Open access peer-reviewed chapter

Fatty Acids from Microalgae: Targeting the Accumulation of Triacylglycerides

Written By

Paola Scodelaro Bilbao, Gabriela A. Salvador and Patricia I. Leonardi

Submitted: September 27th, 2016 Reviewed: January 16th, 2017 Published: June 21st, 2017

DOI: 10.5772/67482

Chapter metrics overview

2,835 Chapter Downloads

View Full Metrics

Abstract

Microalgae were originally considered as sources of long-chain polyunsaturated fatty acids (PUFAs), mainly for aquaculture purposes. However, based on the fact that their fatty acids (FA), stored as triacylglycerides (TAG), can be converted into biodiesel via a transesterification reaction, several microalgal species have emerged over the last decade as promising feedstocks for biofuel production. Elucidation of microalgae FA and TAG metabolic pathways is therefore becoming a cutting-edge field for developing transgenic algal strains with improved lipid accumulation ability. Furthermore, many of the biomolecules produced by microalgae can also be exploited. In this chapter, we describe recent advances in the field of FA and TAG pathways in microalgae, focusing in particular on the enzymes involved in FA and TAG synthesis, their accumulation in lipid droplets, and their degradation. Mention is made of potentially high-value products that can be obtained from microalgae, and possible molecular targets for enhancing FA and TAG production are outlined. A summary is provided of transcriptomics, proteomics, and metabolomics of the above-mentioned pathways in microalgae. Understanding the relation between anabolic and catabolic lipid enzyme pathways will provide new insights into biodiesel production and other valuable biomolecules obtained from microalgae.

Keywords

  • fatty acids
  • triacylglycerides
  • lipid metabolism
  • microalgae

1. Introduction

Despite the drop in crude oil prices over the last few years, global efforts to develop alternative renewable energy sources continue to be driven by increasing air pollution and growing energy consumption. Extensive research is therefore being conducted in the field of biofuels [1], which are derived from renewable biological sources. Biodiesel is the main substitute for diesel fuel and can be produced from both edible and non-edible oils. The use of edible oils has generated controversy because of the negative impact on food availability and the environment [2, 3]. As a consequence of these ethical considerations, non-food crops have emerged as a viable alternative for the production of biodiesel [46]. However, since non-food crops do not produce sufficient biomatter to feasibly cover the fuel requirements of the world’s transport sector, attention is turning to oleaginous microalgae which are able to produce and accumulate large amounts of fatty acids (FA) in the form of triacylglycerides (TAG) that can be converted into biodiesel through a transesterification reaction [2, 3, 7]. Furthermore, some species of oleaginous microalgae can also produce high-value products such as long-chain polyunsaturated fatty acids (docosahexaenoic (DHA) and eicosapentaenoic (EPA) acids), carbohydrates (cellulose, starch), proteins, and other high-value compounds, such as pigments, antioxidants (i.e., β-carotene, astaxanthin), and vitamins, which may have commercial application in various industrial sectors [2, 3, 8, 9]. In addition to their potential as biological factories, the advantage of these photosynthetic microorganisms is that their simple growing requirements (light, CO2, and nutrients) offer several environmental benefits such as high solar energy conversion efficiency, utilization of saline water, CO2 sequestration from the air and self-purification if coupled with wastewater treatment [10].

Despite the wide range of metabolites able to be synthesized by microalgae, little is known about the regulation of FA and TAG biosynthetic pathways and their storage and turnover in microalgae. In this chapter, we therefore describe recent advances in these fields and possible high-value co-products that could render the production of biodiesel from microalgae more sustainably. Recent studies on the transcriptomics, proteomics, and metabolomics of the above-mentioned pathways are also outlined. Understanding these metabolic pathways will accelerate the availability of biodiesel and other valuable biomolecules obtained from microalgae.

Advertisement

2. FA and TAG biosynthetic pathways in microalgae

Fatty acids are organic acids containing a carboxylic functional group with an aliphatic chain that can be saturated (SFA), monounsaturated (MUFA), or polyunsaturated (PUFA). The number of carbon atoms can vary, generating short-chain, medium-chain, or long-chain FA.

In plants, the FA biosynthetic pathway occurs in the chloroplasts (Figure 1).

Figure 1.

Simplified overview of the pathways involved in FA synthesis in plants. Enzyme abbreviations: ACCase, acetyl-CoA carboxylase; MCAT, malonyl-CoA:Acyl Carrier Protein (ACP) transacylase; KAS, ketoacyl-ACP synthases.

As shown in Figure 1, the first step in the pathway involves the acetyl-CoA carboxylase (ACCase) which catalyzes the formation of malonyl-CoA from acetyl-CoA and bicarbonate [11]. There is evidence suggesting the presence of genes encoding this enzyme (accA and accD) in Chlorella pyrenoidosa. In fact, the transcription of these genes showed to be up-regulated under lipid accumulating conditions [12]. Moreover, a marked increase in the level of acetyl-CoA together with a moderate augmentation of malonyl-CoA and CoA was detected in the green microalgae Chlorella desiccata, Dunaliella tertiolecta, and Chlamydomonas reinhardtii under stress conditions, denoting increased activity of ACCase in these strains [13].

The next step in the FA synthesis is mediated by the malonyl-CoA:Acyl Carrier Protein (ACP) transacylase (MCAT) which transfers the malonyl group from malonyl-CoA to malonyl-ACP [11]. A putative MCAT was identified as a part of the FA biosynthetic pathway in Nannochloropsis oceanica [14]. In Haematococcus pluvialis, the genes encoding ACP were up-regulated under TAG accumulating conditions (high temperature, high salinity, and nitrogen deficiency) together with other genes involved in FA biosynthesis [15]. In addition, proteomic studies on Neochloris oleoabundans revealed an augmented expression of ACP, among other enzymes of the lipid synthesis, under nitrogen starvation [16].

Acyl-ACP is the carbon source or substrate for the elongation of FA. This reaction is catalyzed by enzymes known as ketoacyl-ACP synthases (KASIII, KASI, and KASII). After each condensation, a reduction, dehydration, and second reduction occur. These steps are catalyzed by enzymes known as the FAS complex: beta-ketoacyl-ACP reductase (KAR), hydroxyacyl-ACP dehydrase (HAD), and enoyl-ACP reductase (EAR), respectively [11]. Transcriptome analysis of the diatom Chaetoceros sp. GSL56 helped to identify putative enzymes of the FA synthesis pathway. In addition, replacement of ketoacyl-ACP synthase of Synechococcus 7002 with Chaetoceros ketoacyl-ACP synthase III induced FA synthesis [17]. In line with this, TAG accumulating conditions increased the levels of transcripts for KAS in H. pluvialis [15].

The de novo resulting FA often with 16 or 18 carbon atoms can undergo the action of elongases and desaturases that add carbon or double bonds, respectively [11]. Particularly, desaturases and elongases are being intensively studied to achieve transgenic long-chain PUFA production [18, 19].

Some reports suggest the presence of both enzyme types in microalgae. In the marine microalgae Pavlova sp. and Isochrysis sp., two genes encoding elongases that catalyze the elongation of eicosapentaenoic acid (EPA) to docosahexaenoic acid (DHA) have been reported [20]. In the diatom Thalassiosira pseudonana, the genes encoding elongases that mediate the formation of DHA from EPA were successfully overexpressed, thus inducing an increase in DHA content [19]. A delta 5 desaturase was also identified, characterized and overexpressed in the diatom Phaeodactylum tricornutum inducing a significant increase in the unsaturated fatty acids [21].

Upon completion of elongation, FAs are transported to the cytoplasm to act as substrates of the acyl transferases involved in the TAG synthesis. TAG are neutral lipids formed by the esterification of one molecule of glycerol with three FAs. Because of their energy-rich acyl chains, they are the dominant form of stored energy in microalgae. Cellular stresses, such as nutrient deprivation (carbon dioxide, nitrogen, silica, and phosphorous), temperature fluctuation, or high light exposure trigger their formation [2228]. It has been demonstrated that lipid biosynthetic pathways are induced under these conditions to potentiate the lipid storage (30–60% of dry cell weight), and this mechanism is thought to play a role in microalgae adaptation and survival [24, 2939]. It has further been reported that multiple stressors have no additive effect on lipid accumulation [24, 40].

