Open Access
Review
Numéro
OCL
Volume 20, Numéro 6, November-December 2013
Numéro d'article D606
Nombre de pages 8
Section Dossier: Les micro-organismes producteurs de lipides / Producing micro-organisms lipids
DOI https://doi.org/10.1051/ocl/2013031
Publié en ligne 22 novembre 2013

© Y. Li-Beisson et al., published by EDP Sciences, 2013

Licence Creative Commons
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

1 Introduction

1.1 Biofuel

With the rapid decrease in fossil fuel reserves, the increasing demand in energy, particularly for transportation and the rising concerns about global warming and other related social-economic issues, there is a world-wide urge to develop renewable platforms for fuel production. The solutions to meet the energy demand could be many-fold, ranging from solar, water, wind, nuclear energy to bioenergy. Studies of energy production based on biomass have gained increasing attention because the knowledge generated will not only allow production of fuels but also other high-value chemical molecules essential to the oleo-chemical industries which at the moment are still heavily dependent on fossil fuel. Three generations of biofuels have emerged. The 1st generation biofuel is based on edible plant parts (oilseeds, grains, etc.); the 2nd refers to energy production from non-edible plants or non-edible parts of plants; and the 3rd is based on energy production from photosynthetic microorganisms such as microalgae.

1.2 Advantages and limits of the 3rd generation biofuel

Microalgae have very high surface productivity, and can be cultivated on non-arable land (therefore not competing with food production). Because microalgae are photosynthetic (i.e. CO2-fixing) organisms, their use provides greenhouse gas mitigation benefits. Depending on the species, microalgae can grow on fresh, brackish, sea, or even waste water and can accumulate up to 60% oil per dry weight under stress conditions (Chisti, 2007). Therefore, microalgae have attracted increasing attention for their potential as producers of biodiesel or other lipid-based biofuels (Beer, et al., 2009; Scott, et al., 2010; Wijffels and Barbosa, 2010). Costs of cultivation in photobioreactors or open ponds, biomass harvest and oil extraction from algal biomass remain very high however (Grima, et al., 2003). In addition, many microalgal species are not suitable for industrial cultures, fatty acid composition of microalgal lipids may not be optimal for use as biofuel, and stress conditions needed to accumulate lipids result in arrest of cell growth and division, causing a strong limitation of biomass productivity (Siaut, et al., 2011). Therefore, biofuel production from microalgae is not yet sustainable, and we have not yet come up with one algal strain which can be called “an algal crop” for biofuel production. Intensive research efforts are needed in both strain development and technology innovations (Delrue, et al., 2012). Here we review key advances on the biotechnological aspects of microalgal oil research.

thumbnail Fig. 1

Structure of a triacylglycerol and its conversion to biodiesel via acid- or based-catalyzed transmethylation reaction.

2 From microalgal lipids to biodiesel

Microalgae have been found to synthesize a large variety of fatty acids and lipids (Hu, et al., 2008; Harwood and Guschina, 2009), the composition of which often reflects adaptation to environmental conditions (Harwood and Guschina, 2009). In photosynthetic organisms, chloroplast is the central compartment of fatty acid synthesis. Fatty acids produced in the plastid are building blocks of all membrane and storage lipids. Upon stresses, almost all microalgae species can produce oil, i.e. triacylglycerols (TAGs), the major form of storage lipids in eukaryotic cells. One triacylglycerol molecule is made of three often different fatty acids which are esterified to the 3 hydroxyl groups of a glycerol backbone. Biological oils are precursors to diesel, and can be converted to biodiesel via a simple transesterification procedure (Fig. 1). This chemical process liberates fatty acid methyl esters, which are the component of biodiesel. Acyl chains determine the structural thus physical properties of the oil and the biodiesel. Modification of fatty acids (e.g. degree of unsaturation, chain length, introduction of functional groups such as hydroxyl or epoxy, etc.) has been one of the major targets of genetic engineering in plants and microalgae (Shanklin and Cahoon, 1998). Fatty acids produced by microalgae, thus, represent a potential alternative not only to fossil fuels but also to other petroleum derivatives (synthons for green chemistry).

3 Major biotechnological challenges

3.1 Decoupling oil synthesis from arrest of cell division

The amount of oil produced by a microalga is dependent on species and cultivation conditions (Sheehan, 1998; Hu, et al., 2008). Substantial oil accumulation almost always requires stress conditions. One of the most potent stresses to trigger oil accumulation is nitrogen deprivation. Under such culture conditions, some microalgae such as Chlorella species can accumulate up to 60% of oils in its biomass, explaining the considerable regain for microalgae as a biofuel (biodiesel) feedstock (Chisti, 2007). However, nitrogen-starvation limits the overall productivity of the system (Hu, et al., 2008). Maximal lipid yields obtained so far in large scale cultivation systems are 10 to 20 times lower than the theoretical maximum (5000–15 000 gallons per acre per year) (Sheehan, 1998). To circumvent the dependence on stress, dissecting the cellular processes of response to nutrient status, cell division and carbon storage through the study of mutant strains is required.

