Skip to main content

Microalgal upgrading of the fermentative biohydrogen produced from Bacillus coagulans via non-pretreated plant biomass



Hydrogen is a promising source of alternative energy. Fermentative production is more feasible because of its high hydrogen generation rate, simple operating conditions, and utilization of various organic wastes as substrates. The most significant constraint for biohydrogen production is supplying it at a low cost with fewer impurities.


Leaf biomass of Calotropis procera was used as a feedstock for a dark fermentative production of hydrogen by Bacillus coagulans AH1 (MN923076). The optimum operation conditions for biohydrogen production were 5.0% substrate concentrationand pH 9.0, at 35 °C. In which the biohydrogen yield was 3.231 mmol H2/g dry biomass without any pretreatments of the biomass. A freshwater microalga Oscillatroia sp was used for upgrading of the produced biohydrogen. It sequestrated 97 and 99% % of CO2 from the gas mixture when it was cultivated in BG11 and BG11-N media, respectively After upgrading process, the residual microalgal cells exhibited 0.21mg/mL of biomass yield,high content of chlorophyll-a (4.8 µg/mL) and carotenoid (11.1 µg/mL). In addition to Oscillatroia sp residual biomass showed a lipid yield (7.5–8.7%) on the tested media.


Bacillus coagulans AH1 is a promising tool for biohydrogen production avoiding the drawbacks of biomass pretreatment. Oscillatroia sp is encouraged as a potent tool for upgrading and purification of biohydrogen. These findings led to the development of a multiproduct biorefinery with zero waste that is more economically sustainable.

Graphical Abstract


Investigation of new alternative energy sources are promising strategies to address the energy shortage and environmental pollution that threaten social and economic developments [1, 2]. Hydrogen is one of the most valuable renewable carriers, can deliver or store an enormous amount of energy. It can be used in fuel cells to generate electricity, or heat. Hydrogen is characterized as clean, pollution-free and high efficiency and density energy, high, zero-or near zero-emissions operation and reduce greenhouse gases emission [3, 4]. Numerous applications, such as distributed or combined heat and power, backup power, systems for storing and enabling renewable energy, and portable power can be powered by hydrogen and fuel cells. Moreover, hydrogen and fuel cells were used as auxiliary power for transportation, petroleum refining and fertilizer production [4]. As a result, there is much potential for this form of hydrogen production to replace fossil fuels [5]. Biological hydrogen production via fermentative or photosynthetic process is among the diverse technologies for hydrogen production. However, the fermentative production is more feasible as it has a fast rate of hydrogen production, simple operation conditions, and consumption of various organic wastes as substrates [6, 7]. Moreover, fermentative production of hydrogen not only treats the organic wastes but also provides extensive clean energy with low cost [5, 8]. Dark fermentation is the bioconversion of wastes (organic substrates) such as agricultural, and industrial effluents into hydrogen through a series of biochemical reactions catalyzed by bacteria under certain environmental conditions [9]. The major advantage of biohydrogen production through dark fermentation is the consumption of many organic wastes such as cellulosic and lignocellulosic as a substrate and it does not require light. Substrates have a major influence on biohydrogen productivity due to their different biodegradability [10, 11]. Due to their capacity to produce beneficial biopolymers, the latex-bearing plant could be used for many applications [12]. Furthermore, many members of these plants yield active chemicals that are frequently employed in traditional medical practices or toxin production in many countries [13]. Calotropis procera (giant milkweed) is a member of the family Asclepiadaceae that has been studied primarily because of its variable medicinal properties [14]. It has evergreen leaves, extensively disseminated in the world [15, 16]. Other research have revealed that C. procera contains significant levels of hydrocarbons. [16,17,18]. Calotropis procera was a potential plant for bioenergy and biofuel production in semiarid regions of the world [19, 20] as it is a fast-growing plant with easily accessible biomass that is not normally fed by cattle due to latex toxicity [15].

The limitation of biohydrogen production via dark fermentation is the release of minor impurities such as CO2, H2S, N2 and water vapour that reduce the quality of produced hydrogen and constrain its industrial applications. Carbon dioxide which included in biogas reduces the burning velocity which consequently impacts the performance of the engine as well as decreases the peak pressure and the maximum power inside the cylinder [3, 21]. Biogas containing more than 45% carbon dioxide causes harsh and irregular running of the engine. Therefore, reduction of carbon dioxide content will certainly improve the quality of biogas [22, 23]. Microalgae and cyanobacteria are believed to solve the problem of biohydrogen gas impurities due to their potential for CO2 sequestration [24]. They remove CO2 from the atmosphere with greater efficacy than conventional energy crops. The CO2 efficacy may reach 99%, which corresponds to 1.8 kg of CO2 per kilogram of dried biomass [25, 26]. In addition, they are anticipated to serve as the raw material for upcoming multiproduct biorefineries. Owing to their rapid growth, substantial metabolite composition, stress-tolerance mechanism, efficacy in remediation of wastewater, highest CO2 fixation rate, and production of valuable metabolites with industrial, environmental, and pharmaceutical significance [27,28,29,30] The majority of microalgal biomass (> 75%) is composed of lipids and carbohydrates. Carbohydrates derived from microalgae contribute equally to the biofuel industry through fermentation or acid treatment to produce bioethanol. Oleaginous microalgae can store 20–50% of their desiccated cell mass in oil, which is 300 times more than conventional energy crops [31, 32]. Several microalgal species can be stimulated to produce lipids by depriving themselves of nutrients [33]. After lipid extraction, the desiccated biomass can be utilized for bioethanol production, biochar production, and biofertilizer carriers [34]. Moreover, the pigments produced from microalgae can be used for industrial purposes [35]. Therefore, the main objectives of this study are (i) microbial screening and isolation of biohydrogen producing bacteria via non-pretreated biomass of Calotropis procera, (ii) biohydrogen production using Bacillus coagulans AH1 including optimization of fermentation conditions, (iii) upgrade of biohydrogen using microalgal fermentation, iv) assessment of microalgal usability for lipids, pigments.

Materials and methods

Samples collection

Leaves biomass of C. procera were collected from Wadi- Al-Assiuty-Assiut governorate, Egypt and prepared as described by [19]. The granular sludge sample was collected from the wastewater treatment plant in Arab Al-Madabegh region, Assiut-Egypt. The samples were kept in plastic sacs and used as a source of bacterial inoculum.

Enrichment, isolation and purification of hydrogen-producing bacteria

The fermentative biohydrogen-producing bacteria were isolated from the collected sludge sample. About 10% w/v of sludge was seeded in a sterilized anaerobic medium that was prepared according to the method described by [36]. The medium was supplemented with 10% of dried leaves biomass of C. procera as a substrate for biohydrogen production at pH 7.0 under strictly anaerobic conditions. It was incubated at 35 °C for 10 days under aseptic dark conditions with continuous shaking at 120 rpm to provide better contact with the substrate. The produced gas was received and collected in the graduated sterilized syringe. The previous step was repeated three times by placing 10% of the growing bacteria in a new anaerobic media to perform enrichment under the same previously mentioned conditions.

After the third enrichment step, a volume of 5 mL of the fermented mixture was re-seeded into a 50 mL serum bottle consisting of broth reinforced clostridial medium (RCM) [37, 38] for isolation of the potential hydrogen-producing isolates under anaerobic conditions. The composition of RCM per 1000 mL was: 3.0 g yeast extract, 10.09 g beef extract, 10.0 g peptone, 3.0 g sodium acetate, 5.0 g sodium chloride, 5.0 g glucose, 1.0 g soluble starch and 0.5 g cysteine-hydrochloride, pH was adjusted to 6.8. The bottles were incubated at 35 °C with continuous shaking at 120 rpm for 24 h. After that 100 µl was streaked onto RCM agar covered with sterilized paraffin oil and incubated at 35 °C for 24 h. The pure bacterial isolates were obtained by repeated streaking on RCM agar covered with sterilized paraffin oil. The purified isolates were separately preserved for further use.

