Abstract
Chlorella represents a group of eukaryotic green microalgae that has been receiving increasing scientific and commercial interest. It possesses high photosynthetic ability and is capable of growing robustly under mixotrophic and heterotrophic conditions as well. Chlorella has long been considered as a source of protein and is now industrially produced for human food and animal feed. Chlorella is also rich in oil, an ideal feedstock for biofuels. The exploration of biofuel production by Chlorella is underway. Chlorella has the ability to fix carbon dioxide efficiently and to remove nutrients of nitrogen and phosphorous, making it a good candidate for greenhouse gas biomitigation and wastewater bioremediation. In addition, Chlorella shows potential as an alternative expression host for recombinant protein production, though challenges remain to be addressed. Currently, omics analyses of certain Chlorella strains are being performed, which will help to unravel the biological implications of Chlorella and facilitate the future exploration of industrial applications.
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1 Introduction
Chlorella is a group of eukaryotic green microalgae with high photosynthesis ability. Through efficient photosynthesis, Chlorella is able to reproduce itself within several hours, requiring only sunlight, carbon dioxide, water, and a small amount of nutrients. Chlorella is easy to grow, has a simple life cycle and metabolic pathways similar to higher plants, and thus has long been employed as a model organism to investigate the mechanisms of photosynthesis and carbon dioxide assimilation (Calvin–Benson cycle). Due to its high protein content and richness in carotenoids, vitamins, and minerals, Chlorella has also long been proposed as a food substitute for humans and is now widely produced as health food in Germany, China, Japan, and several other Asian countries. Recently, because of the energy crisis and public interest in green renewable fuels, Chlorella has been cited as a promising candidate feedstock for biofuel production and has gained increasing scientific and industrial attention in that it grows fast, has high oil content, and outdoor mass cultures are easy to maintain. This chapter gives an overview of Chlorella, the industrially important microalgal genus, covering its taxonomy, growth physiology, cellular chemical composition, mass cultivation, and potential products and applications. An integrated production of biofuels and other bioproducts coupled with the treatment of greenhouse gas and wastewater is proposed, which may offset the Chlorella-based production cost while providing significant environmental benefits.
2 Morphology, Ultrastructure, and Taxonomy
Chlorella is a genus of unicellular and nonmobile green algae. Chlorella vulgaris is the type species of this genus, which was first described by M.W. Beijerinck in 1890. Commonly, Chlorella cells are spherical or ellipsoidal and the cell size may range from 2 to 15 μm in diameter. They are widely distributed in diverse habits such as freshwater, marine water, soil, and are even symbiotic with lichens and protozoa. Chlorella has no sexual life cycle and reproduces itself through asexual autospore production. When mature, autospores are simultaneously released via rupture of the mother cell wall. The number of autospores derived from a single mother cell may vary greatly from 2 to 16.
The ultrastructure of Chlorella has been studied extensively in past years. Under transmission electron microscopy, the visible structures within the cell include the chloroplast, nucleus, mitochondria, vacuole, starch, lipid bodies, and so on [1–3]. Generally, Chlorella has a cup-shaped chloroplast located peripherally in the cytoplasm. The nucleus situates near the cytoplasmic membrane, and the mitochondria are closely associated with the chloroplast. Pyrenoid, a conspicuous and easily recognizable structure, is present in most of the Chlorella species [2–4]. Usually, the pyrenoid is centrally located in the chloroplast and surrounded by the starch sheath [3]. In some Chlorella species, the pyrenoid contains many lipid-containing globules that are known as pyrenoglobuli and may function as secondary storage products [3, 4]. Chlorella has a thick and rigid cell wall, but the cell-wall structure may differ greatly across the species [2, 5]. When transferred to stress conditions (e.g., nitrogen starvation), the cell wall thickens and the chloroplast begins to regress to the proplastid stage with a gradual reduction in thylakoid number, accompanied by the accumulation of lipid bodies in the cytoplasm [3].
To date, there are hundreds of Chlorella strains reported in the literature, but the classification of Chlorella has been problematic due to the lack of conspicuous morphological characters. Kessler [6] proposed a sound taxonomic method for Chlorella based on multiple biochemical and physiological characters, that is, hydrogenase, secondary carotenoids, acid or salt tolerance, lactic acid fermentation, nitrate reduction, thiamine requirement, and the GC content of DNA. By comparing these characters, 77 Chlorella strains from the Collection of Algae at Göttingen (SAG, Germany) were assigned to 12 taxa and Chlorella was suggested to be an assembly of morphologically similar species of polyphyletic origin. Afterwards Kessler and Huss [7] examined 58 Chlorella strains from the Culture Collection of Algae at the University of Texas (UTEX, USA) according to the above-mentioned biochemical and physiological characters and reassigned them to 10 well-established species. The sugar composition of the cell wall (either glucosamine or glucose and mannose) has also been used for Chlorella classification [8, 9]. In addition, Huss et al. [10] examined the Chlorella genus by using a phylogenic approach based on complete 18S rRNA sequences, and considered it as a polyphyletic assemblage dispersed over two classes of Chlorophyta, that is, Chlorophyceae and Trebouxiophyceae; only four species were suggested to belong to this genus: Chlorella vulgaris, Chlorella sorokiniana, Chlorella kessleri, and Chlorella lobophora. Recently, based on the sequences of 18S rRNA and ITS2 region, Krienitz et al. [11] further investigated the phylogenesis of Chlorella and suggested the exclusion of Chlorella kessleri from the Chlorella genus. In this chapter, we regard Chlorella to be the Chlorella sensu lato and include the data of Chlorella species that may have been excluded from the genus by the above-mentioned studies.
3 Growth Physiology
Chlorella is able to convert solar light energy to chemical energy through efficient photosynthesis. Similar to C3 higher plants, photosynthesis in Chlorella consists of light-dependent reactions and carbon dioxide fixation. The light-dependent reactions produce high-energy molecules ATP and NADPH, which are utilized in the Calvin–Benson cycle to fix carbon dioxide. Chlorella performs photosynthesis efficiently at a relatively low light intensity and becomes saturated when the light intensity reaches a certain value, which may range from 80 to 400 µE m−2s−1on a per cell basis [12, 13]. Higher light intensity above the saturation value may inhibit the photosynthesis of the algae and even cause the destruction of chlorophylls (photobleaching) and cell deaths. The incident intensities of solar light can reach up to 2,500 µE m−2s−1, much higher than that required for photosynthesis saturation. Most of the sunlight energy is lost as heat and only a small portion can be converted to chemical energy by photosynthesis. Although the theoretical maximum solar energy conversion efficiency of oxygenic photosynthesis is thought to be around 8–10 % [14], the outdoor culture of Chlorella can only achieve a low photosynthetic efficiency (PE), for example, 2.7 % in full sunlight [15]. The yet to be enhanced PE remains a big challenge for outdoor mass cultures of algae and presents a promising direction toward the increase of biomass production.
Chlorella growth requires nutrients including carbon, nitrogen, phosphorus, sulfur, and metals. Carbon is the predominant element of Chlorella and carbon dioxide is the primary carbon source for photoautotrophic growth of Chlorella. Chlorella utilizes carbon dioxide principally in the undissociated form of CO2 or H2CO3 [16]. The atmospheric air contains only 0.04 % CO2, which is not sufficient to maintain rapid growth of Chlorella for high cell density. Therefore, a supply of air enriched with CO2 at the concentration of 1–5 % is usually provided to Chlorella cultures [17, 18]. Higher levels of CO2 may cause a decrease in pH of the medium and thus inhibit or even block the algal growth [19, 20]. Nevertheless, the high-CO2–tolerant Chlorella species as reported by Papazi et al. [21] and Sakai et al. [22] can grow well in the presence of up to 40 % CO2, although the optimal growth is obtained under lower CO2 concentrations. Nitrogen is the second most important element in Chlorella. Generally, Chlorella is able to utilize nitrate, ammonia, and organic sources of nitrogen such as urea, glycine, and amino acids [17, 23, 24]. Both nitrate-N and urea-N cannot be directly incorporated into organic compounds by Chlorella and first have to be reduced to ammonia-N. Ammonia and urea are economically more favorable than nitrate as nitrogen sources in that the latter is more expensive per unit N. The uptake of ammonia may result in acidification of the medium, nitrate may cause alkalinization, whereas urea leads to only minor pH changes [25]. In this context, urea is the better choice of nitrogen source, avoiding a large pH shift of unbuffered medium. Different Chlorella species may favor different nitrogen sources for growth, for example, Chlorella pyrenoidosa prefers urea to nitrate or glycine for boosting biomass production and Chlorella protothecoides gives higher biomass yield when using nitrate rather than urea as the nitrogen source [23, 26]. Nitrogen concentration in the culture medium plays an important role in regulating algal growth and metabolism. Nitrogen deficiency/starvation retards the growth of Chlorella, causes the decrease of protein levels, and promotes the accumulation of lipids within cells (Illman et al. [27]; Ördög et al. [28, 29]). Phosphorus is the third essential nutrient required for normal growth of Chlorella. It is involved in the formation of nucleic acid and cell membrane, as well as of ATP that provides energy for cellular metabolism. The most used phosphorus source for algal cultivation is phosphate, either as H2PO4 1− or HPO4 2−. Sulfur is an indispensable constituent of some essential amino acids, vitamins, and sulfolipids. Usually, it is provided in the form of sulfate for algal growth. Other inorganic nutrients include K, Ca, Mg, Fe, Cu, Zn, Mn, and Mo, among others, needed in trace amounts.
In addition to photoautotrophy, Chlorella is able to utilize organic carbon sources alone or together with CO2 and light for heterotrophic or mixotrophic growth (Table 1). Sugars are the most conventional organic carbon sources widely used for Chlorella fermentation [30–32]. Liu et al. [30] surveyed the utilization of various monosaccharides and disaccharides by Chlorella zofingiensis, indicating that glucose, fructose, mannose, and sucrose were able to be efficiently consumed by C. zofingiensis for fast growth whereas lactose and galactose were poorly assimilated. The growth of C. zofingiensis was inhibited when the sugar concentration exceeded 20 g L−1 as indicated by the decreased specific growth rate [33]. Raw materials rich in carbohydrates such as artichoke tubers and sorghum stems have also been reported as carbon sources to feed heterotrophic Chlorella cells [34, 35]. Other organic carbon sources that can be used for heterotrophic Chlorella growth include acetate, glycerol, lactate, glutamate, and methanol (Table 1). The high concentrations of these organic carbons, however, may confer an inhibitive effect on Chlorella growth; for example, glycerol above 2 % was reported to inhibit the growth of Chlorella vulgaris [36] severely, and methanol above 1 % was lethal to Chlorella minutissima [37].
4 Mass Cultivation
The outdoor mass cultivation of Chlorella started in the late 1940s with the almost concurrent launch in the United States, Germany, and Japan [38]. Afterwards the mass cultivation of algae became one of the hottest topics in algal biotechnology leading to the development of diverse culture systems for mass culture applications [39–42]. Generally, algal cultures can be grown photoautotrophically, heterotrophically, or mixotrophically, in open or closed culture systems.