thumbnail Fig. 2

Lipid biosynthesis in an algal cell, major pathways and cellular locations. CoA, coenzyme A; DAG, diacylglycerol; DGAT, diacylglycerol acyltransferase; ER, endoplasmic reticulum; GPAT, glycerol-3-phosphate acyltransferase; FAS, fatty acid synthesis; G3P, glycerol-3-phosphate; LPAT, lysophosphatidic acid acyltransferase; LPA, lysophosphatidic acid; MAG, monoacyglycerol; PA, phosphatidic acid; PAP, phosphatidic acid phosphatise; PL, phospholipids; PDAT, phospholipid:diacylglycerol acyltransferase.

3.2 Harnessing the complexity of lipid metabolism

Our current understanding of oil biosynthesis in microalgae is still rather limited, although rapid progress has been made lately (Merchant, et al., 2012; Liu and Benning, 2013). The current pathway involves three major spatially separated biochemical steps, i.e. plastidial de novo fatty acid synthesis, acylation of fatty acids to glycerol, and deposition as oil bodies, the sub-cellular compartment destined for oil storage (Fig. 2). In plants, this seemingly simple route requires the co-ordinated actions of several hundred proteins, and regulation across three sub-cellular compartments (Li-Beisson, 2010). For oil synthesis alone in micro algae, three distinct pathway have been proposed (Fig. 2). In plans, the best known is an acyl-CoA-dependent pathway, catalyzed by ER membrane bound enzymes (Riekhof, et al., 2005), similarly to what occurs in plants. An alternative route to oil synthesis is present in both plants and yeast, and is catalyzed by phospholipid:diacylglycerol acyltransferase (PDAT) contributing to the synthesis of triacylglycerol using phosphatidylcholine as an acyl donor and sn-1,2-diacylglycerol as an acyl acceptor. A homolog of this enzyme is present in some sequenced algal genomes including C. reinhardtii where a mutant of PDAT is found to accumulate 30% less oil than its wild-type (Yoon, et al., 2012), thus establishing the contribution of this pathway to oil synthesis in the green algal lineage. A third pathway has recently been proposed by Fan and Xu (Fan, et al., 2011) who has demonstrated that at least part of the triacylglycerol synthetic pathway is present in the plastid of C. reinhardtii. They have further observed under transmission electron microscope (TEM) that some of the oil droplets formed in the plastid can be secreted and re-located to the cytosol. This was later confirmed by another study (Goodson, et al., 2011). These observations point to the fact that, in C. reinhardtii at least, some triacylglycerols are made in the plastid. Thus, a deeper knowledge of the underlying biochemistry, cell biology and genetics of lipid metabolism in this group of organisms is needed.

The champions of current agriculture, for example modern maize, wheat, etc., have been obtained through millennium years of domestication (i.e. selective breeding). None of microalgae species so far have yet been subjected to domestication (selection or breeding) for oil production. The goals of microalgae lipid biologists include construction of industrially robust hyper oil-accumulators via genetic engineering, thus providing the society and industry with designer algal “crops” (Fig. 3). These goals can only be met through combining comprehensive knowledge on lipid metabolic pathways with development of novel and sophisticated genetic and genomic tools.

thumbnail Fig. 3

Domestication of plant crops, and the possible future of an algal crop.

4 Current and emerging algal models for research on lipid-based biofuels

Microalgae are microscopic algae present in freshwater as well as marine systems. It encompasses the largest biodiversity with 200 000–800 000 species exist of which about 40 000 species are described (Andersen, 1992), thus constituting one of the least explored biological resources. Several high-value compounds [for example carotenoids, ω-3 polyunsaturated fatty acids (PUFAs), antioxidants, etc.] are currently commercially produced from industrially cultivated species (Wijffels and Barbosa, 2010). An algal crop destined for biofuel production is yet to emerge. Understanding of the function of biological systems in any given group of organism has largely depended on the development of models. Lately, due to the easy access to genome sequencing technologies, more than 10 microalgal species have been sequenced, and many more are in the pipeline. Intensive efforts have been put in place for the development of molecular genetic tools, thus allowing genetic manipulation of any given algal lineage. This development together with our knowledge gained through examining model systems should aid in the master design of an ideal algal cell factory for production of industrially desirable molecules (including diesel). The algae field is still lacking essential models to accommodate its great diversity. Lately, several models for different phyla are emerging and are used for fundamental research in many laboratories. A few of these models are described briefly below.

4.1 Chlamydomonas reinhardtii

Among the diversity of microalgal species, the Chlorophyceae Chlamydomonas reinhardtii is the best studied microalga at the physiological, as well as genetic and genomic level. The success of C. reinhardtti as a model is largely due to the possibility to perform genetic analysis through sexual crosses. It has been used to study various fundamental biological processes including photosynthesis, chloroplast biogenesis, flagella function and assembly, starch accumulation, photobiological hydrogen production, and more recently on lipid accumulation (Harris, 2001; Merchant, et al., 2012).

