WO2012109642A1 - Aqueous extraction methods for high lipid microorganisms - Google Patents

Abstract

Systems and methods for obtaining an extracted lipid fraction from microorganisms are described. The method includes growing the microorganisms in aqueous culture under conditions to provide microorganisms having a high level of lipids. The culture of microorganisms is then dewatered to provide a concentrated culture having an increased amount of solids by weight while still including a substantial amount of water. A controlled disruption of the concentrated culture is then carried out to provide a disrupted culture, which is then separated under aqueous conditions to provide an extracted lipid fraction.

Description

AQUEOUS EXTRACTION METHODS FOR HIGH LIPID MICROORGANISMS

RELATED APPLICATION DATA

[0001] This application claims the benefit of U.S. Provisional Application Serial No. 61/442,197, filed February 12, 2011, which is incorporated by reference herein.

GOVERNMENT FUNDING

[0002] This work was supported, at least in part, by grant number DE-FE0001888 from the Department of Energy. The United States government has certain rights in this invention.

TECHNICAL FIELD

[0003] The present application relates generally to extracting lipids from microorganisms under aqueous conditions. More specifically, the application relates to growing microorganisms such as algae in culture so that they include high levels of lipids, disrupting the microorganisms to release the lipids, and then separating the lipids from the disrupted microorganisms.

BACKGROUND OF THE INVENTION

[0004] Biofuels have an increasing role in the United States energy market as energy prices increase, political emphasis on establishing national energy independence intensifies and apprehension about climate change continues to grow. The price of petroleum has fluctuated dramatically, reaching record highs of more than US $140 per barrel in 2008. In part, those price increases reflected economic, political, and supply chain uncertainties. Political concerns about the availability of petroleum supplies have led to the realization that the United States' energy independence is of critical strategic importance, both economically and strategically. The release of C0₂ from fossil fuel combustion may substantially contribute to global warming, also intensifying efforts to develop biofuels. The United States has responded by issuing a renewable fuel standards update that encourage a shift to more advanced biofuels in the market. For example, Renewable Fuel Standard 2 (RFS2) requires 36 million gallons of renewable fuel to be blended into transportation fuel by 2022. Recently the EPA updated the mandated renewable fuel allotments in an amendment to the RFS2, these revisions are anticipated to happen annually in the future. Additionally, many states have responded by enacting their own renewable portfolio standards mandating electricity providers obtain a certain percentage of their power from renewable energy sources. As a result of these concerns and RFS requirements, domestically produced biofuels have become an increasingly attractive alternative to foreign fossil fuels.

[0005] Microalgae are some of the most productive and therefore, desirable sources for biofuel feedstocks. The Department of Energy (DOE) determined that biofuel yield per acre from microalgal culture exceeds that of competing organisms and land crops. Between the late 1970s and 1990s, the DOE's National Renewable Energy Laboratory (NREL) evaluated the economic feasibility of producing biofuels from a variety of aquatic and terrestrial photosynthetic organisms. Sheehan et ah, Close Out Report, Aquatic Species Program, NREL/TP380-24190 (1998). Biofuel production from microalgae was determined to have the greatest yield per acre potential of any of the organisms screened. Microalgal biofuel production was estimated to be 8 to 24 fold greater than the best terrestrial biofuel production systems. Current estimates of the potential productivity for algal biofuel production range from 2,000 to 10,000 gallons/acre (18,707 to 93,540 L/ha). According to the DOE, microalgae yield "30 times more energy per acre than land crops such as soybeans." Although existing technologies are promising, there is a significant need for systems and methods that create even greater efficiencies in biofuel production from microalgae in order to meet economic targets needed for successful commercialization.

[0006] One of the unit processes of algal biofuel production that can benefit the most from improved efficiencies is the oil extraction process. Extraction is the process by which lipids contained within the algal cells are liberated and purified. Traditional methods of lipid extraction are costly, since they rely on relatively dry feedstock (<10% moisture), solvent, and distillation of a solvent/lipid miscella to recover the lipid and allow recycle of the solvent. Drying and distillation are very energy intensive and make these standard methods impractical for application in the production of algal biofuels. This is particularly true because algal biofuels must compete with traditional sources of fuel. In addition, these older methods produce low lipid yields and extract unwanted oil impurities, leaving much room for improvement. Furthermore, because algae is grown at very low concentrations (typically less than 1% solids), these methods also require the algal feedstock to be significantly dewatered prior to drying which is a time-consuming and costly process.

[0007] Existing methods of extraction include: (1) pressing and expelling, (2) solvent extraction using hexane, and (3) supercritical fluid extraction. A. Demirbas, Production of Biodiesel from Algae Oils, Energy Sources, Part A, 31:163-168, (2009). The efficiency of extraction through pressing or expelling is limited, between 70 and 75% of the oils contained in the algae. To employ this method, the algae also have to be dried, a time-consuming and energy-intensive process.

[0008] Solvent extraction of dried feedstocks (< 10% moisture) presents similar limitations. The most common chemical used in this type of extraction is hexane. The efficiency of traditional solvent extraction methods with hexane is compromised when applied to materials with more than 10% water content. As discussed previously, dewatering is time-consuming and energy-intensive and any process that decreases the need for dewatering is a strong driver for cost containment.

[0009] Using hexane and similar solvents can also lead to the formation of lipid micelles. After lysing, the cellular components can act as a surfactant causing the water and the lipids to form a stable emulsion. The lipids trapped in the emulsion are either lost or must be recovered by further processing. If these trapped lipids cannot be recovered, large refining losses will result which compromise the efficiency of the process. Hy et al, JAOCS, Vol. 83, no. 5 (2006), 457-460. This increases the overall cost of the biofuel produced.

[0010] The resulting oil from a typical, solvent-based extraction process also contains relatively large amounts of phospholipids and sulfur as well as chlorophyll and other pigments, because the solvent extracts these materials from the algal cells. Before the algal oil can be used as a biofuel, it must be further processed to remove these extracted impurities which are not suitable for the particular fuel application.

[0011] Supercritical fluid extraction utilizes the enhanced solvating power of fluids above their critical point. Although this method can achieve extraction yields of close to 100%, economical production of bio fuels from oleaginous microalgae via supercritical processing is challenged by issues of energy-intensive processing and scaling up the process. Supercritical fluid extraction requires high pressure equipment that is expensive and energy intensive. This makes it impractical for usage in biofuel production.

[0012] Given the drawbacks of these existing extraction methods, there is a need for a more practical, economical extraction method for obtaining biofuel from microorganisms. Ideally, this method would rely less on high energy input steps and expensive and energy-intensive equipment, while requiring little dewatering.

SUMMARY OF THE INVENTION

[0013] The present invention provides systems and methods for improving the process of producing biofuels from microorganisms by eliminating the cost of drying the biomass and improving lipid extraction efficiency and lipid purity. This is achieved by performing an aqueous extraction on microorganisms cultured to have high lipid content, using growth in a combination of phototrophic, mixotrophic and/or heterotrophic conditions. Embodiments of the method can provide an extracted lipid fraction that can serve as a biofuel precursor that is relatively free of phospholipids, chlorophyll, sulfur, and other contaminants, thereby avoiding further purification and processing costs. The invention can obviate the need for costly further processing of the extracted lipid such as distillation. Aspects of the present application increase the efficiency in the lipid extraction process by achieving the isoelectric point of the culture containing the microorganisms before lysing them, thus reducing the formation of lipid-water-cell debris emulsions.

[0014] Accordingly, in one aspect, the present invention provides a method of obtaining an extracted lipid fraction from microorganisms that includes growing the microorganisms in aqueous culture under conditions to provide microorganisms having greater than about 45% of their dry weight being lipids; dewatering the culture of microorganisms to provide a concentrated culture having greater than about 15% solids by weight; mixing the concentrated culture with a partitioning agent; conducting a controlled disruption of the concentrated culture including a partitioning agent to provide a disrupted culture; and separating an extracted lipid fraction from the disrupted culture by phase separation.

[0015] Embodiments of the method include use of a first stage that includes phototrophic, mixotrophic, or a combination of phototrophic and mixotrophic conditions, followed by a second stage that includes different conditions selected from mixotrophic or heterotrophic conditions, or a combination thereof. Use of two different growth environments can provide an efficient route to obtain microorganisms that include high lipid levels.

