Keywords

1 Introduction

Hydrogen gas (H2) is one of the potential future sustainable and clean energy carriers that may substitute the use of fossil resources including fuels since it has a high energy content (heating value of 141.65 MJ/kg) when compared to traditional hydrocarbon fuels [1]. Water is a byproduct of combustion being a most significant advantage of H2 being environmentally friendly with the capacity to reduce global greenhouse gas emissions. H2 is used in various applications. It generates electricity in fuel cells, including applications in transportation, and can be applied as fuel in rocket engines [2]. Moreover, H2 is an important gas and raw material in many industrial applications. However, the high cost of the H2 production processes requiring the use of other energy sources is a significant disadvantage. At present, H2 can be prepared in many conventional ways, such as steam reforming, electrolysis, and biohydrogen production processes. Steam reforming uses high-temperature steam to produce hydrogen gas from fossil resources including natural gas. Electrolysis is an electrolytic process to decompose water molecules into O2 and H2. However, both these two methods are energy-intensive and producing hydrogen from natural gas, which is mostly methane (CH4) and in steam reforming generates CO2 and pollutants as by-products. On the other hand, biological hydrogen production is more environmentally sustainable and less energy intensive than thermochemical and electrochemical processes [3], but most concepts are not yet developed to production scale.

The production of H2 using microorganisms has attracted public interest due to its potential as a renewable energy carrier that can be produced using nature’s plentiful resources. There are various approaches for biological H2 production using microorganisms such as green algae, cyanobacteria, photosynthetic anoxic bacteria, and dark fermentative bacteria. These microorganisms are physiologically very diverse, occupy different ecological niches, and use distinct metabolic pathways generating H2. Cyanobacteria, a group of microorganisms performing an oxygenic photosynthesis, can be utilized for H2 production via biophotolysis [4,5,6,7,8]. They are autotrophic organisms and thereby fix CO2 from the atmosphere as carbon source. In addition, many strains have the capacity to reduce atmospheric N2. This chapter addresses and discusses H2 metabolic pathways involved in cyanobacterial H2 production and summarizes available and future potential strategies for H2 yield improvements. The focus is on metabolic manipulation and genetic engineering approaches and on immobilization technologies for enhancing H2 productivity in cyanobacteria.

2 Biophotolysis and H2 Metabolism in Cyanobacteria

Cyanobacteria are photoautotrophic organisms that use sunlight as energy source together with atmospheric CO2 and water for growth. The thylakoid membranes in the cytoplasm of cyanobacteria contain pigment molecules such as chlorophyll a, phycocyanin, phycoerythrin, and allophycocyanin used to absorb light energy (i.e., photons) for oxygenic photosynthesis. The photosynthetic electron transfer reaction is divided into two parts, the light and dark reaction, respectively. The light reaction is involved in transferring electrons through an electron transport chain from PSII to plastoquinone (PQ) pool, cytochrome b6f complex (Cyt b6f), photosystem I (PSI), and ferredoxin (Fd), respectively, for generating ATP and reductants, NAD(P)H. For the dark reaction, CO2 is fixed and reduced into organic compounds using chemical energy obtained from the light reaction [2].

Cyanobacteria constitute a highly diverse group of prokaryotes that have different morphologies, unicellular to heterocystous and non-heterocystous filamentous forms. They are potential microbial chassis for H2 production by biophotolysis [9]. Biophotolysis is a process that involves the use of water as an electron donor, leading to the generation of O2 and H2 in the biological systems in a photosynthetic process. It can be divided into two pathways: direct and indirect biophotolysis pathways (Fig. 1). During direct biophotolysis, H2 is derived from the electrons generated by water splitting at PSII, whereas for indirect biophotolysis, protons and electrons are mainly supplied for hydrogen generation by degradation of intracellular carbon compound(s), the so-called fermentation [3].