C. reinhardtii can be grown either photoautotrophically, mixotrophically or heterotrophically (Harris, 2001). Photosynthetic function in C. reinhardtii is thus dispensable, a feature which has originally been used to isolate mutants defected in photosynthetic apparatus (Bennoun and Levine, 1967). During its vegetative phase, Chlamydomonas nuclear genome is haploid thus allowing identification of mutant phenotypes at the first generation, which facilitated forward genetic approaches. All three genomes (nuclear, plastid and mitochondrion) have been sequenced, and genetic transformation is possible for all three genomes (Rochaix, 2002). Substantial literature and infrastructure is in place, thus opens doors to new discoveries.

Many laboratories have adopted C. reinhardtii as a reference organism for studying TAG accumulation (Merchant, et al., 2012), although there has been some debates regarding if Chlamydomonas is or not an oleaginous alga. Stressed Chlamydomonas cells accumulate oils in oil bodies (Wang, et al., 2009; Moellering and Benning, 2010; Siaut, et al., 2011). Depending on the strains used and stress conditions, the oil content can reach 50% of dry biomass in starch-less mutants in response to N starvation (Li, et al., 2010). With the versatile molecular genetic tools available (Harris, 2001), Chlamydomonas thus serves as an excellent model organism for addressing fundamental biological questions related to oil synthesis and degradation.

If Chlamydomonas has been widely and successfully used to develop forward genetic approaches to study different processes (photosynthesis, flagellar movement, starch metabolism...) reverse genetic approaches are more limited in this model species. Major drawbacks working with Chlamydomonas are the fact that transgene expression is at low efficieng and that no efficient transformation of the nuclear genome based on homologous recombination has been demonstrated. Isolation of mutants affected in a given gene employing PCR-based techniques is feasible but tedious (Gonzalez-Ballester, et al., 2011). With the development of relatively cheap high-throughput genome sequencing (Dutcher, et al., 2012), tagged mutant collections start to emerge (personal communication S. Merchant). Another major concern over a model organism like C. reinhardtii is often the fact that it is not an industrial alga and has not historically been cultivated at large scale. But a possible solution to this drawback has lately been provided by Microphyt, a French company located in Montpellier, who has developed technologies for industrial cultivation of C. reinhardtii in tubular photobioreactors.

4.2 Chlorella sp.

Chlorella is a genus of single-cell green microalgae, belonging to the same phylum Chlorophyta as Chlamydomonas. It is spherical in shape, about 2–10 μm in diameter, and can be distinguished from Chlamydomonas because it does not have flagella. Chlorella consists of over 80 species, isolated from either freshwater or marine environment. It is an attractive food producer because it is high in protein and other essential nutrients; several species are known sources of ω-3 PUFAs (Tokusoglu and Unal, 2003) and astaxanthin (Campenni et al. 2013). It is robust and widely cultivated commercially. One species has been sequenced (Blanc, et al., 2010), and genetic manipulation is demonstrated for three species (C. vulgaris, C. ellipsoidea, and C. kessleri) (Leon and Fernandez, 2007; Gong, et al., 2011; Niu, et al., 2011; Chia, et al., 2013). Chlorella is an emerging model for study of photosynthetic carbon fixation in the green lineage of eukaryotic microalgae.

4.3 Ostreococcus tauri

The marine pico-eukaryote Ostreococcus belongs to Chlorophyta, and is the smallest photosynthetic unicellular eukaryote (Palenik, et al., 2007). It has an extremely small genome i.e. 12 Mb which is 10 times less than C. reinhardtii. Ostreococcus offers great advantages for functional genetic studies because it is thought to represent the minimal genes required to carry out essential biological functions. Ostreoccocus has emerged as a powerful “ready-to-use” algal model because a set of genetic tools have been developed including high efficiency gene overexpression and gene targeting by homologous recombination (personal communication F.Y Bouget), which makes it more amenable to many functional genetic studies than Chlamydomonas. Ostreococcus as a model for marine species is, thus, highly complementary to the freshwater green alga Chlamydomonas. Finally, substantial amount of literature is present which allows us minimal understanding of its physiology.

4.4 Phaeodactylum tricornutum

Diatoms are responsible for a significant proportion of primary marine biomass production. Phaeodactylum tricornutum has been developed as a model for diatom research. It is the only species in the genus Phaeodactylum. Unlike most other diatoms P. tricornutum can grow in the absence of silicon, and the biogenesis of silicified frustules is facultative. P. tricornutum can accumulate significant amount of lipids under silicon absence (Sheehan, 1998), and the fact that it can synthesis high amount of ω-3 PUFAs have brought considerable interests in research on lipid metabolism in this diatom. These features together with its high biomass productivity, high lipid content, a fully sequenced genome (Bowler, et al., 2008), and the versatile molecule toolboxes developed (Siaut, et al., 2007), made it a prominent model system for both biodiesel and ω-3 PUFA production. Nonetheless, one of the major issues associated with P. tricornutum as a model is that there is no easy way to knock out genes, knock down being mostly available (De Riso, et al., 2009).