[0016] In many embodiments of the invention, the microorganisms being used are algae. Γη further embodiments, the algae can be of the genus Chlorella. In yet further embodiments, the algae are selected from the species Chlorella protothecoides, C. kessleri, C. vulgaris, C. sorokiniana, C. zoflngiensis, C. minutissima, C. regularis, and C. variabilis.

[0017] Embodiments of the method can include specific process conditions. In some embodiments, the concentrated culture and the disrupted culture always include at least about 10% water. In additional embodiments, the pH of the concentrated culture is adjusted to the isoelectric point prior to cell disruption. In further embodiments, the ionic strength of the concentrated culture is adjusted prior to cell disruption. Yet further embodiments include the step of diluting the disrupted culture with water or salt water to decrease the effect of the partitioning agent and better affect separation of the lipid fraction.

[0018] Additional embodiments include adding a partitioning agent to enhance the extraction of lipids from algal cells. The partitioning agent, which reduces the surface tension between the oil and aqueous phase and in addition reduces the interactions between the oil and cell debris, thereby reduces emulsions containing broken biomass. The partitioning agent is preferably an amphiphilic chemical with oil dispersing properties, such as acetone. If a portioning agent is used, the method can also include the step of recovering the partitioning agent from the disrupted culture using evaporation, stripping, or a combination thereof.

[0019] Embodiments of the invention provide an extracted lipid fraction having various forms of improved purity or other characteristics. In some embodiments, the extracted lipid fraction includes at least about 70% triglycerides. In further embodiments, the extracted lipid fraction includes about 10% or less phospholipids, i yet further embodiments, the extracted lipid fraction includes about 5% or less chlorophyll pigment.

[0020] Another aspect of the invention provides a method of obtaining an extracted lipid fraction from microorganisms that includes growing the microorganisms in aqueous culture under conditions to provide microorganisms having greater than about 45% of their dry weight being lipids; dewatering the culture of microorganisms to provide a concentrated culture having greater than about 15% solids by weight while still including a substantial amount of water; conducting a controlled disruption of the concentrated culture to provide a disrupted culture; and separating an extracted lipid fraction f om the disrupted culture that includes about 60% or more of the lipid present in the microorganisms before extraction. Embodiments of this method can include the step of diluting the disrupted culture with salt water. In further embodiments, the step of growing the microorganisms in culture includes a first stage that includes phototrophic, mixotrophic, or a combination of phototrophic and mixotrophic conditions, followed by a second stage that includes different conditions selected from mixotrophic or heterotrophic conditions, or a combination thereof.

[0021] Another aspect of the invention provides a system for producing lipid from microorganisms that includes a first culturing apparatus for culturing microorganisms under phototrophic, mixotrophic, or a combination of phototrophic and mixotrophic conditions; a second culturing apparatus that receives cultured microorganisms from the first culturing apparatus for culturing the microorganisms under different conditions selected from mixotrophic or heterotrophic conditions, or a combination thereof to provide an oleaginous culture of microorganisms having greater than about 45% of their dry weight being lipids; a dewatering apparatus to remove a portion of the water from the oleaginous culture to provide a concentrated culture having greater than about 15% solids by weight; a disruption apparatus for disrupting the cell structure of the microorganisms in the concentrated culture to provide a disrupted culture; and a separation apparatus for separating a substantial portion of the lipid from the disrupted culture. Embodiments of this system can include a concentrating apparatus to concentrate the microorganism culture from the first culturing apparatus before it is provided to the second culturing apparatus.

BRIEF DESCRIPTION OF THE FIGURES

[0022] The present invention may be more readily understood by reference to the following figures wherein:

[0023] Figure 1 provides a schematic representation of the aqueous extraction process used to obtain an extracted lipid fraction from microorganisms.

[0024] Figure 2 provides a schematic representation of a system for producing lipid from microorganisms. [0025] Figure 3 provides an additional schematic representation of a system for producing lipid from microorganisms that includes additional processing steps.

[0026] Figure 4 provides a picture showing the results of extraction using ethanol or acetone as the partitioning agents.

[0027] Figure 5 provides a picture showing the results of extraction with isopropyl alcohol as the partitioning agent followed by cell disruption using a Hockmeyer® immersion mill (HM Micro Mill).

[0028] Figure 6 provides a picture showing the results of separation after adding salt to the water prior to phase separation. The tube labeled "M" shows separation with no salt, while the tube labeled "O" shows separation with salt added.

[0029] Figure 7 provides a picture showing the number of cells that survive when the bead beater time and number of microfluidizer passes are varied.

[0030] Figure 8 provides a bar graph showing how the amount of oil mass extracted varies depending on the amount of time the sample is subjected to bead beating (left 3 bars) or the number of microfluidizer passes (right four bars).

[0031] Figure 9 provides a picture showing the oil recovery in samples after each pass (increasing from left to right) in the microfluidizer.

[0032] Figure 10 provides a picture showing the oil recovery in each sample (increasing from left to right) treated by the bead beater for increasing 15 increment amounts of time.

DETAILED DESCRIPTION OF THE INVENTION

DEFINITIONS

[0033] Unless otherwise defined, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which this application pertains. Although methods and materials similar or equivalent to those described herein can be used in the practice or testing of the exemplary embodiments, suitable methods and materials are described below. In case of conflict, the present specification, including definitions, will control. In addition, the materials, methods, and examples are illustrative only and not intended to be limiting.

[0034] The terminology as set forth herein is for description of the embodiments only and should not be construed as limiting the application as a whole. Unless otherwise specified, "a," "an," "the," and "at least one" are used interchangeably. Furthermore, as used in the description of the application and the appended claims, the singular forms "a", "an", and "the" are inclusive of their plural forms, unless contraindicated by the context surrounding such. The singular "alga" is likewise intended to be inclusive of the plural "algae."

[0035] The recitations of numerical ranges by endpoints include all numbers subsumed within that range (e.g., 1 to 5 includes 1, 1.5, 2, 2.75, 3, 3.80, 4, 5, etc.).

[0036] Algae microorganisms, or "microalgae," and in particular the various genera and species of these organisms have been defined in various manners through the ages. It is therefore prudent to make reference to a fixed reference from which microalgae in their various forms can be defined. For example, the genus Chlorella has recently been split into a number of new genera (Var et al, 1999 J Phycology 35:587-598) but for the purposes of this application A xenochlorella, Parachlorella, and any new genera created prior to 1990 from the genus Chlorella are contemplated herein. This would include for example Chlorella protothecoides, C. kessleri, C. vulgaris, and C. soroldniana. For the purposes of this patent application, microalgae include the traditional groups of algae described in Van Den Hoek et al, Algae: An Introduction to Phycology (1995). Additionally, genera and associated species are in a taxonomic flux so unless specifically referenced the Van den Hoek reference will be used as the standard.

[0037] The Heteroboost™ process is defined as inoculating a microorganism biomass into a bioreactor where the microorganism biomass was previously grown phototrophically or mixotrophically, then partially harvested or concentrated. It is then placed in a bioreactor where the microorganism biomass utilizes a fixed carbon source to increase lipid content, either mixotrophically or heterotrophically.

[0038] Microorganisms can be grown under phototrophic, mixotrophic, or heterotrophic conditions. Photoautotrophy means that light is used as the only source of energy for the organism utilizing simple compounds such as carbon dioxide as a mean for capture and storage of the energy that is developed in excess of cell maintenance. The terms phototrophy and photoautotrophy are often indistinguishably used, but strictly speaking they are different. For phototrophy, light can be used as a partial energy source for growth of the organism, but other sources of energy can also be used. Mixotrophy is where an organism uses light and fixed carbon sources for energy to survive simultaneously, and therefore, a form of phototrophy. Heterotrophy, on the other hand, is where the organism's energy is solely derived from fixed carbon compounds.

[0039] "High lipid content" refers to microorganisms having a higher than normal lipid content, and is further defined herein. Generally algae grown in phototrophic or mixotrophic ponds have low amounts of lipid (ranging from 3-15% total lipids), unless special steps are taken to produce higher levels of lipid (from 15-85%). For the purposes of this application high lipid concentrations are those exceeding around 40% total lipid (including all classes of lipid).

[0040] "Aqueous extraction" refers to an extraction process with significant amounts of water present in the feedstocks being extracted, and water is the predominant solvent present. For the purposes of this application feedstocks with greater than 10% moisture content should be considered an aqueous extraction.

