Discovery of a new species of Heterococcus and analysis of its lifecycle, genome, and lipid production A DISSERTATION SUBMITTED TO THE FACULTY OF THE GRADUATE SCHOOL OF THE UNIVERSITY OF MINNESOTA BY David Roy Nelson IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY Paul A. Lefebvre September 2012 © David Roy Nelson, 2012 i Acknowledgements Paul (Pete) Lefebvre (advisor), Lai-wa Tam, Sinafik Mengistu, Paul Ranum, Carolyn Silflow, Doug Mashek, Mara Mashek, Chlamydomonas Center, Matt Laudon, Michael Sadowsky, Ana Muñoz, Doctoral Thesis Committee: Pete Lefebvre, David Marks, Michael Sadowsky, Ford Denison, Carolyn Silflow, University of Minnesota Imaging Center: Mark Sanders and Gail Celio, Minnesota Supercomputing Institute (MSI): Zheng Jin Tu and Ying Zhang, Funding provided by the Minnesota Futures grant and IREE (Institue for renewable energy and the environment). ii Dedication This dissertation is dedicated to my parents, Jerry Nelson and Andrea Hemphill. iii Table of Contents List of Tables ……………………………………………v List of Figures……………………………………………vi Chapter 1…………………………………………………1 Chapter 2…………………………………………………24 Chapter 3…................................................................48 Bibliography…………………………………………… 113 iv List of Figures Ch. 2 Figure 1……………………………………...……………….…………..33 Fig. 2……………………………………………………………………..57 Fig. 3……………………………………………………………………..36 Fig. 4…………………………………………………….……………….37 Fig. 5……………………………………………………..………………38 Fig. 6…………………………………………….……………………….40 Fig. 7……………………………………………………………………..42 Fig. 8……………………………………………………………………..42 Fig. 9………………………………………………………..……………43 Fig. 10……………………………………………………………………44 Ch. 3 Fig. 1……………………………………...……...……………...………55 Fig. 2……………………………………………………………….…….57 Fig. 3……………………………………………………..………………58 Fig. 4……………………………………………………………………..61 Fig. 5……………………………………………………………………..62 Ch. 4 Fig. 1……………………………………………………………………..79 v Fig. 2…………………………………………………………………….80 Fig. 3…………………………………………………………………….82 Fig. 4…………………………………………………………………….83 Fig. 5…………………………………………………………………….84 Fig. 6…………………………………………………………………….85 Fig. 7…………………………………………………………………….91 Fig. 8…………………………………………………………………….92 Fig. 9…………………………………………………………………….93 Fig. 10…………………………………………………………………...94 Fig. 11…………………………………………………………………..98 Fig. 12…………………………………………………………………..99 Fig. 13…………………………………………………………………100 Fig. 14…………………………………………………………………101 Fig. 15…………………………………………………………………102 Fig. 16…………………………………………………………………104 vi List of Tables: Ch. 2 Table 1 ………………………………….……………………………35 Ch. 3 Table 1………………………………………………………..……..67 Ch 4 Table 1……………...………………………………………………74 Table 2………………………………………………...…………….74 Table 3………………………………………………...…………….77 Table 4…………………………………….…………...……………86 Table 5…………………………………………………...………….91 Table 6………………………….………………………...…………97 1 Chapter 1: “Thesis in context” Better living through Biology Increasing population pressure and decreasing reserves of petroleum have provided incentive to discover new sustainable sources of food and fuel. Much attention has been placed on the potential of algae for nutritional purposes and as a feedstock for fuel production (Becker 2007, Simopoulos 1999). The National Renewable Energy Laboratory (NREL) was commissioned to look into this potential in 1978 with the Aquatic Species Program (ASP). Viewed in the context of production cost of algal biofuels vs. cheaper petroleum (at the time), however, the program was discontinued in 1996 due to concerns over production costs. More than 3000 species of algae had been obtained over the duration of the ASP. However, less than 150 species still remain due to limited funding for strain maintenance (Madrigal 2009). Now, as the reality of the world’s limited supply of oil becomes clearer, laboratories and corporations around the globe are revisiting the potential of algae as a fuel source. However, some already economically viable companies know that fuel is just one use for algae. Any company that is in the business of making food products, pigments, or cosmetics from algae is already able to make a good return from their investment. For example, the market for carotenoids, of which algae is the primary producer, was at 1.2 billion USD in 2010 (BCC Research, FOD025D). 2 Given the potential of algal systems to supply products of value there should be tremendous pressure to inventory the algal species available in nature to identify those that can produce useful products. However, it is currently unknown by experts in the field how many species exist in nature that are yet to be discovered. Even an intelligent guess is difficult to make, given the limited environmental sampling that has been performed. More extensive surveys of algal species and their products are necessary to elucidate the actual number of extant algal species and their potentially useful products. If we do not know about so many undiscovered species of algae we cannot find the best strain of algae for any given commercial application. Therefore, strain discovery should be a high priority for future phycology research. Prospecting for new useful species is a relatively untouched area of research, although great progress was made and lost with the ASP. Life near the freezing point An abundance of life exists even in extreme temperatures. Heat-tolerant microorganisms are known as thermophiles while cold-tolerant microorganisms are known as psychrophiles or cryophiles. Psychrophiles have an optimal growth temperature at about 15°C or lower, a maximal temperature for growth at about 20°C and a minimal temperature for growth at 0°C or lower (Moyer et al. 2007). Psychrophiles have been known to grow at temperatures ranging from -20°C to 10°C (D’Amico et al. 2006). Some cold-tolerant organisms, depending on their 3 optimal growth temperature, could also be called either psychrotolerants or psychrotrophs (Morita, 1975). Psychrotrophs can grow at low temperatures but have optimal and maximal growth temperatures above 15 and 20°C, respectively (Moyer et a.l 2007). Psychrophiles are very relevant to applications-based science and biotechnology because of the unique properties of their cellular components including certain enzymes, antifreeze proteins and exopolysaccharides (Chavecchioli et al. 2011, Margesin and Feller 2010, Nicolaus et al. 2010, Paredes et al. 2011, Piette et al. 2011, Wilson and Walker 2010, Xiao 2010). As an example the cellulases from psychrophiles find applications in molecular biology, the food industry, and environmental bioremediation. The metabolism of psychrophiles is much different from that of mesophiles. Psychrophiles raise levels of intracellular adenosine 5'-triphosphate (ATP) as temperatures decline so as to keep available energy high in lieu of decreased molecular motion and thermal energy (Feller 2007). In addition, psychrophiles tend to have less adenosine 5'-monophosphate (AMP) degradation enzymes and more AMP synthetic enzymes than mesophiles (Parry and Shain 2011). Keeping AMP levels high for ATP synthesis is an excellent example of how psychrophiles deal with lower temperatures. In fact, Perry and Shain (2011) found that manipulating AMP metabolism in E. coli led to its growing up to ∼70% faster at low temperatures. 4 In addition to metabolism, a host of other cellular processes have undergone adaptation by psychrophiles to function at low temperatures. The most critically important requirement for life at low temperatures for psychrophiles is the maintenance of functional cellular membranes at exceptionally low temperatures (Thomas and Diekman 2002). Because of the deleterious effects of low temperature on membrane fluidity and enzyme catalysis, the composition of membranes and enzymes must be selected to have favorable characteristics to function at extrememly low temperatures (Goodchild et al, 2004, Mykytczuk et al. 2010, Ratkowsky et al, 2005, Siddiqui and Cavicchioli 2006). Enzymes and chromatin structure must also be folded properly, which mesophillic organisms cannot do at low temperatures, hence a need for psychrophillic adaptations to these components as well (Siddiqui and Cavicchioli 2006). Algae in cold climates Among the myriad species of organisms adapted to cold climates, algae are primary producers of nutrients essential for the other organisms in the ecosystems they inhabit (Arrigo et al. 1997, Bluhm et al. 2008, Cardinale et al. 2011). Algae bring organic carbon into cold ecosystems that can then be used by bacteria, fungi, and higher organisms, although cold ecosystems in general sequester less carbon than most other ecosystems on Earth (Callaghan et al. 2004). The variation of productivity in cold ecosystems is due to the duration/depth of winter snow cover, degree of protection from winter wind 5 damage, variation in soil moisture, soil thaw, and soil temperature (Callaghan et al. 2004). Local variation in these factors can be nearly as great as that across a wide range of latitudes (Shulze et al. 2001). Environmental suitability may be why the same genus of algae (Heterococcus) can be found in Antarctica and Colorado: the climate on the top of a mountain is very similar to some areas in Antarctica. Thus understanding the ecophysiology of arctic algae may help in understanding the ecophysiology of alpine algae. Unfortunately, due to global warming, major shifts in arctic environments have been occurring that may have resulted in massive losses of algal species (Smol et al. 2004). However, vast numbers of algal species still populate northern and southern arctic areas and these areas are the most popular destinations for researchers interested in cold-adapted algae. Algae species in the northern Arctic include Chlamydomonas nivalis, Scotiellopsis sp., Klebsormidium flaccidum, Zygnema sp., Meridion circulare, Tabellaria fenestrata and Fragilaria sp. (Kim et al. 2008). Algae species in the southern arctic (Antarctica) include species such as Pheocystis antarctica and Stichococcus bacillarus (Chen et al. 2012) as well has a number of species of Heterococcus. The species of algae found in cold climates span many diverse taxa. Green algae (Gorton et al. 2010), yellow-green algae (Rybalka et al. 2009), red algae (Lohrmann et al. 2004), golden algae and picobiliphytes (Weber et al. 2012) have all been recovered from cold environments and studied to various extents. 6 Algae that live in cold climates have different responses to environmental changes and different fitness levels depending on the environmental changes involved. Cyanobacteria have different responses to cold, wind, sun, etc. than do eukaryotic algae. Cyanobacteria more readily respond to strong fluctuation of solar radiation levels, desiccation cycles, salinity and freeze-thaw cycles by modulating expression of adaptive genes to mitigate these stresses (Callaghan et al. 2004). Overall, prokaryotic psychrophiles are more resistant to harsh conditions. On the other hand, eukaryotic algae cannot respond as readily to extreme environmental changes but do have higher rates of photosynthesis (Callaghan et al. 2004). Thus given more extreme cold conditions the prokaryotic organisms would out-compete the eukaryotic algae and in more mild conditions the eukaryotic algae would out-compete the prokaryotes. Typical locations of cold-adapted algae are in cold ocean water, snow fields in places such as Antarctica and Siberia, within pack ice, and atop mountains. Algae in arctic deserts can be found under rocks and in crevices (Thomas 2005). Algae in pack ice can even be found in freeboard layers (layers at sea-level covered by snow) but are more likely to be found high in floes where sunlight is more abundant (Brierly and Thomas 2002). Algal growth in these areas in pack ice can be so much as to contribute to ice melting from increased absorption of sunlight (Ackley and Sullivan 1994). Algae can also be found in glaciers, which may explain the vast geographical spread of some families of algae. Glacial algae tend to be found in 7 cryoconite holes, medial moraines, and supraglacial kames (Stibal et al. 2006). In glaciers, the highest density of algae is found in cryoconite holes (holes made from absorption of sunlight by dark sediment) because these locations receive nutrient-rich sediments blown by wind and washed by rains (Stibal et al. 2006). Likewise, any area where nutrients accumulate is likely to support greater populations of algae than exposed surfaces. For example, there is likely to be much more algal growth in the trough of a mountain side than the peak or ridges. Water containing sediment and avian waste is more likely to flow through these trough areas and bring nutrients to algae supporting growth. Increased flooding in snowy upper layers of glaciers has been shown to increase primary production by algae (Arrigo et al. 1997). However flooding does not imply homogenous distributions of algal species in glaciers or pack ice. Large-scale secondary pores such as cryoconite holes may contain highly variant ecosystems unique to each hole. Such uniqe individual sub-ecosystems create a high degree of variation of algae species in geographically similar locations in pack ice (Eicken et al. 1991). Many cold-adapted algae are studied as symbiotes in association fungi and perhaps one or more species of bacteria, commonly known as lichens. Lichens are incredibly cold-resistant (Ascaso and Wierzchos 2002, Cornellisson et al. 2007. In some colder climates lichens provide the dominant food source for some primates (Black-and-white or Yunnan snub-nosed monkeys) specifically because of their abundance at cold temperatures (Grueter et al. 2009). Many 8 lichens have been shown to benefit from being frozen over (Bjerke 2009). Ice encapsulization has been shown to increase productivity and photosynthesis compared to non-encapsulated lichens in the same environment (Bjerke 2009). Alpine algae have not been extensively studied in most alpine areas of the world and are not well characterized (Novis et al. 2011). A few eukaryotic isolates have been studied in New Zealand, however the small number of isolates found are likely only a small percentage of the alpine algae that exist throughout the world (Novis 2002, Novis et al. 2008). In addition to cold-tolerance, alpine algae must also be resistant to the high levels of solar irradiation found above the tree line. Solar radiation has been shown to affect growth, photosynthesis, nitrogen incorporation and enzyme activity in photosynthetic microbes (Hader 2000). Alpine algae must have in place mechanisms to repair or prevent double-strand breaks from higher gamma ray levels and thymine dimers caused by higher-than-normal levels of UV radiation (Marquis et al. 2009, Vlcek et al. 2008). A simple way to deal with this problem among alpine life is simply to increase pigmentation or accumulate specific types of protective pigments including certain carotenoids and xanthins (Gorton et al. 2001, Holzinger et al. 1993, Takaichi 2011). Whatever adaptation acquired, some algae seem to have become impervious to the normally damaging effects of solar radiation. For example, the arctic freshwater alga Zygnema sp. is completely insensitive to experimental UV radiation reaching up to 1300 µmol photons m-2 s-1 (Holzinger et al. 2009). 9 Oil production in psychrophiles In order to keep biological membranes from becoming too rigid in cold temperatures, psychrophiles must maintain a membrane composed of fatty acids with high degree of unsaturation (Quinn 1988). In general the colder the normal environmental temperature for an organism, the more double bonds it will have in its membrane fatty acids and the longer these fatty acids will be. Fatty acids with chain lengths of 16-18 C and 0-3 double bonds comprise the majority of membranes in mesophiles while fatty acids with chain lengths of >20 C and more than 3 double bonds are more abundant in psychrophile membranes. This correlation is directly due to the thermodynamics of membrane structure and hydrocarbon plasticity. In addition, psychrophiles tend to store more oil than mesophiles to compensate for the long and dark winters often found in cold climates (Logue et al. 2000). During the winter carbon stores are depleted. Photosynthesis is limited as day length shortens and the light intensity declines as the sun lowers in the sky. One strategy to deal with seasonal changes is to maintain an annual period of oil accumulation and a period of oil catabolism (Blumental et al. 2011). 10 Yellow-green algae The yellow-green algae, or Xanthophyceae, are a group of algae traditionally containing the orders Botrydiales, Mischococcales, Tribonematales, and Vaucheriales. Yellow-greeen algae are mostly fresh-water algae although some species inhabit marine environments (Stace 1991). Many species inhabit soil or rocky environments (Darling et al. 1987). Yellow-green algae are named for their light yellow green colour. The lightness of the colour is due to the absence of the brown pigment fucoxanthin which gives most other photosynthetic organisms darker hues of green (Stace 1991). Yellow-green algae do not accumulate starch as most other heterokonts do; instead they accumulate the storage polysaccharide crysolaminarin (Stace 1991). Classification of yellow-green algae is currently incomplete even among known species (Adl et al. 2005, Negrisolo et al. 2004). Many groupings are polyphyletic, sometimes even among genus distinctions and it has been suggested that at most two orders should be maintained within the yellow-green algae (Adl et al. 2005, Maistro et al. 2007). Current proposed orders are: Botrydiales, Mischococcales, Tribonematales, and Vaucheriales, with Tribonematales and Botrydiales considered to be polypheletic and Mischococcales considered to be paraphyletic from recent ultrastructural and phylogenetic studies (Adl et al. 2005). Relatives of the yellow-green algae include Chrysophyceae, Eustigmatophyceae, Phaeophyceae, and 11 Synurophycea. The most closely related group to the yellow-green algae are brown algae, or Pheophyceae, including macroalgae such as Ectocarpus siliculosus and Macrocystis pyrofera (Adl et al. 2005). Presented in this thesis is a newly discovered species of algae found in the Colorado Rockies, US. This species of oil-accumulating algae was identified by molecular data and morphology (chapter 2) to belong to the yellow-green algae and more specifically the genus Heteroccocus. Early characterization of yellow-green algae was limited to only very basic life-cycle analysis and culture growth studies (Miller and Fogg 1958). Modus subterraneus was used as a representative yellow-green alga for many of these studies. M. subterraneus was found to grow best on a media designed for blue- green cyanobacteria, Chu M 10 (Chu 1942). No exogenous organic compounds were found to support heterotrophic growth of yellow green algae in the dark. The addition of easily oxidizable organic compounds like sugars provided no boost to the growth of M. subterraneus (Miller and Fogg 1958), Visheria stellate or Pleurochloris communtata (Casselton 1965). However, the addition of peptone preparations like “Bacto-peptone” and “Bacto-tryptone” (Difco) resulted in higher growth rates and higher final cell mass of when added to aerated cultures of M. subterraneus in Chu M 10 medium (Miller and Fogg 1958). Exogenous carbon compounds are used by other yellow-green algae. Glucose is used as a carbon source by Chloridella neglecta, Botrydiopsis arhiza, 12 Chlorocloster solani, Bumilleriopsis peterseniana and Heterococcus caespitosus (Casselton 1965). Heterococcus is rarely found in nature. Even when Heterococcus samples can be obtained, they are only found as single cells or fragments of filaments and are very difficult to culture (Pascher 1939). The distribution of Heterococcus is restricted to alpine and polar regions, providing evidence that this genus is psychrophilic (Darling and Friedmann et al. 1987). Although the genus may be classified as psychrophilic, this newly-discovered strain, which we have named, H. coloradii, grows robustly at 22ºC. The discovered strain of Heterococcus can be characterized as psychrotolerant. Other Heterococcus species may be characterized as psychrotolerant algae as well even though no strains of Heterococcus are found outside of alpine or polar regions. The reason for the restricted biogeography of Heterococcus is probably that Heterococcus has no competitive edge at elevated temperatures. Mesophillic organisms have a steep growth advantage over strains of Heterococcus and would out-compete Heterococcus in any conceivable environment with moderate temperatures. Nevertheless, Heterococcus has previously been found to grow on four different continents: Europe, North America, Antarctica and Zealandia. The previous North American species of Heterococcus, H. canadensis, was the only species reported in North America until the discovery of H. coloradii. 