WO2013055887A1 - Microalgae culture and harvest - Google Patents

Microalgae culture and harvest Download PDF

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WO2013055887A1
WO2013055887A1 PCT/US2012/059707 US2012059707W WO2013055887A1 WO 2013055887 A1 WO2013055887 A1 WO 2013055887A1 US 2012059707 W US2012059707 W US 2012059707W WO 2013055887 A1 WO2013055887 A1 WO 2013055887A1
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microalgae
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method
culture
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Bo Hu
Rongsheng R. RUAN
Jianguo Zhang
Wenguang Zhou
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Regents Of The University Of Minnesota
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    • C02F3/32Biological treatment of water, waste water, or sewage characterised by the animals or plants used, e.g. algae
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    • C12P7/64Fats; Fatty oils; Ester-type waxes; Higher fatty acids, i.e. having at least seven carbon atoms in an unbroken chain bound to a carboxyl group; Oxidised oils or fats
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    • C12P7/64Fats; Fatty oils; Ester-type waxes; Higher fatty acids, i.e. having at least seven carbon atoms in an unbroken chain bound to a carboxyl group; Oxidised oils or fats
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    • C12P7/6445Glycerides
    • C12P7/6463Glycerides obtained from glyceride producing microorganisms, e.g. single cell oil
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Abstract

We disclose herein a method for harvesting cultivated microalgae. In general, the method includes adding at least one filamentous fungus to a culture of microalgae, growing the culture of microalgae under conditions effective to produce a co-culture of the microalgae and the filamentous fungus and to form cell pellets; and harvesting the cell pellets.

Description

MICROALGAE CULTURE AND HARVEST

CROSS-REFERENCE TO RELATED APPLICATION This application claims priority to U.S. Provisional Patent Application Serial No. 61/547,177, filed October 14, 2011.

SUMMARY

In one aspect, this disclosure describes a method for harvesting cultivated microalgae. In general, the method includes adding at least one filamentous fungus to a culture of microalgae, growing the culture of microalgae under conditions effective to produce a co- culture of the microalgae and the filamentous fungus and to form cell pellets; and harvesting the cell pellets.

In some embodiments, the method may be performed using autotrophic algae. In other embodiments, the method may be performed using heterotrophic microalgae.

In some embodiments, the filamentous fungus may be added to the microalgae culture as at least one spore. In other embodiments, the filamentous fungus may be added to the microalgae culture as at least one fungal cell.

In some embodiments, the filamentous fungus cell can express cellulase.

In some embodiments, the filamentous fungus cell can produce oil. In some of these embodiments, the oil produced by the filamentous fungus cell may also be produced by the microalgae.

In some embodiments, harvesting the pellets can include filtering at least a portion of the pellets from the culture.

In some embodiments, the method can further include extracting a bioproduct from the harvested pellets. In some of these embodiments, the bioproducts can include an oil.

Ion another aspect, this disclosure describes a method for treating wastewater. The method generally includes adding at least one filamentous fungus to a culture of microalgae, wherein the culture comprises wastewater, and growing the culture of microalgae under conditions effective to produce a co-culture of the microalgae and the filamentous fungus comprising cell pellets that comprises the microalgae and the filamentous fungus.

The above summary is not intended to describe each disclosed embodiment or every implementation of the present invention. The description that follows more particularly exemplifies illustrative embodiments. In several places throughout the application, guidance is provided through lists of examples, which examples can be used in various combinations. In each instance, the recited list serves only as a representative group and should not be interpreted as an exclusive list.

BRIEF DESCRIPTION OF THE FIGURES FIG. 1. Pelletization of Mucor circinelloides by adding CaC03 (A) and by only adjusting pH during cell growth (B).

FIG. 2. Pelletization of Chlorella vulgaris with assistance from filamentous fungi. (A) Seed culture; (B) Two days after inoculation with fungal spores; (C) Microscopic image of pellet formed by co-culture of microalgae and fungi under lOOx magnification.

FIG. 3. Co-culture of autotrophic microalgae with cellulase-producing fungi.

FIG. 4. Co-culture of Aspergillus niger with C. vulgaris at different initial pH

(autotrophic culture medium A, fungal spore inoculation 7.6 10 6 L, 27°C, 150 rpm, 3 days), measured as: (A) microalgae cell density; (B) microalgae harvest ratio.

FIG. 5. Co-culture of A. niger with C. vulgaris at different initial inoculation level (autotrophic culture medium A, initial pH 4.0, 27°C, 150 rpm, 3 days), measured as: (A) microalgae cell density; (B) microalgae harvest ratio.

FIG. 6. Co-culture of A. niger with C. vulgaris at different initial microalgae concentration (autotrophic culture medium A, initial pH 4.0, fungal spore inoculation

7.6 10 6/L, 27°C, 150 rpm, 3 days), measured as: (A) microalgae cell density; (B) microalgae harvest ratio.

FIG. 7. Co-culture of mixotrotrophic microalgae with oleaginous fungi.

FIG. 8. Co-culture of A. niger with C. vulgaris at different initial pH (heterotrophic culture medium B, fungal spore inoculation 7.6 10 6/L, 27°C, 150 rpm, 2 days), measured as: (A) microalgae cell density; (B) microalgae harvest ratio. FIG. 9. Co-culture of A. niger with C. vulgaris at different initial inoculation level (heterotrophic culture medium B, initial pH 5.0, 27°C, 150 rpm, 2 days), measured as: (A) microalgae cell density; (B) microalgae harvest ratio.

FIG. 10. Co-culture of A. niger with C. vulgaris at different glucose and nitrogen concentration (heterotrophic culture medium B, initial pH 5.0, fungal spore inoculation 7.6 10 6/L, 27°C, 150 rpm, 2 days), measured as microalgae harvest ratio.

FIG. 11. Co-culture of A. niger with C. vulgaris at different initial microalgae concentration (heterotrophic culture medium B, initial pH 4.0, fungal spore inoculation 7.6 10 6/L, 27°C, 150 rpm, 2 days), measured as: (A) microalgae cell density; (B) microalgae harvest ratio.

FIG. 12. Pelletization of various fungi and exemplary fungus-algae combinations.

FIG. 13. Nutrient removal profile in fungi-algae pellets culture system grown on centrate wastewater. A, Ammonia removal profile; B, Total phosphorus removal profile; C, COD removal profile; D, Total Nitrogen removal profile.

FIG. 14. Nutrient removal profile in fungi-algae pellets culture system grown on 20- fold hog manure wastewater. A, Ammonia removal profile; B, COD removal profile; C, Total phosphorus removal profile; D, Total Nitrogen removal profile.

FIG. 15. The color changes of 20x hog manure before and after treatment by fungi- algae pellets overnight. 1, control; 2, overnight treatment (15 hours).

DETAILED DESCRIPTION OF ILLUSTRATIVE EMBODIMENTS

Cultivating microalgae has many commercial applications including, for example, biofuel production and/or wastewater treatment. We describe herein a process that generally involves pelletizing microalgae. Typically, microalgae cells are small and grow individually, which makes harvesting the cells difficult and contributes to 20-30% of the total cost of biomass production. Pelletizing microalgae cells— i.e., inducing the microalgae cells to cluster with one another— during microalgae cultivation can result in more efficient and economical harvesting than harvesting individual cells. We have established an efficient method to introduce a technical platform of cell pelletization to the cultivation of oleaginous microalgae, most of which are not filamentous, by co-culturing filamentous fungi with microalgae so that microalgae cells can be co-pelletized into fungal pellets for easier harvest. This new concept of pelletized cell cultivation will have a direct impact and application on the use of microalgae for, for example, biofuel production and/or wastewater treatment, and it can provide effective, cost-efficient, and environmentally sound industrial processes.