Fig. 1
figure 1

Simplified view of direct and indirect biophotolysis for hydrogen metabolism involving photosynthetic system in thylakoid membrane of cyanobacterial cell. PSII photosystem II, PSI photosystem I, Cyd cytochrome bd quinol oxidase, PQH2/PQ plastoquinol/plastoquinone, Cyt b6f cytochrome b6f complex, PC plastocyanin, Fd ferredoxin, FNR ferredoxin NAD(P) reductase, NDH NAD(P)H dehydrogenase, N2ase nitrogenase, H2ase hydrogenase, H2 hydrogen (This view was modified from previous articles [10, 11])

Full size image

3 H2-Catalyzing Enzymes in Cyanobacteria

In cyanobacteria, there may be three enzymes directly involved in H2 metabolism [5, 6]. (1) Nitrogenase catalyzes the fixation of atmospheric N2 to produce ammonia (NH3) under limiting nitrogen condition and concomitantly produces H2 as a by-product. (2) Uptake (Hup) hydrogenase catalyzes the consumption of the H2 evolved during N2-fixation, which reduces the energy loss during nitrogenase catalysis. (3) Bidirectional (Hox) hydrogenase catalyzes both consumption and production of H2. Both nitrogenase and hydrogenase are highly O2 sensitive and have been a popular target for enzyme improvement. Figure 2 shows an overview of the structural organization of the different hydrogen catalyzing enzymes in cyanobacteria.

Fig. 2
figure 2

The enzymes involved in H2 metabolism in cyanobacteria. Nitrogenase catalyzes N2-fixing from the atmosphere to produce ammonia and H2 as a by-product. The produced H2 is consumed by the uptake Hup-hydrogenase. The bidirectional Hox-hydrogenase can either consume or produce a molecule of H2 depending on the redox potential (Modified from Tamagnini et al. [12])

Full size image

Nitrogenase in N2-fixing, diazotrophic, cyanobacteria is a multiprotein enzyme complex consisting of the dinitrogenase (MoFe protein) and dinitrogenase reductase (Fe protein). The MoFe protein is a heterotetramer α2β2 with a molecular weight of about 220–240 kDa encoded by nifD and nifK for α and β subunits, respectively. However, it has also been found that several strains of Anabaena, including Anabaena variabilis, are able to synthesize an alternative nitrogenase, encoded by the vnf gene cluster, where molybdenum is replaced by vanadium in the active center of the enzyme [13]. The function of dinitrogenase is a reduction of N2 bonds leading to the formation of ammonia (NH3). The Fe protein is a homodimer with a molecular weight of about 60–70 kDa and encoded by nifH. It transfers electrons from the external electron donor to the dinitrogenase protein [13]. This enzyme catalyzes the reduction of atmospheric N2 to NH3 and is also responsible for reducing protons (H+) into H2. However, nitrogenases have a rather low turnover rate [14] and H2 production by nitrogenase requires a considerable number of electrons, reductants, and ATP molecules provided from photosynthesis or by carbohydrate degradation in the cell.

Moreover, nitrogenases are extremely O2 sensitive, and diazotrophic cyanobacteria have evolved several strategies to separate the photosynthetic evolution of O2 from the process of N2 fixation. In filamentous cyanobacteria (e.g., Anabaena variabilis ATCC 29413 and Anabaena sp. PCC 7120), the vegetative cells can differentiate into heterocyst cells (Fig. 3). Mature heterocysts are individual cells providing a microaerobic environment suitable for the enzymes involved in N2 fixation and H2 metabolism. Heterocysts contain a thick cell wall and lack active photosystem II (PSII) complexes resulting in the absence of photosynthetic O2 evolution [16]. The vegetative cells perform photosynthetic and CO2 fixing processes, whereas CO2 fixation is absent in heterocysts due to the lack of the primary CO2 fixing enzyme ribulose bisphosphate carboxylase (Rubisco). Heterocysts import carbohydrates, most likely as sugars, from vegetative cells and use the oxidative pentose phosphate (OPP) pathway for carbohydrate degradation to generate energy and reduce power for nitrogen fixation. In return, the heterocysts export nitrogen in the form of glutamine to the vegetative cells through the GS-GOGAT pathway (Fig. 3) [17, 18]. In some unicellular cyanobacteria, such as Cyanothece sp. and Trichodesmium sp., N2 fixation may be controlled by the circadian clock. They separate the production of O2 and H2 by performing oxygenic photosynthesis during the daytime and nitrogen fixation at night [19,20,21].

