New strategies enhancing feasibility of microalgal cultivations

C H A P T E R

16 New strategies enhancing feasibility of microalgal cultivations Fabrizio Di Caprio, Pietro Altimari, Francesca Pagnanelli* Department of Chemistry, University Sapienza of Rome, Rome, Italy * Corresponding author. e-mail address: [email protected]

1. Introduction Microalgae are unicellular microorganisms that live in nature by exploiting photosynthetic metabolism. Microalgae is the name by which eukaryotic photosynthetic microorganisms are generally indicated, while prokaryotic photosynthetic microorganisms are named cyanobacteria (green-blue algae). For simplicity, in this chapter the term “microalgae” will refer to both. It is estimated that up to 200,000 species of microalgae may exist [1]. They live in nature in a large variety of different environments such as seas, lakes, rivers, glaciers, wastewaters, etc. Exploiting their metabolism, microalgae can 2
use inorganic carbon (CO2, HCOe 3 , and CO3 ), light, water, and simple mineral salts (PO3
4 , 2þ 2þ NO
, Ca , Mg , etc.) to synthetize complex 3 organic molecules such as proteins, fatty acids, polysaccharides, carotenoids, and others [2] (Fig. 16.1). These molecules can be used for a large variety of applications such as biofuels, bioplastics, feed, and food [3]. CO2 and light are renewable resources widely available; consequently, the possibility to

Catalysis, Green Chemistry and Sustainable Energy https://doi.org/10.1016/B978-0-444-64337-7.00016-1

develop an industry able to use these resources as feedstocks to obtain several organic molecules has strongly attracted the interest of scientific researchers and industrialists [4]. Although these general features are owned also by terrestrial plants, microalgae have some properties that make them more promising as future biomass sources with respect to terrestrial plants: -

-

-

Microalgae have higher photosynthetic efficiency (maximum value w 10%) with respect to terrestrial plants (maximum value w 5%) [5e8]. Microalgae can reach higher biomass productivity (50e70 t/ha per year) with respect to terrestrial plants (10e20 t/ha per year) [1,9,10]. Microalgae can grow on nonarable lands and in the sea [11]. Microalgae can grow in closed systems and by using wastewaters and saltwater, thus reducing considerably freshwater consumption [9].

However, despite these potentialities, the biomass produced by terrestrial plant is still the

287

Copyright © 2019 Elsevier B.V. All rights reserved.

288

16. New strategies enhancing feasibility of microalgal cultivations

FIGURE 16.1

Schematic illustration of microalgae photosynthetic metabolism.

main biomass available as feedstock for the industry. The worldwide microalgae biomass production was estimated 5000 t/y for Spirulina in 2013 and 2000 t/y for Chlorella in 2003 [12]. This amount is negligible with respect to biomass coming from agriculture. It corresponds to about 400e500 ha of maize cultivated land, which is comparable with the average production of just two average-sized farms in the United States [13]. This limited production is mainly given by the high cost associated with microalgae biomass production. At the state of the art, with the best available technologies, the minimum cost achievable for microalgae biomass production is between 3 and 10 V per kg of dry microalgae biomass, according to estimates published in 2016 [11,14,15]. Costs lower than 3 V/kg have been also reported in different works; however, these latter derive from assumptions and forecasts that still need to be experimentally verified [14e16]. The elevated cost is given by the application of expensive cultivation systems (mainly open ponds, flat panel and tubular photobioreactors) for reactors installation and for continuous power supply (required for cultivation and harvesting) that

are not balanced by the biomass productivity. The most widespread agricultural crops are less productive with respect to microalgae, but the cultivation systems used are enough cheap to ensure lower production costs. For example, the price associated with some of the main vegetable commodities is about 0.5 V/kg for palm oil, 0.2 V/kg for corn, and 0.3 V/kg for soybean (these three can be considered the main vegetable commodities for lipids, carbohydrates, and proteins, respectively) (Index Mundi, 2018). Due to the high cost associated, currently microalgae biomass production is mainly carried out for high added value products (Table 16.1) such as microalgal powder, tablets, supplements, and ingredients for human and animal nutrition. For example, docosahexaenoic acid (DHA) is an omega-3 fatty acid produced mainly by heterotrophic cultivation of Schizochytrium sp. in different plants located in the United States and France. Haematococcus pluvialis is cultivated by Cyanotech (in Hawaii) and Algatech (Israel) to produce an oleoresin enriched in astaxanthin, which is obtained by CO2 supercritical extraction from dried biomass. Chlorella and Spirulina species

IV. Selected examples and case history

289

2. Development of biorefineries

TABLE 16.1

Main companies involved in industrial production of microalgae biomass and derivates.

Company name Species cultivated

Plant size

Reactor type

References

108 ha Powder, tablets and phycocyanin extracts

Open ponds

http://earthrise. com/

Taiwan Chlorella Chlorella sorokiniana Manufacturing Company

2.6 ha Powder, tablets, and chlorella extracts

Open ponds

http://www. taiwanchlorella.com

Roquette Kl€ otze Chlorella vulgaris GmbH & co. KG

500 km Powder glass tubes

Tubular photobioreactors

www.algomed.de/ en

Tubular photobioreactors

https://www. algatech.com

Open ponds

https://www. cyanotech.com/

Earthrise

Spirulina (Arthrospira platensis)

Products

Algatechnologies Haematococcus pluvialis 600 km Astaxanthin and Ltd. and Phaeodactylum glass fucoxanthin tricornutum tubes Cyanotech

Haematococcus pluvialis 36 ha and Spirulina (Arthrospira platensis)

Spirulina tablets and encapsulated oleoresin enriched in astaxanthin

Phycom

Chlorella sorokiniana and e Chlorella vulgaris

Pellets, powder, and flakes Closed fermenters

http://www. phycom.eu

Solazyme

Different Chlorella sp. strains

e

Oils and several food ingredients

Closed fermenters

http:// solazymeindustrials. com/

Alltech

Schizochytrium sp.

e

DHA oil

Closed fermenters

https://www. alltech.com/

Fermentalg

Schizochytrium sp.

e

DHA oil or pigments

Mixotrophy using mainly heterotrophy (fermenters þ flash)

https://www. fermentalg.com/fr/

are often placed on the market as a powder rich in proteins. Such powders are dried and sold as pressed pills, flakes, or tablets for nutraceutical applications [12]. The powders have also applications as animal protein replacement for vegan food production (e.g., Terranostra Food s.r.l., Italy). In the following sections the new promising strategies to enhance the feasibility of microalgae biomass production at larger industrial scale are described. The described strategies are development of biorefineries (Section 2), innovative photobioreactors (Section 3), wastewater utilization as a source of nutrients (Section 4), and strain improvement (Section 5).

2. Development of biorefineries The aim of this section is to describe the development of a biorefinery, which is one of the most promising approaches to enhance the feasibility of microalgae processes. The concept of a biorefinery is defined in Subsection 2.1, then the different products obtainable from microalgae biomass are described in Subsection 2.2. Subsequently, in Subsection 2.3, the latest innovations in the development of a biorefinery from microalgae are described, illustrating the shifting of the processes from single end products to multiple end products.

IV. Selected examples and case history

290

16. New strategies enhancing feasibility of microalgal cultivations

FIGURE 16.2

Comparison between a conventional process and an example of biorefinery.

2.1 Definition of biorefinery With the term “biorefinery” a facility is indicated, in which microalgae biomass is used as feedstock to produce a large variety of different products. The term derives from the conventional petroleum refineries in which crude oil is used as feedstock to produce a large variety of different petrochemical derivatives (fuels, oils, heat, solvents, plastics, etc.). It is expected that the development of biorefineries may enhance economic and environmental sustainability of microalgae biomass production, through the valorization of different components of the produced biomass. Until now, industrial processes that produce microalgae have been designed to obtain only a single product from a single biomass. The biorefinery aims to change this concept by shifting the process design from one biomass / one product to one biomass / multiple products (Fig. 16.2).

2.2 Potential products obtainable by a microalgae biorefinery Hypothetically, every component of the microalgae biomass may be exploited for a

different product. The main carbohydrates (starch and cellulose) could be used to produce bioplastics, biofuels, or food [17e19], while some other carbohydrates (e.g., b-glucans) could have applications as nutraceuticals or cosmetics [20,21]. Microalgae proteins generally have a good composition in amino acids, so they are considered a promising future source of proteins for human nutrition (especially in replacing animal proteins) and for feed (for livestock and aquaculture) [22,23]. Microalgae proteins can also be exploited to produce bioplastics, for example, by processing directly dried biomass rich in proteins [24], or to synthetize polyurethanes starting from amino acids [25]. Some peptides, with health benefit properties, can be produced by means of specific digestion [26]. Proteins include enzymes, which could be extracted and used as catalyzers for a large variety of different reactions with applications in industry, for example, laccase and esterase [27e29]. Proteins from microalgae showed superior surface activity and gelation behavior when compared with whey proteins, with applications in food industry [30,31]. Lipids from microalgae can be divided in neutral lipids, mainly triacylglycerols (TAGs),

IV. Selected examples and case history

2. Development of biorefineries

and polar lipids, mainly phospholipids [32]. Some microalgae, for example, Botryococcus braunii, can accumulate hydrocarbons too [33]. Fatty acids from microalgae can be used to produce biofuels (biodiesel above all) [34], bioplastics [35], and other products, similarly to what is already done in the industry starting from conventional vegetable oils such as palm oil, soy oil, and others. Glycerol deriving from hydrolysis of TAGs and phospholipids can be recycled as nutrients to improve microalgae productivity [36]. Differently to the widespread vegetable oils, microalgae oils are rich in omega-3 fatty acids (for example DHA), which make them a promising substitute of fish oil for aquaculture and to face the issue of omega-3/omega-6 imbalance in the human diet [37,38]. Pigments such as carotenoids (astaxanthin, b-Carotene), xanthophylls (lutein, zeaxanthin), chlorophylls, and phycocyanin also received attention for their high added value (between 300 and 3000 V/kg) [3]. Some of them are already extracted for nutraceutical application (Table 16.1). They can also be used as pigments for food and feed; for example, astaxanthin is responsible for the red color of salmon [39,40] and lutein of the yellow-orange color of egg yolk [41].

2.3 Shifting microalgae processes from single products to multiple products The shifting from conventional microalgae processes to biorefinery facilities needs several changes and improvements. The main changes required are these: -

-

the shifting of conventional cultivation processes in which a single biochemical target component is maximized to the optimization of processes tailored to achieve a biomass composition balanced for the different biochemical target components; the shifting of the downstream processes of the biomass from separation/purification/

291

transformation of a single target product to multiple target products. The two changes are strongly related because they depend on each other. Technical, economic, and environmental sustainability are factors that determine the number of target products obtainable from a microalgae biorefinery. Currently, downstream treatments of microalgae biomass are carried out by adapting technologies previously developed for other applications. For this reason, the sustainability of the downstream treatment is strongly limited, and a large part of the companies sell biomass that is just dried and pressed (Table 16.1). To extract different compounds from the same starting biomass, the extraction treatments used for every compound should be mild enough to avoid the degradation of the other compounds. Much research has been conducted to optimize the extraction of different compounds from microalgae biomass; however, a large part of these works was carried out by focusing on a single product. For example, a study was carried out for the optimization of lipid extraction and another one for the optimization of protein extraction, instead of carrying out a single study to optimize both. The information produced from these works is essential for the development of a biorefinery, but it is not enough. In fact, the preliminary information achieved by the studies focusing on single products should be successively exploited to carry out further studies in which extraction of different compounds is optimized in a single process. In this direction, difference strategies have been proposed (Fig. 16.3). All the downstream processes have to face initially with the separation of the biomass from water. Water is the solvent in which microalgae carry out their reactions, namely the solvent in which they grow. In photoautotrophic cultivations, water is generally between 99.9% and 99.5% of the microalgae culture [42], while in heterotrophic cultivations, it is generally

IV. Selected examples and case history

292

FIGURE 16.3

16. New strategies enhancing feasibility of microalgal cultivations

Illustration of the main strategies described for separation of different biomolecules from microalgae

biomass.

between 99.1% and 90% [43]. A large part of water can be removed by the preliminary harvesting phase, generally achievable by filtrations, sedimentation, flocculation, or centrifugation [44]. The parameters of the harvesting are mainly determined by the biomass cultivation condition (microalgae strain, final biomass concentration achieved, etc.). Through harvesting, biomass reaches a water content between 99% and 70%, depending on the technology used, with an energy consumption between 0.01 and 8 kWh/kg [15,44]. After harvesting, a large part of the proposed processes includes a drying phase, by which water content is reduced below 5%. Freeze-drying and spray drying are the most used technical methods to dry microalgae biomass before compound extraction. However, the drying process is also a very energy-intensive treatment. Energy consumed for freeze-drying has been estimated by some authors in the range of 5e40 kWh/kg, corresponding to about 1e6 V/kg [45], while some other authors estimated

a lower cost for biomass drying corresponding to about 0.18 V/kg [15]. Due to the impact of the drying phase, there is a recent growing interest toward the development of wet processes in which extractions are carried out directly on wet biomass [46]. The general strategy followed in the biorefinery is the cascade extraction approach, by which different compounds are extracted by using sequential extractions and separation phases. The intracellular molecules have to be extracted and separated by means of a partial or total alteration of cell structure. These alterations can be more or less strong, in function of the used conditions. In general, it can be considered that the kinetic of the extraction increases by increasing the cell destruction efficiency. In fact, when the cells are still intact, or just partially broken, the kinetic is strongly limited by the diffusion gradient inside cells (or cell aggregates) and by cellular layers (cell wall, cell membranes) that obstruct molecule movement.