[0041] The exemplary embodiments of the present application are directed at improving the extraction of desirable compounds from oil enriched cells using feedstock having about 10% or more moisture content. This is particularly applicable to producing feedstock for the production of an extracted lipid fraction useful as algal biofuel or a biofuel precursor. The present application could also be used to extract lipids and biofuels from other microorganisms that can be grown in a number of trophic state combinations of phototrophic, mixotrophic and heterotrophic conditions, including but not limited to species of oleaginous algae, bacteria, protists, and fungi, either in their native form or genetically modified to produce high levels of lipid.

[0042] The present invention provides a method of obtaining an extracted lipid fraction from microorganisms. This method includes first growing the microorganisms in aqueous culture under conditions to provide microorganisms having greater than about 45% of their dry weight being lipids. The culture of microorganisms is then dewatered to provide a concentrated culture having greater than about 15%» solids by weight. The concentrated culture is then mixed with a partitioning agent, and the concentrated culture is subject to a controlled disruption to provide a disrupted culture. Finally, an extracted lipid fraction is separated from the disrupted culture by phase separation.

[0043] Figure 1 provides a schematic representation of the aqueous extraction process. The first step 10 involves growing a microorganism under aqueous culture conditions which provide microorganisms having a high lipid content. The culture is then harvested and dewatered 20 to reduce the water content of the culture. A partitioning agent is then added 30 to the concentrated culture. During this step, it may also be preferable to adjust the pH to the isoelectric point of the concentrated culture. The concentrated culture is then subjected to a controlled disruption 40 that disrupts the microorganisms of the concentrated culture. By controlling the nature of the disruption, the cell debris surface area is minimized. The partitioning agent helps the lipids to enter the aqueous phase during this step. After cell disruption, a phase separation 50 of the disrupted culture is carried out. This can be encouraged by addition of water to dilute the partitioning agent thus reversing its effects and/or the addition of energy in the form of centrifugation or use of a tricanter centrifuge to separate the biomass, aqueous phase (+ partitioning agent), and extracted lipid fraction. The separated phases are then further processed. The extracted lipid phase can be stored 60, further purified, or used directly as a crude oil. The biomass and aqueous phases can be processed to recover the partitioning agent 70 for reuse.

[0044] The first step in the method of obtaining an extracted lipid fraction of the invention is to grow microorganisms in an aqueous culture. Examples of microorganisms suitable for use include algae, yeasts, and fungi. Suitable yeasts include, but are not limited to, Rhodotorula, Saccharomyces, and Apiotrichum strains. Suitable fungi species include, but are not limited to, species of the Mortierella genus.

[0045] The microorganisms can also be algae, and in particular oleaginous algae. An oleaginous alga is an algae species that can, under known conditions, accumulate a significant portion of its biomass as lipid. For example, embodiments of oleaginous algae are algae species that are capable of accumulating at least 10%, at least 20%, at least 30%, at least 40%, or at least 50% of their biomass as lipid. Suitable oleaginous algae species can be found in the Bacillariophyceae, Chlorophyceae, Cyanophyceae, Xanthophyceae, Chrysophyceae, Chlorella, Crypthecodinium, Schizocytrium, Nannochloropsis, Ulkenia, Dunaliella, Cyclotella, Navicula, Nitzschia, Cyclotella, Phaeodactylum, and Thaustochytrid classes and genera. A preferred genus of oleaginous algae is Chlorella, which includes numerous species capable of accumulating about 55% of their total biomass as lipids. See for example Miao & Wu, Journal of Biotechnology, 110, p. 85-93 (2004). Suitable Chlorella species include Chlorella protothecoides, C. kessleri, C. vulgaris, C. sorokiniana, C. zofingiensis, C. minutissima, C. regularis, and C. variabilis.

[0046] The method of the invention also includes growing the microorganisms in aqueous culture. The species of microorganism used form a part of an aqueous culture. The aqueous culture refers to one or more species of microorganism living in an environment that enables their survival and possible growth. The culture may be either an artificial culture found in a biofuel production facility, or it can be a natural culture found in the microorganisms' natural environment. Such strains can also be engineered using modern transgenic techniques to increase their lipid content. The culture conditions required for various microorganisms are known to those skilled in the art. For example it is known that often phototrophic algae do not accumulate lipid unless they are stressed for a period of time (e.g., nitrogen or sulfur stress). Examples of the components of an aqueous culture include water, carbon dioxide, nitrogen, phosphorus, minerals and light. However, the components of the aqueous culture can vary depending on the species of microorganism, and whether or not conditions for autotrophic, mixotrophic, or heterotrophic growth are desired. For autotrophic growth, the culture will require C0₂ and light energy (e.g., sunlight), whereas heterotrophic growth requires organic substrates such as sugar for the growth of the aqueous culture, and can be carried out in the absence of light energy. A mixotrophic culture utilizes both a fixed carbon source and sunlight for growth. Appropriate temperature conditions should be maintained, and preferably that the culture is mixed to provide even access to nutrients and/or light. The aqueous culture is a culture of microorganisms growing in an aqueous environment made up primarily of water. The culture can be an artificial monoculture including a single dominant species, or at least is intended as such, taking into account possible contaminating predators and competitors. Use of such a monoculture makes it easier to provide optimal culture conditions, and can simplify growing and processing the microorganism. However, consortia of microorganisms can also be used, and particularly have a place in an open production system. [0047] The aqueous culture of microorganisms is carried out under conditions to provide microorganisms (e.g., algae) having a high amount of their dry weight as lipids. These organisms can have greater than about 30%, greater than about 40%, greater than about 45%, or greater than 50% of their dry weight being lipids. Use of microorganisms that include a relatively high percentage of lipids facilitates the separation of this lipid from other components of the aqueous culture such as water and cell debris, and minimizes nonpolar lipid loss to emulsions, cell debris, and water. Use of microorganisms having a high amount of lipids also facilitates the use of a method that does not rely on extraction using non-polar solvents. However, an amount of the lipid present in the microorganisms will generally be lost. The amount lost is a function of cell debris surface area and emulsion formation. The higher the starting nonpolar lipid content, the more nonpolar lipid can be recovered.

[0048] Lipids, as defined herein, include naturally occurring fats, waxes, sterols, carotenoids, monoglycerides, diglycerides, triglycerides, and phospholipids. The preferred lipids are fatty acid lipids found in triacylglycerides. Free fatty acids are synthesized in algae through a biochemical process involving various enzymes such as trans-enoyl-acyl carrier protein (ACP), 3-hydroxyacyl-ACP. 3-ketoacyl-ACP, and acyl-ACO. Examples of free fatty acids include fatty acids having a chain length from 14 to 20, with varying degrees of unsaturation. A variety of lipid-derived compounds can also be useful as biofuel and may be extracted from oleaginous algae. These include isoprenoids, straight chain alkanes, and long and short chain alcohols, which short chain alcohols including ethanol, butanol, and isopropanol.

[0049] Microorganisms can be cultured to have a high amount of their dry weight as lipids through various methods. For example, microorganisms can be chosen that exhibit high natural levels of lipid production or microorganisms can be genetically engineered to exhibit high levels of lipid production. See for example U.S. Patent Serial No. 12/743,434, entitled "Molecular Approaches for the Optimization of Biofuel Production," the disclosure of which is incorporated herein by reference. Alternatively, microorganisms could be grown heterotrophically in a bioreactor or fermentor to accumulate large amounts of lipid.

[0050] Microorganisms can also be subjected to stress in order to increase the amount of their lipid dry weight. For example, the microorganisms can be subjected to nutrient stress. Nutrient stress is a condition in which insufficient nutrients are available for the algae to freely proliferate, and typically will result in a decrease in the growth rate of the stressed algae species. Nutrient stress can result from a general unavailability or insufficient quantities of a variety of different nutrients, or it can occur as a result of the absence of a single essential nutrient. Examples of essential nutrients include carbon dioxide, nitrogen, sulfur, molybdenum, magnesium, specific vitamins, and iron. While nutrient deficiency (e.g. nitrogen deficiency) inhibits the cell cycle and production of most algal cellular components, the rate of lipid synthesis remains relatively high, leading to the accumulation of lipids in nutrient-starved algae cells. However, while microorganisms can be stressed by low nutrient availability, other types of stress can be applied as well. Essentially, any significant deviation from the preferred culture system for a microorganism can result in stress. Other sources of stress include too much or too little light, significant deviations in temperature, pH, or salinity from those preferred by the species. The stress should be applied for a number of hours in order to achieve the desired effect of increasing lipid levels. For example, the stress can be applied for a period of about 6 to about 48 hours, for about 10 to 32 hours, or for about 12 to 24 hours. In some embodiments, the microorganisms are stressed by inhibiting nutrient assimilation under nutrient replete conditions, in which nutrients are available but their uptake or use is prevented.