Molecular genetic characterization of the bacterial isolates

The bacterial isolates were genetically characterized based on 16S rRNA gene sequencing after extraction of total genomic DNA [39, 40]. Extraction of the genomic DNA of the isolates was carried out at the Molecular Biology Research Unit, Assiut University. Using universal primers designed to amplify a 1500 bp segment of the 16S rRNA gene, the conserved area of the gene was amplified by polymerase chain reaction (PCR) (Solgent Co., Ltd, Bio-Industry Development Site, 63–10 Hwaan-Dong, Yuseong-Gu, Daejeon, South Korea). The reverse primer was 1492R(AGAGTTTGATCCTGGCTCAG), while the forward primer was 27F (CGGCTACCTTGTTACGACTT). The obtained sequences of the isolates were aligned and compared with the known 16SrRNA gene sequences in the GenBank database using the BLAST search at

A phylogenetic tree was developed to determine the isolate's taxonomic classification using MEGA 4.0's neighbour-joining approach. Phylogenetic trees were derived from 16S rRNA gene sequences which built in the context of 16S rRNA gene sequences from different bacterial strains deposited from GenBank [41, 42].

Optimization of fermentative biohydrogen production

The bacterial cells were previously cultivated in RCM at 35 °C, pH 7.0 and shaking at 120 rpm, under dark strictly anaerobic conditions (by purging with argon). A volume of 20 mL of bacterial suspension was anaerobically placed in a sterilized vail, centrifuged at 5000 rpm for 10 min, and resuspended in phosphate buffer saline (PBS). The batch experiments were performed in 120 mL sterilized serum bottles with a working volume of 50 mL of sterilized anaerobic medium supplemented with 0.5 g dried leaves powder of C. procera after autoclaving and inoculated with 10% v/v pre-growing bacterial cells (OD660 = 1.3). The bottles were sealed well with sterilized air-tight rubber caps and parafilm, and incubated in the dark at 35 °C. Then purged with argon gas to drop the dissolved oxygen concentration down to zero. During the incubation time (10 days), the produced gas was received in a sterilized graduated syringe.

The concentration of produced biohydrogen was determined by gas chromatography (GC) (Shimadzu, GC- 2014) equipped with a thermal conductivity detector (TCD) and Shin Carbon packed column (ST 80/100 2 m, 2 mm ID). Argon was used as carrier gas.

To assess the optimum nutritional and environmental operation conditions for maximum biohydrogen productivity, a conventional one-factor-at-a-time (OFAT) method was applied. The batch experiments were performed as previously mentioned at different substrate concentrations (2.5, 5.0, 7.5, 10.0, 12.5 and 15.0%) with pH 7 at 35 °C. For determination the optimum pH, the medium pH was adjusted to 3.0, 5.0, 7.0, 9.0 or 11.0 before autoclaving using dilute HCl or NaOH 10% of substrate was added at 35 ℃. The fermentative media with a substrate concentration of 10% and pH 7.0 was incubated at (25, 30, 35, 40 and 45 °C) to determine the impact of environmental factors on biohydrogen production. The bottles were incubated for 10 days with continuous shaking at 120 rpm. A control sample was performed using a fermentative medium containing either bacterial inoculum (without substrate supplementation) or C. procera biomass (without bacterial supplementation). An additional blank assay with only an anaerobic medium without substrate or inoculum was also conducted.

After the assesment of the optimum value of each factor, the dark fermentation process was performed to evaluate the amount of producing hydrogen at the optimum conditions.

Biological upgrading of the produced biohydrogen by microalgae

A filamentous microalgal isolate Oscillatoria sp. was previously isolated from the Nile River canal, in Assiut, Egypt. It was deposited in the World Data Center for Microorganisms at Suez Canal University Fungarium collection WDCM-1180 with strain ID: Oscillatoria SCUF0000351. Oscillatoria sp. was enriched in BG11 medium consists per g/L 1.5; NaNO3, 0.04; K2HPO4, 0.075; MgSO4·7H2O, 0.036; CaCl2·2H2O, 0.006; Citric Acid·H2O, 0.006; ferric ammonium citrate, 0.001; Na2EDTA·2H2O, 0.02; Na2CO3 and 1 mL of trace element mixture at pH 7.4 [43]. The microalgal cells were incubated at 35 °C under a white, fluorescent lamp (2000–3000 lx intensity) as a light source.

For determine the gas upgrading potential, the microalgal cells were cultivated in air-tight rubber cap stoppered glass bottles containing three different media: BG11, nitrogen-deprived BG11 medium (BG11- N) or distilled water. The gas mixture (biohydrogen + CO2) that was previously produced from AH1 bacteria at the optimum conditions was then passed over to the cultivated cyanobacterial cells. They were incubated under a white, fluorescent lamp (2000–3000 lx intensity) at 35 °C and 150 rpm in an orbital shaker as illustrated in the flowchart Fig. 1. The impact of the passed gas mixture on the growth indices and the physiological activities of the microalgal cell was investigated.

figure 1

Flowchart for biohydrogen purification by Oscillatoria sp

Determination of the removal efficiency of carbon dioxide

The concentration of gas mixture (H2 and CO2) before (inlet) and after (outlet) passing through Oscillatoria sp. cultures was determined by gas chromatography (GC). The inlet and outlet gases were expressed as mmol (H2 or CO2)/g of microalgal dry weight (DW). The biohydrogen to carbon dioxide ratio (H2/CO2) was estimated before and after the purification process as well as the estimation of the capability of CO2.

The removal efficiency of Oscillatoria sp was expressed by the consumed percentage of the inlet CO2 [44] as the following equation:

$${\mathbf{CO}}_{{\mathbf{2}}} {\mathbf{removal}} \, {\mathbf{efficiency}} = \, [({\text{Inlet CO}}_{{2}} {-}{\text{ Outlet CO}}_{{2}} ) \, /{\text{ Inlet CO}}_{{2}} ] \, \%$$

Determination of physiological and growth indices of Oscillatoria sp.

Determination of growth kinetics of Oscillatoria sp.

A volume of 50 mL of the cyanobacterial cells were harvested by centrifugation for 15 min. at 5000 rpm. The dry weight of cyanobacterial cells was determined after an overnight dry at 60 °C and it was expressed as mg/mL cell suspension. At the termination of each batch culture, the growth kinetics were carried out on a dry mass basis using the gravimetric technique previously used [45]. The biomass productivity (P biomass (mg mL−1d−1) was estimated by applying the following equation:

$${\text{P}}_{{{\text{biomass}}}} = \, \left( {{\text{W}}_{{\text{y}}} - {\text{W}}_{{\text{x}}} } \right)/{\text{reaction volume }}\left( {{\text{mL}}} \right)$$

where Wx and Wy were the initial and the final biomass concentrations at the end of the incubation period.

The specific growth rate (μ) was measured in terms of day−1 via the following equation:

$$\mu \, = {\text{ ln }}\left( {{\text{W}}_{{\text{y}}} /{\text{W}}_{{\text{x}}} } \right)/\left( {{\text{t}}_{{\text{y}}} - {\text{ t}}_{{\text{x}}} } \right)$$

where ty and tx were the time of harvesting and start time of cultivation, respectively. The doubling time (Td) for microalgae could be derived from Eq. (3) as follows:

$${\text{T}}_{{\text{d}}} = \, \mu \, \left( {{\text{t}}_{{\text{y}}} - {\text{ t}}_{{\text{x}}} } \right)/{\text{log2}}\left( {{\text{W}}_{{\text{y}}} /{\text{W}}_{{\text{x}}} } \right)$$

The potential captivity of carbon dioxide by Oscillatoria sp.

In terms of CO2 consumption by the microalgal biomass, the ability of microalgae to fix CO2 for conducting photosynthesis was assessed regularly. According to an equation developed from the average molecular formula of algal biomass CO0.48, H1.83, N0.11, P0.01 [46], the CO2 fixation rate (mg/mL d) was calculated, as follows:

$${\text{PCO}}_{{2}} = { 1}{\text{.883}}\, \times \,{\text{P}}_{{{\text{biomass}}}}$$

Estimation of photosynthetic pigments content in Oscillatoria sp.