13 A total of 62 species of Heterococcus are listed in Algaebase, a comprehensive website/data bank that compiles information regarding the identification, distribution, and relevant literature of land, marine, and freshwater algae (http://www.algaebase.org/). Currently the algae culture collection at UTEX (University of Texas at Austin) holds 9 species of Heterococcus although none of the species are from snow or North America. Algae in industry The goal of the ASP was to identify species of algae that could produce fuel precursors in a marine environment. Actually, algae have been used for hundreds of years to make a variety of products useful to humans (Sanghvi et al. 2011). Now the algae industry uses algae to make products ranging from foodstuffs to medicine to cosmetics (Becker 2007, Christaki et al. 2011, Imhoff et al. 2011, and Sanghvi et al. 2010). Among the metabolites of algae used for industrial purposes are: fatty acids, halogenated compounds, steroids, lectins, carotenoids, polysaccharides, amino acids, polyketides and toxins (Cardozo et al. 2007). Over 15,000 novel compounds have been discovered from various species of algae so far (Cardozo et al. 2007). Algae are desirable to use as feedstock for farm animals because of their nutrient content. Algae grown as a feedstock for animals is high in nutritional fatty acids, vitamins, fiber, minerals, carbohydrates and proteins (Becker 2007, Cardozo et al. 2007, Nakagawa 1997, Simopoulos 1999). Animals that benefit 14 from supplementation include cattle, pigs, fowl and fish (Becker 2007). Since animals fed and supplemented with algae are for human consumption it is ultimately humans who receive the benefits of supplementing animal feed with algae. The nutraceutical industry has seen dramatically increased demand for algae as nutritional supplements, although many countries have been supplementing their diets with algae for centuries (Kahn et al. 2010). This is especially true in Asian countries, where marine algae are commonly eaten daily and for special purposes. In Japan it is very common to eat a seaweed salad before consuming sushi wrapped in Nori, or dried seaweed. In Korea, mothers traditionally consume Miyeok-guk, a hot and spicy algae soup, during the month after delivering their babies. This soup is commonly believed to supply the necessary nutrients to aid in post-natal recovery. The anticancer properties of consumed algae have been reviewed extensively (Chakraborty et al. 2009, Jiang et al. 2011, Liu et al. 2011, Pisani et al. 2002). Palmaria palmate, Laminaria setchelli, Macrocystis integrifolia, Nereocystis leukeana, Udotea flabellum, and Udotea conglutinate are species of algae that have already been shown to inhibit cancer cell proliferation in vitro (Moo-Puc et al. 2009, Yuan and Walsh 2006, Yuan et al. 2005). In addition to anticancer activity, a variety of algae have been shown to have antiviral and antiobesity activity (Kim et al. 2011). 15 Carotenoid production from algae is commercially important since algae make carotenoids in large quantities and carotenoids can be very expensive. Carotenoids made by algae include the valuable pigments astaxanthin and beta- carotene. Astaxanthin is the most expensive and important pigment made by algae (Margalith 1999). Astaxanthin is widely used as a food colorant and has many uses as a nutraceutical for its free-radical scavenging, immunomodulation and cancer prevention properties. Astaxanthin is made by the freshwater flagellate Haematococcus pluvialis (Chlorophyceae) that has red cells due to pigment accumulation. Beta-carotenes can promote vitamin A production and have been shown to prevent cancer, promote ocular health and mitigate some photosensitivy conditions (Mayne 1996). Beta-carotene is made predominantly by Dunaliella salina, an orange-colored species that can also synthesize a number of other valuable compounds (Hosseini et al. 2009). The global market of beta-carotene, dominated by D. salina- produced beta-carotene, is valued at over 261 million USD per year (BCC Research FOD 025D). Obviously the harvest of all of the biological components of an alga used for a commercial biotechnology purpose is highly desirable. Lipid/biofuel production in algae The number of literature reports regarding lipid production in algae has greatly accelerated with the recognition of algae as a potential source of biofuels. Algae are being considered as a source for fatty acids, mostly in the form of 16 triglycerides, to be processed into biodiesel. Biodiesel is the methyl-ester product formed by reacting algal lipids with methanol and sodium or potassium hydroxide. Biodiesel can be used as a drop-in substitute for normal diesel fuels provided functioning conditions are not extreme. For example, biodiesel is not suitable for use at very low temperatures because of its high gelling temperature or in high-performance vehicles because of its low flash point. Nevertheless biodiesel works well as a general purpose fuel and little to no modification is required for optimal performance of diesel engines using biodiesel fuel. For example, undergraduate students at Colorado College routinely use in-house made biodiesel from kitchen oils to power diesel-fueled grounds maintenance machines such as tractors and mowers. High lipid/hydrocarbon-producing species of algae include Botryococcus braunii (Eroglu et al. 2011, Ioki et al. 2011, MacDougal et al. 2011, Niitsu et al. 2011, Yonezawa et al. 2011), Thalassiosira psuedonana, (Jiang et al. 2012), Nannochlopsis sp. (Kilian et al. 2011, Mohammady 2011, Pal et al. 2011, Quinn et al. 2012, Simionato et al. 2011), Dunaliella sp. (Araujo et al. 2011, Chen et al. 2011, Krohn et al. 2011, Rizmani-Yazdi et al. 2011, Yang et al. 2011), and Chlamydomonas reinhardtii (Fan et al. 2011, Kropat et al. 2011, Nguyen et al. 2011, Torri et al. 2011, Wang et al. 2009, Wang et al. 2011) Dunialla tertiolecta and T. psuedonana gave lipid yields of 20-26% (Jiang et al. 2012), Nannochloropsis gave lipid yields of 47.5 ±7.1% (Radakovits et al. 2012), and Chlamydomonas gave lipid yields of 0.9-20% (Ratha et al. 2012). 17 In most algal cells, nitrogen deprivation is a strong signal for lipid production (Araujo et al. 2011, Chen et al. 2011, Jiang et al. 2012, Ratha et al. 2012), although high light (Liu et al. 2012) and sulfur (Cakmak 2012) deprivation have been used with varying effectiveness. The main strategy used for commercial production of lipids is to grow algae past logarithmic stage to stationary phase then remove all nitrogen from the media to produce the highest yields of lipids (Hannon et al. 2012). Botryococcus is a species of green algae (Chlorophyceae) that excretes hydrocarbons into its surrounding media. It has been shown to accumulate up to 55% hydrocarbon fuel precursors by dry weight (Ruangsomboon 2011). One drawback of using Botryococcus for biofuel production is its slow growth rate, as the cells average about one doubling per three days (Ashokkumar et al. 2011). Nannochloropsis is a small yellow-green alga that is widely used as a feedstock for algal biofuel production. Light intensity was not found to be a strong signal for lipid production in Nannochloropsis (Simionato et al 2011), however nitrogen deficiency was found to induce lipid accumulation (Pal et al. 2011). An extremely useful advantage to Nannochloropsis is that it has efficient homologous recombination which is highly advantageous for genetic engineering projects (Kilian et al. 2011). Dunaliella salina and Dunaliella tertiolecta are the most common strains of Dunaliella used for biofuel production. Species of Dunaliella are commonly found in salt-water environments, such as the Dead Sea in Israel or the salt flats of 18 Utah that may reach up to 5.5 M NaCl (Chen et al. 2009). In addition to biofuels, Dunaliella is commonly used for the production of beta-carotene and the production of exopolysaccharides, poly-beta-hydroxyalkanoate bioplastics, and biofuel are being investigated (Oren 2010). Chlamydomonas reinhardtii is a green alga that is a model algal system for studying cilia/flagella function (Silflow and Lefebvre 2001), as well as many other biological processes including circadian clock function (Matsuo 2011), and DNA repair in phototrophic eukaryotes (Vlcek et al. 2008). Lipid production in C. reinhardtii is triggered by nitrogen starvation (Wang et al. 2009) or iron or zinc deficiency (Kropat et al. 2011). Starchless mutants defective in an ADP-glucose pyrophosphorylase hyper-accumulate triglycerides (Li et al. 2010). This hyper- accumulation occurs presumably because energy that would normally be stored in starch is instead shunted to the triglyceride production pathways. Wang et al. (2009) documented the extensive lipid accumulation in a starchless C. reinhardtii mutant deprived of nitrogen. Other efforts to optimize C. reinhardtii for more efficient lipid production include optimizing photosynthetic efficiency. By downregulating the amount of light-harvesting proteins, a 30% increase in photosynthetic efficiency was obtained in C. reinhardtii (Mussgnug et al. 2007). Studies on biodiesel production from algae vary in predicted yields from 80 tonnes per hectare to 125 tonnes per hectare per year (Rhodes 2009). Using the most ideal estimate, 200,000 hectares of land could produce one quad, or 7.5 billion gallons, of biodiesel annually, or 205 million gallons daily. Since the U.S. 19 uses uses an average of 378 million gallons of transportation fuel a day (U.S. E.I.A. 2009), 400,000 total hectares of land for growing algae would be more than enough to supply all of the transportation fuel the U.S. requires. There is no doubt biofuel production from algae is physically possible, however the problem is its market feasibility. Estimates for the cost of biofuel production using conservative figures from the Molina Grima 2003 and Department of Energy reports, cost per gallon estimates of $1300 and $115 were made for closed and open bioreactor systems for photoautotrophically grown algae (Gao et al. 2012). These estimates used an assumed 10% lipid yield and $0.47/kg CO2 which are quite conservative since many algae are known to produce lipids at around 50% of their total mass (Rhodes 2009) However, using the generous assumptions of 30% lipid yield and $0.2/kg CO2, prices were estimated to be under $1.94/gallon (Gao et al. 2012). In order for lipid/biodiesel production to be sustainable and economically viable in algae, production will need to use photoautotrophically grown cells. Although a number of companies are using feedstocks such as acetate or sugars, these carbon compounds can be expensive and are often non- renewable. For example, Solazyme, a biofuel-from-algae company that is now traded publically, uses a patented process to create biofuels that utilizes stressed algae in the dark fed with massive amounts of sugar. A recent estimate puts 75% of the cost of algal biofuels on supplying acetate alone (where acetate is used, Menetrez 2012). Although some scientists believe that carbon feedstocks 20 are necessary to produce high yields of algal biomass (>50 g/L) in liquid media (Algal Biomass Organization, personal correspondence), we have yet to discover the correct organism(s) for photoautotrophic production of biofuels. Using photoautotrophic organisms for the production of biofuels is also important if we are to remove CO2 from the atmosphere using algal growth. In addition to its dangerous effects on global weather patterns (Fiore et al. 2012, Tollefson 2012), global warming, triggered at least in part by CO2 accumulation, has been reported to have significant impacts on human health (Epstein 2000). Use of algae by humans to lower atmospheric CO2 could offset the rise in global warming. Technologies such as the high-rate pond (HRP) method of growing algae have potential to remove CO2 from the atmosphere (Tsai et al. 2011) Researchers are working to develop bioreactors suitable for carbon sequestration (Kumar et al. 2011). Such bioreactors can connect to flue gas outlets from coal-fired power plants and significantly reduce toxic emissions such as CO2, SO(x), and NO(x) that would otherwise promote global warming. Another potential problem with algae biofuel production is the cost and shortage of fertilizers. Although Gao et al. 2012 cited CO2 as the most cost- prohibitive nutrient, other researchers believe that nitrogen and phosphorous are more problematic for large-scale algae biofuel production. Lissa Morgenthaler- Jones, the CEO of the algae company LiveFuels (CA), was quoted as saying “The truth is, neither [Venter nor Keasling] will succeed in replacing petroleum for many reasons, including the fact that [genetically modified organisms] are not as 21 robust as wild species. But what may be the biggest reason was covered by Foreign Policy months ago - the looming phosphate shortage.” However, this problem may be solved by the fact that algae are efficient scavengers of phosphate from waste water, and waste water treatment facilities are looking for ways to remove phosphate from waste water. Algae need nitrogen, phosphorous and carbon to grow and supplying the former two nutrients is so expensive so as to make fuel production cost- prohibitive (U.S. DoE, 2010). Other nutrients required are potassium, iron, manganese, sulfur, zinc, copper, cobalt, silica and calcium. All of the necessary nutrients can be supplied by using wastewater (Knud-Hansen et al. 1998). Biodiesel production from algae is unlikely to be economically feasible in the near future without the use of wastewater. Estimates of algal biomass production using municipal waste per person were between 13 and 59 g per day (Boelee et al. 2012, Young Algeneers Symposium). However, even assuming high lipid yields (%50), this would at most be 30g of lipids for biodiesel per day per person. 30g is hardly a fraction of the average commuter’s daily fuel expenditure of at least 1-2 gallons. Even so, growing algae from wastewater has the added benefit of removing nitrogen and phosphorous which would otherwise cost money. A group from Spain was able to achieve 95% removal of NH4 from wastewater effluent of 35 mg NH4/L in 6-10 days (Ferrer et al. 2012, Young Algeneers Symposium). 22 The United States currently spends $46 billion annually on wastewater treatment (U.S. CBO, 2002). The total reported wastewater flow in the U.S. is 32 billion gallons per day (U.S. EPA, 2008, CleanWatersheds Needs Survey). Assuming a 10% lipid yield (Woertz et al. 2009), a biodiesel density of 0.80 kg l-1 (Vijayaraghavan and Hemanathan, 2009) and 9 months per year operation, an average biodiesel production of 1.7 million gallons per day can be produced from wastewater-treated algae in the U.S. (Christenson and Sims, 2011). The U.S. uses an average of 378 million gallons of transportation fuel a day (U.S. E.I.A. 2009), so biodiesel produced from wastewater-grown algae would still only be less than one percent of the total fuel consumed in the U.S. In order for algal biodiesel to replace a larger fraction of our transportation fuels, non-wastewater grown algae production costs will have to be lowered dramatically and lipid production yields will have to increase dramatically. Given the wide range of estimates for photoautotrophic production of biofuel from algae we should not jump to conclusions lightly, but perservere in research attempts that may have yet unknown benefits. Even if biodiesel from algae is not economically feasible in the near future, there is still high potential for the production of specialized products such as nutritional supplements and medicines from algae. This dissertation describes the discovery and characterization of a new species of algae that may be useful for the production of lipids for biofuels and for long chain polyunsaturated fatty acids for human health. Other implications of 23 this work include the discovery of putative algal antifreeze proteins which may be useful in crop freeze-tolerance experiments. Because currently unknown species of algae exist in the world, bioprospecting and subsequent characterization of new algal species may prove to be valuable in the future. Instead of trying to engineer the perfect strain of algae for a certain commercial application, a company might utilize a strain with the desired properties already existing in nature. 