For example, producing biofuels and bioproducts via microalgae is promising. New technical processes must be developed, however, in order to capitalize on the economically feasible potential of accumulating bioproducts and biofuel inside microalgae biomass. For instance, many microalgae (e.g., C. vulgaris) are capable of accumulating a high content of lipids that can be converted to different forms of "drop-in" fuels such as biodiesel (Fakas and Papanicolaou, 2009 Biomass & Bioenergy 33(4):573-580; Xia et al., 2011 Biotech biofuels 4:15; Heredia- Arroyo et al., 2011 Biomass & Bioenergy 35(5):2245-2253). Microalgae can rapidly accumulate lipids, which fit the industrial needs for biofuel production, with either autotrophic growth or heterotrophic growth mode. For the autotrophic growth mode, microalgae assimilate the carbon dioxide from the atmosphere as their carbon source, and sunlight in most cases as their energy source. The heterotrophic growth of microalgae cells uses organic carbon, for instance glucose, to support their carbon and energy need.

Past studies for large-scale cultivation of algae relied on open-pond systems, which made it difficult to successfully cultivate algae due to the high downstream processing costs. Open-pond cultures are typically commercially viable for producing certain value-added health food supplements such as feed and reagents (Chisti, 2007 Biotechnol Adv 25(3):294- 306). Photobioreactors may achieve higher productivity and can maintain monoculture of algae. The unit cost of microalgae production in enclosed photobioreactors, however, can be much higher than those achievable in open-pond cultures despite photoreactors' higher biomass concentration and better control of culture parameters (Lee, 2001 J Appl Phycology 13(4):307-315).

The algae cell harvest from cultivation broth has been an obstacle for the algae-to-fuel approach. Microalgae cell harvest can be technically challenging, especially considering the low densities (typically in the range of 0.3 g/L to 5 g/L) of microalgae cells, the small size of the oleaginous algal cells (typically in the range of 2 μηι to 40 μπι in diameter), and their similar density to water (Li and Horsman, 2008 Biotechnol Prog 24(4):815-820). Oleaginous microalgae cells are usually suspended in water and do not easily settle by natural gravity force due to their negative surface charges. The recovery of microalgae biomass generally requires one or more solid-liquid separation steps, and can account for 20-30% of the total costs of production (Uduman et al., 2010 J Renew Sustain Energy 2(1)).

Preferred methods for harvesting microalgae cells from cultivation broth can be influenced by, for example, the characteristics of the microalgae such as, for example, size and density (Olaizola, 2003 Biomol Engineer 20(4-6):459-466). Moreover, conventional harvesting methods often involve a separate step after the cell cultivation. Current harvest approaches— e.g., flocculation, flotation, centrifugal sedimentation, and/or filtration— have limitations for effective, cost-efficient production of biofuel (Shelef et al., 1984 "Microalgae harvesting and processing: a literature review," Technion Research and Development Foundation Ltd.: Haifa, Israel, 70 pages). For instance, flotation methods, based on trapping algae cells using dispersed micro-air bubbles, is limited in its technical and economic viability. Most conventional and economical separation methods such as, for example, filtration and gravitational sedimentation are widely applied in wastewater treatment facilities to harvest relatively large (>70 μπι) microalgae such as Coelastrum and Spirulina. However, they are less useful for harvesting algae species approaching bacterial dimensions (<30 μπι) such as Scenedesmus, Diinaliella, and Chlorella (Brennan and Owende, 2010 Renew Sustain Energy Rev 14(2):557-577), to which most oleaginous microalgae species belong.

Centrifugation is widely used to recover microalgae biomass, especially small-sized algae cells. However, centrifugation typically involves intensive energy needs and high equipment maintenance requirements. Thus, its application is often limited to algae cultures for high- value metabolites. While flocculation can sometimes be used to harvest small-sized microalgae cells, it more often is used as a preparatory step involving aggregating the microalgae cells and increasing the particle size so that other harvesting methods such as filtration, centrifugation, or gravity sedimentation can be applied (Molina Grima et al., 2003 Biotechn Adv 20(7-8):491-515). Several flocculants have been developed to facilitate the aggregation of microalgae cells, including multivalent metal salts like ferric chloride (FeCl3), aluminum sulphate (A12(S04)3), ferric sulphate (Fe2(S04)3), and organic polymers such as chitosan and modified starch (Li et al, 2008 Biotechnol Prog 24(4):815-820). Chemical flocculation can be reliably used to remove small algae cells from pond water by forming large (1-5 mm) sized floes (Sharma et al., 2006 J Polym Environ 14(2):195-202). However, besides the high cost of chemical flocculants and possible pollution effects that chemical flocculants may generate, the chemical reactions are highly sensitive to pH and the high doses of flocculants required can produce large amounts of sludge and may leave a residue in the treated effluent. In summary, most technologies including chemical and mechanical methods increase operational costs for algal production and are only economically feasible for production of high-value products (Park et al., 2011 Bioresource Techn 102(l):35-42).

Other microalgae harvesting methods also exhibit certain limitations. Other microalgae harvesting technologies include using the principles of liquid adhesion and capillary action to extract water from dilute microalgae solutions, growing microalgae on the surface of polystyrene foam to simplify the cell harvest (Wilkie and Mulbry, 2002

Bioresource Tech 84(1):81-91 ; Johnson and Wen, 2010 Appl. Microbiol. Biotechnol.

85(3):525-534), and new bioflocculants, which are more environmentally friendly, are also proposed to address the cost and environmental concerns for current flocculation methods (Uduman et al., 2010 J Renew Sustain Energy 2(1)). These methods can decrease the harvest cost to some extent if developed successfully, but each requires heavy investments in equipment and chemical supplies. For example, cell pellets of filamentous fungi Aspergillus flavus were applied as a biological flocculation agent to harvest the microalgae cells. These pellets must first be prepared by fungal cultivation with sugar medium, then the fungal pellets are added to the microalgae culture so they can absorb the microalgae cells into the pellet structure by virtue of differences in the surface charge between the fungal cells and the microalgae cells (Rajab, 2007 "Micro-Algae Removal In Domestic Wastewater Using Aspergillus Flavus Soft Pellets As A Bio-Coagulant," Masters Thesis: Universiti Putra Malaysia).

Enhancing natural algae aggregation to encourage simple gravity settling or filtration is a promising option to achieve both a high-quality treated effluent, in terms of total suspended solids, and an economic recovery of algal biomass for biofuel production and/or other uses (Uduman et al., 2010 J Renew Sustain Energy 2(1)). Natural algae aggregation also can be more environmentally sound than current procedures which may need additives. Many of the algal species in the wastewater treatment processes often form large colonies (e.g., 50 μπι to 200 μπι), and their aggregation can be achieved through nitrogen limitation and/or C02 addition (Park, J. et al, 2011 Bioresource Techn 102(l):35-42). However, most of these microalgae species are not oleaginous species. Methods that induce oleaginous microalgae to aggregate during their cultivation are strategically and urgently needed to develop efficient and economic means of producing biofuels and/or other bioproducts.

We describe herein methods for harvesting unicellular microalgae. Since these microalgae are not filamentous, harvesting methods such as natural algae aggregation, simple gravity settling, or filtration are typically not commercially useful for harvesting their cell biomass. Having a separate harvesting step— whether flocculation, centrifugation, or other method— is often costly and/or creates environmental challenges. Our methods involve pelletizing microalgae cells by adding spores of filamentous fungi to the microalgae culture. Our method shifts harvesting strategies from those that harvest individual cells from a suspension culture to simple, cost-effective methods for harvesting microalgae-filamentous fungi cell pellets. Co-culturing filamentous fungi with microalgae cells so that the filamentous feature of the fungi can be introduced to the microalgae culture induces the cell pelletization. Harvesting cell pellets, which can be done, for example, by simple filtration, can be significantly easier and less costly than harvesting individual cells.

