Maximization Of Scenedesmus Dimorphus Lipid Yield For The Production Of Biodiesel

MAXIMIZATION OF SCENEDESMUS DIMORPHUS LIPID YIELD FOR THE PRODUCTION OF BIODIESEL By Carlos A. Ramos Encarnación Germano Salazar Benítez Gustavo Méndez Santos Sara Currás Medina A Capstone Project Report Submitted in Partial Fulfillment Of the Requirements for the Degree of Bachelor of Science In Chemical Engineering POLYTECHNIC UNIVERSITY OF PUERTO RICO SAN JUAN, PUERTO RICO 2010 Advisor: ___________________________________ i

Abstract ii

Acknowledgments Alessandro Anzalone, Ph.D. – Advisor Chemical Engineering Department Director Prof. Sylvia M. Vélez Villamil Biology Department University of Puerto Rico at Humacao Prof. Edgardo González, Ph.D. Negociado Servicio Forestal, Director Department of Natural and Environmental Resources iii

Table of Contents Abstract.............................................................................................................................. ii Acknowledgments ............................................................................................................ iii Chapter 1: Problem Statement ........................................................................................ 1 1.1 Introduction ........................................................................................................ 1 1.2 Research Description ......................................................................................... 2 1.3 Research Objectives ........................................................................................... 3 1.4 Research Contribution....................................................................................... 3 Chapter 2: Literature Review .......................................................................................... 5 2.1 General Information and Characteristics of Algae ................................................. 5 2.1.1 Biodiesel from Algae ........................................................................................ 5 2.1.2 Macroalgae vs. Microalgae ............................................................................. 7 2.1.3 Classification of Microalgae ............................................................................ 7 2.1.4 Properties of Green Microalgae ..................................................................... 9 2.1.5 Scenedesmus Dimorphus .................................................................................. 9 2.1.6 Production of Lipids from Microalgae ........................................................ 11 2.2 Cultivation and Harvesting of Microalgae.......................................................... 12 2.2.1 Parameters that affect the Cultivation of Microalgae ................................ 12 2.2.1.1 Effect of Light in Microalgae Cultivation ................................................. 13 iv

2.2.1.2 Effect of pH in Microalgae Cultivation..................................................... 13 2.2.1.3 Effect of Agitation and Carbon Dioxide in Microalgae Cultivation ...... 14 2.2.1.4 Effect of Temperature in Microalgae Cultivation ................................... 14 2.2.1.5 Effect of Nutrients in Microalgae Cultivation .......................................... 15 2.2.2 Microalgae Growth Dynamics ...................................................................... 16 2.2.3 Calculating the Cell Concentration of Algae ............................................... 18 2.2.4 Harvesting Algae ............................................................................................ 20 2.3 Oil Extraction ........................................................................................................ 21 2.3.1 Extraction of Lipids (Oil) .............................................................................. 21 2.3.2 Cell Disruption ............................................................................................... 22 2.4 Conversion of Algae Oil to Biodiesel ................................................................... 23 2.4.1 Transesterification ......................................................................................... 23 Chapter 3: Methodology................................................................................................. 26 3.1 Stock culture ..................................................................................................... 26 3.2 CO2 vs. Without CO2 ....................................................................................... 27 3.3 Optimization lipid yield ................................................................................... 29 3.4 Biodiesel............................................................................................................. 34 Bibliography .................................................................................................................... 36 Appendix .......................................................................................................................... 39 v

Hemacytometer............................................................................................................ 40 Colorimetric determination of Urea .......................................................................... 42 vi

List of Figures Figure 2.1: Scenedesmus Dimorphus ................................................................................9 Figure 2.2: Lipid Synthesis in Microalgae .....................................................................12 Figure 2.3: Five growth Phases of Microalgae Cultures ..............................................16 Figure 2.4: Hemacytometer Counting Area ..................................................................18 Figure 2.5: Counting Procedure for Methods A (left panel) and B (right panel) .....19 Figure 2.6: Transesterification Reaction ......................................................................24 Figure 3.1: Maximization of Lipids Experiments ........................................................32 vii

List of Tables Table 2.1: Production of Oil from Different Crops ........................................................6 Table 2.2: Properties of Major Taxonomic Groups of Algae ........................................8 Table 2.3: Algal concentration in some Puerto Rican Lakes in 1977 and 1998 .........10 Table 2.4: Chemical Composition of Algae Expressed on a Dry Matter Basis(%) ...11 Table 2.5: Modified Basal Medium ................................................................................16 viii

Chapter 1: Problem Statement 1.1 Introduction More than 60% of the oil used in the United States of America (USA) is imported. In the USA, about 25 barrels of oil per person are consumed each year, and 16 of those barrels are imported. At an average cost of $90 per barrel, imported oil costs the average person in the USA about $1,400 per year. The USA has just 3% of the known world oil reserves, yet we currently use about 25% of the world's annual oil production [1]. In order to end our country's addiction to oil and combat global warming, society is beginning to look to biofuels as a replacement energy source. Scientists are now mainly focused on photosynthetic organisms which include plants and algae. That’s because in the process of photosynthesis this organism converts sun light and carbon dioxide in to biomass and oxygen. Being the lipids, contained in the biomass of the algae, the raw material needed for the production of biodiesel. Biodiesel is a clean burning alternative fuel, produced from domestic, renewable resources. Its main advantages other than being a renewable energy source, is that it’s environmentally friendlier, burning much cleaner than fossil fuel, and that diesel engines require no modifications to it. Algae presents one of the most exciting possibilities as a future solution to our energy problems, especially that of transportation fuel. So it's hard not to get excited about algae's potential. Its basic requirements are few: carbon dioxide, sun, and water. Algae can flourish in non-arable land or in dirty water, and when it does flourish, its potential oil yield per acre is unmatched by any other terrestrial feedstock. Algae grow rapidly and can have a high percentage of lipids, or oils which conventional petroleum 1

refineries can convert into jet fuel or diesel fuel a product known as "green diesel". Algae organisms use energy from the sun to combine water with carbon dioxide (CO2) to create biomass. They can double their mass several times a day and produce at least 30 times more oil per acre than alternatives such as rapeseed, palms, soybeans, or jatropha. As the slime grows, it makes a kind of vegetable oil, similar to the oil produced from sunflower seeds or soybeans. Because algae can grow under severe conditions - extremes of temperature, pH and salinity, algae-growing facilities can be built on arid coastal land unsuitable for conventional agriculture. Key technical challenges include identifying the strains with the highest oil content and growth rates and developing cost-effective growing and harvesting methods. The hard part about algae production is growing the algae in a controlled way, harvesting it and extracting its oil efficiently [2]. 1.2 Research Description The world is currently immersed in an energy crisis, which is expected to get even worse. This is due to the limited resources of energy that human beings posses to sustain their industrial activities that require vast amounts of energy. Also almost all the energy used in the world comes from three main sources, coal, natural gas, and petroleum. All these sources of energy are available in limited quantities and are highly contaminant to our ecosystem. That’s why the search for alternatives types of energy that are friendly to our environment and come from renewable sources is so important in our times. Microalgae play an important role as one of the possible contributors for the solution of this energetic crisis. Microalgae contains an average of 40% oil per weight and produce more than 30 times the amount of oil than the highest producing plant, oil palm and using just a fraction of the area needed to cultivate the oil palm. Biodiesel from 2

