Microalgae Cultivation Technologies for Bioenergetic System Development: History
View Latest Version
Please note this is an old version of this entry, which may differ significantly from the current revision.
Contributor:
Marcin Dębowski
,
Marcin Zieliński
,
Joanna Kazimierowicz
,
Natalia Kujawska
,
Szymon Talbierz

Microalgal biomass is considered as a sustainable and renewable feedstock for biofuel production (biohydrogen, biomethane, biodiesel) characterized by lower emissions of hazardous air pollutants than fossil fuels. Photobioreactors for microalgae growth can be exploited using many industrial and domestic wastes. It allows locating the commercial microalgal systems in areas that cannot be employed for agricultural purposes, i.e., near heating or wastewater treatment plants and other industrial facilities producing carbon dioxide and organic and nutrient compounds. 

  • microalgal biomass
  • microalgae cultivation
  • biofuels

1. Introduction

Microalgae are single-cell organisms that convert solar radiation energy into chemical energy via photosynthesis [1]. Controlled production of microalgal biomass is a fast-growing technology, as microalgae can be used to produce a wide range of commercially valuable cellular metabolites, including high-quality proteins, lipids, carbohydrates, dyes, and vitamins for the food/feed industry and the broad cosmetic industry (Table 1).
The fact that microalgae represent an alternative and competitive source of biomass is due to their advantage over typical terrestrial and energy plants [2]. Algae possess very high photosynthetic efficiency [3], can relatively fast build biomass [4], are resistant to various contaminants [5], and can be grown on land that is unsuitable for other purposes [6]. Microalgae production systems can also be used in environment-protecting technologies [7], including sewage and leachate treatment [8], neutralization of waste and sludge [9], carbon dioxide biosequestration, biogas upgrading, and flue gas treatment [10] (Table 2). This makes it possible to select and adapt specific strains for individual applications, including energy carrier production, environmental protection, and environmental engineering technologies [11]. Given these considerations, algae may provide a viable alternative to traditional energy crops [12].
Table 1. Venues for microalgal biomass application.

Product

Use

Ref.

Agar

Food ingredient, fruit preserves, hydrocolloids, clarifying brewing agent, paper industry, and others

[13]

Alginate

Food additive, medical, pharmaceutical, paper, cosmetic and fertilizer industries, textile printing

[1]

Antioxidants

Preservatives in cosmetic, chemical, food, and pharmaceutical industries

[14]

Astaxanthin

Food supplement as food dye additive and antioxidant

[15]

Beta-carotene and carotenoids

Precursor for vitamin A and supplement for vitamin C, food additive as coloring agent, and antioxidant

[16]

Bioenergy and biofuels

Biodiesel, bioethanol, biogas, biohydrogen, biomethane, aviation gas, biobutanol, biosyngas, bio-oil, gasoline, solid fuel, jet fuel

[15]

Biochar

Agricultural and sorbent uses, combustion

[13]

Biorefinery

Various chemicals and biofuels

[14]

Biosorbents

Ion exchange materials that bind strongly heavy metal ions

[1]

Carragen or carrageenan

Pet food, food additive, gels, toothpaste

[1]

Catalysts

Catalytic properties

[14]

Chemicals

Industrial and medicinal uses

[1]

Conditioners

Chemical, cosmetic, and farming industries

[15]

Digester residue

Compost or vermicompost

[13]

Extraction of

hydrocolloids or gums

Food industry, phytocolloids such as agar, alginate, and carrageenan

[14]

lipids carbohydrates starch and cellulose

Biogas, biodiesel, gasoline, jet fuel, alcohols, renewable hydrocarbons

minerals and trace elements

Food supplements, glass production, metallurgy

proteins

Fertilizers, industrial enzymes, animal/fish feeds, surfactants, bioplastics

Feed

Animal food

[15]

Fertilizers

N-, P-, and K-rich fertilizers

[1]

Phytosterol

Food supplements

[15]

Pigments

Natural colorants in paper and textile industries

[14]

Production

Cosmetic

Water-binding agents and antioxidants, “skin foods”

[15]

Food and drink

Nori, kombu, wakame, cheese, soup, noodles, pasta, wine, tea, others

Fruit and vegetable preservatives

Food industry

Glass

Glass industry

Paper pulp supplements

Paper industry

Textile

Textile industry

Therapeutic materials

Pharmaceutical industry
Table 2. Applications of microalgae in environment-protecting technologies.

Sector

Use

Ref.

Wastewater treatment

Nitrogen and phosphorus removal from municipal wastewater

[17]

Biodegradation of sparingly degradable pollutants

[18][19]

Treatment of organic wastewater

[20]

Treatment of hard-to-manage wastewater produced by

timber and paper industry

[21][22]

textile industry

[23]

phenol industry

[24][25]

Ethanol and citric acid production

[26]

Removal of heavy metals (copper, nickel, lead) from wastewater

[19][27]

Gas treatment

Reducing emissions of carbon dioxide and other pollutants (nitrogen and sulfur oxides) from waste and exhaust gases

[28][29]

Waste management

Use of waste glycerol as a carbon source in heterotrophic cultivation

[30]

Microalgae cultivation using industrial waste or low-value feedstocks, such as

breadcrumbs

[31]

brewer’s spent yeast

[32]

coconut water

[33]

empty palm fruit bunches

[34]

Leachate treatment

Biodegradation of landfill leachates

[35]

Neutralization of degraded effluent from anaerobic fermentation of sewage sludge

[36]

Biogas upgrading

Biological sequestration of CO2 with photosynthetic microalgae (photosynthesis allows producing biogas with 94% methane content)

[37]
However, the most promising frontiers for microalgae concern their utility for energy purposes, including the production of biogas, biohydrogen, bioethanol, and biodiesel [38]. Microalgal biomass is undoubtedly a promising substrate for energy carrier production, characterized by lower pollutant emission levels compared to conventional fuels [39][40]. By way of example, forecasts for the United States (US) biofuel and biodiesel market [41][42] are shown in Figure 1.
Figure 1. Growth forecasts for the United States of America (USA) algae-based biofuel market by 2025 ((a)—algae biofuel market, (b)—algae oil market).
Systems for producing algal biomass feature high technological efficiency, owing to the significant photosynthesis efficiency of algae and the relatively fast growth of algal biomass [43]. Phototrophic cultivation of microalgae in photobioreactors can process waste from industrial and municipal sources, which means that commercial microalgae cultivation systems can be constructed on land unsuitable for agricultural use, near heating/cogeneration plants, sewage treatment plants, and other industrial facilities that produce carbon dioxide and biogenic compounds [44].
Despite the demonstrated utility of microalgal biomass-based systems for the bioenergy industry, most industrial microalgae cultivation plants established and extensively described in the literature deal mainly with the production of high-quality feed/food additives, precious dyes, or fertilizing substances (Table 1), due to the difficulties in conclusively assessing and balancing methods for microalgal biomass production and technologies for converting it to energy carriers [45]. The majority of studies were carried out in laboratory conditions, with semi-industrial (pilot-scale) projects being a rarity [46].
The commercialization of technology and its transfer from laboratory conditions to a technical scale requires extensive research, conceptual, operational, and marketing works that allow the product to be finally placed on the market. Although relative studies present various models of knowledge and technology commercialization, they also show some similarities, as they involve a certain repetitive group of activities [47]. An important element in the process of making investment decisions regarding the commercialization of innovative products is to assess the maturity of new technologies. This assessment, called “technology readiness assessment” (TRA), should take into account the state of work on the development of a new product/technology, prospects for further development, the amount of funds necessary to invest, and innovative risk. It is a universal metric used to analyze the state of work on technologies and their readiness for commercial implementation. In turn, the “technology readiness level” (TRL) methodology sets nine levels of technology readiness and allows assessing the progress of works on new technologies [48]. It was first used in research and development (R&D) projects carried out by the National Aeronautics and Space Administration (NASA) and the US defense industry. According to TRL, technology maturity is described from the conceptualization phase of a specific solution (TRL 1) to the maturity stage (TRL 9), when this concept (as a result of research and development works) takes the form of a technological solution that can be implemented in practice, e.g., by launching production and marketing [49].
As such, there are very few sources of reliable data for a comprehensive evaluation of the technological, environmental, and economic efficiency of these solutions [50]. Such assessments are further complicated because various researchers have presented contradictory conclusions on microalgal biomass productivity, as well as its actual technological performance and cost-effectiveness. Lardon et al. (2009) unfavorably compared microalgal cultivation with traditional production methods, concluding that it is not a financially viable means of biodiesel production due to very high costs of biomass cultivation, harvesting, and drying, as well as of oil extraction [51]. On the other hand, Clarens et al. (2011) demonstrated the opposite, obtaining a positive energy balance and a beneficial environmental outcome for the biodiesel produced from microalgal biomass. They used exhaust gases and wastewater as sources of carbon dioxide and biogenic compounds for the growth medium [52]. In turn, Frank et al. (2011) used computational software to create a model that demonstrated microalgal fuel production technologies to be less energy-efficient and producing more greenhouse gases than traditional biofuel production methods [53].

