A considerable effort has been made to identify microalgae suitable for the development of new bio products. However, diatoms have been underexplored despite their adaptability, high versatility and high productivity. This review focuses on the potential use of diatoms to develop new bio products for industrial applications. In particular, cellular mechanisms responsible for lipid synthesis and accumulation exist in diatoms and can maximize lipid (oil) production. Since diatoms are present in various types of ecosystems, from rivers to seas and ice sheets, they are very versatile and can adapt to a range of environmental conditions. Understanding the mechanisms behind the response of diatoms to particular water conditions will significantly advance our knowledge of morphogenesis and metabolism in diatoms. In particular, by modifying diatom growth parameters, it is possible to shift their metabolism towards high lipid accumulation and storage. The versatility of diatoms could then be used to promote the development of new bio products for industrial, agricultural and pharmaceutical applications. This review discusses the distinctive features of diatoms and recent advances enabling potential industrial use of diatoms for oil production.

Leterme SC (2015) The Oil Production Capacity of Diatoms. Ann Aquac Res 2(1): 1007.

•    Lipid accumulation
•    Nutrient conditions
•    Cellular mechanisms
•    Algae
•    Adaptation

N: Nitrogen; P: Phosphorus; Si: Silicon; Zn: Zinc; Fe: Iron; PUFAs: Polyunsaturated fatty acids; TAGs: Triacylglycerides; PL: Phospholipids; GL: Glycolipids; SFE: Supercritical fluid extraction; SCCO2: Supercritical carbon dioxide

Diatoms are a major component of the phytoplankton community, accounting for approximately 40% of total primary production in the ocean [1,2]. They contribute to the export of carbon into the deep ocean and are the main players of the biogeochemical cycles of macro-nutrients such as nitrogen (N), phosphorus (P), and silicon (Si) [3-6]. Importantly, they are keystone organisms affecting zooplankton grazing and recruitment [7,8] at the base of the pelagic food chain. Therefore, critically, any change in diatom levels propagates up the pelagic food-chain. As diatoms have the potential for rapid growth and short generation times, they respond quickly to changing environmental conditions. For example, previous work has shown that the nanostructure and Si composition of diatoms’ frustules appear to be influenced by environmental factors such as salinity [9-11].

The uptake of Si needed for the formation of the frustule is related to the presence of other nutrients in the environment as well as to the size of the diatom cells. In particular, nutrients such as N, P, Zn and Fe have been shown to impact on the uptake of silicic acid Si (OH)4 by the diatoms [12,13] which subsequently impacts on the size of the cells present under those conditions and on their storage capabilities. In addition, intermittent fluctuations in nutrient levels can increase nutrient storage abilities [14,15] and has also been shown as increasing lipid synthesis and accumulation [16]. Since cell size and shape plays a role in nutrient uptake and storage, diatoms of different shapes i.e. centric (round) and/ or pennate (elongated) and sizes can be used to monitor the impact of nutrient, light and salinity conditions (i.e. low, high and intermittent), on (i) their growth rate, (ii) the uptake of photo-assimilates and nutrients for proliferation and (iii) their accumulation and storage. The quantification of the influence of size and shape, exposed to variable environmental conditions, is of great importance in mechanistic interpretation of growth rates and storage capacities.

In the past decade, a considerable effort has been made to identify species of microalgae, including diatoms, which are suitable for high lipid production [17,18]. Diatoms have been shown to contain high amounts of lipids and store lipids [19,20] more efficiently than any other macro algae [21]. In particular, the lipid content of diatoms is much higher under nutrient limitations which represent stress conditions occurring at the end of a bloom [18,22]. Diatoms naturally thrive under nutrientreplete conditions, meaning that ?maximal growth rate is obtained in culture conditions, which is an ideal characteristic for a bio fuel production system. In addition, diatoms also grow best under highly mixed conditions, which are desirable for largescale cultivation. Finally, the extraction of lipids from diatoms is easier than with other algae as the individual parts of the diatom frustule can easily be disrupted to allow for the removal of lipids and pigments from the cells via solvent-extraction [23,24]. Diatoms therefore have a great potential as natural bio product producers.

