracts are frequently used in the formulation of cosmetic products. Peptides derived from microalgae have garnered more attention due to their safety as substitutes for bioactive compounds in pharmacology. They are known for their anti-inflammatory, antihypertensive, anticancer, antibacterial, and antioxidant properties [61,62]. These properties can be utilized to improve human health.

Synechocystis is one of the cyanobacteria that can accumulate a significant amount of phycocyanin. Both heme and phycocyanin are present in the bacterium Synechocystis PCC 6803, which can also use heme as a precursor to produce phycocyanin. To reduce carcinogenic effects and ensure food safety, both heme and phycocyanin are effective natural food colorants that could be used instead of traditional colorants such as nitrite and synthetic colorants [63]. In addition, heme could be used in the development of effective anticancer drugs and in the diagnosis of diseases due to its unique photosensitive properties and therapeutic benefits. Phycocyanin has shown strong anticancer, antidiabetic, anti-inflammatory, antioxidant, and immunomodulatory effects [20,64] and has great potential as a new and effective drug for treating a variety of diseases, such as COVID-19 complications, atherosclerosis, multiple sclerosis, and ischemic stroke [65,66]. Phycocyanin has been shown to have a positive effect on the gut, reproductive system, liver, kidney, and brain [20,67,68,69,70]. Moreover, in vitro research has shown that phycocyanin peptides can reduce oxidative and inflammatory cell damage and thereby attenuate pulmonary fibrosis [20]. The biomass of Synechocystis can be a valuable source of proteins rich in essential amino acids [71]. Figure 2 shows the amino acid profile of Synechocystis biomass.

The essential amino acid profile of Synechocystis biomass is compared to that of the well-known edible cyanobacteria Arthrospira platensis, which is currently cultivated worldwide on an industrial scale as a source of protein and phycocyanin. Sulfur-containing amino acids are hardly present in Synechocystis compared to Arthrospira biomass, but in contrast, the presence of lysine is striking, which is much higher than in Arthrospira biomass. Lysine promotes the formation of antibodies, hormones (e.g., growth hormone), and enzymes; it is also necessary for the development and binding of calcium in the bones. Lysine is also important as a precursor of an important vitamin, namely, niacin, vitamin B3, or PP. Therefore, a mixture of both biomasses could lead to a synergistic nutritional effect.

4. Solar H₂ Production Cost

A detailed cost analysis of H₂ production by different STH technologies was recently published by Frowijn and van Sark [52]. According to their cost analysis, the H₂ cost for photovoltaic–electrolysis (PVE) is 9.31 $/kg, while it is about twice as high for DBP. This large difference is due to the low actual efficiency of biophotolysis, which is less than one-tenth of the theoretical efficiency (13.4%) [43]. In the case of biophotolysis, most of the cost is related to the construction of the PBR, which together with nutrients accounts for 63% of the total H₂ cost [52]. Therefore, the cost of H₂ production by biophotolysis could become profitable through the development of genetically modified microalgae and, particularly, the cyanobacterium Synechocystis, which is easier to genetically manipulate. This can be achieved by taking advantage of the progress in synthetic biology and optimizing the use of solar LCE by avoiding the loss of light energy that occurs during most hours of the day due to the saturation of photosynthesis [29,72]. In this regard, important progress can be expected by optimizing the design of PBR, i.e., by introducing geometries that can reduce the incident light on the reactor surface. This can be achieved by increasing the reactor cross-section with respect to the ground surface area occupied by the reactor, so that the incident light can be distributed over a larger surface, the so-called light dilution effect. By reducing the incident light on the surface of the reactor, it is possible to reduce the risk of saturation of photosynthesis and thus the loss of energy through heat (non-photochemical quenching). Recently, a novel PBR design for H₂ production has been proposed [54]. However, it must be pointed out that the advantages obtained by using PBRs that can benefit from a high light dilution rate result in a higher cost for the PBR, as the materials for its construction are more expensive. Therefore, a cost–benefit analysis is required to find the best compromise between the increase in LCE and the cost of the PBR. Figure 3 compares the H₂ production costs in two different situations resulting from the optimization of the LCE in PVE and DBP.