2. Microalgal Biomass as a Source of Biofuels

Microalgae can serve as a potential source of many different types of biofuels (Figure 2). Examples include anaerobic digestion of biomass into biogas, production of biodiesel from lipids stored in algae cells and hydrogen from photobiological conversion, and lastly, gasification, pyrolysis, or direct combustion of the harvested algal biomass [54][55].
Figure 2. Available mechanisms for producing biofuel with microalgae.
The simplest way to use microalgae for fuel purposes involves the combustion or co-combustion of their pre-dried biomass [56]. However, this solution is rarely practiced, most often in cases where the biomass of microalgae cannot be used to produce more advanced biofuels [57]. Biogas and biomethane are produced during controlled, anaerobic degradation of microalgal biomass by fermentation bacteria [58]. Methane fermentation is a cascade of successive biochemical transformations, including hydrolysis, acidogenesis, and methanogenesis, which are carried out by specialized consortia of microorganisms [59]. In turn, biodiesel is produced via the transesterification of bio-oil extracted from microalgal biomass. This process involves the reaction of triglyceride molecules, bio-oil components, with low-molecular-weight alcohols in the presence of catalysts [60]. Hydrogen production by microalgae is based on direct biophotolysis, which involves the photosynthetic production of hydrogen from water, which uses the energy of light to break down the water molecule into hydrogen and oxygen. The process is mediated by hydrogenase—a metal enzyme that catalyzes the reversible oxidation of H2 and releases gaseous hydrogen by reducing protons [61]. The basic technology for bioethanol production from microalgae entails a biochemical process in which bacteria hydrolyze the biomass and then yeast convert the sugars present in the biomass into alcohol, which is then distilled and dehydrated [62]. In turn, syngas and pyrolytic gas are produced via the endothermal conversion of biomass into gas, which mainly consists of hydrogen, carbon monoxide, carbon dioxide, methane, and low-molecular-weight hydrocarbons [63]. The contribution of individual products, including their qualitative composition, depends mainly on the process conditions, such as temperature, reaction time, pressure, and biomass characteristics [64].
There are also reports describing technologies that stimulate and increase fatty compound storage through controlling the concentration of nitrogen compounds in the growth medium, adjusting the supply of light energy, regulating the temperature conditions, and changing the CO2 levels [65][66]. Essential prerequisites for cost-effective biodiesel production include the development of economically feasible technologies for separation/thickening of algal biomass, as well as oil extraction methods [67]. The temperature of the extraction process is a crucial factor that directly affects the quality and quantity of the resultant oil [68]. At temperatures of 60 °C and lower, higher triglyceride levels are achieved, and oil losses are reduced. Although the common practice of lipid extraction is mainly based on the use of organic solvents, some alternative and competitive technologies are still being sought. Other independent methods that aid the extraction process include mechanical, chemical, and biological treatments [69]. Despite being simple, environmentally friendly, and cheap, the mechanical methods offer a low lipid recovery efficiency [70]. Thus, intensive research works are in progress on the use of ionic liquids, supercritical fluids, bio-based extractants, and switchable solvents with simultaneous attention paid to reducing the energy consumption of the process by eliminating the energy-intensive drying process and the integration of multiple downstream processing steps [71]. A prospective solution for lipid recovery is offered by hybrid methods, e.g., enzymatic and mechanical/solvent extraction [72]. The selection of a suitable method for efficient lipid extraction largely depends on the biology and cell-wall characteristics of microalgae [73].
Technologies for converting algal biomass into energy carriers can be divided into two main groups related to thermochemical and biochemical processing [74][75]. Gasification is one of the thermochemical routes, wherein biomass is partially oxidized at temperatures ranging from 800 to 1000 °C [76]. This technological solution entails reacting the biomass with oxygen and water vapor, which directly results in the generation of syngas—a mixture of CO, H2, CO2, N, and CH4 [77]. Syngas has a low calorific value, ranging from 4.0 to 6.0 MJ·m−3, and can be combusted directly or used as a fuel in gas turbines and gas engines [78]. The properties and parameters of the microalgal biomass gasification process have been identified by several researchers. A study by Hirano et al. (1998) examined the gasification of Spirulina sp. algae at temperatures between 850 and 1000 °C and compared the obtained energy value of syngas with that of methanol. The highest operational performance was achieved with a gasification temperature of 1000 °C [79]. Minowa and Sawayama (1999) gasified Chlorella vulgaris algae within a novel technological system, producing high-methane biofuel, as well as a fertilizer rich in ammonium nitrogen [80].
A different technology for obtaining liquid biofuel is based on thermochemical liquefaction of algal biomass [81]. The process is conducted at 300–350 °C and 5.0–20.0 MPa Thermochemical reactions are induced in the presence of hydrogen, which serves as a catalyst [82]. The reactors are complex, both design- and technology-wise, which directly affects the construction and operation costs [83]. Dote et al. (1994) successfully used the featured technology to process Botryococcus braunii algae and obtained an oil yield of 64.0% dry matter of the algae fed into the reactor. The heating value of the bio-oil was 45.9 MJ·kg−1, with a positive energy balance achieved across the entire process [84]. In a similar experiment with Dunaliella tertiolecta, a bio-oil recovery yield reached 42.0% dry algal biomass, and the calorific value of the resulting product was 34.9 MJ·kg−1 [85].
Pyrolysis is yet another technology used to convert algal biomass into biofuel. Compared with the other methods presented in the literature, it has been widely described as a promising technology that yields very good results, inspiring high hopes for application in full-scale installations [86]. Miao and Wu (2004a) used pyrolysis to extract oil from heterotrophic cultures of Chlorella prothothecoides microalgae and achieved a bio-oil yield of 57.9% algal dry matter, with the calorific value of the resultant biofuel averaging 41.0 MJ·kg−1 [87]. By comparison, Miao et al. (2004b) produced bio-oil having a calorific value of 30.0 MJ·kg−1 at a yield of 18.0% dry Chlorella prothothecoides biomass and 29.0 MJ·kg−1 at a yield of 24.0% dry Microcystis aeruginosa biomass. The algae were grown in autotrophic conditions [88].
Demirbas (2006) experimented with the pyrolysis of Chlorella prothotecoides algae, aiming to ascertain how the efficiency of the process changed with temperature. The efficiency of oil recovery from pyrolyzed algal dry matter increased from 5.7% to 55.3% as the temperature rose from 254 to 502 °C. Further increases in temperature led to a direct reduction in production yields. The heating value of the harvested bio-oil peaked at 39.7 MJ·kg−1 [89]. Many of the findings published in the literature seem to indicate that bio-oil extracted from algal biomass is higher in quality than the biofuel obtained through pyrolysis of lignocellulosic plants [89][90].
Algae can also serve as a source of ethyl alcohol. It has been demonstrated that Chlorella sp. algae are viable candidates for effective alcoholic fermentation due to their high starch content (approximately 37.0% dry matter). Experimental data indicate a carbohydrate-to-ethanol conversion rate of 65.0% [91]. Ueno et al. (1998) corroborated the feasibility of ethanol production using microalgae harvested from a heterotrophic culture. The productivity of the alcoholic fermentation process performed at 30 °C was 450 μmol·g−1 dry matter [92]. The research carried out to date confirms that the production of ethyl alcohol from algal biomass can be technologically and commercially viable under specific conditions. In most cases, however, alcoholic fermentation is used as a supplemental technological step for processing algal biomass residues from the oil extraction process [93].
Hydrogen is a naturally occurring molecule that can serve as a clean and efficient energy carrier. Studies have confirmed that microalgae possess the genetic, metabolic, and enzymatic properties required to produce H2 through biochemical conversion [94]. Under anaerobic conditions, eukaryotic algae generate hydrogen as an electron donor in their metabolic pathways as part of the CO2 fixation process. This mechanism has been found to occur both in the light and in the absence of any light sources [95]. During photosynthesis, algae convert the water molecule into a hydrogen ion (H+) and oxygen. The H+ ions are then converted by hydrogenase into molecular hydrogen (H2) under anaerobic conditions [96]. It has been demonstrated that, if photosynthesis is initiated and oxygen is present in the photosynthetic environment, inhibition of the key enzyme (hydrogenase) follows shortly, directly affecting hydrogen production by algae [97].
Most of the scientific publications on this subject reported that the single-cell Chlamydomonas reinhardtii algae, commonly found in soil and saltwater, can produce H2 with high efficiency [98][99]. The hydrogen production capacity of 21 green algae species in an isolated anaerobic environment was also examined. The most productive strains were C. reinhardtii, C. euryale, C. noctigama, C. vectensis, C. pyrenoidosa, Oocystis, D. subspicatus, and P. subcapitata. Publications reported H2 yields of 90–110 cm3 H2·dm−3 for these organisms, with even higher levels of 80–140 cm3 H2·dm−3 reached in some cases [100]. Ample publications have shown that Platymonas subcordiformis algae can be used for the technological production of biohydrogen. The method employs alternating dark and light cycles with external carbon dosing, and it can produce H2 yields of 78.0 cm3 H2·dm−3 to as high as 126 cm3 H2·dm−3 [101][102].
Methane fermentation can also be employed to convert algal biomass into a gaseous energy carrier through biochemical processes. According to available estimates, the conversion of algal biomass into biogas is a highly cost-effective and commercially viable technological solution comparable to cellular lipid extraction in terms of harvested energy [103][104]. In addition to high-energy biogas, the process also produces digestate, which can be used directly as a fertilizer for terrestrial plants or reintroduced into the algal biomass route as a medium component after simple processing [105].
The practical limitations of technological processes involving methane fermentation of algae may stem from their biochemical composition. Algal biomass mostly consists of proteins and, thus, may lead to deficient C:N ratios. This problem can be greatly alleviated through the co-digestion of the algal biomass with organic substrates rich in carbon compounds. Yen and Brune (2007) achieved a substantial increase in methane production by co-digesting cellulose waste with algal biomass. The methane production rate rose to 1170 ± 75 cm3·dm−3·day−1 at a 1:1 ratio of organic waste and algal biomass, as compared to 573 ± 28 cm3·dm−3·day−1 achieved for mono-digestion of algae alone [106].
The high protein content of the algal biomass may lead to an increased production of free ammonia, which is toxic to the methane-fermenting microorganisms. Methanogenesis can also be inhibited by the sodium ions present in the algal biomass from saltwater-based cultivation systems. However, some studies show that anaerobic sludge microorganisms can be adapted and incorporated into the process for the efficient digestion of marine algal biomass [107][108].
Many researchers have argued that methane fermentation is the most promising and effective method for producing energy from algae. Sialve et al. (2009) found that, given suitable operating conditions, methane fermentation as a primary method of algal biomass processing is more economical than systems that incorporate lipid extraction and anaerobic processing of post-extraction residues [104]. Other findings suggest that the balance of methane fermentation unit operations is the most effective in terms of both the economy of the process and the pollution levels [109]. Studies have indicated that methane fermentation may be the most practical means of converting algal biomass into energy. However, Börjesson and Berglund (2006) noted that energy inputs and environmental impact varied greatly between the different methane fermentation technologies [110]. As such, an environmental life-cycle assessment (LCA) is necessary for a complete and objective evaluation of each process [111].
To meet the current challenges related to the circular bioeconomy, it is necessary to change the approach to biorefinery processes [112]. Technological, economic, and environmental efficiency improvements can be achieved by simultaneously producing many high-value products other than biofuels [113][114]. Research and development works must, therefore, be focused on finding new, more complex, and integrated production processes. Although various strategies have been proposed for converting algal biomass into fuel and fine chemicals, none have been proven to be economically viable and energy balanced [115]. Therefore, other, valuable biological products should also be searched for. In this context, the concept of microalgae biorefineries emerged with the concept of recovering multiple products from one operating process. Considering the biorefinery complexity index (BCI) as an indicator of technical and economic risk, one of the most promising seems to be the biorefinery platform based on microalgal biomass conversion into fuels, food, dietary and feed supplements, fertilizers, and pharmaceuticals [116]. A schematic diagram of a comprehensive biorefinery approach to the processing of microalgal biomass is presented below (Figure 3).
Figure 3. A schematic diagram of a comprehensive biorefinery approach to microalgal biomass processing.