24 Chapter 2: New lipid-producing, cold-tolerant species of Heterococcus isolated from the Rocky Mountains of Colorado 1 David R. Nelson2, Sinafik Mengistu, Paul Ranum, Ana Munoz, Gail Celio and Paul A. Lefebvre 1 submitted 12-2-2011 to the Journal of Phycology, accepted xx-xx-xx 2Department of Plant Biology, University of Minnesota, 250 Biological Sciences Building, 1440 Gortner ave. St. Paul, MN. Email: nels5133@umn.edu Phone: (651)335-0422 Fax: (612) 625-5754. Condensed: Novel species of Heterococcus 25 A new species of Xanthophyceae, Heterococcus coloradii, was discovered among snow fields in the Rocky Mountains. The environmental sample containing H. coloradii also contained three other species of algae and several species of fungi and bacteria, all of which were cultured in the laboratory using a minimal salts media. Axenic cultures of H. coloradii were isolated, and their cellular morphology, growth, and accumulation of lipids were characterized. H. coloradii was found to grow at temperatures approaching freezing and to accumulate large intracellular stores of lipids. Of particular interest was the accumulation of several long-chain polyunsaturated fatty acids known to be important for human nutrition. Algae that accumulate lipids in this manner have potential uses as sources of biofuels and poly-unsaturated fatty acids for human nutrition. Key index words: algae, oil, lipids, algal oil, cold-tolerant, desiccation-tolerant INTRODUCTION We describe the discovery of a novel species within the yellow-green algae recovered from the Rocky Mountains of Colorado in the United States. The presented isolate was determined to belong to the genus Heterococcus and was selected for study because of its high degree of lipid accumulation. The genus Heterococcus was established by Chodat in 1908 and encompasses more 26 than 48 species to date, many of which have been found in Antarctica (Rybalka et al. 2009). Species of Heterococcus have been recovered from Europe and Antarctica although no reports describe species of this genus from North America. Isolates are commonly found in cold terrestrial ecosystems (Rybalka et al. 2009), although H. coloradii, the new species described in this report, was cultured directly from an alpine snow sample. MATERIALS AND METHODS Culture: Snow was collected in a small water bottle at about 13,000 ft elevation on Peak 10 of Breckonridge, Colorado. Drops of the melted snow were spread on minimal Chlamydomonas agar media (1.2% agar, Sager and Granick medium I (SGI), Harris 2010) and incubated under constant light at 4 º C. Green colonies of varying morphology were transferred to liquid media using sterile toothpicks and after growth at 4º for 1 week the morphology of resulting cells was examined by phase contrast microscopy. Staining with a lipophilic dye, either Nile Red or Bodipy 505/515, showed that one of the species of algae recovered accumulated large amounts of lipids in abundant spherical intracellular droplets of varying sizes. Each colony with a unique morphology was streaked out and re-tested with Nile Red. The original colony of H. coloradii was isolated as a round colony with cell chains and tested positive for high levels of intracellular lipid stores via Nile Red staining. 27 Axenic cultures were obtained by spray-plating a small amount of liquid culture so that individual droplets formed colonies (Harris, 2010). Briefly, an aerosol of cell cultures was created by blowing filtered air over one end of a 25 µl pipet immersed in the culture solution. Agar plates were passed through the resulting aerosol cloud to allow individual cells to bind to the plate and grow, over the next three days, into colonies. The microdroplets were so small that some individual colonies contained only the single species of interest which separated it from contaminating organisms. As a final step, colonies were streaked out on plates containing three antibiotics: streptomycin (1 g/L, Sigma), ampicillin (100 mg/L, Sigma), timentin (1 g/L, Agri-bio). Colonies were picked after 2-3 weeks of growth on the antibiotic plates to create working cultures of H. coloradii. Identification: H. coloradii was initially classified by its extraordinary life cycle. After adequate cell densities were acquired for DNA extraction, a fragment of 18S rDNA was amplified using polymerase chain reaction (PCR) and the sequence of the resulting PCR product was compared with other nucleotide entries using the BLAST program at the National Center for Biomedical information (NCBI) at the National Library of Medicine. A BLAST (Basic Local Alignment Search Tool) was performed against the NCBI nucleotide database with a 322 base pair PCR-amplified fragment from H. coloradii. The top 4 BLAST results are shown in Table 1. 28 Molecular methods: DNA from the axenic cultures was isolated through either purification using a kit (Gentra) or by simply lysing the cells to release DNA for PCR using a colony PCR method (Cao et al. 2009). 18s rDNA was amplified using PCR. Thermocycling parameters were: 94°C for 5 minutes, then 35 cycles of 1 minute at 95°C, 1 minute at 56°C, and 2 minutes at 72°C. Cycling was followed by a 5 minute extension at 72°C. Pfu Polymerase (Promega) was used along with its buffer to amplify sequence according to the manufacturer’s specifications. The final PCR product was sequenced for comparison with NCBI nucleotide entries, as in Table 1. Primers used for amplification of rDNA were: CCTGCCAGTAGTCATACGCT (forward, nt 5-20, Andreoli et al. 1999) and CCCAGAAATCCAACTACGAG (reverse, nt 667-647, Negrisolo et al. 2004). Cell division and growth: Growth curves were generated for cells grown on different media including Sager and Granick I media (Harris 2010), solid and liquid BG-11, Bristol’s media, Bold’s basal media (all media recipes can be found at: http://www.sbs.utexas.edu/ utex/media.aspx) and other in-house media formulations (Fig. 6, SG1 measurements made by David Nelson, other measurements made by Sinafik Mengistu). Cells were grown under constant light (10,000 lux) and at either 22 or 4º. Growth was measured as an increase in biomass (dry weight). To measure dry weight, cells were inoculated onto 70 mm Whatman filter paper discs on top of the different agar-based media. Each filter was harvested as one sample, and triplicate samples were taken for each day 29 measurements were made. Filter paper discs with algae were baked in a vacuum oven at 80°C for 2 hours before weighing to determine dry mass. Cell counting was performed using a FACSCalibur flow cytometer (FCM) with the CellQuest Pro software. Before each run the FCM was washed with bleach for 10 min. 20 µl of a 2 ml culture was added to 1 ml of FCM running buffer and fed to the FCM with a flow rate of 12 µl/min. Each run was 50 seconds with 3 runs per sample. An FSC amplifier and FL1,FL2 SSC, FL1, FL2, FL3 and FL4 detectors were used. Amplifier/detector settings: FSC: E01v (log), SSC: 415v (log), fl1: 505v (log), fl2: 550v (log), fl3: 650v (log). Fluorescence microscopy: H. coloradii cells were stained with Nile Red in DMSO (Wang et al. 2009) for at least 2 hours prior to imaging. Cells were viewed at 1000x and 400x with a Diaplan differential interference contrast/fluorescence microscope filtered for 468 nm light and captured with a Jenoptic digital camera. Scale bars were created from measurements made with a 10 µm Carl Zeiss stage micrometer. Lipid extraction: Lipids were extracted from H. coloradii using methyl tert-butyl ether (MTBE) according to a protocol developed by Matyash et al. (2008). The authors of this protocol reported more complete yields with MTBE than with traditional hexane extractions (Folch et al. 1957). Lipid extractions were done with more than 10 different cultures that had been growing anywhere from 2 to 30 10 weeks. Cultures were divided into two parts; one part was pelleted and dried for >2 hours in a vacuum oven at 100°C and the other part was used for the MTBE extraction. Dry weights of extracted lipids were compared with dry weight of the total culture. Electron microscopy: Cultures one week old were harvested from SGI plates for observation using electron microscopy. For scanning electron microscopy (SEM), samples were placed in 2% glutaraldehyde and 0.1 M sodium cacodylate buffer for 2 hours, rinsed in 0.1 M sodium cacodylate buffer, then placed in 1% osmium tetroxide and 0.1 M sodium cacodylate buffer for 2 hours. Specimens were rinsed in ultrapure water (NANOpure Infinity®; Barnstead/Thermo Fisher Scientific; Waltham, Maryland) and dehydrated in an ethanol series. After the samples were in 100% ethanol, they were put through two changes of hexamethyldisilazane (HMDS) for 5 min each. Drops of the suspension were placed on individual acetone-cleaned round glass cover slips that had been mounted on aluminum stubs with double-sided carbon adhesive tabs, and allowed to air dry. The specimen stubs were sputter-coated with gold-palladium and observed in a Hitachi S3500N scanning electron microscope at an accelerating voltage of 12 kV. For transmission electron microscopy (TEM), samples were fixed as for SEM. Following the ultrapure water rinse, the samples were embedded in low melting point agarose. The samples were cut into 1-mm3 pieces, dehydrated in 31 an ethanol series, and embedded in Embed 812 resin (Electron Microscopy Sciences, Hatfield, Pennsylvania). Ultrathin sections 80–100 nm thick were cut on a Leica Ultracut UCT microtome using a diamond knife and collected on formvar/carbon-coated copper mesh grids. Sections were post-stained with 3% uranyl acetate followed by Sato’s triple-lead stain (Sato 1968), and examined with an FEI Phillips CM 12 transmission electron microscope operating at 60 kV. Images were recorded with a Maxim DL digital capture system. Desiccation and freezing treatments: SGI Agar plates with healthy H. coloradii cultures were allowed to air-dry over a period of six months. After complete desiccation, cells were scraped from the plates onto fresh SGI plates to test for growth. H. coloradii on BG-11 plates were placed in darkness in -20°C freezers for 2 weeks to test for freezing tolerance. After 2 weeks, plates were taken out to test for growth at either 4°C or 22°C in the light. RESULTS Biological composition of snow sample: Six different species of algae and bacteria were recovered from the snow sample. Two species of bacteria and four species of eukaryotic algae were identified. In addition to H. coloradii, the samples were found to contain: Pseudomonas antarctica, 99% identity from BLAST with PCR-amplified 16s 32 rDNA (Genbank accession: BankIt1479737 seq JN661817); Rhizobium sp., 98% identity from BLAST with PCR-amplified 16s rDNA (Genbank accession: BankIt1479737 seq. JN661816); Chlamydomonas reinhardtii, 100% identity confirmed via mating with a wild-type lab strain of Chlamydomonas; Chlorella sp. 99% identity from BLAST with PCR-amplified 18s rDNA (Genbank accession: BankIt1479737 seq2 JN661815). Classification: H. coloradii was initially classified by the similarity of its morphology to other species of Xanthophyceae. Specifically, the cells had yellow-green chloroplasts, no starch, and swimming zoospores. Swimming cells have two flagella of unequal length (Fig. 1a). The long flagella is 7-10 µm long and the short flagella is 1.5-4 µm long. Morphology of H. coloradii is mostly congruent with reports of other species of Heterococcus from Antarctica and Europe (Lokhorst 1992) except for its unusually high lipid production and size. H. coloradii is the first species of Heterococcus to be discovered in North America. Heterococcus normally grows as cell chains or in a coccoid form (Fig. 1e, 1f, 1g). We use the term “cell chains” instead of “filaments” so as not to be confused with fungal-type hyphae. Heterococcus cells are usually uninucleate (Lokhorst 1996) but our strain is multi-nucleate, containing 1-3 nuclei per cell (fig 2). A closely related genus, Botrydiopsis, is multinucleate and grows as large coccoid cells. The closeness of the morphology and molecular phylogeny of H. coloradii to Botrydiopsis might suggest that H. coloradii is actually a species of 33 Botrydiopsis. However, the presence of cell chains in H. coloradii alone is reason to classify it as a species of Heterococcus. Figure 1: Stereo microscopy (f) and DIC microscopy (a-d, g-h 1000x, e 250x) of H. coloradii cells. a: biflagellate swimming cell b: Coccoid H. coloradii budding off into cell chain growth. c: H. coloradii cell undergoing cytokinesis. d: H. coloradii in environmental sample shown with bacteria. e: H. coloradii cell chains penetrating into agar plates (viewed at 250x). f: extensive cell chain growth after >2 months on an agar plate (viewed at 2x). g: Fresh cell chain growth. h: Giant akinete formed after >3 months. 34 Fig. 2: Thin-section TEM of a) a non-motile coccoid cell and b) a dividing cell. Legend- nuc: nucleus (lower nucleus is labeled, organelle immediately above is also a nucleus), mito: mitochondria, lip bod: lipid body, chl: chloroplast, chrys: chrysolaminarin deposits. DNA sequence analysis was used to confirm that the new species was of the genus Heterococcus (see Molecular Methods). BLAST searches with the PCR-amplified 18S rDNA (Genbank accession: BankIt1479737 seq1 JN661814) returned several species of Heterococcus with very low e values (Table 1). H. chodatii was recovered from Lake Geneva, Switzerland and is synonymous with H. viridis. H. chodatii displays isolated outgrowths of cell chains that are not seen in H. coloradii. H. caespitosis was recovered from Freiburg, Germany in wet, loamy soil. H. caespitosis is very similar to H. coloradii morphologically 35 except for H. caespitosis’s distinct long cell chains that protrude from colonies. H. fuornensis was recovered from Fuorn, Switzerland from forest soil. H. fuornensis produces many immature coccoid cells from the ends of its cell chains that H. coloradii does not produce under any conditions. Although H. coloradii is similar these other species of Heterococcus, it is a unique species due to its large size, multinuclearity, and unusual oil accumulation. The consensus sequence from three of H. coloradii’s closest relative differed from H. coloradii at 26/236 nucleotides (Fig.3). This significant nucleotide divergence of H. coloradii from its closest relatives is not surprising in view of the distance of the location of the recovered species (Colorado) to the published species (Antarctica). However, considering the sequence data 36 together with morphological data we concluded that the novel field isolate was nonetheless a species of Heterococcus. Fig. 3: Alignment of 18S rDNA amplified from genomic H. coloradii DNA with 4 closest relatives according to NCBI BLAST (H. chodatii, H. fuornensis, H. caespitosis and H. pluerococcoides) using ClustalW. Lifecycle: H. coloradii cells can be found in a variety of forms, depending on age of the culture and growth conditions. Individual cells may be coccoid and immotile or may be swimming (figs. 1,5). Cells may be connected in cell chains or form giant akinetes (figs. 1,5). In liquid media H. coloradii divides via mitosis and maintains its coccoid form (Fig.1c). The age of the culture appears to govern what the cells morphology is. For the first week of growth, a newly streaked or inoculated culture will grow as coccoid cells. After one week, cell chains start to emerge. After two weeks, large spherical cells known as akinetes will appear and grow off the cell chains. Lipid accumulation commences after the first week of growth and can continue for up to six months. 37 When unperturbed by mechanical stirring, H. coloradii grows into branching cell chains (Fig.1e). These outgrowths are presumably to extend itself to gather nutrients. The chain growth occurs until the cultures reach higher cell densities. Photosynthetic cells deprived of nutrients lose chlorophyll although the more stable carotenoids will remain (Phadwal et al. 2003). When inoculated into fresh media the cells are dark green but as they use the available nutrients they shift to a bright yellow, perhaps as a consequence of nutrient depletion. As the cells change color, akinetes (Fig.1h) start to emerge. These are the cells that become completely engorged with lipids. Although all of the cell types of H. coloradii accumulate lipid bodies, the akinetes accumulate them to the highest degree and dedicate the largest intracellular volume to these bodies (Fig.4, top right cell in each panel). Fig. 4: 1000x DIC (left) and Nile Red fluorescence (right) images of H. coloradii showing oil accumulating as oil bodies (left and bottom cells) that eventually fill the intracellular space (upper right cell). 38 H. coloradii cells mature from a coccoid form to grow chains and then to form akinetes (Fig.5). Not all cells mature to form akinetes as some of the chain cells simply fill with lipids but never release zoospores. If the akinetes are inoculated into media with higher nitrogen levels they produce swimming zoospores that can number more than fifty per akinete. Zoospores may also contain large lipid bodies. Zoospores eventually stop swimming and mature into coccoid cells and the life cycle of H. coloradii starts anew (Fig.5). Fig. 5: Life cycle of H. coloradii. A: vegetative cell growth in coccoid form. Organelles shown are chloroplasts. B: Cell chain formation. C: akinete. D, 39 E: Cells produce lipid bodies(D) or zoospores (E) depending on nutrient availability. F: Swimming cells (zoospores). Lipid accumulation: Lipid accumulation as visualized by Nile Red fluorescence was shown to fill up a majority of the intracellular volume. Lipids accumulate as spherical droplets that could range in size from 0.1 µm in diameter to 10 µm. Akinetes that had accumulated lipids could grow to larger than 100 microns in diameter (Fig. 1h). Fig. 4 shows mature cells with fluorescent lipid bodies stained with Nile Red. Lipids were determined gravimetrically using an advanced extraction method (see materials and methods). On average a mature culture of H. coloradii is composed of 55% lipids by dry weight. Preliminary studies analyzing lipid composition using GC/MS indicated that certain high-value polyunsaturated fatty acids are present in significant amounts in H. coloradii. Nutrients and Growth: Growth of H. coloradii was measured on several media designed for algal growth. Fig. 6 contains growth curves of H. coloradii for the four most effective media solutions used. Vegetative growth is characterized by cell division and chain growth, while non-vegetative growth is characterized by cell enlargement, akinete formation, and lipid accumulation. We found via flow cytometry that H. coloradii grew poorly in liquid media. Optimal growth was 40 observed on either filter paper or cellophane discs placed on top of the agar media. Although the main media commonly used to grow Heterococcus species is Bold’s Basal Media (BBM, Lokhorst 1992), we found that SGI media was superior to other media tested in promoting rapid growth of H. coloradii (Fig.6). BG-11 media, however, was more effective at keeping cultures greener and viable for a longer time as observed during routine passing of cultures. 41 Fig. 6: Growth of H. coloradii was tested on different media. For SGI (M) chart: ■ = 22°C, ◆ = 4°C. Weight is shown as total dry mass of H. coloradii. Co-cultured bacteria: When H. coloradii was originally isolated as a single colony from the environmental sample, bacterial contamination was observed (Fig.1d, Fig.7). Two strains of bacteria were isolated (see Biological composition of snow sample section) and added back to axenic cultures of H. coloradii. The species of Rhizobium was found to have a significant physical interaction with H. coloradii as observed by SEM microscopy (Fig. 7) and was found to enhance growth of H. coloradii (Fig.8) as well as to increase viability under nitrogen deficient conditions (Fig.9). 