IV. Selected examples and case history

2. Development of biorefineries

The gentlest alterations are those obtained by some selective extractions that can be carried out directly on live cells. In fact, it is possible to use conditions mild enough to maintain the cells alive. This kind of extraction is named milking because of its similarity to the milking of cows. Milking has been used to extract b-carotene from (Dunaliella) salina [47], hydrocarbons from B. braunii [48], and fatty acids from Nannochloropsis sp. [49]. An essential condition for milking is the adoption of biocompatible solvents. Zhang and coworkers found that a partition coefficient >5.5 (log Poct/wat) was required to be biocompatible (nontoxic) for Nannochloropsis sp. cells [49]. However, it should be considered that milking has kinetics and efficiency generally lower than conventional extractions, and it is applicable only for some molecules. In the large parts of the processes investigated, the approach followed was based on sequential selective extractions coupled with treatment for partial or total cell destruction (death of cells). In every extraction condition, some compounds are pulled out with a certain selectivity from the residual biomass, by producing a solution enriched in such compounds. To increase the extraction efficiency, different technical methods have been studied and optimized to achieve cell destruction. They can be classified as mechanical methods such as highpressure homogenization, bead milling, ultrasonication, microwave, and pulsed electric fields and nonmechanical methods such as chemical and enzymatic treatments [45,50]. Chemical and enzymatic treatments are also accompanied by reactions that can induce modification in the chemical properties of the extracted molecules (e.g., hydrolysis). Some extraction solvents, a large part of organic solvents, are able themselves to break down cell membranes by solubilizing membrane phospholipids [51]. Data reported in the literature indicate that the pretreatments carried out for cell disruption require from 0.06 kWh/kg to 150 kWh/kg [50].

293

The large variability reported depends on the technology used, microalgae species, biomass concentration, suspension flow, pH, temperature, and also by the analytical method used to assess cell destruction. Comparing the same operative conditions, beat beater has been used to achieve cell destruction of Chlorella vulgaris, Neochloris oleoabundans, and Tetraselmis suecica with a comparable energy consumption of 0.5 kWh/kg for all three strains [52]. Cell destruction efficiency is generally quantified by measuring cell disappearing by optical counting, flowcytometry, or cell counter [52,53]. However, that means that also with a reported 100% cell destruction, there will be still a certain amount of cell fragments (of variable size) in which different molecules stay linked together, limiting their extraction and separation. Consequently, it is not easy to understand from literature data what is exactly the amount of cell destruction in the extracts, and data obtained can be strongly influenced by the solid/liquid separation method used after treatment. Once the molecules are moved in solution, they can be separated by treatments exploiting conventional chemical and biochemical methods (precipitation, solvent partition, chromatography, etc.). Different studies have investigated the possibility to separate fatty acids and lutein starting from the extract obtained by lipid extraction with organic solvents. These works were based on the different partition coefficient of lutein and TAGs in an ethanol/hexane twophase system [54,55]. Biomass was first treated with a cell destruction treatment coupled with solvent extraction of lipids (e.g., by using ethanol or directly hexane/ethanol mixture). This phase could be coupled with alkaline treatment because high OH
concentrations hydrolyze lutein esters to free lutein, and at the same time, OH
can enhance cell breaking (however OH
should be carefully dosed for exposition time and concentration to avoid lutein degradation) [41]. After lipid extraction, a two-phase ethanol-hexane system can be obtained, in which

IV. Selected examples and case history

294

16. New strategies enhancing feasibility of microalgal cultivations

the lutein goes mainly in the ethanol phase, while the TAGs go mainly in the hexane phase. Pure lutein can be finally obtained from the ethanol phase by means of precipitation induced by water addition [56,57]. TAGs in the hexane extract can be used to synthetize biodiesel by converting fatty acids in their methyl esters by transesterification. The residual defatted biomass rich in proteins and carbohydrates can be further valorized. However, protein and carbohydrate separation from defatted biomass is poorly investigated in the literature. The main alternatives investigated in the literature for the valorization of defatted biomass concern the production of other biofuels: by hydrolysis (chemical or enzymatic) to produce substrates for microbial fermentations for bioethanol or biogas production [58,59] and by thermal and hydrothermal treatments for biochar, gas, and liquid fuels production [60,61]. Other investigated applications include biosorption, by which the defatted biomass was used as biosorbent for heavy metal removal in contaminated water [62]. Extractions carried out by using bead milling in aqueous suspension allowed selective protein extraction, with selectivity between 0.9 and 6.7 for proteins with respect to carbohydrates [52]. But it should be taken into account that this study was conducted on nondefatted biomass. Instead, in the processes investigated by Ref. [63] and by Ref. [64], the microalgae biomass was first treated with organic solvents to extract lipids, and then proteins were extracted from defatted biomass by using alkaline solutions. However, although alkaline proteins extraction was successfully used, for example, to remove proteins from biomasses as rice flour [65,66], when applied to microalgae biomass, it was not very effective (only w 10% protein extraction) [63,64]. Desai and coworkers proposed a process by which astaxanthin and proteins were simultaneously extracted and separated

from Haematococcus pluvialis cells by exploiting a microgel-stabilized ionic liquidewater emulsion. The astaxanthin was extracted in its ionic liquid droplets, with a yield of 62%, while proteins (90%) moved in the aqueous phase [67]. The protein extracts obtained by any extraction process can be processed by conventional downstream treatments such as membrane filtration and precipitation to produce protein concentrates and protein hydrolysate [68,69].

3. Innovative photobioreactors Many research works have been published with the aim to investigate the feasibility of microalgae cultivation, mainly for biofuel production. In these works, different photoautotrophic configurations have been tested for microalgae cultivation: however, not one has reached the hoped performances, and the costs to sustain microalgae biomass production are yet too high for a large part of the expected applications (see Introduction section). The main configurations tested in pilot-scale photoautotrophic plants were open ponds, tubular photobioreactors (vertical and horizontal), and flat panels. For all these tested configurations, the biomass production cost was higher than 3 V/kg. The main factors affecting the cost of produced biomass were characteristics directly linked to reactor design, such as operative costs given by energy consumption and labor and costs for depreciation [11,14e16,42]. Consequently, new reactor configurations can potentially improve the feasibility of microalgae biomass production. The aim of this section is to describe such new reactor configurations. In Subsections 3.1 and 3.2, thin-layer photobioreactors and biofilm reactors are described, respectively. In Subsection 3.3 other new promising configurations are described, which have received less attention.

IV. Selected examples and case history

3. Innovative photobioreactors

3.1 Thin-layer photobioreactors One of the main bottlenecks limiting microalgae productivity in photobioreactors is light limitation. To have high biomass productivity inside reactors, microalgae should grow with a fast growth rate (m w mmax) when they are at high biomass concentration (dX/dt ¼ mX). However, when microalgae grow, increasing their concentration (X), they progressively limit light penetration because of the shading effect. As a consequence, light available for microalgae cells is progressively reduced inside reactors during microalgae growth, and as cell concentration increases the amount of photons supplied per cell becomes progressively lower, making m < mmax [70,71]. The higher the optical path of the reactors is, then the lower the biomass concentration at which light becomes limiting for microalgae growth. The surface to volume ratio (S/V) of photobioreactors decreases by decreasing the optical path. The reactors widely tested for microalgae cultivation at pilot scale have optical paths between 0.02 and 0.3 m, corresponding to S/V ratio between 5 and 100 m2/m3. By using these configurations, microalgae biomass concentrations and biomass productivities typically reached were between 0.2 and 5 g/L and between 0.01 and 1 g/L per day, respectively [14,42,72]. Thin-layer photobioreactors can be defined as reactors having an optical path 0.01 m. These reactors have the potential to reach biomass concentrations and productivities much higher than the widely tested configurations. That is because light decrease inside a reactor follows an exponential trend, so a difference of few millimeters can lead to a relevant gain. However, the realization of reactors with a very low optical path requires one to face new issues that were not encountered with the conventional configurations. For instance, there is no space for the conventional probes used for pH and temperature control.

295

The thin-layer reactor designed by BCS Engineering (Brno, Czech Republic) is a modified open pond in which microalgae suspension is moved by a centrifugal pump instead of a paddle wheel. The suspension is circulated on the surface of a glass sheet (sustained by a steel frame) that is inclined 1.6%e1.7%, 18 m long, and 1 m width. The suspension layer is maintained at an optical path between 6 and 8 mm, with a 0.5 m/s speed. At the end of the sheet, the culture is collected on a tank and recirculated by the pump. During the day, CO2 is supplied to the culture, whereas during the night the whole suspension is maintained in the tank and aerated. Water evaporation is balanced by feeding water to the tank to keep a constant level [73,74]. By using this configuration, microalgae biomass concentrations between 8.9 and 29.7 g/L were achieved in about 15e30 days [73]. A very similar configuration was tested by Apel and coworkers too (5e5.6 mm optical path), by achieving biomass concentrations up to 50 g/L and biomass productivities of 2.4e2.9 g/L per day [75]. AlgoFilm is a photobioreactor with an optical path of 1.5 to 2 mm. It is similar to the reactors tested by Refs. [73,75], but it is, in contrast, a closed system, similar to an inclined flat panel, in which the suspension is continuously recirculated from the bottom part to the top part by using a peristaltic pump. Chlorella vulgaris was cultivated in this configuration by reaching a biomass concentration up to 30 g/L and a biomass productivity up to 7 g/L per day. Gifuni and coworkers investigated the performances of an ultrathin flat photobioreactor (UFP) with 3-mm optical path. The reactor was built up by using three polymethylmethacrylate panels (86  18 cm  0.5 mm) spaced out by two silicone sheets having 3 mm thickness. The panels were assembled in a sandwich structure by obtaining a reactor volume of 0.3 L and a back chamber used for cooling (by water recirculation). Chlorella sorokiniana was cultivated in this

IV. Selected examples and case history

296

16. New strategies enhancing feasibility of microalgal cultivations

latter reactor until reaching 24 g/L of biomass concentration and 11 g/L per day of biomass productivity [76]. The studies carried out prove that by using thin-layer photobioreactors microalgae concentration and productivities comparable to those achievable by heterotrophic cultivations can be attained [43]. This property may reduce the cost of the produced biomass with respect to the conventional technologies, but cost analyses are required from pilot-scale plants to consider also the higher operative and installation costs that will be likely given by the higher S/V ratio. In this direction, preliminary data obtained by Hulatt and Thoma indicate the net energy ratio calculated for biomass produced by flat panel reactors becomes more favorable by optical path reduction [77].

3.2 Biofilm photobioreactors Biofilm bioreactors can be seen as a way to work with an extremely reduced optical path. In these reactors, microalgae cells live attached to a surface within an amount of medium solution that is reduced to a minimum. The main advantages of the biofilm reactors are the high photosynthesis efficiency and microalgae productivity, which are obtained thanks to the facilitated gas exchange and light penetration. Moreover, the costs for harvesting are almost avoided because microalgae grow already separated from the liquid, with water content reduced to w70% [78]. Materials tested as surfaces for microalgae growth were cotton ropes, filter paper or synthetic polymers, stainlesssteel woven meshes, and sanded polycarbonate [79e81]. Biofilm reactors can be divided mainly in two categories: submerged cultures and porous substrate bioreactors (PSBRs). The typical configuration for a submerged culture biofilm reactor is made of a wheel covered with a material on which microalgae can easily grow attached and on which solution

medium can remain impregnated. The well rotates remaining partially (about 40%e50%) submerged in a tank containing medium solution [79,80]. With this configuration, microalgae grow attached to the external surface of the wheel. About a half of the microalgae remain submerged during cultivation, while another half remain constantly covered by a thin water film. Consequently, the configuration if these reactors still requires air/CO2 bubbling inside the cultivation medium (in the tank), and they require a further energy supply for wheel rotation, and about a half of the microalgae are actually under limited condition for light penetration and gas exchange [81]. To overcome these latter issues, PSBRs have been studied recently. These particular biofilm reactors are made of a porous material layer (e.g., filter paper) on which microalgae grow [82]. This layer separates microalgae cells from the cultivation medium. In detail, cells grow on one side of the layer of the porous material and are directly exposed to air, while the cultivation medium is only in contact with the other side of the layer (Fig. 16.4). Water moves from one side of the layer to the other side (the one with cells) by passing through the porous material. The movement of water can be given by different mechanisms such as capillarity, difference of pression, and water evaporation on the surface exposed to air. Mineral salts move in the water by the gradient diffusion between medium and cells. PSBR configuration allows one to maintain the advantages of the reduced optical path, it enhances gas exchange by cell exposition to air, and at the same time, it avoids the use of pumps for air feeding. A pump is required only to move the nutrient solution from the bottom to the top of the bioreactor [83]. The low mobility of the cells in the PSBR has been also indicated as a promising solution to contain contamination, because when that appears, it remains confined to a limited area. Very high productivities have been reached by using PSBRs. In fact, for Scenedesmus obliquus

IV. Selected examples and case history

3. Innovative photobioreactors

FIGURE 16.4

297

Schematic representation of the porous substrate bioreactor (PSBR) configuration.

and Arthrospira platensis, biomass productivities until 80 and 60 g/m2 per day were achieved, respectively [84,85]. The same works measured very high photosynthetic efficiency too, with values greater than 10% [85]. These values are significantly higher with respect to the values typically achieved by using conventional configurations pilot plants, which are between 5 and 20 g/m2 per day. However, these studies were carried out by maintaining the reactors under a closed chamber in which CO2 concentration in the air was strongly enriched (until 2%e5%) with respect to environmental air (0.04%). In contrast, by using the environmental CO2 concentration, a productivity value of 7 g/m2 per day was obtained; this value is comparable with those obtained from conventional configurations [86]. These data indicate that gas exchange from environmental air to the biofilm is the limiting factor

for microalgae growth in PSBRs. Some other relevant factors that can strongly affect the biomass productivity are light dilution generated by the design of PSBRs and biomass loss by respiration in the deep part of the biofilm [82]. More data are required from outdoor pilot plants operated for different seasons to carry out detailed cost analysis comparisons.