Fig. 3
figure 3

Simplified view of heterocyst metabolism and exchange with vegetative cells of filamentous heterocystous cyanobacteria under nitrogen starvation. Carbohydrates are imported from vegetative cells into the heterocyst, where they supply reducing power for N2-fixation. In turn, N2 is bound in glutamine and exported into vegetative cells through the GOGAT pathway. Dotted lines represent a flow of reducing equivalents. PSI photosystem I, OPP oxidative pentose phosphate pathway, RET respiratory electron transport chain, IDH isocitrate dehydrogenase, GS glutamine synthetase, GOGAT glutamate synthase, Fd ferredoxin, Gln glutamine, Glu glutamate, 2-OG 2-oxoglutarate, N2ase nitrogenase, H2ase uptake-Hup-hydrogenase (Modified from Lindberg [15])

Full size image

Uptake (Hup) hydrogenase has been reported for all known N2-fixing cyanobacteria [6, 22]. It is a heterodimeric enzyme consisting of at least two subunits: HupS (encoded by hupS) and HupL (encoded by hupL). The HupS subunit has a molecular weight of about 35 kDa containing three FeS clusters. The HupL subunit containing the active site is about twice as large with about 60 kDa. It consists of four conserved cysteine residues involved in coordinating the metallic NiFe at the center of the active site [23, 24]. The uptake (Hup) hydrogenase is involved in the efficient recycling or consumption of the H2 produced by the nitrogenase. Utilization of H2 in N2-fixing cyanobacteria is associated with (1) providing additional reducing equivalents to PSI and various cell functions, (2) generating ATP from oxyhydrogen reaction, and (3) preventing inactivation of nitrogenase by removing O2 [25]. The structural hupS and hupL genes have been characterized in many cyanobacteria such as Nostoc sp. PCC 73102, Anabaena variabilis ATCC 29413, and Gloeothece sp. ATCC 27152 [26,27,28]. hupS is usually located upstream of hupL. The analysis of gene expression using RT-PCR technique revealed that hupS and hupL are co-transcription and an enhanced transcription level was found when cells were grown under N2-fixing condition or the addition of external Ni2+ in the culture medium [12]. In some N2-fixing cyanobacteria, e.g., Anabaena sp. PCC 7120, hupL in the vegetative cells is interrupted by a DNA element which is excised during heterocyst differentiation by a site-specific recombinase (XisC) resulting in a contiguous hupL (Fig. 4) [22, 29, 30].

Fig. 4
figure 4

Schematic representation of the hupL rearrangement occurring in Anabaena sp. PCC 7120. In the vegetative cell, hupL is interrupted by a 9.5-kb DNA element containing site-specific recombination (xisC). In contrast, the structure of the hupL gene is restored, allowing its expression only in the heterocyst cell. The question marks indicate unclear data explanation (Modified from Tamagnini et al. [12])

Full size image

Bidirectional (Hox) hydrogenase is commonly, though not universally, found in cyanobacteria, catalyzing both consumption and production of molecular H2 [31, 32]. It is a heteropentamer encoded by hoxEFUYH and consists of two protein complexes: a hydrogenase complex (HoxY and HoxH) and a diaphorase complex (HoxE, HoxF, and HoxU). The large subunit, HoxH contains the active metal NiFe center like the uptake hydrogenase (Fig. 2). The physiological role of this enzyme is still under debate. It was found that the bidirectional hydrogenase of Synechocystis sp. PCC 6803 acts as an electron sink, storing excess electrons from PSI in the form of hydrogen [33]. Gutekunst et al. [34] reported that Hox-hydrogenase probably functions as an electron sink for reduced ferredoxin/flavodoxin under mixotrophic and nitrate-limiting condition. In addition, this enzyme has been proposed to be a mediator in the release of excess reducing power under anaerobic conditions [35]. Studies in Synechocystis PCC 6803 found that the enzyme was insensitive to light, reversibly inactivated by O2, and quickly reactivated by NADH or NADPH [36].

4 Strategies for H2 Yield Improvements in Cyanobacteria

Cyanobacteria are potential H2-producers, as they can produce H2 from water as a result of solar energy conversion. However, the main obstacle for the biotechnological process is the low yield of cyanobacteria strains producing H2 (in the range of 0.06–31.8 μmol H2/mg Chl a/h). Increasing the H2-productivity by cell improvement has been widely studied using diverse technologies. This section summarizes recent improvements of H2-metabolism in cyanobacteria by focusing on metabolic manipulation and genetic engineering approaches to understand the metabolic pathways further and increase their respective H2 yields. An overview of selected cyanobacterial strains and their corresponding rates of H2 production are summarized in Tables 1 and 2.