[0051] Culturing the microorganisms to have high lipid levels can be performed by a method using a combination of phototrophic, mixotrophic, and heterotrophic growth conditions. Preferably, the microorganism is first grown under conditions which are at least partially phototrophic to reduce stress and facilitate rapid and inexpensive growth, and then the microorganism is switched to conditions that are at least partially heterotrophic, to trigger an increase in lipid levels. This method of increasing lipid levels can also be referred to herein as the Heteroboost™ process. See for example U.S. Patent Serial No. 12/328,695 for "Optimization of Biofuel Production." In this manner, the method of growing the microorganisms in culture includes a first stage that includes phototrophic, mixotrophic, or a combination of phototrophic and mixotrophic conditions, followed by a second stage that includes different conditions selected from mixotrophic or heterotrophic conditions, or a combination thereof.

[0052] The following describes a more detailed description of how the Heteroboost™ process can be carried out using algae to provide a microorganism having high lipid levels in particular embodiments of the invention. The first stage includes phototrophic or mixotrophic biomass generation. In the case of an algal strain, it can be grown phototrophically in closed or open photobioreactors. It is not necessary at this stage for the cells to contain high lipid content. For example, with Chlorella protothecoides strain KRT1007, lipid concentration is around 7-15% total lipid with the majority being present as phospholipids. In the same open or closed photobioreactors, sugar substrates can be added to induce mixotrophic growth. Under these conditions, nonpolar lipid can quickly accumulate.

[0053] An optional dewatering step can be included between the first and second growth stages. Dewatering, as the term is used herein, refers to a process that decreases, but does not eliminate, water present in the culture. After the first stage of phototrophic or mixotrophic growth, the culture can be dewatered to concentrate the biomass while recovering the water. Dewatering is achieved by any of a number of methods including but not limited to centrifugation, filtration (e.g., ultrafiltration or micro filtration), pressing or other methods that are known to one skilled in the art. The microorganism culture will split into two phases during dewatering; a heavy phase and a light phase. The heavy phase contains the cells, and the light phase contains the process water. The light aqueous phase can be treated, if necessary, and then recycled back into the open pond, bioreactor, or other environment where the microorganisms are grown. The heavy phase generally includes from about 3% -10% solids. However, higher levels of dewatering, from 10% to 30% solids, can be carried out in some embodiments.

[0054] After the first stage, and in some embodiments after the optional dewatering step, a second stage providing a mixotrophic or heterotrophic growth step is carried out. In this stage, the microorganisms (e.g., those recovered in the heavy phase) can be switched to mixotrophic or heterotrophic growth to induce accumulation of lipids. Growing the algae mixotrophically, heterotrophically, or a combination of the two will increase the lipid content and decrease other compounds that could co-purify and contaminate the lipid end product, such as but not limited to chlorophyll. This enhances lipid accumulation by growing the algae in an optimized combination of phototrophic, mixotrophic and/or heterotrophic conditions. A culture medium may be added at this step to facilitate the accumulation of oil in the cell. After the Heteroboost™ process, the algal cells will contain as much as around 70% lipid for Chlorella protothecoides and will vary with species, strain and isolate used. Not only is the lipid content of these cells high, but the composition of the lipid, primarily triacylglycerols (TAGs), is highly desirable for biofuels. In addition to containing large amounts of desirable lipids, the mature cells also contain relatively small amounts of undesirable compounds. For instance, changes in the cell and structural lipids result in a low amount of less desirable lipids such as sterols and phospholipids. The cells also contain a decreased quantity of extractable chlorophyll and other pigments. Avoiding relatively higher levels of these less desirable lipids and pigments avoids the necessity of later removing them from the final product to prepare it for use as a biofuel feedstock.

[0055] Once the algal cells have matured and accumulated lipid, the culture of microorganisms is then dewatered to provide a concentrated culture that still includes a substantial amount of water. Dewatering is carried out as in the previously described optional dewatering step between the first and second stages of the Heteroboost™ process. However, a higher level of dewatering is generally carried out at this step. Dewatering is carried out to provide a concentrated culture that includes at least about 15% solids, or in additional embodiments, about 20%, 25%, or even 30% solids by weight. This dewatered culture is referred to as the concentrated culture, due to this higher level of solids (e.g., algal biomass). Dewatering can be carried out using methods such as centrifugation, filtration (including belt filtration), pressing or other methods that are known to one skilled in the art. While concentration of the culture facilitates later processing steps, the culture should not be excessively dewatered, due to the economic constraints of biofuel production. Accordingly, the concentrated culture should not be dewatered or dried to less than about 10% water content. Obtaining a concentration of 10-30% algal cell dry weight (70-90 % water content) can be achieved by a number of dewatering methods such as belt filtration, centrifugation, filtration, and others known to those skilled in the art.

[0056] After dewatering to provide a concentrated culture, the concentrated culture can be mixed with a partitioning agent. Note that while it is helpful to include a partitioning agent, a partitioning agent is not included in some alternative embodiments of the invention. The partitioning agent is an amphiphilic chemical with oil dispersing properties that functions to reduce the surface tension between lipids and water, thereby allowing the lipids to mix with the aqueous phase. Suitable surfactants, wetting agents, and dispersants can all function as partitioning agents. Examples of partitioning agents include alcohols such as isopropyl alcohol, methanol, sorbitan (e.g., sorbitan monolaurate) and butyl glycol, ketones such as acetone, and weak acids such as succinic acid. Preferably the partitioning agents have a low boiling point to facilitate their later removal, and a relatively small size (e.g., six or few carbons). A preferred partitioning agent is acetone. Suitable partitioning agents also include solvents that are miscible with water and have a dielectric constant equal to or more than 15. Other suitable chemicals have a hydrophilic-lipophilic balance (HLB) (J.T. Davies, Gas/Liquid and Liquid/Liquid Interfaces. Proceedings of 2nd International Congress Surface Activity, Buttersworth, London 1957. 426-438) value between 7 and 9.

[0057] While not intending to be bound by theory, the partitioning agents are believed to act by pulling the algal lipid away from the cell debris into the aqueous phase. In a similar aspect, the partitioning agent acts as a demulsifier where the lipophilic end of the partitioning agent interacts with the lipophilic nonpolar lipid. These interactions between the partitioning agent and the nonpolar lipid block other interactions between the nonpolar lipid and cell debris. The partitioning agent thereby reduces emulsions that can form between the cell debris, water, and lipids during cell disruption. Formation of emulsions is undesirable as emulsions capture lipids in an impure form in combination with other components of the aqueous culture, thereby hindering phase separation and reducing the eventual yield of extracted lipid fraction. The partitioning agent should be completely mixed or dispersed in the mixture when used.

[0058] When a partitioning agent is used, the method of the invention can further include the step of recovering the partitioning agent from the disrupted culture. Recovering the partitioning agent can be beneficial both because it allows the partitioning agent to be reused. The partitioning agent is recovered from the water and biomass through the use of a distillation column, bulk solvent stripper, or any of a number of evaporator systems, such as a rising thin film evaporator (using indirect steam or hot water), or other means known to one skilled in the art. The partitioning agent content of the solution can be further reduced by additional evaporation attained by an evaporator or other means known to one skilled in the art. A simple stripping process can then be used to remove and recover the agent.

[0059] A controlled disruption is then carried out on the concentrated culture, which may or may not also include a partitioning agent at this point. Cell disruption can be achieved using one or more chemical, temperature, or physical methods. Examples of these methods include, but are not limited to, mixing, homogenization, processing with a bead mill, sonic or ultrasonic force, flash freezing, heating, electrochemical treatment, pressure, cavitation, and treatment with cellulose. By "controlled," the inventors intend to exclude highly destructive or "uncontrolled" methods of disrupting microorganisms, such as microfluidizing or homogenizing at high pressures in which the cavitation process breaks the cells into small pieces thus rendering high surface area.