Under the present scenario (blue lines), H₂ production costs are higher (particularly for DBP) as they are obtained with the present STH (solar to H₂) efficiency for both PVE and DBP, while in the prospected scenario (red line), H₂ production costs decrease as the calculation considers the highest potential STH efficiency for each STH method. It is interesting to note that PVE would no longer be the cheapest option. The cheaper option of H₂ production could result in future DBP, which is expected to have a larger gain in terms of LCE compared to PVE [52]. In other words, the scope for increasing the LCE for DBP to the maximum value (13.4%) is significantly greater than for PVE.

5. Challenges and Perspectives

H₂ is essential for the realization of a sustainable society. However, there are two main obstacles to the full use and development of H₂ as part of the energy transition. The first is that fossil fuels are still strongly subsidized, making it difficult for renewables to penetrate the energy markets. Second, renewable energy sources such as wind and solar energy, i.e., the two most widely used sources for renewable H₂ production, are still too expensive [52,54,73,74,75]. Among the methods of photochemical water splitting, PVE is the most advanced system for producing H₂ today. In PVE, H₂ is produced by an indirect method using photovoltaic electricity that powers an electrolyzer. The most common water electrolysis processes are the alkaline electrolyzer and the polymer electrolyte membrane or proton exchange membrane (PEM) electrolyzer [52,76]. DBP, also known simply as biophotolysis, has in common with electrolysis the use of sunlight captured by the photosynthetic apparatus to split water by two nanomachines, the PSII and the hydrogenase (see above for more details). Therefore, both technologies use catalysts that are artificial in the case of electrolysis and natural in the case of biophotolysis. Although electrolysis and biophotolysis are two processes with very different TRLs, close to nine for the first technology and close to four for biophotolysis, a comparison of the two technologies can be very revealing.

Table 6. Comparison between solar to H₂ efficiency of PV-alkaline electrolysis and their actual readiness transfer level (TRL).

Solar to H2 Technology	Solar to H2 Efficiency	Material-Catalysts	TRL	Reference
PV Alkaline Electrolysis	10–12.3%	Material: Perovskite + Alkaline Catalysts: a multilayer anode nickel–iron hydroxide (NiFe) electrocatalyst layer coated on a nickel sulfide (NiSx) layer formed on porous Ni foam (NiFe/NiSx-Ni).	9	[52]
Direct Biophotolysis	1–13.4%	Microalgae: C. reinhardtii; C. reinhardtii D1 mutants; Chlorella sp G-120.; Cyanobacteria; Synechocystis 6803; Synnechocystis mutants Catalysts: Hydrogenases	4–5	[11,43]

summarizes the main characteristics of PVE and DBP when using selected strains of microalgae and cyanobacteria. As indicated in Table 6, the theoretical STH efficiency achieved by biophotolysis compares well with that of water electrolysis. Both technologies achieve efficiencies that are very close to each other (12–13% STH). While these efficiencies are achieved with PVE, DBP still has a long way to go before it will play an important role in the energy transition [52].

It is worth noting that DBP is considered a more sustainable process since it does not require artificial catalysts such as noble metals, rarely found in nature, which could lead to supply problems in countries with low deposits of such important minerals [29,77,78]. Moreover, the two processes also differ in terms of the use of the water source, freshwater in the case of electrolysis and seawater in the case of biophotolysis, which may also rely on the use of wastewater (see Section 2.3). To close the gap in STH efficiency, DBP needs to overcome two important limitations: First, the generation of a hydrogenase that is resistant to the high O₂ produced by the PSII water splitting machinery, which can currently only be circumvented by down-regulating PSII through nutrient limitation. Second, the efficient utilization of solar light irradiance throughout the day to avoid energy loss through non-photochemical quenching, particularly in the middle of the day. Both the inhibition of hydrogenase by O₂ and the increase in STH efficiency are currently the subject of intensive research [44]. To this end, the cyanobacterium Synechocystis PCC 6803 is being extensively subjected to targeted genetic manipulations and metabolic flux adjustments to improve H₂ photoproduction [44].