Data relating to plant FA and TAG metabolism provided the key to identifying possible molecular targets involved in lipid synthesis and accumulation in microalgae [41]. As shown in Figure 2, in plants, the first step of the conventional Kennedy pathway involves the acylation of the glycerol-3-phosphate (G-3-P), catalyzed by the glycerol-3-phosphate acyltransferase (GPAT) to yield lysophosphatidic acid (LPA). GPAT is the rate-limiting step subject to many regulatory controls at the transcriptional and post-transcriptional level and to allosteric mechanisms [42, 43]. Recent studies have revealed the presence of this enzyme in microalgae. In the marine diatom T. pseudonana, a membrane-bound GPAT designated TpGPAT was cloned and characterized. The authors observed that G-3-P was the preferred substrate of TpGPAT [44]. A sequence for GPAT with high homology to that of plants was found in C. reinhardtii, Volvox carteri, Ostreococcus lucimarinus, Ostreococcus tauri, Cyanidioschyzon merolae, and P. tricornutum. As in T. pseudonana, G-3-P and fatty acyl molecules are likely to be the enzyme substrates, as suggested by the residues present in their active sites [45].

Figure 2.

Simplified overview of the pathways involved in TAG synthesis in plants. Enzymes of the conventional Kennedy pathway involved in TAG synthesis and their subcellular localization in plants. Enzyme abbreviations: glycerol-3-phosphate acyltransferase (GPAT); lysophosphatidic acid acyltransferase (LPAAT); phosphatidic acid phosphohydrolase (PAP); diacylglycerol acyltransferase (DAGAT or DGAT). The same enzymes are involved in TAG synthesis in microalgae, but their intracellular localization has not yet been determined.

As described in Figure 2, lysophosphatidic acid acyltransferase (LPAAT) participates in the second step of the Kennedy pathway. This enzyme catalyzes the acylation of the LPA to yield phosphatidic acid (PA) [46]. Candidate LPAATs have been found in some algal genomes including that of H. pluvialis [47, 48], where it has been shown that LPAAT mRNA is induced under high irradiance stress [47]. In addition, it was recently reported that the expression of C. reinhardtii LPAAT (CrLPAAT1) is associated with an increase in lipid synthesis and accumulation under nitrogen starvation [48].

Phosphatidic acid phosphohydrolase (PAP) uses PA as substrate to form diacylglycerol (DAG), a precursor of TAG (Figure 2) [49]. In eukaryotes, PAP enzymes are the members of the evolutionarily conserved lipin protein family whose activity is related to TAG storage [50]. In the green microalga C. reinhardtii, PAP transcripts (named CrPAP2) are induced under stress conditions. In addition, CrPAP2 silencing slightly lowers the lipid content. Thus, in C. reinhardtii, as in other eukaryotes, PAP expression is related to lipid synthesis and accumulation [49].

The last enzyme of the de novo TAG synthesis is acyl-CoA:diacylglycerol acyltransferase (DGAT), which catalyzes the acylation of DAG to yield TAG (Figure 2) [51]. This enzyme employs DAG and acyl-CoA as substrates, so the resulting TAG is formed through an acyl-CoA-dependent pathway [46] and is a key target to increase TAG synthesis and storage through genetic manipulation [52, 53]. In higher plants, three different types of DGATs participate in the formation of TAG: DGAT1, DGAT2, or DGAT3 [54]. Sequences for DGAT1 and DGAT2 isoforms were found in several algal strains [55]. Sequences for DGAT2, but not DAGT1, or DGAT3, were identified in the green microalga O. tauri [56]. DGAT2 was also found in T. pseudonana (TpDGAT2). In addition, the expression of DGAT in a TAG-null yeast mutant restored the synthesis of these neutral lipids [57]. In the oleaginous microalga C. pyrenoidosa grown under stress conditions, a high correlation was found between DGAT and TAG accumulation [58]. Also in N. oceanica IMET1, another oleaginous microalga, seven putative DGAT genes were up-regulated under nitrogen-deficient conditions, when the synthesis of TAG-neutral lipids was significantly increased [59]. In C. reinhardtii dgat1 and dgtt1 to dgtt5 genes encode for DGAT1 and DGAT2, respectively [60, 61]. Increased transcript expression of the genes dgat1 and dgtt1 was detected under stress conditions (less sulfur, phosphorous, iron, zinc, or nitrogen). Once more, the evidence suggests that both DGAT1 and DGAT2 could play a role in TAG synthesis as their expression is induced under TAG-accumulating conditions [62, 63]. In support of this hypothesis, overexpression of a DGAT2 isoform in the marine diatom P. tricornutum stimulated the synthesis of neutral lipids and their accumulation in lipid droplets [64].

As can be observed, much research has focused on the acyl-CoA-dependent reaction catalyzed by DGAT. However, the relative contribution of DGAT1 and DGAT2 isoenzymes to TAG accumulation appears to be species-dependent, so further studies should be performed to gain insight into this aspect.

TAG can be formed by the acyl-CoA-dependent pathway, detailed previously, or through acyl-CoA-independent reactions. Acyl-CoA-independent formation of TAG is mediated by the activities of two types of enzyme: the phospholipid:diacylglycerol acyltransferases (PDAT), which catalyze the formation of TAG using DAG and phosphatidylcholine (PC); and the DAG:DAG transacylases (DGTA) which utilize two molecules of DAG to form TAG and MAG [54, 65].

In fact, in N. oceanica IMET1, it was reported that membrane polar lipids were converted into TAG when the microalgae were grown under nitrogen deficiency [59]. In agreement with this, the gene encoding the acyltransferase PDAT1 was induced under nitrogen starvation in C. reinhardtii. Moreover, TAG content in the C. reinhardtii PDAT-null mutant was 25% lower than in the parent strain. It would thus appear that PDAT has a relevant role in TAG accumulation, stimulating the transacylation pathway in both strains [62]. Furthermore, in C. reinhardtii it was suggested that PDAT functions as a DGTA with acyl hydrolase activity. PDAT might, therefore, mediate membrane polar lipid turnover in a favorable environment whereas under stress conditions it may participate in phospholipid degradation contributing to TAG synthesis [66].

As already mentioned, many aspects of C. reinhartii lipid metabolism have already been characterized, making it the microalga of choice for current purposes [23, 6773]. Nevertheless, Chlamydomonas is a non-oleaginous strain [23]. Other microalgal species with greater potential to yield biodiesel and other high-value products should therefore be more thoroughly investigated.

Advertisement

3. Transcriptomics, proteomics, and metabolomics

A better understanding of the mechanisms involved in TAG enrichment under stress conditions will help to maximize microalgae productivity. However, many biochemical approaches for elucidating molecular pathways depend on the availability of genomic sequence data [29]. Transcriptomics, proteomics, and metabolomics, however, are able to provide a detailed description of cell transcripts (RNA), proteins and metabolites, respectively while completely bypassing the requirement of genomic information [74, 75].

Transcriptome analysis helped to identify sequences of the enzymes involved in the biosynthesis and catabolism of FA, TAG, and starch in D. tertiolecta, revealing that this strain shares genetic information, at least in terms of the mentioned pathways, with closely related microalgae species such as V. carteri and C. reinhardtii [76]. The transcriptome of N. oleoabundans was also determined. In this case, the authors quantified the differences between nitrogen-replete and nitrogen-limiting culture conditions. Under nitrogen deficiency, N. oleoabundans showed higher levels of transcripts of FA and TAG synthesis pathways and inhibition of the FA β-oxidation pathway, compared to nitrogen-replete culture conditions [29]. In agreement with this finding, in C. vulgaris, transcriptomic [31] and proteomic [77] studies revealed an induction of the enzymes of the FA and TAG synthesis machinery under lipid enrichment conditions. Also, transcription factors associated with these metabolic pathways were augmented under the stress condition [77].

The transcriptome of C. reinhardtii showed that genes involved in FA and TAG metabolic pathways and in membrane remodeling were highly induced under neutral lipid accumulation conditions [78]. In this microalga, proteomic studies revealed an augmented rate of lipid synthesis machinery with a concomitant enhancement in FA and TAG; higher levels of starch than under non-stress conditions were also detected by metabolomic analyses. Metabolic pathways such as nitrogen assimilation, amino acid metabolism, oxidative phosphorylation, glycolysis, TCA cycle, and the Calvin cycle suffered adjustments during C. reinhardtii [79, 80].

As in C. vulgaris, nutrient-deprivation stress in C. reinhardtii, D. tertiolecta, and N. oleoabundans induced the expression of genes involved in FA and TAG synthesis pathways in P. tricornutum [81], Chlorella protothecoides [82], and Tisochrysis lutea [83].