microalgae is still in a very early stage, that’s why more research is needed in this area to make the plants more efficient in the production of the biodiesel. The research for this project consist mainly of three parts, finding the optimum conditions and the appropriate days of stress applied to the microalgae culture (days without nutrients), finding a way to harvest the microalgae, and being able to extract oil from the microalgae. All the experiment will be done simulating the conditions of an open pond. 1.3 Research Objectives 1. Select microalgae appropriate to Puerto Rico’s climate. 2. Evaluate how the presence of CO2 affects the microalgae cell growth and lipid yield. 3. Maximize lipid yield by varying urea (nitrogen) concentration. 4. Maximize lipid yield by varying nutrient depravation time. 5. Separate oil from microalgae. 6. Produce biodiesel using the oil from the microalgae. 7. Find the amount of microalgae needed to produce a certain amount of biodiesel. 1.4 Research Contribution Our research is focused on maximizing lipid yield for the microalgae Scenedesmus dimorphus, for the production of biodiesel, chosen specifically for Puerto Rico’s climate. First we will determine if the use of carbon dioxide during algae cultivation is beneficial to its growth and lipid content and then we will maximize the lipid content varying the urea concentration and nutrient depravation time. Based on the lack of experimental information found for the cultivation, 3

harvesting and oil extraction method, one of the biggest contributions will be the detailed documentation of this process. 4

Chapter 2: Literature Review Chapter two summarizes the main studies analyzed for our research and is divided in four main sections which are: general information and characteristics of algae, cultivation and harvesting of microalgae, oil extraction and conversion of algae oil to biodiesel. 2.1 General Information and Characteristics of Algae In section one we discuss the importance of biodiesel made from algae, the differences between using macroalgae and microalgae for the production of biodiesel, the different classifications of microalgae, the properties of green microalgae and Scenedesmus dimorphus, and finally we discuss the production of lipids from microalgae. 2.1.1 Biodiesel from Algae Algae present one of the most exciting possibilities as a future solution to our energy problems, especially that of transportation fuel. Algae are already being used in a wide variety of industries and applications, and many newer applications are being discovered. Such a wide range of end-uses enable companies to produce both fuels and [3] non-fuel products from the same algae feedstock . Biodiesel from algae in itself is not significantly different from biodiesel produced from vegetable/plant oils. All biodiesel essentially are produced using triglycerides (commonly called fats) from the plant/algal oils. Algae produce a lot of polyunsaturated, which may present a stability problem since higher levels of polyunsaturated fatty acids tend to decrease the stability of biodiesel. But polyunsaturated also have much lower melting points than monounsaturated or saturates, thus algal biodiesel should have much better cold weather properties than many other bio-feedstock. Since one of the disadvantages of biodiesel is their relatively poor 5

performance in cold temperatures, it appears that algal biodiesel might score well on this [4] point . Another reason for its preference for use and cultivation is that growing more oilseed crops would displace the food crops grown to feed mankind. Yet the most significant difference is however in the yield of algal oil, as shown on table 2.1, and hence biodiesel. Micro-algae are by a factor of 8 to 25 for palm oil, and a factor of 40 to 120 for rapeseed, the highest potential energy yield temperate vegetable oil crop [5]. Table 2.1: Production of oil from different crops[5] Oil Gallons of Oil per Year per Acre Corn 18 Soybeans 48 Safflower 83 Sunflower 102 Rapessed 127 Oil Palm 635 Microalgae 5000-15000 Some of the main advantages of using algae to produce biodiesel are: • The yields of oil and fuels from algae are much higher (10-100 times) than competing energy crops. • Algae can grow practically anywhere, thus ensuring that there is no competition with food crops. • Algae are excellent bioremediation agents - they have the potential to absorb massive amounts of CO2 and can play an important role in sewage and wastewater treatment. • Certain algal technologies have even been designed to absorb CO2 from smokestacks. 6

• Algae are the only feedstock that has the potential to completely replace world's consumption of transportation fuels. However, the algae energy field is still in its nascence. While people around the world are studying about the energy possibilities from algae and would like to know more, few resources are available that provide an overview of the algae energy industry, its potential, the status of various companies in this industry, and what the future is likely to hold for this industry [3]. 2.1.2 Macroalgae vs. Microalgae The term algae include macroalgae and microalgae. Macroalgae, are multi- cellular fast growing marine and fresh water photosynthetic protists, better known as seaweed, which grow to a considerable size, as much as 60m. In contrast microalgae are microscopic sometimes unicellular aquatic plants that normally grow in suspension within a body of water [6]. Both types of algae grow extremely quickly, this characteristic make them a promising crop for human use. Microalgae in comparison to macroalgae are known to contain large amounts of lipids within their cell structure, being then the microalgae a better choice for cultivation and production of biodiesel. 2.1.3 Classification of Microalgae The classification of algae into taxonomic groups is based upon the same rules used for the classification of land plants; research using electron microscopes has demonstrated differences in features, such as the flagella apparatus, cell division process, and organelle structure and function. The classes are then distinguished by the structure of flagellate cells (e.g., scales, angle of flagella insertion, microtubular roots, and striated roots), the nuclear division process (mitosis), the cytoplasmic division process 7

[7] (cytokinesis), and the cell covering . Table 2.2 demonstrates the different properties of the major groups of microalgae. Table 2.2: Properties of Major Taxonomic Groups of Algae [7] Storage Flagellation and Cell Taxonomy Group Chlorophyll Cartenoids Products Structure β-carotene, 1 apical flagellum in male ± -carotene Chrysolaminarin gametes: Bacillariophyta a, c rarely oils cell in two halves with fucoxanthin elaborate markings β-carotene, ± -carotene Chlorophycophyta rarely 1,2,4 to many equal apical or a, b Starch, oils (green algae) carotene subapical flagella. and lycopene, lutein. 1 or 2 unequal, apical Chrysophycophyta β-carotene, Chrysolaminarin flagella, a, c (golden algae) fucoxanthin oils in some, cell surface covered by characteristic scales Cyanobacteria β-carotene, a, c (blue-green algae) phycobilins β-carotene, Laminarin, Phaeco phycophyta ± soluble a, c 2 lateral flagella (brown algae) fucoxanthin, carbohydrates, violaxanthin oils β-carotene, peridinin, 2 lateral, 1 trailing, 1 girdling Dinophyta neoperididnin flagellum, in most, there is a a, c Starch, oils (dino flagellates) dinoxanthin, longitudinal and transverse neodinoxanth furrow and angular plates. in. β-carotene, Rhodophycophyta Floridean starch a, rarely d zeaxanthin Flagella absent (red algae ) oils ± β carotene Of all of this major taxonomic classes, the algae that are most favored by researchers is the Chlorophyceae (green algae). 8

2.1.4 Properties of Green Microalgae There are approximately 8,000 species of green algae estimated to be in existence. This group has chlorophyll a and chlorophyll b. They use starch as their primary storage component. However, N-deficiency promotes the accumulation of lipids in certain species. Green algae are the evolutionary progenitors of higher plants and, as such, they have received more attention than other groups [8]. Green algae may be unicellular, multi- cellular, colonial (living as a loose aggregation of cells) or coenocytic (composed of one large cell without cross-walls; the cell may be uninucleate or multinucleate). They have membrane-bound chloroplasts and nuclei. Most green algae are aquatic and are found commonly in fresh tropical waters. All green algae are photosynthetic (ie. autotrophic), which means that they get all of their organic carbon (energy) from photosynthesis. Green algae are generally fast growing and sturdy. They reproduce both asexually (by division of cells) and sexually [9]. 2.1.5 Scenedesmus Dimorphus Scenedesmus dimorphus shown in figure 2.1 is a green microalgae, bean shaped of approximately 10µm in size. Categorized as a heavy bacterium, Scenedesmus has a lipid content of 16-40%, being one of the preferred species for oil yield in the production of Biodiesel. One of the problems with this microalga is that it's heavy, and forms thick sediments if not kept in constant agitation. The optimal growth temperature for this strain falls between 30-35 degrees Celsius (86-95 degrees Figure 2.1: Scenedesmus dimorphus [10] 9

