Effects of salinity on the growth and lipid production of ten species of microalgae from the Swartkops saltworks : a biodiesel perspective

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Effects of Salinity on the Growth and Lipid Production of

Ten Species of Microalgae from the Swartkops Saltworks:

A Biodiesel Perspective

By

Martinus Jakobus Sonnekus

Submitted in fulfilment of the requirements for the degree, Magister Scientiae, in the Faculty of Science at the Nelson Mandela Metropolitan

University

January 2010

Supervisor: Dr. D.R. du Preez Co-supervisor: Prof. E.E. Campbell

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Acknowledgements

Many people and organizations have provided advice, finance and support throughout the period of study and I am indebted to all of them. A big thank you to my supervisors, Dr. Derek du Preez and Prof. Eileen Campbell, who have gone the extra mile for me; I have and will continue to learn from you. Research and bursary funds were provided by NMMU and the Swartkops Saltworks.

Thanks to all of the staff and students in the Botany Department at NMMU and Swartkops Saltworks who assisted me in the field and in the laboratory; I would still be counting cells without your help. To my diatom granny Mrs Pat Smailes, thank you for the assistance and an endless mentoring. Your passion for diatoms has definitely inspired me.

To my family and friends, without your support and encouragement I would still be climbing this mountain. A huge thank you goes to my parents, who supported me through the hard times with much love and prayer. I owe you... To Merika, thank you for your continued support in the form of packed lunches, laughter, games, and keeping me company when late nights called.

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List of Tables

Table 2.1. Oil content (%) of various microalgae (Chisti, 2007) ... 31 Table 3.1 Productivity and specific growth rate for six Bacillariophyceae

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7 different salinities (data shown as mean ±1 SD, n = 5, except where

indicated). ... 54 Table 3.2. Productivity and specific growth rate of three species of warm-temperate solar saltworks microalgae belonging to the Chlorophyta , grown at four different salinities (data shown as mean ±1 SD, n = 5, except where indicated). ... 55 Table 4.1 Total lipid content (% DW) for ten species of microalgae isolated from a warm-temperate solar saltworks and grown at four different salinities (data shown as mean ± SD, n = 5, except where indicated). ... 72 Table 5.1. Estimated biomass and oil production for two microalgal species based on current results and general pond features and estimations. ... 82

List of Figures

Figure 3.1. The layout of the Swartkops Saltworks. ... 42 Figure 3.2. Cell density over time at 17 and 35 psu for Nitzschia hungarica. Bars = +1 S.D. ... 46 Figure 3.3. Cell density over time at 17, 35, 52 and 70 psu for Amphora

coffeaeformis var. coffeaeformis. Bars = +1 S.D. ... 47 Figure 3.4. Cell density over time at 17, 35, 52 and 70 psu for Navicula

gregaria. Bars = +1 S.D. ... 48 Figure 3.5. Cell density over time at 17, 35, 52 and 70 psu for Navicula

salinicola. Bars = +1 S.D. ... 49 Figure 3.6. Cell density over time at 17, 35, 52 and 70 psu for Navicula sp. Bars = +1 S.D. ... 50 Figure 3.7. Cell density over time at 17, 35, 52 and 70 psu for Nitzschia

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8 Figure 3.8. Cell density over time at 17, 35, 52 and 70 psu for Thalassiosira sp. Bars = +1 S.D. ... 52 Figure 3.9. Cell density over time at 17, 35, 52 and 70 psu for

Chlamydomonas sp. Bars = +1 S.D. ... 53 Figure 3.10. Cell density over time at 17, 35, 52 and 70 psu for Tetraselmis sp. Bars = +1 S.D. ... 56 Figure 3.11. Cell density over time at 17, 35, 52 and 70 psu for Dunaliella

tertiolecta. Bars = +1 S.D. ... 57 Figure 3.12. Principal Components Analysis (PCA) of species, for specific growth rate, separated on the major ordination axis (x-axis) by specific growth rate and on the y-axis by salinity, (x-axis eigenvalue = 0.948, y-axis

eigenvalue = 0.028. Cumulative percentage variance = 94.8 %, Sum of all eigenvalues = 1.000). ... 59 Figure 3.13. Principal Components Analysis (PCA) of salinities, for specific growth rate, separated on the major ordination axis (x-axis) by specific growth rate and on the y-axis by salinity, (x-axis eigenvalue = 0.948, y-axis

eigenvalue = 0.028. Cumulative percentage variance = 94.8 %, Sum of all eigenvalues = 1.000). ... 60 Figure 4.1 Principal Components Analysis (PCA) of species, for total lipid content (x-axis eigenvalue = 0.792, y-axis eigenvalue = 0.119. Cumulative percentage variance of species data = 91.1. Sum of all eigenvalues = 1.000). ... 74 Figure 4.2 Principal Components Analysis (PCA) of samples indicates the total lipid content with regard to salinity, separated on the major axis (x-axis,) by total amount (% DW) of lipid and on the y-axis by means of salinity (x-axis eigenvalue = 0.792, y-axis eigenvalue = 0.119. Cumulative percentage

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Abstract

Biodiesel from microalgae is a viable alternative for replacing the global demand for petro-diesel. High biomass and lipid production are key desirable characteristics needed in a species to be used for biodiesel production. It has been demonstrated in literature that the increase in salinity can increase the lipid content of microalgae, but lower the growth rate of a species. Therefore the effect that salinity has on the growth and lipid content of ten microalgal species, isolated from a warm temperate solar saltworks, was investigated. The microalgae were cultivated at a temperature of 22°C and at salinities ranging from 17 to 70 psu. It was found that growth and lipid production for all species were influenced to some degree by the salinity. Growth rates greater than 0.6 d-1 showed a decrease with higher salinity. Most (71%) of the growth rates that exceeded 0.6 per day were exhibited by cultures exposed to normal salinity (35 psu). This shift is a good indication that salinity inhibits/slows down growth and that the species in general prefer lower salinity conditions. Growth rates ranged from 0.17 ± 0.05 to 1.19 ± 0.17 d-1. Lipid content for the diatoms (2.78 ± 0.36 to 10.86 ± 4.59 % DW) were lower than expected, whereas the lipid content for the green flagellates (3.10 ± 1.56 to 22.64 ± 1.19 % DW) was on par with that reported in literature. To bring results into perspective a production model was developed to simulate a production scenario at the Swartkops Saltworks. Lipid and productivity results obtained in this study were used to estimate how much oil and biomass can be produced within the ponds of the Swartkops Saltworks. The model showed that although microalgae cultivation for biodiesel is technically feasible, at present it is not economically viable to do so.