Adaptability

Diatoms are single-celled algae encased in silica-based cell walls similar to a Petri dish, with overlapping upper and lower halves called the epitheca and hypotheca, respectively [25]. Their cell wall is reproduced with fidelity through generations by genetically controlled assembly processes and can reach a few hundred microns in size [26]. Each of the estimated 105 diatom species has a specific valve capping the thecae, which is decorated with a unique pattern of nano-sized features [27]. The pores present within the valves are species specific and provide precise features for the identification of diatoms [28]. The energy needed for silicification is provided by aerobic respiration and allows for silicic acid uptake, and deposition even in dark conditions [29]. The strategy of building silica-based cells then contributes to the overall productivity of diatoms as the energetic needs for silica polymerization are much lower than an organically based cell wall [30].

The relatively low surface to volume ratios of diatom cells would normally require nutrient-rich conditions for growth. However, diatoms possess a large nutrient storage vacuole which makes them very efficient at nutrient utilization [31,32]. As a result, diatoms can dominate phytoplankton communities and outcompete other eukaryotes under nutrient-limiting conditions [33] or under conditions of mixing and high turbulence [34]. Diatoms are very efficient in adapting to fluctuating environmental conditions. For example, primary production is considered to be limited by iron availability in the ocean; however, diatom species have evolved to employ different strategies to cope with low iron availability [35, 36].The versatility of diatoms allows them to be present in many different environments (i.e., freshwater, seawater, salterns and sea-ice). Since the development of new bio products is regional, and will be defined by the surrounding environmental conditions, diatoms are the ideal candidate for regional production of bio products including bio fuels, livestock and mariculture feed, thermal energy, bio plastics and feed stocks for the nitrogen chemical industry.

Production of lipids

Diatoms take up macronutrients (i.e., N, P, Si), but also store a suite of essential micronutrients that are present at trace concentrations (<0.1 µM) in seawater [37]. Many trace metals (Fe, Co, Zn) are micronutrients essential for the growth of diatoms, and can shape the diatom community structure and distribution. Previous research has demonstrated that diatoms can still grow when conditions of light, salinity and/or nutrients are not optimal. However, those non-optimal conditions have impacts on (i) the uptake of silicic acid needed to construct the frustule (silica shell), (ii) the accumulation of photo-assimilates and nutrients in the cells, (iii) the cells size and shape, and (iv) the growth rates of the diatoms.

For example, low concentrations of Zn and Fe would lower the uptake of silicic acid Si(OH)4 by the diatoms [12,38]; while high N and P concentrations would favor the growth of species with small cell size [39] and high growth rates [40]. Diatoms are excellent lipid accumulators and a substantial portion of the cells’ volume is often occupied by lipid droplets that accumulate rapidly [26].Modifying nutrient levels could favor high biosynthesis and accumulation of polyunsaturated fatty acids (PUFAs) and triacylglycerides (TAGs). In particular, fluctuating nutrient, light and salinity conditions (i.e., varying intermittently between low and high levels) appear to favor large-celled species with higher nutrient storage abilities [41,42] and can lead to metabolite synthesis and high lipid accumulation [16]. In order to optimize the accumulation of lipids in diatoms, recent work has induced stress in a culture grown under optimal conditions to induce lipid synthesis and demonstrated positive lipid accumulation [16].

Several steps are involved in the production of lipids from microalgae, including flocculation and lipid extraction. Once harvested, the culture needs to be concentrated in order to reduce the costs of downstream processing [43]. Flocculation appears to be the most advantageous way of dewatering the culture due to its low energy requirement [16]. During flocculation, microalgae cells adhere to one another to form heavy aggregates which then settle to become concentrate that is dried and milled into a fine powder. Biological based methods of flocculation use extracellular polymeric substance such as polysaccharides and proteins, originating from microalgae or other microorganisms [44]. On the other hand, Chemical based methods of flocculation use inorganic and organic flocculants such as electrolytes and synthetic polymers [45].Recently, low cost flocculation methods using pH showed flocculation efficiencies >90% when used on high density of freshwater microalgae [46].

Microalgae synthesize several types of lipids which can be classified into two categories based on the polarity of their molecular head group [47]: (1) neutral lipids which comprise glycerols, sterols, fatty acids and TAGs and (2) polar lipids which can be further sub-categorized into phospholipids (PL) and glycolipids (GL). Neutral TAGs are used primarily in microalgal cells for energy storage, while polar lipids are structural lipids packed in parallel to form the bilayer of the diatom cell membranes. TAGs are the lipids of highest value for bio fuel production since they undergo trans-esterification where TAGs react with methanol to produce biodiesel and fatty acid methyl esters [48]. The lipid content of diatoms is species-specific but also depends on the method used for extraction. Traditional methods have been using solvents and a mixture of hexane/ isopropanol (3/2 v/v) has been suggested as a low-toxicity option [49] which is selective towards neutral lipids [50-52].The segregation of neutral lipids (i.e., glycerols, sterols, PUFAs and TAGs) is highly desirable as it would allow microalgal biodiesel production to occur with minimal downstream purification. The use of supercritical fluid extraction (SFE) could be a greener option replacing the traditional use of organic solvent. Supercritical carbon dioxide (SCCO2 ) is the primary solvent used in the majority of SFE [49]. Its moderate critical pressure (72.9 atm) allows for a modest compression cost, while its low critical temperature (31.1 ° C) enables successful extraction of thermally sensitive lipid fractions without degradation. SCCO2 facilitates a safe extraction due to its low toxicity, low flammability, and lack of reactivity [53,54]. 