6. Purple Non-Sulfur Photosynthetic Bacteria for Photofermentation

Purple non-sulfur photosynthetic bacteria (PNSB) are gram-negative bacteria that grow under anaerobic or microaerobic conditions in the form of red colonies. They are often used as model organisms to study the metabolic control of nitrogen and carbon metabolism and can grow in many modes and environments such as aquatic and terrestrial habitats [79]. These bacteria can thrive in a variety of ecological niches, from marine and freshwater to sediments and extreme environments such as soda or hypersaline lakes, cold waters, and hot springs [80]. Thanks to their metabolic versatility, PNSB can inhabit predominantly anoxic niches, but they can also be found in microaerobic or even aerobic conditions. PNSB are anoxygenic photoautotrophs capable of light-dependent CO₂ fixation and phototrophic energy transfer, with bacteriochlorophyll a or b and carotenoids being the major pigments. Depending on the levels of light, carbon, and O₂, they can develop as photoautotrophs (in the presence of light and CO₂), photo-heterotrophs (in the presence of light and organic compounds), or chemo-heterotrophs (in the presence of organic compounds). They have no O₂-generating activity and absorb solar energy in the visible and near-infrared wavelength range. PNSB possess a high substrate conversion efficiency and can use a variety of organic carbon compounds such as pyruvate, acetate, and other organic acids (e.g., butyrate, propionate, malate, and lactate), amino acids, alcohols, and carbohydrates [81]. Aromatic organic compounds can also be used by PNSB as carbon sources [82]. PNSB have the following metabolic pathways: the Tricarboxylic acid cycle, the Calvin–Benson cycle, the Glyoxylic acid cycle, and the Embden–Meyerhof–Parnas pathway. Photophosphorylation and oxidative phosphorylation are the two primary energy metabolisms, both of which are associated with proton transmembrane transfer and electron transport [83]. Their metabolic plasticity makes PNSB attractive for many biotechnological applications. PNSB convert anaerobically organic substrates into molecular H₂ in a process known as photofermentation [84]. For example, Rhodobacter sphaeroides, Rhodopseudomonas palustris, Rhodospirillum rubrum, and Rhodovulum sulfidophilum can utilize organic acids for H₂ production with the help of the enzyme nitrogenase [85,86,87,88]. Organic compounds supply the photosystem (PS) of PNSB with electrons. These are energized using light and channeled through an H₂-pumping electron transport chain, which is moved from the quinone pool through the cytochrome c oxidoreductase (cyt bc1) complex to oxidized ferredoxin (Fd) (Figure 4). The proton motive force generated during this process is used to synthesize adenosine triphosphate (ATP). The electrons of the reduced Fd are transferred to the nitrogenase, which creates H₂ as the only product. To generate H₂, the ATP produced by the photosynthetic process is transferred to the nitrogenase together with protons and electrons. O₂-free and ammonia-limited conditions are required for photofermentative H₂ production, as nitrogenase is inhibited in the presence of O₂ and ammonium salts. Under these conditions, nitrogenase catalyzes proton reduction to generate H₂ at the expense of four moles of ATP, according to the following equation: 8H⁺ + 8e⁻ + 16ATP → 4H₂ + 16ADP + 16P_i. The most used nitrogen source for photofermentative H₂ is glutamate, as it exhibits low nitrogenase inhibition and is rapidly consumed [89,90,91].