42 Fig. 7: SEM image of Rhizobium sp. attached to H. coloradii cell. Figure 8: Growth of axenic H. coloradii compared with H. coloradii with Rhizobium added (appx. one toothpick per 2 ml well) 43 Fig. 9: Chlorophyll content of H. coloradii cells after one month of nitrogen starvation with and without co-cultured Rhizobia. Desiccation and freezing survival: Some species of algae are known to survive long periods of desiccation (Gray et al. 2007). This capability, presumably an adaptive response to environmental conditions faced by the alga, could be useful for various commercial applications. We found that H. coloradii shows substantial tolerance to desiccation and cultures grow readily after desiccation treatments (see Materials and Methods). Desiccated cells are shown in Fig.10 to remain intact and maintain their lipid stores. H. coloradii was found 44 to survive periods of dark at -20°C. However, while H. coloradii could recover from freezing if placed at 4°C, cultures could not recover from freezing if they were thawed at 22°C. Fig. 10: Intact dessicated H. coloradii cell shown at 1000x DIC magnification DISCUSSION Reports on Xanthophyceae species are sparse in the scientific literature and many species are yet uncharacterized or entirely unknown (Ettl 1978). In fact, even the phylogenic distinctions of the genera within the Xanthophyta are dubious, and much debate surrounds the polyphyletic groupings of species (Maistro et al. 2009, Ehara et al. 1997). Despite the ambiguity surrounding the yellow-green algae phylogenetic classification system we have placed the newly discovered species into the genus of Heterococcus and give it the species name of coloradii because of the molecular data and its location of origin. 45 H. coloradii was found to exhibit a variety of morphologies depending on the age of the culture and environmental conditions. As H. coloradii grows it creates structures that withstand harsh environmental conditions such as freezing and/or drying. Once nutrients are restored, spores swim out and away from colonies for dispersion. The new species of Rhizobium found with H. coloradii attaches to the outer cell wall and may promote growth. It is yet unknown exactly how the Rhizobium would promotes the growth of H. coloradii but options to speculate upon include: possible nitrogen fixation by the Rhizobium, possible vitamin supplementation provided by the Rhizobium, or the extra CO2 provided by the Rhizobium is needed in an aquatic environment. Although acetylene reduction assays have been negative, it is possible that the proper environmental conditions for nitrogen fixation by an H. coloradii/Rhizobium consortium have not yet been found. Promotion of algal growth was found in early studies with flow cytometry but has been unable to be reproduced in future studies. We are particularly interested in what H. coloradii’s lipid accumulation could mean in a commercial context. Algae have great potential to produce food, fuel, and medicine (Rasala et al. 2011). Among the most promising commercial applications for algae are the production of lipids for biodiesel production (Christianson et al 2011, Craggs et al. 2011, Hannan et al. 2010, Stephenson et al. 2011, Wiley et al. 2011) and high value lipids for human nutrition (Cockbain et al. 2011, Dangat et al. 2011, Doughman et al. 2007, Dewey et al. 2011, 46 Hegarty et al. 2011, Rhodes et al. 2009, Yeste et al. 2011). H. coloradii produces large amounts of intracellular lipids, making it a candidate for commercial lipid production. In fact, cells appear to fill completely with lipids before dying, leaving intact lipid containers as their remains. Algae that can grow in extreme conditions and still accumulate lipids are of great interest to industry. H. coloradii grows at 4 o C and accumulates large intracellular stores of lipids. Thus, countries in higher latitudes may be able to produce omega-3 fatty acids in the wintertime without fishing by growing H. coloradii. One problem frequently cited in algaculture is the limited size of unicellular algae cells with many species never growing over 10 microns in diameter. H. coloradii matures to create akinetes that can grow to over 100 microns in diameter, thus providing much more cellular volume that can be filled with lipids. In most species of algae, high cellular density causes significant shading due to high amounts of fucoxanthin and other pigments. Shading causes less light to reach individual cells and shaded cells produce less lipids and become malnourished. Xanthophyceae lack fucoxanthin (Stace 1991), thus dense cultures of H. coloradii are remarkably translucent compared with other algae. Light harvest may be more efficient in Xanthophycae cultures for photosynthetic lipid production. Many established algal lipid production systems rely on the alga making lipids from a supplied organic carbon compound (Wang et al. 2009). However 47 supplying exogenous carbon increases cost and it does nothing to sequester atmospheric or power-plant produced CO2. An alga that produces lipids completely photosynthetically would be ideal. H. coloradii photosynthetically produces lipids solely from atmospheric CO2 and does not use any other exogenous carbon compounds for growth or energy storage. It is unknown exactly how many algal species are yet to be discovered but undiscovered species have great potential to possess characteristics useful for commercial applications. Thus more effort should be placed on the discovery and characterization of new strains of algae. 48 Chapter 3: Lipid composition of Heterococcus coloradii and response to variations in temperature and light. David R. Nelson1, Gail Celio, Mara Mashek2, Douglas Mashek2, and Paul A. Lefebvre1 1Department of Plant Biology, University of Minnesota, 250 Biological Sciences Building, 1445 Gortner Ave. St. Paul, MN. Phone: 651-335-0422. Email: nels5133@umn.edu. Fax: (612) 625-5754 2Department of Food Sciences and Nutrition, University of Minnesota index: EPA: eicosapentaenoic acid, DHA: docosahexaenoic acid, PA: palmitoleic acid 49 H. coloradii is a recently discovered species of yellow-green algae (Xanthophyceae) that grows at low temperatures and produces large amounts of intracellular lipids. Lipids accumulated by H. coloradii were analyzed using gas chromatography and mass spectrometry (GC/MS) and found to contain high levels of palmitoleic acid (PA, 16:1) and eicosapentaenoic acid (EPA, C20:5). PA and EPA are known to provide substantial health benefits, making H. coloradii an attractive organism for producing these lipids for human nutrition. We established the environmental conditions needed for production of lipids in H. coloradii. Various environmental stresses including extended periods of darkness or heat lower the proportion of high-value lipids in H. coloradii. H. coloradii produces the highest quantity of high-value lipids when grown undisturbed with high light in low temperatures. Keywords: algae, oil, lipids, algal oil, cold-tolerant, dessication-tolerant INTRODUCTION The world is in need of new sources of omega-3 fatty acids for human health (Whelan and Rust 2006). Omega-3 fatty acids, in particular very-long- chain fatty acids (VLCFAs) such as eicosapentaenoic acid (EPA) and 50 docosahexaenoic acid (DHA), are currently provided to the human population by supplementing diets with fish oil. The fish used for extraction of VLCFAs are almost always ocean fish as farmed fish have much lower levels of VLCFAs (Worm et al. 2009). However, large scale human consumption of ocean fish is not sustainable because of the high levels of contaminants such as mercury in ocean fish and the risk of depleting the world’s fisheries (Whelan and Rust 2006, Worm et al. 2009). The Xanthopyceae have been noted as a group of algae that accumulate high amounts of nutritionally important VLCFAs (Lang et al. 2011). Most species within the Xanthophyceae do not accumulate starch (Belcher and Miller 1960). Instead they store energy as lipids, and some species fill up a large portion of their total volume with lipids (Broady 1976). H. coloradii is a species within the Xanthophyceae that accumulates high amounts of nutritionally important fatty acids when grown in low temperatures. Accumulated lipids in plants and algae are stored as triglycerides in lipid bodies that usually range from 0.5-2.0 µm in diameter (Huang 1996, Tzen et al. 1993, Ting et al. 1996, Shimada TL et al. 2008). We describe the nature of lipid accumulation by H. coloradii and the variance of lipid types with varying environmental conditions as well as the composition of stored triglycerides compared with total lipids. MATERIALS AND METHODS 51 Culture growth: All cultures of H. coloradii were grown on minimal salts media in 12% agar. Media used as indicated was either Minimal media (M) for growing Chlamydomonas reinhardtii with or without nitrogen (Harris 1998) or BG-11 media (cyanobacteria media, http://www- cyanosite.bio.purdue.edu/media/table/BG11.html). All cultures were grown at 4 ºC or 22 ºC and light grown cultures were grown at 1200 lux. Cells were grown on cellophane layered on top of agar plates for ease of harvest. All studies of lipid production in H. coloradii were done from cultures grown on solid media. Lipid extraction/ gravimetric measurement: Lipids were extracted from H. coloradii using methyl tert-butyl ether (MTBE) according to a refined extraction protocol (Matyash et al. 2008). Extracted lipids were weighed and compared to dry weight from an equivalent volume of the same culture for each experiment to determine percentage lipids by dry weight. Fluorescence microscopy: H. coloradii cells were stained with Nile Red in DMSO (Wang et al. 2009) for at least 2 hours prior to imaging. Cells were viewed at 1000x and 400x with a Diaplan differential interference contrast/fluorescence microscope filtered for 468 nm light and captured with a Jenoptic digital camera. Scale bars were created from measurements made with a 10 µm Carl Zeiss stage micrometer. Previous reports indicate that 10 minutes was sufficient to stain lipid bodies in a variety of algae but we found that optimal 52 staining of H. coloradii lipid bodies required at least 2 hours, perhaps because of the large size of H. coloradii lipid bodies. Fatty acid composition analysis (GC/MS): Algal cultures were scraped from plates and pelleted into 15 ml disposable plastic tubes (Falcon) by centrifugation at 2,000 RPMs on an IEC DPR 6000 centrifuge using a 949 rotor. Lipids were then extracted from the pelleted cells (Folch et al. 1957). Dried extracts were resuspended in 1 ml of 5% HCL-methanol and placed in a 90o C water bath for 1 h to saponify and methylate the fatty acids. After allowing the samples to cool to room temperature, 1 ml of water was added and the fatty acid methyl esters (FAME's) were extracted 3 times with 1 ml of hexane prior to GC analysis. Fatty acid methyl esters were extracted with multiple hexane washes followed by GC analyses with a fused silica capillary column (Supelco Omegawax, model 122- 7032), 30 m x 0.25 mm inner diameter (ID) x 0.25 µm film thickness, and Hewlett- Packard Agilent 5890 GC system with flame ionization detector (FID). The temperature program was as follows: 50 °C with a 2 min hold; ramp: 10°C/min to 250 °C with a 15 min hold. Constant pressure of 20 psi was applied throughout the run of 37 min per sample. Analyses were initiated by injection of 1 µL of sample at a split ratio of 20:1 and injector temperature of 250 °C. The FID temperature was set at 300 °C with air and hydrogen flow rates of 433 and 37 ml/min. Specific fatty acid methyl esters were identified based on retention time using a reference standard purchased from Nu-Chek Prep, Inc. Fatty acids are 53 referred to using lipid number nomenclature C:D, where C refers to the carbon chain length and D refers to the number of double bonds within the carbon chain (Rigaudy 1979). Triglyceride composition analysis: 2 grams (dry weight) of lyophilized H. coloradii (equivalent to 96h 4°C light treatment) was sent to Medallion Labs (9000 Plymouth Avenue North, Minneapolis MN 55427. 1-800-245-5615, (763) 764-4453 Fax: (763) 764-4010) for triglyceride analysis. The sample was analyzed using AOCS Method Ca 9f-57 (Table 1). Transmission electron microscopy: Cultures were harvested after one week of growth from SGI plates for observation by electron microscopy. Samples were placed in 2% glutaraldehyde and 0.1 M sodium cacodylate buffer for 2 hours, rinsed in 0.1 M sodium cacodylate buffer, then placed in 1% osmium tetroxide and 0.1 M sodium cacodylate buffer for 2 hours. Specimens were rinsed in ultrapure water (NANOpure Infinity®; Barnstead/Thermo Fisher Scientific; Waltham, Maryland) and dehydrated in an ethanol series. The samples were then embedded in low melting point agarose. The samples were cut into 1-mm3 pieces, dehydrated in an ethanol series, and embedded in Embed 812 resin (Electron Microscopy Sciences, Hatfield, Pennsylvania). Ultrathin sections (80– 100 nm) were cut on a Leica Ultracut UCT microtome using a diamond knife and collected on formvar/carbon-coated copper mesh grids. Sections were post- 54 stained with 3% uranyl acetate followed by Sato’s triple-lead stain (Sato 1968), and examined with an FEI Phillips CM 12 transmission electron microscope operating at 60 kV. Images were recorded with a Maxim DL digital capture system. RESULTS Lipid accumulation: Lipids accumulated in H. coloradii as the cultures grew and matured on agar plates. Small lipid bodies were seen to emerge after roughly 96 hours of growth and these bodies grew steadily for up to 3 months in culture. Accumulation of lipids was found to occur strictly by photosynthesis, as the addition of fixed carbon in the medium in the form of acetate, glucose or malate did not enhance lipid production. In mature cultures (> 1 month old) cells contained very large lipid bodies up to 10 µm in diameter (Fig.1b/f). At what appeared to be maximum lipid accumulation, lipid droplets filled the intracellular volume (Fig. 1). The accumulation of lipids in H. coloradii represents an extreme of algal lipid accumulation in that most cells fill completely with lipids without any special treatments such as nutrient deprivation (-N, -P, -Cu, -Mg, -Fe3+), temperature changes, light schedule changes, or addition of carbon compounds to the growth media. 55 Figure 1: H. coloradii cells viewed by DIC microscopy. a) Fluorescent Nile Red-stained lipid bodies beginning to form (100x). b) Lysed H. coloradii cell filled with lipid viewed (100x) with fluorescence (b) or DIC (f) Chlamydomonas reinhardtii starch-less mutants sta6, marked with stars, are shown for reference. c) 25x view of lipid-filled H. coloradii cells viewed with fluorescence (c) or DIC (g). d) Chain of H. coloradii cells filled with lipid viewed (100x) with fluorescence (d) or DIC (h). e) Group of lipid-filled H. coloradii cells. Cells ruptured easily under pressure on the cover slip and lipid bodies of different sizes and colors were released from H. coloradii cells (Fig. 1b/f). Lipids were not found to be released in the absence of applied mechanical force (e.g. as an inherent biological process). We conclude that the cells accumulate lipids 56 until they perish or become senescent and that the intracellular lipids continue to be stored within the confines of the cell indefinitely. Composition analysis: GC-MS analysis showed that 16:1 and 20:5 fatty acids accumulated in cultures of H. coloradii as the dominant lipid species (Figures 2- 5). To identify optimal conditions for producing these nutritionally-important fatty acids, experiments were conducted to observe how the levels of these fatty acids would change if H. coloradii cells were subjected to various environmental stresses. Among the stresses tested, temperature and light changes showed the most significant effect on fatty acid composition of H. coloradii cultures. 57 Fig. 2: Lipids in H. coloradii as determined by mass spectrometry. Lipid species named by length of carbon chain and number of double bonds: C16:3 has a 16-carbon length chain with 3 double bonds. H. coloradii cells were grown for one month at 4 °C at light levels of 1200 lux. Cells were given one of four treatments as indicated for 96 hours. After treatment, 58 cells were harvested and total lipids were analyzed by GC/MS. Any stress treatment served to lower the amount of EPA while total lipid amounts were not significantly affected. Fig. 3: All stress treatments served to reduce the amount of percentage of EPA (C20:5) as a fraction of total lipids in H. coloradii cultures. 59 The percentage of 16:1, palmitoleic acid (PA), was increased in all treatments. In cultures that received constant light at 4 ºC (control), PA levels stayed at 22%, but cultures that received dark treatment had 55% PA at 4 ºC and 47% PA at 22 ºC. Cultures that were moved to 22 ºC but still received light gained PA to 45% of their total lipids (Fig.2). EPA levels were decreased in all treatments. EPA levels fell from 25% to 10-11% when either moved to 22 ºC, moved to the dark, or received both treatments concurrently. Cultures that received dark treatment at 4 ºC had much lower levels of unidentifiable lipids (2% of total lipids) as opposed to all other cultures analyzed (15-18%). The reason for this drop in unidentifiable lipids is unknown. Total lipid amounts were largely unchanged in all treatments as measured as a percentage of dry weight (55% lipids by dry weight). Also, no degradation of lipid bodies was observed with any treatments amounting to less than one week in duration. Normally when H. coloradii lipid bodies are degraded, pits can be clearly seen on the lipid body surfaces and no pits were seen in these experiments. The relative fluorescence level of Nile Red and Bodipy dyes can be used to quantitate lipid levels but previous studies have found that fluorescence levels do not necessarily correlate well with the fat levels detected by GC-MS (Brooks et al. 2009, Zhang et al. 2010). No environmental treatment was shown to improve lipid accumulation, including high light, nitrogen starvation, bubbling CO2-containing air into liquid cultures, and changing the pH of the media. Treatments of 96 hour heat stress 60 (4 °C to 22 °C), dark stress, or both heat and dark stresses applied concurrently changed fatty acid composition (Fig. 2). In short, EPA levels were significantly reduced with the introduction of stress while the levels of the 16-carbon fatty acids increased. A detailed triglyceride composition analysis was obtained (Medallion Labs) in order to compare fatty acids in triglycerides to the total fatty acids in H. coloradii (Fig. 4). While H. coloradii was found to have 36.9% triglycerides by dry weight, we measured total lipids at 55% lipids by dry weight. Thus 33% of lipids in H. coloradii reside in species other than triglyceride. EPA was found to compose the highest percentage of fatty acids not esterified to a glycerol backbone (Fig. 4). 