Depending on the bio product being developed, further processing might be needed once the lipids are extracted from the diatoms. It is then essential to take the production of biodiesel with the production of other metabolites analyzed for protein, carbohydrates and cellulose content into consideration. In order to produce bio fuel, the extracted lipids have to be subjected to trans-esterification. The purpose of esterification is to lower fatty acid evaporation temperature by changing the lipid functional group into methyl esters (biodiesel fuel) before GC-MS analysis. The trans-esterification methods have been described [55] and successfully used on diatoms [56] to produce biodiesel using methanol (alcohol), sulphuric acid (catalyst) and hexane (solvent). The alcohol and catalyst are needed to change the lipid functional group into methyl esters, while the solvent will allow for the liquid-liquid separation of the biodiesel from the by-products by centrifugation.

Molecular manipulations

The molecular basis for the ecological success of diatoms is largely unknown. The recent sequencing of the whole genome of Thalassiosirapseudonana [19] and Phaeodactylumtricornutum [57] indicates that these organisms have particular metabolic pathways that might partially explain their extraordinary adaptations to a very wide range of habitats and environmental conditions. Successful genetic transformation has been reported for diatoms [58-64] and in many cases the transformation resulted in the stable expression of trans genes from either the nucleus or the plastid [65]. Since one of the strategies to increase lipid accumulation would be to increase lipid catabolism, the genes responsible for the activation of TAG and free fatty acids would have to be over expressed to increase the cellular lipid content. This has recently been tested on P.tricornutum [66]. DNA transformation could be the solution to achieve gene over expression and successful genetic engineering of the diatoms.

The morphological and physiological plasticity of diatoms allows them to respond very quickly to environmental stress. Preliminary work on lipid production in diatoms [18,19] suggests that it might be possible to shift the metabolism of diatoms towards high lipid accumulation and storage, using stress triggers.

Although several microalgal genomes have been sequenced [19,57,67-69], pathways related to algal lipid biosynthesis remain poorly characterized. However, the genes encoding a complete set of enzymes for several types of (PUFA) biosynthesis pathways were identified in two diatoms: T.pseudonana and P.tricornutum [19,70,71] and a TAG pathway has been hypothesized by [72]. These provide a molecular basis for PUFA and TAG biosynthesis in diatoms in the future.

Despite the gap still existing in the knowledge of lipid metabolism at molecular level in diatoms, recent progress in genome sequencing has indicated how their lipid-biosynthesis pathways differ from plants and other model organisms. Current work undertaken on different species of diatoms will help further deciphering the pathway and will provide the framework for metabolic engineering of diatoms for the production of bio products.

I would like to thank J. Jendyk for proofreading the manuscript.

1. Falkowski PG, Barber RT, Smetacek V V. Biogeochemical Controls and Feedbacks on Ocean Primary Production Science. 1998; 281: 200-207.

2. Treguer P, Pondaven P. Global change. Silica control of carbon dioxide Nature. 2000; 406: 358-359.

3. Buesseler KO. The decoupling of production and particulate export in the surface ocean. Global Biogeochem. Cycles. 1998; 12: 297-310.

4. Kolber ZS, Plumley FG, Lang AS, Beatty JT, Blankenship RE, VanDover CL, et al. Contribution of aerobic photoheterotrophic bacteria to the carbon cycle in the ocean. Science. 2001; 292: 2492-2495.

5. Sarthou G, Timmermans KR, Bain S, Tréguer P. Growth physiology and fate of diatoms in the ocean: a review. J. Sea Res. 2005; 53: 25-42.

6. Tréguer P, Nelson DM, Van Bennekom AJ, Demaster DJ, Leynaert A, Quéguiner B. The silica balance in the world ocean: a reestimate. Science. 1995; 268: 375-379.