As used herein, the term "Filamentous fungus" and variations thereof refer to a cell or a spore of a particular fungal species. Thus, for example, "a member of the genus

Aspergillus" refers to a cell or a spore of any fungal species in the genus Aspergillus.

Depending on the application of microalgae cells, different filamentous fungal species can be co-cultured to pelletize microalgae cells for easier harvest. For instance, microalgae species that have cellulose in the cell wall can be co-cultivated with a cellulase-producing fungus. Such a co-culture can induce pelletization of the microalgae and the cellulose in the microalgae cell walls can provide a carbon source that can support the growth of the filamentous fungus without having to add a carbon source to the culture. Moreover, the partial degradation of the microalgae cell wall can also contribute to the extraction of oil or other products inside the microalgae cells.

In another example, if one cultivates microalgae to generate microbial lipids, co- cultxiring an oleaginous fungus can be similarly beneficial. This can be especially true in the heterotrophic cultivation of oleaginous microalgae, where sugars are utilized by microalgae to accumulate oil in the cell biomass. Since the filamentous fungi are oleaginous and their cell biomass, similar to microalgae cells, have a high lipid content that can be extracted for biodiesel production, co-culturing these oleaginous fungi cells will not only facilitate the pelletization of microalgae cells for easier harvest, but the fungal cells can contribute to oil production.

The research represents a highly innovative approach to address both the cost and sustainability issues in microalgae biofuel production, and has the potential to revolutionize the next generation algae biofuel industry. The mixed culture of microalgae and filamentous fungi to pelletize cells provides a novel process with direct commercial application potential. It is a transformative method that can be applied to different microalgae and fungal strains, in some cases by modifying the operational conditions across certain strain combinations.

Introducing a filamentous feature to oleaginous microalgae for better harvest may provide a platform for developing economically feasible processes for microalgae cultivations for a wide variety of industrial applications including, for example, generating biofuel and other products and/or wastewater treatment.

The methods described herein can provide one or more of the following benefits compared to conventional microalgae harvesting methods. While pelletization and granulation of filamentous microbes have been reported for certain commercial production methods, this is the first report of pelletizing non-filamentous microalgae cells by co-culturing filamentous fungi in order to systematically emphasize the separation benefits of the pelletization process. This concept and design can be transformative to other microalgae cultures and can be applied not only for biofuel production, but also in processes for, for example, producing

nutraceuticals or treating wastewater treatment. Compared to other algae harvesting methods, the methods described herein can decrease harvest costs, avoid second pollution from chemical flocculants, and permit high water reuse. Many agricultural wastes have potential to be applied in the bioenergy production, but their application can be limited due to, for example, the relative low-nutrient level of the waste itself and/or, as another example, the high cost of separating the final product can overcome any cost savings from using agricultural waste as a raw material. The methods described herein can facilitate the production of a financially economical, environmentally sustainable, and ecologically stable process. These methods can provide benefits for any microalgae cultivation process that includes harvesting microalgae biomass such as, for example, processes involving wastewater treatment or the production of, for example, bioenergy, nutraceutical oils, food components, fertilizer, bioplastics, or pharmaceuticals. For example, we show fungi-algae pellets cultured on concentrated municipal wastewater for one day effectively remove pollutants from the wastewater (Example 5, FIG. 13). We measured removal of NH4-N, total phosphorus, COD, and total nitrogen in the concentrated municipal wastewater samples. NH4-N was 100% removed after one-day cultivation (FIG. 13 A). This result is better than previous reports using algae alone for wastewater treatment (Li et al., 2011 Bioresource Tech 102:10861-10867; Wang et al., 2010 Appl Biochem Biotechnol 162:1174-1186; Zhou et al., 2011 Bioresour Technol.

102(13):6909-6919). Total phosphorus was reduced from 53.0 mg/L to 5.00 mg/L and stayed at the reduced level through the end of cultivation (FIG. 13B). Maximal removal efficiency reached round 90.0 %, and was thus much higher than those reported in other studies using municipal wastewater (Cheng et al., 2002 Trans. ASAE 45:799-805; Lau et al., 1995

Environmental Pollution 89:59-66; Woertz et al, 2009 J. Envir. Engrg., 135(11):1115-1122). This efficiency suggests a synergetic treatment effect of algae-fungi pellets compared to a sum of the individual effects of algae alone and fungi alone. An increase in pH to 8 can cause aggregation of algae and adsorption of inorganic phosphates. The concentration of COD decreased from 1,600 mg/L to 600 mg/L in 24 hours (FIG. 13C). Since COD is commonly used as an indirect measure of organic compounds in wastewater, our result suggests that organic carbon in the wastewater may be utilized by both algae and fungal species (Zhou et al, 2011 Bioresour Technol. 102(13):6909-6919; Liao et al, 2007 Bioresource Techn 98:3415-3423). Total nitrogen dropped from 90.0 mg/L to 35.0 mg/L in one day (FIG. 13D).

As another example, we used the harvested fungi-algae pellets to treat 20-fold diluted swine manure wastewater. N¾-N, total phosphorus, COD, and total nitrogen in 20-fold diluted swine wastewater for the two-days batch culture is depicted in FIG. 14A-D, respectively. The NH4-N, total phosphorus, and COD dropped from 90 mg/L to 65 mg/L, from 2.00 mg/L to 0.30 mg/L, and from 1000 mg/L to 280 mg/L, respectively, in 48 hours. Total nitrogen was reduced from 130 mg/L to 70.0 mg/L in 24 hours. These results reflect better nutrient removal from swine manure wastewater than previous treatments using algae alone for wastewater treatment. Indeed, after overnight culture in fungi-algae pellets, the color of the swine manure became much more clear than that of initial sample (FIG. 15). Our results confirmed these studies and provided an effective alternative for dark-color animal manure wastewater treatment.

As yet another example, the fungi-algae pellets may be used to treat industrial wastewater contaminated with heavy metals. Some heavy metals such as, for example, aluminum (Al) and copper (Cu) could be used as catalysts for converting the biomass feedstock into bio-oil by hydrothermal technology (Du et al., 2011 Bioresour Technol.

102:4890-4896). Therefore, the fungi-assisted immobilized algal cells with the absorbed metals could be converted to refined bio-oil directly by thermo-chemical processes as both feedstock and catalysts (Du et al, 2011 Bioresour Technol. 102:4890-4896).

In some embodiments, pelletized fungal fermentation can be exploited for microbial oil accumulation. Most oleaginous microalgae are not filamentous and oleaginous microalgae pelletization has not been reported. We investigated inoculating filamentous fungal spores when culturing mixotrophic green algae C. vulgaris and found that pellets clearly formed within two days of culture. The algae solution lost most of its green color, and turned transparent, indicating the majority of microalgae cells were pelletized (FIG. 2). As seen in the image captured via microscope, the skeleton structure of the pellet was still filamentous fungal cells. The microalgae cells, aggregated together with fungal cells, were immobilized in the pellets. This is an innovative technology uniquely addressing the cell harvest of microalgae and has the potential to greatly reduce the algae biofuel cost.

In one aspect, the methods described herein generally include adding at least one filamentous fungus to a culture of microalgae, growing the culture of microalgae under conditions effective to produce a co-culture of the microalgae and the filamentous fungus and to form cell pellets, and harvesting the cell pellets.