61 Fig. 4: Difference in fatty acid composition of total triglycerides (dark bars) compared with total fatty acids (light bars). Ultrastructural analysis: Thin-section TEM revealed that lipid bodies in H. coloradii appear to be surrounded by an intracellular membrane (Fig.5). In other organisms lipid body membranes are composed of a phospholipid monolayer 62 (Shimada et al. 2008). Lipid bodies in H. coloradii did not appear to be associated with any particular organelle. Fig. 5: Thin-section TEM of H. coloradii cell highlighting small lipid bodies. LB: lipid body, Chrys: chrysolaminarin, Mit: mitochondria, Chl: chloroplast 63 DISCUSSION Algal strains for commercial production of biofuel will need to meet three criteria: 1) high growth rate, 2) high lipid content and 3) ease of harvest and extraction (Christiansen and Sims 2011, Craggs et al. 2011, De La Hoz et al. 2011). Often one or more of these criteria are not met for an otherwise desirable strain of algae. One solution is that different strains of algae will be cultured for different purposes. A strain of algae that produces high value omega-3 fatty acids may not be ideal for making jet fuel, and vice-versa. H. coloradii meets two of the three requirements listed above, but it grows fairly slowly. Its high lipid content should produce useful quantities of nutritionally valuable lipids, and its surface-growing lifestyle should be useful for harvesting lipid products without the need for energy-requiring de-watering steps. The accumulation of lipids might serve several functions in H. coloradii. One obvious function is that of energy storage for long, dark, cold winters. Lipids are a particularly useful energy source where a slow, continuous energy supply with duration is needed. H. coloradii cells might spend the summer filling their intracellular spaces with lipids and slowly break them down over the wintertime. Cells might not be able to create photosynthetic energy due to the large amounts of snow coverage known to accumulate in the Rocky Mountains during the wintertime. Also, the extensive lipid accumulation in H. coloradii may act to increase buoyancy. For non-swimming cells, buoyancy may mean the difference between 64 a substantial harvest of photons and CO2 or starvation by darkness/anoxia at the bottom of a small pool of water. We have observed that H. coloradii does not grow well in liquid media. Thus it is possible that nature has selected for buoyant, lipid-rich cells of H. coloradii in aquatic environments. EPA is necessary for membrane stability at lower temperatures (Kawamoto et al. 2009). For example, EPA-deficient Shewanella mutants have been shown significant growth retardation at 4 oC but not at 18 oC. In cold-grown H. coloradii, EPA was found to compose the highest percentage of fatty acids not esterified to a glycerol backbone. There are two possible reasons for this: 1) EPA is maintained in free fatty acid form at high levels or 2) EPA is maintained within the cell membrane as a polar lipid species at high levels. Since EPA is usually found in cell membranes in high levels (Kidd 2007), we presume this is also the case with H. coloradii and that the EPA in the cell membrane supports membrane stability. Many health benefits of consumption of the very-long-chain omega-3 fatty acids (VLCFAs) EPA and docosahexaenoic acid (DHA) have been reported (Dewey et al. 2007, Hegarty and Parker 2011, Rhodes et al. 2009, Stebbins et al. 2010, Walser et al. 2006). These include the prevention of skin cancer (Rhodes et al. 2009), enhanced mood/decreased anxiety (Hegarty and Parker 2011), enhanced cardiovascular function (Stebbins et al. 2010, Walser et al. 2006), amelioration of cancer cachexia (Dewey et al. 2007), faster muscle recovery/anabolic effect (Smith et al. 2011), prevention of colorectal cancer 65 (Cockbain et al. 2011), prevention of atheroschlerotic plaques (Zampelas 2010), improved maternal lactation (Dangat et al. 2011), and improved sperm quality (Yeste et al. 2011). The World Health Organization recommends a dietary intake with a ratio of less than 10:1 of omega-6 fatty acids to omega-3. Microalgae in general are known to have high levels of unsaturated fats that vary widely among species. Among the microalgae, cyanobacteria and green algae (Streptophyta and Chlorophyta) only accumulate low levels of lipids; these lipids are mostly saturated and monounsaturated lipid species. However, the Chromalveolate algae accumulate high levels of PUFAs(Watson 2003) that are comparable to H. coloradii lipid levels. H. coloradii accumulates the fatty acid EPA to as much as 25% of its total lipids. Microalgae usually have a ratio of 1:1 omega-6 fatty acids to omega-3 fatty acids (Lang et al. 2011). Normally EPA and DHA can be supplied with fish oil, but using fish oil for DHA and EPA has a number of negative consequences such as overfishing the oceans and risk of contaminants (Worm et al. 2009, Bistrian 2010, Myers and Worm 2003). Purification of fish oil adds another high-cost process to the manufacture of a DHA/EPA-supplying product. Companies V-pure (Bedford, UK) and Deva (Chelsea AL, US) have already been established to supply DHA and EPA from algae as people are demanding a commercial alternative to fish oil for supplementation with DHA and EPA. Although the human body cannot readily synthesize adequate amounts of DHA or EPA, given enough EPA the body can easily synthesize enough DHA for its needs, and vice-versa (Rhodes et al. 66 2009). Also, there is evidence for specific health benefits for taking either DHA or EPA alone (Rhodes et al. 2009). Thus, it is important to discover species of algae that accumulate EPA and DHA alone. As well as EPA, the fatty acid PA was also found to be abundant in H. coloradii (figs. 2,3, and Table 1). PA has been suggested to be a lipokine that is linked to whole body lipid metabolism (Cao et al. 2006). The authors found that PA was by far the most highly regulated fatty acid involved in adipose tissue metabolism. In adipocyte fractions of adipose tissue, PA, but not other lipids, down-regulated levels of pro-inflammatory cytokines. Thus there is evidence that PA may have therapeutic properties (Cao et al. 2006). 67 Table 1 : Summary of Medallion triglyceride analysis of H. coloradii. In summary, H. coloradii has an excellent lipid profile for potential use as a source of lipids for human nutrition. The high amounts of EPA and PA together make this organism a very attractive potential food source. Mature cells might be harvested with little to no downstream processing and be sold as a nutritional supplement for human use. 68 Chapter 4: Lipid production and cold tolerance genes in Heterococcus coloradii David R. Nelson1, Paul A. Lefebvre1 1 Department of Plant Biology, University of Minnesota, 250 Biological Sciences Building, 1440 Gortner ave. St. Paul, MN. Email: nels5133@umn.edu Phone: (651)335-0422 Fax: (612) 625-5754. A yellow-green alga, Heterococcus coloradii, was isolated from the snows of the Rocky Mountains in Colorado. H. coloradii displays unique properties such as abundant intracellular lipid accumulation and cold tolerance. Here we describe a number of genes discovered from a draft genome constructed from Illumina GAIIx short reads. H. coloradii’s genome is 170 Mb, with 29,080 genes giving hits within NCBI using 69 TeraBlastP. We show that H. coloradii has a large number of genes involved in lipid metabolism and contains the required genes for the biosynthesis of eicosapentaenoic acid, a lipid required for nutrition in humans. A number of putative cold-tolerance-related genes are present in the genome as well. INTRODUCTION A species of Heterococcus was recently discovered that was found to accumulate large intracellular stores of lipids and grow at near-freezing temperatures. This psychrophilic algae accumulates the nutritionally important fatty acids eicosapentaenoic acid (EPA) and palmitoleic acid (PA) as its predominant lipid species, up to 25% and 55%, respectively. Because of interest into this unique organism’s metabolism, a high throughput sequencing project was initiated. Genomes from red, green and brown algae (Tirichine et al. 2011) and a transcriptome from a species of yellow-green algae, Nannochloropsis (Radakovits et al. 2012), have already been sequenced. However, experimentally verified genes in organisms closely related to Heterococcus are limited. Ectocarpus siliculosis, a brown alga, is the closest related organism with a completed genome (Cock et al. 2010). 70 To learn more about the complement of genes in H. coloradii and to characterize the genome of this species we sequenced its DNA. We sought to determine the size of H. coloradii’s genome, make an estimate of the number of genes in H. coloradii, and determine possible biological processes in H. coloradii by homology of found genes with NCBI gene entries. We present here a subset of the discovered genes and how they characterize lipid production and cold tolerance in H. coloradii. MATERIALS AND METHODS H. coloradii cells were grown for 1 month on Chlamydomonas reinhardtii SG1 solid media (1.2% agar, Harris 1989) in 150X15mm agar petri dishes. Genomic DNA was harvested using the Gentra DNA extraction kit under conditions outlined by the manufacturer. More than 15 µg total DNA was extracted from H. coloradii using a Qiagen DNA extraction kit. Genomic DNA samples were quantified using a Pico Green Assay (Invitrogen). DNA samples were prepared for sequencing using the Illumina Genomic DNA Sample Preparation Guide. Library preparation: Genomic DNA was fragmented using a nebulizer and compressed nitrogen. End repair was performed and the 3’ ends were adenylated. Adapters were then ligated and the ligation products were gel 71 purified. Samples were then enriched for DNA fragments and gel purified again. The library was then validated and quantified using an Agilent High Sensitivity chip (Agilent Technologies), Pico Green Assay (Invitrogen) and KAPA qPCR (KAPA BioSystems). Cluster generation: The Illumina cBOT was used for cluster generation. Flow cell and all clustering reagents were acquired from Illumina. The DNA template was immobilized to a random oligo lawn on the surface of a flow cell then amplified, linearized, and blocked. Sequencing primers were then hybridized to the template. Clustered flow cells were then loaded onto the Illumina GAIIx. Data analysis: Read data was stored at the Minnesota Supercomputing Institute (MSI) as fastq files. Low quality scores were filtered and reads were trimmed at scores of 30 or above. Reads were assembled into contigs using the assembly program AbySS. The best assembly was chosen for analysis of gene models. Preference was given to assemblies with the highest average contig size and number. For H. coloradii, an assembly using 54-bp fragments with 5-bp overlaps was found to give the best assembly (170 Mb genome size, 20x coverage). The 170 megabase assembled genome was organized into 78293 scaffolds to be used as a working H. coloradii genome draft (Table 2). 72 The assembled genome was then used for gene prediction using the computer software Genemark (GM, Borodovsky et al. 2011). Genemark ran trained on itself, Chlamydomonas reinhardtii, Arabidopsis thaliana and Medicago truncatula. The final result was that Genemark found 37,288 predicted genes in the assembled genome of H. coloradii. GM’s prediction file (.hmm) was then used for a tera-BLASTp (http://www.ncbi.nlm.nih.gov/) in DeCypher (http://www.timelogic.com/catalog/755) to discover orthologous peptide sequences from NCBI. RESULTS 61423467 total reads were organized into 102,129 contigs and 78,293 scaffolds (Tables 1,2) using the short read assembler ABySS (Simpson et al. 2009). K-mers, n-tuples of nucleic acids used to identify regions for assembly, of 52.5 bp were chosen to give the best assembly. The minimum contig length was 51, the average contig length was 1688.7, and the max contig length was 1688.7 bp. The minimum scaffold length was 52, the average scaffold length was 2213.6, and the maximum scaffold length was 100,893 bp. The genome was found to have a GC content of 48.69% (Table 1). 73 Table 1: Comparision of GC content of H. coloradii with other species. P: Psychrophillic, M: Mesophillic, T: Thermophillic. 74 Table 2: Description of final assembly of H. coloradii genome. This assembly was used for gene predictions using Genemark. In total 37,288 genes were found. 88,079 peptides were predicted from Genemark’s model. 64,303 hits were received from a Tera-BLASTP. 29,080 unique peptides were identified in H. coloradii’s genome to be homologous with peptide sequences in NCBI with low E-values (x<10-15). We found that most hits came from Ectocarpus siliculosus, a brown macroalga (Table 3). Given that little is known about yellow-green algae, the finding of extensive similarities to brown algae may be valuable in analyzing gene function. Other organisms that shared a moderate number of proteins with H. coloradii included Phytophora infestans, or potato blight, with 1601 shared 75 proteins and Thallasiosira pseudonana, a diatom, with 864 shared proteins (Table 3). Only 298 protein hits were found comparing H. coloradii peptides to the the model alga Chlamydomonas reinhardtii, a chlorophyte.. Because the yellow-green algae are not well characterized molecularly we received few hits from this family. 76 Table 3: Comparison of organisms that received peptide matches in a BLASTP with translated nucleotide sequences from H. coloradii. 77 The overall functional gene composition of H. coloradii could be determined by using Gene Ontology (GO) terms. GO terms are descriptors used to describe functionally similar gene products (Torto-Alalibo et al. 2010, Hill et al. 2010). For example, kinases, transporters, etc. Using GO terms is a convenient way of describing biological processes in an organism (Hill et al. 2010). The goal of using GO terms was to categorize gene functions and perform broad functional annotation of H. coloradii’s modeled genes. Functional annotation incorporates experimental results from the research literature. Thus it is possible to functionally annotate predicted genes from H. coloradii using other organisms for which experimental results are available. H. coloradii’s putative proteome was used as source data for Blast2GO, a computer software program designed to annotate BLAST hits with GO terms and KEGG (Kyoto Encyclopedia of Genes and Genomes) metabolic pathway designations. Summaries of H. coloradii’s genome in GO terms can be found in Fig.1. 505 proteins were found to have lipid metabolism-related functions. 78 Figure 1: Composition of H. coloradii’s genome regarding gene functional groupings (GO terms). Proteins from H. coloradii were found to play a role in a number of different metabolic pathways as defined by KEGG. Pathways of interest include: chlorophyll metabolism, methane metabolism, fatty acid biosynthesis, and carbon fixation (Fig.2). H. coloradii appears to have extensive secondary metabolism that may be worthy of future investigation. 79 Fig. 2: KEGG pathway distribution of H. coloradii genes (www.genome.jp/kegg/) Antifreeze proteins: Since H. coloradii is native to a cold climate (0-30ºC) we hypothesized that it must have a number of antifreeze proteins (AFPs) to help it survive during the 80 winter months. A search through the draft genome revealed that there were 3 strong hits (E<-15) to AFPs in H. coloradii (figs. 3-5), one of which displayed the threonine array characteristic of certain ice-binding proteins (Davies 2002). AFPs were found from Stigmatella aurantiaca, Notothenia angustata (Maori Chief, cod icefish), which was a glycoprotein polyprotein, and from the filaria worm, Bruglia malayi. The AFP match from Bruglia malayi contained the threonine array. 81 Fig. 3: H. coloradii peptide sequence (pep48339) aligned with an antifreeze protein (ZP_01462925.1) from Stigmatella aurantiaca (49% identity). 82 Fig. 4: Alignment of H. coloradii peptide sequence (pep35355) with an antifreeze glycoprotein polyprotein (AAM61875.1) from Notothenia angustata (Maori Chief, cod icefish)n (40% identity). 