4.5 Nannochloropsis sp.

Nannochloropsis is a heterokont microalga comprising six species, of which five are marine and one freshwater. Due to its rigorous growth, and high content of ω-3 PUFAs and astaxanthin, it is commonly used in aquaculture applications. Upon stress, it can also accumulate significant amount of neutral lipids thus attracting interests on its development for biofuel production. Draft genome sequences are available for two of the marine species (N. gaditana and N. oceanica) (Radakovits, et al., 2012; Vieler, et al., 2012). Genetic transformation is available, and more importantly homologous recombination has been demonstrated with high efficiencies for N. gaditana (Kilian, et al., 2011). Thus, Nannochloropsis represents an emerging model for lipid research on photosynthetic protists. Substantial collaborative effort at international level is required for this organism to become as the premier model for algal research.

5 Looking through current research approaches into future directions

5.1 Bio-prospecting for high performing algal strains suitable for oil production

Only a handful of the tens of thousands of microalgal species known to exist in nature are currently cultured at large scale and used for commercial applications, among which only a few hundreds have been investigated for potential biotechnological applications. Bio-prospecting for new species from local or extreme environments for oil content or other high value product has been an on-going international effort. One of the largest conducted so far has been the survey carried out by the U.S. Department of Energy (Sheehan, 1998). They studied around 300 species selected from 3000 isolated strains. More recently, several research studies for strain selection and induction of lipid biosynthesis have been carried out (de la Vega, et al., 2011; Do Nascimento et al. 2012). These studies revealed that the intrinsic ability to produce large quantity of oil and lipid is generally species/strain specific, rather than genus specific. Nonetheless, a survey of the literature by Hu et al. (Hu et al. 2008) revealed that species belonging to the group of green microalgae (Chlorophyta) generally accumulate oil to a higher level than species of other algal taxa such as cyanobacteria, brown algae, or red algae. In addition to the ability to produce biomass and accumulate oil, other criteria related to cultivation constraints (strain robustness) or to downstream processes (harvesting, oil extraction...) have to be considered together and has been proposed to impact the final productivity of the system and the economy of microalgae-based biodiesel production (Delrue, et al., 2012). Continuing efforts at bio-prospecting of strains isolated from local environment is a key to isolate strains suitable for one particular geological niche.

thumbnail Fig. 4

Current major approaches applied and future directions.

5.2 Genetic improvement of algal strains

Current crop plants have gone through thousands of years of selective breeding which is a common agricultural practice. However conventional breeding is not suitable for algae strains since sexual cycles are lacking or poorly defined for most species. Metabolic engineering thus represents one of the most promising strategies toward production of suitable algal strains (Rosenberg, et al., 2008). Comparing to classical breeding, genetic engineering is a time-saving technology. Indeed, metabolic engineering has revolutionized the traditional breeding programme and made high contributions to meet our societal needs. One of the most notable examples in successful genetic engineering is probably the creation of “golden rice” via genetic engineering of the rice plant to synthesize beta-carotene via introduction of two enzymes (Ye et al. 2000). Given the latest development in genomics, genetics, and molecular biology, synthetic biology is coming of age technology for addressing basic as well as applied questions in the bio-production of desirable molecules and fuels. For example, genetic engineering has already allowed the isolation of Chlamydomonas mutants with higher ability for the photobiological production of hydrogen (Tolleter, et al., 2011); and mutants with reduced antenna sizes showing improved biomass productivity in photobioreactors (Ort, et al., 2011). Heterologous gene expression has been used successfully to redirect microalgal metabolism. For example, heterologous expression of two thioesterases in Phaeodactylum tricornutum caused accumulation of novel medium chain fatty acids (Radakovits, et al., 2011). Regulatory proteins controlling oil synthesis have also been isolated in the model Chlamydomonas, and overexpression of these proteins led to altered cellular oil content (Yohn, 2011; Boyle, et al., 2012).

Despite the promising achievements, all routine work on metabolic engineering through genetic manipulation remains limited to a few model species where efficient and stable transformation is possible. Common issues associated with other transformation experiments reported in the literature are low efficiency and instability of transgenes introduced. Therefore, there is substantial need for development of molecular tools for other non-model microalgal species. One such achievement is the recent demonstration of high efficiency homologous recombination for an industrially relevant alga N. gaditana (Kilian, et al., 2011). Such genetic tools would allow the metabolic engineering of pathways or physiological processes and therefore allow the production of transgenic microalgae that are genetically tailored for the production of biodiesel.