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Chapter 1 - Introduction

Biodiesel as an alternative to petrodiesel is receiving much attention globally (Sheehan et al., 1998, Miao & Wu, 2006, Chisti, 2007, Duffield, 2007, Chisti, 2008, Demirbas, 2008, Ratledge & Cohen, 2008, Schenk et al., 2008, Song et

al., 2008, Griffiths & Harrison, 2009) and locally (Griffiths & Harrison, 2009). The production of biodiesel is not new as the technology has been around for a few decades (Bartholomew, 1981, Chisti, 2007) with most of the current biodiesel technology based on conventional oil bearing crops such as soybean and sunflower (Chisti, 2007, 2008). The use of conventional oil crops for biodiesel production rather than food caused a global rise in food prices. Global concern over the rise in food prices (Chisti, 2008, Griffiths & Harrison, 2009), the depletion of crude oil reserves, high oil prices and global warming have sparked a search for new, clean, economically viable carbon neutral fuel (Rodolfi et al., 2009). In the search for alternative fuel sources microalgae have shown promise as an alternative (Chisti, 2007). Microalgae have a number of advantages over conventional oil bearing crops in that they grow relatively fast, doubling in biomass every few hours to a few days. The amount of area needed for microalgal cultivation to meet the global biodiesel demand is a small fraction compared to that needed for conventional crops (Chisti, 2007). Although there are numerous advantages to microalgal biodiesel production, the technology and our knowledge are limited.

Research into the use of microalgae started in earnest in the United States in the late 1970’s with the Aquatic Species Program (Sheehan et al., 1998). The Aquatic Species Program was however terminated in 1996 with no recommended biodiesel microalgae. The reason for terminating the Aquatic Species Program was the drop in crude oil prices and thus the need for alternative fuel subsided. The realisation of global warming and the associated global climate change put a renewed haste on the development of clean alternative fuel. The turn of the decade saw a substantial increase in microalgae biodiesel research output when compared to previous decades.

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11 Today microalgae biodiesel projects are in place all over the world, with Germany and Australia being the major contributors. There are however mixed reactions over the feasibility of microalgae for biodiesel production, most authors have a positive outlook (Chisti, 2007, 2008, Schenk et al., 2008, Song et al., 2008) and some are outright pessimistic (Ratledge & Cohen, 2008).

To meet the global demand for biodiesel, large scale cultivation of biomass is needed. Intensive cultivation of microalgae by means of photo-bioreactors will be an expensive exercise whereas extensive cultivation of microalgae using open ponds will be more economical. Solar saltworks, such as the Swartkops Saltworks, uses large open ponds to concentrate salt; these ponds would also be ideal for the cultivation of microalgae and will result in additional income for the saltworks. The Swartkops Saltworks, located outside Port Elizabeth can benefit from such dual functionality. Agricultural and industrial activities within the Swartkops River catchment have resulted in high nutrient concentrations within the estuary (Emmerson, 1985, Scharler & Baird, 2003) from which the Swartkops Saltworks draws its saltwater. These nutrients are detrimental to salt production. However, high nutrient concentrations can be beneficial for cultivation of microalgae and therefore reduce the associated cost of adding additional nutrients. The high nutrient concentration promotes the growth of microalgae, macroalgae and cyanobacteria and leads to increases in organic matter and detritus. This increase in organic matter within the water column is detrimental to salt production (Du Preez, pers comm.). Organic matter combines with the salt crystals in the crystallizer ponds producing a dirty and dark looking salt crystal. Extra production steps are needed to remove the organic matter from the salt. The salt is placed in a furnace to burn off the organic matter and is then washed with clean brine (Du Preez, pers comm). The Swartkops Saltworks can therefore benefit from the following scenario: The cultivation of microalgae within the first three ponds (see Figure 3.1) will absorb the nutrients within the water column and therefore reduce the nutrient load within the saltworks. This in turn will reduce the associated organic matter within the saltworks and this will yield higher quality salt. In order to

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12 achieve this, a very effective harvesting mechanism is needed. Any microalgal cells not removed will result in an increased organic load within the saltworks. Before commencing large scale cultivation, certain requirements need to be met. One such requirement is a good understanding of the biology of the selected species of microalgae.

The present study aims to serve as a baseline study on the growth and lipid productivity of ten species of microalgae isolated from the Swartkops Saltworks. Seven of these species are from the division Heterokontophyta and three from the division Chlorophyta. The effect that salinity has on the growth rate and lipid productivity of ten microalgae species was investigated.

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Chapter 2 - Literature Review

Marine Culture Media

Natural seawater has a highly variable chemical composition (Gagneux-Moreaux et al., 2007), containing more than 50 known elements and a large number of organic compounds. The direct use of natural seawater without addition of nutrients, trace elements and vitamins for algal cultures is seldom recommended (Harrison & Berges, 2005, Gagneux-Moreaux et al., 2007). Marine culture media can be divided into two groups; natural (NW) and artificial seawater (AW). Both natural and artificial seawater are unenriched media and share similar problems with regard to culture requirements; for example macronutrients, such as nitrogen and phosphorus, or trace metals, may be limiting (Harrison & Berges, 2005). This is rectified by adding enrichment solutions that provide for substantial algal yields.

Macronutrients

Nitrogen, phosphorus and silicon are generally considered to be macronutrients. Silicon is only a prerequisite for diatoms, silicoflagellates, and a few chrysophytes. Diatoms generally require nitrogen, silicon and phosphorus in a 16N:16Si:1P ratio (Brzezinski, 1985, Harrison & Berges, 2005). In natural seawater, the ratio of macronutrients is similar to that required by diatoms and other microalgae. The 16N:1P ratio is generally ignored in most media: they generally have a ratio >16N:1P. In such media phosphorus may be limiting (Berges et al., 2001). The same applies to silicon; the N:Si ratio is near 1:1 for most diatoms, but in most culture media the N:Si ratio is not near 1:1 (Berges et al., 2001). Carbon and its relationship to nitrogen and phosphorus is another aspect that needs to be considered when working with culture media. If the ratio of C:N:P is undersupplied then one of these elements may become limiting and inhibit cell growth. An ideal culture

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14 medium will balance these elements in the correct ratio, but this is seldom achieved.

Various chemical compounds can be used as a nutrient source and these are generally made up into working stock solutions (Harrison & Berges, 2005). Nitrate and phosphate are generally added as NaNO3 and NaHPO4.H2O, whereas ammonium and silicate may be added as NH4Cl and Na2SiO3.9H2O, respectively (Harrison & Berges, 2005). The stock solutions should be either autoclaved or filter sterilized to kill and remove bacteria and fungi (Harrison & Berges, 2005).

Trace Metals

Trace metals and vitamins are required in small quantities (Harrison & Berges, 2005). This poses a problem when they are to be weighed and high concentration stock solutions are first prepared to facilitate the weighing of vitamins and trace metals. The desired volume is then added to the appropriate recipe. Care must be taken when storing these stock solutions for a period of time as evaporation and precipitation may occur (Harrison & Berges, 2005).

Stock solutions of chloride or sulphate salts of cobalt, zinc, manganese, selenium and nickel are made up using the chelator, EDTA, added to keep trace metals in solution. Iron can be added in the form of ferric chloride, ferrous sulphate and ferrous ammonium sulphate. In a separate stock solution, the iron is chelated or kept in 10-2 M HCl to avoid precipitation. Boron is only required for artificial seawater as natural seawater contains sufficient amounts of boron (Harrison & Berges, 2005).