83 Fig. 5: Alignment of H. coloradii peptide sequence (pep26099) with an AFP (XP_001892336.1) from the worm Bruglia malayi (65% identity). Lipid metabolism: BLASTP hits of H. coloradii peptides with proteins from other algae, fungi and some bacteria revealed more than 505 protein matches involved with lipid metabolism. BLASTP matches were found in organisms ranging from bacteria to gymnosperms. Table 4 shows a summary of BLASTP matches found from H. coloradii peptide sequences that are involved with lipid synthesis and modification. H. coloradii has 6 peptides that match isoforms of diacylglycerol O-acyltransferase (CBJ30672.1, CBJ27686.1, CBN75192.1, XP_002906877.1, CBN77027.1, ABV91586.1, DGAT, EC:2.3.1.20). DGAT catalyzes the final reaction necessary to make triacylglycerol from diacylglycerol (Fig.6). Triacylglycerol is the main form of fatty acid storage in most organisms (Karantonis et al. 2009). In addition a number of proteins were found that are involved in nearly every step of lipid 84 synthesis from glycerol to mature triglycerides (Tables 4-6). Also included are enzymes that modify lipids by glycosylation or phosphorylation. Fig. 6: Addition of third carbon chain to diacylglycerol by DGAT to form triacylglycerol. Image adopted from KEGG. NCBI accession Protein description CBN77027.1 mono-or diacylglycerol acyltransferase type 2 [Ectocarpus siliculosus] XP_642726.1 diacylglycerol kinase [Dictyostelium discoideum] CBN75121.1 1-acyl-sn-glycerol-3-phosphate acyltransferase [Ectocarpus siliculosus] CBJ28756.1 CDP-diacylglycerol--glycerol-3-phosphate 3- phosphatase [Ectocarpus siliculosus] 85 CBJ28372.1 Monogalactosyldiacylglycerol synthase, family GT28 [Ectocarpus siliculosus] CBJ27648.1 Digalactosyldiacylglycerol synthase, family GT4 [Ectocarpus siliculosus] CBJ28381.1 Monogalactosyldiacylglycerol synthase, family GT28 [Ectocarpus siliculosus] CBN74380.1 diacylglycerol kinase (Partial) [Ectocarpus siliculosus] AAG51624.1 putative phorbol ester / diacylglycerol binding protein [Arabidopsis thaliana] CBN78066.1 1-acyl-sn-glycerol-3-phosphate acyltransferase [Ectocarpus siliculosus] YP_003443466.1 1-acyl-sn-glycerol-3-phosphate acyltransferase [Allochromatium vinosum] NP_001167366.1 CDP-diacylglycerol--glycerol-3-phosphate 3- phosphatase [Salmo salar] CBJ29775.1 1-acyl-sn-glycerol-3-phosphate acyltransferase [Ectocarpus siliculosus] CBJ30672.1 sterol or diacylglycerol O-acyltransferase [Ectocarpus siliculosus] CBJ27686.1 mono-or diacylglycerol acyltransferase type 2 [Ectocarpus siliculosus] 86 CBJ27648.1 Digalactosyldiacylglycerol synthase, family GT4 [Ectocarpus siliculosus] CBJ28381.1 Monogalactosyldiacylglycerol synthase, family GT28 [Ectocarpus siliculosus] XP_002907046.1 1-acyl-sn-glycerol-3-phosphate acyltransferase [Phytophthora infestans] CBN75192.1 mono-or diacylglycerol acyltransferase type 2 [Ectocarpus siliculosus] ABV91586.1 diacylglycerol acyltransferase [Zea mays] XP_002906877.1 diacylglycerol O-acyltransferase, putative [Phytophthora infestans] CBJ25625.1 Glycerol-3-phosphate O-acyltransferase [Ectocarpus siliculosus] CBN74441.1 CDP-diacylglycerol---inositol 3- phosphatidyltransferase [Ectocarpus siliculosus] CBN77837.1 Diacylglycerol O-acyltransferase, type 1 [Ectocarpus siliculosus] CBJ32618.1 diacylglycerol kinase [Ectocarpus siliculosus] CBN74381.1 diacylglycerol kinase [Ectocarpus siliculosus] CBN75531.1 mono-or diacylglycerol acyltransferase type 2 [Ectocarpus siliculosus] CBJ27686.1 mono-or diacylglycerol acyltransferase type 2 87 [Ectocarpus siliculosus] Table 4: Enzymes involved in lipid synthesis and modification found with BLASTP from H. coloradii peptides. Enzymes found that are involved in lipolysis are shown in Table 5. Among the enzymes found, triacylglycerol lipases are notable because they are responsible for the first committed step in triacylglycerol breakdown (Fig.7). Figs. 8-10 demonstrate the similarity between a H. coloradii peptide sequences and a lipases from various other species, including a triacylglycerol lipase (ZP_05344363.3) from the ant Bryantella formatexigens. Other notable enzymes found involved in lipolysis include phospholipid lipases; phospholipid lipases remove fatty acids from the membrane into storage. NCBI accession Protein description CAL50026.1 Predicted lipase (ISS) [Ostreococcus tauri] CBJ26506.1 lipase class 3 [Ectocarpus siliculosus] CBJ26711.1 Putative lysophospholipase, monoglyceride lipase [Ectocarpus siliculosus] CAL50026.1 Predicted lipase (ISS) [Ostreococcus tauri] CBJ26506.1 lipase class 3 [Ectocarpus siliculosus] CBJ27097.1 Lipase domain protein [Ectocarpus siliculosus] CBJ29381.1 phospholipase A2, group VII [Ectocarpus siliculosus] 88 CBJ32519.1 lipase [Ectocarpus siliculosus] CBN73949.1 Putative phospholipase [Ectocarpus siliculosus] CBN75770.1 lipase, putative [Ectocarpus siliculosus] CBN76512.1 similar to phospholipase C, Δ 4 [Ectocarpus siliculosus] CBN76735.1 lipase [Ectocarpus siliculosus] CBN77084.1 similar to Calcium-independent phospholipase A2-gamma [Ectocarpus siliculosus] CBN77733.1 Lipase domain protein, partial [Ectocarpus siliculosus] CBN79491.1 carboxyl-ester lipase [Ectocarpus siliculosus] CBN79984.1 Putative phospholipase [Ectocarpus siliculosus] NP_000291.1 phospholipase A2, membrane associated precursor [Homo sapiens] NP_001135601.1 monoacylglycerol lipase ABHD12 [Xenopus (Silurana) tropicalis] NP_009623.1 Major cell wall mannoprotein with possible lipase [Saccharomyces cerivisiae] NP_631918.3 sn1-specific diacylglycerol lipase beta isoform 1 [Homo sapiens] XP_001178303.1 phospholipase [Strongylocentrotus purpuratus] XP_001377685.1 phospholipase A2 [Monodelphius domestica] XP_001831193.1 lipase [Coprinopsis cinerea okayama7#130] XP_001853511.1 hepatic triacylglycerol lipase [Culex quinquefasciatus] 89 XP_002129990.1 phospholipase C, Δ 4 [Ciona intestinalis] XP_002193959.1 patatin-like phospholipase domain containing 1 [Taeniopygia guttata] XP_002340899.1 lipase/esterase [Talaromyces stipitatus] XP_002748329.1 phospholipase D2 [Callithrix jacchus] XP_002896651.1 patatin-like phospholipase [Phytophthora infestans] XP_002999064.1 patatin-like phospholipase [Phytophthora infestans] XP_003009439.1 cytosolic phospholipase A2 zeta [Verticillium albo-atrum] XP_811442.1 lipase [Trypanosoma cruzi strain CL Brener] YP_422353.1 glyoxysomal fatty acid beta-oxidation multifunctional protein MFP-a [Magnetospirillum magneticum AMB-1] YP_001196580.1 esterase/lipase-like protein [Flavobacterium johnsoniae] YP_001302072.1 putative patatin-like phospholipase [Parabacteroides distasonis] YP_001703547.1 lipase LipH [Mycobacterium abscessus ATCC 19977] YP_002127564.1 phospholipase/Carboxylesterase [Alteromonas macleodii] YP_003249593.1 putative esterase/lipase/thioesterase [Fibrobacter succinogenes] YP_003527017.1 lipase class 3 [Nitrosococcus halophilus Nc4] YP_003713434.1 FliA regulated lipase [Xenorhabdus nematophila] YP_131916.1 putative phospholipase [Photobacterium profundum] 90 YP_159518.1 putative phospholipase [Aromatoleum aromaticum] YP_357523.2 phospholipase D protein, putative [Pelobacter carbinolicus] YP_585935.1 triacylglycerol lipase [Cupriavidus metallidurans] ZP_00683171.1 Triacylglycerol lipase [Xylella fastidiosa Ann-1] ZP_02376724.1 probable phospholipase C (plcA) [Burkholderia ubonensis] ZP_02731213.1 Esterase/lipase [Gemmata obscuriglobus UQM 2246] ZP_05344363.3 triacylglycerol lipase [Bryantella formatexigens] ZP_06488249.1 putative phospholipase accessory protein [Xanthomonas campestris] CAL50026.1 Predicted lipase (ISS) [Ostreococcus tauri] Table 5: Enzymes involved in lipolysis found with BLASTP from H. coloradii peptides. Figure 7: Triacylglycerol lipase (EC: 3.1.1.3) catalyzes the first committed step of triacylglycerol breakdown (Fig.adapted from KEGG) 91 Fig. 8: Alignment of H. coloradii peptide sequence (pep24031) with a triacylglycerol lipase (ZP_05344363.3) from the ant Bryantella formatexigens (38% identity). 92 Fig. 9: Alignment of H. coloradii peptide sequence (pep18689) with a monoacylgylcerol lipase (NP_001135601.1) from the frog Xenopus silurana (32% identity). 93 Fig. 10: Alignment of H. coloradii peptide with an esterase/lipase (EFY91335) from Metarhizium acridum (31% identity). 94 Several enzymes were found that are involved in the initial steps of lipid synthesis. Several fatty acid desaturases and elongases were also found. These are shown listed in Table 6. In addition, several long-chain fatty acyl-CoA synthetases were found in the genome of H. coloradii. NCBI accession Protein description CBN79823.1 Acetyl-coenzyme A synthetase [Ectocarpus siliculosus] YP_628900.1 fatty acid desaturase family protein [Myxococcus xanthus] CBJ29129.1 fatty acid desaturase [Ectocarpus siliculosus] CBJ26568.1 Fatty acid elongase [Ectocarpus siliculosus] YP_001831427.1 fatty acid desaturase [Beijerinckia indica] YP_002361678.1 fatty acid desaturase [Methylocella silvestris] CBJ49159.1 Fatty acid desaturase [Ectocarpus siliculosus] CBN78890.1 Fatty acid elongase [Ectocarpus siliculosus] XP_002508575.1 fatty acid desaturase [Micromonas sp. RCC299] YP_003277402.1 long-chain-fatty-acid--CoA ligase [Comamonas testosteroni CNB-2] P12276.5 Fatty acid synthase [Gallus gallus] ADG36330.1 Δ-4 fatty acid desaturase [Pavlova viridis] CBJ28321.1 cyclopropane-fatty-acyl-phospholipid synthase/ oxidoreductase 95 [Ectocarpus siliculosus] YP_143714.1 long-chain-fatty-acid--CoA ligase [Thermus thermophillus HB8] CBJ26568.1 Fatty acid elongase [Ectocarpus siliculosus] CBN76062.1 fatty acid-ACP thioesterase [Ectocarpus siliculosus] CBJ26764.1 polyunsaturated fatty acids Δ-6-elongase [Ectocarpus siliculosus] YP_001846474.1 long-chain fatty acid ABC transporter [Acinetobacter baumannii] CBN74970.1 Δ-6 fatty acid desaturase [Ectocarpus siliculosus] XP_001260605.1 bifunctional fatty acid transporter/acyl-CoA synthetase [Neosartorya fischeri] ZP_05095815.1 Cyclopropane-fatty-acyl-phospholipid synthase [marine gamma proteobacterium] CAD53323.1 Δ 5 fatty acid desaturase [Phytophthora megasporidium] YP_001519835.1 long-chain-fatty-acid-CoA ligase, putative [Acaryochloris marina] ZP_01221855.1 omega-3 polyunsaturated fatty acid synthase PfaA [Photobacterium profundum] YP_002361678.1 fatty acid desaturase [Methylocella silvestris] 96 ZP_05000329.1 fatty acid desaturase [Streptomyces sp. Mg1] CBJ31067.1 Fatty acid desaturase [Ectocarpus siliculosus] ADG36330.1 Δ-4 fatty acid desaturase [Pavlova viridis] YP_001474935.1 omega-3 polyunsaturated fatty acid synthase PfaB [Shewanella sediminis] XP_002774616.1 Long-chain-fatty-acid--CoA ligase, putative [Perkinsus marinus] XP_002508575.1 fatty acid desaturase [Micromonas sp. RCC299] AAF70457.1 Δ-5 fatty acid desaturase [Homo sapiens] Table 6: Enzymes involved in desaturation/elongation found with BLASTP from H. coloradii peptides. EPA synthesis: Every fatty acid synthase needed to make EPA (Fig.11) was found in H. coloradii. Matches of H. coloradii peptides with enzymes in this pathway include a Δ 9 desaturase: (pep34467/ZP_06711707.1) from Streptomyces sp. e14, a Δ 12 desaturase : (pep9713/AAR20443.1) from Saprolegnia diclina, a Δ 6 desaturase: (pep44208/CBN74970.1), from Ectocarpus siliculosus, a PUFA Δ-6- elongase: (pep84363/CBJ26764.1) from Ectocarpus siliculosus, a Δ 5 desaturase : (pep49169/CAD53323.1), from Phytophthora megasperma, and an 97 Omega-3 fatty acid desaturase: (pep71855/AFJ69342.1)from Nannochloropsis gaditana. Alignments with close matches are shown in figures 12-15. Fig. 11: Biosynthetic pathways of erucic acid, arachidonic acid (AA) and eicosapentaenoic acid (EPA). LA linoleic acid, ALA alpha-linoleic acid, GLA gamma-linolenic acid, SDA stearidonic acid, DGLA dihomo- gamma- linolenic acid, ETA eicosatetraenoic acid, AA arachidonic acid, EPA eicosapentaenoic acid. 98 Fig. 12: Alignment of an H. coloradii peptide (pep84363) with a Δ-6- elongase from Ectocarpus siliculosus (CBJ26764.1) (58% identity). 99 Fig. 13: Alignment of H. coloradii peptide (pep49169) with a Δ 5 fatty acid desaturase (Phytophthora, CAD53323.1) (51% identity). 100 Fig. 14: Alignment of H. coloradii peptide (pep71855) with a Δ-12 desaturase (CBJ31067.1) from E. siliculosus (60% identity). 101 Fig. 15: Alignment of H. coloradii peptide (pep71855) with an omega-3 desaturase from Nannochloropsis gaditana (AFJ69342.1) (44% identity). 102 Virus Proteins H. coloradii had 75/240 unique peptide-coding regions found with homology to EsV-1 proteins. EsV-1 is a lysogenic dsDNA virus E. siliculosis belonging to the super family of nucleocytoplasmic large DNA viruses (NCLDV) (Cock et al. 2010). Also found were 19 unique peptides from the Feldmannia irregularis virus and: 4 unique peptides from the Marseillevirus.df Beta-Tubulin H. coloradii was found to have a phenylalanine to tyrosine polymorphism in two copies of beta tubulin that differ from E. siliculosus (Fig. 16). This polymorphism is the only nucleotide change between the H. coloradii and E.siliculosus beta-tubulin genes. 103 Figure 16: Alignment of beta-tubulin from H. coloradii and E. siliculosus. DISCUSSION Cold tolerance: Many cold tolerant organisms have lower GC content to help them replicate and utilize DNA sequences (Casana and Gullati 2011). H. coloradii had a GC content of 48.69%, which is much higher than its close Mesophillic relative E. siliculosis with its GC content of 30.7% (Table 1). Therefore, H. 104 coloradii must have unique cellular mechanisms that help it adapt to cold climates, since it grows well at 4ºC. H. coloradii was found to have 3 AFPs, which are essential for the survival of many organisms in cold climates. AFPs reduce the harm caused by the crystallization of water to cells (Carvajal-Rondanelli et al. 2010). Normally when ice expands it pushes proteins out of its way and away from its expanding front. AFPs are different in that they adsorb to the surface of the expanding ice and bind it (Carvajal-Rondanelli et al. 2010). An array of threonine residues within certain AFPs are known to aid proper binding to ice (Davies 2002), for example. In this case water molecules that are to become ice must bind in-between amino acids. The expansion of ice through narrow channels rather than as a uniform front is thermodynamically unfavorable. Thus the expansion of the intracellular ice is retarded or completely stopped by the AFPs (Wilson et al. 1993). Algae AFPs have been implicated in changing the structure of sea ice on a macroscopic level (Raymond 2011). It is unknown why the AFPs from H. coloradii would match AFP from such a divergent group of organisms (figs. 3-5). While relatives of H. coloradii such as E. siliculosus may have lost such AFP, a common ancestor may have passed certain AFPs down to H. coloradii and organisms like the cold-tolerant frogs and worms represented in figs. 3-5. Evolutionary convergence may also be cited as the cause of H. coloradii’s putative AFPs and the matches found in NCBI, however this is doubtful because of the similarity of the peptide chains. 105 However, since in AFPs the peptide chain composition is tied tightly to its function, as in the threonine residue arrays, evolutionary convergence may well explain the homology between H. coloradii’s putative AFP sequences and those found in the NCBI nucleotide database. The single nucleotide polymorphism between H. coloradii and E. siliculosus is interesting because it is the only difference in a structural gene between a mesophilic and psychrophilic organism. Phenylalanine to tyrosine mutations in beta-tubulin have been shown to be involved in antibiotic resistance in helminthes (Niciura et al. 2012). Other polymorphisms in beta-tubulin confer cold resistance because they promote elasticity of the peptide chain (Chiappori et al. 2012) Lipid metabolism: With 505 genes found to potentially be involved in lipid metabolism in H. coloradii, this sequencing effort represents a major step forward in describing lipid metabolism in H. coloradii. All genes involved in the EPA synthesis pathway have BLAST hits in the H. coloradii genome, although these newly discovered genes remain to be experimentally verified. Studying the enzymes involved in EPA synthesis may provide information useful for heterologous expression of these enzymes for EPA production in other 106 organisms, or even to increase the amount of EPA increased in H. coloradii. For example, genes encoding a Δ6 desaturase, Δ6 fatty acid elongase, and Δ5 desaturase from the alga, Phaeodactylum tricornutum, were co-expressed in Pichia pastoris to produce arachidonic acid (ARA; 20:4 Δ(5, 8, 11, 14)) and eicosapentaenoic acid (EPA; 20:5 Δ(5, 8, 11, 14, 17)). In these studies each of these genes was introduced into P. tricornutum by transformation. The resulting transgenic P. tricornutum could produce up to 4x more EPA than the wild type P. tricornutum. End EPA levels were up to 0.1% EPA in transgenic P. tricornutum containing double copies of the Δ6 desaturase, Δ6 fatty acid elongase, and Δ5 desaturases while wild type P. tricornutum could produce up to 0.05% EPA of total fatty acids (Li et al. 2009). While P. tricornutum is not widely used in the biotechnology industry, Saccharomyces cerevisiae is frequently used for production of a number of valuable biocompounds in addition to its wide use in food production (Keasling 2010, Yu et al. 2011). Thus, S. cerevisiae is an attractive target for trans genes leading to the production of EPA. Transformation of S. cerevisiae with heterologous Δ5 desaturases from the ciliate protozoan Paramecium tetraurelia and from the microalgae Ostreococcus tauri and Ostreococcus lucimarinus allowed S. cerevisiae to produce EPA using glucose as the sole carbon source (Tavares et al. 2011). An experiment could be conceived wherein S. cerevisiae is transformed with an H. coloradii Δ5 desaturase. 107 The breakdown of tryglycerides, including EPA, is catalyzed by lipases. Lipases act on carboxylic esters on glycerol-esterified fatty acids to free the fatty acids from the glycerol backbone (EC.3.1.1.-). The catalytic center contains three residues acting as acatalytic triad: a serine, a glutamate or aspartate and a histidine. These catalytic residues are responsible for the nucleophilic attack on the carbonyl carbon atom of the ester bond. Lipase reactions are critical for tryglyceride breakdown and are possibly responsible for the drop in certain fatty acids seen in H. coloradii upon change of environmental conditions. In addition to lipase reactions, various transferase, desaturase, and elongase reactions are responsible for changing one fatty acid into another as the organism requires. An acetyl-CoA synthetase was found that provides an initial precursor, acetyl-CoA, for fatty acid production. Acetyl-coA synthetase catalyzes the reaction between ATP and acetate to form acetyl-CoA (KEGG). Acetyl-CoA is used to initiate and elongate fatty acid chains as well as provide precursors to a number of other energy-producing molecules (Starai et al. 2004). Several long-chain fatty acyl-CoA synthetases were found in the genome of H. coloradii (Table 6). These enzymes perform important regulatory functions in cellular homeostasis, particularly in lipid metabolism (Kuar et al. 2011). A fatty-acyl CoA synthetase (EC 6.2.1.x) allows an otherwise non-reactive fatty acid to participate in a myriad of metabolic pathways. Fatty-acyl CoA synthetases in their differenct forms can catalyze reactions with short (2-3 carbons), medium (4-12 carbons), or long (>12 carbons) –chain fatty acid 108 substrates (Watkins 2007). Long-chain fatty acyl-CoA synthetases have been shown to be bi-functional enzymes involved in fatty acid transformation as well as transport, and they have several roles in the maintanaince of human health (Watkins 2007). Virus Proteins An adequate number of EsV-1 proteins were found in the H. coloradii genome to conclude that either 1) EsV-1 infected a common ancestor of yellow- green and brown algae or that 2) EsV-1 has broader host specificity that previously thought and can also infect yellow-green algae. Since our study only analyzed translated peptide fragments, more sequencing will need to be done to address these possibilities. Conclusion We present here a draft version of H. coloradii’s putative proteome, analyze its genes’ functions and compare the organism to its closest relatives. The discovered genes offer doorways to several research venues to more specifically study H. coloradii. Overall, H. coloradii appears to be an interesting organism with a diverse collection of genes contributing to its unique lipid accumulation and cold-tolerance phenotypes. 109 References 1. Adl SM, Simpson AG, Farmer MA, Andersen RA, Anderson OR, Barta JR, et al. The new higher level classification of eukaryotes with emphasis on the taxonomy of protists J Eukaryot Microbiol. 2005 Sep-Oct;52(5):399-451. 2. Amato P, Parazols M, Sancelme M, Laj P, Mailhot G, Delort AM. Microorganisms isolated from the water phase of tropospheric clouds at the puy de dome: Major groups and growth abilities at low temperatures FEMS Microbiol Ecol. 2007 Feb;59(2):242-54. 3. Araujo GS, Matos LJ, Goncalves LR, Fernandes FA, Farias WR. Bioprospecting for oil producing microalgal strains: Evaluation of oil and biomass production for ten microalgal strains Bioresour Technol. 2011 Apr;102(8):5248-50. 4. Arrigo KR, Worthen DL, Lizotte MP, Dixon P, Dieckmann G. Primary production in antarctic sea ice Science. 1997 Apr 18;276(5311):394-7. 