7. Irigoien X, Harris RP, Verheye HM, Joly P, Runge J, Starr M, et al. Copepod hatching success in marine ecosystems with high diatom concentrations. Nature. 2002; 419: 387-389.

8. Ianora A, Turner JT, Esposito F, Ylenia Carotenuto, Giuliana d Ippolito, Giovanna Romano, et al. Copepod egg production and hatching success is reduced by maternal diets of a non-neurotoxic strain of the dinoflagellateAlexandriumtamarense. Mar. Ecol. Prog. Ser. 2004; 280: 199-210.

9. Leterme SC, Ellis AV, Mitchell J, Thomas Pollet, Mathilde Schapira, Laurent Seuront. Morphological flexibility of Cocconeis placentula (Bacillariophyceae) nanostructure to changing salinity levels. J. Phycol. 2010; 46: 715-719.

10. Leterme SC, Prime EA, Mitchell J, Melissa H. Browna , Amanda V. Ellisc. The adaptability of diatoms to environmental changes: a case study on two Cocconeis species from hypersaline areas. Diatom Res. 2013; 28: 29-35.

11. La Vars SM, Johnston MR, Hayles J, Gascooke JR, Brown MH, Leterme SC, et al. 29Si{1H} CP-MAS NMR comparison and ATR-FTIR spectroscopic analysis of the diatoms Chaetoceros muelleri and Thalassiosira pseudonana grown at different salinities. Anal Bioanal Chem. 2013; 405: 3359-3365.

12. Rueter JG, Morel FMM. Interactions between zinc deficiency and copper toxicity as it affects the silicic acid uptake in the marine diatom, Thalassiosirapseudonana. Limnol.Oceanogr. 1981; 26: 67-73.

13. De La Rocha CL, Passow U. Recovery of Thalassiosiraweissflogii from nitrogen and silicon starvation Limnol. Oceanogr. 2004; 49: 245-255.

14. Grover JP. Resource competition in a variable environment: Phytoplankton growing according to the variable-internal-stores model. Am. Nat. 1991; 138: 811-835.

15. Stolte W, Riegman R. A model approach for size-selective competition of marine phytoplankton for fluctuating nitrate and ammonium. J. Phycol. 1996; 32: 732-740.

16. Wijffels RH, Barbosa MJ. An outlook on microalgal biofuels. Science. 2010; 329: 796-799.

17. Sheehan J, Dunahay T, Benemann J, Roessler P. Look back at the U.S. department of energy’s aquatic species program: biodiesel from algae. Close-out report. 1998; NREL/TP-580-24190.

18. Hu Q, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert M, et al. Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances. Plant J. 2008; 54: 621-639.

19. Armbrust EV, Berges JA, Bowler C, Green BR, Martinez D, Putnam NH, et al. The genome of the diatom Thalassiosira pseudonana: ecology, evolution, and metabolism. Science. 2004; 306: 79-86.

20. Kroth PG, Chiovitti A, Gruber A, Martin-Jezequel V, Mock T, Parker MS, et al. A model for carbohydrate metabolism in the diatom Phaeodactylum tricornutum deduced from comparative whole genome analysis. PLoS One. 2008; 3: 1426.

21. Borowitzka MA. Fats, oils and hydrocarbons. In Microalgal Biotechnology: Borowitzka MA, Borowitzka LJ. Eds; Cambridge University Press: Cambridge UK. 1988; 257-287.

22. Griffiths MJ, Harrison STL. Lipid productivity as a key characteristic for choosing algal species for biodiesel production. J. Appl. Phycol. 2009; 21: 493-507.

23. McGinnis KM, Dempster TA, Sommerfeld MR. Characterization of the growth and lipid content of the diatom Chaetocerosmuelleri. J. Appl. Phycol. 1997; 9: 19-24.

24. Schlechtriem Ch, Focken U, Becker K. Effect of different lipid extraction methods on delta13C of lipid and lipid-free fractions of fish and different fish feeds. Isotopes Environ Health Stud. 2003; 39: 135-140.

25. Hildebrand M. Diatoms, biomineralization processes, and genomics. Chem Rev. 2008; 108: 4855-4874.

26. Hildebrand M. The place of diatoms in the biofuels industry.Biofuels. 2012; 3: 221-240.

27. Round FE, Crawford RM, Mann DG. The Diatoms: Biology & Morphology of the Genera. Cambridge University Press.1990.

28. Gell P, Tibby J, Fluin J, P Leahy, M. Reid, K. Adamson, et al. Accessing immunological change and variability using fossil diatom assemblages, south-east Australia. River Res. 2005; 21: 257-269.