Vitamins

Most microalgae require vitamins: usually vitamin B12 (cyanocobalamine), B1 (thiamine) and vitamin H (biotin), but not all of them require all three

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15 (Peperzak et al., 2000, Harrison & Berges, 2005). The order of requirement is vitamin B12 > vitamin B1 and vitamin H (Provasoli, 1958, Harrison & Berges, 2005). As with macronutrients and trace metals, vitamins may also become limiting and must be added to the culture media. The vitamins required by microalgae vary between species and it is possible to omit any of the vitamins from the culture media if the target species do not require that specific vitamin (Harrison & Berges, 2005). Vitamins are heat sensitive and therefore should be filter sterilized (Harrison & Berges, 2005). Stock solutions may be frozen, thawed and refrozen for extended periods of time.

Bertrand et al. (2007) demonstrated the importance of vitamins and iron to growth of phytoplankton species. Peperzak et al. (2000) reported that the addition of vitamins B12, B1 and H to Phaeocystis globosa cultures significantly enhanced the biomass yield of this flagellate.

pH Buffers and Chelators

Tris (2-amino-2-[hydroxymethyl]-1-3-propanediol) and glycylglycine are two common pH buffers used in marine culture media (Harrison & Berges, 2005). The main purpose of pH buffers is to prevent or reduce precipitation. It is noted that some pH buffers, e.g. Tris, are toxic to certain species, whereas glycylglycine is non-toxic to all (Harrison & Berges, 2005). Small quantities of buffering agent (1-5 mM) are added to the culture medium.

EDTA is a chelator commonly used to keep iron in solution (Harrison & Berges, 2005) and to alleviate the toxic effects of Cu and Cd (Muggli & Harrison, 1996). EDTA inhibits growth of some oceanic species and care must be taken, especially on recently isolated species (Muggli & Harrison, 1996).

Natural Seawater Enrichment Media

Both broad spectrum and species specific natural seawater enrichment media exist. The most cited broad spectrum media are f/2 medium, different versions

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16 of Erdschreiber medium and ESNW (Berges et al., 2001, Harrison & Berges, 2005). Some media are especially developed for cyanobacteria and these are less commonly used for oceanic species, e.g. BG medium (Rippka et al., 1979, Harrison & Berges, 2005).

Microalgae Isolation Techniques

The traditional means of microalgae isolation was established by the late 1800’s (Andersen & Kawachi, 2005). Some species are easy to isolate and cultivate, while others seem impossible to get into cultivation. For successful isolation and culturing the natural conditions of the microalgae under investigation must be mimicked (Andersen & Kawachi, 2005). Salinity and temperature are important for coastal species, whereas for open ocean species water quality is more important. After this initial step towards successful isolation, the second step is the elimination of contaminants. Techniques of dilution, single-cell isolation by means of micropipetting, and agar streaking are some of the more common and traditional methods used (Andersen & Kawachi, 2005).

The growth of the isolated species is the final step in successful isolation. Some species start well in the initial stages of isolation, but after a series of transfers to fresh media the target species die off. This die-off is an indication that the culture medium lacks a particular element or organic compound that is found in the microalga’s natural environment, but is absent from the culture medium (Andersen & Kawachi, 2005). Build up of metabolic waste may also lead to death and this needs to be avoided (Andersen & Kawachi, 2005).

Sample collection

The proper collection of microalgae is of vital importance because dead and damaged cells will result in failure. The method of sampling varies from species to species, for example Synura petersenii can be concentrated using

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17 a phytoplankton net and kept cool on ice for over 24 hours (Andersen & Kawachi, 2005). However, using the same method for Gonyostomum semen does not result in the same success because when G. semen cells come into contact with each other, a mucocyst is discharged and many cells die (Andersen & Kawachi, 2005).

The two methods used for sample collection are whole water (not concentrated) and concentrated samples (Andersen & Kawachi, 2005). Whole water sampling often results in the most viable cells (Andersen & Kawachi, 2005). However, it is time consuming to find target species in dilute samples. If prior knowledge of the sampling area is lacking then it is recommended to use different sampling methods on a trial and error basis (Andersen & Kawachi, 2005).

Samples should be filtered to remove unwanted organisms such as zooplankton, colonial algae or other unwanted algae. Care must be taken to avoid damage to or desiccation of the target species (Andersen & Kawachi, 2005).

Another aspect to consider is post-collection treatment of samples (Andersen & Kawachi, 2005). Some species die within a short time after collection, while others multiply rapidly for a day or two and then die. Yet others only multiply rapidly after a few months in culture. Time of sampling can also influence the success of microalgae isolation. If the natural population is in a good state of health then isolating and maintaining growth is easy, whereas when the population is in a poor state of health problems with isolation may occur.

Enrichment cultures

Enrichment cultures are used as an initial step towards obtaining single cell isolates (Andersen & Kawachi, 2005). In enrichment cultures, nutrients are added to a natural sample, enabling rapid cell growth. Enrichment sources may be culture media, soil extract, macronutrients (ammonium, nitrate and phosphate) and trace metals. If soil of good quality can be obtained, then soil

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18 extract is a simple and effective method (Andersen & Kawachi, 2005). However, if bacterial numbers are high the enrichment culture may become anoxic and toxic, leading to algal cell death. Additional selective factors that can be used in conjunction with the enrichment solutions to isolate microalgae are pH, temperature, light, salinity, and specific physical conditions (e.g. sediment environment).

Natural samples are often deficient in one or more nutrients and in order to overcome these shortages small amounts of culture or nutrients solutions are added. However, additional nutrients may be detrimental to ‘non-weedy’ species as ‘weedy’ species may reduce target species by outcompeting them. In such cases unenriched solutions are preferred. Enrichment is important and necessary for r-selected species, whereas for k-selected species, minimally, or unenriched media are more appropriate (Andersen & Kawachi, 2005).

Single-cell isolation by micropipette

The goal of micropipette isolation is to pick up a single cell from the sample, and transfer the cell through a series of sterile droplets, thereby ridding the target cell of unwanted contaminants before culturing. An advantage of this technique is that a axenic monoculture is more easily achieved, but a disadvantage is that excessive handling may damage the cell (Andersen & Kawachi, 2005).

Pasteur pipettes or glass capillaries are used in micropipette isolation (Andersen & Kawachi, 2005). A mouthpiece is attached to one end of the tube and the other is loaded with a sterile drop of water, which serves as a buffer. The operator places his/her tongue over the mouth piece; the micropipette is placed next to the target cell. The tongue is then removed from the mouth piece, which causes a capillary action that draws the cell up into the pipette. Ejection of the cell from the micropipette is done with a gentle blow.

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Isolation with the use of agar

The use of agar is an old and commonly used method for isolation of microalgae (Andersen & Kawachi, 2005). For successful isolation onto agar, the algae must be able to grow on this medium. Not all microalgae grow well on agar. Genera that do not grow at all on agar are: Heterosigma,

Pelagomonas and Peridinium, whereas genera such as Chlamydomonas,

Pavlova, Synura, and Tetraselmis grow very well on agar (Andersen & Kawachi, 2005). Coccoid cells, most diatoms and chlorachniophytes grow very well on agar, while dinoflagellates rarely grow on agar. The concentration of the agar is not an important factor although between 0.8% and 1.5% is generally used (Andersen & Kawachi, 2005).