In addition, successful genetic engineering is also dependent on the development of high-throughput screening procedures which allow the rapid and easy screening of a high number of cell lines for desirable phenotypes (high growth rate, high oil content, better fatty acid profile, etc.). For example, in screening of high oil accumulators, the semi-quantitative lipophilic dye Nile red coupled with Flow cytometry has been demonstrated as a powerful tool to isolate mutant with altered oil content (ongoing work in the authors’ laboratory).

5.3 Advancing knowledge on lipid metabolism in microalgae

A comprehensive view of lipid metabolism in microalgae is still in its infancy. Most of the components of lipid biosynthesis are deduced from homologies to proteins of either known plant or yeast lipid synthetic pathways. To aid in genetic engineering and allow intelligent design of strains, a thorough knowledge of lipid metabolism in microalgae is urgently needed. Recent studies dedicated to advance our understanding of oil metabolism in this alga (summarized in Fig. 4) include: (i) detailed characterization of the oil accumulation kinetics in response to nitrogen depletion (Wang, et al., 2009; Siaut, et al., 2011); (ii) testing the potential impact of diverting carbon precursors from starch synthesis to oil accumulation using starchless mutants (Li, et al., 2010; Siaut, et al., 2011); (iii) identification of proteins associated with oil droplets – the major cellular compartment for oil storage (Moellering and Benning, 2010; Nguyen, et al., 2011); (iv) comparative transcriptomic studies on cells before and after nitrogen removal (Miller, et al., 2010); (v) microscopic and biochemical analyses of the possible sub-cellular locations of oil droplets in Chlamydomonas and other algae (Fan, et al., 2011; Goodson, et al., 2011; Fan, et al., 2012); (vi) Forward genetic screening of mutants affected in oil synthesis (Li, et al., 2012). Please note that these are just examples, and are not meant for an exhaustive list. These pioneering works have revealed the potential complexity and divergence of oil metabolism from that of higher plants, and highlights interesting candidate genes for genetic engineering studies. Now it is the right time for verification of gene function using the latest tools developed for Chlamydomonas such artificial microRNA techniques to knock-down specific genes (Molnar, et al., 2009), or isolation of specific mutants using PCR-based screening (Gonzalez-Ballester, et al., 2011). In parallel, the development of forward genetic approaches based on efficient and high-throughput screening of mutant showing interesting properties (such as the ability to accumulate oil in optimal growth conditions), will be important to identify regulatory genes controlling oil accumulation.

6 Conclusion

Photosynthetic microalgae have provided us with food and feed, and now possibly also fuel. Research on microalgae lipids have come a long way starting from the 40’s during the Second World War in Germany. Given the high impact of research on energy, algal fuel development is tightly related to economic, social and world development. Technologies developed today have progressed from manipulation of one or two genes to more system-based approaches. These technologies include high-throughput genome sequencing, and a suit of – omics technologies (genomics, transcriptomics, proteomics, metabolomics, and lipidomics). These powerful tools if used together with transformation and molecular genetic toolboxes developed for particular algal strains provide ample opportunities for lipid scientists to redesign algal metabolism toward production of oils or other chemical molecules useful for industrial applications.

Acknowledgments

The authors thank funding by ANR project DIESALG and we acknowledge DGA for financial support of the project CAER. They also thanks the support provided by the HélioBiotec platform, funded by the European Union (ERDF), the Région Provence Alpes Côte d’Azur, the French Ministry of Research (ANR) and CEA.