[0060] These controlled methods of breaking the cells are preferably carried out in a manner resulting in the formation of relatively large cell debris particles, for which the total cell debris surface area is decreased. Preferably the large cell debris particles have a size no smaller than 10% of the size of the original cell; i.e., the particle size is defined by the size of the originating algal cell but is around 10% to 100% the size of the original cell. The cell debris includes components of the cell membrane and cell organelles such as phospholipids, proteins, and carbohydrates. Lower mixing speeds, less ultrasonic force, and lower pressure can be used, for example, to provide relatively large cell debris particles. For instance, if a homogenizer is used, thermocouples and pressure gauges tied to a central processor will be used to monitor liquid temperature and pressure controls to ensure that large cell debris particle sizes are achieved and to control the cell breakage percentage. For instance, if a bead mill is used, bead size, bead composition, impeller speed, and residence time in the mill can be controlled to yield large cell debris particle sizes.

[0061] Breaking the microorganisms into very small pieces during cell disruption is undesirable, and conversely there are numerous advantages to obtaining larger particle sizes. Again, while not wishing to be bound by theory, larger particles have less surface area for solvent and lipid to bind to, so recoverable lipid yields are increased. This also improves solvent recovery. Solid-liquid separation is also improved, thereby increasing the purity of the extracted lipid fraction. The resulting extracted lipid f action will also have a lower phospholipid and chlorophyll content that improves yield and oil stability. Using less force also decreases the energy requirements of the process, improving economics. Larger cell particles can also be further processed so that their nutrients can be recycled.

[0062] In some embodiments of the invention, it may be preferable to further treat the concentrated culture before carrying out the controlled disruption step to provide a neutralized, concentrated culture. Prior to cell disruption, lipid interactions with cell debris during disruption can be reduced by adjusting the pH of the mixture to the isoelectric point where the charged moieties in the cell debris and the phospholipids are neutralized. Further, the double layers surrounding the charged moieties can be compressed by adding counter- ions in the form of salts. For example, it may be desirable to adjust the pH of the concentrated culture to the isoelectric point of the mixture prior to cell disruption. Alternately, or in addition, it may also be desirable to adjust the ionic strength of the mixture prior to cell disruption. The isoelectric points for specific molecules have been determined and can be readily identified by those skilled in the art. The charge on molecules and the double layer around the charge on molecules correlated to the zeta potential and can be measured with electrophoresis techniques or specifically tracking the particles through a microscope as they migrate in a voltage field. For example, they can be identified using suitable products from Zeta-Meter®, Inc. Both the pH and the ionic strength are adjusted to neutralize charge and thus reduce the zeta potential to a minimum.

[0063] A neutralized, concentrated culture can be attained through the addition of pH adjusting or ionic strength adjusting chemicals. The partitioning agents help prevent emulsion formation during various stages of the method, such as cell disruption and phase separation. Most cell membranes have a negative charge due to the phosphate portion of the phospholipids and other charged groups contained in the cell's membrane. The presence of this negative charge causes water to orient itself around the charge, creating a water shell or a double layer around the phosphate group of the phospholipid. At the same time, the fatty end of the phospholipid from the cell membrane attracts lipids. These interactions can result in the formation of a stable lipid-water emulsion, which as previously noted results in an eventual decreased yield of extracted lipid fraction. Adjusting the pH varies the amount of hydrogen ions in the aqueous culture, which affects the influence of the negative charge. The double layer can be decreased or compressed by adding salts that contain mono-, di-, and trivalent cations. Examples of suitable salts include NaCl, CaCl₂, and A1C1₃. Likewise, the double layers produced by positive ions can be decreased or compressed by adding salts that contain mono-, di-, and trivalent anions. Examples of suitable salts include NaCl, Na₂S0₄, and NaH₂P0₄. Also a mixture of salts from the ocean such as "sea salt," and sea salt also provides an inexpensive source for many particular salts that are suitable for use in the invention. The decreased size of the water shell that results from salt addition allows for reduction of phospholipid interactions with water, which reduces oil-water-cell debris emulsions and allows for better separation of polar components (e.g., water and partitioning agent) and non-polar components (e.g., the lipid fraction).

[0064] After cell disruption, water or salt water can be added to assist the separation of the partitioning agent from the lipid. Adding water serves to dilute the partitioning agent, thereby reversing the oil dispersing effects of the partitioning agent. While it is preferable to provide a partitioning agent for the controlled disruption, for the reasons provided herein, it is preferable to remove the partitioning agent after the phase separation is carried out, to decrease emulsion formation during all phases. Adding salt to the water assists with the expelling of the lipid from the aqueous phase, and can be helpful for the process even if a partitioning agent was not used. The types of salts that can be added are the same as those described for adjusting the ionic strength. The salts should used in a concentration from about 0.5 to 5 molar, which concentrations of 1-2 molar being preferred.

[0065] After the concentrated culture has been disrupted as described, the extracted lipid fraction can be separated from the other components of the disrupted culture. Separation can be carried out by simply allowing the disrupted culture to settle under gravity, or more preferably by centrifugation, which accelerates the separation. Separation is usually complete within 5 minutes or less, with times of a minute or less being preferred. The extracted lipid fraction can be removed from the other, lower layers of the separated culture by methods known to those skilled in the art, such as siphoning or decanting. A preferred method for use with continuous centrifugation is the use of a "weir," which is a small dam present at the top of the centrifugation device over which the uppermost layer of liquid, which will be the extracted lipid faction, continuously overflows for collection.

[0066] The amount of extracted lipid fraction obtained during separation can vary from about 60% to about 95%, compared to the levels of lipid originally present in the cell, depending on the particulars of the method used. Embodiments of the method can provide about 80% or more, about 90% or more, or about 95% or more of the lipid in the extracted lipid fraction. The extracted lipid fraction includes primarily monoglycerides, diglycerides, and triglycerides, though other lipids can also be present. For example, in one embodiment, the invention provides lipids including greater than 60% triglycerides, diglycerides, monoglycerides; less than 10% free fatty acids; and less than 10% phospholipids. The lipid fraction obtained is typically immediately sequestered for storage, but alternately the lipid can be immediately directed to other uses or for immediate further purification. In comparison to traditional extraction processes with hexane or other generally biocompatible non-polar solvents, a non-polar solvent is not present in the disrupted culture and a distillation step is not required to recover the extracted lipid fraction. Since the extracted lipid fraction is not exposed to high temperatures in a distillation process, thermal degradation of the lipids can be avoided. Polar lipids, which are generally considered to be contaminants, primarily remain in the aqueous phase.

[0067] The extracted lipid fraction obtained using the method of the invention can have high purity and other beneficial characteristics. For example, the extracted lipid fraction can include high levels of triglycerides. For example, the extracted lipid fraction can include about 50% or more of triglycerides by weight, or can include about 70% or more triglycerides by weight. It is preferable to include high levels of triglycerides, since they are good biofuel precursors. The extracted lipid fraction can also include low levels of impurities. Impurities include phospholipids and chlorophyll pigment. For example, the extracted lipid fraction can include about 10% or less phospholipids, more preferably about 5% or less phospholipids, or in further embodiments, about 2% or less phospholipids. Alternately, or in addition, the extracted lipid fraction can include about 5% or less chlorophyll, about 3% or less chlorophyll, or about 1% or less chlorophyll.

[0068] After phase separation, the culture will include three different phases: a biomass concentrate (biomeal), an aqueous mixture, and an extracted lipid fraction. "Biomeal" is disrupted or lysed cellular biomass where the lipid has been largely removed, and is synonymous herein with lipid extracted algae (LEA). The main product desired at this point is the extracted lipid fraction. However, proteins and carbohydrates can be recovered from the biomeal by heating with ammonia to a basic pH, filtering, and cooling down to an acidic pH to precipitate the protein. If desired, any remaining solids could then be sent to a desolventizer or stripper to recover the partitioning agent. This biomass can also be sent to an anaerobic digestor to create methane gas or used for any number of other secondary product production methods. Such processes could be pyrolysis, bioproduct extraction, fermentation, hydrolysis, and etc.