3.3 Other innovative solutions to improve phototrophic cultivation Besides the configurations described in Sections 3.1 and 3.2, there are some other systems that are being investigated to improve microalgae cultivation sustainability. One recent example is given by the liquid foam-bed photobioreactor [87]. This kind of reactor is based on the idea of growing microalgae on the surface

IV. Selected examples and case history

298

16. New strategies enhancing feasibility of microalgal cultivations

of foam bubbles. The advantages are similar to those of biofilm reactors. Microalgae are directly exposed to air, so they can grow faster by exploiting the low light path, and harvesting can be more efficient thanks to the higher biomass concentration. Microalgae are inoculated on a suspension containing a surfactant besides nutrients. The suspension is fed at the bottom of a photobioreactor together with a gas flux. By passing through a gas distributor plate, the foam is generated, filling the reactor to the top. Then the foam goes from the top of the reactor to a foam breaker (made, for example, of a fixed bed filled with plastic beads), and the medium is recirculated by a pump. A key factor that can influence the performance of the reactor is given by the surfactant. Ideally, it should have good foaming properties (e.g., foam stability and bubble size), it should be nonbiodegradable, chemically stable, biocompatible with algae, cheap, and it should allow a good repartition of microalgae to the foam phase. By comparing several surfactants, Pluronic F68 and P84 were found to be the best after the evaluation of different criteria [88]. When bovine serum albumin (BSA) was used as surfactant, a time of operativity of only 8 h was maintained, due to the instability of BSA [87]. In a first study, a lower growth rate with respect to conventional configurations was obtained due to the dark volume of the culture, which accounted for about 30% of total volume [87]. However, with an improved version of the liquid foam-bed photobioreactor, in which Pluronic F68 was used as surfactant and in which liquid recirculation was introduced, higher growth performances were achieved. Chlorella vulgaris and Chlorella sorokiniana were cultivated in continuous mode, for 500 h, maintaining biomass concentration around 20e25 g/L and reaching growth rate values to 0.1 h
1 [89]. To reduce water and energy consumption for temperature control in a photobioreactor, a solar control infrared blocking film (purchased by 3M) was tested [90]. The film prevented

infrared light penetration, reducing maximum temperature achieved by 33% and increasing by 52% and 64% biomass production and phycocyanin content, respectively, with respect to reactors without any temperature control system. The growth performances were comparable with respect to those achieved by using conventional temperature control systems as water jacket. Other innovative materials can be exploited to obtain reactor surfaces able to convert a part of the inactive region of the solar spectrum to wavelengths that can be actively used by photosynthesis [8]. Some authors investigated the efficacy of stacked waveguide reactors, which have micropatterned waveguide surfaces. These reactors exploit evanescent fields near the surface to enhance microalgae growth, which was increased two- to fourfold [91,92]. In some studies, the enhancement of microalgae productivity by fighting grazers was investigated. Grazers are organisms that can contaminate photoautotrophic microalgae cultures (especially open ponds) by decreasing biomass productivity, generating culture crashes. Following the same approach used to fight pests in terrestrial cultivations, several chemical compounds were studied for their effects on grazers and on microalgae; among these, 21 chemicals effective against grazers and compatible with Chlorella sp. were found. Benzalkonium chloride was tested in open pond, proving its ability in killing grazers and preventing negative effects on microalgae growth [93].

4. Wastewaters utilization as a source of nutrients The integration of microalgae cultivation with wastewater treatment is a promising approach that can improve the feasibility of microalgae biomass production. The main features that

IV. Selected examples and case history

4. Wastewaters utilization as a source of nutrients

may potentially increase the feasibility are as follows: -

-

-

-

Wastewater addition can reduce nutrient consumption, which is an issue both for economic (cost for nutrient purchase) and environmental reasons (resource depletion). Wastewater treatment can enhance economic sustainability by making the pollution treatment an added value of the process. Wastewater treatment can enhance environmental sustainability by reducing pollutants spreading in the environment. Organic molecules in wastewaters can work as substrate for mixotrophic and heterotrophic cultivations, which can increase remarkably biomass productivity and biomass concentration with respect to photoautotrophic conditions.

The treatment of several wastewaters has been tested for integration with microalgae cultivation. These include municipal wastewaters [94,95], manure wastewaters [96e98], digestate from anaerobic treatments [99,100], meat processing wastewater [101], cheese whey [102,103], olive mill wastewater [104,105], and textile wastewaters [106]. The aim of this section is to describe the latest innovations and the potentials of wastewater utilization in microalgae processes. In Subsection 4.1 the potential of wastewaters in nutrients supplementation is described, while in Subsection 4.2 the performances of microalgae processes in pollutants removal from wastewater are TABLE 16.2

299

reported. Subsequently, in Section 4.3, the room of improvement that can be achieved by the exploitation of organic substrates from wastewater by microalgae heterotrophic metabolism is outlined.

4.1 Wastewater as a source of nutrients for microalgae growth Photosynthesis is the metabolic pathway by which microalgae convert H2O and CO2 to organic compounds such as sugars (CH2O)n. However, H2O and CO2 are not the only nutrients required for microalgae growth. Other nutrients are required to supply elements such as N, P, S, Mg, and others, which are essential for the production of new microalgae cells [2]. The empirical formula for microalgae cells has been calculated in some different works, and the values determined are summarized in Table 16.2. For a sustainable, large-scale production of microalgae, it is essential that all the nutrients are used in a sustainable way to minimize resource depletion from the earth. It has been calculated that if the biodiesel required to support the European transportation market is produced entirely by microalgae, the amount of P and N required would be twice the amount that is presently produced as fertilizer in Europe [9]. Consequently, recycling nutrients is essential for a sustainable production of microalgae. Several studies have been conducted with the aim of studying the recycling of nutrients

Empirical formula for microalgae biomass.

Microalgae

Empirical formula

Source

-

CH2.4ON0.15P0.009

[107]

T. obliquus

CH1.9O0.6N0.13P0.012K0.01Mg0.006S0.003Ca0.0004

[108]

C. vulgaris

CH2O0.5N0.17P0.014K0.01Mg0.003

[43]

-

CH2.5ON0.15P0.009

[109]

IV. Selected examples and case history

300

16. New strategies enhancing feasibility of microalgal cultivations

coming from wastewaters in place of synthetic nutrients. In a large part of the studies carried out, the wastewaters were diluted with distilled water, tap water, or synthetic culture medium. That is because 100% of wastewaters often generate issues given by the high concentration of substances limiting microalgae growth by reducing light penetration or by their antimicrobial effects (such as the case of phenols and NHþ 4) [110,111]. Municipal wastewater (MW) is one of the most studied as a source of nutrients to support microalgae growth. MW has an N/P w 3 [112], lower than the ratio typically required for microalgae growth, which is w15 (Table 16.2). Therefore, nitrogen is the limiting nutrient in such wastewaters. Nitrogen concentration ranges between 10 and 100 mg/L in MW. Such a concentration can support a biomass production to 0.1e1 g/L before becoming limiting [113]. Generally, phosphorus is not a limiting nutrient; however, it should be considered that at some Ca2þ concentrations, which may be found in tap water, phosphorus can becomes easily limiting because of phosphates precipitation [114]. Digestate effluents originating from anaerobic treatments have higher nutrients concentrations with respect to MW. Nitrogen can vary in the range of 100e3500 mg/L and phosphorus between 20 and 400 mg/L. The N/P ratio is a bit more favorable too, about 3.6e4.3 [112]. However, nitrogen is present mainly as NHþ 4 , which has toxic effects for microalgae at too high concentrations [115]. There is not a fixed threshold limit for NHþ 4 concentration because its effect can vary depending on the microalgae strain and cultivation conditions. However, the reported values over which negative effects were detected ranged between 50 and 300 mg/L [116]. For this latter reason, digestates need to be diluted to be used as a nutrients source for microalgae cultivation. Moreover, it should be considered that microalgae alkalinize the pH by the photosynthetic activity. Therefore, if pH is not controlled the large part of the nitrogen could

be lost by means of NH3 evaporation þ (NHþ 4 þ H2O 4 NH3 þ H3O , pKa ¼ 9.25). Olive mill wastewater (OMW) is produced during the milling of olives to produce olive oil. OMW is characterized by a high content of phosphate and other mineral salts, but it has a low content of nitrogen available for microalgae growth [114,117]. To support microalgae growth, OMW typically requires an addition of nitrogen [118,119] that can be made also by mixing OMW with other wastewaters like urban wastewaters [120]. Cheese whey (CW), which is produced from cheese production plants, is rich in nitrogen and phosphorus. These nutrients are both in organic (mainly proteins) and inorganic forms (P-PO3
and NeNHþ 4 4 ). Proteins are generally removed from whey by filtrations to isolate whey proteins, which have high added value [103]. Inorganic nitrogen and inorganic phosphorus range between 65 and 160 mg/L and 300e400 mg/L, respectively [121]. The N/P ratio is w0.3, about 10-fold lower than MW and digestate effluents. It has been proven that CW can be used to sustain microalgae growth by replacing the addition of inorganic salts; however, only a few strains are able to use lactose as a substrate [103], which is mainly responsible for the high COD and BOD5 content in CW [121]. The organic substrates that can be found in wastewaters (generally measured as COD, BOD5, or TOC) can furnish further nutrients for microalgae growth. They can be a source of energy and mass (C, H, O, N, S, P) for microalgae metabolism. The exploitation of this latter class of nutrients is described in detail in Section 4.3. Recently, the interest about the possibility to recycle directly nutrients from microalgae biomass is growing too. When the target products from microalgae are, for example, TAGs and starch, N and P are not included in the products; consequently, their recycling may be possible in an efficient process. Microalgae biomass can be used as substrate for anaerobic digestion, allowing one to recover about 40 g of

IV. Selected examples and case history

4. Wastewaters utilization as a source of nutrients

N per kg of dry biomass (69%e86%) and 3.8 g of P per kg of dry biomass (9%e49%) [122]. Hydrothermal liquefaction can be also used to recycle nutrients from microalgae biomass, allowing one to recover about 26 g of N per kg of dry biomass (15%e84%) and 6.8 g of P per kg of dry biomass (20%e85%) [122].

4.2 Wastewater treatment by microalgae Microalgae cultivation can reduce the polluting load of wastewaters by removing organic and inorganic molecules. Such removal can be obtained by several mechanisms, such as enzymatic conversion to other molecules, storage inside cells (bioaccumulation), adsorption on cell surface (bioadsorption), variation of chemicalephysical parameters (e.g., pH, O2 concentration), and action of bacteria that live in association with microalgae. For example, inorganic N and P can be accumulated inside cells and then used as a substrate by the enzymes to produce proteins, membranes, and nucleic acids. Heavy metals can be bioaccumulated and/or adsorbed [123,124]. COD can be reduced by converting organic molecules to CO2 and H2O (mineralization) or to other organic molecules with a lower oxygen demand. The pH increase induced by photosynthetic activity can boost phosphates and metals removal by precipitation and adsorption [114,123,125]. From MWs, high phosphorus and nitrogen removal, between 80% and 100%, has been usually obtained. These results were achieved by using several microalgae species (e.g., Chlorella sp. and Scenedesmus sp.) and several reactor configurations (open ponds, photobioreactors, immobilized cells) [126e129]. COD, TOC, and BOD removal has been also obtained at comparable high values, close to 100%. However, for these latter parameters, which are linked to the organic compounds, a main role is played by the bacteria communities that live in symbiosis with

301

microalgae [130]. Bacteria contamination is a limited issue for this kind of wastewater because the initial COD concentration is quite low (between 100 and 500 mg/L) [131]. Several studies have been conducted to exploit a synergic interaction between microalgae and bacteria. It should be considered that in conventional wastewater treatment plants, the aeration required to support aerobic bacteria can account for 45% e75% of the energy consumption [132]. By using a mixed culture of microalgae and bacteria, microalgae can produce O2 by photosynthesis, reducing the energy supply for aeration and furnishing at the same time O2 for COD reduction by bacteria. Bacteria oxidation of organic molecules converts the O2 to CO2, which can be used again by microalgae metabolism in a closed cycle [133,134]. Digestate produced from anaerobic fermentation can be also efficiently treated by microalgae culture to reduce the concentration of pollutants. Removal efficiencies of N and P to 70%e100% have been obtained for digestate produced from swine manure, dairy manure, MWs, poultry wastewaters, and others [116]. Because of the higher pollutant concentration in anaerobic digestates with respect to MWs, they often require a pretreatment before microalgae cultivation. Generally, the pretreatments carried out are dilution, filtration, precipitation, sterilization, and centrifugation [116]. The removal of COD can achieve efficiencies to 50%e90%. However COD initial concentration has higher values with respect to MWs, generating a relevant issue by biologic contamination [116]. In anaerobic digestate, the high content of metal ions (Cu2þ, Pb2þ, Zn2þ, Co2þ, and others) is also a relevant issue. Microalgae growth can reduce significantly the concentration of metals and heavy metals; this has been proven, for example, for Zn2þ and Cd2þ [135]. However the large part of the studies carried out on this field did not evaluate this aspect. The direct cultivation of microalgae in agroindustrial wastewaters such as CW has been

IV. Selected examples and case history

302

16. New strategies enhancing feasibility of microalgal cultivations

proven, allowing lactose removal to 64% [102] and nitrogen removal to 100% [103]. However, lactose can be used as substrate only by some microalgae strains [103]. Moreover, in the reported studies, all the experiments were carried out under fully sterilized conditions, which are hardly applicable to large-scale cultivation. Microalgae cultivation on diluted OMW was efficient in phenols removal to 50%e60% [105,110,114], COD reduction to 30%e40% [118] and phosphates removal to 91% [114].