4.1 Photoautotrophy
Chlorella possesses high PE and is commonly cultured outdoors driven by the sunlight in open ponds or closed photobioreactors (PBRs). The popularly used open ponds include circular ponds and raceway ponds. Circular ponds were first built in Japan and then introduced to China and are now widely employed for mass cultivation of Chlorella in Asia. The capacity scale-up of circular ponds can be achieved by increasing the pond diameter, which can reach up to 50 m [40]. The system has several disadvantages including requirement of expensive structures of heavy reinforced concrete, high energy consumption for continuous stirring, inefficiency in land use, and so on. The raceway pond is another popular open culture system in the world. The raceway pond was initially used for commercial production of Spirulina and is now also employed for mass culture of Chlorella. The raceways are typically made from poured concrete, or they are simply dug into the earth and lined with a plastic liner to prevent the ground from soaking up the liquid. A raceway pond appears to be a single unit or in the form of a meandering channel assembled by individual raceways [39]. Generally, the cultures in open ponds are kept shallow at a depth of no more than 30 cm to facilitate the penetration of sunlight. Overall, the cell density achieved in both circular and raceway ponds is relatively low and commonly less than 1 g dry weight L−1. In contrast, the cascade system, a newly developed open culture system by Czech researchers, is capable of reaching a high cell density of up to 40 g L−1 [41, 43, 44]. This system is characterized by the thin layer of suspension (ca. 6 mm), highly turbulent flow, and high ratio of exposed surface area to total volume. Although the cascade system can achieve high volumetric cell density, its overall areal productivity is just comparable to that of open ponds [43].
Open systems cost less to construct and maintain and are thus regarded more economically favorable than closed PBRs, but they have substantial intrinsic disadvantages including low cell density and biomass productivity, rapid water loss due to evaporation, easy contamination by other microorganisms, and difficulty in managing culture temperature and efficient CO2 delivery. The ease of contamination is a common problem encountered in outdoor cultures. Chlorella culture in open ponds is susceptible to other unwanted algae including diatoms and cyanobacteria, and protozoans such as rotifers, ciliates, and amoebae that feed on the algae, resulting in greatly reduced production of Chlorella biomass. Developing a best management practices plan for prevention and treatment of contamination may represent a feasible approach toward increased production economics of Chlorella by open systems.
Closed PBRs have the potential to overcome the problems of evaporation, contamination, and low biomass productivity encountered in open systems. PBRs are made of transparent materials with a large ratio of surface area to volume. Tubular PBR is one of the most popular designs. It can be arranged as straight tubes horizontally placed parallel to each other, α-type cross tubes at an angle with the horizon, or coiled tubes helically surrounding a supporting frame [39, 45–47]. Schenk et al. [47] reported a large tubular PBR system installed in Germany, which consists of 500 km of tubes arranged as fences in a north/south direction. This system has a capacity of 700 cm3 with the annual production of Chlorella biomass up to 100 t. Panel PBR is another popular design arranged either vertically or inclined to the ground [48–51]. Compared with tubular PBR, the panel design needs less capital for construction and operation, offers less dark volume, accumulates less dissolved oxygen, and so on. To date, the Arizona Center for Algae Technology and Innovation (AzCATI) at Arizona State University (United States) has launched a 0.5 acre of a panel PBR system for investigating the biomass production potential of Chlorella cultures.
4.2 Mixotrophy
Many Chlorella strains are able to grow robustly under mixotrophic conditions utilizing both CO2 and organic carbons in the presence of light (Table 1). Usually, Chlorella grows better under mixotrophic conditions than under autotrophic conditions [32, 36, 52]. Therefore, in some cases, organic carbons are added into open ponds or PBRs to achieve mixotrophic production of Chlorella biomass [40, 53]. However, the Chlorella cultures in open ponds supplemented with organic carbons, sugars in particular, are highly susceptible to bacterial contamination. To reduce the chance of contamination, stepwise feeding of acetate but not sugars is employed. Inasmuch as the PBR system is closed, it offers better performance than an open system to maintain a monoculture under mixotrophic conditions. There was a report of successful maintenance of Chlorella monoculture mixotrophically grown in outdoor PBRs supplemented with sugars [54]. But the monoculture was achieved only in small-volume PBRs (i.e., 10 L) and it got contaminated and crashed when the culture volume scaled up to 300 L. The fermenter system that was conventionally used for fermentation of nonphotosynthetic organisms such as bacteria, yeasts, and animal cells, has also been proposed to mixotrophically grow Chlorella by providing sugars and artificial light sources [55]. The fermenter design, however, is less efficient in light usage and mixotrophic growth, particularly when using large-volume fermenters for high density of cultures.
4.3 Heterotrophy
Heterotrophic growth of Chlorella in fermenters has a long history and is now gaining increasing attention [30, 32, 40, 56, 57]. Fermenters are usually placed indoors without provision of light. Almost all Chlorella species reported are able to grow robustly under heterotrophic conditions with the addition of organic carbon sources, sugars in particular (Table 1). These organic carbon sources, including sugars, hydrolyzed carbohydrates, acetate, and glycerol, serve as the solo carbon and energy sources to support the growth of Chlorella. Chlorella protothecoides and Chlorella zofingiensis are the most well-studied Chlorella species for fermentation. Li et al. [58] reported a scale-up of heterotrophic production of C. protothecoides in an 11,000-L fermenter with the cell density reaching up to 13 g L−1, which is comparable to that achieved in a 5-L fermenter. The competitiveness of heterotrophic production of Chlorella over photoautotrophic production rests largely with high cell density and great biomass productivity, elimination of the light requirement, ease of control for monocultures, and low-cost biomass harvesting [59]. The high cell density and biomass productivity of heterotrophic Chlorella can be achieved by the employment of fed-batch, continuous, and cell-recycle culture strategies that are well developed for the fermentation of bacteria or yeasts [31, 32, 56, 60, 61]. With the optimized fermentation conditions, heterotrophic Chlorella was reported to achieve as high as 100 g cell dry weight per liter with an average biomass productivity of 13 g L−1 day−1 [61]. Although Chlorella fermentation has been gaining the increasing attention of industry, it is regarded economically favorable only for high-value products but not for the low-cost commodity products such as biofuels, because of the relatively high production cost.
Each culture strategy mentioned above has its own advantages and disadvantages. The choice of cultivation methods depends on Chlorella species/strains, locations of culture systems, production capacities, desired products, and so on. In some cases, hybrid systems (e.g., PBR-pond and fermenter-PBR) instead of a sole culture system can be employed. In a PBR-pond system, Chlorella cells are cultured in PBRs for rapid growth, which serve as the inoculation seed in raceway ponds for large-scale biomass production. In a fermenter-PBR system, Chlorella cells are first grown heterotrophically in fermenters for accumulation of high-density biomass, which are then transferred to thin PBRs with high light for induction of desired products, for example, oils or astaxanthin. Regardless of algal strains and culture systems, the key to optimizing a production system lies in the cost balance of output to input.
5 Potential Applications
Chlorella is the sunlight-driven single-cell factory for protein, lipids, carbohydrates, pigments, vitamins, and minerals. It has long been used as health food and additives for human consumption, as well as animal feed in aquaculture. In addition, the green alga proves to be beneficial to environmental cleanup such as bioremediation of industrial flue gases and wastewater. Recently, due to the blooming of renewable energy, Chlorella has attracted unprecedented interest as a feedstock for biofuels, biodiesel in particular. Although the microalgal expression system does not reach a stage as mature as bacteria, yeast, mammalian cell, or plant systems, it shows substantial advantages and has been used increasingly for expression of recombinant proteins.
5.1 Chlorella as Human Food and Animal Feed
Chlorella is abundant in protein (up to 68 %) and contains all the essential amino acids. It is also rich in fatty acids, dietary fibers, carotenoids, vitamins, minerals, and other bioactive compounds, enabling the alga an attractive foodstuff of high nutritional quality. The use of Chlorella as nutritional food has a long history and can be traced back to food shortage periods during the World Wars. Japan and China are the main Chlorella-producing countries, with an annual production of over 3,500 t of biomass in 2005 [40]. Yaeyama Chlorella (Japan), Sun Chlorella (Japan), and Taiwan Chlorella are the most popular companies for Chlorella production. The produced Chlorella is commercialized mainly in the form of dried powder, tablets, or capsules for human consumption. Other forms of products include Chlorella growth factor (CGF), Chlorella tea, Chlorella noodles, and the like. CGF is a hot-water extract of Chlorella and represents a mixture of proteins, nucleic acids, polysaccharides, and a variety of minerals [40]. Administration of Chlorella or Chlorella extracts has been shown to play positive roles in health care and disease prevention, such as boosting immune functions [62, 63], preventing tumors and cancers [64, 65], enhancing hypoglycemic effects [66, 67], attenuating cognitive decline in age-dependent dementia [68], and lowering blood pressure [69]. Chlorella is also used as a natural color additive for human food because of its high levels of pigments [70, 71]. As stated by [70], the addition of Chlorella biomass gave cookies an attractive and innovative appearance and higher textural characteristics.
The use of Chlorella as animal feed is more recent. Nutritional Chlorella biomass can be used directly as feed or to enrich protozoa such as rotifers that serve as feed in aquaculture [72, 73]. Feeding of Chlorella proves beneficial to the growth and nutritional improvement of fish. The level of skin pigmentation is one of the most important quality criteria determining the market value of fish, ornamental fish in particular. They are unable to synthesize carotenoids de novo and have to feed on the carotenoid-containing organisms (e.g., microalgae) to achieve their natural pigmentation. Chlorella is rich in pigments and even keto-carotenoids depending on species and therefore is popularly used in aquaculture for coloring ornamental fish [74–76]. In addition, Chlorella shows promising applications in poultry, for example, feeding to hens to color their egg yolks [77]. Because Chlorella has a tough rigid cell wall, proper pretreatment is commonly necessary to facilitate the digestion and assimilation of nutrients from Chlorella [78, 79].
5.2 Chlorella as a Source of Carotenoids
Carotenoids commonly found in Chlorella include α- and β-carotenes, lutein, zeaxanthin, violaxanthin, and neoxanthin. Some keto-carotenoids such as canthaxanthin and astaxanthin are also found in certain Chlorella species [33]. Figure 1 shows the schematic pathway of carotenoid biosynthesis in Chlorella. Generally, hydroxylation of the C-3 and C-3′ positions of β-carotene and α-carotene results in the formation of zeaxanthin and lutein via β-cryptoxanthin and α-cryptoxanthin, respectively. The subsequent epoxidation of zeaxanthin leads to the production of violaxanthin which is further converted to neoxanthin. In Chlorella zofingiensis additional keto-carotenoid biosynthetic pathways are present, involving several oxygenation and hydroxylation steps that lead to the formation of astaxanthin from β-carotene [80]. Carotenoids have important applications in food, feed, nutraceutical, and pharmaceutical industries because of their strong coloring ability, powerful antioxidative activity, and beneficial effects on human health [81]. Using Chlorella as producers of lutein and astaxanthin has been proposed [31, 32, 82].
Shi et al. [83] analyzed seven Chlorella strains for lutein production and the results suggested that Chlorella protothecoides CS-41 was a potential producer of lutein inasmuch as it accumulated the highest level of lutein (4.5 mg g−1 dry weight). Later, Shi and Chen [82] investigated the growth and lutein production of C. protothecoides under both heterotrophic and mixotrophic culture conditions; mixotrophic cultures produced more biomass and higher amounts of lutein than heterotrophic ones. It was revealed that C. protothecoides was able to utilize various nitrogen sources including nitrate, ammonia, and urea, but urea proved to be superior to the other two nitrogen sources for growth and lutein production [24]. In order to increase lutein production, Shi and Chen [84] adopted a fed-batch culture strategy to grow C. protothecoides with urea as the nitrogen source, achieving a lutein yield up to 225 mg L−1 with the maximal productivity of 48 mg L−1 day−1. The comparable lutein productivity was obtained when the fed-batch process was scaled up to a 30-L culture volume [84], indicating the potential of using heterotrophic C. protothecoides for scalable production of lutein.