Unfortunately, agar is also a good medium for bacterial and fungal growth (Andersen & Kawachi, 2005). Care must be taken to avoid fungal growth on the plates, as they grow and reproduce quickly. If aseptically streaked, microalgal colonies will be formed, which can be picked up and restreaked on fresh agar or transferred to a dilute culture media. A bacterial loop is loaded with a small amount of sample, and then the sample is spread across the agar using the loop. The origin of the streak contains a high number of cells and at some stage along the streak, the cells are spread out sufficiently to deliver single cells. After streaking, the agar plate is incubated under favourable conditions until colonies of cells appear. Isolated colonies are then transferred to new agar or introduced to a suitable culture medium (Andersen & Kawachi, 2005).

Agar pour plates

Not all algae grow on the surface of agar but will grow when embedded in the agar. For example, the cyanobacterial genus Synecococcus was successfully isolated using the agar pour method (Brahamsha, 1996, Toledo & Palenik, 1997). Sample or enrichment cultures are mixed with non-solidified agar, and then poured into agar plates where the agar solidifies. Care must be taken when pouring the sample or enrichment culture into the agar; the agar must

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20 not be so hot as to kill the microalgae. The agar is then incubated and colonies are transferred to fresh agar or introduced to the desired culture medium.

Dilution Techniques

The dilution method was first successfully used by Allen and Nelson (1910). Butcher (1952), Gross (1937) and Parke (1949) also successfully isolated species of the marine phytoplankton in this way. With the dilution method the goal is to deposit a single cell into a culture chamber thereby establishing a monoculture. A drawback of the dilution method is that axenic cultures are less likely to be obtained as bacterial numbers are far greater than microalgal numbers (Droop, 1954, Andersen & Kawachi, 2005).

Culturing of Microalgae in Outdoor Ponds

For microalgal products to be economically viable large quantities of biomass must be produced. This requires large volumes of culture media and in order to achieve this most commercial scale culture is done in outdoor ponds (Richmond, 1999, Tredici, 2004). Several species are currently being grown in outdoor ponds: Chlorella spp., Spirulina platensis, Spirulina maxima,

Dunaliella salina, Haematococcus pluvialis and Nannochloropsis spp. On a smaller scale, examples are Porphyridium spp. and Scenedesmus obliquus that have been successfully used (Borowitzka, 2005).

An advantage of using open ponds is that they are cheaper to construct and operate compared to photobioreactors. Disadvantages are that open pond culture does not work for all species (Borowitzka, 1999), they are also easily contaminated by unwanted species and are restricted to a few geographical areas (low rainfall and cloud cover). Successful cultivation requires a good understanding of the physiology and ecology of the specific species of interest so that the setup meets the specific requirements of the target species (Borowitzka, 2005).

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Types of open ponds

A few basic criteria must be kept in mind when designing an open pond, for example construction and maintenance costs should be kept to a minimum while at the same time providing optimal conditions for growth. Borowitzka (1999, 2005) recommends different types of open pond systems. The four basic types are shallow big ponds, tanks, circular ponds and raceways.

Very large, shallow, unlined and unmixed ponds are unsuitable for most species of microalgae (Borowitzka, 2005). These types of system are referred to as “natural ponds” (Tredici, 2004). “Natural ponds” are extensively used in Australia for culturing of Dunaliella salina (Borowitzka, 1999, Tredici, 2004, Borowitzka, 2005). Pond areas range from 1 ha to more than 200 ha with an average depth of 20 to 30 cm. These large open ponds are prone to contamination (Borowitzka, 2005) and productivity is low (generally not exceeding 1 g m-2 d-1;Tredici, 2004).

For small-scale production of microalgae, deep ponds or tanks may be used (Borowitzka, 2005). Mixing of the water column is done with aeration. Deep ponds are generally smaller than 10 m2 and the depth is greater than 0.5 m. Although very inefficient these systems are easy and cheap to operate.

Circular ponds, similar to those found at wastewater treatment works, are used in Japan and Taiwan for cultivating Chlorella spp. (Borowitzka, 2005), but these are not recommended for commercial cultivation (Tredici, 2004). The rotating mixing arm limits the size of these to about 10 000 m2, because even mixing of the water column is not possible in larger ponds (Borowitzka, 2005).

Single, rectangular ponds fitted with a paddle (raceways) are commonly used for the production of Spirulina spp., Dunaliella salina and Haematococcus

pluvialis. The paddle wheel is the driving force for water circulation. This design appears to be the most efficient, and ponds may be up to 1 ha in size (Borowitzka, 2005). Microalgal productivity reported for raceways is as high as

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22 40 g m-2 d-1 but typically 12-13 g m-2 d-1 is achieved (Tredici, 2004). The principles for design and construction of large raceway ponds were reviewed by Oswald (1988).

Selecting pond types

A consideration when selecting pond type is to use several kinds of pond, rather than only one to minimize losses in case of culture problems (Borowitzka, 2005). Small ponds are more expensive per unit area than larger ponds. The size of the pond affects water circulation and this in turn affects the design and operating cost (Borowitzka, 2005).

Climatic conditions also need to be considered. If the climate only allows for production to occur for part of the year, then more intensive culture methods should be considered. The price of land may also influence whether intensive or extensive systems are going to be used, low land prices will favour more extensive culturing (Borowitzka, 2005).

Strain selection

A productive strain should be selected for the specific conditions of the pond. General characteristics of strains are growth rate, biochemical composition, temperature tolerance, and resistance to mechanical and physiological stress. The selection process starts within the laboratory, with subsequent testing in open ponds for optimal growth conditions. The pond conditions are then manipulated in such a way as to achieve the highest production (Borowitzka, 2005).

Scale-up

Scaling cultures up to the volumes required for commercial production is one of the most difficult tasks in outdoor microalgal mass culture (Borowitzka,

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23 2005). The process is prone to contamination from other algae and bacteria due to the dilute inoculum.

There are two methods for scaling cultures up to production volumes. The first is to take the laboratory culture through a series of dilutions until the volumes of the production pond are achieved. In the second method the inoculum is derived from existing culture ponds. The latter method is preferred because it is quicker compared to the up-scaling from a 20 ml flask to 10 000 m3 raceway – this may take 8 to 9 weeks (Borowitzka, 2005).

Culture Medium

The same culture medium that was used in the laboratory is used in large scale culture setups with slight modification (Borowitzka, 2005). The choice of culture medium depends on several factors: the growth requirements of the algae, how the medium affects the final product quality and the cost. If high quality algae (health food and nutraceutical applications) are required then high-grade chemicals must be used. For lower quality algae animal feed and cheaper industrial grade chemicals can be used (Borowitzka, 2005).

Pond management

Outdoor algal ponds are exposed to a variety of factors: climatic/environmental factors and contamination (Borowitzka, 1999, Borowitzka, 2005). Light and temperature show diel and seasonal variation. The difference in night and day temperature can significantly influence production; large variations can lead to photoinhibitory stress and the resultant loss of production (Lu & Vonshak, 1999, Masojídek et al., 1999). Seasonal variation in temperature and light may also inhibit algal growth and hence algae farms are most suited for subtropical areas (Borowitzka, 2005).