Fahrenheit). Scenedesmus will use any and all light it is given and should be further researched for use in mass production [10]. In Puerto Rico not only is this microalgae preferred for use due to the fact that its optimal growth temperature is favored by its climate, but it has also been found that this algae lives and grows naturally in the fresh water bodies of the island as shown by table 2.3 by a study conducted during the years 1977 and 1998. Table 2.3: Algal concentration in some Puerto Rican Lakes in 1977 and 1998 [11] Lake Sample Date Total Type and % of %Blue- % % Other Algae Algae in Highest Green Diat Algae (cells/mL) Concentration Algae oms Carraizo 1 8-1977 18.2 Actinastrum (31%) 6 37 57 Stephanodiscus 2 8-1977 18.1 18 53 29 (32%) Cylindrospermun 3 8-1977 23.1 26 38 36 (23%) 1 8-1998 201.2 Anabaena 42 59 10 31 Scenedesmus 2 8-1998 156.7 10 36 54 (51%) Dos 1 12-1977 19.8 Staurastrum (35%) 3 27 70 Bocas 2 12-1977 13.6 Staurastrum (20%) 5 55 40 2 12-1977 8.8 Staurastrum (20%) 4 35 61 Stephanodiscus 4 12-1977 9.1 6 47 47 (26%) 1 12-1977 15.9 Navicula (50%) 0 63 37 Scenedesmus 2 12-1998 101.5 7 61 32 (24%) 3 12-1998 65.9 Staurastrum (37%) 12 49 39 All algae primarily comprise of the following, in varying proportions: Proteins, Carbohydrates, Fats and Nucleic Acids. While the percentages vary with the type of algae, there are algae types that are comprised up to 40% of their overall mass by fatty acids. It is this fatty acid (oil) that can be extracted and converted into biodiesel. Table 2.4 demonstrates and compares the chemical compositions of different strains of algae [12] . 10

Table 2.4: Chemical Composition of Algae Expressed on a Dry Matter Basis (%) [12] Strain Protein Carbohydrate Lipid Content Nucleic Acid Scenedesmus obliquus 50-56 10-17 12-14 3-6 Scenedesmus quadricauda 47 - 1.9 - Scenedesmus dimorphus 8-18 21-52 16-40 - Chlamydomonas - rheinhardii 48 17 21 Chlorella vulgaris 51-58 12-17 14-22 4-5 Chlorella pyrenoidosa 57 2 2 - Spirogyra sp. 6-20 33-64 11-21 - Dunaliella bioculata 49 4 8 - Dunaliella salina 57 32 6 - Euglena gracilis 39-61 14-18 14-20 - Prymnesium parvum 28-45 25-33 22-38 1-2 Tetraselmis maculata 52 15 3 - Porphyridium cruentum 28-39 40-57 9-14 - Spirulina platensis 46-63 8-14 4-9 2-5 Spirulina maxima 60-71 13-16 6-7 3-4.5 Synechoccus sp. 63 15 11 5 Anabaena cylindrica 43-56 25-30 4-7 - 2.1.6 Production of Lipids from Microalgae The production of biodiesel is based upon the quantity of fatty acids (lipids) that the alga reaches to produce during its growth. Microalgae are autotrophic organisms and as such they do photosynthesis which allows them to create their own food. It is during photosynthesis that microalgae consumes carbon dioxide in the presence of sunlight to grow its biomass and produce oxygen, biomass is composed of carbohydrates, proteins and lipids. Lipids are esters, composed largely of carbon, oxygen, and hydrogen. They store energy and are essential for cell growth; its chain length is usually between C12 to C20. The main importance in lipid production is that they can be converted via a process called transesterification, into biodiesel fuel. The following figure demonstrates the synthesis in which microalgae produce lipids and carbohydrates. 11

Figure 2.2: Lipid Synthesis in Microalgae 2.2 Cultivation and Harvesting of Microalgae In section two we discuss the parameters that affect the cultivation of microalgae which are: light, pH, temperature, agitation, carbon dioxide and nutrients. We also discuss the growth dynamics of the microalgae, a way to calculate the cell concentration of algae using a hemacytometer, and finally a way to harvest the microalgae using chemical flocculation. 2.2.1 Parameters that affect the Cultivation of Microalgae The most important parameters regulating algal growth are light, pH, temperature, agitation, nutrients and carbon dioxide. The most optimal parameters as well as the tolerated ranges are species specific. Also, the various factors may be interdependent and 12

a parameter that is optimal for one set of conditions is not necessarily optimal for another [13] . 2.2.1.1 Effect of Light in Microalgae Cultivation As with all plants, micro-algae photosynthesize assimilating inorganic carbon for conversion into organic matter. Light is the source of energy which drives this reaction and in this regard intensity, spectral quality and photoperiod need to be considered. Light intensity plays an important role, but the requirements vary greatly with the culture depth and the density of the algal culture: at higher depths and cell concentrations the light intensity must be increased to penetrate through the culture. Too high light intensity (e.g. direct sun light, small container close to artificial light) may result in photo-inhibition [13]. Photoinhibition is light-induced reduction in the photosynthetic capacity of the algae. In most algal-cultivation systems, light only penetrates the top 3 inches (7.6 cm) to 4 inches (10 cm) of the water. This is because as the algae grow and multiply, they become so dense that they block light from reaching deeper into the pond or tank. Algae only need about 1/10th the amount of light they receive from direct sunlight. Direct sunlight is often too strong for algae, so in order to have ponds that are deeper than 4 inches algae growers use various methods to agitate the water in their ponds, thus circulating the algae so that it does not remain on the surface, which would cause it to be over-exposed [14]. 2.2.1.2 Effect of pH in Microalgae Cultivation Maintenance of an acceptable pH range throughout culturing is of the outmost importance as it impacts all aspects of media biochemistry. The pH range for most cultured algal species is between 7 and 9, with the optimum range being 8.2-8.7. Complete culture collapse due to the disruption of many cellular processes can result 13

from a failure to maintain an acceptable pH. The latter is accomplished by aerating the culture. In the case of high-density algal culture, the addition of carbon dioxide allows to correct for increased pH, which may reach limiting values of up to pH 9 during algal growth [13]. 2.2.1.3 Effect of Agitation and Carbon Dioxide in Microalgae Cultivation One of the problems as mentioned before of our microalgae Scenedesmus dimorphus is that it's heavy, and forms thick sediments if not kept in constant agitation. Agitation is not only necessary to prevent sedimentation of the algae, but also ensures that all cells of the population are equally exposed to the light and nutrients, and also improves gas exchange between the culture medium and the air. The latter is of primary importance as the air contains the carbon source for photosynthesis in the form of carbon dioxide. For very dense cultures, the CO2 originating from the air (containing 0.03% CO2) bubbled through the culture is limiting the algal growth and pure carbon dioxide may be supplemented to the air supply (e.g. at a rate of 1% of the volume of air). CO2 addition furthermore buffers the water against pH changes as a result of the CO2/HCO3- balance [13]. 2.2.1.4 Effect of Temperature in Microalgae Cultivation Most commonly cultured species of micro-algae tolerate temperatures between 16 and 27°C. Temperatures lower than 16°C will slow down growth, whereas those higher than 35°C are lethal for a number of species. If necessary, algal cultures can be cooled by a flow of cold water over the surface of the culture vessel or by controlling the air temperature with refrigerated air - conditioning units [13]. The optimal growth temperature varies between strains, for our strain Scenedesmus dimorphus its optimal growth 14