Table 1 H2 production in different cyanobacteria and their optimum environmental condition for enhanced H2 production
Full size table
Table 2 H2 production comparison among cyanobacteria using immobilization techniques vs. cell suspension (free cell)
Full size table

4.1 Metabolic Manipulation Approaches

4.1.1 Physiochemical Parameters Affecting H2 Production

Several parameters may enhance H2 production, such as nutrient and culture compositions, inorganic mineral supplements, the pH and temperature of culture media, and light intensity. Carbon (C), nitrogen (N), phosphorus (P), and sulfur (S) are all required nutrients for cyanobacterial growth and have been examined for optimizing cellular H2 production by various microalgae, see Table 1. Changes in the composition of nutrients affect the H2 production rates. Addition of a carbon source supports by providing energy for cell metabolism. Some cyanobacteria can consume organic carbon sources such as glucose, fructose, galactose, lactose, mannitol, sorbitol, sucrose [39, 44, 49], acetate, succinate, and malate [55] having an effect on hydrogenase or nitrogenase activity and thus on H2 production. In Synechocystis PCC 6803 it was shown that addition of glucose increases the level of reduced NAD(P) which is beneficial for bidirectional Hox-hydrogenase activity, resulting in enhanced H2 production [49]. Besides, in Anabaena sp. PCC 7120, fructose mediated an increase of H2-production with increased nitrogenase activity and nifD expression, in conjunction with elevated electron flow from utilization of fructose through the oxidative pentose phosphate pathway [39].

Although nitrogen and sulfur are essential nutrients for microbial growth, an enhanced H2 production rate was detected when cells were grown in the nitrogen- or sulfur-deprived condition. This phenomenon was observed in several cyanobacteria such as Aphanothece halophytica [40], Anabaena siamensis [38], Arthrospira sp. PCC8005 [43], Gloeocapsa alpicola, and Synechocystis sp. PCC 6803 [46]. Furthermore, exogenously added nitrogen sources inhibit nitrogenase activity [13]. Phosphorus (P) is an essential heteroelement in compounds such as ATP, NAD(P)H, nucleic acids, and sugar phosphates, all of which play important roles in photosynthesis. NAD(P)H is the electron donor to the bidirectional Hox-hydrogenase in cyanobacteria [56].

Generally, trace elements act as essential cofactors, which play an important role in activities of both hydrogenase and nitrogenase enzymes involved in H2 evolution. For example, a culture of Fischerella muscicola TISTR 8215 grown with higher levels of Mo6+ showed increased nitrogenase activity leading to increased H2 production [45]. Additionally, the relevance of concentrations of Fe3+, Ni2+, and Mo2+ ions for H2 production has been investigated and optimized for several strains of cyanobacteria [38, 42, 43, 57], with results suggesting that availability of these elements is a critical factor in controlling H2 production and N2 fixation, including effects on expression of hydrogenase and nitrogenase genes.

Furthermore, pH and temperature are crucial parameters influencing the H2 production process. The pH ranges from 6 to 9 were examined for enhanced H2 production in several cyanobacteria. In tests using Lyngbya perelegans the highest H2 production was obtained at pH 8.0 [47]. Regarding the temperature, the optimum temperature for H2 production for most cyanobacteria varies between 23 and 40°C but with differences between strains. Nostoc muscorum and Lyngbya perelegans showed optimum hydrogen production at 40°C [47, 48] whereas in Arthrospira sp. PCC 8005 the maximum rate of H2 production was observed at 30°C [42]. Moreover, Calothrix sp., Nodularis sp., and Microcystis sp. showed optimum H2 production at 23°C [58].

Light intensity is a most critical factor affecting the efficiency of cyanobacterial H2 production. Under artificial illumination, microalgal cultivation under different light intensities alters the metabolic capacity of the cells. Photobiological H2 production in microalgae and cyanobacteria results from the contribution of a direct and an indirect electron transfer pathway [59,60,61]. The direct biophotolysis involves a PSII-dependent pathway, which links water-splitting activity to H2 production. In indirect biophotolysis, electrons, which are derived from the degradation of stored carbohydrates entering the electron chain at the plastoquinone pool are hereafter transferred to the hydrogenase to produce H2 [62]. Previous studies reported that the impact of light intensity varies among different species and strains. The heterocystous cyanobacteria Nostoc muscorum and Anabaena PCC 7120 produce H2 from nitrogenase in heterocysts under light conditions [48, 63]. Enhanced light intensity resulted in increased H2 production in A. siamensis TISTR 8012 with a saturation at 200 μE/m2/s of light intensity after 12 h. The cells generated less H2 above 200 E/m2/s, along with decreased chlorophyll a and cell lysis [37].