In conclusion, these assembled transcriptomes, proteomes, and metabolomes offer valuable approaches for improving microalgal productivity, providing possible targets for molecular engineering that could enhance microalgae-derived products.

Advertisement

4. Molecular targets for enhancing lipid biosynthesis

Genetic strain modification to improve microalgal productivity and accelerate the industrialization of algal-derived products is a major challenge [84]. Reflecting the fact that enhancement of the FA synthesis pathway had little effect on total lipid content in some plants [85, 86], a growing body of research now focuses on overexpression of the enzymes or heterologous expression of genes involved in the TAG biosynthetic pathway. Table 1 provides an outline of some of the genetic manipulations performed on several microalgal strains, leading to an improvement in their TAG content.

Enzymes overexpressed or heterologously expressedOrganismEffect on lipid production (changes over control condition)References
ME of Phaeodactylum tricornutumChlorella pyrenoidosa18.7%[87]
GPAT of Thalassiosira pseudonanaYeast GPAT-deficient mutantRestored TAG formation[44]
GPAT of Lobosphaera incisaChlamydomonas reinhardtii50%[88]
GPAT of Phaeodactylum tricornutumPhaeodactylum tricornutum2-fold[89]
G3PDH, GPAT, DGAT, LPAAT and PAP of Saccharomyces cerevisiae and Yarrowia lipolyticaChlorella minutissima2-fold[90]
LPAAT of Chlamydomonas reinhardtiiChlamydomonas reinhardtii20%[91]
DGAT2 of Nannochloropsis oceanicaNannochloropsis oceanica3.5-fold[92]
DGAT1 and DGAT2 of Myrmecia incisaS. cerevisiae lipid deficient mutantRe-stored TAG formation[93]
DGAT2 of Neochloris oleoabundansS. cerevisiae DGAT deficient mutantRe-stored TAG formation[94]
DGAT 1 of Phaeodactylum tricornutumS. cerevisiae DGAT deficient mutantRe-stored TAG synthesis and lipid body formation[63]
DGAT 2 of Phaeodactylum tricornutumPhaeodactylum tricornutum35%[64]
Various DGAT type 2Chlamydomonas reinhardtii20–44%[95]

Table 1.

Some of the genetic modifications performed on metabolic pathways related to lipid synthesis in microalgae and their effect on lipid enrichment.

Enzyme abbreviations: ME, malic enzyme; DGAT, diacylglycerol acyltransferase; G3PDH, glycerol-3-phosphate-dehydrogenase; GPAT, glycerol-3-phosphate acyltransferase; LPAAT, lysophosphatidic acid acyltransferase; PAP, phosphatidic acid phosphatase.

Advertisement

5. TAG-accumulation in lipid droplets

Lipid droplets (LDs) are cell organelles that are currently the subject of in-depth study in various organisms. These lipid globules not only act as a reservoir of cell carbon and energy, they may also have a role in lipid homeostasis, signaling, trafficking, and interorganelle communications [96, 97]. As previously mentioned, under stress conditions microalgae synthesize TAG and store them as cytoplasmic LDs [2228], which can vary in size, shape, and function depending on the cell type and the environmental conditions (Figure 3) [98]. In eukaryotic cells, LD structure consists of a TAG-rich hydrophobic core surrounded by surface polar glycerolipids into which proteins of the perilipin (Plin) (animal cells) or oleosin and caleosin (plants) families are embedded [99102]. In microalgae, LD structure is conserved from eukaryotes but different LD proteins have been identified. The analysis of C. reinhardtii LDs recognized 16 proteins related to lipid metabolism and a major lipid droplet protein (named MLDP) was identified. MLDP silencing increased the size of the LD, without modifying LD TAG content [68]. In the green microalga, Nannochloropsis sp., a hydrophobic lipid droplet surface protein, named LDSP, was identified. The expression of LDSP increased concomitantly with TAG content under oil-accumulating conditions [99]. In H. pluvialis, seven proteins were found to be associated with LDs. The most abundant of these, Haematococcus Oil Globule Protein (HOGP), was homologous to the MLDP of C. reinhardtii and its expression was induced under TAG accumulating conditions [103]. LD-associated proteins may also help in the accumulation of TAG in the green microalga Myrmecia incisa [104]. Moreover, LDs from C. reinhardtii showed the presence of enzymes involved in TAG synthesis (GPAT, and PDAT) and in sterol synthesis, lipid signaling, and trafficking [69]. Further in-depth research should be able to determine the proteins associated with LDs and their role in TAG metabolism in microalgae.

Figure 3.

Schematic representation of a cytoplasmic lipid droplet (LD) from microalgae.

In the oleaginous diatom Fistulifera sp., two proteins located in the oil bodies were also detected in the endoplasmic reticulum (ER), suggesting that oil bodies might originate in the ER [105]. The same authors found a signal sequence typical of ER localization in an LD protein called diatom-oleosome-associated-protein 1 (DOAP1) in Fistulifera solaris JPCC DA0580 [106]. Related to these findings, the induction of ER stress leads to LD formation in C. reinhardtii and C. vulgaris [107]. In addition, LDs from C. reinhardtii were associated not only with the ER membrane but also with the outer membrane of the chloroplasts [108]. Available data therefore suggest that in microalgae, cytoplasmic LDs are produced in the ER. However, additional studies are required to arrive at a better understanding of the mechanism of LD formation in the ER, and to determine whether chloroplasts play a role in this process.

Advertisement

6. TAG degradation pathways in microalgae

As previously mentioned, the economic feasibility of using microalgae as a source of FA for biodiesel depends to a great extent on improvements in the production process, one of the most significant challenges being to increase lipid yields. The selection of oleaginous strains and the search for different culture strategies to increase lipid biosynthesis constitute viable approaches; blocking the competing pathways of carbohydrate formation may be another. However, both the approaches give rise to a decrease in strain growth [22]. Lipid catabolism has largely been ignored as a relevant pathway for engineering, despite being a competing pathway to lipid biogenesis [109]. However, lipases were identified in C. reinhardtii [66, 72, 73] and T. pseudonana [110]. In the case of C. reinhardtii, CrLIP1 could restore the lipase activity in a Saccharomyces cerevisiae lipase-null strain. In addition, C. reinhardtii TAG content decreased with increasing expression of CrLIP1 under stress conditions, hydrolyzing mainly DAG and polar lipids [72]. In agreement with this, a galactoglycerolipid lipase was found in C. reinhardtii. The main substrates of the enzyme are galactoglycerolipids and the main products are FAs employed for TAG synthesis [74]. In C. reinhardtii, phospholipid:diacylglycerol acyltransferase (PDAT) demonstrated both transacylation and acyl hydrolase activities, and could mediate membrane lipid turnover and TAG synthesis [66]. The activity of a multifunctional lipase/phospholipase/acyltransferase of T. pseudonana lowered lipid content under both normal and stress conditions [110]. A single gene for PDAT was identified in H. pluvialis, though no functional analysis was performed for the gene in this strain [47]. Further studies are required to gain insight into the molecular mechanisms involved in TAG degradation, which could be the key to increased lipid yields in microalgae.

Advertisement

7. Microalgae-based biorefineries

In the context of improving the economic feasibility of microalgae-based biodiesel, a closer look should be taken at the large amounts of TAG produced in some oleaginous microalgae alongside high-value products such as carbohydrates (cellulose and starch); proteins and other high-value compounds like pigments, antioxidants (i.e., β-carotene, astaxanthin), and vitamins [2, 3, 8, 9], all of which may have commercial application in different industrial sectors. Some potentially high-value products found in microalgae are described in Table 2.