temperature falls between 30-35ºC (86-95ºF) , which is also in the range of temperatures year round in Puerto Rico. 2.2.1.5 Effect of Nutrients in Microalgae Cultivation Algal nutrient solutions are made up of a mixture of chemical salts and water. Sometimes referred to as "Growth Media", nutrient solutions (along with carbon dioxide and light), provide the materials needed for algae to grow. Nutrient solutions, as opposed to fertilizers, are designed specifically for use in aquatic environments and their composition is much more precise [15]. Based on experiments, done by the University of Setsunan, in Japan, on the algae Scenedesmus dimorphus, its results show that nitrogen and phosphorous contents, [16] especially nitrogen content, are the most important factors regulating its growth . Nitrogen source and concentration in the growth media greatly influence algae lipid yield [17] . In nitrogen limited situations, algae lipid content usually increases as a mechanism of survival, which makes cells stop its divisions and start to store energy in the form of lipids. There are many reports that suggest that the content of dry lipids in various strains may duplicate during nitrogen deprivation. Biomass growth is often inhibited in nitrogen- lacking situations, so there is usually a lipid yield peak for each algal strain at certain [17] nitrogen types and concentrations . Each strain of alga prefers different kind of [18] nitrogen sources . For our strain, Scenedesmus dimorphus, it has been proven by experimental data that the best source for its growth is Urea. Urea is an organic compound with the chemical formula (NH2)2CO. The experiment carried out by the Kansas State University proved effective the use of Basal Medium slightly modified, 15

replacing the nitrogen source which is KNO3 for 1.8g of Urea [(NH2)2CO]. Each liter of water must contain the following quantities stated by table 2.4. Table 2.5: Modified Basal Medium [17] Chemical Quantity (mg) (NH2)2CO 1800 KH2PO4 1250 MgSO4*7H2O 1000 EDTA 500 H3BO3 114.2 CaCl2*2H2O 111 FeSO4*7H2O 49.8 ZnSO4*7H2O 88.2 MnCl2*4H2O 14.2 CuSO4*5H2O 15.7 Co(NO3)2*6H2O 4.9 2.2.2 Microalgae Growth Dynamics The growth of an axenic culture (contains only one microbial species) of microalgae is characterized by five phases as shown on the following figure. Figure 2.3: Five growth phases of micro-algae cultures [13] 16

1. Lag or Induction Phase; This phase, during which little increase in cell density occurs, is relatively long when an algal culture is transferred from a plate to liquid culture. Cultures inoculated with exponentially growing algae have short lag phases, which can seriously reduce the time required for up scaling. The lag in growth is attributed to the physiological adaptation of the cell metabolism to growth, such as the increase of the levels of enzymes and metabolites involved in cell division and carbon fixation. 2. Exponential Phase; During the second phase, the cell density increases as a function of time t according to a logarithmic function: Ct = C0.emt (Equation 1) with Ct and C0 being the cell concentrations at time t and 0, respectively, and m = specific growth rate. The specific growth rate is mainly dependent on algal species, light intensity and temperature. 3. Phase of Declining Growth Rate; Cell division slows down when nutrients, light, pH, carbon dioxide or other physical and chemical factors begin to limit growth. 4. Stationary Phase; In the fourth stage the limiting factor and the growth rate are balanced, which results in a relatively constant cell density. 5. Death or “Crash” Phase; During the final stage, water quality deteriorates and nutrients are depleted to a level incapable of sustaining growth. Cell density decreases rapidly and the culture eventually collapses. 17

In practice, culture crashes can be caused by a variety of reasons, including the depletion of a nutrient, oxygen deficiency, overheating, pH disturbance, or contamination. The key to the success of algal production is maintaining all cultures in the exponential phase of growth. Moreover, the nutritional value of the produced algae is inferior once the culture is beyond phase 3 due to reduced digestibility, deficient composition, and possible production of toxic metabolites [13]. 2.2.3 Calculating the Cell Concentration of Algae The exact number of cells in a culture or preparation may be very important for many reasons, consistency being a major one. One of the easiest ways to count cells is under a light microscope with the hemacytometer. The hemacytometer is basically a thick glass slide with a counting chamber. The central portion of this chamber is the counting platform with one or two extremely precise grids. The counting platform is covered by an even surfaced cover slip. This gives a very precise volume in the space delimited by the grid and the cover slip as shown in Figure 2.4 [19]. Figure 2.4: Hemacytometer Counting Area [20] Here we have two simple methods for counting cells based on the surface area of the hemacytometer used to determine cell count. Other counting schemes are acceptable 18

also. The choice of methods depends upon the cell concentration - the accuracy of the procedure depends upon the number of cells counted. When cell concentration is low, one should count more grids [21]. Method A Count the number of cells in the 4 outer squares (see the left panel of Figure 2.5). The cell concentration is calculated as follows: Cell concentration per milliliter = Total cell count in 4 squares x 2500 x dilution factor Example: If one counted 450 cells after diluting an aliquot of the cell suspension 1:10, the original cell concentration = 450 x 2500 x 10 = 11,250,000/ml Method B Estimate cell concentration by counting 5 squares in the large middle square (see the right panel in Figure 2.5). The cell concentration is calculated as follows: Cell concentration per milliliter = Total cell count in 5 squares x 50,000 x dilution factor Example: If one counted 45 cells after diluting an aliquot of the cell suspension 1:10, the original cell concentration = 45 x 50,000 x 10 = 22,500,000/ml Figure 2.5: Counting procedure for Methods A (left panel) and B (right panel) [21] 19

2.2.4 Harvesting Algae Chemical Flocculation Algal harvesting is one of the major factors that must be overcome in order for algae to be used as a fuel source. The problem is that microalgae mass cultures are dilute, typically less than 500 mg/l on a dry weight organic basis, and the cell are very small. In order to be processed into biodiesel the algae must be in the form of a paste that is 15 % solids [22]. Many different algae harvesting processes have been studied between them we have; centrifugation, chemical flocculation, ultrasound and froth flotation. Chemical flocculation is the action of polymers to form bridges between the flocs and bind the particles into large agglomerates or clumps. Bridging occurs when segments of the polymer chain adsorb on different particles and help particles aggregate. An anionic flocculant will react against a positively charged suspension, absorbing on the particles and causing destabilization either by bridging or charge neutralization. In this process it is essential that the flocculating agent be added by slow and gentle mixing to allow for contact between the small flocs and to agglomerate them into larger particles. The newly formed agglomerated particles are quite fragile and can be broken apart by shear forces during mixing. Care must also be taken to not overdose the polymer as doing so will cause settling/clarification problems. Anionic polymers themselves are lighter than water. As a result, increasing the dosage will increase the tendency of the floc to float and not settle. Once suspended particles are flocculated into larger particles, they can usually be removed from the liquid by sedimentation, provided that a sufficient density difference exists between the suspended matter and the liquid. Such particles can also be removed or separated by media filtration, straining or floatation. The flocculation 20