29. Martin-Jézéquel V, Hildebrand M, Brzezinski MA. Silicon metabolism in diatoms: implications for growth. J. Phycol. 2000; 36: 821-840.

30. Raven JA. The transport and function of silicon in plants. Biol.Rev. 1983; 58: 179-207.

31. Eppley RW, Rogers JN. Inorganic nitrogen assimilation of Ditylum bright wellii, a marine plankton diatom. J. Phycol. 1970; 6: 344-351.

32. Raven JA. The role of vacuoles. New Phytol. 1987; 106: 357-422.

33. Amano Y, Takahashi K, Machida M. Competition between the cyanobacterium Microcystis aeruginosa and the diatom Cyclotella sp. under nitrogen-limited condition caused by dilution in eutrophiclake. J. Appl. Phycol. 2012; 24: 965-971.

34. Tozzi S, Schofield O, Falkowski P. Historical climate change and ocean turbulence as selective agents for two key phytoplankton functional groups. Mar. Ecol. Prog. Ser. 2004; 274: 123-132.

35. Strzepek RF, Harrison PJ. Photosynthetic architecture differs in coastal and oceanic diatoms. Nature. 2004; 431: 689-692.

36. Kustka AB, Allen AE, Morel FMM. Sequence analysis and transcriptional regulation of iron acquisition genes in two marine diatoms. J. Phycol. 2007; 43: 715-729.

37. Morel FM, Price NM. The biogeochemical cycles of trace metals in the oceans. Science. 2003; 300: 944-947.

38. De La Rocha C, Brzezinski MA, DeNiro MJ. A first look at the distribution of the stable isotopes of silicon in natural waters. Geochim. Cosmochim. Acta. 2000; 64: 2467-2477.

39. Banse K. Cell volumes, maximal growth rates of unicellular algae and ciliates, and the role of ciliates in marine pelagial. Limnol.Oceanogr. 1982; 26: 1059-1071.

40. Klausmeier CA, Litchman E, Daufresne T, Levin SA. Optimal nitrogento-phosphorus stoichiometry of phytoplankton. Nature. 2004; 429: 171-174.

41. Grover JP. Dynamics of competition in avariable environment – experiments with 2 diatom species. Ecology. 1988; 69: 408-417.

42. Stolte W, Riegman R. A model approach for size- selective competition of marine phytoplankton for fluctuationg nitrate and ammonium. J. Phycol. 1996; 32: 732-740.

43. Danquah MK, Gladman B, Moheimani N, Forde G.M. Microalgal growth characteristics and subsequent influence on dewatering efficiency. Chem. Eng. J. 2009; 151: 73-78.

44. Nie M, Yin X, Jia J, Wang Y, Liu S, Shen Q, et al. Production of a novel bioflocculant MNXY1 by Klebsiella pneumoniae strain NY1 and application in precipitation of cyanobacteria and municipal wastewater treatment. J Appl Microbiol. 2011; 111: 547-558.

45. Zheng H, Gao Z, Yin J, Tang X, Ji X, Huang H. Harvesting of microalgae by flocculation with poly (γ-glutamic acid). Bioresour Technol. 2012; 112: 212-220.

46. Liu J, Zhu Y, Tao Y, Zhang Y, Li A, Li T, et al. Freshwater microalgae harvested via flocculation induced by pH decrease. Biotechnol Biofuels. 2013; 6: 98.

47. Kates M. Definition and classification of lipids. Techniques of lipidology isolation, analysis, and identification of lipids. Amsterdam: Elsevier Science Publisher. 1986.

48. Chisti Y. Biodiesel from microalgae. Biotechnol Adv. 2007; 25: 294- 306.

49. Halim R, Gladman B, Danquah MK, Webley PA. Oil extraction from microalgae for biodiesel production. Bioresour Technol. 2011; 102: 178-185.

50. Guckert JB, Cooksey KE, Jackson LL. Lipid solvent systems are not equivalent for analysis of lipid classes in the microeukaryotic green alga, Chlorella. J. Microbiol. Meth. 1988; 8: 139-149.

51. Lee SJ, Yoon BD, Oh HM. Rapid method for the determination of lipid from the green algae Botryococcusbraunii. Biotech.Tech. 1998; 7: 553-556.