ProductMicroalgal strainApplicationsReferences
Carbohydrates
ExopolysaccharidesNavicula cinctaPharmaceutics and agronomics[111]
Starch, glucose, cellulloseChlorella vulgaris FSP-EBioethanol production[112, 113]
Sulfated extracellular polisaccharidesGraesiella sp.Pharmaceutics[114]
Lipids
Phytosterols; linoleic (C18:2n6) and alpha linolenic (C18:3n3) fatty acidsHaematococcus pluvialisHuman dietary supplement, nutraceutics[115]
PhytosterolsDunaliella tertiolecta, D. salinaNutraceutics[116]
Omega-3 long chain-PUFAIsochrysis, Nannochloropsis, Phaeodactylum, Pavlova, and ThalassiosiraFunctional food[117]
Proteins
ProteinsChlorella pyrenoidosaNutraceutics[118]
ProteinsChlorella vulgaris, Nannochloris bacillaris, Tetracystis sp., Micractinium reisseriAnimal feed[119]
Vitamins
TocopherolNannochloropsis oculata, Tetraselmis suecicaHuman dietary supplement, nutraceutics[120]
Tocopherol, pigments, phenolic compoundsDesmodesmus sp.Human dietary supplement, nutraceutics[121]
Pigments
AstaxanthinHaematococcus pluvialisAntioxidant, cosmetics, pharmaceutics[122]
LuteinDunaliella salinaFunctional food, animal feed[123]
CarotenoidsPhaeodactylum tricornutumCosmetics, pharmaceutics, animal feed[124]
CarotenoidsDunaliella salinaCosmetics, pharmaceutics, animal feed[125]
Others
SilicaNavicula cinctaAbrassive products, insecticides[111]

Table 2.

Recent advances in microalgal-derived high-value products.

Advertisement

8. Conclusion

Oleaginous microalgae grown under stress conditions can synthesize and accumulate large quantities of FA, mainly in the form of TAG, which can then be converted into biodiesel. Although microalgae constitute a promising source of clean energy, knowledge gaps continue to abound in almost all aspects of FA and TAG metabolism for these microorganisms, including the precise identity of enzymatic machinery, the relative contributions of each enzyme and their precise regulation. Further studies are therefore required to establish the exact metabolic pathways involved in FA and TAG synthesis, accumulation, and degradation in order to develop genetic engineering strategies to obtain microalgal strains with improved capacity to convert their biomass into TAG and other valuable co-products.

Advertisement

Acknowledgments

The authors are grateful for research funds provided by the Consejo Nacional de Investigaciones Científicas y Técnicas de la República Argentina (CONICET); Agencia Nacional de Promoción Científica y Tecnológica, PICTs 2014-0893, 2013-0987, and 2015-0800; and the Secretaría de Ciencia y Tecnología de la Universidad Nacional del Sur, PGIs 24/B226 and 24/B196. Paola Scodelaro Bilbao, Gabriela Salvador and Patricia Leonardi are Research Members of CONICET.