5. Ashokkumar V, Rengasamy R. Mass culture of botryococcus braunii kutz. under open raceway pond for biofuel production Bioresour Technol. 2011 Nov 6. 6. Becker EW. Micro-algae as a source of protein Biotechnol Adv. 2007 Mar- Apr;25(2):207-10. 7. BELCHER JH, MILLER JD. Studies on the growth of xanthophyceae in pure culture. IV. nutritional types amongst the xanthophyceae Arch Mikrobiol. 1960;36:219-28. 8. Bistrian B. By the way, doctor. omega-3 fats may be good for you, but I worry about overfishing. is there a way of getting omega-3s without contributing to this problem? Harv Health Lett. 2010 Nov;36(1):8. 9. Bjerke JW. Ice encapsulation protects rather than disturbs the freezing lichen Plant Biol (Stuttg). 2009 Mar;11(2):227-35. 10. Bluhm BA, Gradinger R. Regional variability in food availability for arctic marine mammals Ecol Appl. 2008 Mar;18(2 Suppl):S77-96. 11. Blumenthal S, Morgan-Boyd R, Nelson R, Garshelis DL, Turyk ME, Unterman T. Seasonal regulation of the growth hormone-insulin-like growth factor-I axis in the american black bear (ursus americanus) Am J Physiol Endocrinol Metab. 2011 Oct;301(4):E628-36. 110 12. Bonente G, Formighieri C, Mantelli M, Catalanotti C, Giuliano G, Morosinotto T, et al. Mutagenesis and phenotypic selection as a strategy toward domestication of chlamydomonas reinhardtii strains for improved performance in photobioreactors Photosynth Res. 2011 Sep;108(2-3):107-20. 13. Borodovsky M, Lomsadze A. Eukaryotic gene prediction using GeneMark.hmm-E and GeneMark-ES Curr Protoc Bioinformatics. 2011 Sep;Chapter 4:Unit 4.6.1-10. 14. Brierley AS, Thomas DN. Ecology of southern ocean pack ice Adv Mar Biol. 2002;43:171-276. 15. Brooks KK, Liang B, Watts JL. The influence of bacterial diet on fat storage in C. elegans PLoS One. 2009 Oct 21;4(10):e7545. 16. Cakmak T, Angun P, Ozkan AD, Cakmak Z, Olmez TT, Tekinay T. Nitrogen and sulfur deprivation differentiate lipid accumulation targets of chlamydomonas reinhardtii Bioengineered. 2012 Nov 1;3(6). 17. Calder PC. Fatty acids and inflammation: The cutting edge between food and pharma Eur J Pharmacol. 2011 Jul 28. 18. Callaghan TV, Bjorn LO, Chernov Y, Chapin T, Christensen TR, Huntley B, et al. Responses to projected changes in climate and UV-B at the species level Ambio. 2004 Nov;33(7):418-35. 19. Canizares-Villanueva RO, Rios-Leal E, Olvera Ramirez R, Ponce Noyola T, Marquez Rocha F. Microbial sources of pigments Rev Latinoam Microbiol. 1998 Jan-Jun;40(1- 2):87-107. 20. Cao H, Gerhold K, Mayers JR, Wiest MM, Watkins SM, Hotamisligil GS. Identification of a lipokine, a lipid hormone linking adipose tissue to systemic metabolism Cell. 2008 Sep 19;134(6):933-44. 21. Cardinale BJ, Matulich KL, Hooper DU, Byrnes JE, Duffy E, Gamfeldt L, et al. The functional role of producer diversity in ecosystems Am J Bot. 2011 Mar;98(3):572-92. 22. Cardozo KH, Guaratini T, Barros MP, Falcao VR, Tonon AP, Lopes NP, et al. Metabolites from algae with economical impact Comp Biochem Physiol C Toxicol Pharmacol. 2007 Jul-Aug;146(1-2):60-78. 23. Carvajal-Rondanelli PA, Marshall SH, Guzman F. Antifreeze glycoprotein agents: Structural requirements for activity J Sci Food Agric. 2011 Nov;91(14):2507-10. 111 24. Casselton P. Chemo-organotrophic growth of xanthophycean algae. Dept Botany, Birkbeck College, University of London PhD dissertation. 1965. 25. Cavicchioli R, Charlton T, Ertan H, Omar SM, Siddiqui KS, Williams TJ. Biotechnological uses of enzymes from psychrophiles Microb Biotechnol. 2011 Jul;4(4):449-60. 26. Chakraborty C, Hsu CH, Wen ZH, Lin CS. Anticancer drugs discovery and development from marine organism Curr Top Med Chem. 2009;9(16):1536-45. 27. Chen H, Jiang JG. Osmotic responses of dunaliella to the changes of salinity J Cell Physiol. 2009 May;219(2):251-8. 28. Chen M, Tang H, Ma H, Holland TC, Ng KY, Salley SO. Effect of nutrients on growth and lipid accumulation in the green algae dunaliella tertiolecta Bioresour Technol. 2011 Jan;102(2):1649-55. 29. Chiappori F, Pucciarelli S, Merelli I, Ballarini P, Miceli C, Milanesi L. Structural thermal adaptation of ß-tubulins from th... [proteins. 2012] - PubMed - NCBI Proteins. 2012(9/18/2012). 30. Christaki E, Florou-Paneri P, Bonos E. Microalgae: A novel ingredient in nutrition Int J Food Sci Nutr. 2011 May 16. 31. Christenson L, Sims R. Production and harvesting of microalgae for wastewater treatment, biofuels, and bioproducts Biotechnol Adv. 2011 Jun 2. 32. Chu SP. The influence of the mineral composition of the medium on the growth of planktonic algae. part I. methods and culture media. J Ecol ,(30):284-325. 33. Cock JM, Coelho SM, Brownlee C, Taylor AR. The ectocarpus genome sequence: Insights into brown algal biology and the evolutionary diversity of the eukaryotes New Phytol. 2010 Oct;188(1):1-4. 34. Cockbain AJ, Toogood GJ, Hull MA. Omega-3 polyunsaturated fatty acids for the treatment and prevention of colorectal cancer Gut. 2011 Apr 13. 35. Cornelissen JH, Lang SI, Soudzilovskaia NA, During HJ. Comparative cryptogam ecology: A review of bryophyte and lichen traits that drive biogeochemistry Ann Bot. 2007 May;99(5):987-1001. 36. Craggs RJ, Heubeck S, Lundquist TJ, Benemann JR. Algal biofuels from wastewater treatment high rate algal ponds Water Sci Technol. 2011;63(4):660-5. 112 37. Danielewicz MA, Anderson LA, Franz AK. Triacylglycerol profiling of marine microalgae by mass spectrometry J Lipid Res. 2011 Nov;52(11):2101-8. 38. Darling RB, Friedmann EI, Broady PA. Heterococcus endolithicus sp. nov. (xanthophyceae) and other terrestrial heterococcus species from antarctica: Morphological changes during life history and response to temperature J Phycol. 1987;23:598-607. 39. Davies PL, Baardsnes J, Kuiper MJ, Walker VK. Structure and function of antifreeze proteins Philos Trans R Soc Lond B Biol Sci. 2002 Jul 29;357(1423):927-35. 40. De la Hoz Siegler H, Ben-Zvi A, Burrell RE, McCaffrey WC. The dynamics of heterotrophic algal cultures Bioresour Technol. 2011 May;102(10):5764-74. 41. Delaroque N, Muller DG, Bothe G, Pohl T, Knippers R, Boland W. The complete DNA sequence of the ectocarpus siliculosus virus EsV-1 genome Virology. 2001 Aug 15;287(1):112-32. 42. Doughman SD, Krupanidhi S, Sanjeevi CB. Omega-3 fatty acids for nutrition and medicine: Considering microalgae oil as a vegetarian source of EPA and DHA Curr Diabetes Rev. 2007 Aug;3(3):198-203. 43. Ehara M, Hayashi-Ishimaru Y, Inagaki Y, Ohama T. Use of a deviant mitochondrial genetic code in yellow-green algae as a landmark for segregating members within the phylum J Mol Evol. 1997 Aug;45(2):119-24. 44. Eiken H, Lange MA. Image analysis of sea ice thin sections: A step towards automated texture classification. . . 1991(15):204-9. 45. Elster Chapter 17 J. Geoecology of antarctic ice-free coastal landscapes; ecological classification of terrestrial algal communities in polar environments . 2002;154:303 326. 46. Epstein PR. Is global warming harmful to health? Sci Am. 2000 Aug;283(2):50-7. 47. Eroglu E, Okada S, Melis A. Hydrocarbon productivities in different botryococcus strains: Comparative methods in product quantification J Appl Phycol. 2011 Aug;23(4):763-75. 48. Ettl H. Xanthophyceae.in EttlH, GerloffJ, heynig H. susswasserflora von mitteleuropa bd.3.,1.teil. Gustav Fischer Verlag,Stuttgart. 1978. 113 49. Ewart KV, Lin Q, Hew CL. Structure, function and evolution of antifreeze proteins Cell Mol Life Sci. 1999 Feb;55(2):271-83. 50. Fan J, Andre C, Xu C. A chloroplast pathway for the de novo biosynthesis of triacylglycerol in chlamydomonas reinhardtii FEBS Lett. 2011 Jun 23;585(12):1985-91. 51. Feller G. Life at low temperatures: Is disorder the driving force? Extremophiles. 2007 Mar;11(2):211-6. 52. Fiore AM, Naik V, Spracklen DV, Steiner A, Unger N, Prather M, et al. Global air quality and climate Chem Soc Rev. 2012 Aug 6. 53. Folch J, Lees M, Sloane SG. A simple method for the isolation and purification of total lipids from animal tissues. J Biol Chem. 1957;226(1):497-509. 54. Fritsen CH, Lytle VI, Ackley SF, Sullivan CW. Autumn bloom of antarctic pack-ice algae Science. 1994 Nov 4;266(5186):782-4. 55. Gachon CM, Sime-Ngando T, Strittmatter M, Chambouvet A, Kim GH. Algal diseases: Spotlight on a black box Trends Plant Sci. 2010 Nov;15(11):633-40. 56. Geesey GG, Morita RY. Some physiological effects of near-maximum growth temperatures on an obligately psychrophilic marine bacterium Can J Microbiol. 1975 Jun;21(6):811-8. 57. Gorton HL, Williams WE, Vogelmann TC. The light environment and cellular optics of the snow alga chlamydomonas nivalis (bauer) wille Photochem Photobiol. 2001 Jun;73(6):611-20. 58. Gray DW, Lewis LA, Cardon ZG. Photosynthetic recovery following desiccation of desert green algae (chlorophyta) and their aquatic relatives Plant Cell Environ. 2007 Oct;30(10):1240-55. 59. Grueter CC, Li D, Ren B, Wei F, Xiang Z, van Schaik CP. Fallback foods of temperate-living primates: A case study on snub-nosed monkeys Am J Phys Anthropol. 2009 Dec;140(4):700-15. 60. Guinotte JM, Fabry VJ. Ocean acidification and its potential effects on marine ecosystems Ann N Y Acad Sci. 2008;1134:320-42. 61. Hader DP. Effects of solar UV-B radiation on aquatic ecosystems Adv Space Res. 2000;26(12):2029-40. 114 62. Hammond EM, Kaufmann MR, Giaccia AJ. Oxygen sensing and the DNA-damage response. Curr Opin Cell Biol. 2007 Dec;19(6):680-4. 63. Hannon M, Gimpel J, Tran M, Rasala B, Mayfield S. Biofuels from algae: Challenges and potential Biofuels. 2010 Sep;1(5):763-84. 64. Harris E. The chlamydomonas sourcebook. A comprehensive guide to biology and laboratory use. Academic Press, San Diego, CA,. 1989;xiv:780. 65. Hegarty BD, Parker GB. Marine omega-3 fatty acids and mood disorders--linking the sea and the soul. 'food for thought' I Acta Psychiatr Scand. 2011 Jul;124(1):42-51. 66. Heinze VM, Actis AB. Dietary conjugated linoleic acid and long-chain n-3 fatty acids in mammary and prostate cancer protection: A review Int J Food Sci Nutr. 2011 Jul 15. 67. Hemschemeier A, Fouchard S, Cournac L, Peltier G, Happe T. Hydrogen production by chlamydomonas reinhardtii: An elaborate interplay of electron sources and sinks. Planta. 2008 Jan;227(2):397-407. 68. Heredia-Arroyo T, Wei W, Hu B. Oil accumulation via heterotrophic/mixotrophic chlorella protothecoides Appl Biochem Biotechnol. 2010 Nov;162(7):1978-95. 69. Hewitson KS, McNeill LA, Elkins JM, Schofield CJ. The role of iron and 2- oxoglutarate oxygenases in signalling. Biochem Soc Trans. 2003 Jun;31(Pt 3):510-5. 70. Hewitson KS, McNeill LA, Riordan MV, Tian YM, Bullock AN, Welford RW, et al. Hypoxia-inducible factor (HIF) asparagine hydroxylase is identical to factor inhibiting HIF (FIH) and is related to the cupin structural family. J Biol Chem. 2002 Jul 19;277(29):26351-5. 71. Hill DP, Berardini TZ, Howe DG, Van Auken KM. Representing ontogeny through ontology: A developmental biologist's guide to the gene ontology Mol Reprod Dev. 2010 Apr;77(4):314-29. 72. Hoffmann M, Marxen K, Schulz R, Vanselow KH. TFA and EPA productivities of nannochloropsis salina influenced by temperature and nitrate stimuli in turbidostatic controlled experiments Mar Drugs. 2010 Sep 27;8(9):2526-45. 73. Holzinger A, Roleda MY, Lutz C. The vegetative arctic freshwater green alga zygnema is insensitive to experimental UV exposure Micron. 2009 Dec;40(8):831-8. 74. Hon WC, Wilson MI, Harlos K, Claridge TD, Schofield CJ, Pugh CW, et al. Structural basis for the recognition of hydroxyproline in HIF-1 alpha by pVHL. Nature. 2002 Jun 27;417(6892):975-8. 115 75. Hoshino Y, Shioji K, Nakamura H, Masutani H, Yodoi J. From oxygen sensing to heart failure: Role of thioredoxin. Antioxid Redox Signal. 2007 Jun;9(6):689-99. 76. Hosseini Tafreshi A, Shariati M. Dunaliella biotechnology: Methods and applications J Appl Microbiol. 2009 Jul;107(1):14-35. 77. Huang AH. Oleosins and oil bodies in seeds and other organs Plant Physiol. 1996 Apr;110(4):1055-61. 78. Huang LE, Arany Z, Livingston DM, Bunn HF. Activation of hypoxia-inducible transcription factor depends primarily upon redox-sensitive stabilization of its alpha subunit. J Biol Chem. 1996 Dec 13;271(50):32253-9. 79. Hughes BT, Espenshade PJ. Oxygen-regulated degradation of fission yeast SREBP by Ofd1, a prolyl hydroxylase family member. EMBO J. 2008 May 21;27(10):1491-501. 80. Imhoff JF, Labes A, Wiese J. Bio-mining the microbial treasures of the ocean: New natural products Biotechnol Adv. 2011 Sep-Oct;29(5):468-82. 81. Ioki M, Baba M, Nakajima N, Shiraiwa Y, Watanabe MM. Transcriptome analysis of an oil-rich race B strain of botryococcus braunii (BOT-22) by de novo assembly of pyrosequencing cDNA reads Bioresour Technol. 2011 Aug 27. 82. Jaakkola P, Mole DR, Tian YM, Wilson MI, Gielbert J, Gaskell SJ, et al. Targeting of HIF-alpha to the von hippel-lindau ubiquitylation complex by O2-regulated prolyl hydroxylation. Science. 2001 Apr 20;292(5516):468-72. 83. Jeong BE, Ko EJ, Joo HG. Cytoprotective effects of fucoidan, an algae-derived polysaccharide on 5-fluorouracil-treated dendritic cells Food Chem Toxicol. 2012 Feb 1;50(5):1480-4. 84. Jervis AJ, Green J. In vivo demonstration of FNR dimers in response to lower O(2) availability. J Bacteriol. 2007 Apr;189(7):2930-2. 85. Jiang Y, Yoshida T, Quigg A. Photosynthetic performance, lipid production and biomass composition in response to nitrogen limitation in marine microalgae Plant Physiol Biochem. 2012 May;54:70-7. 86. John RP, Anisha GS, Nampoothiri KM, Pandey A. Micro and macroalgal biomass: A renewable source for bioethanol Bioresour Technol. 2011 Jan;102(1):186-93. 87. Jolivet P, Roux E, D'Andrea S, Davanture M, Negroni L, Zivy M, et al. Protein composition of oil bodies in arabidopsis thaliana ecotype WS Plant Physiol Biochem. 2004 Jun;42(6):501-9. 116 88. Kasana RC, Gulati A. Cellulases from psychrophilic microorganisms: A review J Basic Microbiol. 2011 Mar 24. 89. Kassis NM, Gigliotti JC, Beamer SK, Tou JC, Jaczynski J. Characterization of lipids and antioxidant capacity of novel nutraceutical egg products developed with omega-3- rich oils J Sci Food Agric. 2011 Jul 18. 90. Kaur J, Tiwari R, Kumar A, Singh N. Bioinformatic analysis of leishmania donovani long-chain fatty acid-CoA ligase as a novel drug target Mol Biol Int. 2011;2011:278051. 91. Kawamoto J, Kurihara T, Yamamoto K, Nagayasu M, Tani Y, Mihara H, et al. Eicosapentaenoic acid plays a beneficial role in membrane organization and cell division of a cold-adapted bacterium, shewanella livingstonensis Ac10 J Bacteriol. 2009 Jan;191(2):632-40. 92. Keasling JD. Manufacturing molecules through metabolic engineering Science. 2010 Dec 3;330(6009):1355-8. 93. Khan S, Kong C, Kim J, and Kim S. Protective effect of amphiroa dilatata on ROS induced oxidative damage and MMP expressions in HT1080 cells. Biotechnol Bioprocess Eng. 2010;15:191-8. 94. Kidd PM. Omega-3 DHA and EPA for cognition, behavior, and mood: Clinical findings and structural-functional synergies with cell membrane phospholipids Altern Med Rev. 2007 Sep;12(3):207-27. 95. Kilian O, Benemann CS, Niyogi KK, Vick B. High-efficiency homologous recombination in the oil-producing alga nannochloropsis sp Proc Natl Acad Sci U S A. 2011 Nov 28. 96. Kim S, Pangestuti R. Potential role of maine algae on female health, beauty and longevity. Adv Food Nutr Res;64:41-55. 97. Kim GH, Klochkova TA, Kang SH. Notes on freshwater and terrestrial algae from ny-alesund, svalbard (high arctic sea area) J Environ Biol. 2008 Jul;29(4):485-91. 98. Kim NJ, Li H, Jung K, Chang HN, Lee PC. Ethanol production from marine algal hydrolysates using escherichia coli KO11 Bioresour Technol. 2011 Aug;102(16):7466-9. 99. Kim SK, Pangestuti R. Potential role of marine algae on female health, beauty, and longevity Adv Food Nutr Res. 2011;64:41-55. 100. Knud-Hansen CF, McElwee K, Baker J, Clair D. Pond fertilization: Ecological approach 117 and practical application. Pond Dynamics/Aquaculture Collaborative Research Support Program, Oregon State University. 1998. 101. Kropat J, Hong-Hermesdorf A, Casero D, Ent P, Castruita M, Pellegrini M, et al. A revised mineral nutrient supplement increases biomass and growth rate in chlamydomonas reinhardtii Plant J. 2011 Jun;66(5):770-80. 102. Kumagai H, Hakoyama T, Umehara Y, Sato S, Kaneko T, Tabata S, et al. A novel ankyrin-repeat membrane protein, IGN1, is required for persistence of nitrogen-fixing symbiosis in root nodules of lotus japonicus Plant Physiol. 2007 Mar;143(3):1293-305. 103. Kumar K, Dasgupta CN, Nayak B, Lindblad P, Das D. Development of suitable photobioreactors for CO2 sequestration addressing global warming using green algae and cyanobacteria Bioresour Technol. 2011 Apr;102(8):4945-53. 104. Lake SL, Matthews JB, Kaplan RM, Hodgkinson JE. Determination of genomic DNA sequences for beta-tubulin isotype 1 from multiple species of cyathostomin and detection of resistance alleles in third-stage larvae from horses with naturally acquired infections Parasit Vectors. 2009 Sep 25;2 Suppl 2:S6. 105. Lang I, Hodac L, Friedl T, Feussner I. Fatty acid profiles and their distribution patterns in microalgae: A comprehensive analysis of more than 2000 strains from the SAG culture collection BMC Plant Biol. 2011 Sep 6;11:124. 106. Lebar MD, Heimbegner JL, Baker BJ. Cold-water marine natural products Nat Prod Rep. 2007 Aug;24(4):774-97. 107. Lee SH, Stephens JL, Paul KS, Englund PT. Fatty acid synthesis by elongases in trypanosomes Cell. 2006 Aug 25;126(4):691-9. 108. Li Y, Chen YF, Chen P, Min M, Zhou W, Martinez B, et al. Characterization of a microalga chlorella sp. well adapted to highly concentrated municipal wastewater for nutrient removal and biodiesel production Bioresour Technol. 2011 Apr;102(8):5138-44. 109. Li Y, Han D, Hu G, Dauvillee D, Sommerfeld M, Ball S, et al. Chlamydomonas starchless mutant defective in ADP-glucose pyrophosphorylase hyper-accumulates triacylglycerol Metab Eng. 2010 Jul;12(4):387-91. 110. Li YT, Li MT, Fu CH, Zhou PP, Liu JM, Yu LJ. Improvement of arachidonic acid and eicosapentaenoic acid production by increasing the copy number of the genes encoding fatty acid desaturase and elongase into pichia pastoris Biotechnol Lett. 2009 Jul;31(7):1011-7. 118 111. Liu J, Yuan C, Hu G, Li F. Effects of light intensity on the growth and lipid accumulation of microalga scenedesmus sp. 11-1 under nitrogen limitation Appl Biochem Biotechnol. 2012 Apr;166(8):2127-37. 112. Liu M, Hansen PE, Lin X. Bromophenols in marine algae and their bioactivities Mar Drugs. 2011;9(7):1273-92. 113. Logue JA, Howell BR, Bell JG, Cossins AR. Dietary n-3 long-chain polyunsaturated fatty acid deprivation, tissue lipid composition, ex vivo prostaglandin production, and stress tolerance in juvenile dover sole (solea solea L.) Lipids. 2000 Jul;35(7):745-55. 114. Lohrmann NL, Logan BA, Johnson AS. Seasonal acclimatization of antioxidants and photosynthesis in chondrus crispus and mastocarpus stellatus, two co-occurring red algae with differing stress tolerances Biol Bull. 2004 Dec;207(3):225-32. 