C. zofingiensis is the only Chlorella species known to synthesize keto-carotenoids including astaxanthin. The astaxanthin production potential of C. zofingiensis has been studied under photoautotrophic, heterotrophic, and mixotrophic conditions [33, 55, 85, 86]. Under photoautotrophic conditions, stresses such as high light intensity and/or nitrogen starvation are needed to induce astaxanthin accumulation in C. zofinginesis [87]. These stresses, however, are unfavorable for algal growth and biomass production. In addition, the attenuated light absorption caused by mutual shading of cells severely affects the productivity and quality of algal biomass and products. In contrast, heterotrophic cultivation of C. zofingiensis feeding on an organic carbon source can boost algal growth as well as astaxanthin accumulation, eliminating the need for light and light-associated growth issues [33]. It has been reported that heterotrophic cultures of C. zofingiensis could achieve a comparable astaxanthin yield to H. pluvialis on a volumetric basis, much higher than that under photoautotrophic conditions [87, 88]. In this context, heterotrophic culture mode is regarded to be more feasible than autotrophic mode for astaxanthin production by C. zofingiensis. Sugars, glucose in particular, are commonly used for heterotrophic production of astaxanthin from C. zofingiensis, making the production relatively expensive and thus hampering its commercial application to some extent. To reduce the production cost, waste sugars such as cane molasses have been proposed to replace glucose for astaxanthin accumulation [31, 32]. Cane molasses is a by-product of the sugar industry consisting mainly of sucrose, glucose, and fructose, and is much cheaper than glucose. It has been proved that molasses gave a comparable astaxanthin productivity to glucose [32], opening up a possibility of using industrially cheap organic carbons toward large-scale and cost-saving production of astaxanthin. Although C. zofingiensis can achieve high cell density under heterotrophic conditions, the intracellular astaxanthin content is relatively low compared to that under phototrophic conditions with high light and nitrogen starvation, offsetting in part the production economics of astaxanthin.
We have developed a heterotrophic–phototrophic two-stage culture strategy to grow C. zofingiensis to improve astaxanthin production: C. zofingiensis was first cultured in the presence of glucose in the dark for rapid accumulation of biomass, which was then transferred to high light conditions for induction of astaxanthin. The new culture strategy greatly enhanced the intracellular accumulation of astaxanthin to 3.5 mg g−1 of dry weight, which is 3.2 times the astaxanthin content obtained under heterotrophic conditions (unpublished data). The drastic increase in astaxanthin content may lie in that the alga needs more astaxanthin to cope with the light-associated adverse effects. Additionally, a record high astaxanthin productivity of 4.7 mg L−1 day−1 was achieved, which is 2.9- and 2.4-fold higher than that under heterotrophic and phototrophic conditions, respectively (unpublished data). The newly developed heterotrophic–phototrophic two-stage culture strategy combines the advantages of both heterotrophic and phototrophic modes and eliminates the possible contamination associated with mixotrophic growth, which can significantly enhance astaxanthin production and may open up the possibility of substituting the currently used heterotrophic culture method for commercial production of astaxanthin at large scale.
Another strategy to overcome the low astaxanthin content is to manipulate the carotenoid biosynthetic pathway in C. zofingiensis through genetic engineering. Aside from astaxanthin, C. zofingiensis contains substantial amounts of canthaxanthin and adonixanthin [89], suggesting that CHYb may not accept canthaxanthin as a substrate to produce astaxanthin and BKT might be insufficient to catalyze the formation of astaxanthin from adonixanthin in C. zofingiensis. It has been reported that CHYb from H. pluvialis can utilize canthaxanthin as the substrate for efficient synthesis of astaxanthin [90] and BKT from C. reinhardtii has a high activity of converting adonixanthin to astaxanthin [91]. Therefore, the manipulation of specific astaxanthin biosynthetic steps by introducing these two genes into C. zofingiensis may provide a pulling force for astaxanthin synthesis at the cost of both canthaxanthin and adonixanthin, which, when coupled with the pushing force from PDS overexpression, may represent a feasible strategy to increase astaxanthin content and purity further. We have developed a sophisticated transformation system for C. zofingiensis [80], whereby the genetic engineering for astaxanthin enhancement is possible.
5.3 Chlorella for CO2 Biomitigation and Wastewater Bioremediation
Global warming caused by the increasing greenhouse gases in the atmosphere has attracted great concern by the public and scientific community. Carbon dioxide is the principal greenhouse gas mainly released through burning of fossil fuels. The biomitigation of CO2 by autotrophs such as microalgae is a promising strategy proposed for fixing CO2 and attenuating the greenhouse effect. Chlorella is one of the most commonly used genera of algae for sequestration of CO2 due to its high growth rate and strong CO2 fixation ability [19, 20, 92–94]. Chlorella is able to fix CO2 from different sources, which can be simply classified as air, CO2-enriched air, and industrial exhaust gases such as flue gas. The CO2 fixation rate is associated with the CO2 concentration provided for Chlorella growth (Fig. 2). When provided with atmospheric air (ca. 0.04 % CO2), Chlorella grows poorly and shows low fixation abilities due to the mass transfer limitation. Therefore, to facilitate algal growth, the CO2- enriched air at the concentration of 1–5 % is usually provided, accompanied by the increased CO2 fixation rate. However, when the CO2 content is over 5 %, the fixation rate drops down significantly (Fig. 2).This may be mainly attributed to the decreased medium pH at high CO2 concentration that inhibits the growth of Chlorella. The flue gases contain up to 15 % CO2 and are responsible for more than 7 % of the total world CO2 emissions [95]. It will be environmentally and cost beneficial if Chlorella can directly use flue gases for CO2 fixation, which requires strains tolerant to high CO2 concentration as well as to relatively high temperature. High-CO2 tolerant Chlorella strains have been reported, some of which were able to grow in the presence of up to 40 % CO2 at 42 °C without significant growth inhibitory effects [22, 96].
The tolerance to both high CO2 and high temperature enables these Chlorella strains to be potential cellular reactors for the biomitigation of CO2 from flue gases. For example, Chlorella sp. UK001, one of the CO2 tolerant strains, showed a CO2 fixation rate of more than 1 g L−1 day−1 when aerated with 15 % CO2 [97], much higher than that of regular Chlorella strains. In addition, the choice of culture system affects the capacity of CO2 fixation. Doucha et al. [98] reported the CO2 fixation of flue gas from a natural gas-fired boiler by Chlorella cultivated in an outdoor open cascade system with a culture area of 55 m2. The inhibition of algal growth caused by sulphur and nitrogen oxides (SOx, NOx) that exists in flue gas [99] was not observed in this study. The biomass productivity and PAR utilization of Chlorella cultures saturated with flue gas were 19.4–22.8 g m−2 day−1 and 5.58–6.94 % respectively, comparable to that with pure CO2. Later, Douskova and Livansky [100] investigated the CO2 fixation rate by Chlorella vulgaris in aerated columns with flue gas or CO2-enriched air. Flue gas-aerated Chlorella cultures exhibited an even higher CO2 fixation rate (4.4 g L−1 day−1) than that aerated with CO2-enriched air (3.0 g L−1 day−1). Recently, there have been increasing reports of using Chlorella wild-type or mutant strains to sequestrate CO2 from industrial flue gas [92, 101–103].
It is noteworthy that a high concentration of CO2 generally leads to low efficiency of CO2 removal; for example, the removal efficiency by Chlorella sp. in the presence of 15 % CO2 is only 16 %, much lower than that in the presence of 2 % CO2 (58 %, [19]), indicating that a major portion of CO2 is released from the cultures. This can be overcome in part by passing the gases through sequential culture units where CO2 is resequestrated toward less emission. For example, the fixation efficiency of flue gas CO2 achieved by Chlorella cultures in sequential bioreactors reaches up to 85.6 %, greatly higher than that obtained in a single bioreactor [104]. As suggested by Doucha et al. [98], the daily fixed CO2 per m2 is around 34.4 g, with the simultaneous production of 20 g algal biomass. Assuming the culture season lasts for 150 days, one hectare of Chlorella cultures is able to sequestrate 21 t CO2 and produce around 12 t biomass per year.
Bioremediation using microalgae has long been recognized as an environmentally sound approach for wastewater treatment. Chlorella is one of the microalgal genera widely used in the biological treatment of wastewater and has proven abilities of removing nutrients (N and P), organic contaminants, and heavy metals (Table 2). Generally, pretreatments of wastewater such as settling, activated sludge process, or dilution are needed before supplying to Chlorella for biological treatment [105–108]. Wang et al. [109] intensively investigated the growth of Chlorella sp. on wastewater sampled from four different points of the treatment process flow of a municipal wastewater treatment plant for the removal of nitrogen, phosphorus, and chemical oxygen demand (COD) as well as metal ions. The four types of wastewater are classified as wastewater before primary settling (#1), wastewater after primary settling (#2), wastewater after activated sludge tank (#3), and concentrate (#4). The growth rate (0.95 day−1) and COD removal rate (83.0 %) of Chlorella for wastewater #4 were much higher than those for wastewater #1 and #2 and the removal rates of nitrogen (78.3 %) and phosphorus (85.6 %) were comparable; Chlorella in wastewater #3 showed the lowest growth rate and removal rates of nitrogen, phosphorus, and COD. The efficient removal of nutrients and organic contaminants by Chlorella from wastewater was also demonstrated in other studies [107, 110–114]. These results suggest that growing Chlorella on wastewater seems to be a feasible strategy to reduce the released amounts of organic and inorganic nutrients into natural waters, thus preventing the eutrophication problem.
Although the suspended Chlorella cultures exhibited their potential use in secondary or tertiary steps for wastewater treatment, one of the major and practical limitations is separation of the algal biomass from the treated wastewater, which requires capital-intensive steps such as flocculation, flotation, filtration, or centrifugation. In this context, using immobilized Chlorella cells for wastewater treatment is advantageous in that no harvest step is required [106, 115–117]. The removal efficiency of nutrients by immobilized Chlorella is influenced by culture density, pH, and immobilizing matrix [115, 118]. In some cases, Chlorella was cocultured with other microalgae for wastewater treatment [113]. In addition to removal of nutrients and organic compounds, Chlorella is also able to be used for the biodegradation of toxics [119] and removal of metal ions [109, 115].
The biosorption of Chlorella for removing metals from wastewater involves adsorption of metal ions onto the cell surface and binding to the intracellular molecules such as cytoplasmic ligands, phytochelatins, and metallothioneins (for details see the review by Mehta and Gaur [120]). Chlorella has been reported to remove a wide range of metals, including Al, Ca, Cd, Cu, Fe, Mg, Mn, Ni, Ur, and Zn [109, 121–125]. Considering the acceptable growth and lipid production of Chlorella on wastewater, the integration of biofuel production with wastewater treatment is proposed [109, 111, 112, 114, 126].
5.4 Chlorella as Feedstock for Biofuels
Petroleum fuels are recognized to be unsustainable due to their depleting supplies and release of greenhouse gas [127]. Renewable biofuels are promising alternatives to petroleum, among which biodiesel has attracted unprecedentedly increasing attention in recent years [128]. Compared with traditional fuels, the carbon-neutral biodiesel releases fewer gaseous pollutants and is considered environmentally beneficial. Currently, biodiesel is produced mainly from vegetable oils, animal fats, and waste cooking oils. Plant-oil–derived biodiesel, however, cannot realistically meet the existing need for transport fuels as immense arable lands have to be occupied in cultivating oil crops, causing food–fuels conflicts [39].