A variety of photosynthetic bacteria such as Rhodobacter sphaeroides and Rhodopseudomonas palustris, as well as organic acids such as acetate and lactate, have been used for photofermentation [92,93]. Numerous articles have discussed the production of photofermentative H₂ in the field and in the laboratory [94,95,96]. Photofermentative H₂ production depends on the conditions of cell growth, such as the composition of the growth medium and light, temperature, and pH conditions [95]. The photofermentation process can be enhanced by the addition of chemicals to the growth medium, such as nitrogenase co-factors (e.g., iron, molybdenum, and nickel), ethylenediaminetetraacetic acid (which increases the solubility and availability of metals), and yeast extract [97].

Photosynthetic bacteria have been investigated for the synthesis of H₂ using wastewater, single organic acids, or mixtures of organic acids as carbon sources [90,93,98,99]. The type of carbon source affects the efficiency of H₂ production, which is due to the different electron transfer capabilities of photosynthetic bacteria that differ in their metabolic pathway. Waste materials can be used to feed photosynthetic bacteria, enhancing the cost-effectiveness of the photofermentative process by simultaneously producing H₂ and utilizing waste. The optimal waste sources for H₂ production by photofermentation are those rich in organic acids [81]. Several WW and industrial wastes have already been tested as substrates, such as palm oil mill effluent [97], molasses [100], cheese whey [101], brewery wastewater [102], winery wastewater [103], and food waste [104]. One strategy to increase H₂ yield is based on the co-cultivation of biological systems to optimize the conversion of organic material. The substrates used by the dark fermentative bacteria are converted into volatile fatty acids, which are metabolized by PNSB and lead to the production of H₂. This process can be performed as a two-phase method or alternatively with the integration of mixed cultures, which could lead to positive results in terms of both productivity and economics [105]. Several studies have investigated co-cultures of purple photosynthetic bacteria and dark fermentative bacteria for H₂ production [106,107,108]. Table 7 summarizes recent works on photofermentative H₂ production by photosynthetic bacteria.

Optimization of the Photofermentation Process with Immobilization

Suspension cultures are most used for photofermentative H₂ production [95,109]. The immobilization process is a useful tool in biotechnology because it can improve system stability and enable continuous batch operation for long-term use [99,110,111]. Compared to suspension culture, cell immobilization offers several advantages that can reduce system costs. These advantages include improved stability and resistance to mechanical stress, increased cell biomass, nutritional supplementation without the need to harvest cells, and reduced risk of cell contamination. In addition, immobilization provides a much more stable microenvironment for the cells. The repeated use of the same material over a period of time is one of the great advantages of immobilization. Immobilization, a technique extensively used for photosynthetic bacterial cells, could be used to stabilize and improve photofermentative H₂ production [91,111,112,113,114]. Photofermentative H₂ production by immobilized photosynthetic bacteria has been the subject of numerous studies (Table 8). In Figure 5, PBRs with a suspended (left) and an immobilized (right) culture of photosynthetic bacteria are reported.

Entrapment, biofilms, adsorption, and encapsulation are common immobilization strategies used in photofermentation, allowing high yields and reusability [91,99,114,115]. Photosynthetic bacterial cells have been immobilized with materials such as glass beads, alginate, agar, chitosan hydrogels, and polyvinyl alcohol cryogels [116,117]. Alginates are unbranched polysaccharides obtained from brown algae and consist of residues of b-D-mannuronic acid and (1-4)-linked a-L-guluronic acid in varying amounts [91,114]. Ion exchange between divalent cations such as Ca²⁺ and Na⁺ from guluronic acid salts causes the alginate to gel. Transparency, a gentle encapsulation process, a relatively inert aqueous environment within the gel, a high porosity that allows gas and nutrient exchange between the gel and its environment, and the reversibility of the gelation process are some of these properties. Immobilized Rhodopseudomonas palustris in sodium alginate and in PVA cryogel produced a higher amount of H₂ than the suspended photofermentation [113]. The high biomass concentration in suspended cultures can increase the effects of mutual shading, which is attenuated by the immobilized cells. Researchers reported the feasibility of sequential microaerobic dark- and photofermentative H₂ production by Rhodobacter capsulatus immobilized in agar [118]. Despite the high H₂ production by immobilized cultures, some factors that could affect these systems were reported by Sagir and Alipour [114]: (i) the porosity, texture, pore size, and thickness of the immobilized matrix affect the transport and consumption of substrates by bacteria; (ii) substrates, as utilization and conversion efficiency differ among them; (iii) pH and temperature, which affect enzyme activities; (iv) light source and intensity; (v) the mode of operation and the type of processes (e.g., single-stage, two-stage, sequential, or combined), which affect the efficiency and cost of the system. Overall, immobilization approaches help to overcome the challenges of bacterial cell density to achieve higher H₂ yield, synthesis, and rate. This could improve the economic viability of bio-H₂ production [95].