reaction not only increases the size of the floc particles to settle them faster, but also affects the physical nature of the floc, making these particles less gelatinous and thereby easier to dewater [23]. For microalgae harvesting the most used chemical are; lime, alum, or chitosan. When added to the algae pond solution it causes charge neutralization of the algae, [22] resulting in the algae clumping together . For our investigation we will be working with aluminum sulfate as our flocculating agent. Aluminium sulfate is sometimes incorrectly referred to as alum, its chemical formula is Al2(SO4)3, and is used industrially as a flocculating agent in the purification of drinking water and waste water treatment plants [24]. 2.3 Oil Extraction In section three we discuss the importance and difficulty of extracting the lipids from the microalgae and a chemical method for the cell disruption and extraction of the oil using solvents. 2.3.1 Extraction of Lipids (Oil) Oil extraction from algae is a hotly debated topic currently because this process is one of the most costly processes which can determine the sustainability of algae-based biodiesel. In terms of the concept, the idea is quite simple: Extract the algae from its growth medium (using an appropriate separation process), and use the wet algae to extract the oil [25]. Yet in reality to achieve this is quite difficult. The aim of all extraction procedures is to separate cellular or fluid lipids from the other constituents, proteins, polysaccharides, small molecules (amino acids, sugars...) but also to preserve these lipids for further analyses or uses. There is a great diversity of methodologies because 21

biological tissues are not similar when considering their structure, texture, sensitivities and lipid contents. Removing the non-lipids without losing some lipids is a complex [26] challenge . To extract the oil from the algae there are a few methods which includes mechanical, chemical or a combination of both. 2.3.2 Cell Disruption Solvent Extraction (Chemical Method) A lot of biological molecules are inside the cell, and they must be released from it. This is achieved by cell disruption (lysis). Cell disruption is a sensitive process because of the cell wall's resistance to the high osmotic pressure inside them. Furthermore, difficulties arise from a non-controlled cell disruption that results from an unhindered release of all intracellular products (proteins nucleic acids, cell debris) as well as the requirements for cell disruption without the desired product's denaturation. There are mechanical and non-mechanical cell disruption methods. The results of these methods are often evaluated in terms of the activity level of a cellular enzyme released to the disrupted suspension, combining the efficiency of the disrupting process with an estimate of the degree of cell disruption. Many chemical methods have been employed in order to extract intra cellular components from micro organisms by permeabilizing the outer-wall barriers. It can be achieved with organic solvents that act by creation of canals through the cell membrane: toluene, ether, phenyl ethyl alcohol DMSO, benzene, methanol, [27] chloroform, isopropyl and others can achieve this . For our case algal oil can be extracted using chemicals, the type and quantity of solvent varies between algae strains. The downside to using solvents for oil extraction and not other methods like mechanical ones is the inherent dangers involved in working with chemicals. Among hydrocarbons, 22

hexane is the most popular solvent, is relatively inexpensive, but is a good solvent only for lipids of low polarity. Its main use is to extract neutral lipids from mixtures of water [26] with alcohols . Hexane is an alkane hydrocarbon with the chemical formula C6H14. Hexane isomers are largely unreactive, and are frequently used as an inert solvent in [28] organic reactions because they are very non-polar . Alcohols are also very good solvents for most lipids, one of its most common is isopropyl alcohol, commonly known as isopropyl, a colorless, flammable chemical compound with a strong odor. It has the molecular formula C3H7OH and is the simplest example of a secondary alcohol, where the alcohol carbon is attached to two other carbons. It is an isomer of propanol. Isopropyl alcohol is cheaply available. Like acetone, it dissolves a wide range of nonpolar compounds. It is also relatively nontoxic and dries (evaporates) quickly. Thus it is used widely as a solvent and as a cleaning fluid [29]. 2.4 Conversion of Algae Oil to Biodiesel In our fourth and final section we discuss a process called transesterification; this process is used to convert our algae oil into biodiesel. 2.4.1 Transesterification Almost all biodiesel is produced using base catalyzed transesterification as it is the most economical process requiring only low temperatures and pressures and producing a 98% conversion yield. The transesterification process is the reaction of a triglyceride (fat/oil) with an alcohol to form esters and glycerol. A triglyceride has a glycerine molecule as its base with three long chain fatty acids attached. The characteristics of the fat are determined by the nature of the fatty acids attached to the glycerine. The nature of the fatty acids can in turn affect the characteristics of the 23

biodiesel. During the transesterification process, the triglyceride is reacted with alcohol, in the presence of a catalyst, usually a strong alkaline like sodium hydroxide. The alcohol reacts with the fatty acids to form the mono-alkyl ester, or biodiesel and crude glycerol. In most production methanol or ethanol is the alcohol used (methanol produces methyl esters, ethanol produces ethyl esters) and is a base catalysed by either potassium or sodium hydroxide. Potassium hydroxide has been found to be more suitable for the ethyl ester biodiesel production, either base can be used for the methyl ester. The figure below shows the chemical process for methyl ester biodiesel. The reaction between the fat or oil and the alcohol is a reversible reaction and so the alcohol must be added in excess to drive the reaction towards the right and ensure complete conversion. Figure 2.6: Transesterification Reaction [30] The products of the reaction are the biodiesel itself and glycerol. A successful transesterification reaction is signified by the separation of the ester and glycerol layers after the reaction time. The heavier, co-product, glycerol settles out and may be sold as it is or it may be purified for use in other industries, e.g. the pharmaceutical, cosmetics etc. Straight vegetable oil (SVO) can be used directly as a fossil diesel substitute however using this fuel can lead to some fairly serious engine problems. Due to its relatively high viscosity SVO leads to poor atomization of the fuel, incomplete combustion, coking of the fuel injectors, ring carbonization, and accumulation of fuel in the lubricating oil. The 24

best method for solving these problems is the transesterification of the oil. The engine combustion benefits of the transesterification of the oil are: • Lowered viscosity • Complete removal of the glycerides • Lowered boiling point • Lowered flash point • Lowered pour point [30] 25

Chapter 3: Methodology Chapter three describes a step by step procedure needed to accomplish our research objectives. 3.1 Stock culture 3.1.1 Configuration and growth 3.1.1.1 Prepare the growth media in an Erlenmeyer of 1000mL. 3.1.1.1.1 water 200 mL aprox. 3.1.1.1.2 1250 mg KNO3 3.1.1.1.3 1250 mg KH2PO4 3.1.1.1.4 1000 mg MgSO4.7H2O 3.1.1.1.5 500 mg EDTA 3.1.1.1.6 114.2 mg H3BO3 3.1.1.1.7 111 mg CaCl2.2H2O 3.1.1.1.8 49.8 mg FeSO4.7H2O 3.1.1.1.9 88.2 mg ZnSO4.7H2O 3.1.1.1.10 14.2 mg MnCl2.4H2O 3.1.1.1.11 15.7 mg CuSO4.5H2O 3.1.1.1.12 4.9 mg Co(NO3)2.6H2O 3.1.1.1.13 Fill the Erlenmeyer to the top (1000mL) with water. 3.1.1.2 Add the algae from the stock culture. 3.1.1.3 Leave the algae to grow and reproduce for about 7 days before using it. 26