Chlorophyll was investigated to determine the photosynthetic efficiency while carotenoid pigment was investigated to detect the potential of biotechnological applications. They were extracted by absolute methanol and quantified according to [47]. A volume of 2 mL of the cell suspension was centrifuged at 5000 rpm for 10 min to obtain the cell pellet. Then pigments were resuspended in absolute methanol for 10 min for pigment extraction. The sample was then kept for 24 h at 4 °C in the dark. It was centrifuged for 10 min at 5000 rpm to remove the cell debris. The supernatant was determined at 665 nm and 470 nm against blank (methanol). The chl-a and total carotenoid contents were estimated based on the following:

$${\text{Chl}} - {\text{a }}\left( {\mu {\text{g}}/{\text{mL}}} \right) \, = { 13}.{43 }\left( {{\text{A665}}} \right){\text{ v}}/{\text{b V}}$$
$${\text{Carotenoids }}\left( {\mu {\text{g}}/{\text{mL}}} \right) \, = { 4}.{4 }\left( {{\text{A47}}0} \right){ 1}0)/{\text{b V}}$$

where A665 and A470 are the absorbance values at 665 nm and 470 nm, respectively, against a blank; v was the volume of applied solvent (mL), b was the spectrophotometric cell length (1 cm), and V is the sample volume (mL).

Determination of the potential biodiesel production from Oscillatoria sp.

After the upgrading process, the potential lipid content produced form Oscillatoria sp. was determined according to [48]. A weight of 50 mg of Oscillatoria sp. biomass was harvested and homogenized in 4 mL ice—chilled chloroform–methanol (2:1) to release lipids. After that it was separated into chloroform and aqueous methanol layers by the addition of 4 mL ice–chilled 1M MgCl2 solution. The lower chloroform layer which contained most of the algal lipids was drawn off from the tube via a long syringe. Lipids were transferred into dry test tube and then evaporated and weighted to obtain the lipid yield and content. The lipid yield calculated as % (w/w) culture.,

$${\text{Lipid content }}\left( \% \right) \, = \, \left[ {\left( {{\text{mg lipid }}/{\text{ mg dry mass}}} \right) \, *{1}00} \right]$$

Statistical analysis

All experiments were performed in triplicate sets. Statistical analysis of data was performed using a one-way ANOVA test (Analysis of variance) by SPSS program version 25. Dunken value was determined at 0.05 level.


Isolation, molecular identification, and phylogenetic analysis of the H2-producing isolate

Ten anaerobic bacteria isolates were isolated from the sludge sample. The strain AH1 exhibited a high biogas productivity. Therefore, it was selected for further studies. The nucleotide sequence of the hydrogen-producing bacterial isolate has been deposited in the GenBank nucleotide sequence database under the name Bacillus coagulans strain with the accession Number MN923076. As analyzed by BLAST, the sequence of the 16S rRNA gene of the isolated strain AH1 showed high identity (99.93%) with Bacillus coagulans MT463837 in the GenBank as illustrated in Fig. 2.

Fig. 2
figure 2

Phylogenetic relationship between the strains AH1 and other 16S rRNA gene sequences of published Bacillus coagulans strains

Several sequences relevant to various Bacillus species were chosen from the Genbank database for the building of the phylogenetic trees to corroborate the location of the strain AH1 in the phylogeny. The tree revealed that B. coagulans and strain AH1 belonged to the same clade cluster Fig. 2. As a result, Bacillus coagulans was determined to be strain AH1. The strain AH1 was deposited in at Suez Canal University Fungarium culture collection WDCM-1180 under strain ID: Bacillus SCUF0000352.

Determination of the operation conditions for biohydrogen production

Impact of substrate concentration on Biohydrogen productivity

The concentration of the produced biohydrogen was investigated at different concentrations of C. procera leaves powder (2.5, 5.0, 7.5, 10.0, 12.5, and 15.0%). The amount of producing hydrogen from B. coagulans AH1 was 0.48, 2.94, 1.09, 0.73, 0.49, and 0.26 mmol H2/ g dry biomass, respectively. It was observed that the optimum production of biohydrogen was significantly (p ≤ 0.05) determined at 5.0% of substrate concentration and above this concentration the biohydrogen productivity dramatically reduced as illustrated in Fig. 3a.

Fig. 3
figure 3

Biohydrogen production from Bacillus coagulans using Calotropis procera different substrate concentrations a, different pH b and different temperatures c. *; represents a significantly different at (p ≤ 0.05) based on Duncan’s multiple range test

Impact of initial pH on biohydrogen productivity

Various pH values 3.0, 5.0, 7.0, 9.0 and 11.0 were assessed to investigate the optimum value for biohydrogen production at 35 °C, and 5% of substrate concentration. The concentrations of the produced biohydrogen were 0.0, 0.2, 0.7, 1.12, and 0.41 mmol H2/g dry biomass at the assessed pH values, respectively. It was noticed that the acidic pH of 3.0 significantly (p ≤ 0.05) inhibited the biohydrogen productivity however, the optimum productivity was recorded at an alkaline pH value; of 9.0 as shown in Fig. 3b.

Impact of temperature on biohydrogen productivity

The biohydrogen production was investigated at different temperatures (25, 30, 35, 40, and 45 °C) at pH 7.0 and 10% of the substrate concentration the concentration of produced biohydrogen by AH1 was 0.14, 0.50, 0.74, 0.18, and 0.0 mmol H2/g dry biomass at the tested temperatures, respectively. It was noticed that the highest (p ≤ 0.05) biohydrogen productivity was estimated at 35 °C while the productivity was completely inhibited at 45 °C as illustrated in Fig. 3c. On the other hand, the low temperature 25 °C significantly (p ≤ 0.05) showed reduced productivity compared to the high one Fig. 3c.

Biohydrogen productivity under optimal operation conditions

Based upon the previous optimization results, the optimum conditions that achieved highest biohydrogen productivity were 5% of substrate, pH 5.0 and 35 ℃. The concentration of the biohydrogen under the optimum conditions was 3.231 mmol H2/g dry biomass.

Biological upgrading of biogas using Oscillatoria sp.

Biohydrogen to carbon dioxide ratio before and after the upgrading process

The results in Fig. 4 displayed that the amount of inlet biohydrogen to the inlet carbon dioxide ratio (H2/CO2) was 0.34, 0.50 and 0.47 while the amount of outlet H2/CO2 ratio was 13, 21 and 3, for the microalgal culture BG11+g, BG11-N+g and dist. H2O+g, respectively. This result showed that there was a significant difference (p ≤ 0.05) between the H2/CO2 ratio before and after passing gases on microalgal cultures, where the outlet ratio exhibited a significant (p ≤ 0.05) higher than the inlet one. It was also noticed that the highest H2/ CO2 ratio was detected in BG11-N medium, while the lowest H2/ CO2 ratio was detected in dist. H2O microalgal medium despite it was still higher than the inlet ratio.

Fig. 4
figure 4

Biohydrogen to carbon dioxide ratio before and after purification using different algal cultures media

Determination of certain concentrations of the inlet and outlet gases

Carbon dioxide bio-fixation using microalgal-based technology is an economically feasible method for upgrading the biohydrogen. The capability of Oscillatoria sp. to purify and upgrade the biohydrogen produced by anaerobic bacterial fermentation via capturing of CO2 was investigated. The results in Fig. 5a&b illustrated (inlet) gases through each microalgal media and the outlet gases after the microalgal upgrading process. The inlet CO2 was 2.59, 1.51, 1.79 mmol CO2/g microalgal DW, while after passing the gases through the Oscillatoria sp., the outlet CO2 was 0.064, 0.017, 0.093 mmol CO2/g microalgal DW for BG11+g, BG11-N+g, and dist. H2O+g culture media, respectively. It was noticed that there was a significant (p ≤ 0.01) reduction in the amount of outlet CO2 compared with the outlet CO2 Fig. 5a.