Microalgae have been considered as the promising alternative feedstock for biodiesel production because of their rapid growth and high oil content [39, 128, 129]. Furthermore, unlike oil crops, microalgae can be easily cultured in outdoor ponds or bioreactors, making them superior to oil crops in biomass production. Chlorella represents a group of green microalgal species that grow fast and are easily able to achieve and maintain mass cultures. There have been many research studies focusing on using Chlorella for biodiesel production in the past years, as shown in Table 3. Under optimal conditions Chlorella usually synthesizes a relatively low content of lipids (25 % on average) which can be greatly increased up to 66 % by stress conditions such as nitrogen starvation [17]. The stress conditions also favor the accumulation of neutral lipids, in particular, triacylglycerols (TAGs) that deposit in the cytosol. TAGs are considered to be superior to polar lipids (phospholipids and glycolipids) for biodiesel production. In this context, Chlorella can be first cultured under favorable conditions to maximize biomass production and then exposed to stress conditions to stimulate the accumulation of lipids including TAGs. The important properties of biodiesel such as cetane number, viscosity, cold flow, and oxidative stability are largely determined by the composition and structure of fatty acyl esters which in turn are determined by the characteristics of fatty acids of biodiesel feedstock, for example, carbon chain length and unsaturation degree [130]. The synthesized fatty acids in Chlorella are mainly of medium length, ranging from 16 to 18 carbons, despite the great variation in fatty acid composition (Table 4). Generally, saturated fatty esters possess a high cetane number and superior oxidative stability whereas unsaturated, especially polyunsaturated, fatty esters have improved low-temperature properties [131]. It is suggested that the modification of fatty esters, for example, enhancing the proportion of oleic acid (C18:1) ester, can provide a compromise solution between oxidative stability and low-temperature properties and therefore promote the quality of biodiesel [132]. In this regard, C. protothecoides, which has the highest proportion of oleic acid (71.6 %), may be better than other Chlorella species as biodiesel feedstock [34]. The properties of C. protothecoides derived biodiesel were assessed and most of them proved to comply with the limits established by American Society for Testing and Materials (ASTM), including density, viscosity, flash point, cold filter plugging point, and acid value.
Aside from employing photoautotrophic Chlorella cells, fermentation by feeding organic carbons has also been proposed by some research groups to enrich heterotrophic biomass as biodiesel feedstock. C. protothecoides is the most studied species heterotrophically grown for biodiesel production (Table 5). It could achieve very high cell densities, biomass productivities, and lipid productivities. Glucose is the most widely used carbon source to feed Chlorella for boosting biomass production and lipid accumulation. To reduce the production cost, alternative low-cost carbon sources such as hydrolysates of crude carbohydrates, waste molasses, or glycerol were used [34, 35, 61, 133–136]. Molasses proved to be a promising alternative to feed C. protothecoides for biodiesel, with the biomass yield, biomass productivity, and lipid productivity being 97.1 g L−1, 12.8, and 7.3 g L−1 day−1, respectively [61]. However, the conversion ratio of sugar to biomass in these heterotrophic cultures was restricted to 0.5, which means 2 t of sugar are required for producing 1 t of biomass and 1 t of CO2 is released during this process. In this regard, fermentation is neither economically viable nor environmentally friendly for the production of biomass for biodiesel as compared with photoautotrophy.
Transesterification is needed to convert Chlorella oil to biodiesel. It is a chemical conversion process involving reacting triglycerides catalytically with a short-chain alcohol (typically methanol or ethanol) to form fatty acyl esters (biodiesel) and glycerol. This reaction occurs stepwise with the first conversion of triglycerides to diglycerides and then to monoglycerides and finally to glycerol. Considering the reaction is reversible, a large excess of alcohol is used in industrial processes to ensure the direction of fatty acid esters. Methanol is the preferred alcohol for industrial use because of its low cost, although other alcohols including ethanol, propanol, and butanol are also commonly used. In addition to heat, a catalyst is needed to facilitate the transesterification. The transesterification of triglycerides can be catalyzed by acids, alkalis, or enzymes [137–139]. Currently, alkali (sodium hydroxide and potassium hydroxide) is the preferred type of catalyst for industrial production of biodiesel.
Chlorella is also capable of producing other biofuels such as hydrogen, ethanol, methane, and biocrude [140–144]. The biohydrogen production from microalgae has long been recognized. This process involves hydrogenase, an enzyme highly sensitive to O2. During photosynthesis, hydrogen evolution is transient due to the strong inhibition of hydrogenase by photosynthetically evolved O2 [145]. The sustainable hydrogen production from Chlorella, therefore, requires the maintenance of an anaerobic environment, which can be achieved through the inhibition of O2 evolution by sulfur deficiency [146]. It has been estimated that green algae could theoretically produce a maximum of 20 g hydrogen per m2 per day [145]. In addition to lipids, Chlorella biomass contains a substantial amount of carbohydrates (starch and cellulose) that can be used for ethanol production through technologies such as saccharification and fermentation [142, 147, 148]. Chlorella is also capable of producing ethanol through self-fermentation of intracellular starch under dark and anaerobic conditions, although the conversion efficiency is relatively low [142]. Anaerobic digestion of biomass results in the production of methane that can be used as a heat source or for electricity generation. The raw Chlorella biomass can be directly subjected to digestion, thus avoiding the biomass-harvest and oil-extraction processes used in algal biodiesel production and significantly bringing down the production cost and energy debt. But the digestion efficiency as stated by Ras et al. [143] was restricted to 50 %, indicating the need of proper pretreatments of the raw biomass for complete digestion. Hydrothermal liquefaction, which requires moderate temperatures as compared to the processes of pyrolysis and gasification, is commonly used for biocrude production from Chlorella biomass [141, 149].
The production of biofuels from microalgae is still far from commercialization. There are significant technical challenges yet to be addressed. A promising strategy is to integrate the production of biodiesel with other biofuels and high-value products, as well as the applications of treating flue gas and wastewater (Fig. 3).
5.5 Chlorella as Cell Factories for Recombinant Proteins
Chlorella offers substantial advantages as a promising alternative to currently well-established expression systems of bacteria, yeast, and mammalian cells. In contrast to bacteria, Chlorella belongs to eukaryotic organisms and therefore can perform correct posttranscriptional and posttranslational modifications essential for the production of functional eukaryotic proteins. Being easy, rapid, and inexpensive to grow and maintain on a large scale both indoors and outdoors, Chlorella is more cost effective as compared with the capital-intensive and time-consuming expression systems of mammalian cells, large farm animals, and higher plants. In addition, Chlorella has long been approved and used as health food for human consumption, suggesting the biological safety of engineered proteins from Chlorella. These characteristics enable Chlorella to be potentially useful as a bioreactor for synthesizing engineered products of interest such as enzymes, vaccines, monoclonal antibodies, and growth factors.
Nevertheless, the utilization of Chlorella for heterologous expression is hampered by the lack of a sophisticated genetic toolbox. The genetic engineering of Chlorella has achieved some success during the past 20 years (Table 6). The first report was conducted by Jarvis and Brown [150] who introduced a firefly luciferase gene into Chlorella ellipsoidea for heterologous expression; the luciferase activity, however, was not stable and disappeared after a few days. Later, Maruyama et al. [151] performed the transient expression of the β-glucoronidase (GUS) gene in transgenic Chlorella saccharophila. The transient expression of GUS was also reported in transgenic Chlorella ellipsoidea [152, 153] and Chlorella sp. [154]. These results suggested the feasibility of using Chlorella as an expression system for recombinant protein production. A dominant selectable marker is essential for easy and reliable selection of target Chlorella transformants. The frequently used selectable markers include bleomycin binding protein (Ble, resistant to phleomycin), chloramphenicol acetyltransferase (Cat, resistant to chloramphenicol), hygromycin B phosphotransferase (Hpt, resistant to hygromycin), and neomycin phosphotransferase II (NptII, resistant to kanamycin and geneticin), derived from either Escherichia coli or Streptoalloteichus rimosus (Table 6). Chlorella, however, is not so sensitive to these antibiotics and may need high concentrations to inhibit its growth completely [155]. In addition, Chlorella harboring these bacterial genes may be subject to biological safety problems when used as food or pharmaceuticals for human beings. Therefore, endogenous genes are advantageous as selectable markers for Chlorella transformation. Dawson et al. [156] reported the use of the nitrate reductase (NR) gene from Chlorella vulgaris to rescue the NR-deficient Chlorella sorokiniana mutants, resulting in stable transformants. Wang et al. [157] also reported the stable transformation of Chlorella ellipsoidea using an endogenous nitrate reductase gene to complement the NR-deficient mutant. The limitation of these trials lies in that a NR-deficient mutant is required as the expression host. Very recently, a phytoene desaturase (PDS) from a norflurazon-resistant mutant of Chlorella zofingiensis was proposed as a dominant selectable marker for stable transformation of Chlorella [80, 89, 158]. The Chlorella mutant harbored a point mutation on its PDS, which showed great resistance to norflurazon as well as significantly enhanced desaturation activity. It has been demonstrated that as low as 0.25 μg mL−1 of norflurazon is sufficient to select target transformants of Chlorella zofingiensis transformed with the mutated PDS gene. The transformants retained the norflurazon resistance after more than 100 times of subculture without selection of norflurazon, suggesting the great potential of using the endogenously derived PDS gene mutant as an effective and efficient selectable marker for the stable transformation of Chlorella zofingiensis as well as other Chlorella species.
Genetic manipulation of Chlorella for heterologous gene expression is still in its infant stage. The introduced foreign genes are subject to instability if not lack of expression because of various possible reasons including unstable nuclear integration, position effects, inefficient transcription from heterologous promoters, inaccurate RNA processing, and codon usage bias. Nevertheless, there have been several reports of using Chlorella for expressing commercially interesting proteins such as rabbit neutrophil peptide-1 [159–161], human growth hormone [155], flounder growth hormone [162], mercuric reductase from Bacillus megaterium [163], and trypsin-modulating oostatic factor from mosquitos [164]. Challenges for getting stable transgene integration and expression in Chlorella transformants will require further investigations to develop sophisticated genetic toolboxes with a powerful expression cassette for this small-sized and cell-wall–tough species.
6 Conclusions and Future Prospects
Chlorella is a sunlight-driven single-cell bioreactor that converts carbon dioxide to potential proteins, lipids, carbohydrates, and high-value biocompounds. It is among the most well-studied genera of microalgae for mass cultures. The abundance in protein and other nutritional elements, biological safety, and feasibility of growing and maintaining outdoors on a large scale enable Chlorella to be a good source of health food for human consumption. Chlorella is also considered as a potential source of microalgal oils for biofuel production. There are still substantial challenges involved in the biofuel production pipeline such as mass cultivation, harvest and drying, biomass disruption for oil extraction and conversion, and recycling of water and nutrients, making the Chlorella-derived biofuels currently capital intensive and far from economically viable as compared with fossil fuels. The integrated production of biofuels and other potential high-value products, coupled with the environmentally beneficial applications such as flue gas biomitigation and wastewater treatment represent a promising direction toward a cost-effective production process of Chlorella, which requires close collaboration between biologists and engineers. As interest in Chlorella increases, comprehensive analyses of certain potential strains are underway via genomic, transcriptomic, proteomic, lipidomic, and metabolomic approaches. The availability of those omics data will uncover the biological implications and facilitate the tailored manipulation of Chlorella for broader industrial applications.