Competition for nutrients and space can also play a vital role in the productivity of a pond. The higher the cell concentration the greater the

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24 competition and for this reason keeping the cells at an optimal concentration will benefit production. This concentration is in the log growth phase where maximum growth rates are obtained (Borowitzka, 2005).

It is impossible to prevent contamination in open ponds. Algal cultures generally have contaminants such as bacteria, viruses, fungi, other algae, zooplankton, insects, leaves and other airborne material. It is necessary to control these contaminants to acceptable levels. Contaminants may be removed by placing suitable screens in the water flow, which trap them (Borowitzka, 2005).

Culture monitoring

To be successful, cultures should be monitored constantly (Borowitzka, 2005). The easiest method is to microscopically examine a sample to observe any morphological changes in the algae as well as the presence of unwanted organisms. Routine testing of nutrient levels is also important for successful production. Temperature, O2-concentration and pH should be checked on a daily basis (Borowitzka, 2005). New techniques, such as pulse amplitude modulated fluorometry (PAM) can be used to monitor the physiological state of microalgae cultures for any signs of deterioration (Torzillo et al., 1998, Lippemeier et al., 2001).

The production of microalgae in large open ponds is economically feasible for a limited number of species. A good knowledge of species requirements and management experience are essential for successful culturing (Borowitzka, 2005).

Photobioreactors

The cultivation of microalgae in open ponds has been well researched and developed, however only a few species can be successfully cultured in this way (Molina Grima et al., 2001). Use of photobioreactors has increased the

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25 number of species available for culture compared to open ponds and raceways (Molina Grima et al., 2001, Tredici, 2004, Chisti, 2007). A photobioreactor permits monoculture as the system is closed and so reduces the chance of external contamination (Molina Grima et al., 2001, Chisti, 2007). Small to large scale microalgal production is possible using highly controlled photobioreactors. Energy is supplied by natural and/or artificial light. The main advantage of photobioreactors is that the production method may be cost effective and consistent in quality from batch to batch by controlling and optimizing of the culture parameters (Behrens, 2005). Not all species are suited for use in photobioreactors, delicate and fragile species can be damaged by the circulation system and others may adhere to the surface of the bioreactor (Borowitzka, 1999, Chisti, 2007), Construction, maintenance and operation costs are high (Borowitzka, 1999, Chisti, 2007). The designs of photobioreactors differ, depending on the ultimate goal, which may be large quantities of algae of a low value or small quantities of a high value (Behrens, 2005).

Reactor design

Light is the most crucial design consideration for a photobioreactor (Behrens, 2005). High surface-to-volume ratio allows for adequate light penetration, which in turn enhances photosynthetic efficiency. Other factors to be considered in the design process are circulation, inorganic carbon supply, oxygen removal, and pH and temperature control. Circulation is important for optimum illumination of the algae, adequate gas exchange and temperature and pH control. The choice of materials for construction of the photobioreactor is important as well. Glass and acrylic are widely used, especially UV-stabilized acrylic as it is light and durable. Various shapes and sizes have been developed to date and these can be grouped into three basic categories: tubular, flat and fermenter-type. The tubular and flat reactors are designed to harvest natural light, while fermenter-reactors require artificial illumination, heterotrophic organisms and a carbon source, e.g. sugar, is used as the

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26 primary energy source (Borowitzka, 1999, Behrens, 2005, Carvalho et al., 2006). The most popular are the tubular and flat reactor types (Borowitzka, 1999) as no expensive illumination is needed. Several layouts have been used with success (Tredici, 2004). A basic system consists of a light harvesting unit and a gas exchange unit. The light harvesting unit employs small diameter tubing to increase the surface-to-volume ratio, while in the gas exchange unit CO2 is supplied to the growth medium. A pump or airlift is used to circulate the culture between the two units. However, care must be taken when a pump is used as shear force can damage cells (Garcia Camacho et

al., 1999, Molina Grima et al., 2001, Tredici, 2004, Carvalho et al., 2006, Chisti, 2007).

Tubular reactors

Four basic types of tubular reactors are used: vertical, horizontal, helical and α-shape reactors (Lee et al., 1995, Tredici, 2004, Carvalho et al., 2006). The name of the basic types is derived from the angle of the tubes with respect to the horizontal. The α-shape is so-call as it resembles the α-sign in side view (Lee et al., 1995).

In vertical tubular reactors, air is pumped in at the bottom and allowed to rise up the tubes. The rising of the air allows for mixing of the culture as well as the removal of excess O2 and supply of CO2. Air lift and bubble reactors are examples of vertical tubular reactors. Polyethylene bags make for an inexpensive vertical bioreactor, as they are cheap, sterile and highly transparent. Frequently employed by the aquaculture industry, they are capable of reaching cell concentrations three times that of open ponds (Tredici, 2004, Carvalho et al., 2006). Productivity ranges from 0.06 to 0.5 g L -1

d-1 for Haematococcus pluvialis and Phaeodactylum tricornutum, respectively (Alías et al., 2004, Lopez et al., 2006).

Horizontal tubular reactors differ from vertical reactors in the following ways: greater surface-to-volume ratio, the amount of gas in dispersion, the gas-liquid

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27 mass transfer characteristics, how the fluid moves through the system and the amount of internal irradiance received (Sánchez Mirón et al., 1999). Horizontal tubular reactors are gaining popularity with growers as they are capable of handling large culture volumes (Tredici, 2004, Carvalho et al., 2006). A drawback of such a system is that temperature fluctuates between day and night, high culture temperatures can be obtained and this may damage cells (Tredici, 2004, Carvalho et al., 2006, Chisti, 2007). Expensive cooling systems must be employed to cool down the culture.

The horizontal tubes are best placed in a north-south orientation to maximise sunlight capture (Grima et al., 1999, Sánchez Mirón et al., 1999, Chisti, 2007). The most common arrangement of the horizontal tubes is that they are placed on the ground. In an attempt to save space the tubes are sometimes arranged in a fence-like structure. Other arrangements are based on small modifications of the above two designs, e.g. near-horizontal tubular reactors designed by Tredici & Chini Zittelli (1998). Large systems (up to 8000 L) have been used by various authors (Carvalho et al., 2006) and productivity in horizontal tubular reactors ranges from 0.32 to 1.5 g L-1 d-1 (Richmond et al., 1993, Grima et al., 1994, Barbosa et al., 2004).

An alternative to vertical and horizontal tubular reactors are helical tubular reactors. The basic design of a helical tubular reactor is a polyethylene pipe, coiled in an open circular framework. The coil is attached to a degassing unit where the temperature, CO2, O2 and pH are regulated. The culture is circulated through the coil using either an airlift or a mechanical pump. For

Spirulina platensis a productivity of 0.9 g L-1 d-1was achieved in a very stable 120 L bubble reactor (Tredici & Chini Zittelli, 1998). Productivity was lower in a 21 L helical tubular photobioreactor with a maximum productivity of 0.4 g L-1 d -1

(Travieso et al., 2001).

Lee et al. (1995) designed a α-shape reactor with a 300 L capacity and a footprint of 12 m 2. The design was found to have several advantages: a one-way flow of culture, high flow rates with minimum air supply and the 45° angle of the tubes allowed for more efficient light harvesting. Productivity obtained

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28 for Chlorella sp. was 2.88 g L-1 d-1 [72 g m-2 footprint d-1] (Lee et al., 1995), far greater than any other tubular reactor. The only drawback of the design was that scale up to a commercial scale is very difficult (Carvalho et al., 2006).