3.2 CO2 vs. Without CO2 For this experiment we want to determine if the use of carbon dioxide during algae cultivation is beneficial to its growth and lipid content. 3.2.1 Configuration and growth 3.2.1.1 Prepare the growth media in an Erlenmeyer of 1000mL. 3.2.1.1.1 water 200 mL aprox. 3.2.1.1.2 1.8g/L urea 3.2.1.1.3 1250 mg KH2PO4 3.2.1.1.4 1000 mg MgSO4.7H2O 3.2.1.1.5 500 mg EDTA 3.2.1.1.6 114.2 mg H3BO3 3.2.1.1.7 111 mg CaCl2.2H2O 3.2.1.1.8 49.8 mg FeSO4.7H2O 3.2.1.1.9 88.2 mg ZnSO4.7H2O 3.2.1.1.10 14.2 mg MnCl2.4H2O 3.2.1.1.11 15.7 mg CuSO4.5H2O 3.2.1.1.12 4.9 mg Co(NO3)2.6H2O 3.2.1.1.13 Fill the Erlenmeyer to the top (1000mL) with water. 3.2.1.2 Using two beakers of 250 mL prepare two cultures in a growth media at 90˚C. 3.2.1.2.1 The first one will contain CO2. 3.2.1.2.1.1 The CO2 will be injected using a diffuser. 3.2.1.2.1.2 Also the CO2 agitation will prevent the sedimentation of the culture. 3.3.1.2.1.3 Fill a 250ml beaker to 227ml with the growth media. 27

3.3.1.2.1.4 Add 23ml from the stock solution. 3.2.1.2.2 The second one will be without using CO2. 3.2.1.2.2.1 To prevent sedimentation the culture will be place in a shaker table. 3.3.1.2.2.2 Fill a 250ml beaker to 227ml with the growth media. 3.3.1.2.2.3 Add 23ml from the stock solution. 3.2.1.2.3 Measure the growing rate using a hemacytometer every day. 3.2.1.2.4 Measure the nitrogen consumption by the culture every day. 3.2.1.2.5 Record the data for 17 days. 3.2.2 Lipid Yield 3.2.2.1 After 17 days of algae culturing, extract 100 mL from the culture and transfer it to a 250 mL beaker. 3.2.2.2 Using aluminum sulfate as flocculant, flocculate the sample. 3.2.2.3 With a 1.5µm filter and vacuum, filtrate the sample using a vacuum Erlenmeyer of 1000 ml. 3.2.2.4 Measure and record the weight of a 500 mL bottle with cap. 3.2.2.5 Deposit the biomass contained in the filter in to the weighted bottle and measure again. 3.2.2.6 For every gram of algae biomass add 18 ml of a mixture of Hexane/Isopropyl (3:2) (solvent) and agitate manually until the biomass is dissolved. 3.2.2.7 Measure the weight of a vacuum Erlenmeyer of 1000 mL. 3.2.2.8 With a 1.5µm filter and vacuum, filtrate the sample using the previously weighed Erlenmeyer. 3.2.2.9 Put the Erlenmeyer with the filtrated solution in the hood. 28

3.2.2.10 Using a heat plate, apply heat to the solution (95˚C) to evaporate the solvent solution. 3.2.2.11With the solution dry, measure the weight and evaluate the results. 3.2.3 Dry weight biomass (DW) 3.2.3.1 After 17 days put a filter paper in an oven at 75˚C for 5 hours. 3.2.3.2 Measure the weight of the paper. 3.2.3.3 Take 30 mL from the culture and filtrate in a vacuum Erlenmeyer of 1000mL. 3.2.3.4 After filtrating the sample put the filter paper in the oven at 75˚C for 5 hours. 3.2.3.5 Leave the sample overnight in the desecator. 3.2.3.6 Measure the weight of the sample. 3.2.3.7 Compare the weight of the dry lipids vs. the dry biomass. 3.2.4 Comparing results 3.2.4.1 Using the results obtained in the previous analysis determine the effect of using CO2 in the growing period. Choose the better way to growth the culture, with CO2 or without CO2. 3.3 Optimization lipid yield For this experiment we want to maximize the lipid yield varying the urea concentration and nutrient depravation time. 3.3.1 Configuration and growth 3.3.1.1 Prepare 15 cultures in 15 beakers of 250 mL. (fig. 3.1) 3.3.1.1.1 First block (5 beakers) 3.3.1.1.1.1 Prepare the growth media in an Erlenmeyer of 1000mL. 3.3.1.1.1.1.1 Distillate water 200 mL aprox. 29

3.3.1.1.1.1.2 1.2g/L urea 3.3.1.1.1.1.3 1250 mg KH2PO4 3.3.1.1.1.1.4 1000 mg MgSO4.7H2O 3.3.1.1.1.1.5 500 mg EDTA 3.3.1.1.1.1.6 114.2 mg H3BO3 3.3.1.1.1.1.7 111 mg CaCl2.2H2O 3.3.1.1.1.1.8 49.8 mg FeSO4.7H2O 3.3.1.1.1.1.9 88.2 mg ZnSO4.7H2O 3.3.1.1.1.1.10 14.2 mg MnCl2.4H2O 3.3.1.1.1.1.11 15.7 mg CuSO4.5H2O 3.3.1.1.1.1.12 4.9 mg Co(NO3)2.6H2O 3.3.1.1.1.1.13 Fill the Erlenmeyer to the top (1000mL) with water. 3.3.1.1.1.2 Fill a 250ml beaker to 227ml with the growth media. 3.3.1.1.1.3 Add 23ml from the stock solution. 3.3.1.1.2 Second block (5 beakers) 3.3.1.1.2.1 Prepare the growth media in an Erlenmeyer of 1000mL. 3.3.1.1.2.1.1 Distillate water 200 mL aprox. 3.3.1.1.2.1.2 1.8g/L urea 3.3.1.1.2.1.3 1250 mg KH2PO4 3.3.1.1.2.1.4 1000 mg MgSO4.7H2O 3.3.1.1.2.1.5 500 mg EDTA 3.3.1.1.2.1.6 114.2 mg H3BO3 3.3.1.1.2.1.7 111 mg CaCl2.2H2O 30

3.3.1.1.2.1.8 49.8 mg FeSO4.7H2O 3.3.1.1.2.1.9 88.2 mg ZnSO4.7H2O 3.3.1.1.2.1.10 14.2 mg MnCl2.4H2O 3.3.1.1.2.1.11 15.7 mg CuSO4.5H2O 3.3.1.1.2.1.12 4.9 mg Co(NO3)2.6H2O 3.3.1.1.2.1.13 Fill the Erlenmeyer to the top (1000mL) with water. 3.3.1.1.2.2 Fill a 250ml beaker to 227ml with the growth media. 3.3.1.1.2.3 Add 23ml from the stock solution. 3.3.1.1.3 Third block (5 beakers) 3.3.1.1.3.1 Prepare the growth media in an Erlenmeyer of 1000mL. 3.3.1.1.3.1.1 Distillate water 200 mL aprox. 3.3.1.1.3.1.2 2.4 g/L urea 3.3.1.1.3.1.3 1250 mg KH2PO4 3.3.1.1.3.1.4 1000 mg MgSO4.7H2O 3.3.1.1.3.1.5 500 mg EDTA 3.3.1.1.3.1.6 114.2 mg H3BO3 3.3.1.1.3.1.7 111 mg CaCl2.2H2O 3.3.1.1.3.1.8 49.8 mg FeSO4.7H2O 3.3.1.1.3.1.9 88.2 mg ZnSO4.7H2O 3.3.1.1.3.1.10 14.2 mg MnCl2.4H2O 3.3.1.1.3.1.11 15.7 mg CuSO4.5H2O 3.3.1.1.3.1.12 4.9 mg Co(NO3)2.6H2O 3.3.1.1.3.1.13 Fill the Erlenmeyer to the top (1000mL) with water. 31