3. Systems of Microalgae Species Cultivation for Biofuel

The growth rate of microalgae and their composition is influenced by the growth conditions and the species employed [117][118][119]. Many classification schemes categorize methods and technologies used to cultivate algae for biofuel [120][121]. Due to the specific nature of microalgae, the most important scheme divides the systems on the basis of the nutrient source and the type of biochemical processes used to grow the algal biomass rapidly. With this criterion in mind, cultures can be divided into four main types: photoautotrophic, heterotrophic, mixotrophic, and photoheterotrophic [122].
In a photoautotrophic culture, microalgae use light as their source of energy, as well as carbon dioxide and water to synthesize organic compounds [123]. This type of algae culture is most commonly used for commercial applications [124]. Studies have shown photoautotrophic cultures to exhibit great variability in algal biomass lipid content, with values ranging from 5% to 68% depending on the tested strain. A study with Chaetoceros calcitrans CS 178 showed a lipid production rate of rLIP = 17.6 mg·dm−3·day−1 and a final lipid content of 39.8% dry matter [125]. In contrast, a Botryococcus braunii UTEX 572 culture ended in the lipid production yield of rLIP = 5.5 mg·dm−3·day−1 [124]. The highest productivity was obtained in a study that tested the impact of high concentrations of CO2 on biomass growth and lipid synthesis in Chlorella sp. culture. The final lipid concentration reached 32–34% cell dry matter, with a maximum lipid production rate of rLIP = 179.8 mg·dm−3·day−1 [126].
Like bacteria and fungi, some microalgae species are capable of heterotrophic growth using organic carbon sources, such as glucose and glycerol [127][128]. Heterotrophic cultivation can be used to avoid the problem endemic to photoautotrophic systems, i.e., overgrown photobioreactor surfaces and the microalgal growth blocking its own light source, thus limiting the energy supply necessary for efficient photosynthesis, biomass growth, and lipid synthesis [123]. Heterotrophic cultures are characterized by higher growth rates and final biomass/lipid concentrations than the phototrophic or mixotrophic cultures. For example, a heterotrophic culture of Crypthecodinium cohnii—a strain known for its ability to biosynthesize omega-3 acids—grown on a complex medium of glucose, acetic acid, and yeast extract, produced final concentrations of 109 g·dm−3 dry biomass and 61 g·dm−3 lipids in the culture [129].
Changing the culture conditions from photoautotrophic to heterotrophic can increases lipid content per cell dry matter for some microalgal strains. For example, a 40% increase in lipid content was observed in a Chlorella protothecoides culture after the cultivation scheme was changed from photoautotrophic to heterotrophic [130]. In another study, changing the conditions from phototrophic to heterotrophic led to an over tenfold reduction in the final biomass concentration in a C. vulgaris ESP-31 culture [131]. In the lipid analysis of Chlorella protothecoides cultures, Caporgno et al. (2019) achieved fatty acid contents at 11.8% ± 0.1% dry weight (DW) and below 6% DW under heterotrophic and photoautotrophic conditions, respectively [132]. Sim et al. (2019) also observed an increased lipid production by Chlorella protothecoides. It reached 18.4% ± 0.4% DW under conditions of the heterotrophic culture and 15.1% ± 0.3% DW under photoautotrophic conditions [133]. Shen et al. (2019) demonstrated an increase in fatty acid production by Chlorella vulgaris that ranged from 14.9 ± 2.1 mg·dm−3·day−1 under photoautotrophic conditions to 51.4 ± 14.6 mg·dm−3·day−1 in the heterotrophic culture [134]. Li et al. (2016) obtained maximum biomass production in the photoautotrophic culture of Chlorella sorokiniana, reaching 0.36 ± 0.01 g·dm−3 at a specific growth rate of 0.60 ± 0.01 day−1. Under heterotrophic conditions, the respective values were 2.78 ± 0.06 g·dm−3 and 1.56 ± 0.02 day−1 [135]. In turn, Zheng et al. (2012) proved that the growth rate, cell density, and productivity of heterotrophic Chlorella sorokiniana were 3.0, 3.3, and 7.4 times higher than their phototrophic counterpart, respectively [136]. Lastly, Li et al. (2014) achieved the lipid content at 9.0% DW in the photoautotrophic culture of Chlorella sorokiniana and at 6.2% to 17.6% DW in the heterotrophic cultures [137].
Microalgae have been shown to take up many different organic carbon sources, including glucose, acetate, glycerol, fructose, sucrose, lactose, galactose, and mannose [138][139]. De Swaaf (2003) presented a study examining the use of different organic substrates in a heterotrophic culture, utilizing acetic acid and its feeding regime in a pH-controlled culture to grow Crypthecodinium cohnii [129]. This technological solution resulted in very high values of the final productivity parameters, i.e., final cell dry matter concentration at 109 g·dm−3 and 61 g·dm−3 lipids in the culture. Other studies showed Chlorella protothecoides to be capable of growth in a batch culture with crude glycerol as the sole carbon source in the medium, with the final biomass concentration at 23.5 g·dm−3 and the final lipid concentration at 14.6 g·dm−3 after a 6 day cultivation [140]. In turn, a semi-continuous batch-fed regime allowed increasing the lipid production rate to 3 g·dm−3·day−1 [140].
However, heterotrophic cultivation certainly has its disadvantages, including the frequent contamination of the culture with other strains of microalgae, fungi, and bacteria, reducing the final productivity of the technology and, in some cases, inhibiting fermentation [130][141][142]. One instance of this problem was described by Zhang et al. (2012) who investigated the impact of bacterial contamination on the dry biomass yield and lipid productivity in a heterotrophic culture of Chlorella pyrenoidosa, with soybean-processing wastewater used as a medium. On the one hand, the introduction of bacteria improved nitrogen and phosphorus degradation rates while reducing the chemical oxygen demand. On the other hand, the bacteria also reduced the final concentrations of microalgal biomass and lipids [143]. One of the methods used to avoid contamination of heterotrophic microalgal cultures entails spiking the medium with antibiotics, such as chloramphenicol [144].
In the mixotrophic cultivation, microalgal cells perform photosynthesis with simultaneous uptake of organic and inorganic carbon substrates [145]. Microalgae absorb organic compounds, and the CO2 released through respiration is captured and reused as a substrate for photosynthesis [146]. Unlike phototrophic and heterotrophic systems, the mixotrophic cultivation is rarely employed for the production of microalgae-derived bio-oil. One example of a mixotrophic culture was found in a study by Bhatnagar et al. (2011), who examined the growth rates of Chlamydomonas globosa, Chlorella minutissima, and Scenedesmus bijuga in the three most common cultivation modes. Supplementing Chlamydomonas globosa, Chlorella minutissima, and Scenedesmus bijuga cultures with 1% (w/v) glucose was found to increase mixotrophic biomass yields 9.4, 6.7, and 5.8 times (respectively) compared to the phototrophic culture and 3.0, 2.0, and 4.4 times compared to the heterotrophic culture [147]. Yu et al. (2009) obtained similar results, demonstrating that the growth rates of Nostoc flagelliforme biomass in glucose-amended media were the highest in the mixotrophic culture, with productivity values 5.0 and 2.3 times those obtained in the phototrophic and heterotrophic cultures, respectively [148].
Though microalgal oil yields are in large part determined by the choice of strain, the heterotrophic cultivation is the most effective solution in terms of the final operational performance, i.e., the biomass concentration in the system and lipid content in cells. As such, the heterotrophic method has generated strong interest among companies involved in the commercialization of bioenergy technologies and research teams working to develop such systems [149]. The most serious drawback of this scheme is the risk of culture contamination with other microorganisms, including other microalgae, which leads to severe complications with the operation of industrial-scale installations [144]. Moreover, the high cost of pure organic carbon sources limits the utility of this cultivation mode to the production of secondary or primary metabolites with a high market value [150].
Photoautotrophic cultures are the most widespread mode of cultivation, easy to scale up through the use of open or hybrid systems [151]. It is also a promising method, due to the capability of photoautotrophic microalgae for the uptake of waste CO2, such as that generated by cogeneration plants, breweries, or biogas plants. However, the oil yields produced via this method are usually vastly inferior to the heterotrophic cultivation, with slow cell growth and low biomass productivity as the main reasons. Nevertheless, with this mode being cheaper to scale up, it is highly attractive to investors despite the flaws.
The defining feature of photoheterotrophic cultivation is the use of light, required for the absorption and decomposition of organic carbon. The main difference between mixotrophic and photoheterotrophic modes is that the latter requires light as an energy source, whereas mixotrophic cultivation uses organic compounds for the same purpose. Therefore, photoheterotrophic cultivation requires a combined supply of carbohydrates and light [127]. Although photoheterotrophic systems can be used to increase the production of certain expensive secondary metabolites, the method has not found use in the production of biodiesel, as is the case with mixotrophic microalgal cultures [152].
Prior to undertaking any metabolic engineering work in microalgae, it is necessary to understand the key enzymes involved in the metabolic pathway and the rate-limiting enzymes. Many advances have been made toward understanding lipid metabolism and regulatory factors in soybean and rapeseed, but the lipid production in microalgae at a molecular level is currently very poorly understood. The first step in de novo synthesis of triacylglycerol in microalgae starts in the plastid, where pyruvate is produced from glycolysis and the Calvin cycle. The pyruvate is converted into acetyl-CoA by the pyruvate dehydrogenase complex (PDC). Acetyl-CoA is converted into malonyl-CoA by acetyl-CoA carboxylase (ACCase). Acetyl-CoA carboxylase is the rate limiting enzyme for lipid biosynthesis [153]. Malonyl-CoA is converted into malonyl-ACP by malonyl-CoA transacylase (MAT) [154]. Malonyl-ACP and acyl-ACP are converted into 3-ketoacyl-ACP by 3-ketoacyl-ACP reductase (KAS) in the fatty acid synthesis cycle. 3-Ketoacyl-ACP is converted into 3-hydroxyacyl-ACP by 3-ketoacyl-ACP reductase (KAR). 3-Hydroxyacyl-ACP is converted into trans-enoyl-ACP by 3-hydroxyacyl-ACP dehydratase (HD). trans-Enoyl-ACP is converted into acyl-ACP by enoyl-ACP reductase (ENR). Acyl-ACP is converted into free fatty acids (FFAs) by fatty acyl-ACP thioesterase (FAT) [155][156]. The FFAs are transferred into the cytosol and then endoplasmic reticulum for conversion into triacylglycerol (TAG) in the microalgae. The free fatty acids are converted into acyl-CoA by long-chain acyl-CoA synthetase. Acyl-CoA and glycerol 3-phosphate are converted into lysophosphatidic acid by glycerol 3-phosphate acyltransferase (GPAT). Lysophosphatidic acid is converted to phosphatidic acid by lysophosphatidic acid acyltransferase (LPAT). Phosphatidic acid is converted into diacylglycerol by phosphatidic acid phosphatase (PAP). Diacylglycerol is converted into triacylglycerol (TAG) by diacylglycerol acyltransferase (DGAT). Triacylglycerol forms the TAG lipid body [156][157].
Hydrogen production in biological processes conducted by algae is based on the direct biophotolysis, which consists of the photosynthetic production of hydrogen from water, in which the energy of light is used to break the water molecule into hydrogen and oxygen [158]. It takes place mainly due to hydrogenase, which catalyzes the reversible oxidation of H2 and releases gaseous hydrogen by reducing protons [159][160]. Two transmembrane peptide complexes are responsible for hydrogen production in the photolysis process by microalgae: photosystem I (PSI) and photosystem II (PSII). The exposure of both complexes to solar radiation results in a water molecule breakdown. Then, O2 is produced by PSII, while PSI uses the electrons generated in this process to reduce CO2 and build cellular material (aerobic conditions), or the electrons are transferred by ferredoxin to hydrogenase and used for hydrogen production [161][162]. Another biochemical process led by algae to produce hydrogen is indirect biophotolysis. It has been proven to occur in the organisms of cyanobacteria, which accumulate carbohydrates resulting from CO2 reduction as a result of photosynthesis, which in turn are decomposed by fermentation mediated by photosystem I. The PSI proteins transfer electrons to ferredoxin using light energy [158][162]. In the indirect biophotolysis process, an important role is played by carbon dioxide, which is a carrier of electrons and protons formed during the water molecule degradation, and by enzymes, including two NiFe hydrogenases and nitrogenase, which catalyze atmospheric nitrogen reduction to ammonia with simultaneous proton reduction and hydrogen release [163][164].