In some of embodiments, the filamentous fungus can include at least one spore and/or at least one filamentous fungus cell. Exemplary filamentous fungi include, for example, members of the genus Aspergillus (e.g., A. oryzae, A. flavus, and/or A. niger), members of the genus Mucor (e.g., M. circinelloides), members of the genus Phanerochaete, members of the genus Leucogyrophana, members of the genus Rhizopus (e.g., Rhizopus oryzae, and/or Rhizopus chinentis), members of the genus Absidia (e.g., Absidia coerulea), members of the genus Penicillium, members of the genus Trichoderma, and members of the genus Fusarium (e.g., Fusarium venenatum), although the method may be practiced using other filamentous fungi. In some embodiments, the filamentous fungi can include a mixture of two or more filamentous fungus species. Also, the filamentous fungus can include a mixture of spores and filamentous fungus cells.

In some embodiments, the microalgae can include at least one autotroph and/or at least one heterotroph. Exemplary autotrophic microalgae include, members of the genus Chlorella, members of the genus Dunaliella, members of the genus Scenedesmus, members of the genus Schizochytrium, and members of the genus Ourococcus, although other autotrophic microalgae species exist and may be used to practice the methods described herein.

Exemplary heterotrophic microalgae include, for example, members of the genus Chlorella, members of the genus Crypthecodinium, members of the genus Scenedesmus, members of the genus Chlamydomonas, members of the genus Micractinium, and members of the genus Euglena, although other heterotrophic microalgae species exist and may be used to practice the methods described herein

When the microalgae includes an autotrophic microalgae, the co-culture conditions can include a maximum pH of no greater than pH 7 such as, for example, a pH of no greater than pH 7.0, no greater than pH 6.9, no greater than pH 6.8, no greater than pH 6.7, no greater than pH 6.6, no greater than pH 6.5, no greater than pH 6.4, no greater than pH 6.3, no greater than pH 6.2, no greater than pH 6.1, no greater than pH 6.0, no greater than pH 5.9, no greater than pH 5.8, no greater than pH 5.7, no greater than pH 5.6, no greater than pH 5.5, no greater than pH 5.4, no greater than pH 5.3, no greater than pH 5.2, no greater than pH 5.1, no greater than pH 5.0, no greater than pH 4.9, no greater than pH 4.8, no greater than pH 4.7, no greater than pH 4.6, no greater than pH 4.5, no greater than pH 4.4, no greater than pH 4.3, no greater than pH 4.2, no greater than pH 4.1 , no greater than pH 4.0, no greater than pH 3.9, no greater than pH 3.8, no greater than pH 3.7, no greater than pH 3.6, no greater than pH 3.5, no greater than pH 3.4, no greater than pH 3.3, no greater than pH 3.2, no greater than pH 3.1, or no greater than pH 3.0.

In some of these embodiments, co-culture including a autotrophic microalgae can include a minimum pH of no less than pH 3 such as, for example, a pH of no less than pH 3.0, no less than pH 3.1, no less than pH 3.2, no less than pH 3.3, no less than pH 3.4, no less than pH 3.5, no less than pH 3.6, no less than pH 3.7, no less than pH 3.8, no less than pH 3.9, no less than pH 4.0, no less than pH 4.1, no less than pH 4.2, no less than pH 4.3, no less than pH 4.4, no less than pH 4.5, no less than pH 4.6, no less than pH 4.7, no less than pH 4.8, no less than pH 4.9, no less than pH 5.0, no less than pH 5.1, no less than pH 5.2, no less than pH 5.3, no less than pH 5.4, no less than pH 5.5, no less than pH 5.6, no less than pH 5.7, no less than pH 5.8, no less than pH 5.9, no less than pH 6.0, no less than pH 6.1, no less than pH 6.2, no less than pH 6.3, no less than pH 6.4, no less than pH 6.5, no less than pH 6.6, no less than pH 6.7, no less than pH 6.8, no less than pH 6.9, or no less than pH 7.0.

The pH of the co-culture also can be characterized by any range that includes, as endpoints, any combination of minimum pH value identified above and maximum pH value identified above that is greater than the minimum pH value.

Also when the microalgae includes an autotroph, the filamentous fungus spores may be added to a minimum final concentration of at least 103 spores per liter such as, for example, at least 103 spores per liter, at least 104 spores per liter, at least 105 spores per liter, or at least 106 spores per liter. The filamentous fungus also may be added to the microalgae culture to a maximum final concentration of no more than 10 spores per liter such as, for example, no more than 10 spores per liter, no more than 10 spores per liter, or no more than 104 spores per liter. The amount of spores added to the microalgae culture also can be characterized by any range that includes, as endpoints, any combination of a minimum final concentration and a maximum final concentration that is greater than minimum final concentration. The precise amount of spores added to the microalgae culture can depend, at least in part, on the cell concentration of the microalgae when the spores are added.

In some embodiments, the filamentous fungus may be added to a culture that includes an autotrophic microalgae when the microalgae culture possesses a cell density of at least 4 x 106 cells per milliliter and no more than 10 x 106 cells per milliliter such as, for example, a cell density of at least 6 x 106 cells per milliliter and no more than 8 x 106 cells per milliliter. In some embodiments, the filamentous fungus may be added to the microalgae culture during mid-log phase.

When the microalgae culture includes at least one heterotroph, the co-culture conditions can include a minimum pH of no less than pH 3 such as, for example, a pH of no less than pH 3.0, no less than pH 3.1, no less than pH 3.2, no less than pH 3.3, no less than pH 3.4, no less than pH 3.5, no less than pH 3.6, no less than pH 3.7, no less than pH 3.8, no less than pH 3.9, no less than pH 4.0, no less than pH 4.1, no less than pH 4.2, no less than pH 4.3, no less than pH 4.4, no less than pH 4.5, no less than pH 4.6, no less than pH 4.7, no less than pH 4.8, no less than pH 4.9, no less than pH 5.0, no less than pH 5.1, no less than pH 5.2, no less than pH 5.3, no less than pH 5.4, no less than pH 5.5, no less than pH 5.6, no less than pH 5.7, no less than pH 5.8, no less than pH 5.9, no less than pH 6.0, no less than pH 6.1 , no less than pH 6.2, no less than pH 6.3, no less than pH 6.4, no less than pH 6.5, no less than pH 6.6, no less than pH 6.7, no less than pH 6.8, no less than pH 6.9, no less than pH 7.0, no less than pH 7.1, no less than pH 7.2, no less than pH7.3, no less than pH 7.4, no less than pH 7.5, no less than pH 7.6, no less than pH 7.7, no less than pH 7.8, or no less than pH 7.9.

In some of these embodiments, co-culture including a heterotrophic microalgae can include a maximum pH of no greater than pH 8 such as, for example, a pH of no greater than pH 8.0, no greater than pH 7.9, no greater than pH 7 •8, no greater than pH 7-7, no greater than pH 7.6, no greater than pH 7.5, no greater than pH 7, ■4, no greater than pH 7.3, no greater than pH 7.2, no greater than pH 7.1, no greater than pH 7, .0, no greater than pH 6.9, no greater than pH 6.8, no greater than pH 6.7, no greater than pH 6, .6, no greater than pH 6.5, no greater than pH 6.4, no greater than pH 6.3, no greater than pH 6. .2, no greater than pH 6.1, no greater than pH 6.0, no greater than pH 5.9, no greater than pH 5. 8, no greater than pH 5.7, no greater than pH 5.6, no greater than pH 5.5, no greater than pH 5, ■4, no greater than pH 5.3, no greater than pH 5.2, no greater than pH 5.1, no greater than pH 5. ■0, no greater than pH 4.9, no greater than pH 4.8, no greater than pH 4.7, no greater than pH 4. •6, no greater than pH 4.5, no greater than pH 4.4, no greater than pH 4.3, no greater than pH 4. .2, no greater than pH 4.1, no greater than pH 4.0, no greater than pH 3.9, no greater than pH 3. .8, no greater than pH 3.7, no greater than pH 3.6, no greater than pH 3.5, no greater than pH 3. ■4, no greater than pH 3.3, no greater than pH 3.2, no greater than pH 3.1, or no greater than pH 3.0.