4.1.2 Cell Immobilization for Reduced O2 and Cell-Stacking Effects

Hydrogenase catalyzes the incorporation of two protons and two electrons to form H2, which is the smallest molecule in the universe. H2 is produced inside the cytoplasm of the cell, diffuses toward liquid broth through the lipid bilayer cell membrane, and finally to the headspace, driven by the partial pressure of the gas. The cytoplasm has the highest H2 partial pressure, followed by liquid broth and headspace, accordingly. Consequently, the H2 yield can be determined by quantifying the gas in the headspace using, e.g., gas chromatography. O2, the strong competitive hydrogenase inhibitor generated in PSII by the water-splitting reaction, has highly similar physicochemical properties, which makes it challenging to separate both gases. Apart from being a strong inhibitor for most H2 producing enzymes, it also forms an explosive mixture with H2 (Knallgas reaction) and thus poses a significant safety issue if it comes to scale-up. The amount of molecular oxygen released by the photosynthetic activity depends on the cellular respiration process consuming O2 as the final electron acceptor. Therefore, one strategy to keep O2 levels low in H2 producing cultures is the balancing of PSII activity and cellular respiration [41].

Apart from the parameters discussed above also the cell concentration in the culture was reported to affect the H2 yield, which is decreasing with increasing cell densities [45, 51]. Too dense cells culture led to the so-called cell-stacking effect, in which cells shade each other and thus run into a light limitation (Fig. 5), which is also difficult to solve by vigorous shaking or mixing [64]. Especially for filamentous cyanobacteria strong mixing is not an option, as the filaments prevent a homogenous mixing, and will also be negatively impacted by high shear forces. Cell immobilization may be a promising solution to relieve the problem of cell-stacking.

Fig. 5
figure 5

Schematic shows light penetration power to stacked cells/precipitated cells (a), shaking cells (b), and immobilized cells (c). Dotted lines represent the light penetration path from the surface toward the center of a container

Full size image

Cell immobilization is an essential technique for reducing the cell-stacking effect. For immobilization, the cells are embedded into a supporter material, which is polymerizing during the process. Depending on the physicochemical properties and the concentration of the supporter material, in situ gas removal strategies (like O2 or H2) can be implemented into the system. To obtain the highest H2 yield many studies have been performed with production cycles. Several common biomaterials have been used as cell support. These include carrageenan [53], agar [45, 51, 53, 54], agarose [53], and alginate [52, 65,66,67,68]. After immobilizing the cells in a selected support, the mixtures can be molded into different shapes. Thin films [52, 67], cubes [45, 53, 54], and beads [51, 65, 66, 68] are commonly used. Reported yields and corresponding production system parameters are summarized in Table 2.

Rashid et al. [54] demonstrated that the unicellular cyanobacterium Microcystis aeruginosa immobilized in 1.5% agar in cubes can sustain a hydrogen production phase for up to 95 h with a yield of about 65–70 mL H2/L culture. This can be increased by the addition of glucose to the culture, which may be degraded through glycolytic pathways, generating the reducing equivalent NADH, which support the flow of electrons to the plastoquinone pool between PSII and PSI and thereby increase the yield of H2. Wuthithien et al. [45] also reported that immobilizing cells of the N2-fixing filamentous cyanobacterium Fischerella muscicola TISTR 8215 in 1.5% agar improved the H2 yield significantly, and increasing the Mo6+ ion concentration also resulted in an increase in H2 production rate. It seems that stimulation of nitrogenase activity occurs through an addition of molybdenum into their active site [38].

The beneficial effect of immobilization on H2 production may be explained as follows: (1) The immobilization matrix reduced the O2 concentration in the direct environment of the cells [69]. (2) The cell-stacking effect was reduced by the immobilization resulting in improved light supply. (3) Optimized mass-transfer between nutrients from broth to cells [70]. (4) The initial cell numbers to agar concentration was appropriate for increasing agar mechanical stability [70].

Pansook et al. [53] compared different materials for immobilization and reported that immobilized unicellular cyanobacterium Aphanothece halophytica in 3% (w/v) agar showed the highest H2 production compared to carrageenan, agarose, and free cells. Carrageenan might encounter the problem of low stability during gel formation due to the presence of NaCl, as previously reported [71], whereas agarose showed lower stability than agar in agreement with Semenchuk et al. [72]. The high production rate in cells immobilized in agar was related to: (1) improved cell survival rate and mechanical stability, (2) better nutrient diffusion rate from broth to cells, and (3) small size of the immobilization particles (0.125 cm3) facilitating H2 and O2 diffusion from cells toward the bulk.