Fig. 5
figure 5

The concentration of inlet (black column) and outlet (white column) CO2 (a) and H2 (b) through upgrading process by Oscillatoria sp. that grown on the microalgal cultures BG11, BG11– N and dist. H2O. *; represents a significantly different at (p ≤ 0.05) based on Duncanꞌs multiple range test. Tables represent the performance of biohydrogen upgrading process; % removal of CO2 (a) and % loss of H2(b)

On the other hand, the amount of inlet and outlet biohydrogen Fig. 5b. The inlet biohydrogen was 0.88, 0.76, 0.85 mmol H2/g microalgal DW while the outlet biohydrogen was 0.83, 0.35, 0.28 mmol H2/g microalgal DW for the microalgal media BG11+g, BG11-N+g and dist. H2O+g culture media, respectively. It was observed that the BG11 medium exhibited a conserved medium that did not negatively impact the amount of biohydrogen.

Performance of biohydrogen upgrading process

The results in Fig. 5 illustrated the performance of the upgrading process that was estimated based upon the inlet and outlet gases results. The cultivation of microalgae in BG11-N medium significantly (p ≤ 0.05) enhanced the removal of CO2 by 99% compared with H2O-grown microalgal isolate 94.8%. It was observed that there was a missing percentage of biohydrogen during the upgrading process in different microalgal culture media. The microalgal dist.H2O culture medium exhibited a significant (P ≤ 0.05) increase in the biohydrogen loss (67.1%), while the microalgae cultivated in BG11 medium exhibited a non-significant loss (5.7%) in the amount of biohydrogen compared to the inlet percentage. However, it exhibited a significant decrease in the loss of biohydrogen when compared with BG11-N and H2O microalgal media Fig. 5a, b.

Impact of upgrading process on the microalgal physiological activities

The results in Table.1 showed that the maximum biomass production of 0.21 mg/mL, 0.13 and 0.18 g L- 1 (dry mass basis) were observed when the microalgae were cultured in BG11+g, BG11-N+g and H2O+g, respectively, which indicated that Oscillatoria sp. performed better growth in BG11, which is often noticed in cyanobacterial isolates. On the other hand, the bottles that were not exposed to the gas mixture showed a negative impact on the growth and biomass production of microalgae in three culture media.

Table1 The growth parameters, carbon dioxide fixation rate and pigmentation of microalgal gown on different culture media

The maximum specific growth rate (0.023/d) and doubling time (0.076 d) were observed in microalgae growing in BG11+g Table 1. The rate of carbon dioxide fixation was detected in three tested culture media, the results showed that the BG11+g exhibited significant (p ≤ 0.05) maximum CO2 fixation rate (0.42 mg/mL/d) when compared with BG11-N+g or H2O+g.

Microalgal photosynthetic pigments play a role in industrial applications. The results in Table 1 showed that the cultivation of Oscillatoria sp. in BG11+g enhanced the production of both chlorophyll-a and carotenoid pigments to be 4.8 and 11.1 µg/mL when the content of the initial pigment before passing of gas mixture was 1.1 and 2.5 µg/mL, respectively. These results were significantly (p ≤ 0.05) high when compared with other tested microalgal culture media. The microalgae that were cultured in H2O+g showed no significant increase in the chlorophyll-a and carotenoid content (1.13 and 2.5 µg/mL) when compared with the initial ones 1.1 and 2.5 µg/mL, respectively.After the end of the purification process, the microalgal cells grown in different culture media used for biohydrogen upgrading could be reused for another process or extraction of lipids for prospective biodiesel production. The results in Table 2 showed that the lipid content was 8.7, 7.5 and 7.4% for the microalgal cultures BG11+g, BG11-N+g and dist.H2O+g respectively. Lipid yield was 2.9, 2.6 and 2.3 µg/mL respectively.

Table 2 Lipid yield and Lipid content which extracted from Oscillatoria sp. from the used cultures media BG11+g, BG11—N+g and dist. H2O+g


The fermentation method of producing biohydrogen is proven to be efficient since it is safe for the environment, reduces the need for fossil fuel consumption, and minimizes pollutants. However, biohydrogen production via dark fermentation is complex and influenced by many factors including type and pretreatment of substrate, type and source of inoculum, pH and temperature [2]. Regarding the inoculum type, B. coagulans AH1 (MN923076) is a gram-positive, facultative anaerobic bacteria isolated from wastewater sludge. As mentioned by [50], the facultative anaerobic fermentative microorganisms were cost effective during the biohydrogen production compared with strictly anaerobes. As they can enhance the anaerobic condition of the medium by fast utilization of the dissolved oxygen. In addition, many literatures discussed the effective role of sludge inhabiting bacteria for the fermentative production of hydrogen [2, 51]. In addition, the deceasing in the biohydrogen productivity when the biomass exceeded 5% may be attributed to the enhancement of metabolic and enzymatic activities of the fermentative bacteria under this concentration (5%) [4, 7, 52]. However, at high substrate concentrations, it might be rapidly converted to hydrogen and /or volatile organic acids that cause the drop in pH, consequently decreasing the metabolic activity of fermentative bacteria and reducing the produced H2. Moreover, it caused high hydrogen partial pressure in the fermentation medium that would restrict the hydrogen production efficiency [53]. In addition, an excessive amount of the substrate increased osmotic pressure and thus inhibited the growth of hydrogen-producing microbes [4, 54]. On the other hand, at low substrate concentrations, a large amount of it may be consumed for microbial growth rather than hydrogen production [4]. Notably, in this study, the biomass of C. procera was directly supplemented as a substrate for bacterial dark fermentation for H2 production without any prior treatment. Based upon our previous study, the leaf biomass of C. procera was used as a main substrate for the production of bioethanol after acid-alkaline pretreatment as well and it was reported to be applied as a promising feedstock for biodiesel production [19]. However, in the current study, the direct application of plant biomass led to avoiding the negatives of the different pretreatment strategies (i.e. high cost, energy and time consumption) [49].

The most crucial variables in the anaerobic fermentation process are temperature and pH. These variables affect the synthesis of biohydrogen and microorganisms that produce H2 [55]. In the current study, the amount of biohydrogen reduced at low pH may be due to the inhibition of hydrogenase activity, which is a key enzyme in the biohydrogen production process or inhibition of the metabolic activity of bacteria [55,56,57,58]. Moreover, [59] reported that the low pH value stimulated the solventogenesis and methanogenesis that suppress the production of hydrogen during fermentation. The optimum pH 9.0 of AH1 strains for maximum production of hydrogen was compatible with that recorded by [60, 61] using glucose as the sole substrate, but it was higher than the value of H2 at pH 7.5 that demonstrated by [55] using molasses. According to [62], pH levels above 10.0 had a toxicity impact on the microorganisms, which explained the decrease of biohydrogen production at pH 11.0 in the current study.

Besides that, dark fermentative biohydrogen production could be conducted at a broad range of temperatures, 25–80 °C [63]. The ambient temperature average for hydrogen production is from 30 to 49 °C and it is also preferred in terms of expenditures and other technical features [22]. Biohydrogen production under mesophilic conditions is cost effective, less energy consumption and easy to regulate on a large scale [52]. The production under mesophilic conditions (30–35 °C) has been reported by [60, 64] which was agreed with the current results. A lower culture or higher culture temperature deactivated or denaturized the microbial enzyme system and consequently decreased the biohydrogen production [65, 66]. The optimum conditions for biohydrogen production from Bacillus coagulans IIT-BT S1 was reported by (Kotay, and Das, 2007) were pH 6.5, temperature 37 °C and initial glucose concentration of 2% (w/v), to produce 2.28 molH2/mol glucose. The data in Table 3 showed the optimal conditions of B. coagulans, for biohydrogen production from C. procera leaves compared to those reported in other studies.