References

  1. 1. Misra N, Kumar Panda P, Kumar Parida B, Kanta Mishra B. Phylogenomic study of lipid genes involved in microalgal biofuel production: candidate gene mining and metabolic pathway analyses. Evol Bioinform Online. 2012; 8: 545-564. doi: 10.4137/EBO.S10159.
  2. 2. Brennan L, Owende P. Biofuels from microalgae: a review of technologies for production, processing, and extractions of biofuels and co-products. Renew Sust Energy Rev. 2010; 14: 557-577. doi: 10.1016/j.rser.2009.10.009.
  3. 3. Mata TM, Martinsa AA, Caetano NS. Microalgae for biodiesel production and other applications: a review. Renew Sust Energy Rev. 2012; 14: 217-232. doi: 10.1016/j.rser.2009.07.020.
  4. 4. Foild N, Foild G, Sánchez M, Mittelbach M, Hackel S. J. curcas L. as a source for the production of biofuel in Nicaragua. Bioresour Technol. 1996; 58: 77-82.
  5. 5. Rashid U, Anwar F, Moser BR, Knothe G. Moringa oleifera oil: a possible source of biodiesel. Bioresour Technol. 2008; 99: 8175-8179. doi: 10.1016/j.biortech.2008.03.066.
  6. 6. Ahmad M, Zafar M, Azam A, Sadia H, Khan MA, Sultana S. Techno-economic aspects of biodiesel production and characterization. Energy sources. 2011; 6: 166-177.
  7. 7. Yu WL, Ansari W, Schoepp NG, Hannon MJ, Mayfield SP, Burkart MD. Modifications of the metabolic pathways of lipid and triacylglycerol production in microalgae. Microb Cell Fact. 2011; 10: 91. doi: 10.1186/1475-2859-10-91.
  8. 8. Williams PJ le B, Laurens LML. Microalgae as biodiesel and biomass feedstocks: review and analysis of the biochemistry, energetics and economics. Energy Environ. Sci. 2010; 3: 554-590. doi: 10.1039/B924978H.
  9. 9. Borowitzka MA. High-value products from microalgae—their development and commercialization. J Appl Phycol. 2013; 25: 743-756. doi:10.1007/s10811-013-9983-9.
  10. 10. Lam MK, Lee KT. Microalgae biofuels: a critical review of issues, problems and the way forward. Biotechnol Adv. 2012; 30: 673-690. doi: 10.1016/j.biotechadv.2011.11.008.
  11. 11. Brown AP, Slabas AR, Rafferty JB. Fatty acid biosynthesis in plants-metabolic pathways, structure and organization. Lipids in Photosynthesis. 2010; 30: 11-34. doi: 10.1007/978-90-481-2863-1_2.
  12. 12. Fan J, Cui Y, Wan M, Wang W, Li Y. Lipid accumulation and biosynthesis genes response of the oleaginous Chlorella pyrenoidosa under three nutrition stressors. Biotech Biofuels. 2014; 7: 17. doi: 10.1186/1754-6834-7-17.
  13. 13. Avidan O, Brandis A, Rogachev I, Pick U. Enhanced acetyl-CoA production is associated with increased triglyceride accumulation in the green alga Chlorella desiccate. J Exp Botany. 2015; 66: 1-11. doi:10.1093/jxb/erv166.
  14. 14. Chen JW, Liu W-J, Hu D-X, Wang X, Balamurugan S, Alimujiang A, Yang W-D, Liu J-S, Li H-Y. Identification of a malonyl CoA-acyl carrier protein transacylase and its regulatory role in fatty acid biosynthesis in oleaginous microalga Nannochloropsis oceanica. Biotech Appl Biochem. 2016. doi: 10.1002/bab.1531.
  15. 15. Lei A, Chen H, Shen G, Hu Z, Chen L, Wang J. Expression of fatty acid synthesis genes and fatty acid accumulation in Haematococcus pluvialis under different stressors. Biotechnol Biofuels. 2012; 5: 18. doi: 10.1186/1754-6834-5-18.
  16. 16. Morales-Sánchez D, Kyndtb J, Ogdenc K, Martinez A. Toward an understanding of lipid and starch accumulation in microalgae: a proteomic study of Neochloris oleoabundans cultivated under N-limited heterotrophic conditions. Algal Res. 2016; 20: 22-34. doi: 10.1016/j.algal.2016.09.006.
  17. 17. Gu H, Jinkerson RE, Davies FK, Sisson LA, Schneider PE, Posewitz MC. Modulation of medium-chain fatty acid synthesis in Synechococcus sp. PCC 7002 by replacing FabH with a Chaetoceros Ketoacyl-ACP synthase. Front Plant Sci. 2016; 7: 690. doi: 10.3389/fpls.2016.00690.
  18. 18. Peng KT, Zheng CN, Xue J, Chen XY, Yang WD, Liu JS, Bai W, Li HY. Delta 5 fatty acid desaturase upregulates the synthesis of polyunsaturated fatty acids in the marine diatom Phaeodactylum tricornutum. J Agric Food Chem. 2014; 62: 8773-8776. doi: 10.1021/jf5031086.
  19. 19. Petrie JR, Liu Q, Mackenzie AM, Shrestha P, Mansour MP, Robert SS, Frampton DF, Blackburn SI, Nichols PD, Singh SP. Isolation and characterisation of a high-efficiency desaturase and elongases from microalgae for transgenic LC-PUFA production. Mar Biotechnol. 2010; 12: 430-438. doi: 10.1007/s10126-009-9230-1.
  20. 20. Pereira SL, Leonard AE, Huang YS, Chuang LT, Mukerji P. Identification of two novel microalgal enzymes involved in the conversion of the omega3-fatty acid, eicosapentaenoic acid, into docosahexaenoic acid. Biochem J. 2004; 384: 357-366.
  21. 21. Cook O, Hildebrand M. Enhancing LC-PUFA production in Thalassiosira pseudonana by overexpressing the endogenous fatty acid elongase genes. J Appl Phycol. 2016; 28: 897-905.
  22. 22. Hu Q, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert M, Darzins A. Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances. Plant J. 2008; 54: 621-639. doi: 10.1111/j.1365-313X.2008.03492.x.
  23. 23. Liu B, Benning C. Lipid metabolism in microalgae distinguishes itself. Curr Opin Biotechnol. 2013; 24: 300-309. doi: 10.1016/j.copbio.2012.08.008.
  24. 24. Damiani MC, Popovich CA, Constenla D, Leonardi PI. Lipid analysis in Haematococcus pluvialis to assess its potential use as a biodiesel feedstock. Bioresour Technol. 2010; 101: 3801-3807. doi: 10.1016/j.biortech.2009.12.136.
  25. 25. Popovich CA, Damiani C, Constenla D, Leonardi PI. Lipid quality of Skeletonema costatum and Navicula gregaria from South Atlantic Coast (Argentina): evaluation of its suitability as biodiesel feedstock. J Appl Phycol. 2012; 24: 1-10. doi:10.1007/s10811-010-9639-y.
  26. 26. Popovich CA, Damiani MC, Constenla D, Martínez AM, Freije H, Giovanardi M, Pancaldi S, Leonardi PI. Neochloris oleoabundans grown in enriched natural seawater for biodiesel feedstock: evaluation of its growth and biochemical composition. Bioresour Technol. 2012; 114: 287-293. doi: 10.1016/j.biortech.2012.02.121.
  27. 27. Bongiovani N, Popovich C, Martínez A, Freije H, Constenla D, Leonardi PI. In vivo measurements to estimate culture status and neutral lipid accumulation in Nannochloropsis oculata CCALA 978: implications for biodiesel oil studies. Algological Studies. 2013; 142: 3-16 doi: 10.1127/1864-1318/2013/0104.
  28. 28. Martín L, Popovich C, Martinez A, Damiani C, Leonardi PI. Oil assessment of Halamphora coffeaeformis diatom growing in a hybrid two-stage system for biodiesel production. Renewable Energy. 2016; 92: 127-135. doi: 10.1016/j.renene.2016.01.078
  29. 29. Rismani-Yazdi H, Haznedaroglu BZ, Hsin C, Peccia J. Transcriptomic analysis of the oleaginous microalga Neochloris oleoabundans reveals metabolic insights into triacylglyceride accumulation. Biotechnol Biofuels. 2012; 5: 74. doi: 10.1186/1754-6834-5-74.
  30. 30. Yu ET, Zendejas FJ, Lane PD, Gaucher S, Simmons BA, Lane TW. Triacylglycerol accumulation and profiling in the model diatoms Thalassiosira pseudonana and Phaeodactylum tricornutum (Bacillariophyceae) during starvation. J Appl Phycol. 2009; 21: 669-681. doi: 10.1007/s10811-008-9400-y.
  31. 31. Guarnieri MT, Nag A, Smolinski SL, Darzins A, Seibert M, et al. Examination of triacylglycerol biosynthetic pathways via de novo transcriptomic and proteomic analyses in an unsequenced microalga. PLoS One. 2011; 6: e25851. doi: 10.1371/journal.pone.0025851.
  32. 32. Pal D, Khozin-Goldberg I, Cohen Z, Boussiba S. The effect of light, salinity, and nitrogen availability on lipid production by Nannochloropsis sp. Appl Microbiol Biotechnol. 2011; 90: 1429-1441. doi:10.1007/s00253-011-3170-1.
  33. 33. Tang H, Chen M, Garcia ME, Abunasser N, Ng KY, Salley SO. Culture of microalgae Chlorella minutissima for biodiesel feedstock production. Biotechnol Bioeng. 2011; 108: 2280-2287. doi: 10.1002/bit.23160.