This entry is adapted from the peer-reviewed paper 10.3390/su12239980

References

  1. Deviram, G.; Mathimani, T.; Anto, S.; Ahamed, T.S.; Ananth, D.A.; Pugazhendhi, A. Applications of microalgal and cyanobacterial biomass on a way to safe, cleaner and a sustainable environment. J. Clean. Prod. 2020, 253, 119770.
  2. Kamani, M.H.; Eş, I.; Lorenzo, J.M.; Remize, F.; Roselló-Soto, E.; Barba, F.J.; Clark, J.H.; Khaneghah, A.M. Advances in plant materials, food by-products, and algae conversion into biofuels: Use of environmentally friendly technologies. Green Chem. 2019, 21, 3213–3231.
  3. Patil, S.; Prakash, G.; Lali, A.M. Reduced chlorophyll antenna mutants of Chlorella saccharophila for higher photosynthetic efficiency and biomass productivity under high light intensities. J. Appl. Phycol. 2020, 32, 1559–1567.
  4. Santhakumaran, P.; Ayyappan, S.M.; Ray, J.G. Nutraceutical applications of twenty-five species of rapid-growing green-microalgae as indicated by their antibacterial, antioxidant and mineral content. Algal Res. 2020, 47, 101878.
  5. Tolboom, S.N.; Carrillo-Nieves, D.; de Jesús Rostro-Alanis, M.; de la Cruz Quiroz, R.; Barceló, D.; Iqbal, H.M.N.; Parra-Saldivar, R. Algal-based removal strategies for hazardous contaminants from the environment—A review. Sci. Total. Environ. 2019, 665, 358–366.
  6. Ziolkowska, J.R. Chapter 1—Biofuels technologies: An overview of feedstocks, processes, and technologies. In Biofuels for a More Sustainable Future; Elsevier: Amsterdam, The Netherlands; Oxford, UK; Cambridge, MA, USA, 2020; pp. 1–19.
  7. Stiles, W.A.V.; Styles, D.; Chapman, S.P.; Esteves, S.; Bywater, A.; Melville, L.; Silkina, A.; Lupatsch, I.; Fuentes, C.; Lovitt, R.; et al. Using microalgae in the circular economy to valorise anaerobic digestate: Challenges and opportunities. Bioresour. Technol. 2018, 267, 732–742.
  8. SundarRajan, P.; Gopinath, K.P.; Greetham, D.; Antonysamy, A.J. A review on cleaner production of biofuel feedstock from integrated CO2 sequestration and wastewater treatment system. J. Clean. Prod. 2019, 210, 445–458.
  9. Nawaz, T.; Rahman, A.; Pan, S.; Dixon, K.; Petri, B.; Selvaratnam, T. A review of landfill leachate treatment by microalgae: Current status and future directions. Processes 2020, 8, 384.
  10. Nagarajan, D.; Lee, D.J.; Chang, J.S. Biogas Upgrading by Microalgae: Strategies and Future Perspectives. In Microalgae Biotechnology for Development of Biofuel and Wastewater Treatment; Alam, M., Wang, Z., Eds.; Springer: Singapore, 2019.
  11. Rahman, A.; Agrawal, S.; Nawaz, T.; Pan, S.; Selvaratnam, T. A review of algae-based produced water treatment for biomass and biofuel production. Water 2020, 12. 2351.
  12. Sahu, S.K.; Mantri, V.A.; Zheng, P.; Yao, N. Chapter 1 Algae Biotechnology. Current Status, Potential and Impediments. In Encyclopedia of Marine Biotechnology; John Wiley & Sons Ltd: Hoboken, NJ, USA, 2020.
  13. Vassilev, S.V.; Vassileva, C.G. Composition, properties and challenges of algae biomass for biofuel application: An overview. Fuel 2016, 181, 1–33.
  14. Barsanti, L.; Gualtieri, P. Is exploitation of microalgae economically and energetically sustainable? Algal Res. 2018, 31, 107–115.
  15. Bhalamurugan, G.L.; Valerie, O.; Mark, L. Valuable bioproducts obtained from microalgal biomass and their commercial applications: A review. Environ. Eng. Res. 2018, 23, 229–241.
  16. Priyadarshani, I.; Rath, B. Commercial and industrial applications of micro algae—A review. J. Algal Biomass Util. 2012, 3, 89–100.
  17. Gomez Villa, H.; Voltolina, D.; Nieves, M.; Pina, P. Biomass production and nutrient budget in outdoor cultures of Scenedesmus obliquus (chlorophyceae) in artificial wastewater, under the winter and summer conditions of Mazatla´ n, Sinaloa, Mexico. Vie Et Milieu 2005, 55, 121–126.
  18. Mùnoz, R.; Guieysse, B. Algal-bacterial processes for the treatment of hazardous contaminants: A review. Water Res. 2006, 40, 2799–2815.
  19. Chojnacka, K.; Chojnacki, A.; Gorecka, H. Biosorption of Cr3+, Cd2+ and Cu2+ ions by blue–green algae Spirulina sp.: Kinetics, equilibrium and the mechanism of the process. Chemosphere 2005, 59, 75–84.
  20. Mùnoz, R.; Köllner, C.; Guieysse, B. Biofilm photobioreactors for the treatment of industrial wastewaters. J. Hazard. Mater. 2009, 161, 29–34.
  21. Yewalkar, S.N.; Dhumal, K.N.; Sainis, J.K. Chromium (VI)-reducing Chlorella spp. isolated from disposal sites of paper-pulp and electroplating industry. J. Appl. Phycol. 2007, 19, 459–465.
  22. Tarlan, E.; Dilek, F.B.; Yetis, U. Effectiveness of algae in the treatment of a wood-based pulp and paper industry wastewater. Bioresour. Technol. 2002, 84, 1–5.
  23. Acuner, E.; Dilek, F.B. Treatment of tectilon yellow 2G by Chlorella vulgaris. Process. Biochem. 2004, 39, 623–631.
  24. Essam, T.; Magdy, A.A.; El Tayeb, O.; Mattiasson, B.; Guieysse, B. Solar-based detoxification of phenol and p-nitrophenol by sequential TiO2 photocatalysis and photosynthetically aerated biological treatment. Water Res. 2007, 41, 1697–1704.
  25. Lima, S.A.C.; Raposo, M.F.J.; Castro, P.M.L.; Morais, R.M. Biodegradation of p-chlorophenol by a microalgae consortium. Water Res. 2004, 38, 97–102.
  26. Valderramaa, L.T.; Del Campoa, C.M.; Rodrigueza, C.M.; de-Bashan, L.E.; Bashan, Y. Treatment of recalcitrant wastewater from ethanol and citric acid production using the microalga Chlorella vulgaris and the macrophyte Lemna minuscule. Water Res. 2002, 36, 4185–4192.
  27. Tien, C.J. Biosorption of metal ions by freshwater algae with different surface characteristics. Process. Biochem. 2002, 38, 605–613.
  28. Jacob-Lopes, E.; Scoparo, C.H.G.; Queiroz, M.I.; Franco, T.T. Biotransformations of carbon dioxide in photobioreactors. Energy Convers. Manag. 2010, 51, 894–900.
  29. De Morais, M.G.; Costa, J.A.V. Isolation and selection of microalgae from coal fired thermoelectric power plant for biofixation of carbon dioxide. Energy Convers. Manag. 2007, 48, 2169–2173.
  30. Lam, M.K.; Lee, K.T.; Rahman, M.A. Current status and challenges on microalgae-based carbon capture. Int. J. Greenh. Gas Control. 2012, 10, 456–469.
  31. Thyagarajan, T.; Puri, M.; Vongsvivut, J.; Barrow, C.J. Evaluation of Bread Crumbs as a Potential Carbon Source for the Growth of Thraustochytrid Species for Oil and Omega-3 Production. Nutrients 2014, 6, 2104–2114.
  32. Ryu, B.; Kim, K.; Kim, J.; Han, J.; Yang, J. Use of organic waste from brewery industry for high-density cultivation of docosahexaenoic acid-rich microalga Aurantochytrium sp. KRS101. Bioresour. Technol. 2012, 129, 351–359.
  33. Unagul, P.; Assantachai, C.; Phadungruengluij, S.; Suphantharika, M.; Tanticharoen, M.; Verduyn, C. Coconut water as a medium additive for the production of docosahexaenoic acid (C22:6 n3) by Schizochytrium mangrovei Sk-02. Bioresour. Technol. 2007, 98, 281–287.