Table 3 Biohydrogen production from C. procera leaves under the optimal conditions of the bacterial strain AH1 compared to the other bacterial cultures that reported in other studies

Carbon dioxide including biohydrogen represents the major obstacle to application of biogas as an alternative fuel [2, 3]. In this study a blue-green microalgae Oscillatoria sp was used for the sequestration of carbon dioxide from the biohydrogen gas produced from bacterial dark fermentation. The produced gas mixture (H2 + CO2) passed through three different culture media containing microalgal cells. The increase in hydrogen to carbon dioxide ratio after passing of the gas mixture (outlet H2/CO2) through the microalgal media was due to the reduction in the amount of carbon dioxide, indicating its utilization by the microalgae and the occurrence of the purification process. Many literatures discussed the rate and capability of blue-green microalgae to fix CO2 via the photosynthesis process. The BG11-N media exhibited a significant (p ≤ 0.05) high ratio of outlet H2/CO2 when compared with other culture media. It may be attributed to the microalgae exploiting CO2 in the gas mixture to perform the photosynthesis process to provide energy for the production of nitrogenase enzyme and fix nitrogen to compensate for nitrogen deprivation [67]. On the other hand, dist.H2O media exhibited a significant (p ≤ 0.05) reduction in the outlet H2/CO2 may be due to the mineral deficiency and the formation of H2CO3 (CO2 + H2O) led to of that reduced the pH level and retarded the microalgal growth and metabolism [68] mentioned that the optimum pH for the growth of many microalgal species ranged from 7.2 to 8.0. These results were confirmed by determining the inlet and outlet concentration of CO2 (Fig. 5a). The amount of outlet CO2 was significantly (p ≤ 0.05) less than the inlet one, which means that the microalgal cells consumed an adequate amount of CO2 during the upgrading process, which suggested that the gas mixture had been purified and the quality of biohydrogen has been improved. According to [69], Oscillatoria sp. and other microalgae exhibited high potential of CO2 fixation, and have been broadly used for biogas upgrading and biogas slurry nutrient reduction. The appropriate CO2 level depends on the species of microalgae, the layout of the system, and the operational circumstances [70].

On the other hand, the result in (Fig. 5b) showed a decrease in the amount of outlet H2 after upgrading in BG11-N+g and dist. H2O+g may be due to the activity of the uptake hydrogenase enzyme or the consumption of H2 as an electron donor. The uptake H2ase (Hup) found in cyanobacteria absorbs H2, and interfered with nitrogenase-based H2 production [71]. It was mentioned by [72] that green algae (under anaerobic conditions) can either use H2 as an electron donor in the CO2-fixation process or evolve H2 in both the dark and the light. The losing hydrogen in the case of BG11-N medium may be due to inhibition of photosystem II that is under nitrogen deprivation conditions which increases the anaerobic conditions that activate the hydrogenase enzyme. The latter may activate in the reverse direction and increase the uptake of hydrogen. under hydrogen photoproduction conditions (N-poor medium without fixed nitrogen, anaerobic conditions) the degradation of photosystem II proteins of Lyngbya sp. was observed [73]. However, in the case of dist.H2O as a culture medium, the losing hydrogen may be due to the absence of nutrients and the use of hydrogen by Oscillatoria sp. as an energy source. The main physiological function of the uptake hydrogenase is to reutilize and regain the H2/electrons. These results came in context with the recent study of [74] that reported the role of Chlorella vulgaris in upgrading methane via decarbonation and desulfurization. Many literatures have been discussed the role of microalgae in purification of biogas from impurities as summarized in Table 4, however, to our knowledge, the highest upgrading percentage is mentioned in the current study (95–98% removal of CO2).

Table 4 The CO2 sequestration and upgrading efficiency % from biofuels by different microalgae-based technologies

Cyanobacteria are a multipurpose feedstock with several biotechnological and biorefining potential [45]. In the current study, the increase in microalgal biomass after the upgrading process may pave the way to apply the current microalgal isolate as a feedstock for biotechnological application [70]. The resulting microalgal biomass can be further utilized to produce biofuels or other value-added products via biorefinery strategies. The biomass production was significantly higher (p ≤ 0.05) in the BG11 medium when compared with other media (Table 1). These results agreed with [83], Chlorella 359 (a mutant strain) achieved maximum biomass (1.99 g/L) when it was cultivated in a media supplemented with 5% CO2. Similarly, the cultivation of Synechocystis in 0.6 g L−1 of sodium bicarbonate produced 2.24 g L−1 of biomass with 0.22 g L−1 day- 1 of CO2 fixation rate [84]. Cultivation of Dunaliella salina in 5 g L−1 NaHCO3 produced 3.17 g L−1 of biomass [85].

Three primary photosynthetic pigments that stand out in cyanobacteria Chlorophylls, carotenoids, and phycobiliproteins, play a critical role in cyanobacterial cell protection. Due to their bioactive properties, such as antioxidant, antitumoral, antiviral, etc., which can be used in the pharmaceutical, feed, and cosmetic industries as well as functional ingredients in food, these pigments have significant biotechnological significance [46]. In addition, their naturally bright colors are quite appealing to the food colorant and textile industries [86]. In the current study, the BG11 medium of Oscillatoria sp stimulated high induction of chlorophyll-a and carotenoid pigment. These results come in context with [45]. They documented that microalgal Plectonema terebrans BERC10 exhibited high production of chl-a and carotenoid after 15 days of incubation in BG11 medium.

Lipids extracted from algae are a highly valued metabolite due to their unique ability to produce biodiesel [87]. The microalgae Oscillatoria sp isolated in the current study could be used as feedstock for biodiesel production and as third-generation feedstock. it was reported that the biodiesel production is nutrient dependent, a high nutrients enhanced the lipid content of P. terebrans BERC10 from 33 to 41% [45].


It could be concluded that, the pure bacterial strains Bacillus coagulans (AH1) could produce biohydrogen by using Calotropis procera as a substrate, and their production was influenced by the three variables (concentration of C. procera as a substrate, pH values, and temperature levels). The hydrogen yield was 3.23mmol H2/g dry biomass at pH 9.0 with a substrate concentration of 5% at temperature of 35 °C, which is relatively higher than other previously mentioned pure cultures. A biological upgrade of biohydrogen was achieved using Oscillatoria sp. The CO2 removal efficiency percentage was recorded 98, 99 and 95% for the three algal cultures media BG11, BG11-N and dist.H2O, respectively. The remaining microalgal biomass exhibited pigmentation production capability and biomass feedstock for promising biodiesel production.

Availability of data and materials

The authors confirm that the data of the current study are available from the corresponding author upon reasonable request.


  1. Zhang L, Chung J, Ren N, Sun R. Effects of the ecological factors on hydrogen production and [Fe–Fe]-hydrogenase activity in Ethanoligenens harbinense YUAN-3. Int J Hydrogen Energy. 2015;40:6792–7.

    CAS  Google Scholar 

  2. Soares JF, Confortin TC, Todero I, Mayer FD, Mazutti MA. Dark fermentative biohydrogen production from lignocellulosic biomass: Technological challenges and future prospects. Renew Sustain Energy Rev. 2020;117: 109484.

    CAS  Google Scholar 

  3. Bari S. Effect of carbon dioxide on the performance of biogas/diesel duel-fuel engine. Renewable Energy. 1996;9:1007–10.

    CAS  Google Scholar 

  4. Alavi-Borazjani SA, da Cruz Tarelho LA, Capela M. Parametric optimization of the dark fermentation process for enhanced biohydrogen production from the organic fraction of municipal solid waste using Taguchi method. Int J Hydr Energy. 2021.

    Article  Google Scholar 

  5. Ahmed SF, Rafa N, Mofijur M, Badruddin IA, Inayat A, Ali MS, Farrok O, Yunus Khan TM. Biohydrogen production from biomass sources: metabolic pathways and economic analysis. Front Energy Res. 2021.

    Article  Google Scholar 

  6. Li Y, Zhang Z, Xia C, Jing Y, Zhang Q, Li S, Zhu S, Jin P. Photo-fermentation biohydrogen production and electrons distribution from dark fermentation effluents under batch, semi-continuous and continuous modes. Bioresour Technol. 2020;311: 123549.

    CAS  PubMed  Google Scholar 

  7. Yang G, Wang J. Biohydrogen production by co-fermentation of sewage sludge and grass residue: effect of various substrate concentrations. Fuel. 2019;237:1203–8.

    CAS  Google Scholar 

  8. Baeyens J, Zhang H, Nie J, Appels L, Dewil R, Ansart R, Deng Y. Reviewing the potential of bio-hydrogen production by fermentation. Renew Sustain Energy Rev. 2020;131: 110023.