References

  • Andersen RA. 1992. Diversity of eukaryotic algae. Biodivers. Conserv. 1: 267–292. [CrossRef] [Google Scholar]
  • Beer LL, Boyd ES, Peters JW, Posewitz MC. 2009. Engineering algae for biohydrogen and biofuel production. Curr. Opin. Biotechnol. 20: 264–271. [CrossRef] [PubMed] [Google Scholar]
  • Bennoun P, Levine RP. 1967. Detecting Mutants That Have Impaired Photosynthesis by Their Increased Level of Fluorescence. Plant Physiol. 42: 1284–1287. [CrossRef] [PubMed] [Google Scholar]
  • Blanc G, Duncan G, Agarkova I, Borodovski M, Gurnon J, Kuo A, et al. 2010. The Chlorella variabilis NC64A Genome Reveals Adaptation to Photosymbiosis, Coevolution with Viruses, and Cryptic Sex. Plant Cell 22: 2943–2955. [CrossRef] [PubMed] [Google Scholar]
  • Bowler C, Allen AE, Badger JH, Grimwood J, Jabbari K, Kuo A, et al. 2008. The Phaeodactylum genome reveals the evolutionary history of diatom genomes. Nature 456: 239–244. [CrossRef] [PubMed] [Google Scholar]
  • Boyle NR, Page MD, Liu B, Blaby IK, Casero D, Kropat J, et al. 2012. Three Acyltransferases and Nitrogen-responsive Regulator Are Implicated in Nitrogen Starvation-induced Triacylglycerol Accumulation in Chlamydomonas. J. Biol. Chem. 287: 15811–15825. [CrossRef] [PubMed] [Google Scholar]
  • Campenni L, Nobre BP, Santos CA, Oliveira AC, Aires-Barros MR, Palavra AM, et al. 2013. Carotenoid and lipid production by the autotrophic microalga Chlorella protothecoides under nutritional, salinity, and luminosity stress conditions. Appl. Microbiol. Biotechnol. 97: 1383–1393. [CrossRef] [PubMed] [Google Scholar]
  • Chia MA, Lombardi AT, Melão MdGG, Parrish CC. 2013. Lipid composition of Chlorella vulgaris (Trebouxiophyceae) as a function of different cadmium and phosphate concentrations. Aquat. Toxicol. 128–129: 171–182. [CrossRef] [PubMed] [Google Scholar]
  • Chisti Y. 2007. Biodiesel from microalgae. Biotechnol. Adv. 25: 294–306. [CrossRef] [PubMed] [Google Scholar]
  • de la Vega M, Diaz E, Vila M, Leon R. 2011. Isolation of a new strain of Picochlorum sp and characterization of its potential biotechnological applications. Biotechnol. Progress 27: 1535–1543. [CrossRef] [Google Scholar]
  • De Riso V, Raniello R, Maumus F, Rogato A, Bowler C, Falciatore A. 2009. Gene silencing in the marine diatom Phaeodactylum tricornutum. Nucleic Acids Res. 37: e96 [CrossRef] [PubMed] [Google Scholar]
  • Delrue F, Setier PA, Sahut C, Cournac L, Roubaud A, Peltier G, et al. 2012. An economic, sustainability, and energetic model of biodiesel production from microalgae. Bioresour. Technol. 111: 191–200. [CrossRef] [PubMed] [Google Scholar]
  • Do Nascimento M, Ortiz-Marquez JCF, Sanchez-Rizza L, Echarte MM, Curatti L. 2012. Bioprospecting for fast growing and biomass characterization of oleaginous microalgae from South-Eastern Buenos Aires, Argentina. Bioresour. Technol. 125: 283–290. [CrossRef] [PubMed] [Google Scholar]
  • Dutcher SK, Li L, Lin H, Meyer L, Giddings TH, Kwan AL, et al. 2012. Whole-genome sequencing to identify mutants and polymorphisms in Chlamydomonas reinhardtii. G3 (Bethesda) 2: 15–22. [Google Scholar]
  • Fan J, Yan C, Andre C, Shanklin J, Schwender J, Xu C. 2012. Oil accumulation is controlled by carbon precursor supply for fatty acid synthesis in Chlamydomonas reinhardtii. Plant Cell Physiol. 53: 1380–1390. [CrossRef] [PubMed] [Google Scholar]
  • Fan JL, Andre C, Xu CC. 2011. A chloroplast pathway for the de novo biosynthesis of triacylglycerol in Chlamydomonas reinhardtii. Febs Lett. 585: 1985–1991. [CrossRef] [PubMed] [Google Scholar]
  • Gong YM, Hu HH, Gao Y, Xu XD, Gao H. 2011. Microalgae as platforms for production of recombinant proteins and valuable compounds: progress and prospects. J. Ind. Microbiol. Biotechnol. 38: 1879–1890. [CrossRef] [PubMed] [Google Scholar]
  • Gonzalez-Ballester D, Pootakham W, Mus F, Catalanotti C, Magneschi L, et al. 2011. Reverse genetics in Chlamydomonas: a platform for isolating insertional mutants. Plant Methods 7: 24. [PubMed] [Google Scholar]
  • Goodson C, Roth R, Wang ZT, Goodenough U. 2011. Structural Correlates of Cytoplasmic and Chloroplast Lipid Body Synthesis in Chlamydomonas reinhardtii and Stimulation of Lipid Body Production with Acetate Boost. Eukaryotic Cell 10: 1592–1606. [CrossRef] [PubMed] [Google Scholar]
  • Grima EM, Belarbi EH, Fernandez FGA, Medina AR, Chisti Y. 2003. Recovery of microalgal biomass and metabolites: process options and economics. Biotechnol. Adv. 20: 491–515. [CrossRef] [PubMed] [Google Scholar]
  • Harris E. 2001. Chlamydomonas as a model organism. Annu. Rev. Plant Physiol. Plant Mol. Biol. 52: 363–406. [CrossRef] [PubMed] [Google Scholar]
  • Harwood JL, Guschina IA. 2009. The versatility of algae and their lipid metabolism. Biochimie 91: 679–684. [CrossRef] [PubMed] [Google Scholar]
  • 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. [CrossRef] [PubMed] [Google Scholar]
  • Kilian O, Benemann CSE, Niyogi KK, Vick B. 2011. High-efficiency homologous recombination in the oil-producing alga Nannochloropsis sp. Proc. Natl. Acad. Sci. USA 108: 21265–21269. [CrossRef] [Google Scholar]
  • Leon R, Fernandez E. 2007. Nuclear transformation of eukaryotic microalgae - Historical overview, achievements and problems. Ado. Exp. Med. Biol. 616: 1–11. [CrossRef] [Google Scholar]
  • Li-Beisson Y, Shorrosh B, Beisson F, et al. 2010. Acyl lipid metabolism In R Last, ed, The Arabidopsis book. American Society of Plant Biologists Rockville, MD. [Google Scholar]
  • Li X, Moellering ER, Liu B, Johnny C, Fedewa M, Sears BB, Kuo M-H, 2012. A Galactoglycerolipid Lipase Is Required for Triacylglycerol Accumulation and Survival Following Nitrogen Deprivation in Chlamydomonas reinhardtii. Plant Cell 24: 4670–4686. [CrossRef] [PubMed] [Google Scholar]