Table 7. Photofermentative H₂ production by photosynthetic bacteria.

PBR Type (Volume)	Strain	Carbon Source (g/L)	Rate (mL/L/h)	Reference
Schott bottle (100 mL)	R. sphaeroides NCIMB8253	Palm Oil Mill Effluent/pulp and paper mill effluent	64.9	[97]
Serological bottles (120 mL)	R. capsulatus ATCC 17015	Acetate (1.0), Butyrate (11.6), Propionate (1.76)	19.67	[119]
Cylindrical glass (220 mL)	Rhodopseudomonas sp.	Acetate (4.0)	19.6	[90]
Glass bottle (260 mL)	R. palustris PB-Z	Glucose (12.6)	78.7	[120]
Flat panel (4.0 L)	R. capsulatus hup-	Sugar Beet Thick Juice	25.01	[121]
Tubular (50 L)	R. palustris 42OL	Malate (4.0)	27.2	[122]
Tubular (70 L)	R. sphaeroides HY01	Glucose (5.4)	37.6	[123]
Tubular (20 L)	R. capsulatus YO3	Molasses	15.45	[100]

Table 7. Photofermentative H₂ production by photosynthetic bacteria.

PBR Type (Volume)	Strain	Carbon Source (g/L)	Rate (mL/L/h)	Reference
Schott bottle (100 mL)	R. sphaeroides NCIMB8253	Palm Oil Mill Effluent/pulp and paper mill effluent	64.9	[97]
Serological bottles (120 mL)	R. capsulatus ATCC 17015	Acetate (1.0), Butyrate (11.6), Propionate (1.76)	19.67	[119]
Cylindrical glass (220 mL)	Rhodopseudomonas sp.	Acetate (4.0)	19.6	[90]
Glass bottle (260 mL)	R. palustris PB-Z	Glucose (12.6)	78.7	[120]
Flat panel (4.0 L)	R. capsulatus hup-	Sugar Beet Thick Juice	25.01	[121]
Tubular (50 L)	R. palustris 42OL	Malate (4.0)	27.2	[122]
Tubular (70 L)	R. sphaeroides HY01	Glucose (5.4)	37.6	[123]
Tubular (20 L)	R. capsulatus YO3	Molasses	15.45	[100]

Table 8. Photofermentative H₂ production by immobilized photosynthetic bacteria.

PBR Type (Volume)	Strain	Carbon Source (g/L)	Matrix	Rate (mL/L/h)	Reference
Roux bottle (200 mL)	Rhodobacter capsulatus YO3	Acetate (3.6)	Agar	48.9	[110]
Flat panel (1400 mL)	Rhodobacter capsulatus YO3	Acetate (3.6)	Agar	31.2	[111]
Roux bottle (200 mL)	Rhodobacter capsulatus DSM 1710	Acetate (3.6)	Agar	18.6	[110]
Flat panel (1400 mL)	Rhodobacter capsulatus DSM 1710	Acetate (3.6)	Agar	18.0	[111]
Cylindrical glass bottle (200 mL)	Rhodopseudomonas sp. S16-VOGS3	Acetate (2.0)	Ca-alginate	14.96	[91]
Cylindrical glass bottle (200 mL)	Rhodopseudomonas sp.	Acetate (2.0)	Ca-alginate	10.2	[99]
Flat Roux glass bottle (600 mL)	Rhodopseudomonas sp. S16-VOGS3	Acetate (2.0)	Ca-alginate	2.58	[91]

Table 8. Photofermentative H₂ production by immobilized photosynthetic bacteria.