[0069] As described previously herein, the method of obtaining an extracted lipid fraction can be carried out either with or without a partitioning agent. While use of a partitioning agent is helpful, high yields of extracted lipid fraction can be obtained even in the absence of a partitioning agent. When carried out in this fashion, the method includes growing the microorganisms in aqueous culture under conditions to provide microorgamsms having a high amount of lipids; e.g., greater than about 30%, greater than about 40%, greater than about 45%, or greater than 50% of their dry weight being lipids. The high amount of lipid can result from use of, for example, a Heteroboost™ method involving differential growth conditions between the first and second stage. The culture of microorganisms is then dewatered to provide a concentrated culture having 15-30% solids by weight, while still including a substantial amount of water. The concentrated culture is then subject to a controlled disruption of to provide a disrupted culture; and an extracted lipid fraction is then separated from the disrupted culture that includes about 50%, about 60%, or about 70% or more of the lipid present in the microorganisms before extraction. This method can also include the step of diluting the disrupted culture with salt water, as described herein.

[0070] As already described herein, the present invention provides a method for obtaining an extracted lipid fraction from microorgamsms (e.g., algae or yeast). Another aspect of the invention provides a system for carrying out this method. A schematic representation of a system of the present invention is provided by Figure 2. The system represents the combination of various apparatus that can be used to carry out the steps of the invention, and is a system for producing lipid from microorgamsms. In some embodiments, the apparatus are directly in communication with one another; while in other embodiments it may be necessary to transport the output from one apparatus for use as an input for the apparatus used in the next step of the invention.

[0071] In a preferred embodiment of the invention, the system includes an apparatus for growing microorganisms 100 under conditions to provide high lipid levels. For example, the system can include an apparatus configured for growing algae under Heteroboost™ conditions using differential growth conditions including a first culturing apparatus for culturing microorgamsms under phototrophic, mixotrophic, or a combination of phototrophic and mixotrophic conditions and a second culturing apparatus 110 that receives cultured microorgamsms from the first culturing apparatus for culturing the microorgamsms under different conditions selected from mixotrophic or heterotrophic conditions, or a combination thereof.

[0072] Apparatii for culturing microorganisms under phototrophic conditions are well-known to those skilled in the art. Examples of growth environments that can be used for phototrophic growth include bioreactors, open ponds having various shapes and configurations such as raceway ponds, and greenhouse environments such as enclosed pools. A raceway-type pond, which is commonly used, is divided into a rectangular grid, with each rectangle containing one channel in the shape of an oval, like an automotive raceway circuit. Each rectangle also typically includes a paddlewheel to provide continuous water flow around the circuit. Apparatii for phototrophic growth include air circulation, possible supplemented with industrial sources, to provide carbon dioxide, an aqueous culture for suspending the microorganisms, and sufficient light to support photosynthetic growth. Phototrophic growth apparatus are useful for growing a large mass of photosynthetic microorganisms quickly and cheaply as possible.

[0073] Once a sufficient amount of growth has occurred in the first growth apparatus 100, the microorganisms can be transferred to a second growth apparatus 110 for carrying out either mixotrophic and/or heterotrophic growth. If the first growth apparatus uses mixotrophic growth, the second apparatus should use heterotrophic growth, or at least mixotrophic growth with a much higher aspect of heterotrophic growth than what was used in the first growth apparatus. Preferably, the microorganisms are substantially concentrated before being introduced into the second growth apparatus.

[0074] Apparatus for carrying out heterotrophic growth are also well known in the art. Preferably the apparatus for carrying out heterotrophic growth is an enclosed, opaque bioreactor. In a heterotrophic growth bioreactor, access to sunlight is prevented and carbon sources other than carbon dioxide are used. Examples of alternate carbon sources include glycerol and various sugars. Other nutrients such as magnesium, potassium, and nitrogen may be varied to encourage growth that leads to high lipid levels. A heterotrophic growth apparatus includes features and controls that allow H, temperature, and dissolved oxygen to be measured and controlled. It also allows effective mass transfer of oxygen and sugar into the cells, and maintenance of C0₂ egress from the cells. This can be done with bubble column mixing or impeller mixing or a combination of the two.

[0075] Apparatus for carrying out mixotrophic growth can be used in either the first or second stage of growth of the microorganisms. An apparatus for mixotrophic growth provides alternate carbon sources, but also provides carbon dioxide and access to sunlight. For example, an open pond provided with an alternate carbon source such as glycerol represents an example of a mixotrophic growth apparatus, as does a bioreactor that includes access to light and carbon dioxide as well as a sugar growth source. [0076] The system of the invention also includes a dewatering apparatus 120 to remove a portion of the water from the oleaginous culture to provide a concentrated culture. Dewatering can be carried out between the first and second stages of microorganism growth, and can also be carried out after microorganism growth is complete. A dewatering device is capable of splitting the culture into two phases; a heavy phase that the bulk of the microorganisms, and a light phase. Dewatering is achieved by any of a number of methods including but not limited to centrifugation, flocculation, settling, filtration {e.g., ultrafiltration or microfiltration), pressing or other methods that are known to one skilled in the art. Specific commercial sources for suitable centrifuges include Alfa Laval®, GEA Westfalia®, and Flottweg®. Plate and frame filters can be obtained from Andritz®, and ultra and microfilters can be obtained from GE®, Pall®, and Koch Membrane Systems®.

[0077] The system for producing an extracted lipid fraction from microorganisms also includes a disruption apparatus 130. The disruption apparatus is used for disrupting the cell structure of the microorganisms in the concentrated culture to provide a disrupted culture. In particular, it is preferable to carry out the disruption in a controlled manner to avoid fragmenting the cells into very small pieces. Examples of apparatus suitable for carrying out controlled disruption are homogenizers, bead mills, presses, microfluidizers, and cavitation apparatus. Bead mills, can be obtained from Netzsch® Premier Technologies, Hochmeyer® Equipment Corp., Glen Mills® Inc., Union Process®, Buhler®; homogenizers, which can be obtained from GEA® and Niro Soavi®, and Microfluidizers which can be obtained from Microfluidics® Corp.

[0078] The system also includes a separation apparatus 140. The separation apparatus is used to separate a substantial portion of the lipid from the disrupted culture. The separation apparatus can be as simple as a container in which the disrupted culture is allowed to settle by gravity such as a settling tank. However, preferable apparatus also include equipment to enable fractions such as the extracted lipid fraction to be withdrawn, and centrifuge equipment to increase the speed of the separation. In some embodiments, two centrifuges can be used in tandem, where one centrifuge removes the biomeal (clarifier centrifuge) and the other centrifuge separates the lipid fraction from the aqueous phase (separator centrifuge). A tricanter centrifuge can also be used to separate all three phases in a single process or a decanter can be used to just separate the lipid fraction from the aqueous phase and biomeal. Suitable centrifuges can be obtained from Alfa Laval®, GEA Westfalia®, and Flottweg®. [0079] Figure 3 provides an additional schematic representation of an additional embodiment of system for producing lipid from microorganisms. The system includes a first apparatus for growing microorganisms 100 such as an open pond, a covered pond, or closed photobioreactors for phototrophic or mixotrophic growth. The system also includes a first dewatering apparatus 105 to concentrate culture from the apparatus for growing microorganisms 100. The system then includes a second growth apparatus 110 that provides either mixotrophic or heterotrophic growth conditions. This can be as elaborate as a standard fermentor or as simple as an enclosed pond. For mixotrophic growth this would require a light source so a closed photobioreactor could be a preferred growth apparatus at this stage. A surge tank 115 can also be included prior to aqueous extraction. A second dewatering apparatus 120 is included to concentrate culture to greater than 15% solids by weight. A first mix tank 125 is also included to add the partitioning agent as well pH and ionic strength adjusting chemicals. A cell disruption apparatus 130 is included, which directs its output to a second mix tank 135 to add water to reduce the effects of the partitioning agent and start the phase separation process. A separation apparatus 140 is included such as centrifuge. A storage tank 150 can be included to hold the extracted lipid fraction. A distillation system 160 can be included to recover partitioning agent from biomeal and/or aqueous phase, and a partition agent tank 170 can be included to hold recovered partitioning agent until it is ready for delivery to the second mixing tank 135.

[0080] The following examples are included for purposes of illustration and are not intended to limit the scope of the invention.

EXAMPLES

Example 1: Aqueous extraction of algal oil from Chlorella protothecoides KRT1007.

[0081] The following is an example of a system implementing the present application. This process can be run as a steady state continuous flow process. The algae are first to be grown in open or closed photobioreactors. The broth contained in these ponds consists of roughly 0.10 % solids and 0.07% biomass. Approximately 1-30% of the biomass is lipid.