4.3 Enhanced microalgae growth by exploiting organic substrates from wastewaters The large part of wastewaters contains significant amounts of organic substrates. Many microalgae can use such organic substrates, by means of a heterotrophic metabolism, as a source of carbon and energy, in addition to light and CO2 exploited by the photoautotrophic metabolism. When the heterotrophic metabolism is operated concurrently with photoautotrophic metabolism, the microalgae metabolism is called mixotrophic (heterotrophic þ photoautotrophic). This further source of energy and carbon gives a room of improvement for microalgae biomass production processes. In this subsection, such possibility of improvement is described. 4.3.1 Microalgae growth on organic substrates The organic molecules that are present in wastewaters have the possibility to boost microalgae productivity by feeding their heterotrophic metabolism. Heterotrophic metabolism is independent of light penetration inside reactors; consequently, it could allow achieving high biomass concentration and biomass productivity by cultivating microalgae in reactors with a low S/V ratio. Microalgae can use a large variety of organic molecules as source of carbon and energy;

however, this ability can vary from one strain to another strain. Glucose is likely the most tested substrate, and it has been used for the cultivation of a large number of different strains including Chlorella protothecoides, Chlorella regularis, Chlorella vulgaris, Chromochloris zofingiensis, Crypthecodinium cohnii, Dunaliella sp., Euglena gracilis, Galdieria sulphuraria, Scenedesmus sp., and many others [43,136]. Other sugars have been studied less. Sucrose is generally not useable directly by microalgae because of the absence of invertase [137]. Lactose can be used only by a few strains [102,103]. Glycerol and acetate have been widely used to support the growth of several microalgae species [43,136,138]. Other substrates that have been also used for microalgae cultivations are some amino acids (such as glutamate, aspartate, asparagine), butyric acid, ethanol, galactose, mannose, and fructose [43,136,139]. When cultivated under a heterotrophic regime, microalgae have yield factors that usually range from 0.3 to 0.7 g of dry weight biomass per gram of organic substrate [43]. The yield factor varies depending on the strain, the kind of organic substrate, and the cultivation conditions. Considering that several wastewaters have concentrations of organic compounds to w50 g/L, it can be deduced that more than 30 g/L of microalgae biomass may be obtained by adding wastewater as a nutrient source. Moreover, because of the independence of the heterotrophic metabolism from the light supply, it should be possible to obtain also high biomass productivities by working with low-cost configurations (low S/V ratio). However, similar improvements in microalgae biomass production and productivity have been registered only for microalgae cultivated in fully sterilized conditions [43]. That is because, without an accurate sterilization process and without an axenic microalgae culture, the organic substrate is used more efficiently by the contaminant bacteria [116,140]. Bacteria can easily outclass microalgae growth in the

IV. Selected examples and case history

4. Wastewaters utilization as a source of nutrients

presence of high substrate concentration because their maximum growth rate (w20 d
1) is about one order of magnitude higher than microalgae (w2 d
1). Several studies have been also conducted by using unsterilized wastewaters, namely by working with a consortia of bacteria and microalgae. However, in these cases the attention was mainly focused on wastewater pollutants removal, and the initial COD concentrations were generally low (<1 g/L) because of the kind of wastewater used (e.g., MWs) or because of wastewater dilution [129,134,141,142]. In these conditions, contamination is strongly limited by the low COD, but at the same time, microalgae growth is mainly phototrophic, without any relevant advantage given by the organic substances. Only in some cases, unsterilized wastewater was used to exploit directly the advantage given by the supplemented organic substrates. For example, OMW was added in a two-stage phototrophic-heterotrophic cultivation process. In such a process, microalgae were first cultivated in a photoautotrophic regime, to reach nitrogen depletion, and then the wastewater was added generating biomass production and productivity comparable to the photoautotrophic control [19,105]. Although there was not an improvement in biomass growth with respect to the control, the possibility to obtain a comparable growth in a reactor with lower S/V ratio was already a gain. The use of wastewaters as a source of organic substrates to improve microalgae biomass productivity has not yet been proven in a way that is technically promising. The issue of bacterial contamination to nonaxenic conditions is the main limit that needs to be solved. 4.3.2 Innovative strategies for organic substrate addition Some innovative strategies have been investigated with the aim to control bacteria growth in nonaxenic microalgae cultivations. A factor that

303

can be exploited to favoring microalgae growth over bacteria is the different ability to store nutrients [109]. Microalgae growth is generally described by a Droop model [113,143], which describes the ability of microalgae to store nitrogen and to grow in N-starvation condition by using the internal nitrogen quota. Instead, a large part of bacteria show a reduced ability to store nutrients, and their growth is generally described by a Monod model, which means that growth is stopped as soon as nutrients are depleted in the cultivation media [140]. This difference was exploited by a mixotrophic cultivation in which glucose and nitrate were added alternatively [144]. In the described alternate feeding strategy, a fixed amount of glucose and nitrate were added separately and alternatively every 2 days, under mixotrophic regime. The strategy was compared with a mixotrophic control in which glucose and nitrate were added simultaneously every 2 days. The authors found that with the alternate strategy, bacterial contamination was strongly reduced. During the alternate strategy, there was an alternance of low glucose/high nitrate concentration condition and low nitrate/high glucose concentration condition. Microalgae could grow under the low nitrate concentration condition (N-starvation) by exploiting the internal nitrogen quota, and under the low glucose concentration condition by using the extra inorganic carbon (CO2) by means of photosynthesis. The efficacy of a similar strategy was assessed also in heterotrophic condition [177]. In this latter case, the ability of microalgae to store organic substrates was exploited in addition to the nitrogen storage. In fact, in heterotrophic condition, the growth of microalgae under high nitrate/low glucose concentration condition was only possible by exploiting the microalgae ability to use solely the internal quota of organic molecules stored as a source of energy (mainly starch and lipids). In this strategy, named uncoupled heterotrophic strategy (UHS), glucose and nitrate were added alternatively,

IV. Selected examples and case history

304

16. New strategies enhancing feasibility of microalgal cultivations

by tailoring their initial concentrations as a function of the biomass concentration and of the internal quota of organic compounds stored. Moreover, the time between nitrate addition and the subsequent glucose addition was tailored as a function of the consumption kinetic. These adjustments on glucose and nitrate addition avoided low biomass growth or high bacterial contamination given by underestimation or overestimation of the substrate consumption in every phase. With the UHS strategy, a strong reduction in bacteria concentration was achieved, and at the same time, 6 g/L per day and 26 g/L of biomass productivity and biomass concentration were achieved, respectively.

producing bigger fruits and seeds instead of in defense against the environment. The differences between maize and its ancestor teosinte (Zea mays ssp. mexicana) can be considered an example [146]. The large part of the current agricultural production derives from the genetic improvements obtained by the “green revolution” [145]. Analogous to what was done with terrestrial plants, the future of the cultivation of microalgae will be likely strongly related to the genetic improvements. A “green revolution” is expected to be a key factor to improve the feasibility of microalgae cultivation processes [147]. In the following subsections, the main metabolic pathways that have been manipulated in microalgae by means of genetic modification are described.

5. Strain improvement Microalgaecultivationcanbeseenasanewkindof agriculture. The conventional agriculture that we knowistheresultofthousandsofyearsofimprovements. These changes have not been directed only byagronomicpractices,butakeyandfundamental rolewasalsoplayedbythegeneticimprovementof the species cultivated [145]. The reason at the base of this latter fact is that plants in nature are the results of millions of years of evolution, but this evolution was mainly directed by their fitness, which means by their ability to reproduce themselves in the environmental conditions in which they live and in the new environmental conditions with which they had to adapt during the years. Consequently, the parameters that have been improved in plants living in nature were, for example, the production of resistant seeds, leaves, and stems to face herbivorous and competitors. By means of agriculture, several species were genetically improved toward the characteristics requested for cultivation. The natural environment was replaced by the agricultural field, a more comfortable environment, generated by the human work, in which plants could grow using more energy in

5.1 Improvement of photosynthetic efficiency Photosynthetic efficiency of microalgae is measured as the ratio between energy supplied to microalgae by light illumination and the energy content of the microalgae biomass produced by photosynthesis. It should be considered that from the atmosphere to the ground, only a fraction of the light energy, estimated between 10% and 34%, reaches the external surface of the reactors [8]. The energy is lost due to atmospheric scattering, weather conditions, latitude, reactor orientation, and the limited photosynthetic active region. Then, other losses can be given by light saturation of cells and by energy consumption of the microalgae metabolism. Considering only the incident light on the reactor surface, the typical photosynthetic efficiency registered is around 0.1%e10% and the theoretical maximum value is w12% [8]. A factor that can reduce photosynthetic efficiency is photoinhibition, which can occur when microalgae are exposed to too high a flux of photons. In this condition, the rate of photon absorption by light harvesting antenna becomes higher

IV. Selected examples and case history

5. Strain improvement

than the rate at which photons are utilized by metabolism. The excess of photons (to 80%) is lost as heat, damaging the photosystems [148]. To overcome this issue, some genetically modified strains of Chlamydomonas reinhardtii have been produced, which have a truncated light harvesting antenna [149e151]. These strains have reduced the amount of chlorophyll for light harvesting, making them less sensitive to photoinhibition. Moreover, their reduced light adsorption per unit cell reduces the self-shading effect inside photobioreactors too.

5.2 Improvement in target compound accumulation As well as maize that was genetically improved to accumulate more starch in bigger cobs with respect to its wild ancestor, the same approach can be followed to induce microalgae to accumulate higher amounts of target compounds. In this direction, several microalgae strains able to accumulate higher amounts of TAG have been obtained by inhibiting the synthesis of starch (starchless mutants). Most microalgae accumulate organic carbon during N-starvation both as TAG and as starch, which are two competitive pathways. Limiting the starch pathway, the flux of carbon assimilation is directed mainly to TAG synthesis. Starchless mutant strains have been obtained for Chlorella pyrenoidosa, Chlamydomonas reinhardtii, and Tetradesmus obliquus, showing a higher level of TAG accumulation [152,153]. Other pathways can be also changed to improve lipid accumulation; for example, in Phaeodactylum tricornutum the neutral lipid content was increased by 82% by inactivating pyruvate dehydrogenase kinase [154]. For the same species, an increase in lipid content up to 2.5-fold was obtained with a transgenic strain that overexpressed malic enzyme [155]. Chlamydomonas reinhardtii mutants with a more than twice improved starch accumulation

305

ability, with respect to wild-type control, were obtained by using gamma irradiation [156]. For the same species, a mutant deficient for a dualspecificity tyrosine phosphorylation-regulated kinase showed a higher level of starch accumulation with respect to wild type [157]. Some researchers attempted to enhance accumulation of pigments too. By expressing a phytoene synthase gene (CzPSY) from Chlorella zofingiensis to Chlamydomonas reinhardtii, 2.0- and 2.2-fold increases were obtained for violaxanthin and lutein accumulation, respectively [158].

5.3 Improvement in strain resistance to adverse conditions The environment inside bioreactors is controlled within certain values to maintain the main environmental parameters around the optimal values. However, two things should be considered: (1) it is impossible to control every parameter, and there will be invariably some parameters that may vary until producing stressful conditions for microalgae. (2) Every control system has a cost that affects the economic sustainability of the cultivation process. Therefore, the ability of microalgae strains to resist to the adverse/nonoptimal conditions encountered during cultivation is a key factor to consider in strain selection and improvement. Without any control system, temperature can vary inside a photobioreactor in the range of below 0e50 C, affecting strongly microalgae growth. A lot of energy is required to cool down water temperature during warm seasons and to raise water temperature during winter seasons [159]. For this reason, many efforts have been made looking for strains more tolerant to temperature fluctuations. A mutant of Chlorella sp., obtained by chemical mutagenesis, showed higher tolerance to high temperature (to 45 C) with respect to wild type [160]. Similarly, a thermotolerant mutant of

IV. Selected examples and case history

306

16. New strategies enhancing feasibility of microalgal cultivations

Chlorella pyrenoidosa (NCIM 2738) was cultivated under subtropical outdoor conditions corresponding to 43e45 C during the day and 27e30 C during the night [161]. Five mutant strains of Arthrospira were obtained by ethyl methanesulfonate mutagenesis. These strains grew better than the wild-type strain at low temperature, by achieving biomass production 10-fold higher than wild type at 16 and 18 C [162]. Improvement of Chlamydomonas growth was obtained at 10 C by using a diploid strain [163] and at 4 C by using AMP deaminase suppression [164]. The overexpression of hspA and osmotin genes in Synechococcus elongatus induced a coupled improved tolerance to high temperature (45 C), high salt concentration (3.5%), and high illumination (300 mmol/m2 per second) [165]. Another important aspect is CO2 concentration. CO2 is often a limiting nutrient in photobioreactors; consequently, to obtain higher biomass productivities, the photobioreactors could be supplied with higher CO2 concentrations in the feeding gas. However, CO2 concentrations higher than 10% can have toxic affects on microalgae [166,167]. For this reason, different works have been conducted to isolate microalgae strains more tolerant to high CO2 levels. Strains able to tolerate CO2 concentrations to 20%e30% have been obtained [167,168]. Apart from the main environmental parameters considered during microalgae cultivation, several pollutants present in liquid and air streams feeding the reactors can affect microalgae growth too. Utilization of flue gas in place of pure CO2 to supply photobioreactors can potentially reduce production costs. However, flue gas contains other compounds beyond CO2, such as NOx and SOx, that can limit microalgae growth [169]. A Chlorella sp. strain tolerant to flue gas was obtained by 46 cycles of adaptive evolution, showing the same growth on synthetic flue gas compared to pure CO2 [170]. Several microalgae strains resistant to pollutants found in different wastewaters

have been selected, for example, microalgae strains more tolerant to phenols [171], to pesticides [93,172], to heavy metals [173], to extremely acid pH [174,175], and to high salinity [165,176].