  34. 34. Breuer G, Lamers PP, Martens DE, Draaisma RB, Wijffels RH. Effect of light intensity, pH, and temperature on triacylglycerol (TAG) accumulation induced by nitrogen starvation in Scenedesmus obliquus. Bioresour Technol. 2013; 143: 1-9. doi: 10.1016/j.biortech.2013.05.105.
  35. 35. Gong Y, Guo X, Wan X, Liang Z, Jiang M. Triacylglycerol accumulation and change in fatty acid content of four marine oleaginous microalgae under nutrient limitation and at different culture ages. J Basic Microbiol. 2013; 53: 29-36. doi: 10.1002/jobm.201100487.
  36. 36. Yu SY, Li H, Tong M, Ouyang LL, Zhou ZG. Identification of a Δ6 fatty acid elongase gene for arachidonic acid biosynthesis localized to the endoplasmic reticulum in the green microalga Myrmecia incisa Reisigl. Gene. 2012; 493: 219-227. doi: 10.1016/j.gene.2011.11.053.
  37. 37. Aguirre AM, Bassi A. Investigation of biomass concentration, lipid production, and cellulose content in Chlorella vulgaris cultures using response surface methodology. Biotechnol Bioeng. 2013; 110: 2114-2122. doi: 10.1002/bit.24871.
  38. 38. Simionato D, Block MA, La Rocca N, Jouhet J, Maréchal E, Finazzi G, Morosinotto T. The response of Nannochloropsis gaditana to nitrogen starvation includes de novo biosynthesis of triacylglycerols, a decrease of chloroplast galactolipids, and reorganization of the photosynthetic apparatus. Eukaryot Cell. 2013; 12: 665-676. doi: 10.1128/EC.00363-12.
  39. 39. La Russa M, Bogen C, Uhmeyer A, Doebbe A, Filippone E, Kruse O, Mussgnug JH. Functional analysis of three type-2 DGAT homologue genes for triacylglycerol production in the green microalga Chlamydomonas reinhardtii. J Biotechnol. 2012; 162: 13-20. doi: 10.1016/j.jbiotec.2012.04.006.
  40. 40. Roleda MY, Slocombe SP, Leakey RJ, Day JG, Bell EM, Stanley MS. Effects of temperature and nutrient regimes on biomass and lipid production by six oleaginous microalgae in batch culture employing a two-phase cultivation strategy. Bioresour Technol. 2013; 129: 439-449. doi: 10.1016/j.biortech.2012.11.043.
  41. 41. Adarme-Vega TC, Lim DKY, Timmins M, Vernen F, Li Y, Schenk PM. Microalgal biofactories: a promising approach towards sustainable omega-3 fatty acid production. Microb Cell Fact. 2012; 11: 96. doi: 10.1186/1475-2859-11-96.
  42. 42. Coleman RA, Lee DP. Enzymes of triacylglycerol synthesis and their regulation. Prog Lipid Res. 2004; 43: 134-176. doi: 10.1016/S0163-7827(03)00051-1.
  43. 43. Courchesne NMD, Parisien A, Wang B, Lan CQ. Enhancement of lipid production using biochemical, genetic and transcription factor engineering approaches. J Biotechnol. 2009; 141: 31-41. doi: 10.1016/j.jbiotec.2009.02.018.
  44. 44. Xu J, Zheng Z, Zou J. A membrane-bound glycerol-3-phosphate acyltransferase from Thalassiosira pseudonana regulates acyl composition of glycerolipids. Botany. 2009; 87: 544-551. doi: 10.1139/B08-145.
  45. 45. Misra N, Panda PK, Parida BK. Agrigenomics for microalgal biofuel production: an overview of various bioinformatics resources and recent studies to link OMICS to bioenergy and bioeconomy. OMICS. 2013; 17: 537-549. doi: 10.1089/omi.2013.0025.
  46. 46. Chapman KD, Ohlrogge JB. Compartmentation of triacylglycerol accumulation in plants. J Biol Chem. 2012; 287: 2288-2294. doi: 10.1074/jbc.R111.290072.
  47. 47. Gwak Y, Hwang YS, Wang B, Kim M, Jeong J, Lee CG, Hu Q, Han D, Jin E. Comparative analyses of lipidomes and transcriptomes reveal a concerted action of multiple defensive systems against photooxidative stress in Haematococcus pluvialis. J Exp Bot. 2014; 65: 4317-4334. doi: 10.1093/jxb/eru206.
  48. 48. Yamaoka Y, Achard D, Jang S, Legéret B, Kamisuki S, Ko D, Schulz-Raffelt M, Kim Y, Song WY, Nishida I, Li-Beisson Y, Lee Y. Identification of a Chlamydomonas plastidial 2-lysophosphatidic acid acyltransferase and its use to engineer microalgae with increased oil content. Plant Biotechnol J. 2016; 14: 2158-2167. doi: 10.1111/pbi.12572.
  49. 49. Deng XD, Cai JJ, Fei XW. Involvement of phosphatidate phosphatase in the biosynthesis of triacylglycerols in Chlamydomonas reinhardtii. J Zhejiang Univ Sci B. 2013; 14: 1121-1131. doi: 10.1631/jzus.B1300180.
  50. 50. Csaki LS, Dwyer JR, Fong LG, Tontonoz P, Young SG, Reue K. Lipins, lipinopathies, and the modulation of cellular lipid storage and signaling. Progr Lipid Res. 2013; 52: 305-316. doi: 10.1016/j.plipres.2013.04.001.
  51. 51. Radakovits R, Jinkerson RE, Darzins A, Posewitz MC. Genetic engineering of algae for enhanced biofuel production. Eukaryot Cell. 2010; 9: 486-501. doi: 10.1128/EC.00364-09.
  52. 52. Taylor DC, Katavic V, Zou J, MacKenzie SL, Keller WA, An J, Friesen W, Barton DL, Pedersen KK, Giblin EM, et al. Field testing of transgenic rapeseed cv. Hero transformed with a yeast sn-2 acyltransferase results in increased oil content, erucic acid content and seed yield. Mol Breed. 2002; 8: 317-322. DOI: 10.1023/A:1015234401080.
  53. 53. Lardizabal KD, Thompson GA, Hawkins D. Diacylglycerol acyl transferase proteins. In Official Gazette of the United States Patent and Trademark Office Patents Edited by: Office TUSPaT. USA, 2006.
  54. 54. Lung SC, Weselake RJ. Diacylglycerol acyltransferase: a key mediator of plant triacylglycerol synthesis. Lipids. 2006; 41: 1073-1088.
  55. 55. Chen JE, Smith AG. A look at diacylglycerol acyltransferases (DGATs) in algae. J Biotechnol. 2012; 162: 28-39. doi: 10.1016/j.jbiotec.2012.05.009.
  56. 56. Wagner M, Hoppe K, Czabany T, Heilmann M, Daumb G, Feussner I, Fulda M. Identification and characterization of an acyl-CoA:diacylglycerol acyltransferase 2 (DGAT2) gene from the microalga O. tauri. Plant Physiol Biochem. 2010; 48: 407-416. doi: 10.1016/j.plaphy.2010.03.008.
  57. 57. Xu J, Kazachkov M, Jia Y, Zheng Z, Zou J. Expression of a type 2 diacylglycerol acyltransferase from Thalassiosira pseudonana in yeast leads to incorporation of docosahexaenoic acid β-oxidation intermediates into triacylglycerol. FEBS J. 2013; 280: 6162-6172. doi: 10.1111/febs.12537.
  58. 58. Fan J, Cui Y, Wan M, Wang W, Li Y. Lipid accumulation and biosynthesis genes response of the oleaginous Chlorella pyrenoidosa under three nutrition stressors. Biotechnol Biofuels. 2014; 7: 17. doi: 10.1186/1754-6834-7-17.
  59. 59. Xiao Y, Zhang J, Cui J, Feng Y, Cui Q. Metabolic profiles of Nannochloropsis oceanica IMET1 under nitrogen-deficiency stress. Bioresour Technol. 2013; 130: 731-738. doi: 10.1016/j.biortech.2012.11.116.
  60. 60. Miller R, Wu G, Deshpande RR, Vieler A, Gärtner K, Li X, Moellering ER, Zäuner S, Cornish AJ, Liu B, Bullard B, Sears BB, Kuo M-H, Hegg EL, Shachar-Hill Y, Shiu S-H, Benning C. Changes in transcript abundance in Chlamydomonas reinhardtii following nitrogen deprivation predict diversion of metabolism. Plant Physiol. 2010; 154: 1737-1752. doi: 10.1104/pp.110.165159.
  61. 61. Msanne J, Xu D, Konda AR, Casas-Mollano JA, Awada T, Cahoon EB, Cerutti H. Metabolic and gene expression changes triggered by nitrogen deprivation in the photoautotrophically grown microalgae Chlamydomonas reinhardtii and Coccomyxa sp. C-169. Phytochemistry. 2012; 75: 50-59. doi: 10.1016/j.phytochem.2011.12.007.
  62. 62. Boyle NR, Page MD, Liu B, Blaby IK, Casero D, Kropat J, Cokus SJ, Hong-Hermesdorf A, Shaw J, Karpowicz SJ, Gallaher SD, Johnson S, Benning C, Pellegrini M, Grossman A, Merchant SS. Three acyltransferases and nitrogen-responsive regulator are implicated in nitrogen starvation-induced triacylglycerol accumulation in Chlamydomonas. J Biol Chem. 2012; 287: 15811-1525. doi: 10.1074/jbc.M111.334052.
  63. 63. Guihéneuf F, Leu S, Zarka A, Khozin-Goldberg I, Khalilov I, Boussiba S. Cloning and molecular characterization of a novel acyl-CoA:diacylglycerol acyltransferase 1-like gene (PtDGAT1) from the diatom Phaeodactylum tricornutum. FEBS J. 2011: 278: 3651-3666. doi: 10.1111/j.1742-4658.2011.08284.x.