[0082] A portion of the pond's volume will then be dewatered to produce two phases. The heavy phase, consisting of roughly 5.2% dissolved solids and 5.0% biomass, will continue to the Heteroboost™ process where the cells are grown heterotrophically in a bioreactor on sugar substrates. The light phase, consisting of 2.0% dissolved solids, will be recycled back into the pond.

[0083] In the Heteroboost™ process, the algal cells will mature and accumulate oil. It is important that during this transfer to the Heteroboost™ process that: (1) shear is minimized; (2) the culture is not subjected to chemical shock; and (3) a proper temperature range is maintained. A nutrient rich medium will be introduced to the culture to enhance maturation and lipid accumulation. This will provide a high carbon content source (such as sugar), essential amino acids, but shift the balance of nutrients to favor lipid accumulation rather than additional cell growth. For example, a shift up in the C N ratio will force the cells to limit cell growth. This mimics the nitrogen starvation seen in the open ponds and is used to stimulate lipid accumulation. Alternative stressors and inducers are also possible. This process can be run in a continuous or batch mode. In a continuous mode, every 24 hours approximately 25% of the total volume of the growing culture is removed and processed.

[0084] After lipid accumulation, the resultant broth is concentrated by dewatering with a belt filter press, centrifuge, plate and frame filter, or other devices known to those skilled in the art. At this point in the process, the algal culture is dewatered to approximately 80 g L to 300 g/L of dry weight algal mass.

[0085] After dewatering, isopropyl alcohol, a partitioning agent, is then added to the heavy phase. This can be but is not limited to a ratio of about 1:1 ΓΡΑ to concentrate. After addition of ΓΡΑ, concentrated NaOH is used to adjust the broth to the isoelectric point which is near a pH of 7. In addition, ammonium sulfate is added to compress the double layers of any remaining charged moieties. At this point, roughly 32% of the broth is solids and the algal cells, which represent a majority of those solids, contain between about 40-70%) lipid. Next, the cells are disrupted using a homogenizer. The process will operate at 50 - 60 °C and a pressure of between about 1,000 and 1,400 bar - this will vary with microorganism strain being used. The temperature and pressure will be closely monitored using thermocouples and pressure gauges, respectively, to ensure that large particle sizes are obtained.

[0086] To the slurry of disrupted cells, liberated lipid, water, and DP A, additional water is added to induce a phase separation. Approximately, but not limited to, a ratio of about 1:1 water to slurry can be gently mixed into the slurry. The liberated oil will begin to phase separate from the aqueous phase; this separation is accelerated by centrifugation. The result is a light layer of algal lipid, a middle layer of mostly water and IP A, and a heavy layer of lipid extracted algae. To recover the ΓΡΑ for reuse, the water and IPA can be processed with a fractional distillation column. To reduce costs, the purity of the output of the ΓΡΑ at the output of the distillation column can be the azeotrope or 85% ΓΡΑ and 15% water. The resultant ΓΡΑ-water mixture can be used as the partitioning agent for future extractions.

Example 2. Extraction with ethanol and acetone with test tube beat beating

[0087] Amphiphilic solvents such as ethanol or acetone can be used as the partitioning agent. Algal oil was successfully extracted and separated from algal cells by carrying out the following steps.

1. Grew algal biomass where the algal cell mass contains 55% lipid by dry weight.

2. Concentrated the cells to 300 g/L

3. Took a 4 mL sample of the algal concentrate and add 3.6 mL of ethanol or acetone.

4. Adjusted the pH to 7.

5. Bead beat for 1 hour with three 6 mm beads and a 1 mL volume of 0.5 mm beads.

This was performed in a vortex mixer.

6. Added 5 mL of water or 1M NaCl

7. Centrifuged for 10 min at 2000 rpm.

[0088] In Figure 4, the tube labeled "Eth" shows the results where ethanol was used as the partitioning agent, and the tube labeled "AC" shows the results where acetone was used as the partitioning agent. The extracted algal oil has phase separated and is the top layer in the test tubes.

Example 3. Extraction with isopropyl alcohol with a Hockmeyer Immersion Mill

1. Grew algal biomass where the algal cell mass contains 55% lipid by dry weight.

2. Concentrated the cells to 450 g/L

3. To 600 g of concentrate 444 mL of isopropyl alcohol was added and mixed into a slurry.

4. The initial temperature and pH of the slurry were 32° C and 5.8, respectively.

5. The slurry was inserted into a Hockmeyer immersion mill for 30 minutes with an impeller rate of 5,000 rpm. The bead size was 0.4 mm.

6. The final temperature and pH were 56.5° C and 6.75, respectively. 7. A 10 mL sample of the disrupted biomass was collected where 15 mL of deionized water was added.

8. The disrupted biomass slurry and water were centrifuged for 15 min at 4,000 rpm.

[0089] Figure 5 illustrates the extracted algal oil from the cells where the oil has phase separated and was evident at the top of the test tube.

Example 3. Addition of salt water to induce phase separation

[0090] To reduce emulsion formation and thus oil lost to the emulsion layer, salt was added to the water prior to phase separation. The following procedure was followed:

1. Grew the K T 1007 phototrophically in open ponds, then harvest and induce algal lipid accumulation in Phycal's Heteroboost™ process

2. Concentrated the resultant broth to 350 g/L

3. Adjusted pH of biomass slurry to about 7.0.

4. Added five 6 mm glass beads and 1 mL 0.5 mm silica beads to 50 mL centrifuge tubes with biomass.

5. Added 4 mL of pH adjusted biomass to 50 mL centrifuge tubes from step 4.

6. Added PA to pH adjusted biomass slurry to produce a ratio of 1 :2 (Biomass slurry:IPA)

7. Vortexed on high for one hour

8. Added salt solution or DI water at a 1 : 1 biomass slurry to solution/water ratio

9. Mixed gently 5 times

10. Centrifuged 5 minutes at 1,800 RPM

[0091] Table 1 and Figure 6 represent the resultant centrifuge tubes after step 10.

Table 1

Example 4. Controlled disruption

[0092] Cell disruption is a critical step in this extraction process. This example illustrates proper and improper methods for performing the controlled cell disruption to minimize cell debris surface area and to maximize algal lipid recovery. For Figures 7 through 10, the same algal biomass was processed separately in either a Biospec® bead beater or a microfluidizer. Phycal's batch number for the sample was 20111108-K T1009 Ferm A, which was prepared by growing Chlorella strain KRT1009 phototrophically in an open photobioreactor which was then harvested, concentrated, and inserted into Phycal's Heteroboost™ process

1 Obtained two 100 mL samples of concentrate from the Heteroboost™ reactor.

2 Added 1:1 EPA.

3 The first sample was bead beat and sampled every 15 minutes of which a picture was taken for a total 60 minutes; the second sample was processed in a microfluidizer and sampled every pass where a picture was taken for a total of 4 passes.

Figure 7 provides pictures taken of the disrupted cells

[0093] With every pass through the microfluidizer more cells were disrupted, but the way they were disrupted hinders the lipid extraction efficiency. As the cells were disrupted from left to right, the total cell debris that is shown is reduced. This illustrates that the cells were exploding due to the cavitations encountered in the microfluidizer and producing cell debris so small that they were not visible in the microscope. The parameters used to disrupt the cells through the microfluidizer produced much more surface area and can be shown in the amount of lipid extracted in the table below. In the bead beater, cells were cracked in a more controlled manner, rendering large pieces of cell debris at 15 minutes of processing time. This particular extraction time of 15 minutes provided the most lipid recovered compared with other times as shown. As the cells were disrupted for 30 minutes and 45 minutes, most of the cells were cracked releasing the lipid, but the cell debris was continually broken into small pieces which increased the surface area of the cell debris. Thus as the cells were disrupted for longer periods of time, the lipid recovery was reduced.

[0094] To obtain the data shown in Figure 8, the 10 mL samples from the disruption methods and times above were mixed with 15 mL of 2M NaCl, then centrifuged for 10 min at 4000 rpm. The algal lipid separated as the top phase and was removed and weighed.

[0095] As evident from the data, the most algal lipid was recovered at 15 minutes with the bead beater. As the algae spent more time in the bead beater, the cell debris surface area was increased and less lipid was recovered. As for the microfluidizer, more lipid was recovered with every pass, but due to the small cell debris created by the conditions of the cavitation disruption mechanism, only 65% of the lipid was recovered from the fourth pass in the microfluidizer compared to the lipid recovered from 15 minutes in the bead mill. See Figure 9, which shows the oil recovery from each pass (increasing from left to right) in the microfluidizer, and Figure 10, which shows oil recovery from each 15 minute sample (increasing from left to right) in the bead beater for further information on this recovery.