6. Conclusions and future trends It can be concluded that although the cost of microalgae biomass production is limiting for a large part of the possible industrial applications, this technology has a large room for improvement. The future trends expected for the scientific and industrial research in microalgae processes development are as follows: (a) the development of new reactors by which energy and cost associated with the cultivation of microalgae can be reduced; (b) the development of new microalgae strains for a new “green revolution” by which the efficiency of target compounds production in the reactor environment can be maximized; (c) the development of cultivation processes integrated with wastewater treatment for the exploitation of organic substrates and for nutrient recycling; (d) the development of tailored systems to control biologic contaminants; (e) the development of biorefinery facilities by which several end products can be produced starting from microalgae biomass.

List of abbreviations and acronyms Adenosine monophosphate Biochemical oxygen demand (measured after 5 days of incubation) BSA Bovine serum albumin COD Chemical oxygen demand CW Cheese whey DHA Docosahexaenoic acid MW Municipal wastewater OMW Olive mil wastewater PSBR Porous substrate bioreactors

AMP BOD5

IV. Selected examples and case history

307

References

TAGs UFP

Triacylglycerols Ultrathin flat photobioreactor [13]

References [1] E. Walz, S. Energy, E. Walz, Biotech’s green gold? Nature Biotechnology 27 (2009) 15e18, https://doi.org/ 10.1038/nbt0109-15. [2] G. Markou, D. Vandamme, K. Muylaert, Microalgal and cyanobacterial cultivation: the supply of nutrients, Water Research 65 (2014) 186e202, https://doi.org/10.1016/j.watres.2014.07.025. [3] P. Spolaore, C. Joannis-Cassan, E. Duran, A. Isambert, Commercial applications of microalgae, Journal of Bioscience and Bioengineering 101 (2006) 87e96, https://doi.org/10.1263/jbb.101.87. [4] J.A. Garrido-Cardenas, F. Manzano-Agugliaro, F.G. Acien-Fernandez, E. Molina-Grima, Microalgae research worldwide, Algal Research 35 (2018) 50e60, https://doi.org/10.1016/j.algal.2018.08.005. [5] K.M. Weyer, D.R. Bush, A. Darzins, B.D. Willson, Theoretical maximum algal oil production, Bioenergy Research 3 (2010) 204e213, https://doi.org/10.1007/ s12155-009-9046-x. [6] X.G. Zhu, S.P. Long, D.R. Ort, What is the maximum efficiency with which photosynthesis can convert solar energy into biomass? Current Opinion in Biotechnology 19 (2008) 153e159, https://doi.org/10.1016/ j.copbio.2008.02.004. [7] M.R. Tredici, Photobiology of microalgae mass cultures: understanding the tools for the next green revolution, Biofuels 1 (2010) 143e162, https:// doi.org/10.4155/bfs.09.10. [8] M.D. Ooms, C.T. Dinh, E.H. Sargent, D. Sinton, Photon management for augmented photosynthesis, Nature Communications 7 (2016) 12699, https:// doi.org/10.1038/ncomms12699. [9] R.H. Wijffels, M.J. Barbosa, An outlook on microalgal biofuels, Science 329 (2010) 796e799, https:// doi.org/10.1126/science.1189003. [10] K.S. Dhugga, Maize biomass yield and composition for biofuels, Crop Science 47 (2007) 2211e2227, https://doi.org/10.2135/cropsci2007.05.0299. [11] M.R. Tredici, L. Rodolfi, N. Biondi, N. Bassi, G. Sampietro, Techno-economic analysis of microalgal biomass production in a 1-ha Green Wall Panel (GWPÒ) plant, Algal Research 19 (2016) 253e263, https://doi.org/10.1016/j.algal.2016.09.005. [12] M. Vigani, C. Parisi, E. Rodríguez-Cerezo, M.J. Barbosa, L. Sijtsma, M. Ploeg, C. Enzing, Food and feed products from micro-algae: market

[14]

[15]

[16]

[17]

[18]

[19]

[20]

[21]

[22]

opportunities and challenges for the EU, Trends in Food Science and Technology 42 (2015) 81e92, https://doi.org/10.1016/j.tifs.2014.12.004. J.M. MacDonald, P. Korb, R.A. Hoppe, Farm Size and the Organization of U.S. Crop Farming, ERR-152, 2013. http://www.ers.usda.gov/media/1156726/ err152.pdf. F.G. Acien, J.M. Fernandez, J.J. Magan, E. Molina, Production cost of a real microalgae production plant and strategies to reduce it, Biotechnology Advances 30 (2012) 1344e1353, https://doi.org/10.1016/ j.biotechadv.2012.02.005. J. Ruiz, G. Olivieri, J. de Vree, R. Bosma, P. Willems, J.H. Reith, M.H.M. Eppink, D.M.M. Kleinegris, R.H. Wijffels, M.J. Barbosa, Towards industrial products from microalgae, Energy and Environmental Science 9 (2016) 3036e3043, https://doi.org/10.1039/ C6EE01493C. N.H. Norsker, M.J. Barbosa, M.H. Vermuë, R.H. Wijffels, Microalgal production e a close look at the economics, Biotechnology Advances 29 (2011) 24e27, https://doi.org/10.1016/ j.biotechadv.2010.08.005. C.Y. Chen, X.Q. Zhao, H.W. Yen, S.H. Ho, C.L. Cheng, D.J. Lee, F.W. Bai, J.S. Chang, Microalgae-based carbohydrates for biofuel production, Biochemical Engineering Journal 78 (2013) 1e10, https://doi.org/ 10.1016/j.bej.2013.03.006. S. Pierucci, J.J. Klemes, L. Piazza, S. Bakalis, I. Gifuni, G. Olivieri, I.R. Krauss, G. D’errico, A. Pollio, A. Marzocchella, Microalgae as new sources of starch: isolation and characterization of microalgal starch granules, Chemical Engineering Transactions (2017), https://doi.org/10.3303/CET1757238. F. Di Caprio, A. Visca, P. Altimari, L. Toro, B. Masciocchi, G. Iaquaniello, F. Pagnanelli, Two stage process of microalgae cultivation for starch and carotenoid production, Chemical Engineering Transactions 49 (2016) 415e420, https://doi.org/10.3303/ CET1649070. C. Schulze, M. Wetzel, J. Reinhardt, M. Schmidt, L. Felten, S. Mundt, Screening of microalgae for primary metabolites including b-glucans and the influence of nitrate starvation and irradiance on b-glucan production, Journal of Applied Phycology 28 (2016) 2719e2725, https://doi.org/10.1007/s10811-016-0812-9. H.M.D. Wang, C.C. Chen, P. Huynh, J.S. Chang, Exploring the potential of using algae in cosmetics, Bioresource Technology 184 (2015) 355e362, https://doi.org/10.1016/j.biortech.2014.12.001. T. Chiong, C. Acquah, S.Y. Lau, E.H. Khor, M.K. Danquah, Microalgal-based protein by-products:

IV. Selected examples and case history

308

[23]

[24]

[25]

[26]

[27]

[28]

[29]

[30]

16. New strategies enhancing feasibility of microalgal cultivations

extraction, purification, and applications, in: Protein Byprod. Transform. From Environ. Burd. Into ValueAdded Prod., 2016, pp. 213e234, https://doi.org/ 10.1016/B978-0-12-802391-4.00012-4. M. Hayes, H. Skomedal, K. Skjånes, H. MazurMarzec, A. Toru nska-Sitarz, M. Catala, M. Isleten Hosoglu, M. García-Vaquero, Microalgal proteins for feed, food and health, in: Microalgae-Based Biofuels Bioprod. From Feed. Cultiv. to End-Products, 2017, pp. 347e368, https://doi.org/10.1016/B978-0-08101023-5.00015-7. M.A. Zeller, R. Hunt, A. Jones, S. Sharma, Bioplastics and their thermoplastic blends from Spirulina and Chlorella microalgae, Journal of Applied Polymer Science 130 (2013) 3263e3275, https://doi.org/10.1002/ app.39559. S. Kumar, E. Hablot, J.L.G. Moscoso, W. Obeid, P.G. Hatcher, B.M. Duquette, D. Graiver, R. Narayan, V. Balan, Polyurethanes preparation using proteins obtained from microalgae, Journal of Materials Science 49 (2014) 7824e7833, https://doi.org/ 10.1007/s10853-014-8493-8. C.A. Ovando, J.C. de Carvalho, G. Vinícius de Melo Pereira, P. Jacques, V.T. Soccol, C.R. Soccol, Functional properties and health benefits of bioactive peptides derived from Spirulina: a review, Food Reviews International 34 (2018) 34e51, https://doi.org/ 10.1080/87559129.2016.1210632. P. Chiaiese, F. Palomba, F. Tatino, C. Lanzillo, G. Pinto, A. Pollio, E. Filippone, Engineered tobacco and microalgae secreting the fungal laccase POXA1b reduce phenol content in olive oil mill wastewater, Enzyme and Microbial Technology 49 (2011) 540e546, https://doi.org/10.1016/ j.enzmictec.2011.06.002. S. Godet, C. Loiseau, G. Pencreac’h, F. Ergan, J. Herault, Isolation and sequence analysis of a cdna encoding a novel putative esterase from the marine microalga isochrysis galbana (prymnesiophyceae, haptophyta), Journal of Phycology 46 (2010) 679e684, https://doi.org/10.1111/j.15298817.2010.00838.x. S. Godet, J. Herault, G. Pencreac’h, F. Ergan, C. Loiseau, Isolation and analysis of a gene from the marine microalga Isochrysis galbana that encodes a lipase-like protein, Journal of Applied Phycology 24 (2012) 1547e1553, https://doi.org/10.1007/s10811012-9815-3. E. Suarez Garcia, J. van Leeuwen, C. Safi, L. Sijtsma, M.H.M. Eppink, R.H. Wijffels, C. van den Berg,

[31]

[32]

[33]

[34]

[35]

[36]

[37]

[38]

[39]

Selective and energy efficient extraction of functional proteins from microalgae for food applications, Bioresource Technology 268 (2018) 197e203, https:// doi.org/10.1016/j.biortech.2018.07.131. E.S. Garcia, J.J.A. Van Leeuwen, C. Safi, L. Sijtsma, L.A.M. Van Den Broek, M.H.M. Eppink, R.H. Wijffels, C. Van Den Berg, Techno-Functional properties of crude extracts from the green microalga Tetraselmis suecica, Journal of Agricultural and Food Chemistry 66 (2018) 7831e7838, https://doi.org/ 10.1021/acs.jafc.8b01884. F. Di Caprio, F. Pagnanelli, R.H. Wijffels, D. Van der Veen, Quantification of Tetradesmus obliquus (Chlorophyceae) cell size and lipid content heterogeneity at single-cell level, Journal of Phycology 54 (2018) 187e197, https://doi.org/10.1111/jpy.12610. C. Largeau, E. Casadevall, C. Berkaloff, P. Dhamelincourt, Sites of accumulation and composition of hydrocarbons in Botryococcus braunii, Phytochemistry 19 (1980) 1043e1051, https://doi.org/ 10.1016/0031-9422(80)83054-8. J. Kim, G. Yoo, H. Lee, J. Lim, K. Kim, C.W. Kim, M.S. Park, J.W. Yang, Methods of downstream processing for the production of biodiesel from microalgae, Biotechnology Advances 31 (2013) 862e876, https://doi.org/10.1016/ j.biotechadv.2013.04.006. P. Roesle, F. Stempfle, S.K. Hess, J. Zimmerer, C. Ríobartulos, B. Lepetit, A. Eckert, P.G. Kroth, S. Mecking, Synthetic polyester from algae oil, Angewandte Chemie International Edition 53 (2014) 6800e6804, https://doi.org/10.1002/anie.201403991. B.S. Grama, S.N. Agathos, C.S. Jeffryes, Balancing photosynthesis and respiration increases microalgal biomass productivity during photoheterotrophy on glycerol, ACS Sustainable Chemistry and Engineering 4 (2016) 1611e1618, https://doi.org/10.1021/ acssuschemeng.5b01544. O.P. Ward, A. Singh, Omega-3/6 fatty acids: alternative sources of production, Process Biochemistry 40 (2005) 3627e3652, https://doi.org/10.1016/ j.procbio.2005.02.020. T.C. Adarme-Vega, D.K.Y. Lim, M. Timmins, F. Vernen, Y. Li, P.M. Schenk, Microalgal biofactories: a promising approach towards sustainable omega-3 fatty acid production, Microbial Cell Factories 11 (2012), https://doi.org/10.1186/1475-2859-11-96. R.R. Ambati, P.S. Moi, S. Ravi, R.G. Aswathanarayana, Astaxanthin: sources, extraction, stability, biological activities and its commercial

IV. Selected examples and case history

309

References

[40]

[41]

[42]

[43]

[44]

[45]

[46]

[47]

[48]

[49]

applications e a review, Marine Drugs 12 (2014) 128e152, https://doi.org/10.3390/md12010128. F. Di Caprio, P. Altimari, L. Toro, F. Pagnanelli, Effect of lipids and carbohydrates extraction on astaxanthin stability in Scenedesmus sp, Chemical Engineering Transactions 43 (2015) 205e210, https://doi.org/ 10.3303/CET1543035. J.H. Lin, D.J. Lee, J.S. Chang, Lutein production from biomass: marigold flowers versus microalgae, Bioresource Technology 184 (2015) 421e428, https:// doi.org/10.1016/j.biortech.2014.09.099. R. Bosma, J.H. de Vree, P.M. Slegers, M. Janssen, R.H. Wijffels, M.J. Barbosa, Design and construction of the microalgal pilot facility AlgaePARC, Algal Research 6 (2014) 160e169, https://doi.org/ 10.1016/j.algal.2014.10.006. F. Bumbak, S. Cook, V. Zachleder, S. Hauser, K. Kovar, Best practices in heterotrophic high-celldensity microalgal processes: achievements, potential and possible limitations, Applied Microbiology and Biotechnology 91 (2011) 31e46, https://doi.org/ 10.1007/s00253-011-3311-6. M.L. Gerardo, S. Van Den Hende, H. Vervaeren, T. Coward, S.C. Skill, Harvesting of microalgae within a biorefinery approach: a review of the developments and case studies from pilot-plants, Algal Research 11 (2015) 248e262, https://doi.org/10.1016/ j.algal.2015.06.019. M. Al Hattab, A. Ghaly, Microalgae oil extraction pretreatment methods: critical review and comparative analysis, Journal of Fundamentals of Renewable Energy and Applications 5 (2015) 1e26, https:// doi.org/10.4172/2090-4541.1000172. T. Dong, E.P. Knoshaug, P.T. Pienkos, L.M.L. Laurens, Lipid recovery from wet oleaginous microbial biomass for biofuel production: a critical review, Applied Energy 177 (2016) 879e895, https:// doi.org/10.1016/j.apenergy.2016.06.002. M.A. Hejazi, E. Holwerda, R.H. Wijffels, Milking microalga Dunaliella salina for b-Carotene production in two-phase bioreactors, Biotechnology and Bioengineering 85 (2004) 475e481, https://doi.org/10.1002/ bit.10914. N.R. Moheimani, R. Cord-Ruwisch, E. Raes, M.A. Borowitzka, Non-destructive oil extraction from Botryococcus braunii (Chlorophyta), Journal of Applied Phycology 25 (2013) 1653e1661, https:// doi.org/10.1007/s10811-013-0012-9. F. Zhang, L.H. Cheng, X.H. Xu, L. Zhang, H.L. Chen, Screening of biocompatible organic solvents for enhancement of lipid milking from Nannochloropsis sp, Process Biochemistry 46 (2011) 1934e1941, https://doi.org/10.1016/j.procbio.2011.06.024.