  • Li Y, Han D, Hu G, Sommerfeld M, Hu Q. 2010. Inhibition of starch synthesis results in overproduction of lipids in Chlamydomonas reinhardtii. Biotechnol. Bioeng. 107: 258–268 [CrossRef] [PubMed] [Google Scholar]
  • Liu B, Benning C. 2013. Lipid metabolism in microalgae distinguishes itself. Curr. Opin. Biotechnol. 24: 300–309. [CrossRef] [PubMed] [Google Scholar]
  • Merchant SS, Kropat J, Liu B, Shaw J, Warakanont J. 2012. TAG, You’re it! Chlamydomonas as a reference organism for understanding algal triacylglycerol accumulation. Curr. Opin. Biotechnol. 23: 352–363. [CrossRef] [PubMed] [Google Scholar]
  • Miller R, Wu GX, Deshpande RR, Vieler A, Gartner K, Li XB, et al. 2010. Changes in transcript abundance in Chlamydomonas reinhardtii following nitrogen deprivation predict diversion of metabolism. Plant Physiol. 154: 1737–1752. [CrossRef] [PubMed] [Google Scholar]
  • Moellering ER, Benning C. 2010. RNA interference silencing of a major lipid droplet protein affects lipid droplet size in Chlamydomonas reinhardtii. Eukaryotic Cell 9: 97–106. [CrossRef] [PubMed] [Google Scholar]
  • Molnar A, Bassett A, Thuenemann E, Schwach F, Karkare S, et al. 2009. Highly specific gene silencing by artificial microRNAs in the unicellular alga Chlamydomonas reinhardtii. Plant J. 58: 165–174. [CrossRef] [PubMed] [Google Scholar]
  • Nguyen HM, Baudet M, Cuiné S, Adriano J-M, Barthe D, Billon E, et al. 2011. Proteomic profiling of oil bodies isolated from the unicellular green microalga Chlamydomonas reinhardtii: With focus on proteins involved in lipid metabolism. Proteomics 11: 4266–4273. [CrossRef] [PubMed] [Google Scholar]
  • Niu YF, Zhang MH, Xie WH, Li JN, Gao YF, Yang WD, et al. 2011. A new inducible expression system in a transformed green alga, Chlorella ulgaris. Genet. Mol. Res. 10: 3427–3434. [CrossRef] [PubMed] [Google Scholar]
  • Ort DR, Zhu XG, Melis A. 2011. Optimizing Antenna Size to Maximize Photosynthetic Efficiency. Plant Physiol. 155: 79–85. [CrossRef] [PubMed] [Google Scholar]
  • Palenik B, Grimwood J, Aerts A, Rouzé P, Salamov A, Putnam N, et al. 2007. The tiny eukaryote Ostreococcus provides genomic insights into the paradox of plankton speciation. Proc. Natl. Acad. Sci. USA 104: 7705–7710. [CrossRef] [Google Scholar]
  • Radakovits R, Eduafo PM, Posewitz MC. 2011. Genetic engineering of fatty acid chain length in Phaeodactylum tricornutum. Metab. Eng. 13: 89–95. [CrossRef] [PubMed] [Google Scholar]
  • Radakovits R, Jinkerson RE, Fuerstenberg SI et al. 2012. Draft genome sequence and genetic transformation of the oleaginous alga Nannochloropis gaditana. Nat. Commun. 3: 686 [CrossRef] [PubMed] [Google Scholar]
  • Riekhof WR, Sears BB, Benning C. 2005. Annotation of genes involved in glycerolipid biosynthesis in Chlamydomonas reinhardtii: Discovery of the betaine lipid synthase BTA1(Cr). Eukaryotic Cell 4: 242–252. [CrossRef] [PubMed] [Google Scholar]
  • Rochaix JD. 2002. The three genomes of Chlamydomonas. Photosynth. Res. 73: 285–293. [CrossRef] [PubMed] [Google Scholar]
  • Rosenberg JN, Oyler GA, Wilkinson L, Betenbaugh MJ. 2008. A green light for engineered algae: redirecting metabolism to fuel a biotechnology revolution. Curr. Opin. Biotechnol. 19: 430–436. [CrossRef] [PubMed] [Google Scholar]
  • Scott SA, Davey MP, Dennis JS et al. 2010. Biodiesel from algae: challenges and prospects. Curr. Opin. Biotechnol. 21: 277–286. [CrossRef] [PubMed] [Google Scholar]
  • Shanklin J, Cahoon EB. 1998. Desaturation and related modifications of fatty acids. Ann. Rev. Plant Physiol. Plant Mol. Biol. 49: 611–641. [CrossRef] [PubMed] [Google Scholar]
  • Sheehan J, Dunahay T, Benemann J, Roessler PG. 1998. A look back at the US Department of Energy’s aquatic species program – biodiesel from algae. In US Department of Energy’s Office of Fuels Development, Golden, CO: National Renewable Energy Laboratory. [Google Scholar]
  • Siaut M, Cuine S, Cagnon C et al. 2011. Oil accumulation in the model green alga Chlamydomonas reinhardtii: characterization, variability between common laboratory strains and relationship with starch reserves. BMC Biotechnology 11: 7. [CrossRef] [PubMed] [Google Scholar]
  • Siaut M, Heijde M, Mangogna M et al. 2007. Molecular toolbox for studying diatom biology in Phaeodactylum tricornutum. Gene 406: 23–35. [CrossRef] [PubMed] [Google Scholar]
  • Tokusoglu O, Unal MK. 2003. Biomass nutrient profiles of three microalgae: Spirulina platensis, Chlorella vulgaris, and Isochrisis galbana. J. Food Sci. 68: 1144–1148. [CrossRef] [Google Scholar]
  • Tolleter D, Ghysels B, Alric J et al. 2011. Control of Hydrogen Photoproduction by the Proton Gradient Generated by Cyclic Electron Flow in Chlamydomonas reinhardtii. Plant Cell 23: 2619–2630. [CrossRef] [PubMed] [Google Scholar]
  • Vieler A, Wu G, Tsai C-H et al. 2012. Genome, Functional Gene Annotation, and Nuclear Transformation of the Heterokont Oleaginous Alga Nannochloropsis oceanica CCMP1779. PLoS Genet. 8: e1003064. [CrossRef] [PubMed] [Google Scholar]
  • Wang ZT, Ullrich N, Joo S, Waffenschmidt S, Goodenough U. 2009. Algal lipid bodies: stress induction, purification, and biochemical characterization in wild-type and starchless Chlamydomonas reinhardtii. Eukaryot Cell 8: 1856–1868. [CrossRef] [PubMed] [Google Scholar]
  • Wijffels RH, Barbosa MJ. 2010. An Outlook on Microalgal Biofuels. Science 329: 796–799. [CrossRef] [PubMed] [Google Scholar]
  • Ye X, Al-Babili S, Klöti A, Zhang J, Lucca P, Beyer P, et al. 2000. Engineering the Provitamin A (β-Carotene) Biosynthetic Pathway into (Carotenoid-Free) Rice Endosperm. Science 287: 303–305. [CrossRef] [PubMed] [Google Scholar]
  • Yohn C MM, Behnke C, Brand A. 2011. Stress-induced lipid trigger. US Patent. [Google Scholar]
  • Yoon K, Han D, Li Y, Sommerfeld M, Hu Q. 2012. 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 24: 3708–3724. [CrossRef] [PubMed] [Google Scholar]