[0096] This example does not illustrate that cell disruption in a bead beater was better than cell disruption in a microfluidizer. It demonstrates that the cell disruption step must be tempered or more precisely controlled to control the cell debris surface area. By reducing the microfluidizer pressure and adjusting the concentration of algae being processed, similar results to the bead beater can be obtained. Therefore, a variety of cell disruption equipment can be used for the present invention.

Example 5. Extraction using water as the partitioning agent

Water can potentially be used as the partitioning agent. This example illustrates how water or salt water with the addition of heat can reduce the viscosities of the algal lipid and induce a phase separation to recover the lipid after cell disruption. The below procedure is for lipid extraction with salt water without heat.

1. Grow the KRT 1007 phototrophically in open ponds, then harvest and induce algal lipid accumulation in Phycal's Heteroboost™ process.

2. Concentrate the resultant broth to 350 g/L.

3. Adjust the pH of biomass slurry to about 7.0.

4. Dilute with a 1 : 1 ratio of 1M NaCl solution.

5. Perform cell disruption with beads and vortexor in centrifuge tube.

6. Add more 1M NaCl and perform phase separation in centrifuge tube.

7. Recover the light phase oil layer.

The below procedure is for lipid extraction with salt water with heat before and after the cell disruption step.

1. Grow the KRT 1007 phototrophically in open ponds, then harvest and induce algal lipid accumulation in Phycal's Heteroboost™ process.

2. Concentrate the resultant broth to 350 g/L.

3. Adjust the pH of biomass slurry to about 7.0. 4. Dilute with a 1 : 1 ratio of 1M NaCl solution.

5. Heat to 70° C (Heat Before).

6. Perform cell disruption with beads and vortex in centrifuge tube.

7. Heat to 70° C (Heat After).

8. Add more 1M NaCl and perform phase separation in centrifuge tube.

9. Recover the light phase oil layer.

Table 2

[0097] When a 1M NaCl solution is added as the partitioning agent then the mixture is heated to 70° C prior to cell disruption or the mixture is heated to 70° C after cell disruption, the lipid recovery yield from the cells is increased. In particular, when heat is added after the cell disruption during the phase separation 100% of the lipid compared to a Bligh and Dyer extraction was recovered. This method eliminates solvents as the partitioning agent and thus eliminates the need to recover them and the costs of distillation.

Example 6. Reduction of chlorophyll during Heteroboost™ process

[0098] In addition to Phycal's algae growth methods that produce algae containing greater than 45% nonpolar lipid as a percentage of the cellular dry weight, the Heteroboost™ process also reduces the chlorophyll content of the cells. Since the chlorophyll is degraded due to the nonphototrophic growth, the chlorophyll does not get extracted with the lipid and therefore produces an algal oil product low in chlorophyll. The chlorophyll degradation could be seen visually, where Chlorella protothecoides was grown in open pond photobioreactors, then transferred to a bioreactor where it was grown heterotrophically. During this transformation the chlorophyll is catabolized where the phytol chain is removed, the magnesium ion is released and the chlorophyll tetrapyrole ring structure is opened. Oil from algae that have undergone the Heteroboost™ process does not exhibit a green color from chlorophyll that is typical of algal oil extracted via traditional methods using dried biomass and hexane in a soxhlet extractor. Typically, oil from the Heteroboost process possesses less than 5% chlorophyll content.

Example 7. Removal of partitioning agent from the aqueous phase.

[0099] Since this process describes an aqueous extraction where 50 - 80% of the algal concentrate can be water, the partitioning agent is mixed with this slurry, diluted, and must be recovered for reuse. Typically, the boiling point to the partitioning agent is different enough from water to be able to fractionate the partitioning agent using distillation methods. Membrane separation is also a feasible process to recover the partitioning agents.

[00100] The complete disclosure of all patents, patent applications, and publications, and electronically available material cited herein are incorporated by reference. The foregoing detailed description and examples have been given for clarity of understanding only. No unnecessary limitations are to be understood therefrom. The invention is not limited to the exact details shown and described, for variations obvious to one skilled in the art will be included within the invention defined by the claims.

Claims

CLAIMS What is claimed is:

1. A method of obtaining an extracted lipid fraction from microorganisms, comprising: growing the microorganisms in aqueous culture under conditions to provide microorganisms having greater than about 45% of their dry weight being lipids;

dewatering the culture of microorganisms to provide a concentrated culture having greater than about 15% solids by weight;

mixing the concentrated culture with a partitioning agent;

conducting a controlled disruption of the concentrated culture including a partitioning agent to provide a disrupted culture; and

separating an extracted lipid fraction from the disrupted culture by phase separation.

2. The method of claim 1, wherein growing the microorganisms in culture comprises: a first stage that includes phototrophic, mixotrophic, or a combination of phototrophic and mixotrophic conditions, followed by

a second stage that includes different conditions selected from mixotrophic or heterotrophic conditions, or a combination thereof.

3. The method of claims 1 or 2, wherein the microorganisms are algae.

4. The method of claim 3, wherein the algae are from the genus Chlorella.

5. The method of claim 4, wherein the algae are selected from the species Chlorella protothecoid.es, C, kessleri, C. vulgaris, C. sorokiniana, C. zofingiensis, C. minutissima, C. regularis, and C. variabilis.

6. The method of claims 1 or 2, wherein the concentrated culture and the disrupted culture comprise at least about 10% water.

7. The method of claims 1 or 2, wherein the partitioning agent is an amphiphilic chemical with oil dispersing properties.

8. The method of claims 1 or 2, wherein the partitioning agent is acetone.

9. The method of claims 1 or 2, further comprising adjusting the pH of the concentrated culture to the isoelectric point prior to cell disruption.

10. The method of claims 1 or 2, further comprising adjusting the ionic strength of the concentrated culture prior to cell disruption.

11. The method of claims 1 or 2, further comprising the step of diluting the disrupted culture with water or salt water to decrease the effect of the partitioning agent and better affect separation of the lipid fraction.

12. The method of claims 1 or 2, further comprising the step of recovering the partitioning agent from the disrupted culture using evaporation, stripping, or a combination thereof.

13. The method of claims 1 or 2, wherein the extracted lipid fraction comprises at least about 70% triglycerides.

14. The method of claims 1 or 2, wherein the extracted lipid fraction comprises about 10% or less phospholipids.

15. The method of claims 1 or 2, wherein the extracted lipid fraction comprises about 5% or less chlorophyll pigment.

16. A method of obtaining an extracted lipid fraction from microorganisms, comprising: growing the microorganisms in aqueous culture under conditions to provide microorganisms having greater than about 45% of their dry weight being lipids;

dewatering the culture of microorganisms to provide a concentrated culture having greater than about 15% solids by weight while still including a substantial amount of water; conducting a controlled disruption of the concentrated culture to provide a disrupted culture; separating an extracted lipid fraction from the disrupted culture that includes about 60% or more of the lipid present in the microorganisms before extraction.

17. The method of claim 16, further comprising the step of diluting the disrupted culture with salt water.

18. The method of claim 16, wherein growing the microorganisms in culture comprises: a first stage that includes phototrophic, mixotrophic, or a combination of phototrophic and mixotrophic conditions, followed by

a second stage that includes different conditions selected from mixotrophic or heterotrophic conditions, or a combination thereof.

19. A system for producing lipid from microorganisms, comprising:

a first culturing apparatus for culturing microorganisms under phototrophic, mixotrophic, or a combination of phototrophic and mixotrophic conditions;

a second culturing apparatus that receives cultured microorganisms from the first culturing apparatus for culturing the microorganisms under different conditions selected from mixotrophic or heterotrophic conditions, or a combination thereof to provide an oleaginous culture of microorganisms having greater than about 45% of their dry weight being lipids;

a dewatering apparatus to remove a portion of the water from the oleaginous culture to provide a concentrated culture having greater than about 15% solids by weight;

a disruption apparatus for disrupting the cell structure of the microorganisms in the concentrated culture to provide a disrupted culture; and

a separation apparatus for separating a substantial portion of the lipid from the disrupted culture.

20. The system of claim 19, further comprising a concentrating apparatus to concentrate the microorganism culture from the first culturing apparatus before it is provided to the second culturing apparatus.