Flat Plate Reactors

Flat plate reactors are narrow panels that allow for a large area-to-volume ratio and consequently efficient utilization of sunlight. Although simple in design, not many are used on a commercial scale (Tredici, 2004). Richmond and Cheng-Wu (2001) developed a 1 000 L reactor, consisting of five 200 L units fitted together. Mixing and degassing occurred by allowing compressed air flow through a perforated pipe that extended the length of the reactor. A cooling system was employed to control temperature. Refrigerated water was sprayed on the top of the reactor and allowed to run down the sides to be collected in furrows and recycled. The maximum productivity obtained for a

Nannochloropsis species in such a system was 0.85 g L-1 d-1.

Alveolar panels are another type of flat plate reactor with high productivity (2.15 g L-1 d-1 for Spirulina platensis) successfully cultivated on a commercial scale (Tredici, 2004). Advantages of the flat plate reactor design are: uniform light distribution, high productivity, the bubbling of gas in the systems means the absence of mechanical pumps and associated cell damage (Carvalho et

al., 2006) and an energy efficient system (Sierra et al., 2008).

Fermenter type reactors

Fermenter type reactors have a small surface to area ratio; hence the harvesting of sunlight is inefficient. Light within fermenters is supplied via sophisticated systems of internal illumination (Carvalho et al., 2006). The main feature of fermenters is that organic carbon is supplied to the microalgae in the form of glucose or any other carbohydrates, thus making the heterotrophic cultivation of microalgae possible. The basic operation of fermenters is the

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29 same as for photobioreactors. Fermenter type reactors are beyond the scope of this project and literature review. The cheapest way to produce oil for biodiesel production is to utilize a cheap energy source and sunlight provides the cheap alterative. Fermenters are expensive to operate. For a discussion on fermenter type reactors see Banerjee et al. (2002), Behrens (2005) and Carvalho et al. (2006).

Design considerations

Light

As stated earlier, light is the most important variable in the design and construction of a photobioreactor, as light is the primary energy source for microalgae (Behrens, 2005, Carvalho et al., 2006). Light can be supplied either continually or in light-dark cycles. When light is limiting or of poor quality then microalgal growth will be limited. In an ideal setup, with the optimal nutrient concentrations, including carbon, light must be the only limiting factor for growth (Carvalho et al., 2006). For net growth to occur microalgae must receive sufficient light to exceed their light compensation point (the light intensity at which photosynthesis balances respiration). If light is insufficient then respiration will result in loss of carbon. Excessive light will lead to photoinhibition. The high extinction coefficient of chlorophyll does not allow light to travel large distances through cultures and light becomes limiting if the path-length through the culture is too great (Behrens, 2005).

As light travels from dense (e.g. water) to less dense (e.g. air) medium and vice versa light is refracted and reflected respectively. The surface of the photobioreactor should be designed to minimize reflection and refraction (Behrens, 2005).

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Gas exchange

Photosynthesis generates O2 and high levels of O2 inhibit algal growth (Behrens, 2005, Carvalho et al., 2006). Therefore excess O2 should be removed to optimise productivity. Nearly 50% of microalgal biomass consists of carbon and this element is required in large quantities (Morita et al., 2001, Carvalho et al., 2006). The simplest approach to supply the necessary carbon and remove the excess O2 is to enrich air with CO2and supply it to the culture medium. The bubbling of the enriched air will enhance the gas exchange and therefore lower the O2 levels.

Carbon dioxide is the preferred inorganic carbon source for microalgae (Grima

et al., 1999). The method of supply ranges from bubbling to hollow membrane transfers (Carvalho et al., 2006). The CO2 should make up 0.2 -5% of the total gas flow. However, care must be taken to ensure that the CO2 input does not lower the pH of the culture as added CO2acidifies the culture medium. Two methods for O2 removal are described by Behrens (2005): first, oxygen can be removed by periodic purges with nitrogen. Nitrogen purging has the added advantage of allowing the CO2to be used more efficiently and maintaining the oxygen level below that of ambient air. The second method is the use of a chemical system whereby, pure H2 is added to a catalyst and the excess O2 to form water (Behrens, 2005). Grima et al. (1999) provides an in-depth discussion on the light regime and gas transfer/exchange of photobioreactors.

Temperature

The inefficiency of photosynthetic systems to convert light into stored chemical energy leads to heat production (Morita et al., 2001). The theoretical conversion of red light into chemical energy (NADPH) is only 31% and the rest is lost as heat, so cooling of the culture is necessary (Behrens, 2005). Cooling of a photobioreactor can be achieved by means of a heat exchange system. A simple stainless steel rod that connects the culture medium to a refrigeration unit can be used as an effective temperature control method (Behrens, 2005).

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Harvesting methods

Even if algal cultures are growing at high concentrations, they are still quite dilute. The most common harvesting approaches are flocculation, filtering, sedimentation and centrifugation (Grima et al., 2004, Behrens, 2005). Flocculation is the collection of microalgal cells by the addition of a polymer that causes the cells to aggregate (Grima et al., 2004). Although flocculants enable and facilitate the concentration of algal biomass, centrifugation is still usually necessary to ensure a suitable volume reduction. Small scale recovery can easily be done in large centrifuge bottles, whereas larger scale cultures are best done with a continuous flow centrifuge (Behrens, 2005).

The potential of biodiesel from microalgae

Microalgae are a diverse group of eukaryotic cells producing a variety of commercially interesting secondary compounds such as lipids, hydrocarbons and other complex oils (Borowitzka, 1988, Sawayama et al., 1995, Sheehan

et al., 1998, Banerjee et al., 2002, Metzger & Largeau, 2005, Guschina & Harwood, 2006, Chisti, 2007), sugars and functional bioactive compounds (Song et al., 2008). Not all oils produced by microalgae are suitable for biodiesel production, but these are uncommon.

Microalgae exhibit properties that make them suitable for large-scale biodiesel production (Sheehan et al., 1998). Microalgae have the advantage of being able to grow fast. Harvesting of microalgal crops can occur more often, from 1 to 10 days, compared to conventional crops that are harvested once or twice per year (Schenk et al., 2008). Some microalgae may even double their biomass every 3.5 hours (Miao et al., 2004). Oil content in microalgae may reach 90% of dry mass if conditions are favourable (Spolaore et al., 2006), but commonly oil content ranges from 20 to 50% of dry mass (Chisti, 2007), with a composition similar to that of vegetable oil (Borowitzka, 1988). Table 1 is a breakdown of percentage oil content found in different taxa. The growth rate

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32 and the oil content of the microalgae determine how much oil is produced per day. Oil productivity from microalgae is calculated as the mass of oil produced per unit volume of culture (Chisti, 2007).