References
Bar E, Rise M, Vishkautsan M, Arad S (1995) Pigments and structural changes in Chlorella zofingiensis upon light and nitrogen stress. J Plant Physiol 146:527–534
Gerken H, Donohoe B, Knoshaug E (2013) Enzymatic cell wall degradation of Chlorella vulgaris and other microalgae for biofuels production. Planta 237:239–253
Goncalves E, Johnson J, Rathinasabapathi B (2013) Conversion of membrane lipid acyl groups to triacylglycerol and formation of lipid bodies upon nitrogen starvation in biofuel green algae Chlorella UTEX29. Planta 238:895–906
Ikeda T, Takeda H (1995) Species-specific differences of pyrenoids in Chlorella (Chlorophyta). J Phycol 31:813–818
Yamada T, Sakaguchi K (1982) Comparative studies on Chlorella cell walls: induction of protoplast formation. Arch Microbiol 132:10–13
Kessler E (1976) Comparative physiology, biochemistry, and the taxonomy of Chlorella (Chlorophyceae). Plant Syst Evol 125:129–138
Kessler E, Huss VAR (1992) Comparative physiology and biochemistry and taxonomic assignment of the Chlorella (Chlorophyceae) strains of the culture collection of the University of Texas at Austin. J Phycol 28:550–553
Takeda H (1991) Sugar composition of the cell wall and the taxonomy of chlorella (Chlorophyceae). J Phycol 27:224–232
Takeda H (1993) Chemical-composition of cell-walls as a taxonomical marker. J Plant Res 106:195–200
Huss VAR, Frank C, Hartmann EC, Hirmer M, Kloboucek A, Seidel BM, Wenzeler P, Kessler E (1999) Biochemical taxonomy and molecular phylogeny of the genus Chlorella sensu lato (Chlorophyta). J Phycol 35:587–598
Krienitz L, Hegewald EH, Hepperle D, Huss VAR, Rohr T, Wolf M (2004) Phylogenetic relationship of Chlorella and Parachlorella gen. nov. (Chlorophyta, Trebouxiophyceae). Phycologia 43:529–542
Lloyd NDH, Canvin DT, Culver DA (1977) Photosynthesis and photorespiration in algae. Plant Physiol 59:936–940
Winokur M (1948) Photosynthesis relationships of Chlorella species. Am J Bot 35:207–214
Melis A (2009) Solar energy conversion efficiencies in photosynthesis: minimizing the chlorophyll antennae to maximize efficiency. Plant Sci 177:272–280
Wassink EC, Kok B, van Oorschot P (1964) The efficiency of light-energy conversion in Chlorella cultures as compared with higher plants. In: Burlew JS (ed) Algal culture: from laboratory to pilot plant. Carnegie Institution of Washington Publication, Washinton, D.C., pp 55–62
Shelp BJ, Canvin DT (1980) Utilization of exogenous inorganic carbon species in photosynthesis by Chlorella pyrenoidosa. Plant Physiol 65:774–779
Hsieh C-H, Wu W-T (2009) Cultivation of microalgae for oil production with a cultivation strategy of urea limitation. Bioresour Technol 100:3921–3926
Ong S-C, Kao C-Y, Chiu S-Y, Tsai M-T, Lin C-S (2010) Characterization of the thermal-tolerant mutants of Chlorella sp. with high growth rate and application in outdoor photobioreactor cultivation. Bioresour Technol 101:2880–2883
Chiu SY, Kao CY, Chen CH, Kuan TC, Ong SC, Lin CS (2008) Reduction of CO2 by a high-density culture of Chlorella sp in a semicontinuous photobioreactor. Bioresour Technol 99:3389–3396
Fulke AB, Mudliar SN, Yadav R, Shekh A, Srinivasan N, Ramanan R, Krishnamurthi K, Devi SS, Chakrabarti T (2010) Bio-mitigation of CO2, calcite formation and simultaneous biodiesel precursors production using Chlorella sp. Bioresour Technol 101:8473–8476
Papazi A, Makridis P, Divanach P, Kotzabasis K (2008) Bioenergetic changes in the microalgal photosynthetic apparatus by extremely high CO2 concentrations induce an intense biomass production. Physiol Plantarum 132:338–349
Sakai N, Sakamoto Y, Kishimoto N, Chihara M, Karube I (1995) Chlorella strains from hot springs tolerant to high temperature and high CO2. Energy Convers Manage 36:693–696
Davis EA, Dedrick J, French CS, Milner HW, Myers J, Smith JHC, Spoehr HA (1964) Laboratory experiments on Chlorella culture at the Carnegie Institution of Washington department of plant biology. In: Burlew JS (ed) Algal culture: from laboratory to pilot plant. Carnegie Institution of Washington Publication, Washington, D.C., pp 105–153
Shi X-M, Zhang X-W, Chen F (2000) Heterotrophic production of biomass and lutein by Chlorella protothecoides on various nitrogen sources. Enzyme Microb Tech 27:312–318
Goldman JC, Brewer PG (1980) Effect of nitrogen source and growth rate on phytoplankton-mediated changes in alkalinity. Limnol Oceanogr 25:352–357
Shen Y, Yuan W, Pei Z, Mao E (2010) Heterotrophic culture of Chlorella protothecoides in various nitrogen sources for lipid production. Appl Biochem Biotechnol 160:1674–1684
Illman AM, Scragg AH, Shales SW (2000) Increase in Chlorella strains calorific values when grown in low nitrogen medium. Enzyme Microb Tech 27:631–635
Ördög V, Stirk W, Bálint P, van Staden J, Lovász C (2012) Changes in lipid, protein and pigment concentrations in nitrogen-stressed Chlorella minutissima cultures. J Appl Phycol 24:907–914
Ramazanov A, Ramazanov Z (2006) Isolation and characterization of a starchless mutant of Chlorella pyrenoidosa STL-PI with a high growth rate, and high protein and polyunsaturated fatty acid content. Phycol Res 54:255–259
Liu J, Huang J, Fan KW, Jiang Y, Zhong Y, Sun Z, Chen F (2010) Production potential of Chlorella zofingienesis as a feedstock for biodiesel. Bioresour Technol 101:8658–8663
Liu J, Huang J, Jiang Y, Chen F (2012) Molasses-based growth and production of oil and astaxanthin by Chlorella zofingiensis. Bioresour Technol 107:393–398
Liu J, Sun Z, Zhong Y, Gerken H, Huang J, Chen F (2013) Utilization of cane molasses towards cost-saving astaxanthin production by a Chlorella zofingiensis mutant. J Appl Phycol 25:1447–1456
Ip PF, Chen F (2005) Production of astaxanthin by the green microalga Chlorella zofingiensis in the dark. Process Biochem 40:733–738
Cheng Y, Zhou W, Gao C, Lan K, Gao Y, Wu Q (2009) Biodiesel production from Jerusalem artichoke (Helianthus Tuberosus L.) tuber by heterotrophic microalgae Chlorella protothecoides. J Chem Technol Biotechnol 84:777–781
Gao C, Zhai Y, Ding Y, Wu Q (2010) Application of sweet sorghum for biodiesel production by heterotrophic microalga Chlorella protothecoides. Appl Energy 87:756–761
Liang Y, Sarkany N, Cui Y (2009) Biomass and lipid productivities of Chlorella vulgaris under autotrophic, heterotrophic and mixotrophic growth conditions. Biotechnol Lett 31:1043–1049
Kotzabasis K, Hatziathanasiou A, Bengoa-Ruigomez MV, Kentouri M, Divanach P (1999) Methanol as alternative carbon source for quicker efficient production of the microalgae Chlorella minutissima: role of the concentration and frequence of administration. J Biotechnol 70:357–362
Burlew JS (1964) Algal culture: from laboratory to pilot plant, 4th edn. Carnegie Institution of Washington Publication, Washington, D.C.