3.3.1.1.3.2 Fill a 250ml beaker to 227ml with the growth media. 3.3.1.1.3.3 Add 23ml from the stock solution. 3.3.1.2 Measure the growing rate using a hemacytometer every day. 3.3.1.3 Measure the nitrogen consumption by the culture every day. Figure 3.1: Maximization of Lipids Experiments 3.3.1.3.1 Measuring the nitrogen consumption, we are looking to analyze the starvation days. When we identify that the nitrogen is consumed, we will examine the first beaker in this row. For example, if the first row to consume the nitrogen is row 1, we will begin examining the beaker 1A. The next day we will look for the next one, 1B then the 1C, 1D and finally 1E. This procedure will be done with all the rows. 3.3.2 Lipid Yield 3.3.2.1 After nitrogen consumption; using aluminum sulfate as flocculant, flocculate the sample. 3.3.2.2 With a 1.5µm filter and vacuum, filtrate the sample using a vacuum Erlenmeyer of 1000 mL. 32

3.3.2.3 Measure the weight of a 500 mL bottle with cap. 3.3.2.4 Deposit the biomass contained in the filter in to the weighted bottle and measure again. 3.3.2.5 For every gram of biomass add 18 ml of a mixture of Hexane/Isopropyl (3:2) (solvent) and agitate manually until the biomass is dissolved. 3.3.2.6 Measure the weight of a vacuum Erlenmeyer of 1000 mL. 3.3.2.7 With a 1.5µm filter and vacuum, filtrate the sample using the measured Erlenmeyer. 3.3.2.8 Transfer the Erlenmeyer with the filtrated solution to the hood. 3.3.2.9 Using a heat plate, apply heat to the solution (95˚C) to evaporate the solvent solution. 3.3.2.10 With the dry solution measure the weight and evaluate the results. 3.3.3 Dry weight biomass (DW) 3.3.3.1 After 17 days put the filter paper in an oven at 75˚C for 5 hours. 3.3.3.2 Measure the weight of the paper. 3.3.3.3 Take 30 mL from the culture and filtrate in a vacuum Erlenmeyer of 1000mL. 3.3.3.4 After filtrate the sample put the filter paper in the oven at 75˚C for 5 hours. 3.3.3.5 Leave overnight the sample in the desecator. 3.3.3.6 Measure the weight of the sample. 3.3.4 Comparing results 3.3.4.1 Using the results obtained in the previous analysis determinate the best urea concentration, and the optimal starving period for the algae growth. 33

3.4 Biodiesel 3.4.1 Configuration and growth 3.4.1.1 Using a 1000 mL beaker prepare a culture using all the parameters that have been determined in the previous procedures. 3.4.1.2 Measure the growing rate using a hemacytometer every day. 3.4.1.3 Measure the nitrogen consumption by the culture every day. 3.4.1.4 When the culture is prepared to be processes, using aluminum sulfate as flocculant, flocculate the sample. 3.4.1.5 With a 1.5µm filter and vacuum, filtrate the sample using a vacuum Erlenmeyer of 1000 mL. 3.4.1.6 Measure the weight of a 500 mL bottle with cap. 3.4.1.7 Deposit the biomass contained in the filter in to the weighted bottle and measure again. 3.4.1.8 For every gram of biomass add 18 ml of a mixture of Hexane/Isopropyl (3:2) (solvent) and agitate manually until the biomass is dissolved. 3.4.1.9 With a 1.5µm filter and vacuum, filtrate the sample using the measured Erlenmeyer. 3.4.1.10 Separate by distillation the solvent from the oil. 3.4.1 Transterification 3.4.1.1 Reaction 3.4.1.1.1 Add the extracted algae oil to a 1 liter flask 34

3.4.1.2.1 In another flask, add 1% by weight of oil of KOH and ethanol in a quantity that corresponds to 100% excess of the stoichiometric amount required for the transesterification. 3.4.1.3.1 Heat the ethanol if needed, in order to be able to dissolve the KOH completely. 3.4.1.4.1 Add the ethanol and the dissolved catalyst to the oil and start the agitation. 3.4.1.5 .1 Extremely vigorous agitation is needed to maintain the reaction. 3.4.1.2 Phase Separation 3.4.1.2 .1 After 120 minutes of reaction time, the reaction is stopped. 3.4.1.2 .2 The reaction mixture is allowed to stand overnight while phase separation occurs. 3.4.1.2 .3 Decant the ester phase from the equilibrium mixture. 3.4.1.3 Washing 3.4.1.3.1 Wash the excess alcohol and residual catalyst from the ester. 3.4.1.3.2 Place the ester phase in a glass column. 3.4.1.3.3 Spray water into the top of the glass column. 3.4.1.3.4 The excess alcohol and catalyst were removed by the water as it percolated throught the column. 3.4.1.3.5 A period of 24-48 hours is required for the water phase containing alcohol, catalyst and emulsified ester to settle and the ester phase to become clear. 35

Bibliography 1. “Oil Crisis”. Health and Energy. 13 July 2009 <http://healthandenergy.com/oil_crisis.htm>. 2. “Biodiesel Fuel”. SECO: State Energy Conservation Office. 13 July 2009 <http://www.seco.cpa.state.tx.us/re_biodiesel.htm>. 3. “Why are algae so exciting from a renewable energy standpoint?” Oilgae. 2006. 10 June 2009 < http://www.oilgae.com/ref/report/digest/digest.html>. 4. “Algal Biodiesel Characteristics & Properties”. Oilgae. 2006. 10 June 2009 <http://www.oilgae.com/algae/oil/biod/char/char.html>. 5. Riesing, Thomas F. “Cultivating Algae for Liquid Fuel Production”. Permaculture Activism. 29 March 2007. 10 June 2009 <http://www.oakhavenpc.org/cultivating_algae.htm>. 6. Chang, H. “Marine Biodiversity and Systematics”. Research Programmes. 2007. 11 June 2009 <http://www.niwa.cri.nz/rc/prog/marinebiodiversity/obj7>. 7. “Classification of Algae”. Oilgae. 2006. 11 June 2009 <http://www.oilgae.com/algae/cla/cla.html >. 8. Sheehan, John, Terri Dunahay, John Benemann, and Paul Roessler. A Look Back at the U.S. Department of Energy’s Aquatic Species Program—Biodiesel from Algae. Colorado: National Renewable Energy Laboratory, 1998. 9. Guiry, Michael. “Chlorophyta: Green Algae”. Welcome to Michael Guiry's Seaweed Site. 9 June 2009. 12 June 2009 <http://www.seaweed.ie/algae/chlorophyta.html>. 10. “Scenedesmus Dimorphus-Algae Culture”. Algae Depot. 2009. 12 June 2009 <http://www.algaedepot.com/servlet/the-1/Scenedesmus-dimorphus--dsh-- Algae/Detail>. 11. Martínez, Gustavo A. PhD. Determination of Numeric Nutrient Target Criteria in Lakes and Reservoirs in Puerto Rico. Rio Piedras, Puerto Rico: University of Puerto Rico, 2004. 12. “Algal Chemical Composition”. Oilage. 2006. 12 June 2009 <http://www.oilgae.com/algae/comp/comp.html>. 13. Lavens, Patrick, and Patrick Sorgeloos. Manual on the Production and Use of Live Food for Aquaculture. Ghent, Belgium: University of Ghent, 1996. 36