  64. 64. Niu YF, Zhang MH, Li DW, Yang WD, Liu JS, Bai WB, Li HY. Improvement of neutral lipid and polyunsaturated fatty acid biosynthesis by overexpressing a type 2 diacylglycerol acyltransferase in marine diatom Phaeodactylum tricornutum. Mar Drugs. 2013; 11: 4558-4569. doi: 10.3390/md11114558.
  65. 65. Dahlqvist A, Stahl U, Lenman M, Banas A, Lee M, Sandager L, Ronne H, Stymne S. Phospholipid:diacylglycerol acyltransferase: an enzyme that catalyzes the acyl-CoA-independent formation of triacylglycerol in yeast and plants. Proc Natl Acad Sci USA. 2000; 97: 6487-6492. doi: 10.1073/pnas.120067297.
  66. 66. Yoon K, Han D, Li Y, Sommerfeld M, Hu Q. Phospholipid:diacylglycerol acyltransferase is a multifunctional enzyme involved in membrane lipid turnover and degradation while synthesizing triacylglycerol in the unicellular green microalga Chlamydomonas reinhardtii. Plant Cell. 2012; 24: 3708-3724. doi: 10.1105/tpc.112.100701.
  67. 67. La Russa M, Bogen C, Uhmeyer A, Doebbe A, Filippone E, Kruse O, Mussgnug JH. Functional analysis of three type-2 DGAT homologue genes for triacylglycerol production in the green microalga Chlamydomonas reinhardtii. J Biotechnol. 2012; 162: 13-20. doi: 10.1016/j.jbiotec.2012.04.006.
  68. 68. Moellering ER, Benning C. RNA interference silencing of a major lipid droplet protein affects lipid droplet size in Chlamydomonas reinhardtii. Eukaryot Cell. 2010; 9: 97-106. doi: 10.1128/EC.00203-09.
  69. 69. Nguyen HM, Baudet M, Cuiné S, Adriano JM, Barthe D, Billon E, Bruley C, Beisson F, Peltier G, Ferro M, Li-Beisson Y. Proteomic profiling of oil bodies isolated from the unicellular green microalga Chlamydomonas reinhardtii: with focus on proteins involved in lipid metabolism. Proteomics. 2011; 11: 4266-4273. doi: 10.1002/pmic.201100114.
  70. 70. Kato N, Dong T, Bailey M, Lum T, Ingram D. Triacylglycerol mobilization is suppressed by brefeldin A in Chlamydomonas reinhardtii. Plant Cell Physiol. 2013; 54: 1585-1599. doi:10.1093/pcp/pct103.
  71. 71. Merchant SS, Kropat J, Liu B, Shaw J, Warakanont J. TAG, you're it! Chlamydomonas as a reference organism for understanding algal triacylglycerol accumulation. Curr Opin Biotechnol. 2012; 23: 352-363. doi: 10.1016/j.copbio.2011.12.001.
  72. 72. Li X, Benning C, Kuoc M-H. Rapid Triacylglycerol Turnover in Chlamydomonas reinhardtii Requires a Lipase with Broad Substrate. Eukaryot Cell. 2012; 11: 1451-1462. doi: 10.1128/EC.00268-12.
  73. 73. Li X, Moellering ER, Liu B, Johnny C, Fedewa M, Sears BB, Kuo MH, Benning C. A galactoglycerolipid lipase is required for triacylglycerol accumulation and survival following nitrogen deprivation in Chlamydomonas reinhardtii. Plant Cell. 2012; 24: 4670-4686. doi: 10.1105/tpc.112.105106.
  74. 74. Guarnieri MT, Pienkos PT. Algal omics: unlocking bioproduct diversity in algae cell factories. Photosynth Res. 2014; 123: 255-263. doi: 10.1007/s11120-014-9989-4.
  75. 75. Jamers A, Blust R, De Coen W. Omics in algae: paving the way for a systems biological understanding of algal stress phenomena? Aquat Toxicol. 2009; 92: 114-121. doi: 10.1016/j.aquatox.2009.02.012.
  76. 76. Rismani-Yazdi H, Haznedaroglu BZ, Bibby K, Peccia J. Transcriptome sequencing and annotation of the microalgae Dunaliella tertiolecta: pathway description and gene discovery for production of next-generation biofuels. BMC Genomics. 2011; 12: 148. doi: 10.1186/1471-2164-12-148.
  77. 77. Guarnieri MT, Nag A, Yang S, Pienkos PT. Proteomic analysis of Chlorella vulgaris: potential targets for enhanced lipid accumulation. J Proteomics. 2013; 93: 245-253. doi: 10.1016/j.jprot.2013.05.025.
  78. 78. Lv H, Qu G, Qi X, Lu L, Tian C, Ma Y. Transcriptome analysis of Chlamydomonas reinhardtii during the process of lipid accumulation. Genomics. 2013; 101: 229-237. doi: 10.1016/j.ygeno.2013.01.004.
  79. 79. Wase N, Black PN, Stanley BA, DiRusso CC. Integrated quantitative analysis of nitrogen stress response in Chlamydomonas reinhardtii using metabolite and protein profiling. J Proteome Res. 2014; 13: 1373-1396. doi: 10.1021/pr400952z.
  80. 80. Valledor L, Furuhashi T, Recuenco-Muñoz L, Wienkoop S, Weckwerth W. System-level network analysis of nitrogen starvation and recovery in Chlamydomonas reinhardtii reveals potential new targets for increased lipid accumulation. Biotechnol Biofuels. 2014; 7: 171. doi: 10.1186/s13068-014-0171-1.
  81. 81. Yang ZK, Ma YH, Zheng JW, Yang WD, Liu JS, Li HY. Proteomics to reveal metabolic network shifts towards lipid accumulation following nitrogen deprivation in the diatom Phaeodactylum tricornutum. J Appl Phycol. 2014; 26: 73-82. doi: 10.1007/s10811-013-0050-3.
  82. 82. Li Y, Han F, Xu H, Mu J, Chen D, Feng B, Zeng H. Potential lipid accumulation and growth characteristic of the green alga Chlorella with combination cultivation mode of nitrogen (N) and phosphorus (P). Bioresour Technol. 2014; 174: 24-32. doi: 10.1016/j.biortech.2014.09.142.
  83. 83. Garnier M, Carrier G, Rogniaux H, Nicolau E, Bougaran G, Saint-Jean B, Cadoret JP. Comparative proteomics reveals proteins impacted by nitrogen deprivation in wild-type and high lipid-accumulating mutant strains of Tisochrysis lutea. J Proteomics. 2014; 105: 107-120. doi: 10.1016/j.jprot.2014.02.022.
  84. 84. Guihéneuf F, Khan A, Tran L-S. Genetic Engineering: a promising tool to engender physiological, biochemical, and molecular stress resilience in green microalgae. Front Plant Sci. 2016; 7: 400 doi: 10.3389/fpls.2016.00400.
  85. 85. Roesler K, Shintani D, Savage L, Boddupalli S, Ohlrogge J. Targeting of the Arabidopsis homomeric acetyl-coenzyme A carboxylase to plastids of rapeseeds. Plant Physiol. 1997; 113:75-81.
  86. 86. Dehesh K, Tai H, Edwards P, Byrne J, Jaworski JG. Overexpression of 3-ketoacyl-acyl-carrier protein synthase IIIs in plants reduces the rate of lipid synthesis. Plant Physiol. 2001; 125:1103-14.
  87. 87. Xue J, Wang L, Zhang L, Balamurugan S, Li D-W, Zeng H, Yang W-D, Liu J-S, Li H-Y. The pivotal role of malic enzyme in enhancing oil accumulation in green microalga Chlorella pyrenoidosa. Microb Cell Fact. 2016; 15: 120. doi: 10.1186/s12934-016-0519-2.
  88. 88. Iskandarov U, Sitnik S, Shtaida N, Cohen SD,Stefan, Khozin-Goldberg L, Cohen Z, Boussiba S. Cloning and characterization of a GPAT-like gene from the microalga Lobosphaera incisa (Trebouxiophyceae): overexpression in Chlamydomonas reinhardtii enhances TAG production. J Appl Phycol. 2016; 28: 907-919. doi:10.1007/s10811-015-0634-1.
  89. 89. Niu Y-F, Wang X, Hu D-X, Balamurugan S, Li D-W, Yang W-D, Liu J-S, Li H-Y. Molecular characterization of a glycerol-3-phosphate acyltransferase reveals key features essential for triacylglycerol production in Phaeodactylum tricornutum. Biotechnol Biofuels. 2016; 9: 60. doi: 10.1186/s13068-016-0478-1.
  90. 90. Hsieh H-J, Su C-H, Chien L-J. Accumulation of Lipid Production in Chlorella minutissima by triacylglycerol biosynthesis-related genes cloned from Saccharomyces cerevisiae and Yarrowia lipolytica. J Microbiol. 2012; 50: 526-534. doi: 10.1007/s12275-012-2041-5.
  91. 91. Yamaoka Y, Achard D, Jang S, Legéret B, Kamisuki S, Ko D, Schulz‐Raffelt M, Kim Y, Song W-Y, Nishida I, Li‐Beisson Y, Lee Y. Identification of a Chlamydomonas plastidial 2‐lysophosphatidic acid acyltransferase and its use to engineer microalgae with increased oil content. Plant Biotechnol J. 2016; 14: 2158-2167. doi: 10.1111/pbi.12572
  92. 92. Li DW, Cen SY, Liu YH, Balamurugan S, Zheng XY, Alimujiang A, Yang WD, Liu JS, Li HY. A type 2 diacylglycerol acyltransferase accelerates the triacylglycerol biosynthesis in heterokont oleaginous microalga Nannochloropsis oceanica. J Biotechnol. 2016; 229: 65-71. doi: 10.1016/j.jbiotec.2016.05.005.
  93. 93. Chen C-X, Sun Z, Cao H-S, Fang F-L, Ouyang L-L, Zhou Z-G. Identification and characterization of three genes encoding acyl-CoA: diacylglycerol acyltransferase (DGAT) from the microalga Myrmecia incisa. Algal Res. 2015; 12: 280-288. doi: 10.1016/j.algal.2015.09.007.