    CAS  Google Scholar 

  9. Sinha P, Pandey A. An evaluative report and challenges for fermentative biohydrogen production. Int J Hydrogen Energy. 2011;36:7460–78.

    CAS  Google Scholar 

  10. Orozco RL, Redwood MD, Leeke GA, Bahari A, Santos RCD, Macaskie LE. Hydrothermal hydrolysis of starch with CO2 and detoxification of the hydrolysates with activated carbon for bio-hydrogen fermentation. Int J Hydrogen Energy. 2012;37:6545–53.

    CAS  Google Scholar 

  11. Chen H, Wu J, Huang R, Zhang W, He W, Deng Z, Han Y, Xiao B, Luo H, Qu W. Effects of temperature and total solid content on biohydrogen production from dark fermentation of rice straw: performance and microbial community characteristics. Chemosphere. 2022;286: 131655.

    CAS  PubMed  Google Scholar 

  12. Hayashi Y. Production of natural rubber from Para rubber tree. Plant Biotechnol. 2009;26:67–70.

    CAS  Google Scholar 

  13. Traore AS. Biogas production from Calotropis procera: a latex plant found in West Africa. Biores Technol. 1992;41:105–9.

    CAS  Google Scholar 

  14. Khan K, Jan G, Irfan M, Jan FG, Hamayun M, Ullah F, Bussmann RW. Ethnoveterinary use of medicinal plants among the tribal populations of District Malakand, Khyber Pakhtunkhwa Pakistan. Ethnobotany Res Appli. 2023;25:1–24.

    Google Scholar 

  15. Mahmoud O, Adam S, Tartour G. The effects of Calotropis procera on small ruminants: I. Effects of feeding sheep with the plant. J Comp Pathol. 1979;89:241–50.

    CAS  PubMed  Google Scholar 

  16. Behera B, Arora M, Sharma D. Studies on biotransformation of Calotropis procera latex-A renewable source of petroleum, value-added chemicals, and products. Energy Sources. 2000;22:781–807.

    CAS  Google Scholar 

  17. Yoganandam K, Ganeshan P, NagarajaGanesh B, Raja K. Characterization studies on Calotropis procera fibers and their performance as reinforcements in epoxy matrix. J Nat Fibers. 2019.

    Article  Google Scholar 

  18. Mukhopadhyay S, Dutta R, Das P. A critical review on plant biomonitors for determination of polycyclic aromatic hydrocarbons (PAHs) in air through solvent extraction techniques. Chemosphere. 2020;251: 126441.

    CAS  PubMed  Google Scholar 

  19. Mahmoud AH, El-Bery HM, Ali MM, Aldaby ES, Mawad AM, Shoreit A. Latex-bearing plant (Calotropis procera) as a biorefinery for bioethanol production. Biomass Conv Bioref. 2021.

    Article  Google Scholar 

  20. Kumar A. Global warming, climate change and greenhouse gas mitigation. Biofuels. 2018.

    Article  Google Scholar 

  21. Harari P, Banapurmath N, Yaliwal V, Khan TY, Soudagar MEM, Sajjan A. Experimental studies on performance and emission characteristics of reactivity controlled compression ignition (RCCI) engine operated with gasoline and Thevetia Peruviana biodiesel. Renewable Energy. 2020;160:865–75.

    CAS  Google Scholar 

  22. Chozhavendhan S, Rajamehala M, Karthigadevi G, Praveenkumar R, Bharathiraja B. A review on feedstock, pretreatment methods, influencing factors, production and purification processes of bio-hydrogen production. Case Studies Chem Environ Eng. 2020;2: 100038.

    Google Scholar 

  23. Dunikov D, Borzenko V, Blinov D, Kazakov A, Lin C-Y, Wu S-Y, Chu C-Y. Biohydrogen purification using metal hydride technologies. Int J Hydrogen Energy. 2016;41:21787–94.

    CAS  Google Scholar 

  24. Yoo C, Jun S-Y, Lee J-Y, Ahn C-Y, Oh H-M. Selection of microalgae for lipid production under high levels carbon dioxide. Biores Technol. 2010;101:S71–4.

    CAS  Google Scholar 

  25. Barati B, Zeng K, Baeyens J, Wang S, Addy M, Gan S-Y, El-Fatah Abomohra A. Recent progress in genetically modified microalgae for enhanced carbon dioxide sequestration. Biomass Bioenerg. 2021;145: 105927.

    CAS  Google Scholar 

  26. Schenk PM, Thomas-Hall SR, Stephens E, Marx UC, Mussgnug JH, Posten C, Kruse O, Hankamer B. Second generation biofuels: high-efficiency microalgae for biodiesel production. Bioenergy Res. 2008;1:20–43.

    Google Scholar 

  27. Choi YY, Patel AK, Hong ME, Chang WS, Sim SJ. Microalgae bioenergy with carbon capture and storage (BECCS): an emerging sustainable bioprocess for reduced CO2 emission and biofuel production. Biores Technol Rep. 2019;7: 100270.

    Google Scholar 

  28. Valdovinos-García EM, Barajas-Fernández J, MdlÁ O-A, Petriz-Prieto MA, Guzmán-López A, Bravo-Sánchez MG. Techno-economic study of CO2 capture of a thermoelectric plant using Microalgae (Chlorella vulgaris) for Production of feedstock for bioenergy. Energies. 2020;13:413.

    Google Scholar 

  29. Deviram G, Mathimani T, Anto S, Ahamed TS, Ananth DA, Pugazhendhi A. Applications of microalgal and cyanobacterial biomass on a way to safe, cleaner and a sustainable environment. J Clean Prod. 2020;253: 119770.

    CAS  Google Scholar 

  30. Aldaby E, Mawad A. Pyrene biodegradation capability of two different microalgal strains. Global NEST J. 2019;21:291–6.

    Google Scholar 

  31. Chia SR, Ong HC, Chew KW, Show PL, Phang S-M, Ling TC, Nagarajan D, Lee D-J, Chang J-S. Sustainable approaches for algae utilisation in bioenergy production. Renewable Energy. 2018;129:838–52.

    CAS  Google Scholar 

  32. Gill SS, Mehmood MA, Rashid U, Ibrahim M, Saqib A, Tabassum MR. Waste-water treatment coupled with biodiesel production using microalgae: a bio-refinery approach. Pak J Life Soc Sci. 2013;11:179–89.

    Google Scholar 

  33. Fields MW, Hise A, Lohman EJ, Bell T, Gardner RD, Corredor L, Moll K, Peyton BM, Characklis GW, Gerlach R. Sources and resources: importance of nutrients, resource allocation, and ecology in microalgal cultivation for lipid accumulation. Appl Microbiol Biotechnol. 2014;98:4805–16.

    CAS  PubMed  PubMed Central  Google Scholar 

  34. Gill SS, Mehmood MA, Ahmad N, Ibrahim M, Rashid U, Ali S, Nehdi IA. Strain selection, growth productivity and biomass characterization of novel microalgae isolated from fresh and wastewaters of upper Punjab Pakistan. Front Life Sci. 2016;9:190–200.

    CAS  Google Scholar 

  35. Morocho-Jácome AL, Ruscinc N, Martinez RM, de Carvalho JCM, Santos de Almeida T, Rosado C, Costa JG, Velasco MVR, Baby AR. Technological aspects of microalgae pigments for cosmetics. Appl Microbiol Biotechnol. 2020;104:9513–22.

    PubMed  Google Scholar 

  36. Grosser A. Determination of methane potential of mixtures composed of sewage sludge, organic fraction of municipal waste and grease trap sludge using biochemical methane potential assays. A comparison of BMP tests and semi-continuous trial results. Energy. 2018;143:488–99.

    CAS  Google Scholar 

  37. Soutschek E, Winter J, Schindler F, Kandler O. Acetomicrobium flavidum, gen. nov., sp. nov., a thermophilic, anaerobic bacterium from sewage sludge, forming acetate, CO2 and H2 from glucose. Syst Appl Microbiol. 1984;5:377–90.

