**5. Conversion processes for microbial lipid production**

#### **5.1 Autotrophic microalgae cultivation and lipid accumulation**

Three cultivation processes were designed to culture microalgae and other oleaginous cells, including open-pond system, photobioreactor, and fermentation. The open-pond system is typically a closed loop with a pump to create microalgae flow in the channel. The channel is 0.2-0.5 meter deep, and the pump keeps the microalgae cell well mixed for continuous growth. This type of open-pond system has been used for several years because it is easy to operate and inexpensive to maintain. The open-pond system also can be upgraded to large-scale microalgae production. On the other hand, a high cell density of microalgae cannot be reached because of limited capability to assimilate the sun light and low carbon dioxide concentration of air. Increasing the CO2 concentration by using

Microbial Biodiesel Production - Oil Feedstocks Produced from Microbial Cell Cultivations 101

Table 3. Comparison of different algae cultivation systems (Alabi. 2009)

which two steps are integrated into one.

Table 3 compares three cultivation methods and shows that the process cost of fermentation can be high due to its requirement of raw materials and oxygen, and sterilization of culture media during the cell growth. It is readily available both in the lab and in the industry, but is only suitable to produce high-valued products, of which biofuel products are not. The key barriers to apply this technology to biofuel production is the cost and availability of raw materials. Considering the competition with human diet, sugars cannot serve as the raw material for biofuel production; and alternative materials such as lignocellulosic materials should be used for the heterotrophic oil production. If the oleaginous cells are capable of generating the hydrolytic enzymes for lignocelluloses degradation, it will be the big plus for biodiesel production via oleaginous fermentation the overall system. Otherwise, external hydrolytic enzymes have to be used to release the monosugar, followed by lipid accumulation via olgeaginous microorganisms. Separated hydrolysis and fermentation (SHF) is a common working model to have these two steps separated. Two bioreactors will be necessary because the hydrolytic degradation of lignocellulose is preferred at 50ºC, while the oleaginous microorganisms grow at much lower temperature (28⁰C to 30⁰C for most of the fungus). Simultaneous saccharification and fermentation is another working model currently under intense investigation, in

flue gas instead of air can increase the microagale cell concentrations, but the final cell density is still limited to the mutual shading effects where light cannot penetrate through dense microalgae cell broth. Another problem is biological contamination during the long period of cultivation. The bacteria contamination or other non-oleaginous microalgae invasion can occur in stressed cultural conditions, where lipid accumulation usually is stimulated, such as nitrogen depletion or other nutrient imbalance. There is now extensive evidence that open-pond systems can operate for more than six months without significant contamination using a wide range of microalgae. Prolific strains of *Chlorella*, for example, are often dominant because they outgrow their competitors (and indeed can often be contaminants themselves in *Arthrospira* cultures or other microalgal strains). Extreme halophiles, such as *Dunaliella salina*, are also dominant in their optimal environments because they do not encounter much competition at high salinities. However, in the context of the wider microalgal industry, contamination issues are still of significant interest.

To enhance the productivity of microalgae, closed photobioreactor systems (tubular flat plate, Orcolumn are designed to increase the surface of microalgae broth exposed to sunlight. Closed photobioreactors are more costly than open-pond systems, but they have potential for higher productivity of cell biomass with less chance of contamination. The flat plate photobioreactors can receive greater sunlight for microalgae growth although there is potential for cell mixing. The microalgae cell density could reach up to 80g/L dry cell weight, significantly higher than the cell density of a pond system, which ranges within several g/L (Hu, 1998). Another design for the photoreactor is the tubular photoreactor, made with a diameter less than 0.1 m to maximize the sunlight harvest by microalgae. The tubular reactor can also expose the microalgae cells to sunlight from all the directions (Miron, Gomez et al. 1999; Ugwu, Ogbonna et al. 2002). There are a few reports about scaleup test of tubular photoreactor, such as the one in Hawaii with a size of 25M3 (Olaizola 2000), and 700 M3 in Germany (Pulz 2001). However, the tubular photobioreactors cannot scale-up indefinitely because of oxygen accumulation, carbon dioxide limitation, and pH changes (Eriksen 2008). The third type of photoreactor is the column photoreactor, the most controllable type among three because it most closely resembles the traditional bioreactor. The column containing microalgae is vertical, and the air is bubbled from the bottom. Sunlight is provided horizontally (Eriksen 2008).