Another commonly used immobilization matrix is alginate. It is a water-soluble carbohydrate polymer, which will polymerize when interacting with CaCl2 in solution. Ca2+ will replace Na+ ions and cross-interact with carboxylate groups (-COO−) and negatively polar groups (-OH), leading to carbohydrate strand incorporations and gel formation over time (Fig. 6). Immobilized filamentous cyanobacterium Lyngbya perelegans in 4% agar cubes was studied and compared to alginate beads [51]. There were only slight differences between the two materials tested, but both showed about 1.8 times higher H2 productivity than free cells. Interestingly, this immobilized culture was used to investigate the impact of various gas mixtures, and it was found that a CH4:Ar (11:2) mix resulted in the highest productivities [51]. Furthermore, Leino et al. [52] screened for H2 producing cyanobacteria from the University of Helsinki Culture Collection and identified the N2-fixing heterocystous filamentous cyanobacterium Calothrix 336/3 as the strain with the highest H2 production. Immobilized in a Na-Alginate thin film, it showed a maximum H2 production rate higher than the rate from free cells. In a study using the model strain Anabaena PCC 7120 wildtype and a ΔhupL mutant of the same strain, there were only small differences between free cells and cells immobilized in alginate films. The study also found that periodically purging the system with CO2 balanced Ar led to increased H2 yield as CO2 was used as a signal to enhance N2 fixation [52], consequently prolonging H2 production.

Fig. 6
figure 6

Cells immobilized in calcium alginate beads prepared using sodium alginate dissolved in algal medium, and Ca-alginate formation from Na-alginate. Negatively charges of Na-alginate chains repulse each other, leading to a uniform structure. Once Ca2+ ions are present, positively charges attempt to combine each negative strand close to each other, forming a gel structure (Modified from Touloupakis et al. [50])

Full size image

One of the essential parameters for H2 production is the photosynthetic activity since PSII generates O2 with an electron flow to the bidirectional hydrogenase and nitrogenase. Restoring photosynthetic activity between the production cycles thus plays a vital role in prolonging H2 production as observed in Calothrix 336/3, Anabaena PCC 7120 wildtype, and its ΔhupL strain [67].

Finally, cell immobilization enables cell retention and the recycling of the cellular biocatalyst for multiple batches. This facilitates process optimization, as various reaction conditions can be tested with one batch of biocatalysts, like purging with inert gas or gas mixtures, applying different media, etc. Furthermore, it allows to operate the reactors in a continuous or semi-continuous mode, positively influencing process economics.

In summary, cyanobacterial immobilization is an interesting option to enhance H2 production and process stability by facilitating gas (H2 and O2) removal from the cultures. However, optimal conditions in terms of immobilization material and reaction environment will differ from strain to strain and we are still far from defining general process parameters for optimized H2 production, as up to now only case (strain) specific examples are reported and general operation protocols are missing.

4.2 Genetic Engineering Approaches

4.2.1 Eliminating of Electron Competing Pathways for Promoting H2 Metabolism

The principal reason for H2 metabolism through bidirectional Hox-hydrogenase in cyanobacteria may be a disposal of excess reducing equivalents during fermentative metabolism associated with photosynthesis or/and dark anaerobic conditions. Therefore, the bidirectional Hox-hydrogenase requires numerous electrons and reductants as substrates supporting its activities. However, electrons generated through oxygenic photosynthesis are under most conditions not primarily shuttled to H2 metabolism. Instead, these electrons can be transferred to other competing pathways, such as the respiratory electron transport chain, nitrogen assimilation, and carbohydrate metabolism, shown in Fig. 7. Therefore, diverse genetic engineering strategies for enhanced H2 production by re-direction of electrons flow toward H2 metabolism have been extensively examined (Table 3). In Synechocystis sp. PCC 6803, interruption of all respiratory terminal oxidases (ΔctaI, ΔctaII, and Δcyd) induce the bidirectional Hox-hydrogenase activity leading to a higher H2-production rate than in wildtype cells under light condition [73]. Moreover, inactivation of type I NADPH-dehydrogenase complex (NDH I) by deleting the large subunit NdhB in a mutant Synechocystis strain M55 resulted in prolonged H2-production and a lower level of O2 being produced under light condition [36]. Engineering strains with disrupted nitrate assimilation, either nitrate reductase (ΔnarB) or nitrite reductase (ΔnirA) or both genes (ΔnarB/ΔnirA), in Synechocystis sp. PCC 6803 were found to induce significantly higher H2 production than in wildtype cells [74]. In addition, a mutant Synechococcus sp. PCC 7002 (ΔldhA), lacking the enzyme for the NADH-dependent reduction of pyruvate to D-lactate, showed an increased ratio of NADPH to NADP+ and a five-times higher H2-production when compared with wildtype cells [75]. This work supported that by eliminating competing fermentative carbon metabolism such as the pathway to produce lactate it may be possible to redirect the electron flux to H2 metabolism in cyanobacteria. Accordingly, an engineering approach by eliminating competitive electron pathways is an effective and promising method to improve cyanobacteria potential for H2 production, which should be further explored.