  34. Hong, W.; Yu, A.; Heo, S.; Oh, B.; Kim, C.; Sohn, J.; Yang, J.W.; Kondo, A.; Seo, J.W. Production of lipids containing high levels of docosahexaenoic acid from empty palm fruit bunches by Aurantiochytrium sp. KRS101. Bioprocess. Biosyst. Eng. 2013, 36, 959–963.
  35. Lin, L.; Chan, G.Y.S.; Jiang, B.L.; Lan, C.Y. Use of ammoniacal nitrogen tolerant microalgae in landfill leachate treatment. Waste Manag. 2007, 27, 1376–1382.
  36. Jedynak, P.; Burczyk, J.; Borowski, S.; Kaszycki, P.; Hałat-Łaś, M.; Kędra, M.; Mungunkhuyag, K.; Malec, P. Use of microalgae for treatment of post-fermentation effluent from biogas production. Przemysł Chem. 2018, 97, 2106–2109.
  37. Yan, C.; Zheng, Z. Performance of photoperiod and light intensity on biogas upgrade and biogas effluent nutrient reduction by the microalgae Chlorella sp. Bioresour. Technol. 2013, 139, 292–299.
  38. Kumar, M.; Sun, Y.; Rathour, R.; Pandey, A.; Thakur, I.S.; Tsang, D.C.W. Algae as potential feedstock for the production of biofuels and value-added products: Opportunities and challenges. Sci. Total. Environ. 2020, 716, 137116.
  39. Callegari, A.; Bolognesi, S.; Cecconet, D.; Capodaglio, A.G. Production technologies, current role, and future prospects of biofuels feedstocks: A state-of-the-art review. J. Crit. Rev. Environ. Sci. Technol. 2020, 50, 384–436.
  40. Walsh, M.J.; van Gerber Doren, L.; Sills, D.L.; Archibald, I.; Beal, C.M.; Lei, X.G.; Greene, C.H. Algal food and fuel coproduction can mitigate greenhouse gas emissions while improving land and water-use efficiency. Environ. Res. Lett. 2016, 11, 114006.
  41. Market Analysis Report. Algae Biofuel Market Size, Share & Trend Analysis By Application (Transportation, Others), By Region (North America, Europe, Asia Pacific, ROW), By Country, And Segment Forecasts, 2018–2025. Grand View Research. 2017. Available online: https://www.grandviewresearch.com/industry-analysis/algae-biofuel-market (accessed on 4 August 2020).
  42. Market Analysis Report. Biodiesel Market Analysis by Feedstock , By Application (Fuel, Power Generation), And Segment Forecasts, 2018–2025. Grand View Research. 2017. Available online: https://www.grandviewresearch.com/industry-analysis/biodiesel-market (accessed on 5 August 2020).
  43. Fu, W.; Nelson, D.R.; Mystikou, A.; Daakour, S.; Salehi-Ashtiani, K. Advances in microalgal research and engineering development. Curr. Opin. Biotechnol. 2019, 59, 157–164.
  44. Yadav, G.; Sen, R. Sustainability of Microalgal Biorefinery: Scope, Challenges, and Opportunities. In Sustainable Energy Technology and Policies; Green Energy and, Technology, De, S., Bandyopadhyay, S., Assadi, M., Mukherjee, D., Eds.; Springer: Singapore, 2018.
  45. Anto, S.; Mukherjee, S.S.; Muthappa, R.; Mathimani, T.; Deviram, G.; Kumar, S.S.; Verma, T.N.; Pugazhendhi, A. Algae as green energy reserve: Technological outlook on biofuel production. Chemosphere 2020, 242, 125079.
  46. Borowitzka, M.A.; Vonshak, A. Scaling up microalgal cultures to commercial scale. Eur. J. Phycol 2017, 52, 407–418.
  47. Webster, A.; Gardner, J. Aligning technology and institutional readiness: The adoption of innovation. Technology Anal. Strateg. Manag. 2019, 31, 1229–1241.
  48. Ozdemir, H.I.; Pinto, C.A.; Unal, R.; Keating, C.B.; Britcher, C. Supporting technology selection via portfolio readiness level and technology forecasting. In Proceedings of the International Annual Conference of the American Society for Engineering Management; Huntsville, Philadelphia, PA, USA, 24–26 October 2019.
  49. Bates, C.A.; Clausen, C. Engineering Readiness: How the TRL Figure of Merit Coordinates Technology Development. Eng. Stud. 2020, 12, 9–38.
  50. Okoro, V.; Azimov, U.; Munoz, J.; Hernandez, H.H.; Phan, A.N. Microalgae cultivation and harvesting: Growth performance and use of flocculants—A review. Renew. Sustain. Energy Rev. 2019, 115, 109364.
  51. Lardon, L.; He´lias, A.; Sialve, B.; Steyer, J.; Bernard, O. Life-cycle assessment of biodiesel production from microalgae. Environ. Sci. Technol. 2009, 43, 6475–6481.
  52. Clarens, A.F.; Nassau, H.; Resurreccion, E.P.; White, M.A.; Colosi, L.M. Environmental Impacts of Algae-Derived Biodiesel and Bioelectricity for Transportation. Environ. Sci. Technol. 2011, 45, 7554–7560.
  53. Frank, E.D.; Han, J.; Palou-Rivera, I.; Elgowainy, A.; Wang, M.Q. User Manual for algae life-cycle analysis with GREET: Version 0.0; ANL/ESD/11-7; Argonne National Laboratory: Lemont, IL, USA, 2011.
  54. Ganesan, R.; Manigandan, S.; Samuel, M.S.; Shanmuganathan, R.; Brindhadevi, K.; Chi, N.T.L.; Duc, P.A.; Pugazhendhi, A. A review on prospective production of biofuel from microalgae. Biotechnol. Rep. 2020, 27, e00509.
  55. Peng, L.; Fu, D.; Chu, H.; Wang, Z.; Qi, H. Biofuel production from microalgae: A review. Environ. Chem. Lett. 2020, 18, 285–297.
  56. Coimbra, R.N.; Escapa, C.; Otero, M. Comparative Thermogravimetric Assessment on the Combustion of Coal, Microalgae Biomass and Their Blend. Energies 2019, 12, 2962.
  57. Panahi, H.K.S.; Tabatabaei, M.; Aghbashlo, M.; Dehhaghi, M.; Rehan, M.; Nizami, A.-S. Recent updates on the production and upgrading of bio-crude oil from microalgae. Bioresour. Technol. Rep. 2019, 7, 100216.
  58. Feng, R.; Zaidi, A.A.; Zhang, K.; Shi, Y. Optimization of microwave pretreatment for biogas enhancement through anaerobic digestion of microalgal biomass. Period. Polytech. Chem. Eng. 2018, 63, 65–72.
  59. Córdova, O.; Chamy, R. Chapter 15—Microalgae to Biogas: Microbiological Communities Involved. In Microalgae Cultivation for Biofuels Production; Elsevier: London, UK; San Diego, CA, USA; Oxford, UK, 2020; pp. 227–249.
  60. Li, F.; Hülsey, M.J.; Yan, N.; Dai, Y.; Wang, C.-H. Co-transesterification of waste cooking oil, algal oil and dimethyl carbonate over sustainable nanoparticle catalysts. Chem. Eng. J. 2020, 405, 127036.
  61. Wang, Y.; Yang, H.; Zhang, X.; Han, F.; Tu, W.; Yang, W. Microalgal Hydrogen Production. Small Methods 2020, 4.
  62. Özçimen, D.; Koçer, A.T.; İnan, B.; Özer, T. Chapter 14—Bioethanol production from microalgae. In Handbook of Microalgae-Based Processes and Products; Elsevier: London, UK; San Diego, CA, USA; Cambridge, MA, USA; Oxford, UK, 2020; pp. 373–389.
  63. Chernova, N.I.; Kiseleva, S.V.; Larina, O.M.; Sytchev, G.A. Manufacturing gaseous products by pyrolysis of microalgal biomass. Int. J. Hydrog. Energy 2019, 45, 1569–1577.
  64. Lee, X.J.; Ong, H.C.; Gan, Y.Y.; Chen, W.-H.; Mahlia, T.M.I. State of art review on conventional and advanced pyrolysis of macroalgae and microalgae for biochar, bio-oil and bio-syngas production. Energy Convers. Manag. 2020, 210.