In addition to individual open-pond system and closed photobioreactor, the hybrid system of microalgae cultivation is currently under intense investigation because it combines the open-pond system and closed photobioreactor to increase the cell productivity and reduce the cost. The first stage is autotrophy to avoid biological contamination, and the second stage is heterotrophy, which provides the stress condition for lipid accumulation.

#### **5.2 Oleaginous microbial fermentation and lipid accumulation**

Besides the pond system and photobioreactor, heterotrophic cell cultivation, including heterotrophic microalgae, oleaginous yeast and fungi, is usually limited to industrial fermentation tanks. These cells can be cultured in a dark fermentor with optional sunlight, and can usually reach a very high cell density (up to 200g/L). Due to its excellent controllability of all operation parameters, most of the industrial microalgae cultivations for nutraceutical production (e.g., polyunsaturated fatty acid) have been switched to the heterotrophic fermentations where sugar is provided to produce high-valued products, such as Docosahexaenoic acid (DHA).

flue gas instead of air can increase the microagale cell concentrations, but the final cell density is still limited to the mutual shading effects where light cannot penetrate through dense microalgae cell broth. Another problem is biological contamination during the long period of cultivation. The bacteria contamination or other non-oleaginous microalgae invasion can occur in stressed cultural conditions, where lipid accumulation usually is stimulated, such as nitrogen depletion or other nutrient imbalance. There is now extensive evidence that open-pond systems can operate for more than six months without significant contamination using a wide range of microalgae. Prolific strains of *Chlorella*, for example, are often dominant because they outgrow their competitors (and indeed can often be contaminants themselves in *Arthrospira* cultures or other microalgal strains). Extreme halophiles, such as *Dunaliella salina*, are also dominant in their optimal environments because they do not encounter much competition at high salinities. However, in the context of the wider microalgal industry, contamination issues are still of

To enhance the productivity of microalgae, closed photobioreactor systems (tubular flat plate, Orcolumn are designed to increase the surface of microalgae broth exposed to sunlight. Closed photobioreactors are more costly than open-pond systems, but they have potential for higher productivity of cell biomass with less chance of contamination. The flat plate photobioreactors can receive greater sunlight for microalgae growth although there is potential for cell mixing. The microalgae cell density could reach up to 80g/L dry cell weight, significantly higher than the cell density of a pond system, which ranges within several g/L (Hu, 1998). Another design for the photoreactor is the tubular photoreactor, made with a diameter less than 0.1 m to maximize the sunlight harvest by microalgae. The tubular reactor can also expose the microalgae cells to sunlight from all the directions (Miron, Gomez et al. 1999; Ugwu, Ogbonna et al. 2002). There are a few reports about scaleup test of tubular photoreactor, such as the one in Hawaii with a size of 25M3 (Olaizola 2000), and 700 M3 in Germany (Pulz 2001). However, the tubular photobioreactors cannot scale-up indefinitely because of oxygen accumulation, carbon dioxide limitation, and pH changes (Eriksen 2008). The third type of photoreactor is the column photoreactor, the most controllable type among three because it most closely resembles the traditional bioreactor. The column containing microalgae is vertical, and the air is bubbled from the bottom.