115. Lorincz Z, Preininger E, Kosa A, Ponyi T, Nyitrai P, Sarkadi L, et al. Artificial tripartite symbiosis involving a green alga (chlamydomonas), a bacterium (azotobacter) and a fungus (alternaria): Morphological and physiological characterization Folia Microbiol (Praha). 2010 Jul;55(4):393-400. 116. MacDougall KM, McNichol J, McGinn PJ, O'Leary SJ, Melanson JE. Triacylglycerol profiling of microalgae strains for biofuel feedstock by liquid chromatography-high-resolution mass spectrometry Anal Bioanal Chem. 2011 Nov;401(8):2609-16. 117. Madrigal A. How algal biofuels lost a decade in the race to replace oil. Wired. 2009. 118. Maistro S, Broady PA, Andreoli C, Negrisolo E. Molecular phylogeny and evolution of the order tribonematales (heterokonta, xanthophyceae) based on analysis of plastidial genes rbcL and psaA Mol Phylogenet Evol. 2007 May;43(2):407-17. 119. Manes SS, Gradinger R. Small scale vertical gradients of arctic ice algal photophysiological properties Photosynth Res. 2009 Oct;102(1):53-66. 120. Margalith PZ. Production of ketocarotenoids by microalgae Appl Microbiol Biotechnol. 1999 Apr;51(4):431-8. 121. Margesin R, Feller G. Biotechnological applications of psychrophiles Environ Technol. 2010 Jul-Aug;31(8-9):835-44. 122. Marquis O, Miaud C, Ficetola GF, Boscher A, Mouchet F, Guittonneau S, et al. Variation in genotoxic stress tolerance among frog populations exposed to UV and pollutant gradients Aquat Toxicol. 2009 Nov 8;95(2):152-61. 119 123. Matsuo T, Ishiura M. Chlamydomonas reinhardtii as a new model system for studying the molecular basis of the circadian clock FEBS Lett. 2011 May 20;585(10):1495-502. 124. Matyash V, Liebisch G, Kurzchalia TV, Shevchenko A, Schwudke D. Lipid extraction by methyl-tert-butyl ether for high-throughput lipidomics J Lipid Res. 2008 May;49(5):1137-46. 125. Mayne S. Beta-carotene, carotenoids, and disease prevention in humans. FASEB J. 1996:690-701. 126. Menetrez MY. An overview of algae biofuel production and potential environmental impact Environ Sci Technol. 2012 Jul 3;46(13):7073-85. 127. Metzker ML. Sequencing technologies - the next generation Nat Rev Genet. 2010 Jan;11(1):31-46. 128. Miller J, Fogg G. Studies on the growth of xanthophyceae in pure culture. . Archiv fur Mikrobiologie. 1958;Bd. 30:1-13. 129. Miller JR, Koren S, Sutton G. Assembly algorithms for next-generation sequencing data Genomics. 2010 Jun;95(6):315-27. 130. Mohammady NG. Characterization of the fatty acid composition of nannochloropsis salina as a determinant of biodiesel properties Z Naturforsch C. 2011 Jul-Aug;66(7- 8):328-32. 131. Moo-Puc R, Robledo D, Freile-Pelegrin Y. In vitro cytotoxic and antiproliferative activities of marine macroalgae from Yucata´n, mexico. Cienc. 2009:245-58. 132. Morita RY. Psychrophilic bacteria Bacteriol Rev. 1975 Jun;39(2):144-67. 133. Murphy CF, Allen DT. Energy-water nexus for mass cultivation of algae Environ Sci Technol. 2011 Jul 1;45(13):5861-8. 134. Mussgnug JH, Thomas-Hall S, Rupprecht J, Foo A, Klassen V, McDowall A, et al. Engineering photosynthetic light capture: Impacts on improved solar energy to biomass conversion Plant Biotechnol J. 2007 Nov;5(6):802-14. 135. Myers RA, Worm B. Rapid worldwide depletion of predatory fish communities Nature. 2003 May 15;423(6937):280-3. 120 136. Nagao M. Klebsormidium flaccidum, a charophycean green alga, exhibits cold acclimation that is closely associated with compatible solute accumulation and ultrastructural changes. Plant, cell and environment. 2008;31(6):872-85. 137. Nakajima K, Yamashita T, Kita T, Takeda M, Sasaki N, Kasahara K, et al. Orally administered eicosapentaenoic acid induces rapid regression of atherosclerosis via modulating the phenotype of dendritic cells in LDL receptor-deficient mice Arterioscler Thromb Vasc Biol. 2011 Aug 4. 138. Negrisolo E, Maistro S, Incarbone M, Moro I, Dalla Valle L, Broady PA, et al. Morphological convergence characterizes the evolution of xanthophyceae (heterokontophyta): Evidence from nuclear SSU rDNA and plastidial rbcL genes Mol Phylogenet Evol. 2004 Oct;33(1):156-70. 139. Nguyen HM, Baudet M, Cuine S, Adriano JM, Barthe D, Billon E, et al. Proteomic profiling of oil bodies isolated from the unicellular green microalga chlamydomonas reinhardtii: With focus on proteins involved in lipid metabolism Proteomics. 2011 Nov;11(21):4266-73. 140. Niciura SC, Veríssimo CJ, Gromboni JG, Rocha MI, de Mello SS, Barbosa CM, Chiebao DP, Cardoso D, Silva GS, Otsuk IP, Pereira JR, Ambrosio LA, Nardon RF, Ueno TE, Molento MB. F200Y polymorphism in the β-tubulin gene in fi... [vet parasitol. 2012] - PubMed - NCBI Vet Parasit. ;2012(9/18/2012). 141. Nicolaus B, Kambourova M, Oner ET. Exopolysaccharides from extremophiles: From fundamentals to biotechnology Environ Technol. 2010 Sep;31(10):1145-58. 142. Niitsu R, Kanazashi M, Matsuwaki I, Ikegami Y, Tanoi T, Kawachi M, et al. Changes in the hydrocarbon-synthesizing activity during growth of botryococcus braunii B70 Bioresour Technol. 2011 Aug 25. 143. Oren A. Industrial and environmental applications of halophilic microorganisms Environ Technol. 2010 Jul-Aug;31(8-9):825-34. 144. Ott DW. Endosymbiotic bacteria in vaucheria (xanthophyceae): Association with cytoplasmic microtubules in vaucheria sessilis Cytobios. 1979;24(95 96):185-94. 145. 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 May;90(4):1429-41. 146. Paredes DI, Watters K, Pitman DJ, Bystroff C, Dordick JS. Comparative void- volume analysis of psychrophilic and mesophilic enzymes: Structural bioinformatics of 121 psychrophilic enzymes reveals sources of core flexibility BMC Struct Biol. 2011 Oct 20;11(1):42. 147. Park Y, Kim GD, Choi TJ. Molecular cloning and characterization of the DNA adenine methyltransferase gene in feldmannia sp. virus Virus Genes. 2007 Apr;34(2):177-83. 148. Parry BR, Shain DH. Manipulations of AMP metabolic genes increase growth rate and cold tolerance in escherichia coli: Implications for psychrophilic evolution Mol Biol Evol. 2011 Jul;28(7):2139-45. 149. Pascher A. Kryptogamenflora von deutschland, osterreich und der schweiz, vol. II. Akademische Verlagsgesellschaft M B H , Leipzig. 1939:1092. 150. Pearce DA. Climate change and the microbiology of the antarctic peninsula region Sci Prog. 2008;91(Pt 2):203-17. 151. Phadwal K, Singh PK. Effect of nutrient depletion on beta-carotene and glycerol accumulation in two strains of dunaliella sp Bioresour Technol. 2003 Oct;90(1):55-8. 152. Piette F, Struvay C, Feller G. The protein folding challenge in psychrophiles: Facts and current issues Environ Microbiol. 2011 Mar 1. 153. Pisani P, Bray F, Parkin D. Estimates of the world-wide prevalence of cancer for 25 sites in the adult population. Int J Cancer. 2002;97:72-81. 154. Quinn JC, Turner CW, Bradley TH. Scale-up of flat plate photobioreactors considering diffuse and direct light characteristics Biotechnol Bioeng. 2012 Feb;109(2):363-70. 155. Quinn PJ. Effects of temperature on cell membranes Symp Soc Exp Biol. 1988;42:237-58. 156. Radakovits R, Jinkerson RE, Fuerstenberg SI, Tae H, Settlage RE, Boore JL, et al. Draft genome sequence and genetic transformation of the oleaginous alga nannochloropis gaditana Nat Commun. 2012 Feb 21;3:686. 157. Rasala BA, Mayfield SP. The microalga chlamydomonas reinhardtii as a platform for the production of human protein therapeutics Bioeng Bugs. 2011 Jan-Feb;2(1):50-4. 158. Ratha SK, Babu S, Renuka N, Prasanna R, Prasad RB, Saxena AK. Exploring nutritional modes of cultivation for enhancing lipid accumulation in microalgae J Basic Microbiol. 2012 Jun 26. 122 159. Ratkowsky DA, Olley J, Ross T. Unifying temperature effects on the growth rate of bacteria and the stability of globular proteins J Theor Biol. 2005 Apr 7;233(3):351-62. 160. Raymond JA. Algal ice-binding proteins change the structure of sea ice Proc Natl Acad Sci U S A. 2011 Jun 14;108(24):E198. 161. Raymond JA, DeVries AL. Adsorption inhibition as a mechanism of freezing resistance in polar fishes Proc Natl Acad Sci U S A. 1977 Jun;74(6):2589-93. 162. Remias D, Karsten U, Lutz C, Leya T. Physiological and morphological processes in the alpine snow alga chloromonas nivalis (chlorophyceae) during cyst formation Protoplasma. 2010 Jul;243(1-4):73-86. 163. Rhodes CJ. Oil from algae; salvation from peak oil? Sci Prog. 2009;92(Pt 1):39-90. 164. 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 Mar 14;12:148. 165. Rousseau F, Burrowes R, Peters AF, Kuhlenkamp R, de Reviers B. A comprehensive phylogeny of the phaeophyceae based on nrDNA sequences resolves the earliest divergences C R Acad Sci III. 2001 Apr;324(4):305-19. 166. Ruangsomboon S. Effect of light, nutrient, cultivation time and salinity on lipid production of newly isolated strain of the green microalga, botryococcus braunii KMITL 2 Bioresour Technol. 2011 Jul 19. 167. Rybalka N, Andersen RA, Kostikov I, Mohr KI, Massalski A, Olech M, et al. Testing for endemism, genotypic diversity and species concepts in antarctic terrestrial microalgae of the tribonemataceae (stramenopiles, xanthophyceae) Environ Microbiol. 2009 Mar;11(3):554-65. 168. Sanghvi AM, Lo YM. Present and potential industrial applications of macro- and microalgae Recent Pat Food Nutr Agric. 2010 Nov 1;2(3):187-94. 169. Schulze E-. Global biogeochemical cycles in the climate system San Diego, Calif.: Academic Press; 2001. 170. Shimada TL, Hara-Nishimura I. Oil-body-membrane proteins and their physiological functions in plants Biol Pharm Bull. 2010;33(3):360-3. 123 171. Shimada TL, Shimada T, Takahashi H, Fukao Y, Hara-Nishimura I. A novel role for oleosins in freezing tolerance of oilseeds in arabidopsis thaliana Plant J. 2008 Sep;55(5):798-809. 172. Siddiqui KS, Cavicchioli R. Cold-adapted enzymes Annu Rev Biochem. 2006;75:403-33. 173. Silflow CD, Lefebvre PA. Assembly and motility of eukaryotic cilia and flagella. lessons from chlamydomonas reinhardtii Plant Physiol. 2001 Dec;127(4):1500-7. 174. Simionato D, Sforza E, Corteggiani Carpinelli E, Bertucco A, Giacometti GM, Morosinotto T. Acclimation of nannochloropsis gaditana to different illumination regimes: Effects on lipids accumulation Bioresour Technol. 2011 May;102(10):6026-32. 175. Simopoulos AP. New products from the agri-food industry: The return of n-3 fatty acids into the food supply Lipids. 1999;34 Suppl:S297-301. 176. Simpson JT, Wong K, Jackman SD, Schein JE, Jones SJ, Birol I. ABySS: A parallel assembler for short read sequence data Genome Res. 2009 Jun;19(6):1117-23. 177. Smith GI, Atherton P, Reeds DN, Mohammed BS, Rankin D, Rennie MJ, et al. Dietary omega-3 fatty acid supplementation increases the rate of muscle protein synthesis in older adults: A randomized controlled trial Am J Clin Nutr. 2011 Feb;93(2):402-12. 178. Smith GI, Atherton P, Reeds DN, Mohammed BS, Rankin D, Rennie MJ, et al. Omega-3 polyunsaturated fatty acids augment the muscle protein anabolic response to hyperinsulinaemia-hyperaminoacidaemia in healthy young and middle-aged men and women Clin Sci (Lond). 2011 Sep 1;121(6):267-78. 179. Smol JP, Wolfe AP, Birks HJ, Douglas MS, Jones VJ, Korhola A, et al. Climate- driven regime shifts in the biological communities of arctic lakes Proc Natl Acad Sci U S A. 2005 Mar 22;102(12):4397-402. 180. Stace CA. Plant Taxonomy and Biosystematics Cambridge University Press. 1991. 181. Starai VJ, Escalante-Semerena JC. Acetyl-coenzyme A synthetase (AMP forming) Cell Mol Life Sci. 2004 Aug;61(16):2020-30. 182. Stebbins CL, Hammel LE, Marshal BJ, Spangenberg EE, Musch TI. Effects of dietary omega-3 polyunsaturated fatty acids on the skeletal-muscle blood-flow response to exercise in rats Int J Sport Nutr Exerc Metab. 2010 Dec;20(6):475-86. 124 183. Stephenson PG, Moore CM, Terry MJ, Zubkov MV, Bibby TS. Improving photosynthesis for algal biofuels: Toward a green revolution Trends Biotechnol. 2011 Jul 18. 184. Stibal M, Sabacka M, Kastovska K. Microbial communities on glacier surfaces in svalbard: Impact of physical and chemical properties on abundance and structure of cyanobacteria and algae Microb Ecol. 2006 Nov;52(4):644-54. 185. Sun Z, Liu J, Zeng X, Huangfu J, Jiang Y, Wang M, et al. Protective actions of microalgae against endogenous and exogenous advanced glycation endproducts (AGEs) in human retinal pigment epithelial cells Food Funct. 2011 May 25;2(5):251-8. 186. Takaichi S. Carotenoids in algae: Distributions, biosyntheses and functions Mar Drugs. 2011;9(6):1101-18. 187. Tavares S, Grotkjaer T, Obsen T, Haslam RP, Napier JA, Gunnarsson N. Metabolic engineering of saccharomyces cerevisiae for production of eicosapentaenoic acid, using a novel {delta}5-desaturase from paramecium tetraurelia Appl Environ Microbiol. 2011 Mar;77(5):1854-61. 188. Thomas DN. Photosynthetic microbes in freezing deserts Trends Microbiol. 2005 Mar;13(3):87-8. 189. Ting JT, Lee K, Ratnayake C, Platt KA, Balsamo RA, Huang AH. Oleosin genes in maize kernels having diverse oil contents are constitutively expressed independent of oil contents. size and shape of intracellular oil bodies are determined by the oleosins/oils ratio Planta. 1996;199(1):158-65. 190. Tirichine L, Bowler C. Decoding algal genomes: Tracing back the history of photosynthetic life on earth Plant J. 2011 Apr;66(1):45-57. 191. Tollefson J. Heatwaves blamed on global warming Nature. 2012 Aug 7;488(7410):143-4. 192. Torri C, Samori C, Adamiano A, Fabbri D, Faraloni C, Torzillo G. Preliminary investigation on the production of fuels and bio-char from chlamydomonas reinhardtii biomass residue after bio-hydrogen production Bioresour Technol. 2011 Sep;102(18):8707-13. 193. Torto-Alalibo T, Collmer CW, Gwinn-Giglio M, Lindeberg M, Meng S, Chibucos MC, et al. Unifying themes in microbial associations with animal and plant hosts described using the gene ontology Microbiol Mol Biol Rev. 2010 Dec;74(4):479-503. 125 194. Tronelli D, Maugini E, Bossa F, Pascarella S. Structural adaptation to low temperatures--analysis of the subunit interface of oligomeric psychrophilic enzymes FEBS J. 2007 Sep;274(17):4595-608. 195. Tsai DD, Ramaraj R, Chen PH. Growth condition study of algae function in ecosystem for CO2 bio-fixation J Photochem Photobiol B. 2012 Feb 6;107:27-34. 196. Tzen J, Cao Y, Laurent P, Ratnayake C, Huang A. Lipids, proteins, and structure of seed oil bodies from diverse species Plant Physiol. 1993 Jan;101(1):267-76. 197. Tzen JT, Lie GC, Huang AH. Characterization of the charged components and their topology on the surface of plant seed oil bodies J Biol Chem. 1992 Aug 5;267(22):15626- 34. 198. Van Etten JL. Unusual life style of giant chlorella viruses Annu Rev Genet. 2003;37:153-95. 199. Van Etten JL, Gurnon JR, Yanai-Balser GM, Dunigan DD, Graves MV. Chlorella viruses encode most, if not all, of the machinery to glycosylate their glycoproteins independent of the endoplasmic reticulum and golgi Biochim Biophys Acta. 2010 Feb;1800(2):152-9. 200. van Ginneken VJ, Helsper JP, de Visser W, van Keulen H, Brandenburg WA. Polyunsaturated fatty acids in various macroalgal species from north atlantic and tropical seas Lipids Health Dis. 2011 Jun 22;10:104. 201. Vijayaraghavan K, Hemanathan K. Biodiesel production from freshwater algae. Energy Fuels. 2009;23:5448-53. 202. Vlcek D, Sevcovicova A, Sviezena B, Galova E, Miadokova E. Chlamydomonas reinhardtii: A convenient model system for the study of DNA repair in photoautotrophic eukaryotes Curr Genet. 2008 Jan;53(1):1-22. 203. Walser B, Giordano RM, Stebbins CL. Supplementation with omega-3 polyunsaturated fatty acids augments brachial artery dilation and blood flow during forearm contraction Eur J Appl Physiol. 2006 Jun;97(3):347-54. 204. Wang H, Alvarez S, Hicks LM. Comprehensive comparison of iTRAQ and label- free LC-based quantitative proteomics approaches using two chlamydomonas reinhardtii strains of interest for biofuels engineering J Proteome Res. 2011 Dec 1. 205. Watkins PA. Very-long-chain acyl-CoA synthetases J Biol Chem. 2008 Jan 25;283(4):1773-7. 126 206. Weber F, Del Campo J, Wylezich C, Massana R, Jurgens K. Unveiling trophic functions of uncultured protist taxa by incubation experiments in the brackish baltic sea PLoS One. 2012;7(7):e41970. 207. Whelan J. RC. Innovative dietary sources of N-3 fatty acids. Annu Rev Nutr. 2006;26:75-103. 208. Wiley PE, Campbell JE, McKuin B. Production of biodiesel and biogas from algae: A review of process train options Water Environ Res. 2011 Apr;83(4):326-38. 209. Wilson PW, Beaglehole D, Devries AL. Antifreeze glycopeptide adsorption on single crystal ice surfaces using ellipsometry Biophys J. 1993 Jun;64(6):1878-84. 210. Wilson SL, Walker VK. Selection of low-temperature resistance in bacteria and potential applications Environ Technol. 2010 Jul-Aug;31(8-9):943-56. 211. Woertz I, Feffer A, Lundquist T, Nelson Y. Algae grown on dairy and municipal wastewater for simultaneous nutrient removal and lipid production for biofuel feedstock. Environ Eng. 2009;135:1115-22. 212. Worm B, Hilborn R, Baum JK, Branch TA, Collie JS, Costello C, et al. Rebuilding global fisheries Science. 2009 Jul 31;325(5940):578-85. 213. Xiao N, Suzuki K, Nishimiya Y, Kondo H, Miura A, Tsuda S, et al. Comparison of functional properties of two fungal antifreeze proteins from antarctomyces psychrotrophicus and typhula ishikariensis FEBS J. 2010 Jan;277(2):394-403. 214. Yang C, Jia L, Chen C, Liu G, Fang W. Bio-oil from hydro-liquefaction of dunaliella salina over Ni/REHY catalyst Bioresour Technol. 2011 Mar;102(6):4580-4. 215. Yang ZH, Miyahara H, Hatanaka A. Chronic administration of palmitoleic acid reduces insulin resistance and hepatic lipid accumulation in KK-ay mice with genetic type 2 diabetes Lipids Health Dis. 2011 Jul 21;10(1):120. 216. Yeste M, Barrera X, Coll D, Bonet S. The effects on boar sperm quality of dietary supplementation with omega-3 polyunsaturated fatty acids differ among porcine breeds Theriogenology. 2011 Jul 1;76(1):184-96. 217. Yonezawa N, Matsuura H, Shiho M, Kaya K, Watanabe MM. Effects of soybean curd wastewater on the growth and hydrocarbon production of botryococcus braunii strain BOT-22 Bioresour Technol. 2011 Aug 3. 127 218. Yu KO, Jung J, Ramzi AB, Kim SW, Park C, Han SO. Improvement of ethanol yield from glycerol via conversion of pyruvate to ethanol in metabolically engineered saccharomyces cerevisiae Appl Biochem Biotechnol. 2011 Dec 13. 219. Yuan YV, Carrington MF, and Walsh NA. Extracts from dulse (palmaria palmata) are effective antioxidants and inhibitors of cell proliferation in vitro. Food Chem Toxicol. 2005;43:1073-81. 220. Yuan Y, Walsh N. Antioxidant and antiproliferative activities of extracts from a variety of edible seaweeds. Food Chem Toxicol. 2006;44:1144-50. 221. Zhang SO, Trimble R, Guo F, Mak HY. Lipid droplets as ubiquitous fat storage organelles in C. elegans BMC Cell Biol. 2010 Dec 8;11:96. 222. Zhou XR, Green AG, Singh SP. Caenorhabditis elegans {delta}12-desaturase FAT- 2 is a bifunctional desaturase able to desaturate a diverse range of fatty acid substrates at the {delta}12 and {delta}15 positions J Biol Chem. 2011 Oct 31.