The pH of the co-culture also can be characterized by any range that includes, as endpoints, any combination of minimum pH value identified above for a culture that includes a heterotroph and any maximum pH value identified above for a culture that includes a heterotroph that is greater than the minimum pH value. For example, in some embodiments, the conditions can include a pH of from pH 5 to pH 6.

In some embodiments, the conditions can include no more than 25 g/L of sugar as a carbon source, such as, for example, no more than 20 g/L of sugar, no more than 15 g/L of sugar, no more than 10 g/L of sugar, no more than 5.0 g/L of sugar, no more than 3.0 g/L of sugar, no more than 2.5 g/L of sugar, no more than 2.0 g/L of sugar, no more than 1.5 g/L of sugar, no more than 1.0 g/L of sugar, no more than 0.75 g/L of sugar, no more than 0.5 g/L of sugar, or no more than 0.25 g/L of sugar as a carbon source. Such culture conditions may be viable, in part, because certain filamentous fungi express cellulase, an enzyme that can at least partially degrade cellulose in the cell wall of microalgae. Thus, the filamentous fungi can use, to some extent, cellulose released from the microalgae as a result of the filamentous fungi secreting cellulase into the culture medium. Thus, in one embodiment, the co-culture conditions include 0.25 g/L of sugar as a carbon source.

In embodiments in which the co-culture includes a heterotroph, filamentous fungus spores may be added to a minimum final concentration of at least 10 spores per liter such as, for example, at least 103 spores per liter, at least 104 spores per liter, at least 105 spores per liter, or at least 106 spores per liter. The filamentous fungus also may be added to the microalgae culture to a maximum final concentration of no more than 10 spores per liter such as, for example, no more than 106 spores per liter, no more than 105 spores per liter, or no more than 104 spores per liter. The amount of spores added to the microalgae culture also can be characterized by any range that includes, as endpoints, any combination of a minimum final concentration and a maximum final concentration that is greater than minimum final concentration. The precise amount of spores added to the microalgae culture can depend, at least in part, on the cell concentration of the microalgae when the spores are added.

In some embodiments, the filamentous fungus may be added to a culture that includes a heterotrophic microalgae when the microalgae culture possesses a cell density of at least 1 x 106 cells per milliliter and no more than 8 x 106 cells per milliliter such as, for example, a cell density of at least 2 x 106 cells per milliliter and no more than 6 x 106 cells per milliliter. In some embodiments, the filamentous fungus may be added to the microalgae culture early log phase.

In some embodiments, the filamentous fungus may be selected because it can produce a bioproducts in addition to inducing the formation of cell pellets. For example, certain filamentous fungi are oleaginous - i.e., they are capable of producing oil. Exemplary oleaginous filamentous fungi include, for example, Mucor circinelloides. Thus, in certain embodiments, the filamentous fungus may be selected to produce an oil that is also produced by microalgae in the co-culture. The cell pellets may be harvested by any conventional method such as, filtration. As noted above, filtration of the pellets formed during co-culture can be easier and less costly than filtration of microalgae suspended in culture.

In some embodiments, the method can include additional steps for extracting a bioproducts from the harvested cell pellets. As discussed above, one exemplary bioproducts can include oil such as, for example, for biofuel production. Alternative exemplary bioproducts include, for example, nutraceutical oils, food components, fertilizers, bioplastics, or pharmaceuticals.

In submerged cultures, many filamentous fungi tend to aggregate and grow as pellets or granules. These fungal cell pellets can be spherical or ellipsoidal masses of hyphae with variable internal structures, ranging from loosely packed hyphae, forming "fluffy" pellets, to tightly packed, compact, dense granules (Hu and Chen, 2007 Int J Hydrogen Energy

32(15):3266-3273; Hu and Chen, 2008 Appl Biochem Biotechn 148(1 -3):83-95; Hu et al., 2009 Environ Prog Sustain Energy 28(1):60-71; Xia et al., 2011 Biotechn Biofuels 4:15). Beyond benefits from cell immobilization, other advantages for microalgae oil production include increasing the ease of harvesting cells and the ability to reuse pond water (Johnson and Wen, 2010 Appl. Microbiol. Biotechnol. 85(3):525-534; Xia et al, 2011 Biotechn Biofuels 4:15).

Our data show conditions for cell pelletization seem to be strain-specific— i.e., not all the filamentous fungal strains can form pellets during their growth. For example, Aspergillus oryzae is a commercial fungal strain able to produce cellulase. A. oryzae can form relatively large, stable, homogeneous cell pellets during the cultivation without any inducing

approaches.

As another example, Mortierella isabellina, on the other hand, is less able to form stable pellets. During cultivation in flasks, pellets were formed in the first day of culture of M. isabellina, but all of the pellets disappeared after four days of cultivation. M. isabellina is a promising oil-producing fungus, with 40-60% content of biomass as oil. Thus, stable pelletization strategies for M. isabellina are still being explored. In the meantime, M.

isabellina may be used in pelletizing methods that can exploit the transient pellet-forming nature of the fungus. Conventional pelletization strategies involving M. circinelloides can include induction with CaC03 powder in the early stage of its cultivation. In such methods, cell pellets can be formed homogeneously, lasting for the entire cultivation cycle. However, such approaches can be costly and cause solid waste disposal issues. Thus, we have investigated alternative ways to induce pelletization of M. circinelloides. We successfully induced pelletization of M. circinelloides by adjusting the pH of the culture (FIG. 1). We discovered that changing conditions during cell cultivation can force fungal cells to aggregate and form pellets (Xia et al, 2011 Biotechn Biofuels 4:15).

In the preceding description, particular embodiments may be described in isolation for clarity. Unless otherwise expressly specified that the features of a particular embodiment are incompatible with the features of another embodiment, certain embodiments can include a combination of compatible features described herein in connection with one or more embodiments.

For any method disclosed herein that includes discrete steps, the steps may be conducted in any feasible order. And, as appropriate, any combination of two or more steps may be conducted simultaneously.

As used in the preceding description and the following Examples and claims, the term "and/or" means one or all of the listed elements or a combination of any two or more of the listed elements; the terms "comprises" and variations thereof do not have a limiting meaning where these terms appear in the description and claims; unless otherwise specified, "a," "an," "the," and "at least one" are used interchangeably and mean one or more than one; and 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.).

The present invention is illustrated by the following examples. It is to be understood that the particular examples, materials, amounts, and procedures are to be interpreted broadly in accordance with the scope and spirit of the invention as set forth herein. EXAMPLES

General Methods

Microbial species

Filamentous oleaginous fungi A. niger Ted S-OSU and M. circinelloides were chosen to test their pelletization for the microalgae cells. M. circinelloides is a filamentous oleaginous fungus with limited capability to produce cellulase. Due to this fungus's features for lipid accumulation and biodiesel production, the U.S. Department of Energy has decided to sequence the whole genome through its bioenergy program at the Joint Genome Institute. M circinelloides is mostly filamentous, but has a yeast-like morphology at conditions with low oxygen or high substrate concentrations. It has a moderate level of lipids in the mycelium (around 25% dry mass in wild-type strains and over 60% in mutated strains), and good biomass production during submerged batch cultivation in bioreactors (Aggelis, 1996 Folia Microbiologica 41(3):254-256; Wynn et al., 2001 Microbiology-Sgm 147:2857-2864).

Filamentous fungi A. niger Ted S-OSU is a well-known cellulase-producing strain, and our lab found that it is relatively easy to form pellets during its liquid fermentation. Other filamentous fungal species are also tested on their pelletization capability when co-culturing with microalgae.