In addition to individual open-pond system and closed photobioreactor, the hybrid system of microalgae cultivation is currently under intense investigation because it combines the open-pond system and closed photobioreactor to increase the cell productivity and reduce the cost. The first stage is autotrophy to avoid biological contamination, and the second

Besides the pond system and photobioreactor, heterotrophic cell cultivation, including heterotrophic microalgae, oleaginous yeast and fungi, is usually limited to industrial fermentation tanks. These cells can be cultured in a dark fermentor with optional sunlight, and can usually reach a very high cell density (up to 200g/L). Due to its excellent controllability of all operation parameters, most of the industrial microalgae cultivations for nutraceutical production (e.g., polyunsaturated fatty acid) have been switched to the heterotrophic fermentations where sugar is provided to produce high-valued products, such

stage is heterotrophy, which provides the stress condition for lipid accumulation.

**5.2 Oleaginous microbial fermentation and lipid accumulation** 

significant interest.

Sunlight is provided horizontally (Eriksen 2008).

as Docosahexaenoic acid (DHA).



Table 3 compares three cultivation methods and shows that the process cost of fermentation can be high due to its requirement of raw materials and oxygen, and sterilization of culture media during the cell growth. It is readily available both in the lab and in the industry, but is only suitable to produce high-valued products, of which biofuel products are not. The key barriers to apply this technology to biofuel production is the cost and availability of raw materials. Considering the competition with human diet, sugars cannot serve as the raw material for biofuel production; and alternative materials such as lignocellulosic materials should be used for the heterotrophic oil production. If the oleaginous cells are capable of generating the hydrolytic enzymes for lignocelluloses degradation, it will be the big plus for biodiesel production via oleaginous fermentation the overall system. Otherwise, external hydrolytic enzymes have to be used to release the monosugar, followed by lipid accumulation via olgeaginous microorganisms. Separated hydrolysis and fermentation (SHF) is a common working model to have these two steps separated. Two bioreactors will be necessary because the hydrolytic degradation of lignocellulose is preferred at 50ºC, while the oleaginous microorganisms grow at much lower temperature (28⁰C to 30⁰C for most of the fungus). Simultaneous saccharification and fermentation is another working model currently under intense investigation, in which two steps are integrated into one.

Microbial Biodiesel Production - Oil Feedstocks Produced from Microbial Cell Cultivations 103

very limited in its technical and economic viability. Most conventional and economical separation methods such as filtration and gravitational sedimentation are widely applied in wastewater treatment facilities to harvest relatively large (>70 µm) microalgae such as *Coelastrum* and *Spirulina*. However, they cannot be used to harvest algae species approaching bacterial dimensions (<30 µm) like *Scenedesmus*, *Dunaliella*, and *Chlorella* (Brennan and Owende 2010), to which most oleaginous microalgae species belong. Centrifugation is a method widely used to recover microalgae biomass, especially smallsized algae cells; however, its application is restricted to algae cultures for high-value metabolites due to intensive energy needs and high equipment maintenance requirements. While flocculation is used to harvest small-sized microalgae cells, it is a preparatory step to aggregate the microalgae cells and increase the particle size so that other harvesting methods such as filtration, centrifugation, or gravity sedimentation can be applied (Molina Grima, Belarbi et al. 2003). Several flocculants have been developed to facilitate the aggregation of microalgae cells, including multivalent metal salts like ferric chloride (FeCl3), aluminium sulphate (Al2(SO4)3), and ferric sulphate (Fe2(SO4)3), and organic polymers such as Chitosan (Li, Horsman et al. 2008). Chemical flocculation can be reliably used to remove small algae cells from pond water by forming large-sized (1–5 mm) flocs (Sharma, Dhuldhoya et al. 2006). However, the chemical reactions are highly sensitive to pH and the high doses of flocculants required produce large amounts of sludge and may leave a residue in the treated effluent. In summary, most technologies including chemical and mechanical methods greatly increase operational costs for algal production and are only economically

feasible for production of high-value products (Park, Craggs et al. 2011).

investments on equipment and chemical supplies are still needed.