Fig. 7
figure 7

Different pathways of electron flow involved in H2 metabolism of cyanobacteria. Dotted lines represent electrons that can be transferred to other assimilatory or competing pathways. Cyd quinol oxidase, Cyt b6f cytochrome b6f, Cyt c553 cytochrome c553, Cyt ox cytochrome c oxidase, Fd ferredoxin, FNR ferredoxin-NADP reductase, Hox bidirectional Hox-hydrogenase, Hup uptake Hup-hydrogenase, N2ase nitrogenase, NDH NADPH dehydrogenase, OPP oxidative pentose phosphate pathway, PC plastocyanin, PSI photosystem I, PSII photosystem II, PQ plastoquinone pool, Rubisco ribulose-1,5-bisphosphate carboxylase oxygenase, SDH succinate dehydrogenase (Modified from Khetkorn et al. [10])

Full size image
Table 3 H2 production in engineered cyanobacterial strains using different strategies (Modified from Khetkorn et al. [10])
Full size table

4.2.2 Modifying Heterocyst Frequency for Increased H2 Production

In heterocystous filamentous cyanobacteria, nitrogenase is a key player for H2 production. The heterocyst provides a partially microoxic environment suitable for oxygen-sensitive enzymes such as nitrogenase since it lacks the PSII activity and has an increased respiration rate [84]. Furthermore, it is surrounded by a thick envelope limiting O2 diffusion through the cell wall (Fig. 3). Therefore, increasing the heterocyst frequency may enhance H2 production by promoting nitrogenase activity. The heterocyst differentiation process has been primarily studied in Anabaena sp. PCC 7120 in which it takes approximately 24 h to develop a mature heterocyst from a vegetative cell under nitrogen limited condition [17]. One of the key genes in the regulation of heterocyst pattern formation, hetR, encodes a serine-type protease, which is expressed early during heterocyst differentiation. Inactivation of hetR inhibits early steps in the differentiation process, while overexpression of the gene increases heterocyst frequency [85]. Recently, it was demonstrated that the addition of fructose rapidly induced the development of mature heterocysts and led to upregulation of hetR transcription, resulting in enhanced N2-fixation and H2-production in Anabaena sp. PCC 7120 ΔhupL strain [39]. HetF (a protease) influences heterocyst development by inhibiting hetR expression during cell differentiation [86]. PatA, a response regulator, is also known to effect post-translational modification of HetR [87]. However, a practical study with strains exhibiting a genetically engineered high heterocyst frequency with enhanced H2 production is yet to be reported.

4.2.3 Inactivation of Uptake (Hup) Hydrogenase Function for Enhanced H2 Production

Uptake hydrogenase activity is a major obstacle for enhanced H2-production in N2-fixing cyanobacteria since it catalyzes the consumption of H2 produced by nitrogenase. Therefore, the disruption of uptake hydrogenase function has been widely studied in many N2-fixing cyanobacteria. Generally, the structural genes encoding uptake Hup-hydrogenases are clustered in a similar physical organization forming a transcript unit, hupS being located upstream of hupL (Fig. 4). Inactivation of xisC in Anabaena sp. PCC 7120 resulted in a strain incapable of forming a functional uptake hydrogenase [29]. A mutant strain AMC 414 (ΔxisC) showed high potential for H2-production compared to wildtype strain under higher light intensity [63]. Moreover, target genes (hupS, hupL, and hupW) that affect H2-uptake deficiency in N2-fixing cyanobacteria have been extensively investigated, see Table 3. All generated strains produce H2 at significantly higher rates than their respective wildtype cells. These experiments indicate that the genetic inactivation of hup is an effective strategy for improving cyanobacterial H2 production.