Table 2.1. Oil content (%) of various microalgae (Chisti, 2007)

Phylum Class Species

Oil content (% dry weight) Chlorophyta Chlorophyceae Botryococcus braunii 25–75 Chlorella sp. 28–32 Dunaliella primolecta 23 Nannochloris sp. 20–35 Neochloris oleoabundans 35–54 Prasinophyceae Tetraselmis sueica 15–23

Dinophyta Dinophyceae Crypthecodinium cohnii 20

Haptophyta Haptophyceae Isochrysis sp. 25–33

Heterokontophyta

Bacillariophyceae

Cylindrotheca sp. 16–37

Nitzschia sp. 45–47

Eustigmatophyceae Nannochloropsis sp. 31–68

Microalgae are also very energy efficient at converting raw and simple carbon forms into a complex high density liquid form or oil (Song et al., 2008). The energy efficiency of microalgae will also reduce the amount of fertilizer and nutrients needed, thus reducing pollution (Schenk et al., 2008). Algal culturing facilities do not need arable land and can potentially be built on marginal and non-arable land and even utilize waste water areas (Schenk et al., 2008).

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33 Using marine species for the production of biodiesel will lower the demand on freshwater reserves, since conventional crops consume vast quantities of freshwater (Sheehan et al., 1998, Schenk et al., 2008). The utilization of saline water in non-arable areas will open up economical opportunities to arid areas, for example the west coast of southern Africa.

Commercial production of microalgal biomass will only be viable when the cost of production can be lowered to be competitive with crude oil prices. Two aspects that need to be addressed are the recovery of algal biomass from the dilute medium and the extraction of oil from moist biomass (Chisti, 2008). The current method of biomass recovery is the use of large industrial centrifuges to concentrate the biomass; this biomass then needs to be dried before oil extraction can commence. Both steps are energy intensive and increase the cost of production. Improving photobioreactor design, harvesting and extraction techniques can reduce the cost of biomass production and ultimately be the solution to a sustainable (bio)fuel source.

In South Africa the current rate of diesel consumption is approximately 8 million m3 per annum (Germinshuis, 2006, Van Wyk et al., 2006) and in the United States of America (USA) 530 million m3 of transport fuel is consumed. Replacing the current demand for fuel with biodiesel, produced from oil crops, animal fat and waste cooking oil is not sufficient and alternative fuel sources are needed. The use of renewable energy sources are important to combat climate change (McKendry, 2002a). Christi (2007) uses the USA as an example. If half of the transport fuel is to be replaced with biodiesel, then the total crop area and the average oil yield per hectare for various crops may be calculated. For corn, biodiesel yield is 172 L ha-1 and if corn is to be used as a biodiesel source then 846% of the existing USA cropping area will have to be cultivated just for biodiesel. If oil palm, the crop with the highest oil content, yielding ~5 950 L of oil ha-1, is considered then 61% of the present USA cropping area will have to be committed to oil palm cultivation (Chisti, 2008). By contrast, if microalgae are used, only 1.1% to 2.5% of existing USA cropping area will be needed. The amount of oil produced by microalgae is species dependant, but if 30% and 70% oil from biomass can be obtained

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34 then the total oil yield per hectare is 58 700 L ha-1 and 136 900 L ha-1, respectively. This scenario illustrates that microalgae are the only “crop” that can produce sufficient quantities of biodiesel using small areas. Microalgal biodiesel is the only biodiesel that has the potential to completely replace conventional transport fuel (Chisti, 2008).

Oil extraction from microalgae

Pyrolysis is the conversion of biomass to liquid, solid and gaseous fractions by heating in the absence of oxygen (McKendry, 2002b). This produces fuels with high fuel-to-feed ratio (Demirbas, 2002). The high fuel-to-feed ratio makes it the most efficient process for biomass conversion as well as the most capable method for competing with and eventually replacing non-renewable fossil fuel resources (Demirbas, 2002). Previous studies showed that microalgal oil can be extracted from Chlorella protothecoides and

Spirulina platensis using pyrolysis (Peng et al., 2001, Miao et al., 2004). Rapid pyrolysis produces higher bio-oil yields at better quality compared to slow pyrolysis (Peng et al., 2001, Demirbas, 2002, McKendry, 2002b, Miao & Wu, 2004, Miao et al., 2004).

On a smaller scale, the method of Bligh and Dyer (1959) may be used for lipid extraction from wet biomass and the method described by Zhu et al. (2002) may be used for extraction from dry biomass. The Bligh and Dyer (1959) method uses a single phase solvent, a combination of chloroform and methanol, to extract lipids from biological biomass. The dilution of the single phase with water results in a biphasic solution, with the lower phase containing the chloroform and most of the extracted lipids. Various modifications are made to the Bligh and Dyer (1959) method (Lewis et al., 2000). Modifications include the pre-treatment of samples with enzyme denaturing reagents, (Fishwick & Wright, 1977), the addition of water to the solvent when extracting dry samples (White et al., 1979), acid extraction (Dubinsky & Aaronson, 1979), the sonication of samples (Dunstan et al., 1992), the addition of water to lyophilized samples prior to lipid extraction

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35 (Dunstan et al., 1993), changing the chloroform-methanol ratio (Smedes & Thomasen, 1996) as well as the sequence in which the solvents are added (Lewis et al., 2000).

The pre-treatment of samples with boiling isopropanol effectively denatures potential lipid degrading enzymes and resulted in a more efficient method for lipid extraction (Fishwick & Wright, 1977). The addition of acid to the chloroform-methanol solution resulted in a significant increase in lipid yield (Dubinsky & Aaronson, 1979). The percentage of lipid increase due to acid extraction ranged from 3% to 385% for Chlorella-Euglena and Fragillaria

construens , respectively. Dunstan et al. (1993) demonstrated that the addition of water to lyophilized samples prior to lipid extraction greatly improved lipid yields, especially the yield of triacylglycerols. The order in which the solvents are added also influences the total lipid yield, the addition of solvents in the order chloroform, methanol and water resulted in an increased lipid yield when compared to adding the solvents in reverse (Lewis

et al., 2000). Lewis et al. (2000) also reported that doubling the quantity of methanol used in the extract as well as the sonication of the sample yielded no significant increase in extracted lipid.

A faster and simpler method of lipid extraction is through direct transesterification, a one step process that involves the bypassing of all the extraction and processing steps needed for the analysis of biological specimens by gas-liquid chromatography (GLC) (Lepage & Roy, 1984, 1986). Direct transesterification showed a higher recovery of fatty acids for adipose tissue, milk and bacteria but for plant tissue the recovery of fatty acids did not change (Lewis et al., 2000). A consequence of the direct transesterification method is that lipid class data cannot be obtained (Lewis et al., 2000).

Environmental stress and lipid yield

The production of microalgal biomass for biodiesel must be economically feasible and competitive with liquid fuels. Consequently the success of

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36 biodiesel production from microalgae depends on high biomass productivity and considerable lipid yields [50– 60% of biomass weight in the form of lipids] (Neenan et al., 1986, Liu et al., 2008). Previous studies have shown that when microalgae are subjected to stress in the form of nitrogen deprivation (Piorreck et al., 1984, Illman et al., 2000), silicon deficiency (Lynn et al., 2000), phosphate limitation (Reitan et al., 1994, Lynn et al., 2000), high Fe3+ concentrations [in the initial media] (Liu et al., 2008), high salinity (Takagi et

al., 2006, Rao et al., 2007) and low temperature (Renaud et al., 1995, Renaud

et al., 2002, de Castro Araujo & Garcia, 2005) significant increases in lipid content occurred. This increase in lipid content can make a certain species more favourable for biomass production, e.g. Chlorella sp. (Liu et al., 2008). Although higher lipid content is desirable, high levels of stress cause a reduction in growth rate.