[50]

[51]

[52]

[53]

[54]

[55]

[56]

[57]

[58]

[59]

[60]

E. G€ unerken, E. D’Hondt, M.H.M. Eppink, L. GarciaGonzalez, K. Elst, R.H. Wijffels, Cell disruption for microalgae biorefineries, Biotechnology Advances 33 (2015) 243e260, https://doi.org/10.1016/ j.biotechadv.2015.01.008. R. Halim, M.K. Danquah, P.A. Webley, Extraction of oil from microalgae for biodiesel production: a review, Biotechnology Advances 30 (2012) 709e732, https://doi.org/10.1016/j.biotechadv.2012.01.001. P.R. Postma, E. Suarez-Garcia, C. Safi, G. Olivieri, R.H. Wijffels, Energy efficient bead milling of microalgae: effect of bead size on disintegration and release of proteins and carbohydrates, Bioresource Technology 224 (2017) 670e679, https://doi.org/10.1016/ j.biortech.2016.11.071. M. Wang, W. Yuan, X. Jiang, Y. Jing, Z. Wang, Disruption of microalgal cells using high-frequency focused ultrasound, Bioresource Technology 153 (2014) 315e321, https://doi.org/10.1016/ j.biortech.2013.11.054. F.J. Petracek, L. Zechmeister, Determination of partition coefficients of carotenoids as a tool in pigment analysis, Analytical Chemistry 28 (1956) 1484e1485, https://doi.org/10.1021/ac60117a041. R. Dineshkumar, S.K. Dash, R. Sen, Process integration for microalgal lutein and biodiesel production with concomitant flue gas CO2 sequestration: a biorefinery model for healthcare, energy and environment, RSC Advances 5 (2015) 73381e73394, https://doi.org/10.1039/c5ra09306f. M. D’Este, D. De Francisci, I. Angelidaki, Novel protocol for lutein extraction from microalga Chlorella vulgaris, Biochemical Engineering Journal 127 (2017) 175e179, https://doi.org/10.1016/j.bej.2017.06.019. H. Bin Li, Y. Jiang, F. Chen, Isolation and purification of lutein from the microalga Chlorella vulgaris by extraction after saponification, Journal of Agricultural and Food Chemistry 50 (2002) 1070e1072, https:// doi.org/10.1021/jf010220b. O.K. Lee, A.L. Kim, D.H. Seong, C.G. Lee, Y.T. Jung, J.W. Lee, E.Y. Lee, Chemo-enzymatic saccharification and bioethanol fermentation of lipid-extracted residual biomass of the microalga, Dunaliella tertiolecta, Bioresource Technology 132 (2013) 197e201, https:// doi.org/10.1016/j.biortech.2013.01.007. D. Hernandez, M. Solana, B. Ria~ no, M.C. GarcíaGonzalez, A. Bertucco, Biofuels from microalgae: lipid extraction and methane production from the residual biomass in a biorefinery approach, Bioresource Technology 170 (2014) 370e378, https://doi.org/10.1016/ j.biortech.2014.07.109. M. Francavilla, P. Kamaterou, S. Intini, M. Monteleone, A. Zabaniotou, Cascading microalgae

IV. Selected examples and case history

310

[61]

[62]

[63]

[64]

[65]

[66]

[67]

[68]

[69]

16. New strategies enhancing feasibility of microalgal cultivations

biorefinery: fast pyrolysis of Dunaliella tertiolecta lipid extracted-residue, Algal Research 11 (2015) 184e193, https://doi.org/10.1016/ j.algal.2015.06.017. Y. Zhu, K.O. Albrecht, D.C. Elliott, R.T. Hallen, S.B. Jones, Development of hydrothermal liquefaction and upgrading technologies for lipid-extracted algae conversion to liquid fuels, Algal Research 2 (2013) 455e464, https://doi.org/10.1016/ j.algal.2013.07.003. E.J. Kim, S. Park, H.J. Hong, Y.E. Choi, J.W. Yang, Biosorption of chromium (Cr(III)/Cr(VI)) on the residual microalga Nannochloris oculata after lipid extraction for biodiesel production, Bioresource Technology 102 (2011) 11155e11160, https://doi.org/10.1016/ j.biortech.2011.09.107. J.A. Gerde, T. Wang, L. Yao, S. Jung, L.A. Johnson, B. Lamsal, Optimizing protein isolation from defatted and non-defatted Nannochloropsis microalgae biomass, Algal Research 2 (2013) 145e153, https:// doi.org/10.1016/j.algal.2013.02.001. A. Visca, F. Di Caprio, R. Spinelli, P. Altimari, A. Cicci, G. Iaquaniello, L. Toro, F. Pagnanelli, Microalgae cultivation for lipids and carbohydrates production, Chemical Engineering Transactions 57 (2017) 127e132, https://doi.org/10.3303/CET1757022. N. Lumdubwong, P.A. Seib, Rice starch isolation by alkaline protease digestion of wet-milled rice flour, Journal of Cereal Science 31 (2000) 63e74, https:// doi.org/10.1006/jcrs.1999.0279. S. Lim, J.-H. Lee, D.-H. Shin, H.S. Lim, Comparison of protein extraction solutions for rice starch isolation and effects of residual protein content on starch pasting properties, Starch Staerke 51 (1999) 120e125, https://doi.org/10.1002/(SICI)1521-379X(199904)51: 4<120::AID-STAR120>3.3.CO;2-1. R.K. Desai, H. Monteillet, X. Li, B. Schuur, J.M. Kleijn, F.A.M. Leermakers, R.H. Wijffels, M.H.M. Eppink, One-step mild biorefinery of functional biomolecules from microalgae extracts, Reaction Chemistry and Engineering 3 (2018) 182e187, https://doi.org/10.1039/ c7re00116a. L. Soto-Sierra, P. Stoykova, Z.L. Nikolov, Extraction and fractionation of microalgae-based protein products, Algal Research 36 (2018) 175e192, https://doi.org/10.1016/j.algal.2018.10.023. L. Grossmann, S. Ebert, J. Hinrichs, J. Weiss, Effect of precipitation, lyophilization, and organic solvent extraction on preparation of protein-rich powders from the microalgae Chlorella protothecoides, Algal Research 29 (2018) 266e276, https://doi.org/ 10.1016/j.algal.2017.11.019.

[70]

[71]

[72]

[73]

[74]

[75]

[76]

[77]

[78]

[79]

W. Blanken, P.R. Postma, L. de Winter, R.H. Wijffels, M. Janssen, Predicting microalgae growth, Algal Research 14 (2016) 28e38, https://doi.org/10.1016/ j.algal.2015.12.020. F. Pagnanelli, P. Altimari, F. Trabucco, L. Toro, Mixotrophic growth of Chlorella vulgaris and Nannochloropsis oculata: interaction between glucose and nitrate, Journal of Chemical Technology and Biotechnology 89 (2014) 652e661, https://doi.org/10.1002/ jctb.4179. A. Mazzelli, A. Cicci, G. Franceschini, F.D. Caprio, G. Iaquaniello, P. Altimari, F. Pagnanelli, L. Toro, Investigation of effects of nutrients and external parameters on kinetic growth of outdoor microalgal cultivation, Chemical Engineering Transactions 64 (2018), https://doi.org/10.3303/CET1864116. S. Egloff, F. Tschudi, Z. Schmautz, D. Refardt, Highdensity cultivation of microalgae continuously fed with unfiltered water from a recirculating aquaculture system, Algal Research 34 (2018) 68e74, https:// doi.org/10.1016/j.algal.2018.07.004. J. Doucha, K. Lívanský, High density outdoor microalgal culture, in: Algal Biorefineries, vol. 1, Cultiv. Cells Prod., 2014, pp. 147e173, https://doi.org/ 10.1007/978-94-007-7494-0_6. A.C. Apel, C.E. Pfaffinger, N. Basedahl, N. Mittwollen, J. G€ obel, J. Sauter, T. Br€ uck, D. Weuster-Botz, Open thin-layer cascade reactors for saline microalgae production evaluated in a physically simulated Mediterranean summer climate, Algal Research 25 (2017) 381e390, https://doi.org/ 10.1016/j.algal.2017.06.004. I. Gifuni, A. Pollio, A. Marzocchella, G. Olivieri, New ultra-flat photobioreactor for intensive microalgal production: the effect of light irradiance, Algal Research 34 (2018) 134e142, https://doi.org/ 10.1016/j.algal.2018.07.014. C.J. Hulatt, D.N. Thomas, Energy efficiency of an outdoor microalgal photobioreactor sited at midtemperate latitude, Bioresource Technology 102 (2011) 6687e6695, https://doi.org/10.1016/ j.biortech.2011.03.098. T. Naumann, Z. Çebi, B. Podola, M. Melkonian, Growing microalgae as aquaculture feeds on twinlayers: a novel solid-state photobioreactor, Journal of Applied Phycology 25 (2013) 1413e1420, https:// doi.org/10.1007/s10811-012-9962-6. L.B. Christenson, R.C. Sims, Rotating algal biofilm reactor and spool harvester for wastewater treatment with biofuels by-products, Biotechnology and Bioengineering 109 (2012) 1674e1684, https://doi.org/ 10.2478/mspe-2018-0005.

IV. Selected examples and case history

311

References

[80] W. Blanken, M. Janssen, M. Cuaresma, Z. Libor, T. Bhaiji, R.H. Wijffels, Biofilm growth of Chlorella sorokiniana in a rotating biological contactor based photobioreactor, Biotechnology and Bioengineering 111 (2014) 2436e2445, https://doi.org/10.1002/ bit.25301. [81] P.J. Graham, B. Nguyen, S.C. Pierobon, X. Cheng, E.G. Karakolis, D. Sinton, Emerging microalgae technology: a review, Sustainable Energy and Fuels (2017), https://doi.org/10.1039/C7SE00236J. [82] B. Podola, T. Li, M. Melkonian, Porous substrate bioreactors: a paradigm shift in microalgal biotechnology? Trends in Biotechnology 35 (2017) 121e132, https://doi.org/10.1016/ j.tibtech.2016.06.004. [83] T. Li, G. Lin, B. Podola, M. Melkonian, Continuous removal of zinc from wastewater and mine dump leachate by a microalgal biofilm PSBR, Journal of Hazardous Materials 297 (2015) 112e118, https:// doi.org/10.1016/j.jhazmat.2015.04.080. [84] L. Zhang, L. Chen, J. Wang, Y. Chen, X. Gao, Z. Zhang, T. Liu, Attached cultivation for improving the biomass productivity of Spirulina platensis, Bioresource Technology 181 (2015) 136e142, https:// doi.org/10.1016/j.biortech.2015.01.025. [85] T. Liu, J. Wang, Q. Hu, P. Cheng, B. Ji, J. Liu, Y. Chen, W. Zhang, X. Chen, L. Chen, L. Gao, C. Ji, H. Wang, Attached cultivation technology of microalgae for efficient biomass feedstock production, Bioresource Technology 127 (2013) 216e222, https://doi.org/ 10.1016/j.biortech.2012.09.100. [86] L.K.P. Schultze, M.V. Simon, T. Li, D. Langenbach, B. Podola, M. Melkonian, High light and carbon dioxide optimize surface productivity in a twin-layer biofilm photobioreactor, Algal Research 8 (2015) 37e44, https://doi.org/10.1016/j.algal.2015.01.007. [87] A. Janoska, P.P. Lamers, A. Hamhuis, Y. van Eimeren, R.H. Wijffels, M. Janssen, A liquid foam-bed photobioreactor for microalgae production, Chemical Engineering Journal 313 (2017) 1206e1214, https:// doi.org/10.1016/j.cej.2016.11.022. [88] A. Janoska, M. Vazquez, M. Janssen, R.H. Wijffels, M. Cuaresma, C. Vílchez, Surfactant selection for a liquid foam-bed photobioreactor, Biotechnology Progress 34 (2018) 711e720, https://doi.org/ 10.1002/btpr.2614. [89] A. Janoska, R. Barten, S. de Nooy, P. van Rijssel, R.H. Wijffels, M. Janssen, Improved liquid foam-bed photobioreactor design for microalgae cultivation, Algal Research 33 (2018) 55e70, https://doi.org/ 10.1016/j.algal.2018.04.025.