C. vulgaris and C. protothecoides were our model microalgae species. Our previous research has revealed that C. vulgaris is a potential robust producer of microbial lipids because it is a typical mixotrophic oleaginous microalgae, which can assimilate both organic carbon and sunlight for their cell growth (Heredia-Arroyo et al., 2011 Biomass & Bioenergy 35(5):2245-2253). Compared to C. vulgaris, which has been studied for the potential as food and nutritional sources, C. protothecoides shares many similarities as C. vulgaris, and its capability to accumulate lipids is superior so that it has been widely researched for biofuel production (Heredia-Arroyo et al, 2010 Appl Biochem Biotechn 162(7):1978-1995). The co- pelletization conditions were also applied to test a few other microalgae strains that have commercial applications, such as Scenedesmus obliquus and Dunaliella sp.. Detailed research steps are as follows: Cell cultivation

Cultivation medium

Autotrophic medium A (per L): 1 g KN03, 0.075 g K¾P04, 0.1 g K2HP04, 0.5 g MgS04.2H20, 0.0625 g Ca(N03)2.4H20, 0.0 lg FeS04.7H20, 0.5 g Yeast extract, and 1 mL A5 lmL. A5 (per L): 2.86 g H3B03, 0.039 g Na2Mo4.2H20, 0.222 g ZnS04.7H20, 1.81 g MnCl2.4H20, 0.074 g CuS0.5H20, and 0.03 g CoCl2.

Heterotrophic medium B (per L): 12 g potato dextrose broth and 15 g glucose.

Cultivation of seed C. vulgaris:

C. vulgaris was purchased from UTEX (Austin, TX) and cultivated with 3L culture medium A in a 6L flask with a mechanical stir (100 rpm) under fluorescent light at 25-27°C. The cultivation broth was regularly replaced with fresh media to maintain the active cell growth so that this flask of cultivation broth can be used as the seed for the co-cultivation. Cultivation, harvest and storage of seed fungal spore

Fungal spores were inoculated on sterile petri dish with culture medium B with 2% agar. Then, culture the petri dish in an incubator at 27°C for 7 days. Wash the fungal spores by 10 mL sterile water per petri dish, and put 1 mL spore solution into sterile Eppendorf tube. These spores were stored in -70°C after counting their numbers under microscope and then used as the seed for the co-cultivation.

Co-culture of filamentous fungi with autotrophic or heterotrophic microalgae

100 mL culture medium A was prepared in a 250 mL flask, and then pH adjusted by adding 2 mol/L HCl or 1 mol/L NaOH to the desired pH value. 20 mL microalgae seed broth was transferred into these 250 mL flasks. Finally, fungal spores were inoculated into medium A to reach a spore concentration of 7.6 10 6/L. The inoculated flasks were cultured at 27°C, 150 rpm for three days. Analysis

Glucose analysis:

Glucose concentration was estimated by using DNS method. The detailed

measurement method includes following:

Solution (per L): 10 g Dinitrosalicylic acid, 2 g phenol, 0.5 g sodium sulfite, 10 g sodium hydroxide. Potassium sodium tartrate solution, 40%. Standard Glucose solution: 0.5 g/50 mL, then dilute 20-fold for use.

Procedure: 1 mL of sample was mixed with 1 mL diluted DNS after dilution, then incubated at 100°C for 15 minutes. 0.5 mL 40% potassium sodium tartrate solution was added after reaction, cooled to room temperature, then tested at OD540. The glucose concentration was calculated according the standard curve.

Dry Biomass determination:

The microbial broth was centrifuged at 7,000 rpm for 7 minutes, dried at 105°C for 16 hours and then weighed.

Fatty acid analysis:

About 100 mg dried microbial biomass samples were weighed and put into 25 mL screw-top glass tubes. 10 mL mixture of chloroform, methanol, and concentrated sulfuric acid (prepared in a v/v ratio of 5:4.25:0.75, respectively) was then added into each tube for the transesterification reaction at 90°C water bath (Model 25, Precision Scientific; Chicago, IL) for 90 minutes. 2.5 mL water was added to the reactants and vortexed for one minute, and then the reactant liquid was centrifuged at 7,000 rpm for seven minutes. The upper layer liquid was mixed with 3 mL chloroform and vortexed for one minute. The chloroform layer of the second centrifugation was fed into the tube having the first centrifugation chloroform layer. FAME in chloroform was carefully collected and subjected to GC after filtration (0.22 μπι). The GC was equipped with a flame ionization detector and a HP-5 capillary column. The oven temperature was 80°C, held for five minutes, raised to 290°C at a rate of

4°C/minute, and held at 290°C for five minutes, while the injector and detector temperature were set at 250°C and 230°C, respectively. The carrier gas (helium) was controlled at 1.2 mL/minute. Chromatographic data were recorded and integrated using Agilent data analysis software. The fatty acid was quantified by comparing the peak area with that of the GLC standard mixtures (GLC-10, GLC-40, GLC-80 GLC-100) (Sigma-Aldrich; St. Louis, MO). Photos of pellets:

Fungi-microalgae pellets were poured from the growth flask into the petro dish and photos were taken with digital camera (DSC-T20, Sony). In this case, each petri dish in the photo has all the pellets formed in that flask culture, but not all the supernatant liquid. Microalgae cell number in supernatant:

To count microalgae cells remaining in the supernatant, the supernatant was diluted multiple times until the cell numbers could be counted under microscope.

Total microalgae cell number:

The entire cultivation broth, including pellets and supernatant liquid, was poured from the growth flask into a beaker. The broth was blended to completely break the structure of pellets in order to release microalgae cells from pellets. The blended broth was diluted multiple times until microalgae cell numbers could be counted under a microscope. Harvest ratio

Harvest ratio or pelletization ratio is defined as 100% minus the microalgae cell numbers in supernatant divided by the total microalgae cell numbers in the fermentation broth. Example 1 - Test on different filamentous fungal strains and different microalgae species Different fungal species were tested in our lab for their co-pelletization capability with the green algae C. vulgaris, as shown in Table 1. Several fungal species, such as ATCC 9642, RLG 9902, and A. flavus, can form pellets that can entrap all of the individual microalgae cells, resulting in a clear co-culture broth of those strains (FIG. 12). Similar results, not shown in FIG. 12, were observed for additional fungal species such as, for example, ATCC 11730. These results clearly showed that this co-pelletization can also be applied to many other similar filamentous fungal species (molds). We also tested the possibility of pellet formation in cultures of A. niger with other types of microalgae and FIG. 12 showed that these strains of microalgae can form the pellets when co-culturing with the filamentous fungi.

Table 1. Co-pelletization of fungal species and C. vulgaris after 24 hour heterotrophic condition

Figure imgf000022_0001

Photos of co-pellets of fungal species and C. vulgaris are shown in FIG. 12.

Example 2 - Test on different growth mode of microalgae

Many microalgae can grow on both autotrophic and heterotrophic growth modes. When co-culturing a cellulase producing filamentous fungus, i.e. A. niger, the fungus can either depend on the degradation of microalgae cell wall in the autotrophic culture or depend on the sugar sources in the heterotrophic cultivation. Two exploratory processes were tested by using this approach of cell pelletization to facilitate the cell harvest as shown in Table 2. For the co-culture with autotrophic microalgae, around 60% of algae cells were pelletized with fungal cells. Interestingly, the algae concentration in the supernatant was about same as the control, which did not have fungal inoculation. This clearly showed that the co-cultivation of fungal cells actually stimulated the overall microalgae cultivation and dramatically increased the final algae cell concentration, which was limited by the mutual shading effects. The "extra" algae cells in the co-culture were pelletized so that the suspended algae cell concentration could remain same. For the heterotrophic microalgae culture, since algae growth relies on the sugar, instead of light, microalgae cell concentration increased dramatically, with relatively smaller portion being pelletized. Table 2. Co-culture of A. niger with C. vulgaris at different growth mode (initial pH 4.0 for

Figure imgf000023_0001

Control: 100 mL microalgae seed; autotrophic: 100 mL microalgae seed inoculated with 7.6 10 6/L fungal spore, no fresh medium added; 100 mL microalgae seed inoculated with 7.6 10 6/L fungal spore, 0.05 g glucose powder added.