PBR Type (Volume)	Strain	Carbon Source (g/L)	Matrix	Rate (mL/L/h)	Reference
Roux bottle (200 mL)	Rhodobacter capsulatus YO3	Acetate (3.6)	Agar	48.9	[110]
Flat panel (1400 mL)	Rhodobacter capsulatus YO3	Acetate (3.6)	Agar	31.2	[111]
Roux bottle (200 mL)	Rhodobacter capsulatus DSM 1710	Acetate (3.6)	Agar	18.6	[110]
Flat panel (1400 mL)	Rhodobacter capsulatus DSM 1710	Acetate (3.6)	Agar	18.0	[111]
Cylindrical glass bottle (200 mL)	Rhodopseudomonas sp. S16-VOGS3	Acetate (2.0)	Ca-alginate	14.96	[91]
Cylindrical glass bottle (200 mL)	Rhodopseudomonas sp.	Acetate (2.0)	Ca-alginate	10.2	[99]
Flat Roux glass bottle (600 mL)	Rhodopseudomonas sp. S16-VOGS3	Acetate (2.0)	Ca-alginate	2.58	[91]

7. Conclusions

Green H₂ represents an important opportunity to decarbonize various sectors, particularly transportation and heavy industry. Both biophotolysis and photofermentation could be an option to produce H₂. Direct biophotolysis is particularly attractive as it is a fully environmentally sustainable process. However, as large amounts of water are required for the cultivation of microalgae, this can lead to an increase in production costs. The major water demand occurs when filling the PBR, which requires about 50 L of water per square meter of reactor area, while the H₂ production phase theoretically requires 9 L of water to produce 1 kg of H_2, which is similar to an electrolyzer. However, in both cases, the effective water consumption can reach as much as 20 L per kg of H₂, as water purification and/or process cooling is required. Currently, freshwater from natural sources serves as the primary water supply for H₂ production, particularly for electrolysis. Despite the rapid global expansion of H₂ production through electrolysis, little attention has been paid to the fact that freshwater is the main feedstock for these systems. For both microalgae and photosynthetic bacteria, this problem can be easily solved by using various nutrient-rich wastewater for their growth and cultivation. Some microalgae, in particular the cyanobacterium Synechocystis, have a high metabolic plasticity that allows them to rapidly adapt to environmental stress, including high salinity. This characteristic makes them an ideal organism for H₂ production using seawater and salted water. Moreover, since this organism is relatively easy to genetically manipulate, it may be possible to achieve a strain with high H₂ production efficiency that is also resistant to outdoor environmental stresses, especially high light intensity and high temperatures. These capabilities are extremely important for expanding cultivation to coastal areas and, in general, to areas where conventional agriculture is not economically profitable.

Scalable production systems are made possible by cultivating microalgae in large-scale PBRs using low-cost materials and simple fabrication technologies. The production of H₂ from solar energy without critical raw materials or toxic processes aligns with the EU’s Sustainable Development Goals. By reducing production costs to as low as EUR 5 per kg of H₂, this technology is competitive with current photovoltaic coupled with electrolysis methods. In addition, microalgal cultures can be grown on CO₂-rich biogas effluents from anaerobic digesters, integrating wastewater treatment with H₂ production. This approach not only improves sustainability but also allows the H₂ produced to be used directly to enrich the energy content of the biogas.