[90]

[91]

[92]

[93]

[94]

[95]

[96]

[97]

E.G. Nwoba, D.A. Parlevliet, D.W. Laird, K. Alameh, N.R. Moheimani, Sustainable phycocyanin production from Arthrospira platensis using solar-control thin film coated photobioreactor, Biochemical Engineering Journal 141 (2019) 232e238, https:// doi.org/10.1016/j.bej.2018.10.024. E.E. Jung, A. Jain, N. Voulis, D.F.R. Doud, L.T. Angenent, D. Erickson, Stacked optical waveguide photobioreactor for high density algal cultures, Bioresource Technology 171 (2014) 495e499, https://doi.org/10.1016/ j.biortech.2014.08.093. Y. Sun, Q. Liao, Y. Huang, A. Xia, Q. Fu, X. Zhu, Y. Zheng, Integrating planar waveguides doped with light scattering nanoparticles into a flat-plate photobioreactor to improve light distribution and microalgae growth, Bioresource Technology 220 (2016) 215e224, https://doi.org/10.1016/ j.biortech.2016.08.063. S. Karuppasamy, A.S. Musale, B. Soni, B. Bhadra, N. Gujarathi, M. Sundaram, A. Sapre, S. Dasgupta, C. Kumar, Integrated grazer management mediated by chemicals for sustainable cultivation of algae in open ponds, Algal Research 35 (2018) 439e448, https://doi.org/10.1016/j.algal.2018.09.017. R. Singh, R. Birru, G. Sibi, Nutrient removal efficiencies of Chlorella vulgaris from the urban wastewater for reduced eutrofication, Journal of Environmental Protection 8 (2017) 1e11, https:// doi.org/10.4236/jep.2017.81001. M.P. Caporgno, A. Taleb, M. Olkiewicz, J. Font, J. Pruvost, J. Legrand, C. Bengoa, Microalgae cultivation in urban wastewater: nutrient removal and biomass production for biodiesel and methane, Algal Research 10 (2015) 232e239, https://doi.org/ 10.1016/j.algal.2015.05.011. K. Katam, D. Bhattacharyya, Comparative study on treatment of kitchen wastewater using a mixed microalgal culture and an aerobic bacterial culture: kinetic evaluation and FAME analysis, Environmental Science and Pollution Research 25 (2018) 20732e20742, https://doi.org/10.1007/s11356-0182209-6. C. Gonzalez-Fernandez, B. Ria~ no-Irazabal, B. Molinuevo-Salces, S. Blanco, M.C. GarcíaGonzalez, Effect of operational conditions on the degradation of organic matter and development of microalgae-bacteria consortia when treating swine slurry, Applied Microbiology and Biotechnology 90 (2011) 1147e1153, https://doi.org/10.1007/s00253011-3111-z.

IV. Selected examples and case history

312

16. New strategies enhancing feasibility of microalgal cultivations

[98] X. Han, N. Rusconi, P. Ali, K. Pagkatipunan, F. Chen, Nutrients extracted from chicken manure accelerate growth of microalga Scenedesmus obliquus HTB1, Green and Sustainable Chemistry 7 (2017) 101e113, https://doi.org/10.4236/gsc.2017.72009. [99] M. Franchino, E. Comino, F. Bona, V.A. Riggio, Chemosphere growth of three microalgae strains and nutrient removal from an agro-zootechnical digestate, Chemosphere 92 (2013) 738e744, https:// doi.org/10.1016/j.chemosphere.2013.04.023. [100] A. Ruiz-Martinez, N. Martin Garcia, I. Romero, A. Seco, J. Ferrer, Microalgae cultivation in wastewater: nutrient removal from anaerobic membrane bioreactor effluent, Bioresource Technology 126 (2012) 247e253, https://doi.org/10.1016/j.biortech. 2012.09.022. [101] A.S. Vikneswara, R.M.S. Radin Mohamed, A.A.S. AlGheethi, A.H. Mohd Kassim, N. Othman, Removal of Nutrients from Meat Processing Wastewater Through the Phycoremediation Process. Management of Greywater in Developing Countries, Springer International Publishing, 2019, pp. 245e263, https://doi.org/ 10.1007/978-3-319-90269-2_13. [102] J.M. Girard, R. Tremblay, N. Faucheux, M. Heitz, J.S. Desch^enes, Phycoremediation of cheese whey permeate using directed commensalism between Scenedesmus obliquus and Chlorella protothecoides, Algal Research 22 (2017) 122e126, https://doi.org/ 10.1016/j.algal.2016.12.013. [103] J.M. Girard, M.L. Roy, M. Ben Hafsa, J. Gagnon, N. Faucheux, M. Heitz, R. Tremblay, J.S. Desch^enes, Mixotrophic cultivation of green microalgae Scenedesmus obliquus on cheese whey permeate for biodiesel production, Algal Research 5 (2014) 241e248, https://doi.org/10.1016/j.algal.2014.03.002. [104] F. Di Caprio, P. Altimari, G. Iaquaniello, L. Toro, F. Pagnanelli, T. Obliquus mixotrophic cultivation in treated and untreated olive mill wastewater, Chemical Engineering Transactions 64 (2018), https:// doi.org/10.3303/CET1864105. [105] F. Di Caprio, P. Altimari, F. Pagnanelli, Integrated microalgae biomass production and olive mill wastewater biodegradation: optimization of the wastewater supply strategy, Chemical Engineering Journal 349 (2018) 539e546, https://doi.org/10.1016/ j.cej.2018.05.084. [106] C.J. de Andrade, L.M. de Andrade, Microalgae for bioremediation of textile wastewater: an overview, MOJ Food Processing and Technology 6 (2018), https://doi.org/10.15406/mojfpt.2018.06.00200. [107] L. Xin, H. Hong-ying, G. Ke, S. Ying-xue, Effects of different nitrogen and phosphorus concentrations on the growth, nutrient uptake, and lipid

[108]

[109]

[110]

[111]

[112]

[113]

[114]

[115]

[116]

[117]

[118]

accumulation of a freshwater microalga Scenedesmus sp, Bioresource Technology 101 (2010) 5494e5500, https://doi.org/10.1016/j.biortech. 2010.02.016. R.W. Krauss, W.H. Thomas, The growth and inorganic nutrition of Scenedesmus obliquus in mass culture. 123, Plant Physiology 29 (1954) 205e214, https://doi.org/10.1104/pp.29.3.205. R.W. Sterner, J.J. Elser, Stoichiometry and homeostasis, Ecological Stoichiometry (2002) 1e43, https://doi.org/10.1073/pnas.0703993104. F. Di Caprio, P. Scarponi, P. Altimari, G. Iaquaniello, F. Pagnanelli, The influence of phenols extracted from olive mill wastewater on the heterotrophic and mixotrophic growth of Scenedesmus sp, Journal of Chemical Technology and Biotechnology 93 (2018) 3619e3626, https://doi.org/10.1002/ jctb.5743. D.L. Cheng, H.H. Ngo, W.S. Guo, S.W. Chang, D.D. Nguyen, S.M. Kumar, Microalgae biomass from swine wastewater and its conversion to bioenergy, Bioresource Technology 275 (2019) 109e122, https:// doi.org/10.1016/j.biortech.2018.12.019. T. Cai, S.Y. Park, Y. Li, Nutrient recovery from wastewater streams by microalgae: status and prospects, Renewable and Sustainable Energy Reviews 19 (2013) 360e369, https://doi.org/10.1016/ j.rser.2012.11.030. O. Bernard, Hurdles and challenges for modelling and control of microalgae for CO2 mitigation and biofuel production, Journal of Process Control 21 (2011) 1378e1389, https://doi.org/10.1016/j.jprocont. 2011.07.012. F. Di Caprio, P. Altimari, F. Pagnanelli, Effect of Ca2þ concentration on Scenedesmus sp. growth in heterotrophic and photoautotrophic cultivation, Nature Biotechnology 40 B (2017) 228e235, https:// doi.org/10.1016/j.nbt.2017.09.003. T. K€allqvist, A. Svenson, Assessment of ammonia toxicity in tests with the microalga, Nephroselmis pyriformis, Chlorophyta, Water Research 37 (2003) 477e484, https://doi.org/10.1016/S0043-1354(02)00361-5. A. Xia, J.D. Murphy, Microalgal cultivation in treating liquid digestate from biogas systems, Trends in Biotechnology 34 (2016) 264e275, https://doi.org/ 10.1016/j.tibtech.2015.12.010. G. Hodaifa, M.E. Martínez, S. Sanchez, Influence of temperature on growth of Scenedesmus obliquus in diluted olive mill wastewater as culture medium, Engineering in Life Science 10 (2010) 257e264, https:// doi.org/10.1002/elsc.201000005. F. Di Caprio, P. Altimari, F. Pagnanelli, Integrated biomass production and biodegradation of olive mill

IV. Selected examples and case history

References

[119]

[120]

[121]

[122]

[123]

[124]

[125]

[126]

[127]

wastewater by cultivation of Scenedesmus sp, Algal Research 9 (2015) 306e311. G. Markou, I. Chatzipavlidis, D. Georgakakis, Cultivation of Arthrospira (Spirulina) platensis in oliveoil mill wastewater treated with sodium hypochlorite, Bioresource Technology 112 (2012) 234e241, https://doi.org/10.1016/ j.biortech.2012.02.098.  G. Hodaifa, S. Sanchez, M.E. Martínez, R. Orpez, Biomass production of Scenedesmus obliquus from mixtures of urban and olive-oil mill wastewaters used as culture medium, Applied Energy 104 (2013) 345e352, https://doi.org/10.1016/j.apenergy.2012.11.005. B. Farizoglu, B. Keskinler, E. Yildiz, A. Nuhoglu, Cheese whey treatment performance of an aerobic jet loop membrane bioreactor, Process Biochemistry 39 (2004) 2283e2291, https://doi.org/10.1016/ j.procbio.2003.11.028. Y. Zhang, A. Kendall, J. Yuan, A comparison of on-site nutrient and energy recycling technologies in algal oil production, Resources, Conservation and Recycling 88 (2014) 13e20, https://doi.org/10.1016/ j.resconrec.2014.04.011. F. Di Caprio, P. Altimari, D. Uccelletti, F. Pagnanelli, Mechanistic modelling of copper biosorption by wild type and engineered Saccharomyces cerevisiae biomasses, Chemical Engineering Journal 244 (2014) 561e568. G. Markou, L. Wang, J. Ye, A. Unc, Using agroindustrial wastes for the cultivation of microalgae and duckweeds: contamination risks and biomass safety concerns, Biotechnology Advances 36 (2018) 1238e1254, https://doi.org/10.1016/ j.biotechadv.2018.04.003. F. Di Caprio, P. Altimari, E. Zanni, D. Uccelletti, L. Toro, F. Pagnanelli, Lanthanum biosorption by different saccharomyces cerevisiae strains, Chemical Engineering Transactions 49 (2016) 37e42, https:// doi.org/10.3303/CET1649007. F.Z. Mennaa, Z. Arbib, J.A. Perales, Urban wastewater treatment by seven species of microalgae and analgal bloom: biomass production, N and P removal kinetics and harvestability, Water Research 83 (2015) 42e51, https://doi.org/10.1016/ j.watres.2015.06.007. S.Y. Chiu, C.Y. Kao, T.Y. Chen, Y. Bin Chang, C.M. Kuo, C.S. Lin, Cultivation of microalgal Chlorella for biomass and lipid production using wastewater as nutrient resource, Bioresource Technology 184 (2015) 179e189, https://doi.org/10.1016/ j.biortech.2014.11.080.

313

[128] J.C.M. Pires, M.C.M. Alvim-Ferraz, F.G. Martins, M. Sim~ oes, Wastewater treatment to enhance the economic viability of microalgae culture, Environmental Science and Pollution Research 20 (2013) 5096e5105, https://doi.org/10.1007/s11356-013-1791-x. [129] P. Foladori, S. Petrini, G. Andreottola, Evolution of real municipal wastewater treatment in photobioreactors and microalgae-bacteria consortia using real-time parameters, Chemical Engineering Journal 345 (2018) 507e516, https://doi.org/10.1016/j.cej.2018.03.178. [130] A.L. Gonçalves, J.C.M. Pires, M. Sim~ oes, A review on the use of microalgal consortia for wastewater treatment, Algal Research 24 (2017) 403e415, https://doi.org/10.1016/j.algal.2016.11.008. [131] M.H. Huang, Y.M. Li, G.W. Gu, Chemical composition of organic matters in domestic wastewater, Desalination 262 (2010) 36e42, https://doi.org/10.1016/ j.desal.2010.05.037. [132] L. Peng, H.H. Ngo, W.S. Guo, Y. Liu, D. Wang, S. Song, W. Wei, L.D. Nghiem, B.J. Ni, A novel mechanistic model for nitrogen removal in algal-bacterial photo sequencing batch reactors, Bioresource Technology 267 (2018) 502e509, https://doi.org/ 10.1016/j.biortech.2018.07.093. [133] P. Foladori, S. Petrini, M. Nessenzia, G. Andreottola, Enhanced nitrogen removal and energy saving in a microalgal-bacterial consortium treating real municipal wastewater, Water Science and Technology 78 (2018) 174e182, https://doi.org/10.2166/ wst.2018.094. [134] J. Ye, J. Liang, L. Wang, G. Markou, Q. Jia, Operation optimization of a photo-sequencing batch reactor for wastewater treatment: study on influencing factors and impact on symbiotic microbial ecology, Bioresource Technology 252 (2018) 7e13, https:// doi.org/10.1016/j.biortech.2017.12.086. [135] M.A. Alam, C. Wan, X.Q. Zhao, L.J. Chen, J.S. Chang, F.W. Bai, Enhanced removal of Zn2þ or Cd2þ by the flocculating Chlorella vulgaris JSC-7, Journal of Hazardous Materials 289 (2015) 38e45, https://doi.org/ 10.1016/j.jhazmat.2015.02.012. [136] O. Perez-Garcia, F.M.E. Escalante, L.E. de-Bashan, Y. Bashan, Heterotrophic cultures of microalgae: metabolism and potential products, Water Research 45 (2011) 11e36, https://doi.org/10.1016/ j.watres.2010.08.037. [137] S. Wang, Y. Wu, X. Wang, Heterotrophic cultivation of Chlorella pyrenoidosa using sucrose as the sole carbon source by co-culture with Rhodotorula glutinis, Bioresource Technology 220 (2016) 615e620, https://doi.org/10.1016/j.biortech.2016.09.010.