  65. Patil, R.A.; Kausley, S.B.; Joshi, S.M.; Pandit, A.B. Chapter 27—Process intensification applied to microalgae-based processes and products. In Handbook of Microalgae-Based Processes and Products; Elsevier: London, UK; San Diego, CA, USA; Cambridge, MA, USA; Oxford, UK, 2020; pp. 737–769.
  66. Nagappan, S.; Devendran, S.; Tsai, P.-C.; Dahms, H.-U.; Ponnusamy, V.K. Potential of two-stage cultivation in microalgae biofuel production. Fuel 2019, 252, 339–349.
  67. Fazal, T.; Mushtaq, A.; Rehman, F.; Khan, A.U.; Rashid, N.; Farooq, W.; Rehman, M.S.U.; Xu, J. Bioremediation of textile wastewater and successive biodiesel production using microalgae. Renew. Sustain. Energy Rev. 2018, 82, 3107–3126.
  68. Zhang, L.; Cheng, J.; Pei, H.; Pan, J.; Jiang, L.; Hou, Q. Cultivation of microalgae using anaerobically digested ef fl uent from kitchen waste as a nutrient source for biodiesel production. Renew. Energy 2018, 115, 276–287.
  69. Sati, H.; Mitra, M.; Mishra, S.; Baredar, P. Microalgal lipid extraction strategies for biodiesel production: A review. Algal Res. 2019, 38.
  70. Menegazzo, M.L.; Fonseca, G.G. Biomass recovery and lipid extraction processes for microalgae biofuels production: A review. Renew. Sustain. Energy Rev. 2019, 107, 87–107.
  71. Lee, S.Y.; Khoiroh, I.; Vo, D.N.; Kumar, S.; Loke, P. Show Techniques of lipid extraction from microalgae for biofuel production: A review. Environ. Chem. Lett. 2020, 1–21.
  72. Ranjith Kumar, R.; Hanumantha Rao, P.; Arumugam, M. Lipid extraction methods from microalgae: A comprehensive review. Front. Energy Res. 2015, 2, 61.
  73. Kumar, V.; Arora, N.; Nanda, M.; Pruthi, V. Different Cell Disruption and Lipid Extraction Methods from Microalgae for Biodiesel Production. In Microalgae Biotechnology for Development of Biofuel and Wastewater Treatment; Alam, M., Wang, Z., Eds.; Springer: Singapore, 2019.
  74. Choudhary, P.; Assemany, P.P.; Naaz, F.; Bhattacharya, A.; Castro, J.S.; Couto, E.A.; Calijuri, M.L.; Pant, K.K.; Malik, A. A review of biochemical and thermochemical energy conversion routes of wastewater grown algal biomass. Sci. Total. Environ. 2020, 726, 137961.
  75. Ren, J.; Liu, Y.-L.; Zhao, X.-Y.; Cao, J.-P. Biomass thermochemical conversion: A review on tar elimination from biomass catalytic gasification. J. Energy Inst. 2020, 93, 1083–1098.
  76. Clark, J.; Deswarte, F. Introduction to chemicals from biomass. In Wiley Series in Renewable Resources; Stevens, C.V., Ed.; John Wiley & Sons: Hoboken, NJ, USA, 2008.
  77. Radenahmad, N.; Azad, A.T.; Saghir, M.; Taweekun, J.; Bakar, M.S.A.; Reza, M.S.; Azad, A.K. A review on biomass derived syngas for SOFC based combined heat and power application. Renew. Sustain. Energy Rev. 2020, 119, 109560.
  78. Kousheshi, N.; Yari, M.; Paykani, A.; Saberi Mehr, A.; de la Fuente, G.F. Effect of Syngas Composition on the Combustion and Emissions Characteristics of a Syngas/Diesel RCCI Engine. Energies 2020, 13, 212.
  79. Hirano, A.; Hon-Nami, K.; Kunito, S.; Hada, M.; Ogushi, Y. Temperature effect on continuous gasification of microalgal biomass: Theoretical yield of methanol production and its energy balance. Catal. Today 1998, 45, 399–404.
  80. Minowa, T.; Sawayama, S. A novel microalgal system for energy production with nitrogen cycling. Fuel 1999, 78, 1213–1215.
  81. Patil, V.; Tran, K.-Q.; Giselrød, H.R. Towards sustainable production of biofuels from microalgae. Int. J. Mol. Sci. 2008, 9, 1188–1195.
  82. Ong, H.C.; Chen, W.-H.; Farooq, A.; Gan, Y.Y.; Lee, K.T.; Ashokkumar, V. Catalytic thermochemical conversion of biomass for biofuel production: A comprehensive review. Renew. Sustain. Energy Rev. 2019, 113, 109266.
  83. McKendry, P. Energy production from biomass (part 2): Conversion technologies. Bioresour. Technol. 2002, 83, 47–54.
  84. Dote, Y.; Sawayama, S.; Inoue, S.; Minowa, T.; Yokoyama, S.-Y. Recovery of liquid fuel from hydrocarbon-rich microalgae by thermochemical liquefaction. Fuel 1994, 73, 1855–1857.
  85. Minowa, T.; Yokoyama, S.-Y.; Kishimoto, M.; Okakura, T. Oil production from algal cells of Dunaliella tertiolecta by direct thermochemical liquefaction. Fuel 1995, 74, 1735–1738.
  86. Aravind, S.; Kumar, P.S.; Kumar, N.S.; Siddarth, N. Conversion of green algal biomass into bioenergy by pyrolysis. A review. Environ. Chem. Lett. 2020, 18, 829–849.
  87. Miao, X.L.; Wu, Q.Y. High yield bio-oil production from fast pyrolysis by metabolic controlling of Chlorella protothecoides. J. Biotechnol. 2004, 110, 85–93.
  88. Miao, X.; Wu, Q.; Yang, C. Fast pyrolysis of microalgae to produce renewable fuels. J. Anal. Appl. Pyrolysis 2004, 71, 855–863.
  89. Demirbas, A. Oily products from mosses and algae via pyrolysis. Energy Sources Part A—Recover. Utilization Environ. Effects 2006, 28, 933–940.
  90. Li, F.; Srivatsa, S.C.; Bhattacharya, S. A review on catalytic pyrolysis of microalgae to high-quality bio-oil with low oxygeneous and nitrogenous compounds. Renew. Sustain. Energy Rev. 2019, 108, 481–497.
  91. Hirano, A.; Ueda, R.; Hirayama, S.; Ogushi, Y. CO2 fixation and ethanol production with microalgal photosynthesis and intracellular anaerobic fermentation. Energy 1997, 22, 137–142.
  92. Ueno, Y.; Kurano, N.; Miyachi, S. Ethanol production by dark fermentation in the marine green alga, Chlorococcum littorale. J. Ferment. Bioeng. 1998, 86, 38–43.
  93. Lee, S.Y.; Sankaran, R.; Chew, K.W.; Tan, C.H.; Krishnamoorthy, R.; Chu, D.-T.; Show, P.-L. Waste to bioenergy: A review on the recent conversion technologies. BMC Energy 2019, 1, 4.
  94. Ghirardi, M.L.; Zhang, L.; Lee, J.W.; Flynn, T.; Seibert, M.; Greenbaum, E.; Melis, A. Microalgae: A green source of renewable H2. Trends Biotechnol. 2000, 18, 506–511.
  95. Saratale, G.D.; Saratale, R.G.; Banu, J.R.; Chang, J.-S. Biohydrogen Production From Renewable Biomass Resources. In Biohydrogen; Elsevier: Amsterdam, The Netherlands, 2019; pp. 247–277.
  96. Veras, T.S.; Mozer, T.S.; César, A.S. Hydrogen: Trends, production and characterization of the main process worldwide. Int. J. Hydrogen Energy 2017, 42, 2018–2033.
  97. Razu, M.H.; Hossain, F.; Khan, M. Advancement of Bio-hydrogen Production from Microalgae. In Microalgae Biotechnology for Development of Biofuel and Wastewater Treatment; Springer: Gateway East, Singapore, 2019; pp. 423–462.
  98. Grechanik, V.; Romanova, A.; Naydov, I.; Tsygankov, A. Photoautotrophic cultures of Chlamydomonas reinhardtii: Sulfur deficiency, anoxia, and hydrogen production. Photosynth. Res. 2020, 143, 275–286.
  99. Fakhimi, N.; Tavakoli, O. Improving hydrogen production using co-cultivation of bacteria with Chlamydomonas reinhardtii microalga. Mater. Sci. Energy Technol. 2019, 2, 1–7.
  100. Skjanes, K.; Knutsen, G.; Kӓllqvist, T.; Lindblad, P. H2 production from marine and freshwater species of green algae during sulfur deprivation and considerations for bioreactor design. Int. J. Hydrog. Energy 2008, 33, 511–521.