Griffiths & Harrison (2009) reported lipid content for green algae under nutrient-replete conditions to be in the range of 13 % to 31 % dry weight, with a mean of 23 %, under nutrient-depleted conditions the mean lipid content increased to 41 % dry weight. Diatoms showed a wider range of lipid content, ranging from 11 to 51 % dry weight, with a similar mean (25 % dry weight) to that reported for green algae (Griffiths & Harrison, 2009). Nitrogen deprivation helped to increase the lipid content (27 % dry weight) of diatoms (Griffiths & Harrison, 2009); however, the effect was not of the same magnitude. Nutrient deficiency does not always result in an increase in lipid content, some species such as Dunaliella primolecta, Dunaliella salina and Nitzschia palea, to name a few, had lower lipid content under nutrient depleted conditions (Griffiths & Harrison, 2009). Griffiths and Harrison (2009) give a list of 55 species of microalgae and their mean reported lipid content produced under various nutrient conditions.

Illman et al. (2000) showed that when Chlorella species, including Chlorella

vulgaris Beijerinck and the marine strain of Chlorella minutissima Fott & Nováková, are subject to nitrogen deprivation the lipid content increased together with a reduction in growth rate. The lipid content of C. vulgaris and C.

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37 (Illman et al., 2000). The total lipid increase due to nitrogen deprivation for five

Chlorella sp. ranged between 10% and 122 %. The green algae,

Scenedesmus obliquus (Turpin) Kützing, contained a high percentage of total lipid (45% of biomass) at low nitrogen concentrations, while at high nitrogen concentrations the percentage of total lipid dropped to 20 % of biomass (Piorreck et al., 1984). At low nitrogen concentrations the growth rate of

Chlorella vulgaris dropped from 0.99 d-1 to 0.77 d-1 and showed a four hour increase in doubling time from 17 hours to 21 hours (Illman et al., 2000). The increase in lipid content after nitrogen deprivation is not only confined to the Chlorophyta, other taxa also show evidence of lipid increase when nutrients become limiting (Reitan et al., 1994). The freshwater centric diatom Stephanodiscus minutulus (Kützing) Cleve & Möller also produced more lipids under various nutrient (phosphorous, silicon and nitrogen) limitations. The greatest increase in lipid content was observed with P- and Si-limitations. Under non-limiting nutrient concentrations the lipid content for S. minutulus was 34 % DW, when Si, N, and P were limited the lipid content increased to 51 , 32 and 44 % DW, respectively (Lynn et al., 2000).

Piorreck et al. (1984) and Lynn et al. (2000) point out the increase in neutral lipids such as triacylglycerols (triglyceride) when nutrient limitation occurs. The increase in triacylglycerols is preferred since triacylglycerols are transesterified to produce methyl esters (biodiesel) and glycerol (Fukuda et

al., 2001, Chisti, 2007).

Liu et al. (2008) reported that the addition of 1.2 × 10-5 mol L-1 Fe3+ to

Chlorella vulgaris cultures exhibited fast growth in the exponential phase with the onset of the stationery much sooner than the cultures with lower Fe3+ concentrations. The total lipid content as percentage of dry weight at 1.2 × 10 -5

mol L-1 Fe3+ was 56.6 % at 1.2 × 10-6 mol L-1 Fe3+ the total lipids dropped to 16.5 %. The fast onset of the stationary phase reduces the time between each harvest (Liu et al., 2008). The shortened period between two consecutive harvests coupled with high cell densities and increased lipid will increase the

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38 bio-oil yield and will allow microalgae to be competitive with other fuel sources.

Takagi et al. (2006) investigated the effect of salt stress on the accumulation of lipids, especially triacylglyceride in Dunaliella tertiolecta. A seven percent increase, from 60 % to 67 %, in lipids was observed when the salt concentration doubled from 50 mM to 100 mM. A 50 mM NaCl concentration is equivalent to seawater or 35 psu. Cell growth was noticeably lower at high, 150 mM and 200 mM, NaCl concentrations and peaked at a concentration of 100 mM with a slight decrease in cell densities towards hyposaline conditions. Takagi et al. (2006) also demonstrated that when 50 mM or 100 mM of NaCl were added to 100 mM cultures, at mid-log phase or end of log phase, the intracellular lipids increased to 70% with no marked decrease in cell densities.

The lipid content of Botryococcus braunii increased from 20 % in freshwater to 24-28 % at salinities ranging from 17 mM to 85 mM. Biomass increased with increased salinity with maximum biomass obtained at 17 mM and 34 mM salinity (Rao et al., 2007).

De Castro Araujo and Garcia (2005) found that lipid content for the marine diatom Chaetoceros cf. wighamii was higher at lower temperatures, 20 to 25°C. The lipid content for C. cf. wighamii was highest when the cultures were kept at 25°C and a salinity of 25 ppt with the addition of CO2, via bubbling. However at higher salinities the addition of CO2 to cultures at 25°C resulted in a significant reduction in lipid content. The lipid content of four species of microalgae, Chaetoceros sp., Rhodomonas sp., Cryptomonas sp. and an unidentified prymnesiophyte, was generally higher at lower temperatures (Renaud et al., 2002). Growth rates for the four microalgal species followed the same pattern with faster doubling times observed at lower temperatures; some cultures resulted in a negative growth or worse, died at high temperatures. Renaud et al. (1995) observed that, in general, maximum lipid content coincides with optimal range in growth temperature in many species, and this content is lower at temperatures below and above this range.

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39 Maximum lipid content is expected to coincide with optimal range in growth salinity and behave the same as with temperature. This can be an added advantage as it can be used to increase the lipid yield of microalgae, thus producing more biodiesel production.

Regulation of lipid accumulation in oleaginous

micro-organisms

The regulation of lipid accumulation in oleaginous micro-organisms is described in detail by Ratledge (2002). Under normal, non-limiting growing conditions, microalgae produce glucose; this production continues, even when nitrogen becomes limiting. However, the activity of isocitrate dehydrogenase within the mitochondrion slows to a halt, due to a lack of available AMP. This leads to the accumulation of citrate, which in turns get cleaved into acetyl-CoA by ATP: citrate lyase. The enzyme ATP: citrate lyase is absent in non-oleaginous microalgae and is therefore essential for lipid accumulation (Ratledge, 2002). The presence of ATP: citrate lyase does not, however, explain why different microalgae have different lipid production capacities. How much lipid gets produced by the cell is controlled by the activity of malic enzyme (ME), which acts as the sole source of NADPH for fatty acid synthase (FAS) (Ratledge, 2002). The inhibition or genetic absence of malic enzyme will result in very low lipid productivity (Ratledge, 2002).

Effects of salinity on the growth and lipid production of ten species of microalgae from the Swartkops saltworks : a biodiesel perspective