    CAS  Google Scholar 

  38. Montoya D, Spitia S, Silva E, Schwarz W. Isolation of mesophilic solvent-producing clostridia from Colombian sources: physiological characterization, solvent production and polysaccharide hydrolysis. J Biotechnol. 2000;79:117–26.

    CAS  PubMed  Google Scholar 

  39. Wang Y, He L, Zhang Z, Zhao X, Qi N. Efficiency enhancement of H2 production by a newly isolated maltose-preferring fermentative bio-hydrogen producer of Clostridium butyricum NH-02. J Energy Stor. 2020;30:101426.

    Google Scholar 

  40. Rölleke S, Muyzer G, Wawer C, Wanner G, Lubitz W. Identification of bacteria in a biodegraded wall painting by denaturing gradient gel electrophoresis of PCR-amplified gene fragments coding for 16S rRNA. Appl Environ Microbiol. 1996;62:2059–65.

    PubMed  PubMed Central  Google Scholar 

  41. Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987;4:406–25.

    CAS  PubMed  Google Scholar 

  42. Huson DH, Bryant DJ. Application of phylogenetic networks in evolutionary studies. Mole Biol Evol. 2006;23:254–67.

    CAS  Google Scholar 

  43. Stanier R, Kunisawa R, Mandel M, Cohen-Bazire GJ. Purification and properties of unicellular blue-green algae (order Chroococcales). Bacteriol Rev. 1971;35:171–205.

    CAS  PubMed  PubMed Central  Google Scholar 

  44. Liu X, Chen G, Tao Y, Wang J. Application of effluent from WWTP in cultivation of four microalgae for nutrients removal and lipid production under the supply of CO2. Renew Energy. 2020;149:708–15.

    CAS  Google Scholar 

  45. Shahid A, Malik S, Liu C-G, Musharraf SG, Siddiqui AJ, Khan F, Tarbiah NI, Gull M, Rashid U, Mehmood MA. Characterization of a newly isolated cyanobacterium Plectonema terebrans for biotransformation of the wastewater-derived nutrients to biofuel and high-value bioproducts. J of Water Proc Eng. 2021;39: 101702.

    Google Scholar 

  46. Shahid A, Usman M, Atta Z, Musharraf SG, Malik S, Elkamel A, Shahid M, Abdulhamid Alkhattabi N, Gull M, Mehmood MA. Impact of wastewater cultivation on pollutant removal, biomass production, metabolite biosynthesis, and carbon dioxide fixation of newly isolated cyanobacteria in a multiproduct biorefinery paradigm. Biores Technol. 2021;333: 125194.

    CAS  Google Scholar 

  47. Sarkar S, Manna MS, Bhowmick TK, Gayen K. Extraction of chlorophylls and carotenoids from dry and wet biomass of isolated Chlorella Thermophila: optimization of process parameters and modelling by artificial neural network. Process Biochem. 2020;96:58–72.

    CAS  Google Scholar 

  48. Hara A, Radin NS. Lipid extraction of tissues with a low-toxicity solvent. Anal Biochem. 1978;90:420–6.

    CAS  PubMed  Google Scholar 

  49. Zadeh ZE, Abdulkhani A, Aboelazayem O, Saha B. Recent insights into lignocellulosic biomass pyrolysis: a critical review on pretreatment, characterization, and products upgrading. Processes. 2020;8:799.

    CAS  Google Scholar 

  50. Yokoi H, Ohkawara T, Hirose J, Hayashi S, Takasaki Y. Characteristics of hydrogen production by aciduric Enterobacter aerogenes strain HO-39. J Ferment Bioeng. 1995;80:571–4.

    CAS  Google Scholar 

  51. Han W, Liu DN, Shi YW, Tang JH, Li YF, Ren NQ. Biohydrogen production from food waste hydrolysate using continuous mixed immobilized sludge reactors. Biores Technol. 2015;180:54–8.

    CAS  Google Scholar 

  52. Rajesh Banu J, Mohamed Usman TM, Kavitha S, Yukesh Kannah R, Yogalakshmi KN, Sivashanmugam P. Amit Bhatnagar, Gopalakrishnan Kumar: a critical review on limitations and enhancement strategies associated with biohydrogen production. Int J Hydr Energy. 2021;46:16565–90.

    Google Scholar 

  53. Gokfiliz-Yildiz P, Karapinar I. Optimization of particle number, substrate concentration and temperature of batch immobilized reactor system for biohydrogen production by dark fermentation. Int J Hydr Energy. 2018;43:10655–65.

    CAS  Google Scholar 

  54. Lopez-Hidalgo AM, Alvarado-Cuevas ZD, De Leon-Rodriguez A. Biohydrogen production from mixtures of agro-industrial wastes: chemometric analysis, optimization and scaling up. Energy. 2018;159:32–41.

    CAS  Google Scholar 

  55. RamKumar N, Anupama PD, Nayak T, Subudhi S. Scale up of biohydrogen production by a pure strain; Clostridium butyricum TM-9A at regulated pH under decreased partial pressure. Rene Energy. 2021;170:1178–85.

    CAS  Google Scholar 

  56. Khanal SK, Chen W-H, Li L, Sung S. Biological hydrogen production: effects of pH and intermediate products. Int J hydr Energy. 2004;29:1123–31.

    CAS  Google Scholar 

  57. Yasin NHM, Man HC, Yusoff MZM, Hassan MA. Microbial characterization of hydrogen-producing bacteria in fermented food waste at different pH values. Int J Hydr Energy. 2011;36:9571–80.

    CAS  Google Scholar 

  58. De Gioannis G, Muntoni A, Polettini A, Pomi RJWM. A review of dark fermentative hydrogen production from biodegradable municipal waste fractions. Waste Manage. 2013;33:1345–61.

    Google Scholar 

  59. Kumar G, Zhen G, Sivagurunathan P, Bakonyi P, Nemestóthy N, Bélafi-Bakó K, Kobayashi T, Xu K-Q. Biogenic H2 production from mixed microalgae biomass: impact of pH control and methanogenic inhibitor (BESA) addition. Biofuel Res J. 2016;3:470–4.

    CAS  Google Scholar 

  60. Taroepratjeka DAH, Imai T, Chairattanamanokorn P. Reungsang AJCMJS: biohydrogen production by extremely halophilic bacteria from the salt pan of Samut Sakhon Thailand. Chiang Mai J Sci. 2020;47:378–90.

    CAS  Google Scholar 

  61. Kim IS, Hwang MH, Jang NJ, Hyun SH, Lee ST. Effect of low pH on the activity of hydrogen utilizing methanogen in bio-hydrogen process. Int J Hydrogen Energy. 2004;29:1133–40.

    CAS  Google Scholar 

  62. Zhao Y, Chen Y, Zhang D, Zhu XJ. Technology: waste activated sludge fermentation for hydrogen production enhanced by anaerobic process improvement and acetobacteria inhibition: the role of fermentation pH. Environ sci Technol. 2010;44:3317–23.

    CAS  PubMed  Google Scholar 

  63. Elbeshbishy E, Dhar BR, Nakhla G, Lee H-S. A critical review on inhibition of dark biohydrogen fermentation. Renew Sustain Energy Rev. 2017;79:656–68.

    CAS  Google Scholar 

  64. Kim S-H, Han S-K, Shin H-S. Feasibility of biohydrogen production by anaerobic co-digestion of food waste and sewage sludge. Int J Hydr Energy. 2004;29:1607–16.

    CAS  Google Scholar 

  65. Wang Y, He L, Zhang Z, Zhao X, Qi N, Han T. Efficiency enhancement of H2 production by a newly isolated maltose-preferring fermentative bio-hydrogen producer of Clostridium butyricum NH-02. J Energy Storage. 2020;30: 101426.

    Google Scholar 

  66. Wang Y, Zhou X, Hu J, Jing Y, Wu Q, Chang J, Lu C, Zhang Q. A comparison between simultaneous saccharification and separate hydrolysis for photofermentative hydrogen production with mixed consortium of photosynthetic bacteria using corn stover. Int J Hydrogen Energy. 2017;42:30613–20.