Besides traditional methods mentioned above, there are several new technology developments in this field. DOE-ARPA-E recently funded a research project for Algae Venture Systems (AVS) to develop a Harvesting, Dewatering, and Drying (AVS-HDD) technology by using the principles of liquid adhesion and capillary action to extract water from dilute microalgae solutions. Attached algal culture systems have been developed for growing microalgae on the surface of polystyrene foam (Wilkie and Mulbry 2002) (Johnson and Wen 2010) to simplify the cell harvest. New bioflocculants, which are more environmentally friendly, are also proposed to address the cost and environmental concerns for current flocculation method (Uduman, Qi et al. 2010). All these methods are innovative and will decrease the harvest cost to some extent if developed successfully, but heavy

Dr. Bo Hu's research group at University of Minnesota developed an innovative approach to enhance natural algae aggregation and to encourage simple gravity settling or filtration by co-culturing filamentous fungal cells at the end of the microalgae cultures. Instead of suspended culture, this approach uses pelletized or granulized culture where cells form pellets in culture medium. In submerged cultures, many filamentous microorganisms tend to aggregate and grow as pellets/granules. They are spherical or ellipsoidal masses of hyphae with variable internal structure, ranging from loosely packed hyphae, forming "fluffy" pellets, to tightly packed, compact, dense granules (Hu and Chen 2007; Hu and Chen 2008; Hu, Zhou et al. 2009; Chunjie Xia 2011). Besides merits from the cell immobilization, there are several other advantages, especially for the micro-oil production: a). easy to harvest cells, and b). easy to re-use pond water (Johnson and Wen 2010; Xia 2011). As the first research group to introduce pelletized liquid fermentation (PLF) into biofuel production, this research group at University of Minnesota found key operational conditions that induce the fungal pelletization. They discovered that changing conditions

Different fermentation processes are applied to obtain high productivity of lipids and high conversion ratio of substrate for the fermentation, such as batch cultivation, fed-batch cultivation, and continous cultivation. Fed-batch cultivatioin is a modified batch model that can reach high cell density and it has many applications in the fermentative lipid accumulation process. For example, *Rhorosporidum toruloides* reach much higher cell density with 48% lipid compared to its batch cultivation (Li, Zhao et al. 2007). The high productivity of fed-batch cultivation was conformed by *Phodotorula glutinis* (Xue, Miao et al. 2008), *C. curvatus* (Meesters, Huijberts et al. 1996), and *L. starkeyi* (Yamauchi, Mori et al. 1983). Continuous cultivation has advantages of easy maintenance and time-saving, although it is difficult to control the contamination. It has limited applicatioins in the fermentative lipid accumulation.

Besides the commonly used submerged cultivations, solid state fermentation, as a compact process for lipid production, showed many advantages, such as low requirements to the raw materails; low capital cost; low energy expenditure; less expensive downstream processing; less water usage and low water output; potential higher volumetric productivity; less fermentation space; easy operation and maintenance. The research for *Aspergillus oryzae* growing on rice bran and wheat bran through solid state fermentation resulted to the lipid content of cell biomass at about 10-11% (Da Silveira, Oliveira et al. 2010). The lipid yield reached 62.87 mg/gds in solid state fermentation on the 6th day after Plackett-Burman design (PBD) by *A. oryzae* A-4 (Lin, Cheng et al. 2010) . Currently, the solid state fermentation research is still in its infancy and many barriers are hindering this process from commercilization. The lipid yield is relatively low compared to submerge cultivation. Modern biotechnological approaches, such as heterogenous expression of hydrolytic enzymes and UV radiation, are available to enhance the hydrolytic enzymes production (Li, Yang et al. 2010; Awan, Tabbasam et al. 2011). Semi-solid state fermentation is used to avoid high sugar concentration on the surface of lignocellulose. An oleaginous fungus *M. isabellina*  was cultured at semi-solid state fermentation with the results of 11g oil per 100g sweet sorghum (Economou, Makri et al. 2010) .