IV. Selected examples and case history

314

16. New strategies enhancing feasibility of microalgal cultivations

[138] S. Ethier, K. Woisard, D. Vaughan, Z. Wen, Continuous culture of the microalgae Schizochytrium limacinum on biodiesel-derived crude glycerol for producing docosahexaenoic acid, Bioresource Technology 102 (2011) 88e93, https://doi.org/10.1016/ j.biortech.2010.05.021. [139] C. Baroukh, V. Turon, O. Bernard, Dynamic metabolic modeling of heterotrophic and mixotrophic microalgal growth on fermentative wastes, PLoS Computational Biology (2017) 1e18, https://doi.org/ 10.1371/journal.pcbi.1005590. [140] J.S. Desch^enes, A bacteriostatic control approach for mixotrophic cultures of microalgae, IFAC-PapersOnLine 49 (2016) 1074e1078, https:// doi.org/10.1016/j.ifacol.2016.07.345. [141] P. Bohutskyi, K. Liu, L.K. Nasr, N. Byers, J.N. Rosenberg, G.A. Oyler, M.J. Betenbaugh, E.J. Bouwer, Bioprospecting of microalgae for integrated biomass production and phytoremediation of unsterilized wastewater and anaerobic digestion centrate, Applied Microbiology and Biotechnology 99 (2015) 6139e6154, https://doi.org/10.1007/ s00253-015-6603-4. [142] G. Mujtaba, K. Lee, Treatment of real wastewater using co-culture of immobilized Chlorella vulgaris and suspended activated sludge, Water Research 120 (2017) 174e184, https://doi.org/10.1016/ j.watres.2017.04.078. [143] M.R. Droop, 25 Years of algal growth kinetics, Botanica Marina 26 (1983) 99e112, https://doi.org/ 10.1515/botm.1983.26.3.99. [144] J.S. Desch^enes, A. Boudreau, R. Tremblay, Mixotrophic production of microalgae in pilot-scale photobioreactors: practicability and process considerations, Algal Research 10 (2015) 80e86, https://doi.org/10.1016/j.algal.2015.04.015. [145] G.S. Khush, Green revolution: the way forward, Nature Reviews Genetics 2 (2001) 815e822, https:// doi.org/10.1038/35093585. [146] H.H. Iltis, From teosinte to maize: the catastrophic sexual transmutation, Science 222 (1983) 886e894, https://doi.org/10.1126/science.222.4626.886. [147] P.G. Stephenson, C.M. Moore, M.J. Terry, M.V. Zubkov, T.S. Bibby, Improving photosynthesis for algal biofuels: toward a green revolution, Trends in Biotechnology 29 (2011) 615e623, https:// doi.org/10.1016/j.tibtech.2011.06.005. [148] J.E.W. Polle, S. Kanakagiri, E.S. Jin, T. Masuda, A. Melis, Truncated chlorophyll antenna size of the photosystems e a practical method to improve microalgal productivity and hydrogen production in mass culture, in: Int. J. Hydrogen Energy, 2002,

[149]

[150]

[151]

[152]

[153]

[154]

[155]

[156]

[157]

pp. 1257e1264, https://doi.org/10.1016/S03603199(02)00116-7. P. Altimari, F. Di Caprio, L. Toro, A.L. Capriotti, F. Pagnanelli, Hydrogen photo-production by mixotrophic cultivation of Chlamydomonas reinhardtii: interaction between organic carbon and nitrogen, Chemical Engineering Transactions 38 (2014) 199e204, https://doi.org/10.3303/CET1438034. S.N. Kosourov, M.L. Ghirardi, M. Seibert, A truncated antenna mutant of Chlamydomonas reinhardtii can produce more hydrogen than the parental strain, International Journal of Hydrogen Energy 36 (2011) 2044e2048, https://doi.org/10.1016/ j.ijhydene.2010.10.041. A. Melis, Solar energy conversion efficiencies in photosynthesis: minimizing the chlorophyll antennae to maximize efficiency, Plant Science 177 (2009) 272e280, https://doi.org/10.1016/ j.plantsci.2009.06.005. R. Radakovits, R.E. Jinkerson, A. Darzins, M.C. Posewitz, Genetic engineering of algae for enhanced biofuel production, Eukaryotic Cell 9 (2010) 486e501, https://doi.org/10.1128/EC.00364-09. G. Breuer, L. de Jaeger, R.E. Verbeek, R.B. Draaisma, D.E. Martens, J. Springer, G. Eggink, R.H. Wijffels, Superior triacylglycerol (TAG) accumulation in starchless mutants of Scenedesmus obliquus: (I) mutant generation and characterization, Biotechnology for Biofuels 7 (2014) 69, https://doi.org/10.1186/17546834-7-69. Y.H. Ma, X. Wang, Y.F. Niu, Z.K. Yang, M.H. Zhang, Z.M. Wang, W.D. Yang, J.S. Liu, H.Y. Li, Antisense knockdown of pyruvate dehydrogenase kinase promotes the neutral lipid accumulation in the diatom Phaeodactylum tricornutum, Microbial Cell Factories 13 (2014), https://doi.org/10.1186/s12934-014-0100-9. J. Xue, Y.F. Niu, T. Huang, W.D. Yang, J.S. Liu, H.Y. Li, Genetic improvement of the microalga Phaeodactylum tricornutum for boosting neutral lipid accumulation, Metabolic Engineering 27 (2015) 1e9, https://doi.org/10.1016/j.ymben.2014.10.002. K.M. Koo, S. Jung, B.S. Lee, J.B. Kim, Y.D. Jo, H. Il Choi, S.Y. Kang, G.H. Chung, W.J. Jeong, J.W. Ahn, The mechanism of starch over-accumulation in Chlamydomonas reinhardtii high-starch mutants identified by comparative transcriptome analysis, Frontiers in Microbiology 8 (2017), https://doi.org/10.3389/ fmicb.2017.00858. M. Schulz-Raffelt, V. Chochois, P. Auroy, S. Cuine, E. Billon, D. Dauvillee, Y. Li-Beisson, G. Peltier, Hyper-accumulation of starch and oil in a Chlamydomonas mutant affected in a plant-specific DYRK kinase,

IV. Selected examples and case history

315

References

[158]

[159]

[160]

[161]

[162]

[163]

[164]

[165]

Biotechnology for Biofuels 9 (2016), https://doi.org/ 10.1186/s13068-016-0469-2. B.F. Cordero, I. Couso, R. Le on, H. Rodríguez,  Vargas, Enhancement of carotenoids biosynM.A. thesis in Chlamydomonas reinhardtii by nuclear transformation using a phytoene synthase gene isolated from Chlorella zofingiensis, Applied Microbiology and Biotechnology 91 (2011) 341e351, https://doi.org/ 10.1007/s00253-011-3262-y. P. Perez-L opez, J.H. de Vree, G. Feijoo, R. Bosma, M.J. Barbosa, M.T. Moreira, R.H. Wijffels, A.J.B. van Boxtel, D.M.M. Kleinegris, Comparative life cycle assessment of real pilot reactors for microalgae cultivation in different seasons, Applied Energy 205 (2017) 1151e1164, https://doi.org/10.1016/ j.apenergy.2017.08.102. T.M. Lee, Y.F. Tseng, C.L. Cheng, Y.C. Chen, C.S. Lin, H.Y. Su, T.J. Chow, C.Y. Chen, J.S. Chang, Characterization of a heat-tolerant Chlorella sp. GD mutant with enhanced photosynthetic CO2 fixation efficiency and its implication as lactic acid fermentation feedstock, Biotechnology for Biofuels 10 (2017), https://doi.org/10.1186/s13068-017-0905-y. P. Mehta, R. Rani, R. Gupta, A.S. Mathur, S.K. Puri, Biomass and lipid production of a novel freshwater thermo-tolerant mutant strain of Chlorella pyrenoidosa NCIM 2738 in seawater salinity recycled medium, Algal Research 36 (2018) 88e95, https://doi.org/ 10.1016/j.algal.2018.10.015. J. Guan, S. Shen, H. Wu, X. Liu, W. Shen, Y. He, R. Duan, Biomass and terpenoids produced by mutant strains of Arthrospira under low temperature and light conditions, World Journal of Microbiology and Biotechnology 33 (2017), https://doi.org/ 10.1007/s11274-016-2199-9. M. Kwak, W.K. Park, S.E. Shin, H.G. Koh, B. Lee, B. ryool Jeong, Y.K. Chang, Improvement of biomass and lipid yield under stress conditions by using diploid strains of Chlamydomonas reinhardtii, Algal Research 26 (2017) 180e189, https://doi.org/ 10.1016/j.algal.2017.07.027. S.O. Kotchoni, E.W. Gachomo, K. Slobodenko, D.H. Shain, AMP deaminase suppression increases biomass, cold tolerance and oil content in green algae, Algal Research 16 (2016) 473e480, https:// doi.org/10.1016/j.algal.2016.04.007. H.Y. Su, H.H. Chou, T.J. Chow, T.M. Lee, J.S. Chang, W.L. Huang, H.J. Chen, Improvement of outdoor culture efficiency of cyanobacteria by over-expression of stress tolerance genes and its implication as biorefinery feedstock, Bioresource Technology 244

[166]

[167]

[168]

[169]

[170]

[171]

[172]

[173]

[174]

(2017) 1294e1303, https://doi.org/10.1016/ j.biortech.2017.04.074. D. Tang, W. Han, P. Li, X. Miao, J. Zhong, CO2 biofixation and fatty acid composition of Scenedesmus obliquus and Chlorella pyrenoidosa in response to different CO2 levels, Bioresource Technology 102 (2011) 3071e3076, https://doi.org/10.1016/ j.biortech.2010.10.047. D. Li, L. Wang, Q. Zhao, W. Wei, Y. Sun, Improving high carbon dioxide tolerance and carbon dioxide fixation capability of Chlorella sp. by adaptive laboratory evolution, Bioresource Technology 185 (2015) 269e275, https://doi.org/10.1016/ j.biortech.2015.03.011. P. Varshney, S. Sohoni, P.P. Wangikar, J. Beardall, Effect of high CO2 concentrations on the growth and macromolecular composition of a heat- and highlight-tolerant microalga, Journal of Applied Phycology 28 (2016) 2631e2640, https://doi.org/ 10.1007/s10811-016-0797-4. S. Arata, C. Strazza, A. Lodi, A. Del Borghi, Spirulina platensis culture with flue gas feeding as a cyanobacteria-based carbon sequestration option, Chemical Engineering and Technology 36 (2013) 91e97, https://doi.org/10.1002/ceat.201100722. D. Cheng, X. Li, Y. Yuan, C. Yang, T. Tang, Q. Zhao, Y. Sun, Adaptive evolution and carbon dioxide fixation of Chlorella sp. in simulated flue gas, The Science of the Total Environment 650 (2019) 2931e2938, https://doi.org/10.1016/j.scitotenv.2018.10.070. L. Zhou, Y. Yuan, X. Li, S. Mei, J. Gao, Q. Zhao, W. Wei, Y. Sun, Exploration of phenol tolerance mechanism through antioxidative responses of an evolved strain, Chlorella sp. L5, Journal of Applied Phycology 30 (2018) 2379e2385, https://doi.org/10.1007/ s10811-018-1428-z. A.A. Corcoran, M.A. Saunders, A.P. Hanley, P.A. Lee, S. Lopez, R. Ryan, C.B. Yohn, Iterative screening of an evolutionary engineered Desmodesmus generates robust field strains with pesticide tolerance, Algal Research 31 (2018) 443e453, https://doi.org/ 10.1016/j.algal.2018.02.026. A. Ibuot, A.P. Dean, O.A. McIntosh, J.K. Pittman, Metal bioremediation by CrMTP4 over-expressing Chlamydomonas reinhardtii in comparison to natural wastewater-tolerant microalgae strains, Algal Research 24 (2017) 89e96, https://doi.org/10.1016/ j.algal.2017.03.002. M.C. Ruiz-Domínguez, I. Vaquero, V. Obreg on, B. de la Morena, C. Vílchez, J.M. Vega, Lipid accumulation and antioxidant activity in the eukaryotic acidophilic

IV. Selected examples and case history

316

16. New strategies enhancing feasibility of microalgal cultivations

microalga Coccomyxa sp. (strain onubensis) under nutrient starvation, Journal of Applied Phycology 27 (2015) 1099e1108, https://doi.org/10.1007/s10811014-0403-6. [175] V. L opez-Rodas, F. Marva, E. Costas, A. Flores-Moya, Microalgal adaptation to a stressful environment (acidic, metal-rich mine waters) could be due to selection of pre-selective mutants originating in non-extreme environments, Environmental and Experimental Botany 64 (2008) 43e48, https:// doi.org/10.1016/j.envexpbot.2008.01.001.

[176] N. von Alvensleben, M. Magnusson, K. Heimann, Salinity tolerance of four freshwater microalgal species and the effects of salinity and nutrient limitation on biochemical profiles, Journal of Applied Phycology 28 (2016) 861e876, https://doi.org/10.1007/s10811015-0666-6. [177] F.D. Caprio, P. Altimari, G. Iaquaniello, L. Toro, F. Pagnanelli, Heterotrophic cultivation of T. obliquus under non-axenic conditions by uncoupled supply of nitrogen and glucose, Biochemical Engineering Journal 145 (2019) 127e136, https://doi.org/ 10.1016/j.bej.2019.02.020.

IV. Selected examples and case history