Detailed research was then done on each growth mode as following;

Example 3 - Co-culture of filamentous fungi with autotrophic microalgae

Co-cultivation of a cellulase-producing fungus with autotrophic microalgae cultivation (FIG. 3) can use most current commercial filamentous fungal strains that generate cellulase, which is excreted in the fermentation broth. It is recently reported that external cellulase can be added to the microalgae cultures to hydrolyze the polysaccharides within the microalgae cell walls in order to generate free sugars for bioethanol fermentation (Harun and Danquah, 2011 Chem Engineer J 168(3):1079-1084) and also to facilitate the lipid extraction (Fu et al., 2010 Bioresource Techn 101(22):8750-8754). The carbohydrate composition of microalgae is mainly polysaccharide entrapped in the cell walls and can account for up to 70% dry weight of the microalgae biomass.

The cellulose hydrolysis by external cellulase has been applied to various microalgae cultures, including Chlorella species and Chlorococum humicola, and satisfactory results were reported to break down the microalgae cell walls and facilitate oil extraction (Fu et al., 2010 Bioresource Techn 101(22):8750-8754; Yin et al, 2010 J Food Science 75(9):H317-H323). We tested the co-culture of the filamentous cellulase-producing fungus with autotrophic microalgae. Different cultivation conditions were studied as following: Effect of different initial pH on microalgae pelletization

The results showed that the best initial pH for the co-cultivation was at 4.0. Green pellets were formed during the co-cultivation and almost 60% of algae cells were in the form of pellets. The growth of microalgae cells seemed being stimulated when pellets were formed because the supernatant microalgae cell concentration at initial pH 4.0 were similar or slightly larger than at other pH level, where pellets were not formed. When initial pH was 7.0 or pH 8.0, there were no pellets formed at all, and the color of the cultivation broth turned to yellow green. When the initial pH was 5.0 or pH 6.0, there were a very few irregular cell clumps or pellets formed in the cultivation broth. Results are shown in FIG. 4.

Effect of spore concentration on microalgae pelletization

Fungal spore concentration had some influence on the co-pelletization of microalgae and fungal cells. With the fungal spore inoculation concentration at 7.6 10 6/L, around 60% of microalgae cells were aggregated into the pellets. Results are shown in FIG. 5.

Effect of algae concentration on microalgae pelletization

Different concentration of initial microalgae inoculate significantly affected the overall harvest ratio, although the supernatant microalgae concentration remained relatively constant (FIG. 6). The results showed that the best concentration of microalgae inoculate is almost half concentration of the mature stage of microalgae cultivation, which gave us the direction that it is the best to inoculate the fungal spores in the middle the log phase growth of microalgae.

Example 4 - Co-culture of filamentous fungi with heterotrophic microalgae

Co-cultivation of filamentous fungi with the heterotrophic microalgae can be applied as a modification of a two-stage hybrid cultivation system (Xiong et al., 2008 Appl Microbiol Biotechn 78(l):29-36). Microalgae were cultured in the autotrophic conditions in the first stage to accumulate biomass by using carbon dioxide and sunlight, and then the whole culture was switched to heterotrophic cultivation with external sugars from agricultural biomass added into the system to create nitrogen limited conditions for oil accumulation (FIG. 7). Different cultivation conditions were studied as following:

Effect of different initial pH on microalgae pelletization

Pellet formation was greatest when the pH of the co-culture was adjusted to between pH 5.0 and pH 6.0. Within this range, microalgae growth was stimulated and over 90% of the microalgae cells were pelletized. Low pH (including 3.0 and 4.0) showed inhibition of microalgae growth, and similar phenomena could be seen at higher initial pH levels (7.0 and 8.0) although the harvest ratio at initial pH 7.0 was also around 90%. Results are shown in FIG. 8.

Effect of spore concentration on microalgae pelletization

Enough fungal spore inoculation should be provided to stimulate the co-pelletization process. If the fungal inoculum was too small, there were no pellets formed at all because obviously no significant fungal growth was observed. Results are shown in FIG. 9.

Effect of glucose and nitrogen concentration on microalgae pelletization (fixed C/N)

With fixed C/N ratio, decreasing the glucose input did not significantly affect the harvest ratio although the overall microalgae cells generally decreased due to the limited supply of nutrients, especially nitrogen (FIG. 10). These results are very promising because only 0.25 g/L of sugars is needed to have almost 90% of algae harvest. Results are shown in FIG. 10.

Effect of algae concentration on microalgae pelletization

With higher concentration of microalgae to start with the co-culture, less microalgae cells are pelletized with fungi (FIG. 11). For the heterotrophic microalgae cultivation, the results showed that fungal spores need to be inoculated in the early stage of microalgae cultivation in order to obtain higher pelletization ratio. Results are shown in FIG. 11.

Example 5 - Treatment of concentrated municipal wastewater

Chlorella vulgaris UMN235 were cultivated for 3-4 days, then Aspergillus oryzae spores were added into culture medium and co-cultured for two days, producing fungi-algae pellets. The fungi-algae pellets were washed twice using distilled water before wastewater treatment experiment.

The fungi-algae pellets were cultured on concentrated municipal wastewater for one day to test its capability to remove pollutants. The change of NH4-N, total phosphorus, COD, and total nitrogen in concentrated municipal wastewater is depicted in FIG. 13A-D, respectively. NELt-N was 100% removed, after one-day cultivation (FIG. 13 A). Total phosphorus was reduced from 53.0 mg/L to 5.00 mg/L and remained at the reduced level until the end of cultivation (FIG. 13B). Maximal removal efficiency reached round 90.0 %, and was thus much higher than those reported in other studies using municipal wastewater (Cheng and Liu, 2002 Trans. ASAE 45:799-805; Lau et al, 1995 Environ Pollution 89:59-66; Woertz et al., 2009 J. Envir. Engrg., 135(11):1115-1122). This efficiency suggests the synergetic treatment effect of algae-fungi pellets rather than algae or fungi alone. The concentration of COD, which is commonly used to indirectly measure the amount of organic compounds in wastewater, decreased from 1,600 mg/L to 600 mg/L in 24 hours (FIG. 13C). Total nitrogen dropped from 90.0 mg/L to 35.0 mg/L in one day (FIG. 13D). As shown in Table 3, most of the nitrogen in centrate was ammonium, which was readily available to algae.

Table 3. Characteristics of the wastewater

Parameter Concentration (mg/L) Parameter Concentration (mg/L)

COD 1600 ± 53.3 P04 3" P 53.3± 6.9

TOC 543±26.30 NH3-N 53.7±2.1 pH 6.35 ± 0.19 TK 90.0± 7.3

NO3-N 0.41 ± 0.29 NO2-N <0.04

Al 0.094 As 0.431

B 0.363 Ba 0.082

Be 0.001 Ca 128.49

Cd 0.013 Co 0.045

Note: pH in the concentrated municipal wastewater was 6.35 Example 6 - Treatment of swine manure wastewater

Fungi-algae pellets prepared as described in Example 5 were further applied to treat 20-fold diluted swine manure wastewater. NH4-N, total phosphoras, COD, and total nitrogen in 20-fold diluted swine wastewater for the two days batch culture is depicted in FIG. 14A-D, respectively. The NH4-N, total phosphorus, and COD dropped from 90 mg/L to 65 mg/L, from 2.00 mg/L to 0.30 mg/L, and from 1000 mg/L to 280 mg/L, respectively, in 48 hours. Total nitrogen was reduced from 130 mg/L to 70.0 mg/L in 24 hours.