52. Nagle N, Lemke P. Production of methyl ester fuel from microalgae. Appl. Biochem. Biotech. 1990; 24: 355-361.

53. Macias-Sanchez MD, Mantell C, Rodriguez M, E Martínez de la Ossaa, LM Lubiánb, O Monterob, et al. Supercritical fluid extraction of carotenoids and chlorophyll a from Synechococcus sp. J. Supercritical Fluids. 2007; 39: 323-329.

54. Taylor LT. Supercritical fluid extraction. New York: John Wileys& Sons, Inc. 1996.

55. Johnson MB, Wen Z. Production of biodiesel fuel from the microalga Schizochytriumlimanium by direct transterification of algal biomass. Energy Fuels. 2009; 23: 5179-5183.

56. Rekha V, Gurusamy R, Santhanam P, A Shenbaga, S Ananth. Culture and biofuel production efficiency of marine microalgae Chlorella marina and Skeletonemacostatum. Indian J Mar. Sci. 2012; 41: 152- 158.

57. Bowler C, Allen AE, Badger JH, Grimwood J, Jabbari K, Kuo A, et al. The Phaeodactylum genome reveals the evolutionary history of diatom genomes. Nature. 2008; 456: 239-244.

58. Dunahay TG, Jarvis EE, Roessler PG. Genetic transformation of the diatoms Cyclotellacryptica and Naviculasaprophila. J Phycol. 1995; 31: 1004-1012.

59. Apt KE, Kroth-Pancic PG, Grossman AR. Stable nuclear transformation of the diatom Phaeodactylum tricornutum. Mol Gen Genet. 1996; 252: 572-579.

60. Zaslavskaia LA, Lippmeier JC, Kroth PG, Arthur R. Grossman, Kirk E. Transformation of the diatom Phaeodactylumtricornutum with a variety of selectable marker and reporter genes. J. Phycol. 2000; 36: 379-386.

61. Apt KE, Zaslavkaia L, Lippmeier JC, Lang M, Kilian O, Wetherbee R, et al. In vivo characterization of diatom multipartite plastid targeting signals. J Cell Sci. 2002; 115: 4061-4069.

62. Poulsen N, Kröger N. A new molecular tool for transgenic diatoms: control of mRNA and protein biosynthesis by an inducible promoterterminator cassette. FEBS J. 2005; 272: 3413-3423.

63. Poulsen N, Chesley PM, Kröger N. Molecular genetic manipulation of the diatom Thalassiosirapseudonana (Bacillariophyceae). J. Phycol. 2006; 42: 1059-1065.

64. De Riso V, Raniello R, Maumus F, Rogato A, Bowler C, Falciatore A. Gene silencing in the marine diatom Phaeodactylum tricornutum. Nucleic Acids Res. 2009; 37: 96.

65. Radakovits R, Jinkerson RE, Darzins A, Posewitz MC. Genetic engineering of algae for enhanced biofuel production. Eukaryot Cell. 2010; 9: 486-501.

66. Niu YF, Zhang MH, Li DW, Yang WD, Liu JS, Bai WB, et al. 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.

67. Derelle E, Ferraz C, Rombauts S, Rouzé P, Worden AZ, Robbens S, et al. Genome analysis of the smallest free-living eukaryote Ostreococcus tauri unveils many unique features. Proc Natl Acad Sci U S A. 2006; 103: 11647-11652.

68. Merchant SS, Prochnik SE, Vallon O, Harris EH, Karpowicz SJ, Witman GB, et al. The Chlamydomonas genome reveals the evolution of key animal and plant functions. Science. 2007; 318: 245-250.

69. Price DC, Chan CX, Yoon HS, Yang EC, Qiu H, Weber AP, et al. Cyanophora paradoxa genome elucidates origin of photosynthesis in algae and plants. Science. 2012; 335: 843-847.

70. Guihéneuf F, Leu S, Zarka A, Khozin-Goldberg I, Khalilov I, Boussiba S. Cloning and molecular characterization of a novel acylCoA:diacylglycerol acyltransferase 1-like gene (PtDGAT1) from the diatom Phaeodactylum tricornutum. FEBS J. 2011; 278: 3651-3666.

71. Gong Y, Zhang J, Guo X, Wan X, Liang Z, Hu CJ, et al. Identification and characterization of PtDGAT2B, an acyltransferase of the DGAT2 acyl-coenzyme A: diacylglycerol acyltransferase family in the diatom Phaeodactylum tricornutum. FEBS Lett. 2013; 587: 481-487.

72. Obata T, Fernie AR, Nunes-Nesi A. The central carbon and energy metabolism of marine diatoms. Metabolites. 2013; 3: 325-346.