Cite this article as: Yonghua Li-Beisson, Gilles Peltier. Third-generation biofuels: current and future research on microalgal lipid biotechnology. OCL 2013, 20(6) D606.

All Figures

thumbnail Fig. 1

Structure of a triacylglycerol and its conversion to biodiesel via acid- or based-catalyzed transmethylation reaction.

In the text
thumbnail Fig. 2

Lipid biosynthesis in an algal cell, major pathways and cellular locations. CoA, coenzyme A; DAG, diacylglycerol; DGAT, diacylglycerol acyltransferase; ER, endoplasmic reticulum; GPAT, glycerol-3-phosphate acyltransferase; FAS, fatty acid synthesis; G3P, glycerol-3-phosphate; LPAT, lysophosphatidic acid acyltransferase; LPA, lysophosphatidic acid; MAG, monoacyglycerol; PA, phosphatidic acid; PAP, phosphatidic acid phosphatise; PL, phospholipids; PDAT, phospholipid:diacylglycerol acyltransferase.

In the text
thumbnail Fig. 3

Domestication of plant crops, and the possible future of an algal crop.

In the text
thumbnail Fig. 4

Current major approaches applied and future directions.

In the text

Les statistiques affichées correspondent au cumul d'une part des vues des résumés de l'article et d'autre part des vues et téléchargements de l'article plein-texte (PDF, Full-HTML, ePub... selon les formats disponibles) sur la platefome Vision4Press.

Les statistiques sont disponibles avec un délai de 48 à 96 heures et sont mises à jour quotidiennement en semaine.

Le chargement des statistiques peut être long.