Nutrient removal rate was higher in treatment of municipal wastewater compared with treatment of swine manure wastewater. As shown in Table 3 and Table 4, the pH in concentrated municipal wastewater was 6.35 while pH was 9.02 in 20-fold digested swine manure wastewater. When fungi-algae pellets were grown in culture medium, in which pH was above 7.4, the sphere structure of fungi-algae complex was less stable and more easily broken into small pieces (data not shown), which may partially explain the lower efficiency of nutrient removal when growing on diluted swine manure wastewater. However, if only considering the removal efficiency in two days, these results were much better than our earlier data (Wang et al., 2010 Bioresour Technol. 101 :2623-2628) and others in the literature which used algae alone for wastewater treatment. Table 4. Characteristics of the swine manure wastewater after 20-fold dilution

Nutrients concentration (mg/L)

COD TN NH4-N N03-N N02-N TP

1000 130 90 2.00

Note: pH was 9.02 for 20-fold

After overnight culture in fungi-algae pellets, the color of the swine manure became much more clear than that of initial (FIG. 15). The complete disclosure of all patents, patent applications, and publications, and electronically available material (including, for instance, nucleotide sequence submissions in, e.g., GenBank and RefSeq, and amino acid sequence submissions in, e.g., SwissProt, PIR, PRF, PDB, and translations from annotated coding regions in GenBank and RefSeq) cited herein are incorporated by reference in their entirety. In the event that any inconsistency exists between the disclosure of the present application and the disclosure(s) of any document incorporated herein by reference, the disclosure of the present application shall govern. 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.

Unless otherwise indicated, all numbers expressing quantities of components, molecular weights, and so forth used in the specification and claims are to be understood as being modified in all instances by the term "about." Accordingly, unless otherwise indicated to the contrary, the numerical parameters set forth in the specification and claims are approximations that may vary depending upon the desired properties sought to be obtained by the present invention. At the very least, and not as an attempt to limit the doctrine of equivalents to the scope of the claims, each numerical parameter should at least be construed in light of the number of reported significant digits and by applying ordinary rounding techniques.

Notwithstanding that the numerical ranges and parameters setting forth the broad scope of the invention are approximations, the numerical values set forth in the specific examples are reported as precisely as possible. All numerical values, however, inherently contain a range necessarily resulting from the standard deviation found in their respective testing measurements.

All headings are for the convenience of the reader and should not be used to limit the meaning of the text that follows the heading, unless so specified.

Claims

What is claimed is:
1. A method comprising:
adding at least one filamentous fungus to a culture of microalgae;
growing the culture of microalgae under conditions effective to:
produce a co-culture of the microalgae and the filamentous fungus, and form cell pellets; and
harvesting the cell pellets.
2. The method of claim 1 wherein the microalgae comprises at least one autotroph.
3. The method of claim 2 wherein the autotroph comprises a member of the genus Chlorella, a member of the genus Dunaliella, a member of the genus Scenedesmus, a member of the genus Schizochytrium, or a member of the genus Ourococcus.
4. The method of claim 2 or claim 3 wherein the conditions comprise a pH of no more than pH 7.
5. The method of claim 4 wherein the pH is no less than pH 3.
6. The method of any one of claims 2-5 wherein the filamentous fungus comprises spores, and the spores are added to a final concentration of at least 103 spores per liter.
7. The method of claim 6 wherein spores are added to a final concentration of at most
10 spores per liter.
8. The method of one of claims 2-7 wherein the filamentous fungus is added to the microalgae culture when the microalgae culture comprises a cell density of at least 4 x 106 cells per milliliter and no more than 10 x 106 cells per milliliter.
9. The method of claim 8 wherein the filamentous fungus is added to the microalgae culture when the microalgae culture comprises a cell density of at least 6 x 106 cells per milliliter and no more than 8 x 106 cells per milliliter.
10. The method of any one claims 2-7 wherein the filamentous fungus is added to the microalgae culture during mid-log phase.
11. The method of claim 1 wherein the microalgae comprises at least one heterotroph.
12. The method of claim 11 wherein the heterotroph comprises a member of the genus Chlorella, a member of the genus Crypthecodinium, a member of the genus Scenedesmus, a member of the genus Chlamydomonas, a member of the genus Micractinium, or a member of the genus Euglena.
13. The method of claim 10 or claim 12 wherein the conditions comprise a pH of no less than pH 3 and no greater than pH 8.
14. The method of claim 13 wherein the conditions comprise a pH of from pH 5 to pH 6.
15. The method of any one of claims 11-14 wherein the conditions comprise no more than 25 g/L of sugar as a carbon source.
16. The method of claim 15 wherein the conditions comprise no less than 0.25 g/L of sugar as a carbon source.
17. The method of any one of claims 11-14 wherein the filamentous fungus comprises spores and the spores are added to a final concentration of at least 103 spores per liter.
18. The method of any one of claims 11-18 wherein the filamentous fungus is added to the microalgae culture when the microalgae culture comprises a cell density of at least 1 x 106 cells per milliliter and no more than 8 x 106 cells per milliliter.
19. The method of claim 18 wherein the filamentous fungus is added to the microalgae culture when the microalgae culture comprises a cell density of at least 2 x 106 cells per milliliter and no more than 6 x 106 cells per milliliter.
20. The method of any one of claims 11-19 wherein the filamentous fungus is added to the microalgae culture during early log phase.
21. The method of any preceding claim wherein the filamentous fungus comprises at least one fungal cell.
22. The method of any preceding claim wherein the filamentous fungus comprises a member of the genus Aspergillus, a member of the genus Mucor, a member of the genus Phanerochaete, a member of the genus Leucogyrophana, a member of the genus Rhizopus, a member of the genus Absidia, a member of the genus Penicillium, a member of the genus Trichoderma, or a member of the genus Fusarium.
23. The method of claim 22 wherein the filamentous fungus expresses cellulase.
24. The method of claim 22 or claim 23 wherein the filamentous fungus produces oil.
25. The method of claim 24 wherein the filamentous fungus produces oil that is also produced by the microalgae.
26. The method of any preceding claim wherein harvesting the pellets comprises filtering at least a portion of the pellets from the culture.
27. The method of any preceding claim further comprising extracting a bioproducts from the harvested pellets.
28. The method of claim 27 wherein the bioproducts comprises oil.
29. A method comprising:
adding at least one filamentous fungus to a culture of microalgae, wherein the culture comprises wastewater; and
growing the culture of microalgae under conditions effective to produce a co-culture of the microalgae and the filamentous fungus comprising cell pellets that comprises the microalgae and the filamentous fungus.
30. The method of claim 29 wherein the filamentous fungus comprises spores, and the spores are added to a final concentration of at least 10 spores per liter.
31. The method of claim 30 wherein spores are added to a final concentration of at most
7
10 spores per liter.
32. The method of one of claims 29-31 wherein the filamentous fungus is added to the microalgae culture when the microalgae culture comprises a cell density of at least 4 x 106 cells per milliliter and no more than 10 x 106 cells per milliliter.
33. The method of claim 32 wherein the filamentous fungus is added to the microalgae culture when the microalgae culture comprises a cell density of at least 6 x 106 cells per milliliter and no more than 8 x 106 cells per milliliter.
34. The method of any one claims 29-33 wherein the filamentous fungus is added to the microalgae culture during mid-log phase.
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