Chisti Y (2007) Biodiesel from microalgae. Biotechnol Adv 25:294–306
Lin L-P (2005) Chlorella: its ecology, structure, cultivation, bioprocess and application. Yi Hsien Publishing, Taipei, Taiwan
Masojidek J, Kopecky J, Giannelli L, Torzillo G (2011) Productivity correlated to photobiochemical performance of Chlorella mass cultures grown outdoors in thin-layer cascades. J Ind Microbiol Biotechnol 38:307–317
Moheimani N (2013) Inorganic carbon and pH effect on growth and lipid productivity of Tetraselmis suecica and Chlorella sp (Chlorophyta) grown outdoors in bag photobioreactors. J Appl Phycol 25:387–398
Doucha J, Lívanský K (2006) Productivity, CO2/O2 exchange and hydraulics in outdoor open high density microalgal (Chlorella sp.) photobioreactors operated in a Middle and Southern European climate. J Appl Phycol 18:811–826
Douskova I, Doucha J, Livansky K, Machat J, Novak P, Umysova D, Zachleder V, Vitova M (2009) Simultaneous flue gas bioremediation and reduction of microalgal biomass production costs. Appl Microbiol Biotechnol 82:179–185
Borowitzka MA (1999) Commercial production of microalgae: ponds, tanks, tubes and fermenters. J Biotechnol 70:313–321
Lee Y-K, Ding S-Y, Low C-S, Chang Y-C, Forday W, Chew P-C (1995) Design and performance of an α-type tubular photobioreactor for mass cultivation of microalgae. J Appl Phycol 7:47–51
Schenk P, Thomas-Hall S, Stephens E, Marx U, Mussgnug J, Posten C, Kruse O, Hankamer B (2008) Second generation biofuels: high-efficiency microalgae for biodiesel production. BioEnergy Res 1:20–43
Hu Q, Kurano N, Kawachi M, Iwasaki I, Miyachi S (1998) Ultrahigh-cell-density culture of a marine green alga Chlorococcum littorale in a flat-plate photobioreactor. Appl Microbiol Biotechnol 49:655–662
Liu J, Sommerfeld M, Hu Q (2013) Screening and characterization of Isochrysis strains and optimization of culture conditions for docosahexaenoic acid production. Appl Microbiol Biotechnol 97:4785–4798
Zemke P, Sommerfeld M, Hu Q (2013) Assessment of key biological and engineering design parameters for production of Chlorella zofingiensis (Chlorophyceae) in outdoor photobioreactors. Appl Microbiol Biotechnol 97:5645–5655
Zhang CW, Richmond A (2003) Sustainable, high-yielding outdoor mass cultures of Chaetoceros muelleri var. subsalsum and Isochrysis galban in vertical plate reactors. Mar Biotechnol 5:302–310
Lee Y-K (2001) Microalgal mass culture systems and methods: their limitation and potential. J Appl Phycol 13:307–315
Lee Y-K (1997) Commercial production of microalgae in the Asia-Pacific rim. J Appl Phycol 9:403–411
Lee Y-K, Ding S-Y, Hoe C-H, Low C-S (1996) Mixotrophic growth of Chlorella sorokiniana in outdoor enclosed photobioreactor. J Appl Phycol 8:163–169
Ip PF, Wong KH, Chen F (2004) Enhanced production of astaxanthin by the green microalga Chlorella zofingiensis in mixotrophic culture. Process Biochem 39:1761–1766
De la Hoz Siegler H, Ben-Zvi A, Burrell RE, McCaffrey WC (2011) The dynamics of heterotrophic algal cultures. Bioresour Technol 102:5764–5774
Xiong W, Li X, Xiang J, Wu Q (2008) High-density fermentation of microalga Chlorella protothecoides in bioreactor for microbio-diesel production. Appl Microbiol Biotechnol 78:29–36
Li X, Xu H, Wu Q (2007) Large-scale biodiesel production from microalga Chlorella protothecoides through heterotrophic cultivation in bioreactors. Biotechnol Bioeng 98:764–771
Chen F (1996) High cell density culture of microalgae in heterotrophic growth. Trends Biotechnol 14:421–426
Chen Y-H, Walker TH (2012) Fed-batch fermentation and supercritical fluid extraction of heterotrophic microalgal Chlorella protothecoides lipids. Bioresour Technol 114:512–517
Yan D, Lu Y, Chen Y-F, Wu Q (2011) Waste molasses alone displaces glucose-based medium for microalgal fermentation towards cost-saving biodiesel production. Bioresour Technol 102:6487–6493
Cheng FC, Lin A, Feng JJ, Mizoguchi T, Takekoshi H, Kubota H, Kato Y, Naoki Y (2004) Effects of chlorella on activities of protein tyrosine phosphatases, matrix metalloproteinases, caspases, cytokine release, B and T cell proliferations, and phorbol ester receptor binding. J Med Food 7:146–152
Suárez ER, Kralovec JA, Noseda MD, Ewart HS, Barrow CJ, Lumsden MD, Grindley TB (2005) Isolation, characterization and structural determination of a unique type of arabinogalactan from an immunostimulatory extract of Chlorella pyrenoidosa. Carbohydrate Res 340:1489–1498
Morimoto T, Nagatsu A, Murakami N, Sakakibara J, Tokuda H, Nishino H, Iwashima A (1995) Anti-tumour-promoting glyceroglycolipids from the green alga, Chlorella vulgaris. Phytochem 40:1433–1437
Sheng J, Yu F, Xin Z, Zhao L, Zhu X, Hu Q (2007) Preparation, identification and their antitumor activities in vitro of polysaccharides from Chlorella pyrenoidosa. Food Chem 105:533–539
Cherng J-Y, Shih M-F (2005) Potential hypoglycemic effects of Chlorella in streptozotocin-induced diabetic mice. Life Sci 77:980–990
Cherng J-Y, Shih M-F (2006) Improving glycogenesis in Streptozocin (STZ) diabetic mice after administration of green algae Chlorella. Life Sci 78:1181–1186
Nakashima Y, Ohsawa I, Konishi F, Hasegawa T, Kumamoto S, Suzuki Y, Ohta S (2009) Preventive effects of Chlorella on cognitive decline in age-dependent dementia model mice. Neurosci Lett 464:193–198
Merchant RE, Andre CA (2001) A review of recent clinical trials of the nutritional supplement Chlorella pyrenoidosa in the treatment of fibromyalgia, hypertension, and ulcerative colitis. Altern Ther Health Med 7:79–91
Gouveia L, Batista AP, Miranda A, Empis J, Raymundo A (2007) Chlorella vulgaris biomass used as colouring source in traditional butter cookies. Innov Food Sci Emerg 8:433–436
Gouveia L, Raymundo A, Batista A, Sousa I, Empis J (2006) Chlorella vulgaris and Haematococcus pluvialis biomass as colouring and antioxidant in food emulsions. Euro Food Res Technol 222:362–367
Hirayama K, Nakamura K (1976) Fundamental studies on the physiology of rotifers in mass culture—V. Dry Chlorella powder as a food for rotifers. Aquaculture 8:301–307
Işik O, Sarihan E, Kuşvuran E, Gül Ö, Erbatur O (1999) Comparison of the fatty acid composition of the freshwater fish larvae Tilapia zillii, the rotifer Brachionus calyciflorus, and the microalgae Scenedesmus abundans, Monoraphidium minitum and Chlorella vulgaris in the algae-rotifer-fish larvae food chains. Aquaculture 174:299–311
Gouveia L, Choubert G, Pereira N, Santinha J, Empis J, Gomes E (2002) Pigmentation of gilthead seabream, Sparus aurata (L. 1875), using Chlorella vulgaris (Chlorophyta, Volvocales) microalga. Aquac Res 33:987–993
Gouveia L, Gomes E, Empis J (1996) Potential use of a microalga (Chlorella vulgaris) in the pigmentation of rainbow trout (Oncorhynchus mykiss) muscle. Z Lebensm Unters For A 202:75–79
Gouveia L, Rema P, Pereira O, Empis J (2003) Colouring ornamental fish (Cyprinus carpio and Carassius auratus) with microalgal biomass. Aquacult Nutr 9:123–129
Gouveia L, Veloso V, Reis A, Fernandes H, Novais J, Empis J (1996) Chlorella vulgaris used to colour egg yolk. J Sci Food Agri 70:167–172
Janczyk P, Franke H, Souffrant WB (2007) Nutritional value of Chlorella vulgaris: effects of ultrasonication and electroporation on digestibility in rats. Anim Feed Sci Tech 132:163–169
Komaki H, Yamashita M, Niwa Y, Tanaka Y, Kamiya N, Ando Y, Furuse M (1998) The effect of processing of Chlorella vulgaris: K-5 on in vitro and in vivo digestibility in rats. Anim Feed Sci Tech 70:363–366
Liu J, Sun Z, Gerken H, Huang J, Jiang Y, Chen F (2014) Genetic engineering of the green alga Chlorella zofingiensis: a modified norflurazon-resistant phytoene desaturase gene as a dominant selectable marker. Appl Microbiol Biotechnol 98:5069–5079
Fraser PD, Bramley PM (2004) The biosynthesis and nutritional uses of carotenoids. Prog Lipid Res 43:228–265
Shi XM, Chen F (1999) Production and rapid extraction of lutein and the other lipid-soluble pigments from Chlorella protothecoides grown under heterotrophic and mixotrophic conditions. Food Nahrung 43:109–113
Shi X-M, Chen F, Yuan J-P, Chen H (1997) Heterotrophic production of lutein by selected Chlorella strains. J Appl Phycol 9:445–450
Shi X-M, Chen F (2002) High-yield production of lutein by the green microalga Chlorella protothecoides in heterotrophic fed-batch culture. Biotechnol Prog 18:723–727
Del Campo JA, Moreno J, Rodriguez H, Vargas MA, Rivas J, Guerrero MG (2000) Carotenoid content of chlorophycean microalgae: factors determining lutein accumulation in Muriellopsis sp. (Chlorophyta). J Biotechnol 76:51–59
Del Campo JA, Rodriguez H, Moreno J, Vargas MA, Rivas J, Guerrero MG (2004) Accumulation of astaxanthin and lutein in Chlorella zofingiensis (Chlorophyta). Appl Microbiol Biotechnol 64:848–854
Orosa M, Valero JF, Herrero C, Abalde J (2001) Comparison of the accumulation of astaxanthin in Haematococcus pluvialis and other green microalgae under N-starvation and high light conditions. Biotechnol Lett 23:1079–1085
Sun N, Wang Y, Li Y-T, Huang J-C, Chen F (2008) Sugar-based growth, astaxanthin accumulation and carotenogenic transcription of heterotrophic Chlorella zofingiensis (Chlorophyta). Process Biochem 43:1288–1292
Liu J, Zhong Y, Sun Z, Huang J, Sandmann G, Chen F (2010) One amino acid substitution in phytoene desaturase makes Chlorella zofingiensis resistant to norflurazon and enhances the biosynthesis of astaxanthin. Planta 232:61–67
Linden H (1999) Carotenoid hydroxylase from Haematococcus pluvialis: cDNA sequence, regulation and functional complementation. Biochim Biophys Acta 1446:203–212
Huang J, Zhong Y, Sandmann G, Liu J, Chen F (2012) Cloning and selection of carotenoid ketolase genes for the engineering of high-yield astaxanthin in plants. Planta 236:691–699
Borkenstein CG, Knoblechner J, Fruhwirth H, Schagerl M (2011) Cultivation of Chlorella emersonii with flue gas derived from a cement plant. J Appl Phycol 23:131–135
de Morais M, Costa J (2007) Carbon dioxide fixation by Chlorella kessleri, C. vulgaris, Scenedesmus obliquus, Spirulina sp. cultivated in flasks and vertical tubular photobioreactors. Biotechnol Lett 29:1349–1352
Yewalkar S, Li B, Posarac D, Duff S (2011) Potential for CO2 fixation by Chlorella pyrenoidosa grown in oil sands tailings water. Energy Fuel 25:1900–1905
Wang B, Li Y, Wu N, Lan C (2008) CO2 bio-mitigation using microalgae. Appl Microbiol Biotechnol 79:707–718
Michiki H (1995) Biological CO2 fixation and utilization project. Energy Convers Manage 36:701–705
Murakami M, Ikenouchi M (1997) The biological CO2 fixation and utilization project by rite (2) - Screening and breeding of microalgae with high capability in fixing CO2. Energy Convers Manage 38(Supplement):S493–S497
Doucha J, Straka F, Lívanský K (2005) Utilization of flue gas for cultivation of microalgae Chlorella sp. in an outdoor open thin-layer photobioreactor. J Appl Phycol 17:403–412