  101. Guan, Y.; Deng, M.; Yu, X.; Zhang, W. Two-stage photo-biological production of hydrogen by marine green alga Platymonas subcordiformis. Biochem. Eng. J. 2004, 19, 69–73.
  102. Guo, Z.; Chen, Z.; Lu, H.; Fu, Y.; Yu, X.; Zhang, W. Sustained hydrogen photoproduction by marine green algae platymonas subcordiformis integrated with in situ hydrogen consumption by an alkaline fuel cell system. J. Biotechnol. 2008, 136, 558–576.
  103. Chew, K.W.; Yap, J.Y.; Show, P.L.; Suan, N.H.; Juan, J.C.; Ling, T.C.; Lee, D.-J.; Chang, J.-S. Microalgae biorefinery: High value products perspectives. Bioresour. Technol. 2017, 229, 53–62.
  104. Sialve, B.; Bernet, N.; Bernard, O. Anaerobic digestion of microalgae as a necessary step to make microalgal biodiesel. Sustainable. Biotechnol. Adv. 2009, 27, 409–416.
  105. Olguin, E.J. The cleaner production strategy applied to animal production. In Environmental Biotechnology and Cleaner Bioprocesses; Olguín, E.J., Sánchez, G., Hemández, E., Eds.; Taylor and Francis: London, UK, 2000; pp. 107–121.
  106. Yen, H.-W.; Brune, D.E. Anaerobic co-digestion of algal sludge and waste paper to produce methane. Bioresour. Technol. 2007, 98, 130–134.
  107. Klocke, M.; Mahnert, P.; Mundt, K.; Souidi, K.; Linke, B. Microbial community analysis of a biogas-producing completely stirred tank reactor fed continuously with fodder beet silage as mono-substrate. Syst. Appl. Microbiol. 2007, 30, 139–151.
  108. Schlüter, A.; Bekel, T.; Diaz, N.N.; Dondrup, M.; Eichenlaub, R.; Gartemann, K.H.; Krahn, I.; Krause, L.; Kromeke, H.; Kruse, O.; et al. The metagenome of a biogas-producing microbial community of a production-scale biogas plant fermenter analysed by the 454-pyrosequencing technology. J. Biotechnol. 2008, 136, 77–90.
  109. Campbell, J.; Lobell, D.; Field, C. Greater transportation energy and GHG offsets from bioelectricity than ethanol. Science 2009, 324, 1055–1057.
  110. Börjesson, P.; Berglund, M. Environmental systems analysis of biogas systems -part I: Fuel-cycle emissions. Biomass Bioenergy 2006, 30, 469–485.
  111. Ubando, A.T.; Rivera, D.R.T.; Chen, W.H.; Culaba, A.B. A comprehensive review of life cycle assessment (LCA) of microalgal and lignocellulosic bioenergy products from thermochemical processes. Bioresour. Technol. 2019, 291, 121837.
  112. Ubando, A.T.; Felix, C.B.; Chen, W.-H. Biorefineries in circular bioeconomy: A comprehensive review. Bioresour. Technol. 2020, 299, 122585.
  113. Chandra, R.; Iqbal, H.M.N.; Vishal, G.; Lee, H.S.; Nagra, S. Algal biorefinery: A sustainable approach to valorize algal-based biomass towards multiple product recovery. Bioresour. Technol. 2019, 278, 346–359.
  114. Kisielewska, M.; Zieliński, M.; Dębowski, M.; Kazimierowicz, J.; Romanowska-Duda, Z.; Dudek, M. Effectiveness of Scenedesmus sp. Biomass Grow and Nutrients Removal from Liquid Phase of Digestates. Energies 2020, 13, 1432.
  115. Mohan, S.V.; Hemalatha, M.; Chakraborty, D.; Chatterjee, S.; Ranadheer, P.; Kona, R. Algal biorefinery models with self-sustainable closed loop approach: Trends and prospective for blue-bioeconomy. Bioresour. Technol. 2019, 295, 122128.
  116. Mishra, S.; Roy, M.; Mohanty, K. Microalgal bioenergy production under zero-waste biorefinery approach: Recent advances and future perspectives. Bioresour. Technol. 2019, 292, 122008.
  117. Metsoviti, M.N.; Papapolymerou, G.; Karapanagiotidis, I.T.; Katsoulas, N. Comparison of Growth Rate and Nutrient Content of Five Microalgae Species Cultivated in Greenhouses. Plants 2019, 8, 279.
  118. Aziz, M.M.A.; Kassim, A.K.; Shokravi, Z.; Jakarni, F.M.; Liu, H.Y.; Zaini, N. Two-stage cultivation strategy for simultaneous increases in growth rate and lipid content of microalgae: A review. Renew. Sustain. Energy Rev. 2020, 119, 109621.
  119. Sánchez-Bayo, A.; Morales, V.; Rodríguez, R.; Vicente, G.; Bautista, L.F. Cultivation of microalgae and cyanobacteria: Effect of operating conditions on growth and biomass composition. Molecules 2020, 25, 2834.
  120. Neofotis, P.; Huang, A.; Sury, K.; Chang, W.; Joseph, F.; Gabr, A.; Twary, S.; Qiu, W.; Holguine, O.; Polle, J.E.W. Characterization and classification of highly productive microalgae strains discovered for biofuel and bioproduct generation. Algal Res. 2016, 15, 164–178.
  121. Li, P.; Sakuragi, K.; Makino, H. Extraction Techniques in Sustainable Biofuel Production: A Concise Review. Fuel Process. Technol. 2019, 193, 295–303.
  122. Piasecka, A.; Nawrocka, A.; Wiącek, D.; Krzemińska, J. Agro-industrial by-product in photoheterotrophic and mixotrophic culture of Tetradesmus obliquus: Production of ω3 and ω6 essential fatty acids with biotechnological importance. Sci Rep. 2020, 10, 1–11.
  123. Huang, G.H.; Chen, F.; Wei, D.; Zhang, X.W.; Chen, G. Biodiesel production by microalgal biotechnology. Appl. Energy 2010, 87, 38–46.
  124. 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. Bioresour. Technol. 2010, 101, 71–74.
  125. Rodolfi, L.; Zittelli, G.C.; Bassi, N.; Padovani, G.; Biondi, N.; Bonini, G.; Tredici, M.R. Microalgae for oil: Strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnol. Bioeng. 2009, 102, 100–112.
  126. Chiu, S.-Y.; Kaom, C.-J.; Chen, C.-H.; Kuan, T.-C.; Ong, S.-C.; Lin, C.-S. Reduction of CO2 by a high-density culture of Chlorella sp. in a semicontinuous photobioreactor. Bioresour. Technol. 2008, 99, 3389–3396.
  127. Chojnacka, K.; Marquez-Rocha, F.J. Kinetic and stoichiometric relationships of the energy and carbon metabolism in the culture of microalgae. Biotechnology 2004, 3, 21–34.
  128. Morales-Sánchez, D.; Martinez-Rodriguez, O.A.; Martinez, A. Heterotrophic cultivation of microalgae: Production of metabolites of commercial interest. J. Chem. Technol. Biotechnol. 2016, 92, 925–936.
  129. De Swaaf, M.; Sijtsma, L.; Pronk, J. High-cell-density fed-batch cultivation of the docosahexaenoic acid producing microalga Crypthecodinium cohnii. Biotechnol. Bioeng. 2003, 81, 666–672.
  130. Xu, H.; Miao, X.L.; Wu, Q.Y. High quality biodiesel production from a microalga Chlorella protothecoides by heterotrophic growth in fermenters. J. Biotechnol. 2006, 126, 499–507.
  131. Kuei-Ling, Y.; Jo-Shu, C. Effects of cultivation conditions and media composition on cell growth and lipid productivity of indigenous microalga Chlorella vulgaris ESP-31. Bioresour. Technol. 2012, 105, 120–127.
  132. Caporgno, M.P.; Haberkorn, I.; Böcker, L.; Mathys, A. Cultivation of Chlorella protothecoides under different growth modes and its utilisation in oil/water emulsions. Bioresour. Technol. 2019, 288, 121476.
  133. Sim, S.J.; Joun, J.; Hong, M.E.; Patel, A.K. Split mixotrophy: A novel cultivation strategy to enhance the mixotrophic biomass and lipid yields of Chlorella protothecoides. Bioresour. Technol. 2019, 291, 121820.