14. “Algaculture”. Wikipedia the free encyclopedia. 3 June 2009. 16 June 2009 <http://en.wikipedia.org/wiki/Algaculture>. 15. “Algal Nutrients Solutions”. Wikipedia the free encyclopedia. 1 March 2009. 17 June 2009 <http://en.wikipedia.org/wiki/Algal_nutrient_solutions>. 16. Kunikane. S, M. Kakeko, and R. Maehara. Growth and Nutrient Uptake of Green Alga, Scenedesmus Dimorphous , Under a Wide Range of Nitrogen/Phosphorous Ratio-I. Setsunan, Japan: University of Setsunan, 1984. 17. Shen, Ying, Zhijian Pei, Wenqiao Yuan, and Enrong Mao. Effect of Nitrogen and Extraction Method on Algae Lipid Yield. Kansas State: University of Kansas State, 2009. 18. Xiong W, Li X F, and Wu Q Y. High-density fermentation of microalga Chlorella protothecoides in bioreactor for microbio-diesel production. Appl Microbiol Biotechnol, 2000; 27(3-5):312-318. 19. “Cell counting/hemacytometer”. Open WetWare. 14 Sept. 2008. 24 June 2009 <http://openwetware.org/wiki/Cell_counting/hemocytometer>. 20. Rouge, Melissa. “Counting Cells with a Hemacytometer”. Pathophysology of the Reproductive System. 2 Sept. 2002. 24 June 2009 <http://arbl.cvmbs.colostate.edu/hbooks/pathphys/reprod/semeneval/hemacytometer.h tml>. 21. Hansen, P.J. “Use of a Hemacytometer”. Department of Animal Science, University of Florida. 30 Oct. 2001. 24 June 2009 <http://www.animal.ufl.edu/hansen/protocols/hemacytometer.htm>. 22. Sieg, David. Making Algae Biodiesel at Home, 2008. 23. “About Coagulation and Flocculation”. Water Specialties Technologies. 2009. 10 July 2009 <http://www.waterspecialists.biz/html/about_coagulation___flocculati.html>. 24. “Aluminum Sulfate”. Wikipedia the free encyclopedia. 20 June 2009. 10 July 2009 <http://en.wikipedia.org/wiki/Aluminium_sulfate>. 25. “Algae Oil Extraction”. Oilgae. 2006. 6 July 2009 <http://www.oilgae.com/algae/oil/extract/extract.html>. 37

26. Dr. Leray, Claude. “Lipid Extraction”. CyberLipid Center. 13 June 2009. 7 July 2009 <http://www.cyberlipid.org/extract/extr0001.htm>. 27. Tzann, Stelios T. “Non Mechanical Methods”. Tutorial on Cell Disruption. 3 June 1996. 7 July 2009 < http://128.113.2.9/dept/chem-eng/Biotech Environ/DOWNSTREAM/disrupt.htm>. 28. “Hexane”. Wikipedia the free encyclopedia. 15 June 2009. 8 July 2009 <http://en.wikipedia.org/wiki/Hexane>. 29. “Isopropyl Alchohol”. Wikipedia the free encyclopedia. 29 June 2009. 8 July 2009 <http://en.wikipedia.org/wiki/Isopropyl_alcohol>. 30. “Biodiesel Production”. What is Biodiesel? 10 July 2009 <http://www.esru.strath.ac.uk/EandE/Web_sites/0203/biofuels/what_biodiesel.htm#bi o_production>. 38

Appendix 39

Hemacytometer Procedure: 1. Clean the surface of the hemacytometer and coverslip with 70% ethanol. 2. Place the coverslip over the counting platform, pressing on the elevated ridges of the hemacytometer, not the center. (Some place a small amount of water on the ground glass area to improve the seal). 3. Mix the cells thoroughly to disperse clumps and produce a uniform suspension. 4. Either use directly the cell suspension or mix with a staining solution. Staining of cells often facilitates visualization and counting. Either mix cells with an equal volume of trypan blue [0.4% (w/v) trypan blue in PBS] to determine live/dead count (dead cells are blue) or kill cells with 10% formalin and then stain with trypan blue or other stain (to improve visualization of all cells. A maximum cell count of 20 to 50 cells per 1mm2 is recommended. 5. Transfer the sample to the edge of the hemacytometer and let it be drawn under the coverslip by capillarity. Do not overfill nor underfill the chamber or it will alter the distance between the hemacytometer surface and the cover slip and thus an altered volume and calculated concentration. 6. Allow a few minutes for the cells to settle and the viability stain to work before counting. 7. View the slide at 100X magnification (10X ocular with 10X objective). The central area of the grid should occupy the center of the microscope field and cells should be evenly distributed and without any clumps. If cells are not evenly distributed or clumped, wash the hemacytometer and cover slip and start over. 40

8. Use the counter to record the number of cells in the grid. Most often the center field and four corners are counted but certain types of cells/applications use more or less of the 1mm2 squares as needed for the desired precision. The more squares and cells, the better the precision but the more time consuming. 9. Count the cells touching the middle line of the triple line on the top and left of the grid but do not count the cells touching the line on the bottom or right 41

Colorimetric determination of Urea To be able to compare the consumption of nitrogen to estimate the better period for the nutrient depravation time, we need to know the nutrient consumption rate. That is why we use a method for the determination of the urea concentration. A. Method a. Reagents i. Sodium tungstate, 10% w/v 1. Dissolve 10gm of sodium tungstate, C.P., in distilled water and dilute to 100ml. Neutralize the solution with 1N sulfuric acid using phenolphthalein as indicator. ii. Sulfuric acid N/12 1. Prepared by mixing 2.32cc. of conc. H2SO4 C.P., with distilled water and diluting to 1000ml. Normality should be checked. iii. Tungstic acid 1. Mix just before use, 9 parts of N/12 sulfuric acid (2) with 1 part of sodium tungstate 10% (1). iv. Phospohoric acid 60% v/v 1. Mix 60 cc. of phosphoric acid, C.P., (85.0 – 87.0 %) with distilled water and dilute 100ml. This solution is stable indefinitely. v. Diacetyl monixime-thiosemicarbaside (DAM-TSC) 42

1. Dissolve 0.6 gm of DAM (2,3, butanedione -2- monoxime, C.P.) and 0.03 gm of TSC, C.P., with distilled water and dilute to 100ml. this solution is stable at room temperature. A pale yellow color which appears after few days does not interfere with the reaction. vi. Reagent A 1. Just before use, mix 10 parts of phosphoric acid 60% (4) and 2 parts of DAM-TSC (5). This solution is not stable for more than 1 hr. vii. Urea standard solution, 150mg /100ml 1. Dissolve 150mg of urea, C.P. with distilled water and dilute to 100 ml. Add 1 drop of preservative (benkalkonium chloride 12.8 %) per ml is the solution is to be kept. Prepare various diluted solutions from the stosk standard and use like blood specimen to calibrate the spectrophotometer. b. Procedure i. Algae 1. Collect 0.1 cc. of algae with 0.1 cc. pipet and add immediately to a small centrifuge tube containing 0.9 cc. of tungstic acid (Reagent iii). After 5 minutes centrifuge and collect 0.2ml. of supernatant liquid with 43

0.2-ml. pipet, add to 5ml. of Reagent A (No.vi) in a test tube. After vigorous shaking, heat in a boiling water bath for 20 min. + 1 min. Remove immediately and cool in running water to room temperature. Read immediately in a spectrophotometer at 530 mµ or in a colorimeter using a yellow-green filter, adjusting the 100% T with a blank made with Reagent A. 44

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Maximization Of Scenedesmus Dimorphus Lipid Yield For The Production Of Biodiesel

by Carlos Ramos

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Maximization Of Scenedesmus Dimorphus Lipid Yield For The Production Of Biodiesel Document Transcript