4.2.4 Introduction of Non-native Hydrogenase for Enhanced H2 Productivity

Cyanobacteria produce H2 through bidirectional Hox-hydrogenase ([NiFe]-hydrogenase) with a low rate of H2-evolution. Therefore, the expression of non-native hydrogenase has been a focus for improving H2 productivity in cyanobacteria. These include high turnover [FeFe] hydrogenase and some O2-tolerant [NiFe] hydrogenases from other organisms using advanced synthetic biology techniques. However, successful heterologous expression of [FeFe]-hydrogenase in cyanobacteria remains a challenge, and to date, only very few reports are available, Table 3. The first report by Ducat et al. [80] demonstrated the expression of a [FeFe] hydrogenase (HydA) and the accessory HydEFG from the anaerobic fermentative bacterium Clostridium acetobutylicum into Synechococcus elongatus PCC 7942. Interestingly, the results showed both in vitro and in vivo activity of non-endogenous hydrogenase connected to the light-dependent reactions of the electron transport chain. Gärtner et al. [81] have been successfully expressed the FeFe-hydrogenase operon (hydA, hydB, hydE, hydF, hydG) and two additional genes, S03922 and S03924, from Shewanella oneidensis MR-1 into the filamentous cyanobacterium Anabaena sp. PCC 7120. Avilan et al. [88] expressed a clostridial [FeFe]-hydrogenase specifically in the heterocysts together with a GlbN cyanoglobin to decrease the O2 levels in the cell. The obtained strain showed H2 production concomitantly with oxygenic photosynthesis in the vegetative cells of the filaments. Furthermore, Weyman et al. [82] reported expressing [NiFe] hydrogenases from Thiocapsa roseopersicina, as well as hynSL along with 11 adjacent proteins from Alteromonas macleodii in Synechococcus elongatus. The advantage of using [NiFe] homolog over the [FeFe] hydrogenases was their increased half-life and enhanced tolerance toward oxygen stress [89]. The results showed in vitro activity of the expressed protein. Expression of such oxygen-tolerant hydrogenases in photosynthetic systems may open new avenues in cyanobacterial H2 production. Recently, another strategy that circumvents the biological maturation of [FeFe]-hydrogenase by an artificial synthetic activation of a heterologously expressed HydA protein in living cells of, e.g., Synechocystis PCC 6803 was developed. A functional HydA was created by the addition of a synthetic analogue of the [2Fe] subcluster mimicking the active site outside the cells [7]. The experiments showed that the non-native, semisynthetic FeFe-hydrogenase retain its H2 production capacity for several days after synthetic activation with a regulation of activity based on availability of electrons. The artificial activation technology was expanded to a newly discovered [FeFe]-hydrogenase which when expressed in Synechocystis showed stable expression and significant H2 production under different environmental conditions [8]. The developed technology opens up unique possibilities to investigate not only [FeFe]-hydrogenases but also other metalloenzymes in a photosynthetic microbial cell environment, completely bypassing the many challenges of, e.g., biological maturation and regulations.

In another recent development, Lupacchini et al. [83] introduced an O2-tolerant hydrogenase from Ralstonia eutropha (ReSH) into Synechocystis genome. The resulting engineered strain was able to produce H2 in the dark under fermentative conditions, as well as in the light, under conditions promoting intracellular NADH excess. This opens new possibilities for efficient cyanobacterial H2 production also under O2 replete conditions.

5 Conclusions and Perspectives

Due to the growing emphasis on developing renewable energy sources, cyanobacteria have been intensively studied as green cell factories for sustainable H2 production. Researchers are concentrating their efforts on the main native processes of cyanobacterial photosynthesis, fermentative metabolism, and on the enzymes involved in H2-metabolism, which holds great promise in terms of gaining fundamental knowledge and practical applications in biotechnology. The majority of research focuses on applying various metabolic manipulation strategies to enhance H2 yield in cyanobacteria. Additionally, genetic engineering is used to increase the H2 yield as well as the technology of cell immobilization for H2 scale-up challenges. Despite the enormous theoretical potential of cyanobacterial based H2 production, there are still significant barriers to its commercialization. The prospects of the biohydrogen energy sector will be determined by the combined efforts of scientists and engineers, state political support, and substantial R&D efforts.