  134. Shen, X.; Qin, Q.; Yan, S.; Huang, J.L.; Liu, K.; Zhou, S.-B. Biodiesel production from Chlorella vulgaris under nitrogen starvation in autotrophic, heterotrophic, and mixotrophic cultures. J. Appl. Phycol. 2019, 31, 1589–1596.
  135. Li, T.; Kirchhoff, H.; Gargouri, M.; Feng, J.; Cousins, A.B.; Pienkos, P.T.; Gang, D.R.; Chen, S. Assessment of photosynthesis regulation in mixotrophically cultured microalga Chlorella sorokiniana. Algal Res. 2016, 19, 30–38.
  136. Zheng, Y.; Chi, Z.; Lucker, B.; Chen, S. Two-stage heterotrophic and phototrophic culture strategy for algal biomass and lipid production. Bioresour. Technol. 2012, 103, 484–488.
  137. Li, T.; Zheng, Y.; Yu, L.; Chen, S. Mixotrophic cultivation of a Chlorella sorokiniana strain for enhanced biomass and lipid production. Biomass Bioenergy 2014, 66, 204–213.
  138. Liang, Y.; Sarkany, N.; Cui, Y. Biomass and lipid productivities of Chlorella vulgaris under autotrophic, heterotrophic and mixotrophic growth conditions. Biotechnol. Lett. 2009, 31, 1043–1049.
  139. Pang, N.; Gu, X.; Chen, S.; Kirchhoff, H.; Lei, H.; Rojec, S. Exploiting mixotrophy for improving productivities of biomass and co-products of microalgae. Renew. Sustain. Energy Rev. 2019, 112, 450–460.
  140. Chen, Y.H.; Walker, T.H. Biomass and lipid production of heterotrophic microalgae Chlorella protothecoides by using biodiesel-derived crude glycerol. Biotechnol. Lett. 2011, 33, 1973.
  141. Zuccaro, G.; Yousuf, A.; Pollio, A.; Steyer, J.-P. Microalgae Cultivation Systems. In Microalgae Cultivation for Biofuels Production; Elsevier: London, UK; San Diego, CA, USA; Cambridge, MA, USA; Oxford, UK, 2020; pp. 11–29.
  142. Cezare-Gomes, E.A.; del Mejia-da-Silva, L.C.; Pérez-Mora, L.S.; Matsudo, M.C.; Ferreira-Camargo, L.S.; Singh, A.K.; de Carvalho, J.C.M. Potential of Microalgae Carotenoids for Industrial Application. Appl. Biochem. Biotechnol. 2019, 188, 602–634.
  143. Zhang, Y.; Su, H.; Zhong, Y.; Zhang, C.; Shen, Z.; Sang, W.; Yan, G.; Zhou, X. The effect of bacterial contamination on the heterotrophic cultivation of Chlorella pyrenoidosa in wastewater from the production of soybean products. Water Res. 2012, 46, 5509–5516.
  144. Marudhupandia, T.; Sathishkumara, R.; Kumara, T.T.A. Heterotrophic cultivation of Nannochloropsis salina for enhancing biomass and lipid production. Biotechnol. Rep. 2016, 10, 8–16.
  145. Chandra, R.; Arora, S.; Rohit, M.V.; Mohan, S.V. Lipid metabolism in response to individual short chain fatty acids during mixotrophic mode of microalgal cultivation: Influence on biodiesel saturation and protein profile. Bioresour. Technol. 2015, 188, 169–176.
  146. Mata, T.M.; Martins, A.A.; Caetano, N.S. Microalgae for biodiesel production and other applications: A review. Renew. Sustain. Energy Rev. 2010, 14, 217–232.
  147. Bhatnagar, A.; Chinnasamy, S.; Singh, M.; Das, K.C. Renewable biomass production by mixotrophic algae in the presence of various carbon sources and wastewaters. Appl. Energy 2011, 88, 3425–3431.
  148. Yu, H.F.; Jia, S.R.; Dai, Y.J. Growth characteristics of the cyanobacterium Nostoc flagelliformein photoautotrophic, mixotrophic and heterotrophic cultivation. J. Appl. Phycol. 2009, 21, 127–133.
  149. Bailey, R.; DiMasi, D.; Hansen, J.; Mirrasoul, P.; Ruecker, C.; Kaneko, T.; Barclay, W. Enhanced Production of Lipids Containing Polyenoic Fatty Acid by Very High Density Cultures of Eukaryotic Microbes in. Fermentors Patent No.: US 6,607,900 B2, 19 August 2003.
  150. Anand, P.; Saxena, R.K. A comparative study of solvent-assisted pretreatment of biodiesel derived crude glycerol on growth and 1,3-propanediol production from Citrobacter freundii. New Biotechnol. 2011, 29, 199–205.
  151. Jiang, Y.; Yoshida, T.; Quigg, A. Photosynthetic performance, lipid production and biomass composition in response to nitrogen limitation in marine microalgae. Plant Physiol. Biochem. 2012, 54, 70–77.
  152. Ogbonna, J.C.; Ichige, E.; Tanaka, H. Regulating the ratio of photoautotrophic to heterotrophic metabolic activities in photoheterotrophic culture of Euglena gracilis and its application to alpha-tocopherol production. Biotechnol. Lett. 2002, 24, 953–958.
  153. Reverdatto, S.; Beilinson, V.; Nielsen, N.C. A multisubunit acetyl coenzyme A carboxylase fromsoybean. Plant Physiol. 1999, 119, 961–978.
  154. Chen, J.W.; Liu, W.J.; Hu, D.X.; Wang, X.; Balamurugan, S.; Alimujiang, A.; Yang, W.D.; Liu, J.S.; Li, H.Y. Identification of a malonyl CoA-acyl carrier protein transacylase and its regulatory role infatty acid biosynthesis in oleaginous microalga Nannochloropsis oceanic. Biotechnol. Appl. Biochem. 2016, 64, 620–626.
  155. Fan, J.; Andre, C.; Xu, C. A chloroplast pathway for the de novo biosynthesis of triacylglycerol inChlamydomonas reinhardtii. FEBS Lett. 2011, 585, 1985–1991.
  156. Wan, L.; Han, J.; Sang, M.; Li, A.; Wu, H.; Yin, S.; Zhang, C. De novo transcriptomic analysis of anoleaginous microalga: Pathway description and gene discovery for production of next-generationbiofuels. PLoS ONE 2012, 7, e35142.
  157. Lenka, S.K.; Carbonaro, N.; Park, R.; Miller, S.M.; Thorpe, I.; Li, Y. Current advances in molecular, biochemical, and computational modeling analysis of microalgal triacylglycerol biosynthesis. Biotechnol. Adv. 2016, 34, 1046–1063.
  158. Show, K.-Y.; Yan, Y.-G.; Lee, D.-J. Biohydrogen production from algae: Perspectives, challenges, and prospects. In Biofuels from Algae, 2nd ed.; Elsevier: Amsterdam, The Netherlands; Oxford, UK; Cambridge, MA, USA, 2019; pp. 325–343.
  159. Dasgupta, C.N.; Gilbert, J.J.; Lindblad, P.; Heidorn, T.; Borgvang, S.A.; Skjanes, K.; Das, D. Recent trends on the development of photobiological processes and photobioreactors for the improvement of hydrogen production. Int. J. Hydrog. Energy 2010, 35, 10218–10238.
  160. Miyake, J.; Miyake, M.; Asada, Y. Biotechnological hydrogen production: Research for efficient light energy conversion. J. Biotechnol. 1999, 70, 89–101.
  161. Ni, F.M.; Leung, D.Y.C.; Leung, M.K.H.; Sumathy, K. An overview of hydrogen production from biomass. Fuel Process. Technol. 2006, 87, 461–472.
  162. Akhlaghi, N.; Najafpour-Darzi, G. A comprehensive review on biological hydrogen production. Int. J. Hydrog. Energy 2020, 45, 22492–22512.
  163. Das, D.; Veziroglu, T.N. Hydrogen production by biological processes: A survey of literature. Int. J. Hydrog. Energy 2001, 26, 13–28.
  164. Kim, D.H.; Kim, M.S. Hydrogenases for biological hydrogen production. Bioresour. Technol. 2011, 102, 8423–8431.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.
This entry is offline, you can click here to edit this entry!
Video Production Service
Feedback