Negoro M, Shioji N, Miyamoto K, Micira Y (1991) Growth of microalgae in high CO2 gas and effects of SOx and NOx. Appl Biochem Biotechnol 28–29:877–886
Doucha J, Lívanský K (2009) Outdoor open thin-layer microalgal photobioreactor: potential productivity. J Appl Phycol 21:111–117
Cheng J, Huang Y, Feng J, Sun J, Zhou J, Cen K (2013) Mutate Chlorella sp. by nuclear irradiation to fix high concentrations of CO2. Bioresour Technol 136:496–501
He L, Subramanian VR, Tang YJ (2012) Experimental analysis and model-based optimization of microalgae growth in photo-bioreactors using flue gas. Biomass Bioenerg 41:131–138
Kumar K, Banerjee D, Das D (2014) Carbon dioxide sequestration from industrial flue gas by Chlorella sorokiniana. Bioresour Technol 152:225–233
Cheng J, Huang Y, Feng J, Sun J, Zhou J, Cen K (2013) Improving CO2 fixation efficiency by optimizing Chlorella PY-ZU1 culture conditions in sequential bioreactors. Bioresour Technol 144:321–327
Fallowfield HJ, Garrett MK (1985) The photosynthetic treatment of pig slurry in temperate climatic conditions: a pilot-plant study. Agr Wastes 12:111–136
Shi J, Podola B, Melkonian M (2007) Removal of nitrogen and phosphorus from wastewater using microalgae immobilized on twin layers: an experimental study. J Appl Phycol 19:417–423
Tam NFY, Wong YS (1989) Wastewater nutrient removal by Chlorella pyrenoidosa and Scenedesmus sp. Environ Pollut 58:19–34
Yun Y-S, Lee SB, Park JM, Lee C-I, Yang J-W (1997) Carbon dioxide fixation by algal cultivation using wastewater nutrients. J Chem Technol Biotechnol 69:451–455
Wang L, Min M, Li YC, Chen P, Chen YF, Liu YH, Wang YK, Ruan R (2010) Cultivation of breen algae Chlorella sp in different wastewaters from municipal wastewater treatment plant. Appl Biochem Biotechnol 162:1174–1186
Aziz MA, Ng WJ (1992) Feasibility of wastewater treatment using the activated-algae process. Bioresour Technol 40:205–208
Ji M-K, Kim H-C, Sapireddy V, Yun H-S, Abou-Shanab RI, Choi J, Lee W, Timmes T, Jeon Inamuddin B-H (2013) Simultaneous nutrient removal and lipid production from pretreated piggery wastewater by Chlorella vulgaris YSW-04. Appl Microbiol Biotechnol 97:2701–2710
Li Y, Chen Y-F, Chen P, Min M, Zhou W, Martinez B, Zhu J, Ruan R (2011) Characterization of a microalga Chlorella sp. well adapted to highly concentrated municipal wastewater for nutrient removal and biodiesel production. Bioresour Technol 102:5138–5144
Tarlan E, Dilek FB, Yetis U (2002) Effectiveness of algae in the treatment of a wood-based pulp and paper industry wastewater. Bioresour Technol 84:1–5
Yang J, Rasa E, Tantayotai P, Scow KM, Yuan H, Hristova KR (2011) Mathematical model of Chlorella minutissima UTEX2341 growth and lipid production under photoheterotrophic fermentation conditions. Bioresour Technol 102:3077–3082
Mallick N, Rai LC (1994) Removal of inorganic ions from wastewaters by immobilized microalgae. W J Microbiol Biotechnol 10:439–443
Megharaj M, Pearson HW, Venkateswarlu K (1992) Removal of nitrogen and phosphorus by immobilized cells of Chlorella vulgaris and Scenedesmus bijugatus isolated from soil. Enzyme Microb Technol 14:656–658
Tam NFY, Lau PS, Wong YS (1994) Wastewater inorganic N and P removal by immobilized Chlorella vulgaris. Water Sci Technol 30:369–374
Mallick N, Rai LC (1993) Influence of culture density, pH, organic acids and divalent cations on the removal of nutrients and metals by immobilized Anabaena doliolum and Chlorella vulgaris. W J Microbiol Biotechnol 9:196–201
Jin J, Yang LH, Chan SMN, Luan TG, Li Y, Tam NFY (2011) Effect of nutrients on the biodegradation of tributyltin (TBT) by alginate immobilized microalga, Chlorella vulgaris, in natural river water. J Hazard Mater 185:1582–1586
Mehta SK, Gaur JP (2005) Use of algae for removing heavy metal ions from wastewater: progress and prospects. Crit Rev Biotechnol 25:113–152
Chong AMY, Wong YS, Tam NFY (2000) Performance of different microalgal species in removing nickel and zinc from industrial wastewater. Chemosphere 41:251–257
Kalin M, Wheeler WN, Meinrath G (2004) The removal of uranium from mining waste water using algal/microbial biomass. J Environ Radioact 78:151–177
Lau PS, Lee HY, Tsang CCK, Tam NFY, Wong YS (1999) Effect of metal interference, pH and temperature on Cu and Ni biosorption by Chlorella vulgaris and Chlorella miniata. Environ Technol 20:953–961
Mehta SK, Gaur JP (2001) Removal of Ni and Cu from single and binary metalsolutions by free and immobilized Chlorella vulgaris. Euro J Protistol 37:261–271
Sandau E, Sandau P, Pulz O (1996) Heavy metal sorption by microalgae. Acta Biotechnol 16:227–235
Mutanda T, Karthikeyan S, Bux F (2011) The utilization of post-chlorinated municipal domestic wastewater for biomass and lipid production by Chlorella spp. under batch conditions. Appl Biochem Biotechnol 164:1126–1138
Chisti Y (2008) Biodiesel from microalgae beats bioethanol. Trends Biotechnol 26:126–131
Hu Q, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert M, Darzins A (2008) Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances. Plant J 54:621–639
Liu J, Huang J, Chen F (2011a) Microalgae as feedstocks for biodiesel production. Biodiesel—Feedstocks and processing technologies. InTech, Available from http://www.intechopen.com/articles/show/title/microalgae-as-feedstocks-for-biodiesel-production
Knothe G (2005) Dependence of biodiesel fuel properties on the structure of fatty acid alkyl esters. Fuel Process Technol 86:1059–1070
Knothe G (2008) “Designer” biodiesel: optimizing fatty ester composition to improve fuel properties. Energy Fuel 22:1358–1364
Knothe G (2009) Improving biodiesel fuel properties by modifying fatty ester composition. Energy Environ Sci 2:759–766
Cerón-García MC, Macías-Sánchez MD, Sánchez-Mirón A, García-Camacho F, Molina-Grima E (2013) A process for biodiesel production involving the heterotrophic fermentation of Chlorella protothecoides with glycerol as the carbon source. Appl Energy 103:341–349
Espinosa-Gonzalez I, Parashar A, Bressler DC (2014) Heterotrophic growth and lipid accumulation of Chlorella protothecoides in whey permeate, a dairy by-product stream, for biofuel production. Bioresour Technol 155:170–176
O’Grady J, Morgan JA (2011) Heterotrophic growth and lipid production of Chlorella protothecoides on glycerol. Bioprocess Biosyst Eng 34:121–125
Xu H, Miao X, Wu Q (2006) High quality biodiesel production from a microalga Chlorella protothecoides by heterotrophic growth in fermenters. J Biotechnol 126:499–507
Fukuda H, Kondo A, Noda H (2001) Biodiesel fuel production by transesterification of oils. J Biosci Bioeng 92:405–416
Gerpen JV (2005) Biodiesel processing and production. Fuel Process Technol 86:1097–1107
Ranganathan SV, Narasimhan SL, Muthukumar K (2008) An overview of enzymatic production of biodiesel. Bioresour Technol 99:3975–3981
Bala Amutha K, Murugesan AG (2011) Biological hydrogen production by the algal biomass Chlorella vulgaris MSU 01 strain isolated from pond sediment. Bioresour Technol 102:194–199
Duan P, Jin B, Xu Y, Yang Y, Bai X, Wang F, Zhang L, Miao J (2013) Thermo-chemical conversion of Chlorella pyrenoidosa to liquid biofuels. Bioresour Technol 133:197–205
Hirano A, Ueda R, Hirayama S, Ogushi Y (1997) CO2 fixation and ethanol production with microalgal photosynthesis and intracellular anaerobic fermentation. Energy 22:137–142
Ras M, Lardon L, Bruno S, Bernet N, Steyer J-P (2011) Experimental study on a coupled process of production and anaerobic digestion of Chlorella vulgaris. Bioresour Technol 102:200–206
Wang HY, Fan XL, Zhang YT, Yang DW, Guo RB (2011) Sustained photo-hydrogen production by Chlorella pyrenoidosa without sulfur depletion. Biotechnol Lett 33:1345–1350
Melis A, Happe T (2001) Hydrogen production. Green algae as a source of energy. Plant Physiol 127:740–748
Rashid N, Lee K, Mahmood Q (2011) Bio-hydrogen production by Chlorella vulgaris under diverse photoperiods. Bioresour Technol 102:2101–2104
Cheng Y-S, Zheng Y, Labavitch JM, Vander Gheynst JS (2013) Virus infection of Chlorella variabilis and enzymatic saccharification of algal biomass for bioethanol production. Bioresour Technol 137:326–331
Zhou N, Zhang Y, Wu X, Gong X, Wang Q (2011) Hydrolysis of Chlorella biomass for fermentable sugars in the presence of HCl and MgCl2. Bioresour Technol 102:10158–10161
Zhang J, Chen W-T, Zhang P, Luo Z, Zhang Y (2013) Hydrothermal liquefaction of Chlorella pyrenoidosa in sub- and supercritical ethanol with heterogeneous catalysts. Bioresour Technol 133:389–397
Jarvis EE, Brown LM (1991) Transient expression of firefly luciferase in protoplasts of the green alga Chlorella ellipsoidea. Curr Genet 19:317–321
Maruyama M, Horakova I, Honda H, Xing XH, Shiragami N, Unno H (1994) Introduction of foreign DNA into Chlorella saccharophila by electroporation. Biotechnol Tech 8:821–826
Chen Y, Li W, Bai Q, Sun Y (1998) Study on transient expression of gus gene in Chlorelia ellipsoidea (Chlorophyta) by using biolistic particle delivery system. Chin J Oceanol Limn 16:47–49
Wang P, Sun YR, Li X, Zhang LM, Li WB, Wang YQ (2004) Rapid isolation and functional analysis of promoter sequences of the nitrate reductase gene from Chlorella ellipsoidea. J Appl Phycol 16:11–16
Wang C, Wang Y, Su Q, Gao X (2007) Transient expression of the GUS gene in a unicellular marine green alga Chlorella sp. MACC/C95, via electroporation. Biotechnol Bioprocess Eng 12:180–183
Hawkins RL, Nakamura M (1999) Expression of human growth hormone by the eukaryotic alga, Chlorella. Curr Microbiol 38:335–341
Dawson HN, Burlingame R, Cannons AC (1997) Stable transformation of Chlorella: rescue of nitrate reductase-deficient mutants with the nitrate reductase gene. Curr Microbiol 35:356–362
Wang YQ, Chen Y, Zhang XY, Wang P, Geng DG, Zhao SM, Zhang LM, Sun YR (2005) Isolation and characterization of a nitrate reductase deficient mutant of Chlorella ellipsoidea (Chlorophyta). J Appl Phycol 17:281–286
Huang JC, Liu J, Li YT, Chen F (2008) Isolation and characterization of the phytoene desaturase gene as a potential selective marker for genetic engineering of the astaxanthin-producing green alga Chlorella zofingiensis (Chlorophyta). J Phycol 44:684–690
Chen Y, Wang YQ, Sun YR, Zhang LM, Li WB (2001) Highly efficient expression of rabbit neutrophil peptide-1 gene in Chlorella ellipsoidea cells. Curr Genet 39:365–370
Han XM, Li YG, Sun XZ, Wei XD, Sun YR, Wang YQ (2005) Studies on the heterotrophic cultivation of transgenic Chlorella containing the rabbit defensin gene. Process Biochem 40:3055–3060
Wang Y, Chen Y, Bai Q, Zhao S, Li W, Sun Y (2001) Using transgenic Chlorella ellipsoidea as bio-reactor to produce rabbit defensin. High Technol Lett 9:1–5
Kim DH, Kim YT, Cho JJ, Bae JH, Hur SB, Hwang I, Choi TJ (2002) Stable integration and functional expression of flounder growth hormone gene in transformed microalga, Chlorella ellipsoidea. Mar Biotechnol 4:63–73
Huang CC, Chen MW, Hsieh JL, Lin WH, Chen PC, Chien LF (2006) Expression of mercuric reductase from Bacillus megaterium MB1 in eukaryotic microalga Chlorella sp DT: an approach for mercury phytoremediation. Appl Microbiol Biotechnol 72:197–205
Borovsky D (2003) Trypsin-modulating oostatic factor: a potential new larvicide for mosquito control. J Exp Biol 206:3869–3875
Avery SV, Codd GA, Gadd GM (1992) Replacement of cellular potassium by caesium in Chlorella emersonii: differential sensitivity of photoautotrophic and chemoheterotrophic growth. J Gen Microbiol 138:69–76
Oh SH, Kwon MC, Choi WY, Seo YC, Kim GB, Kang DH, Lee SY, Lee HY (2010) Long-term outdoor cultivation by perfusing spent medium for biodiesel production from Chlorella minutissima. J Biosci Bioeng 110:194–200
Yang J, Li X, Hu H, Zhang X, Yu Y, Chen Y (2011) Growth and lipid accumulation properties of a freshwater microalga, Chlorella ellipsoidea YJ1, in domestic secondary effluents. Appl Energy 88:3295–3299
Li T, Zheng Y, Yu L, Chen S (2013) High productivity cultivation of a heat-resistant microalga Chlorella sorokiniana for biofuel production. Bioresour Technol 131:60–67
Miao X, Wu Q (2006) Biodiesel production from heterotrophic microalgal oil. Bioresour Technol 97:841–846
Ruiz NJ, García MDCC, Mirón AS, Haftalaui EHB, Camacho FG, Grima EM (2009) Lipids accumulation in Chlorella protothecoides through mixotrophic and heterotrophic cultures for biodiesel production. New Biotechnol 25:S266–S266
Rai MP, Nigam S, Sharma R (2013) Response of growth and fatty acid compositions of Chlorella pyrenoidosa under mixotrophic cultivation with acetate and glycerol for bioenergy application. Biomass Bioenergy 58:251–257
Running J, Huss R, Olson P (1994) Heterotrophic production of ascorbic acid by microalgae. J Appl Phycol 6:99–104
Endo H, Sansawa H, Nakajima K (1977) Studies on Chlorella regularis, heterotrophic fast-growing strain II. Mixotrophic growth in relation to light intensity and acetate concentration. Plant Cell Physiol 18:199–205
Sansawa H, Endo H (2004) Production of intracellular phytochemicals in Chlorella under heterotrophic conditions. J Biosci Bioenergy 98:437–444
Herrera-Valencia V, Contreras-Pool P, López-Adrián S, Peraza-Echeverría S, Barahona-Pérez L (2011) The green microalga Chlorella saccharophila as a suitable source of oil for biodiesel production. Curr Microbiol 63:151–157
Tan C, Johns M (1991) Fatty acid production by heterotrophic Chlorella saccharophila. Hydrobiologia 215:13–19
Chen F, Johns M (1991) Effect of C/N ratio and aeration on the fatty acid composition of heterotrophic Chlorella sorokiniana. J Appl Phycol 3:203–209
Heredia-Arroyo T, Wei W, Ruan R, Hu B (2011) Mixotrophic cultivation of Chlorella vulgaris and its potential application for the oil accumulation from non-sugar materials. Biomass Bioenergy 35:2245–2253
Zhang K, Sun B, She X, Zhao F, Cao Y, Ren D, Lu J (2014) Lipid production and composition of fatty acids in Chlorella vulgaris cultured using different methods: photoautotrophic, heterotrophic, and pure and mixed conditions. Ann Microbiol 64:1239–1246
Ramos Tercero EA, Sforza E, Morandini M, Bertucco A (2014) Cultivation of Chlorella protothecoides with urban wastewater in continuous photobioreactor: Biomass productivity and nutrient removal. Appl Biochem Biotechnol 172:1470–1485
Wang L, Li Y, Chen P, Min M, Chen Y, Zhu J, Ruan RR (2010) Anaerobic digested dairy manure as a nutrient supplement for cultivation of oil-rich green microalgae Chlorella sp. Bioresour Technol 101:2623–2628
Kobayashi N, Noel EA, Barnes A, Watson A, Rosenberg JN, Erickson G, Oyler GA (2013) Characterization of three Chlorella sorokiniana strains in anaerobic digested effluent from cattle manure. Bioresour Technol 150:377–386
Bertoldi FC, Sant’Anna E, da Costa Braga MV, Oliveira JLB (2006) Lipids, fatty acids composition and carotenoids of Chlorella vulgaris cultivated in hydroponic wastewater. Grasas Y Aceites 57:270–274
Chu W-L, See Y-C, Phang S-M (2009) Use of immobilised Chlorella vulgaris for the removal of colour from textile dyes. J Appl Phycol 21:641–648
Lim S-L, Chu W-L, Phang S-M (2010) Use of Chlorella vulgaris for bioremediation of textile wastewater. Bioresour Technol 101:7314–7322
Abou-Shanab RAI, Hwang J-H, Cho Y, Min B, Jeon B-H (2011) Characterization of microalgal species isolated from fresh water bodies as a potential source for biodiesel production. Appl Energy 88:3300–3306
Cao J, Yuan H, Li B, Yang J (2014) Significance evaluation of the effects of environmental factors on the lipid accumulation of Chlorella minutissima UTEX 2341 under low-nutrition heterotrophic condition. Bioresour Technol 152:177–184
Converti A, Casazza AA, Ortiz EY, Perego P, Del Borghi M (2009) Effect of temperature and nitrogen concentration on the growth and lipid content of Nannochloropsis oculata and Chlorella vulgaris for biodiesel production. Chem Eng Process 48:1146–1151
Feng P, Deng Z, Hu Z, Fan L (2011) Lipid accumulation and growth of Chlorella zofingiensis in flat plate photobioreactors outdoors. Bioresour Technol 102:10577–10584
Liu Z-Y, Wang G-C, Zhou B-C (2008) Effect of iron on growth and lipid accumulation in Chlorella vulgaris. Bioresour Technol 99:4717–4722
Pruvost J, Van Vooren G, Le Gouic B, Couzinet-Mossion A, Legrand J (2011) Systematic investigation of biomass and lipid productivity by microalgae in photobioreactors for biodiesel application. Bioresour Technol 102:150–158
Rodolfi L, Zittelli GC, Bassi N, Padovani G, Biondi N, Bonini G, Tredici MR (2009) Microalgae for oil: Strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnol Bioenergy 102:100–112
Scarsella M, Parisi MP, D’Urso A, De Filippis P, Opoka J, Bravi M (2009) Achievements and perspectives in hetero- and mixotrophic culturing of microalgae. In: Pierucci S (ed) Icheap-9: 9th International Conference on Chemical and Process Engineering, Pts 1-3, vol 17., Chemical Engineering TransactionsAidic Servizi Srl, Milano, pp 1065–1070
Widjaja A, Chien C-C, Ju Y-H (2009) Study of increasing lipid production from fresh water microalgae Chlorella vulgaris. J Taiwan Inst Chem E 40:13–20
Yeh KL, Chang JS, Chen WM (2010) Effect of light supply and carbon source on cell growth and cellular composition of a newly isolated microalga Chlorella vulgaris ESP-31. Eng Life Sci 10:201–208
Wang Y, Rischer H, Eriksen NT, Wiebe MG (2013) Mixotrophic continuous flow cultivation of Chlorella protothecoides for lipids. Bioresour Technol 144:608–614
D’Oca MGM, Viêgas CV, Lemões JS, Miyasaki EK, Morón-Villarreyes JA, Primel EG, Abreu PC (2011) Production of FAMEs from several microalgal lipidic extracts and direct transesterification of the Chlorella pyrenoidosa. Biomass Bioenerg 35:1533–1538
Shekh AY, Shrivastava P, Krishnamurthi K, Mudliar SN, Devi SS, Kanade GS, Lokhande SK, Chakrabarti T (2013) Stress-induced lipids are unsuitable as a direct biodiesel feedstock: a case study with Chlorella pyrenoidosa. Bioresour Technol 138:382–386
Chinnasamy S, Bhatnagar A, Hunt RW, Das KC (2010) Microalgae cultivation in a wastewater dominated by carpet mill effluents for biofuel applications. Bioresour Technol 101:3097–3105
Li Y, Yuan Z, Mu J, Chen D, Feng B (2013) Proteomic analysis of lipid accumulation in Chlorella protothecoides cells by heterotrophic N deprivation coupling cultivation. Energy Fuel 27:4031–4040
Zheng Y, Li T, Yu X, Bates PD, Dong T, Chen S (2013) High-density fed-batch culture of a thermotolerant microalga Chlorella sorokiniana for biofuel production. Appl Energy 108:281–287
Matsumoto M, Sugiyama H, Maeda Y, Sato R, Tanaka T, Matsunaga T (2010) Marine diatom, Navicula sp. strain JPCC DA0580 and marine green alga, Chlorella sp. strain NKG400014 as potential sources for biodiesel production. Appl Biochem Biotechnol 161:483–490
Moazami N, Ranjbar R, Ashori A, Tangestani M, Nejad AS (2011) Biomass and lipid productivities of marine microalgae isolated from the Persian Gulf and the Qeshm Island. Biomass Bioenerg 35:1935–1939
Phukan MM, Chutia RS, Konwar BK, Kataki R (2011) Microalgae Chlorella as a potential bio-energy feedstock. Appl Energy 88:3307–3312
Yeesang C, Cheirsilp B (2011) Effect of nitrogen, salt, and iron content in the growth medium and light intensity on lipid production by microalgae isolated from freshwater sources in Thailand. Bioresour Technol 102:3034–3040
Liu J, Huang J, Sun Z, Zhong Y, Jiang Y, Chen F (2011) Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic Chlorella zofingiensis: assessment of algal oils for biodiesel production. Bioresour Technol 102:106–110
Feng Y, Li C, Zhang D (2011) Lipid production of Chlorella vulgaris cultured in artificial wastewater medium. Bioresour Technol 102:101–105
Yoo C, Jun S-Y, Lee J-Y, Ahn C-Y, Oh H-M (2010) Selection of microalgae for lipid production under high levels carbon dioxide. Bioresour Technol 101:S71–S74
Liang G, Mo Y, Zhou Q (2013) Optimization of digested chicken manure filtrate supplementation for lipid overproduction in heterotrophic culture Chlorella protothecoides. Fuel 108:159–165
Liu L, Wang Y, Zhang Y, Chen X, Zhang P, Ma S (2013) Development of a new method for genetic transformation of the green alga Chlorella ellipsoidea. Mol Biotechnol 54:211–219
El-sheekh MM (1999) Stable transformation of the intact cells of Chlorella kessleri with high velocity microprojectiles. Biol Plantarum 42:209–216
Chow KC, Tung WL (1999) Electrotransformation of Chlorella vulgaris. Plant Cell Rep 18:778–780
Niu Y, Zhang M, Xie W, Li J, Gao Y, Yang W, Liu J, Li H (2011) A new inducible expression system in a transformed green alga, Chlorella vulgaris. Genet Mol Res 10:3427–3434
Cha T, Yee W, Aziz A (2012) Assessment of factors affecting Agrobacterium-mediated genetic transformation of the unicellular green alga, Chlorella vulgaris. World J Microbiol Biotechnol 28:1771–1779
Talebi A, Tohidfar M, Tabatabaei M, Bagheri A, Mohsenpor M, Mohtashami S (2013) Genetic manipulation, a feasible tool to enhance unique characteristic of Chlorella vulgaris as a feedstock for biodiesel production. Mol Biol Rep 40:4421–4428
Xiong W, Gao C, Yan D, Wu C, Wu Q (2010) Double CO2 fixation in photosynthesis-fermentation model enhances algal lipid synthesis for biodiesel production. Bioresour Technol 101:2287–2293
Acknowledgments
This study was partially supported by a grant from the 985 Project of Peking University, by the State Oceanic Administration of China, and by the National Research Foundation of Singapore.
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Liu, J., Chen, F. (2014). Biology and Industrial Applications of Chlorella: Advances and Prospects. In: Posten, C., Feng Chen, S. (eds) Microalgae Biotechnology. Advances in Biochemical Engineering/Biotechnology, vol 153. Springer, Cham. https://doi.org/10.1007/10_2014_286
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DOI: https://doi.org/10.1007/10_2014_286
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