  94. 94. Chungjatupornchai W, Watcharawipas A. Diacylglycerol acyltransferase type 2 cDNA from the oleaginous microalga Neochloris oleoabundans: cloning and functional characterization. J Appl Phycol. 2015; 27: 1499. doi:10.1007/s10811-014-0448-6
  95. 95. Deng X-D, Gu B, Li Y-J, Hu X-W, Guo J-C, Fei X-W. The roles of acyl-CoA:diacylglycerol acyltransferase 2 genes in the biosynthesis of triacylglycerols by the green algae Chlamydomonas reinhardtii. Mol Plant. 2012; 5: 945-947. doi: 10.1631/jzus.B1300180.
  96. 96. Murphy D. The biogenesis and functions of lipid bodies in animals, plants and microorganisms. Prog Lipid Res. 2001; 40, 325-438.
  97. 97. Farese RV Jr, Walther TC. Lipid droplets finally get a little R-E-S-P-E-C-T. Cell. 2009; 139: 855-860. doi: 10.1016/j.cell.2009.11.005.
  98. 98. Fujimoto T, Parton RG. Not just fat: the structure and function of the lipid droplet. Cold Spring Harb Perspect Biol. 2011; 3: a004838. doi: 10.1101/cshperspect.a004838.
  99. 99. Vieler A, Brubaker SB, Vick B, Benning C. A lipid droplet protein of Nannochloropsis with functions partially analogous to plant oleosins. Plant Physiol. 2012; 158: 1562-1569. doi: 10.1104/pp.111.193029
  100. 100. Kimmel AR, Brasaemle DL, McAndrews-Hil M, Sztalryd C, Londos C. Adoption of PERILIPIN as a unifying nomenclature for the mammalian PAT-family of intracellular lipid storage droplet proteins. J Lipid Res. 2010; 51: 468-471. doi: 10.1194/jlr.R000034.
  101. 101. Subramanian V, Rothenberg A, Gomez C, Cohen AW, Garcia A, Bhattacharyya S, Shapiro L, Dolios G, Wang R, Lisanti MP, et al. Perilipin A mediates the reversible binding of CGI-58 to lipid droplets in 3T3-L1 adipocytes. J Biol Chem. 2004; 279: 42062-42071. doi: 10.1074/jbc.M407462200.
  102. 102. Jolivet P, Roux E, D'Andrea S, Davanture M, Negroni L, Zivy M, Chardot T. Protein composition of oil bodies in Arabidopsis thaliana ecotype WS. Plant Physiol Biochem. 2004; 42: 501-509. doi: 10.1016/j.plaphy.2004.04.006.
  103. 103. Peled E, Leu S, Zarka A, Weiss M, Pick U, Khozin-Goldberg I, Boussiba S. Isolation of a novel oil globule protein from the green alga Haematococcus pluvialis (Chlorophyceae). Lipids. 2011; 46: 851-861. doi: 10.1007/s11745-011-3579-4.
  104. 104. Ouyang LL, Chen SH, Li Y, Zhou ZG. Transcriptome analysis reveals unique C4-like photosynthesis and oil body formation in an arachidonic acid-rich microalga Myrmecia incisa Reisigl H4301. BMC Genomics. 12013; 4: 396. DOI: 10.1186/1471-2164-14-396.
  105. 105. Nojima D, Yoshino T, Maeda Y, Tanaka M, Nemoto M, Tanaka T. Proteomics analysis of oil body-associated proteins in the oleaginous diatom. J Proteome Res. 2013; 12: 5293-5301.
  106. 106. Maeda Y, Sunaga Y, Yoshino T, Tanaka T. Oleosome-associated protein of the oleaginous diatom Fistulifera solaris contains an endoplasmic reticulum-targeting signal sequence. Mar Drugs. 2014; 12: 3892-3903. doi: 10.3390/md12073892.
  107. 107. Kim S, Kim H, Ko D, Yamaoka Y, Otsuru M,Kawai-Yamada M, Toshiki Ishikawa T, Oh H-M, Nishida I, Li-Beisson Y, Lee Y. Rapid Induction of Lipid Droplets in Chlamydomonas reinhardtii and Chlorella vulgaris by Brefeldin A. PLoS One 2013; 8: e81978. doi: 10.1371/journal.pone.0081978.
  108. 108. Goodson C, Roth R, Wang ZT, Goodenough U. Structural correlates of cytoplasmic and chloroplast lipid body synthesis in Chlamydomonas reinhardtii and stimulation of lipid body production with acetate boost. Eukariot Cell. 2011; 10: 1592-1606. doi: 10.1128/EC.05242-11.
  109. 109. Solovchenko AE. Physiological role of neutral lipid accumulation in eukaryotic microalgae under stresses. Russian J Plant Physiol. 2012; 59: 167-176. doi:10.1134/S1021443712020161.
  110. 110. Trentacoste EM, Shrestha RP, Smith SR, Glé C, Hartmann AC, Hildebrand M, Gerwick WH. Metabolic engineering of lipid catabolism increases microalgal lipid accumulation without compromising growth. Proc Natl Acad Sci USA. 2013; 110: 19748-19753. doi: 10.1073/pnas.1309299110.
  111. 111. Barnech Bielsa G, Popovich CA, Rodríguez MC, Martín LA, Matulewicz MC, Leonardi PI. Simultaneous production assessment of triacylglycerols for biodiesel and exopolysaccharides as valuable co-products in Navicula cincta. Algal Res. 2016; 15: 120-128. doi: 10.1016/j.algal.2016.01.013.
  112. 112. Ho S-H, Huang S-W, Chen C-Y, Hasunuma T, Kondo A, Chang J-S. Bioethanol production using carbohydrate-rich microalgae biomass as feedstock. Bioresour Technol. 2013; 135: 191-198. Doi: 10.1016/j.biortech.2012.10.015.
  113. 113. Kumar VB, Pulidindi IN, Kinel-Tahan Y, Yehoshua Y, Gedanken A. Evaluation of the potential of chlorella vulgaris for bioethanol production. Energy Fuels. 2016; 30: 3161-3166. doi: 10.1021/acs.energyfuels.6b00253.
  114. 114. Trabelsi L, Chaieb O, Mnari A, Abid-Essafi S, Aleya L. Partial characterization and antioxidant and antiproliferative activities of the aqueous extracellular polysaccharides from the thermophilic microalgae Graesiella sp. BMC Complement Altern Med. 2016; 16: 210. doi: 10.1186/s12906-016-1198-6.
  115. 115. Scodelaro Bilbao PG, Damiani C, Salvador GA, Leonardi PI. Haematococcus pluvialis as a source of fatty acids and phytosterols: potential nutritional and biological implications. J Appl Phycol. 2016; 28: 3283. doi:10.1007/s10811-016-0899-z.
  116. 116. Francavilla M, Trotta P, Luque R. Phytosterols from Dunaliella tertiolecta and Dunaliella salina: a potentially novel industrial application. Bioresour Technol. 2010; 101: 4144-4150. doi: 10.1016/j.biortech.2009.12.139.
  117. 117. Ryckebosch E, Bruneel C, Termote-Verhalle R, Goiris K, Muylaert K, Foubert I. Nutritional evaluation of microalgae oils rich in omega-3 long chain polyunsaturated fatty acids as an alternative for fish oil. Food Chem. 2014; 160: 393-400. doi: 10.1016/j.foodchem.2014.03.087.
  118. 118. Waghmare AG, Manoj K. Salve MK, Jean Guy LeBlanc JG, Shalini S. Arya SS Concentration and characterization of microalgae proteins from Chlorella pyrenoidosa. Biores Bioproc. 2016; 3:16. doi: 10.1186/s40643-016-0094-8.
  119. 119. Tibbetts SM, Whitney CG, MacPherson MJ, Bhatti S, Banskota AH, Stefanova R, McGinn PJ. Biochemical characterization of microalgal biomass from freshwater species isolated in Alberta, Canada for animal feed applications. Algal Res. 2015; 11: 435-447. doi: 10.1016/j.algal.2014.11.011.
  120. 120. Bong SC, Loh SP. A study of fatty acid composition and tocopherol content of lipid extracted from marine microalgae, Nannochloropsis oculata and Tetraselmis suecica, using solvent extraction and supercritical fluid extraction. Int Food Res J. 2013; 20: 721-729.
  121. 121. Safafar H, van Wagenen J, Møller P, Jacobsen C. Carotenoids, phenolic compounds and tocopherols contribute to the antioxidative properties of some microalgae species grown on industrial wastewater. Mar Drugs. 2015; 13: 7339-7356. doi: 10.3390/md13127069.
  122. 122. Shah MR, Liang Y, Cheng JJ, Daroch M. Astaxanthin-producing green microalga Haematococcus pluvialis: from single cell to high value commercial products. Front Plant Sci. 2016; 7: 531. doi: 10.3389/fpls.2016.00531.
  123. 123. Fu W, Paglia G, Magnúsdóttir M, Steinarsdóttir EA, Gudmundsson S, Palsson BO, Andrésson OS, Brynjólfsson S. Effects of abiotic stressors on lutein production in the green microalga Dunaliella salina. Microb Cell Fact. 2014; 13: 3. doi: 10.1186/1475-2859-13-3.
  124. 124. Borodina A, Ladygina LV. The Effect of cultivation conditions on accumulation of carotenoids in Phaeodactylum tricornutum bohl. (Bacillariophyta). Int J Algae. 2013; 15: 274-284. doi: 10.1615/InterJAlgae.v15.i3.70
  125. 125. Borowitzka MA. Carotenoid production using microorganisms. In Cohen Z, Ratledge C (eds). Single Cells Oils: Microbial and Algal Oils. AOCS Press, Urbana. 2010; 225-240.

Written By

Paola Scodelaro Bilbao, Gabriela A. Salvador and Patricia I. Leonardi

Submitted: September 27th, 2016 Reviewed: January 16th, 2017 Published: June 21st, 2017