    CAS  Google Scholar 

  67. Do Nascimento M, Sanchez Rizza L, Arruebarrena Di Palma A, MdlA D, Salerno G, Rubio LM, Curatti L. Cyanobacterial biological nitrogen fixation as a sustainable nitrogen fertilizer for the production of microalgal oil. Algal Res. 2015;12:142–8.

    Google Scholar 

  68. Gatamaneni BL, Orsat V, Lefsrud M. Factors affecting growth of various microalgal species. Environ Eng Sci. 2018;35:1037–48.

    CAS  Google Scholar 

  69. Zhang W, Zhao C, Cao W, Sun S, Hu C, Liu J, Zhao YJES. Removal of pollutants from biogas slurry and CO2 capture in biogas by microalgae-based technology: a systematic review. Environ Sci Pollut Res. 2020;27:28749–67.

    CAS  Google Scholar 

  70. Zhu LJB. Biorefining: microalgal culture strategies for biofuel production: a review. Bioproducts. 2015;9:801–14.

    CAS  Google Scholar 

  71. Sakurai H, Tsygankov AA: Photobiological biohydrogen production. In Second and Third Generation of Feedstocks. Elsevier; 2019: 437–467.

  72. Ghirardi ML, Zhang L, Lee JW, Flynn T, Seibert M, Greenbaum E, Melis AJT. Microalgae: a green source of renewable H2. Trends Biotechnol. 2000;18:506–11.

    CAS  PubMed  Google Scholar 

  73. Kuwada Y, Inoue Y, Koike H, Ohta Y. Functional and structural changes of PSII of Lyngbya sp. under hydrogen-producing conditions. Agric Biol Chem. 1991;55:299–305.

    CAS  Google Scholar 

  74. Xie T, Herbert C, Zitomer D, Kimbell L, Stafford M, Venkiteshwaran K. Biogas conditioning and digestate recycling by microalgae: acclimation of Chlorella vulgaris to H2S-containing biogas and high NH4-N digestate and effect of biogas: digestate ratio. Chem Eng J. 2023;453: 139788.

    CAS  Google Scholar 

  75. Lopez-Hidalgo AM, Sánchez A, De León-Rodríguez A. Simultaneous production of bioethanol and biohydrogen by Escherichia coli WDHL using wheat straw hydrolysate as substrate. Fuel. 2017;188:19–27.

    CAS  Google Scholar 

  76. Arreola-Vargas J, Razo-Flores E, Celis LB, Alatriste-Mondragón F. Sequential hydrolysis of oat straw and hydrogen production from hydrolysates: role of hydrolysates constituents. Int J Hydrogen Energy. 2015;40:10756–65.

    CAS  Google Scholar 

  77. Saratale GD, Kshirsagar SD, Saratale RG, Govindwar SP, Oh M-K. Fermentative hydrogen production using sorghum husk as a biomass feedstock and process optimization. Biotechnol Bioprocess Eng. 2015;20:733–43.

    CAS  Google Scholar 

  78. Kumar N, Das DJPB. Enhancement of hydrogen production by Enterobacter cloacae IIT-BT. Proc Biochem. 2000;08(35):589–93.

    Google Scholar 

  79. Silva JS, Mendes JS, Correia JAC, Rocha MVP, Micoli L. Cashew apple bagasse as new feedstock for the hydrogen production using dark fermentation process. J Biotechnol. 2018;286:71–8.

    CAS  PubMed  Google Scholar 

  80. Muharja M, Junianti F, Ranggina D, Nurtono T, Widjaja A. An integrated green process: subcritical water, enzymatic hydrolysis, and fermentation, for biohydrogen production from coconut husk. Biores Technol. 2018;249:268–75.

    CAS  Google Scholar 

  81. Kirli B, Karapinar I. The effect of HRT on biohydrogen production from acid hydrolyzed waste wheat in a continuously operated packed bed reactor. Int J Hydrogen Energy. 2018;43:10678–85.

    CAS  Google Scholar 

  82. Mullai P, Yogeswari M, Sridevi K. Optimisation and enhancement of biohydrogen production using nickel nanoparticles–a novel approach. Biores Technol. 2013;141:212–9.

    CAS  Google Scholar 

  83. Chou HH, Su HY, Song XD, Chow TJ, Chen CY, Chang JS, Lee TM. Isolation and characterization of Chlorella sp mutants with enhanced thermo- and CO2 tolerances for CO2 sequestration and utilization of flue gases. Biotechnol Biofuels. 2019;12:251.

    PubMed  PubMed Central  Google Scholar 

  84. Ashokkumar V, Chen W-H, Ngamcharussrivichai C, Agila E, Ani FN. Potential of sustainable bioenergy production from Synechocystis sp. cultivated in wastewater at large scale—a low cost biorefinery approach. Energy Conv Manage. 2019;186:188–99.

    CAS  Google Scholar 

  85. Kim G-Y, Heo J, Kim H-S, Han J-I. Bicarbonate-based cultivation of Dunaliella salina for enhancing carbon utilization efficiency. Biores Technol. 2017;237:72–7.

    CAS  Google Scholar 

  86. Assunção J, Amaro HM, Malcata FX, Guedes AC. Chapter 8 - Cyanobacterial pigments: photosynthetic function and biotechnological purposes. In: Lopes G, Silva M, editors. The Pharmacological Potential of Cyanobacteria. Vasconcelos V: Academic Press; 2022. p. 201–56.

    Google Scholar 

  87. Yadav G, Sekar M, Kim S-H, Geo VE, Bhatia SK, Sabir JS, Chi NTL, Brindhadevi K, Pugazhendhi A. Lipid content, biomass density, fatty acid as selection markers for evaluating the suitability of four fast growing cyanobacterial strains for biodiesel production. Biores Technol. 2021;325:124654.

    CAS  Google Scholar 

  88. Tongprawhan W, Srinuanpan S, Cheirsilp B. Biocapture of CO2 from biogas by oleaginous microalgae for improving methane content and simultaneously producing lipid. Biores Technol. 2014;170:90–9.

    CAS  Google Scholar 

  89. Zhang Y, Bao K, Wang J, Zhao Y, Hu C. Performance of mixed LED light wavelengths on nutrient removal and biogas upgrading by different microalgal-based treatment technologies. Energy. 2017;130:392–401.

    CAS  Google Scholar 

  90. Yan C, Zhu L, Wang Y. Photosynthetic CO2 uptake by microalgae for biogas upgrading and simultaneously biogas slurry decontamination by using of microalgae photobioreactor under various light wavelengths, light intensities, and photoperiods. Appl Energy. 2016;178:9–18.

    CAS  Google Scholar 

  91. Zhao Y, Sun S, Hu C, Zhang H, Xu J, Ping L. Performance of three microalgal strains in biogas slurry purification and biogas upgrade in response to various mixed light-emitting diode light wavelengths. Biores Technol. 2015;187:338–45.

    CAS  Google Scholar 

Download references


The authors would like to thank Dr Ibrahim M. A. Nafady the manager of Wadi Al-Asiuty protected area for helping in collecting the Calotropis procera plant. Also, this work was supported by the Science, Technology & Innovation Funding Authority (STIFA) in Egypt as a part of research projects (ID: 41623,43281). The authors are indebted to Prof. Ahmed Geies, former Assiut University president, for his support.


Open access funding provided by The Science, Technology & Innovation Funding Authority (STDF) in cooperation with The Egyptian Knowledge Bank (EKB). This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.

Author information

Authors and Affiliations



AS was responsible for conceptualization and supervising, AMMM and EA were responsible for writing—the original draft; writing—review & editing, HE and AM were responsible for data curation; formal analysis; investigation; and methodology. MA was responsible for reviewing data and statistical analysis.

Corresponding author

Correspondence to Haitham M. El-Bery.

Ethics declarations

Ethics approval and consent to participate

This article does not contain any studies with human participants or animals performed by any of the authors.

Competing interests

All authors confirm that there is no conflict of interest in publishing this manuscript.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Aldaby, E.S.E., Mahmoud, A.H.A., El-Bery, H.M. et al. Microalgal upgrading of the fermentative biohydrogen produced from Bacillus coagulans via non-pretreated plant biomass. Microb Cell Fact 22, 190 (2023).

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: