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A novel method to harvest microalgae via co-culture of filamentous fungi to form cell pellets
Bioresource Technology 114 (2012) 529–535
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A novel method to harvest microalgae via co-culture of filamentous fungi to form cell pellets Jianguo Zhang, Bo Hu ⇑ Department of Bioproducts and Biosystems Engineering, University of Minnesota, USA
a r t i c l e
i n f o
Article history: Received 1 November 2011 Received in revised form 13 March 2012 Accepted 17 March 2012 Available online 28 March 2012 Keywords: Microalgae cultivation Chlorella vulgaris Filamentous fungus Cell pelletization Microalgae cell harvest
a b s t r a c t While current approaches have limitations for efficient and cost-effective microalgal biofuel production, new processes, which are financially economic, environmentally sustainable, and ecologically stable, are needed. Typically, microalgae cells are small and grow individually. Harvest of these cells is technically difficult and it contributes to 20–30% of the total cost of biomass production. A new process of pelletized cell cultivation is described in this study to co-culture a filamentous fungal species with microalgae so that microalgae cells can be co-pelletized into fungal pellets for easier harvest. This new process can be applied to microalgae cultures in both autotrophic and heterotrophic conditions to allow microalgae cells attach to each other. The cell pellets, due to their large size, can be harvested through sieve, much easier than individual cells. This method has the potential to significantly decrease the processing cost for generating microagal biofuel or other products. Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction Producing biofuels and bioproducts via microalgae is promising; however, new technical processes must be developed to capitalize on the economically feasible potential of accumulating bioproducts and biofuel inside microalgae biomass. For instance, many microalgae (e.g., Chlorella 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 et al., 2009; HerediaArroyo et al., 2011). 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 cost. Open-pond cultures are only commercialized to produce some value-added health food supplements such as feed and reagents (Chisti, 2007). Photobioreactors are developed to achieve higher productivity and to maintain monoculture of algae;
⇑ Corresponding author. Address: Department of Bioproducts and Biosystems Engineering, University of Minnesota, 316 BAE, 1390 Eckles Ave., Saint Paul, MN 55108-6005, USA. Tel.: +1 612 625 4215; fax: +1 612 624 3005. E-mail address: [email protected] (B. Hu). URL: http://bohu.cfans.umn.edu/ (B. Hu). 0960-8524/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2012.03.054
however, the unit cost of microalgae production in these enclosed photobioreactors are actually much higher than those achievable in open-pond cultures despite photoreactors’ higher biomass concentration and better control of culture parameters (Lee, 2001). The algae cell harvest from cultivation broth has always been one of the major obstacles for the algae-to-fuel approach. Microalgae cell harvest is technically challenging, especially considering the low concentration (typically in the range of 0.3–5 g/L), the small size of the oleaginous algal cells (typically in the range of 2–40 lm), and their similar density to water (Li et al., 2008). 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 usually accounts for 20–30% of the total costs of production (Uduman et al., 2010a). How to harvest microalgae cells from cultivation broth is dependent on the characteristics of the microalgae, such as size and density (Olaizola, 2003); and harvesting usually requires a separate step after the cell cultivation. All of the available harvest approaches, which include flocculation, flotation, centrifugal sedimentation, and filtration, have limitations for efficient, costeffective production of biofuel (Shelef et al., 1984). 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 filtration and gravitational sedimentation are widely applied in wastewater treatment facilities to harvest relatively large (>70 lm) microalgae such as Coelastrum and Spirulina. However,
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they cannot be used to harvest algae species approaching bacterial dimensions (<30 lm) like Scenedesmus, Dunaliella, and Chlorella (Brennan and Owende, 2010), to which most oleaginous microalgae species belong. Centrifugation is widely used to recover microalgae biomass, especially small-sized algae cells; however, its application is restricted to algae cultures for high-value metabolites due to intensive energy needs and high equipment maintenance requirements. While flocculation is used to harvest small-sized microalgae cells, it is a preparatory step to aggregate the microalgae cells and increase the particle size so that other harvesting methods such as filtration, centrifugation, or gravity sedimentation can be applied (Molina Grima et al., 2003). Several flocculants have been developed to facilitate the aggregation of microalgae cells, including multivalent metal salts like ferric chloride (FeCl3), aluminum sulphate (Al2(SO4)3) and ferric sulphate (Fe2(SO4)3), and organic polymers such as chitosan and modified starch (Li et al., 2008). Chemical flocculation can be reliably used to remove small algae cells from pond water by forming large (1–5 mm) sized flocs (Sharma et al., 2006). However, besides the high cost of chemical flocculants and possible pollution effects that may generate, the chemical reactions are highly sensitive to pH and the high doses of flocculants required produce large amounts of sludge and may leave a residue in the treated effluent. In summary, most technologies including chemical and mechanical methods greatly increase operational costs for algal production and are only economically feasible for production of high-value products (Park et al., 2011). Besides traditional methods mentioned above, there are several new technology developments in this field. DOE-ARPA-E recently funded a research project for Algae Venture Systems (AVS) to develop a Harvesting, Dewatering and Drying (AVS-HDD) technology using the principles of liquid adhesion and capillary action to extract water from dilute microalgae solutions. Attached algal culture systems have been developed for growing microalgae on the surface of polystyrene foam to simplify the cell harvest (Johnson and Wen, 2010; Wilkie and Mulbry, 2002). New bioflocculants, which are more environmentally friendly, are also proposed to address the cost and environmental concerns for current flocculation methods (Uduman et al., 2010a). These methods are innovative and will decrease the harvest cost to some extent if developed successfully, but require heavy investments on equipment and chemical supplies. Enhancing natural algae aggregation to encourage simple gravity settling or filtration appears to be the most 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 use (Uduman et al., 2010a). It will also 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 (50–200 lm), and their cell aggregation can be achieved through nitrogen limitation and CO2 addition (Park et al., 2011); however, most of these microalgae species are not oleaginous species. Methods to enable oleaginous microalgae aggregate during their cultivation are strategically and urgently needed to develop efficient and economic means of biofuel production. In submerged cultures, filamentous microorganisms, including some species of molds and bacteria, tend to aggregate and grow as pellets/granules. These fungal cell pellets are 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, 2008; Hu et al., 2009). Fungal cell pelletization can significantly decrease the viscosity of the fermentation broth; therefore it has been widely researched to increase the cultural performance on the mixing and mass transfer properties. Other advantages of cell pelletization to the micro-oil production process include the ease of
harvesting cells and of re-using pond water (Johnson and Wen, 2010; Xia et al., 2011). Conditions for cell pelletization seem to be strain-specific and that not all the filamentous fungal strains can form pellets during their growth. A recent study at University of Minnesota showed that Mucor circillenous is relatively difficult to form pellets during their cell growth; however, with induction of CaCO3 powder in the early stage of its cultivation, cell pellets can be formed homogeneously, lasting for the entire cultivation cycle. Changing operational conditions during cell cultivation was found to be able to induce fungal cells to aggregate and form pellets (Xia et al., 2011). This method avoids traditional approaches that use CaCO3 powder or other nuclei to induce the fungal pelletization (Liao et al., 2007; Liu et al., 2008a) which are costly and cause solid waste disposal issues. Pelletization is more widely seen in the fungal fermentation process where the microorganisms are filamentous. However, most oleaginous microalgae are not filamentous and oleaginous microalgae pelletization has not been seen in a complete review of current literature. In this study, a preliminary study was conducted to inoculate filamentous fungal spores when culturing mixotrophic green algae C. vulguris and it was found that pellets clearly formed within two days of culture. The microalgae cells, aggregated together with fungal cells, were immobilized in the pellets. This paper described this new technology which uniquely addresses the cell harvest of microalgae and has the potential to greatly reduce the algae biofuel cost. While pelletization and granulation have already been developed for commercial production of numerous products to increase culture conditions (e.g., mass transfer and oxygen transfer), this paper is the first study to investigate co-culturing microalgae with filamentous fungi for possible direct pelletization of microalgae to lower the overall cost of the process. 2. Materials 2.1. Microbial species Filamentous fungi Aspergillus niger Ted S-OSU was tested in this study for its co-cultivation with microalgae, as well as other filamentous fungal strains, which were all purchased from ATCC. C. vulguris was the model microalgae species to test its co-pelletization with filamentous fungi. Previous research has revealed that C. vulguris 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). 2.2. Cell cultivation 2.2.1. Cultivation medium Autotrophic medium A (per L): KNO3 1 g, KH2PO4 0.075 g, K2HPO4 0.1 g, MgSO42H2O 0.5 g, Ca(NO3)24H2O 0.0625 g, FeSO47H2O 0.01 g, Yeast extract 0.5 g, A5 1 ml. A5 (L): H3BO3 2.86 g, Na2Mo42H2O 0.039 g, ZnSO47H2O 0.222 g, MnCl24H2O 1.81 g, CuSO45H2O 0.074 g, CoCl2 0.03 g. Heterotrophic medium B (per L): Potato dextrose broth 12 g, Glucose 15 g. 2.2.2. Cultivation of seed C. vulgaris An autotrophic flask culture of C. vulgaris was maintained in the lab to provide algae seeds for the co-cultural experiments. A 4 L flask was filled with 3 L culture medium A with a magnetic stir (100 rpm) and four fluorescent lights were provided outside the flask to culture the algae cells at 25–27 °C. Fresh cultural medium was routinely added into the flask to maintain the active cell growth.
J. Zhang, B. Hu / Bioresource Technology 114 (2012) 529–535
2.2.3. Preparation for seed fungal spores Fungal spore solution was stored at 70 °C with 25% of glycerol. The stored spores were activated by cultivation on the petri dish with medium B and 2% agar at 27 °C for 7 days. Sterilized DI (10 ml) water was applied on the culture to harvest the spores and the spore solution was used as the inoculation for the co-culture after the number of spores in the solution was counted. 2.2.4. Flask co-culture of filamentous fungi with autotrophic microalgae Prepare 100 ml culture medium A in 250 mL flasks, and then adjust pH by adding 2 mol/L HCl or 1 mol/L NaOH to pH 4.0. Transfer 20 ml microalgae seed broth into these 250 ml flasks. Finally, inoculate fungal spores into medium A to reach certain spore concentration (7.6 10E6/L). These flasks were cultured at 27 °C, 150 rpm for 3 days. 2.2.5. Cellulase degradation of C. vulgaris cell wall Inoculate with 7.6 10E6/L spores of A. niger in a flask with 10 times dilution of medium B and culture at 27 °C, 100 rpm for 3 days to obtain the fungal cultivation broth. The broth was filtered with sterilized filters to remove the fungal cells. C. vulgaris cell broth (1.76 10E10/L, 10 ml) was harvested by centrifuge, then inoculated into 4.5 ml of fungal supernatant solution together with 0.5 ml 500 mmol/L phosphate buffer and 100 ll cellulase (C8546, Sigma, St. L MO. USA), and reacted at 50 °C, for 1 h in water bath (Model 25, Precision scientific Inc. Chicago, IL, USA). 2.2.6. Flask co-culture of filamentous fungi with heterotrophic microalgae Prepare 100 ml culture medium B in 250 mL flasks, and then adjust pH to 5.0. Transfer 20 ml microalgae seed broth and inoculate fungal spores (7.6 10E6/L) into these 250 ml flasks. These flasks were cultured at 27 °C, 150 rpm for 3 days. 2.2.7. Co-culture of C. vulgaris and A. niger in a bioreactor Prepare 1 L of medium B together with 1 L water in the Bioreactor (Bioflo 110, New Brunswick, MA USA), and pH was adjusted to 5.0 after sterilization. Inoculate with C. vulgaris cultivation broth and A. niger spores (7.6 10E6/L) into the bioreactor. The culture condition was 27 °C, 200 rpm. The pH value was maintained by alkaline (1 mmol/L NaOH) addition automatically and the air feeding rate is 1 vvm. 2.3. Analysis At the end of each flask culture, pour all the pellets of the cocultivation of microalgae and filamentous fungi from each flask into the Petri dish (discard extra liquid if necessary) 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. The microalgae cell numbers in the supernatant were measured after diluting the supernatant multiple times until the cell numbers can be counted under microscope. Total microalgae algae cells were measured after the cultivation broth was blended completely to break the structure of pellets in order to release microalgae cells from pellets and then diluting the cell concentrations until the cell numbers can be counted under microscope. 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. Glucose concentration was estimated by using DNS method (Xiong et al., 2008). The total biomass concentration of the flask cultures were determined by the dry cell biomass. Dry biomass was then processed with 10 ml mixture of chloroform and metha-
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nol (2:1 v/v) to extract the microbial oil. Finally fatty acid methyl ester (FAME) was generated via transesterification reaction at 90 °C water bath (Model 25, Precision Scientific, Chicago, IL, USA) for 90 min with concentrated sulfuric acid as the catalyst. FAME in chloroform was carefully collected and subjected to GC after filtration (0.22 lm). The GC was equipped with a flame ionization detector and a HP-5 capillary column. The oven temperature was 80 °C, held for 5 min, raised to 290 °C at a rate of 4 °C/min, and held at 290 °C for 5 min, 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/min. 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, MO). All values were means of triplicate determinations. 3. Results and discussion 3.1. Description of pellets formed in the co-cultivation of C. vulgaris and A. niger Microalgae cells were suspended in the cultivation broth and the solution can be relatively stable and homogeneous. With the inoculation of fungal spores and addition of organic nutrients, the algae solution lost most of its green color within two days of cultivation, and turned transparent, indicating the majority of microalgae cells were pelletized. Pellets were formed in the cultivation broth with almost similar density as the cultivation solution. The pellets are spherical, packed, relatively homogeneous in size, and will settle down once the shaking stops. Significantly different from the color of pellets formed by fungi itself; the pellets in the co-cultivation are green, instead of white or grey. The microscopic image showed that the skeleton structure of the pellets is still filamentous fungal cells; while green algae cells are entrapped inside the fungal mycelia and attached on the fungal cell surface (Supplementary Figure). Since these microalgae cells are not filamentous, natural algae aggregation, simple gravity settling or filtration cannot be applied to these microalgae species for harvesting their cell biomass. Having a separate harvesting step, no matter by using flocculation, centrifugation or other methods, is costly and environmental non-friendly (Olaizola, 2003). Co-culturing filamentous fungi with microalgae cells enabled these microalgae cells to pelletize, and these cell pellets (size at 2–5 mm) can be harvested by a simple filtration with sieves, significantly easier than harvesting individual cells. The process is a totally different method compared to chemical flocculation. Instead of adding a chemical flocculent, which may be expensive, needs large quantity and/or leaves toxic residue to the harvested microalgae pat, fungal spores or hyphae were inoculated into the microalgae cultivation broth and were cultured together with microalgae cells for the harvest purpose. The pellets formed in this co-culture are more packed than ‘‘fluffy’’ floc, which gives more advantages to pellets for cell harvest (Olaizola, 2003). 3.2. Test of co-cultivation of C. vulguris with different filamentous fungal strains Aspergillus oryzae, a commercial fungal strain for cellulase production, can easily form relatively large cell pellets, which are stable and homogenous, during the cell growth without any inducing approaches (Table 1). Same phenomena also showed on many other fungal strains such as ATCC 11730, RLG 9902 and Aspergillus flavus. Mortierella isabellina, on the other hand, is nearly incapable of forming stable pellets (Table 1). During a cell culture in flasks,
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Table 1 Photos of co-pellets of fungal species and Chlorella vulgaris. Fungal species
M. isabellina
M. circinelloides
A. flavus
Fungal Pellets
Co-pellets
RLG 9902
Ted S-OSU
ATCC 9642
No photo
No pellets
pellets were formed in the first day of culture of M. isabellina; while all the pellets disappeared after 4 days of cultivation. M. circillenous is also relatively difficult to form pellets during their cell growth; however, with induction of CaCO3 powder in the early stage of its cultivation, cell pellets can be formed homogeneously, lasting for the entire cultivation cycle. Key operational conditions were revealed in the previous research to induce the pelletization of filamentous fungi M. circillenous (Xia et al., 2011). It was discovered that by only changing conditions during cell cultivation can force fungal cells to aggregate and form pellets. This method avoids traditional approaches that use CaCO3 powder or other nuclei to induce the fungal pelletization, which are costly and cause solid waste disposal issues (Liao et al., 2007; Liu et al., 2008a). Conditions for cell pelletization seem to be strain-specific and that not all the filamentous fungal strains can form pellets during their growth. Different species of fungal spores were tested for their co-pelletization capability with the green algae C. vulguris, as shown in Table 1 and Table 2. Several fungal species, such as ATCC 11730, RLG 9902 and A. flavus, can form pellets that can entrap almost all of the individual microalgae cells, resulting in a clear co-culture broth of those strains (Table 2). As for Mucor circinelloides (Table 2), there are still some C. vulgaris cells in supernatant, although green pellets also formed in their cultures. For Phanerochaete chrysosporium and A. oryzae, even though pellets were still formed in the coculture with microalgae, these pellets were either grey or milky while with almost no algae cells entrapped inside the pellets. There were no harvest effects on these cultures because microalgae cells were predominantly in the supernatant. Strains such as M. isabellina could form unstable pellets on its own; but it could not form any pellets when co-cultured with C. vulguris. The formation of fungal pellets and co-pellets were depending on the strains as well
as the cultivation conditions. Numerous researches have showed some specific methods on inducing the fungal pelletization (Liao et al., 2007; Liu et al., 2008a; Xia et al., 2011), however, there is no generally applicable strategy, especially for the co-pelletization of fungal and algae strains. Detailed study on the co-cultivation conditions is needed in each strain combination in order to induce the pellets formation of these cells. Depending on the application of microalgae cells, different filamentous fungal species can be co-cultured to pelletize microalgae cells for easier harvest. For instance, if the microalgae cells can generate cellulose on their cell wall, co-cultivation of a cellulaseproducing fungus with this microalgae will be beneficial because fungal cells can support themselves by partially degrading the cell wall of microalgae to generate sugars as well as pelletize microalgae cells into pellets for easer harvest. In this case, no external sugars are needed for this process. If the primary purpose of culturing microalgae is to generate microbial lipids, co-culturing an oleaginous fungus can be similarly beneficial. This is 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 high content of lipids that can be extracted for biodiesel production, co-culturing these oleaginous fungi cells will not only help with the oil-producing purpose, but will also facilitate the pelletization of microalgae cells for easier harvest. 3.3. Test of co-cultivation of A. niger with C. vulguris on different growth mode C. vulguris can grow on both autotrophic and heterotrophic growth modes and spores of A. niger were inoculated to both
Table 2 Co-pelletization of fungal species and C. vulgaris after 24 h heterotrophic condition. Fungal species
Pellets
Initial cell number (10 E9/L)
Mortierella isabellina Aspergillus flavus Aspergillus niger Aspergillus versicolor (ATCC 11730) Corynebacterium nephridii (Bacteria, ATCC 11425) Pycnoporus sanguineus (ATCC 24598) Aerococcus viridans (ATCC 9642) Phanerochaete chrysosporium (ATCC 24725) Trametes lilacinogilva (ATCC 46156) Pycnoporus sanguineus (ATCC 18395) Leucogyrophana arizonica (RLG 9902) Mucor circinelloides Aspergillus oryzae Control (No fungi)
p p p
1.80 2.65 3.30 1.85 2.60 2.40 3.95 2.40 2.75 2.35 2.80 2.75 1.75 2.45
p p⁄ p p p⁄
Algae in supernatant (10 E9/L) in 24 h 0 0.65 0 (48 h)
0.25 White pellets
0 6.95 White pellets 20.80
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70 60
Harvest ratio (%)
growth conditions to study the co-pelletization of these strains as shown in Table 3. The initial microalgae cell concentration both started at 6.9 10E9/L with medium A for the autotrophic cultivations (no sugar addition) and medium B for the heterotrophic cultivations. The microalgae final cell concentration reached at (10.5 ± 3.16) 10E9/L for the control tests with no fungal spores inoculated. For the co-culture at autotrophic conditions, the suspended algae concentration in the supernatant was about same as the control ((10.7 ± 0.78) 10E9/L), while a large amount of the microalgae cells were co-pelletized with the fungi, which enlarged the total microalgae cells in the co-culture almost three times more the control. The maximum microalgae cell concentration in autotrophic cultivations is usually limited by the mutual shading effects, where microalgae cells in the pond water cannot access to the sun light if the final algae cells reach to the maximum concentration (Lee and Palsson, 1994). The experiments showed that the co-cultivation of fungal cells actually stimulated the overall microalgae cultivation and dramatically increased the total algae production. Pelletization creates an inhomogeneous cultural system that large amount of algae cells were immobilized inside the pellets while the suspended algae cell density in the culture broth could remain at low level, which allows the penetration of light. For the co-culture with autotrophic microalgae, around 60% of algae cells were pelletized with fungal cells (Table 3 and Fig. 1). No sugar was added to the autotrophic co-cultivation to support the germination and growth of A. niger. Due to the low concentration of fungal cells, there was not any measurable cellulase activity in the co-cultivation broth of autotrophic microalgae and fungi (Data not shown). However, it was confirmed by many researchers that A. niger can produce cellulase during their cell growth (Acharya et al., 2008) and the cellulase is excreted in the fermentation broth to degrade the cellulose to support the cell growth. Test of degradation of C. vulgaris cell wall by external cellulase (Table 4) showed that sugar concentration increased from 1.74 to 2.36 g/L. It was also confirmed by other researchers 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) and also to facilitate the lipid extraction (Fu et al., 2010). The carbohydrates composition of microalgae is mainly polysaccharide, entrapped in the cell walls with up to 70% dry weight. The composition varies among microalgae strains, and cellulose is reported as the main structural component of the cell wall for most microalgae species. 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 the oil extraction (Fu et al., 2010; Yin et al., 2010). This experiment may provide some supports that fungal cells grew on the microalgae cell degradation, but further research is needed in the future to provide direct evidence. For the heterotrophic co-culture, the supernatant microalgae cell concentration reached to (72.50 ± 21.21) 10E9/L and the total microalgae cells reached to (95.75 ± 5.30) 10E9/L. Since algae
50 40 30 20 10 0 Autotrophic condition
Heterotrophic condition
Fig. 1. Harvest ratio of C. vulgaris at autotrophic and heterotrophic growth condition.
Table 4 Test of cellulase degradation of C.vulgaris cell wall. Cellulase addition
Initial
Fungal supernatant
Microalgae solution
Sugar concentration (g/L)
2.5
1.74 ± 0.03
2.36 ± 0.07
growth in heterotrophic conditions relied on the available external sugar, instead of light, microalgae cell concentration increased dramatically, with relatively smaller portion being pelletized (Fig. 1). Total cell biomass in the heterotrophic co-cultivation was significantly higher than other conditions because sugars were provided and consumed both by the fungus and microalga. A. niger, which only generated 17.69% of its biomass as oil (measured based on Fatty Acid Methyl Ester FAME), is not an oleaginous strain (Table 3). This explained that the oil content (FAME) of total biomass from co-cultivations was lower than the purely cultured C. vulguris. Although the fungus’s fatty acid profile was significantly different from C. vulguris (with relatively higher content of C18:2, less content of C16:1), the fatty acid profile of total biomass from co-cultivations were not much different than pure culture of C. vulguris (Fig. 2). The predominant fatty acid components of C. vulguris include C16:0, C16:1, C18:0, C18:1 and C18:2. Since A. niger cells did not contribute significantly to the total oil extracted from cocultivation biomass, the fatty acid composition was primarily determined by the algae. When co-culturing an oleaginous filamentous fungus, the fungal cells will contribute to the total oil production and the fatty acid profile of total biomass may change more significantly.
Table 3 Co-culture of A. niger with C. vulguris at different growth. C. vulgaris Volume (ml) Initial microalgae (10E9/L) Initial spores (10E6/L) Microalgae in supernatant (10E9/L) Total microalgae cell (10E9/L) Biomass (g/L) Fatty acid (%) Fatty acid (mg/L)
100 6.9 10.5 ± 3.16 10.5 ± 3.16 0.142 ± 0.01 37.40 ± 1.50 53.11 ± 1.51
A. niger
Autotrophic
Heterotrophic
100 0 7.6 0 0 0.215 ± 0.02 17.69 ± 3.08 38.03 ± 10.05
100 6.9 7.6 10.7 ± 0.78 29 ± 3.50 0.304 ± 0.02 32.58 ± 3.26 99.04 ± 16.30
100 6.9 7.6 72.50 ± 21.21 95.75 ± 5.30 0.968 ± 0.129 28.50 ± 1.93 275.88 ± 52.40
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Chrolella vulgaris Aspergillus niger Autotrophic condition Heterotrophic condition
40
Percentage (%)
30
20
10
0 C14
C15 C16:1 C16 C18:3 C18:2 C18:2 C18:1 C18
C20
C22
Fatty acid Fig. 2. Fatty acid profiles of co-cultivated C. vulguris and A. niger.
Experiments in the bioreactor containing C. vulgaris and A. niger showed typical filamentous growth, which pure culture of microalgae C. vulgaris should not have, although some cells form pellet like aggregates while other cells attached on the mixer or other surface inside the bioreactor. With the co-culture of fungal cells, the supernatant microalgae cells dropped to 0.2E9/L from 7.8E9/L at initial inoculations while the total microalgae reached to 10.4 10E9/L, which means that 98.1% microalgae cells were entrapped in the fungal clumps (Fig. 3). Factors to affect the pelletization process and how to scale up the co-cultivation should be investigated in details in the future.
3.4. Mechanism of co-pelletization of fungal and microalgae cells
12
100
10
DO (%)
80
8 60 6 40 4 DO Glucose Microalgae
20
2
0
0 0
10
20
30
40
50
Time (h) Fig. 3. Co-culture of C. vulgaris and A. niger in the bioreactor.
Microalgae supernatant (10E9/L) Glucose (g/L)
Fungal cell pelletization has been studied for decades and widely applied in the industrial processes to produce organic acid, pharmaceuticals, enzymes and other high-valued fermentation products, and wastewater treatment to remove pollutants. The research results showed conditions for cell pelletization seem to be strain-specific and that not all the filamentous fungal strains can form pellets during their growth. The capability of cell aggregation of filamentous fungal cells is predominately rooted to the production of hydrophobic proteins (referred to as hydrophobins), a family of low molecular weight amphipathic proteins. For those dimorphic fungal species, hydrophobin were detected on the hyphal surface while missing when the cells were grown in the yeast
form (Linder, 2009). One of the biological functions of these hydrophobic proteins is to coordinate the adherence of hyphae to solid substrates, which facilitates biofilm formation (attaching on a solid surface) and cell pelletization/granulation/aggregation (attaching on each other) (Feofilova, 2010). Numerous theories have been proposed to explain the fungal pelletization process, which can be categorized into two types of mechanisms (Liu et al., 2008b; Ryoo and Choi, 1999). The first one is coagulative mechanism, where spores coagulate in the early stage of cultivation and develop into pellets through their intertwining hyphae. The second one is non-coagulative mechanism, where the spores germinate into hyphae, and then intertwine into pellets. Many genus of fungus such as Aspergillus, basidiomycete, Phanerochaete chrysosporium follows the coagulative mechanism where fungal spores conglomerate at an early stage of development and then each pellet may arise from each spore aggregate. While other genus of fungus, such as Rhizopus spp, Mucor spp. Penicillium chrysogenum, follows the non-coagulative mechanism. The experiment results showed that co-pelletization of fungal and microalgae cells only occurred on certain filamentous fungal strains (Table 2) and the detailed mechanisms are still not clear. The cell interaction information could have significant potential impacts in inducing the co-pelletization so that co-pelletization conditions can be refined for the large-scale production of microalgae-fungal pellets as a biofuel feedstock. One microscopic photo taken on the co-pellets of A. niger hyphae and C. vulgaris grinded by a blender clearly showed that the microalgae cells were attracted on the filamentous hyphae (Supplementary Figure). The mutual attraction of microalgae and filamentous cells may be attributed to several possible reasons, but the primary reason might be due to the surface charge. Most microalgae cells carry negative charges on their cellular surfaces and therefore are capable of forming stable suspensions in the cultivation pond water. The stability of these microalgal suspensions depends on the forces that interact between the cells themselves and between the cells and water. Hence they are considered as hydrophilic bio-colloids (Uduman et al., 2010b). Current flocculation method to harvest microalgae is to add flocculants such as metals ions or organic polymers that have counterions to charge neutralisation and polymer bridging, and then allow cells to aggregate to form floc (Pieterse and Cloot, 1997). Fungal hyphae and mycelia contain polysaccharide with active sites that may enable their surface capability of bioadsorption and also enable the fungal cells to be charged (Tan et al., 2004). In a similar published research study to search for a biological flocculation agent for microalgae cell harvest (Rajab, 2007), A. flavus were grown to form cell pellets, and then applied to the microalgae cultural broth to flocculate the microalgae cells. The fungal cell pellets did absorb the microalgae cells, and their zeta potential measurement revealed the average number for microalgae as 23.7 millivolt while A. flavus as +46.1 mV. The charge difference between the microalgae and fugal cells indicated the capability of A. flavus to capture microalgae, and it provides some clue that surface charge might be the reason of co-pelletization of microalgae and fungal cell cultures.
4. Conclusions This paper studied the possibility to co-culture non-filamentous microalgae cells with filamentous fungi to form cell pellets for easier harvest. The co-culture of filamentous fungi to enable filamentous features on non-filamentous microalgae cultures will potentially provide an innovative process with direct commercial application potential on microalgae production of biofuel and other products. Compared to other algae harvesting methods, this method will potentially decrease harvest costs, avoid second pollu-
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tion from chemical flocculants and stimulate the algae production, although further research is needed to study the detailed pelletization conditions in the larger scale industrial applications. Acknowledgements Dr. Jianguo Zhang’s research was supported by Dr. Bo Hu’s faculty seed money program at University of Minnesota. We appreciate Dr. Jonathan Schilling at University of Minnesota for providing some fungal strains. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.biortech.2012. 03.054. References Acharya, P.B., Acharya, D.K., Modi, H.A., 2008. Optimization for cellulase production by Aspergillus niger using saw dust as substrate. Afr. J. Biotechnol. 7, 4147–4152. Brennan, L., Owende, P., 2010. Biofuels from microalgae – A review of technologies for production, processing, and extractions of biofuels and co-products. Renew. Sustain. Energy Rev. 14, 557–577. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294–306. Fakas, S., Papanikolaou, S., Batsos, A., Galiotou-Panayotou, M., Mallouchos, A., Aggelis, G., 2009. Evaluating renewable carbon sources as substrates for single cell oil production by Cunninghamella echinulata and Mortierella isabellina. Biomass Bioenergy 33, 573–580. Feofilova, E.P., 2010. The fungal cell wall: modern concepts of its composition and biological function. Microbiology 79, 711–720. Fu, C.-C., Hung, T.-C., Chen, J.-Y., Su, C.-H., Wu, W.-T., 2010. Hydrolysis of microalgae cell walls for production of reducing sugar and lipid extraction. Bioresour. Technol. 101, 8750–8754. Harun, R., Danquah, M.K., 2011. Enzymatic hydrolysis of microalgal biomass for bioethanol production. Chem. Eng. J. 168, 1079–1084. Heredia-Arroyo, T., Wei, W., Ruan, R., Hu, B., 2011. Mixotrophic cultivation of Chlorella vulgaris and its potential application for the oil accumulation from non-sugar materials. Biomass Bioenergy 35, 2245–2253. Hu, B., Chen, S.L., 2007. Pretreatment of methanogenic granules for immobilized hydrogen fermentation. Int. J. Hydrogen Energy 32, 3266–3273. Hu, B., Chen, S.L., 2008. Biological hydrogen production using chloroform-treated methanogenic granules. Appl. Biochem. Biotechnol. 148, 83–95. Hu, B., Zhou, X., Forney, L., Chen, S.L., 2009. Changes in Microbial Community Composition Following Treatment of Methanogenic Granules with Chloroform. Environ. Prog. Sustainable Energy 28, 60–71. Johnson, M.B., Wen, Z.Y., 2010. Development of an attached microalgal growth system for biofuel production. Appl. Microbiol. Biotechnol. 85, 525–534.
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Lee, C.G., Palsson, B.O., 1994. High-density algal photobioreactors using lightemitting-diodes. Biotechnol. Bioeng. 44, 1161–1167. Lee, Y.K., 2001. Microalgal mass culture systems and methods: Their limitation and potential. J. Appl. Phycol. 13, 307–315. Li, Y., Horsman, M., Wu, N., Lan, C.Q., Dubois-Calero, N., 2008. Biofuels from microalgae. Biotechnol. Prog. 24, 815–820. Liao, W., Liu, Y., Chen, S.L., 2007. Studying pellet formation of a filamentous fungus Rhizopus oryzae to enhance organic acid production. Appl. Biochem. Biotechnol. 137, 689–701. Linder, M.B., 2009. Hydrophobins: proteins that self assemble at interfaces. Curr. Opin. Colloid Interface Sci. 14, 356–363. Liu, Y., Liao, W., Chen, S., 2008a. Co-production of lactic acid and chitin using a pelletized filamentous fungus Rhizopus oryzae cultured on cull potatoes and glucose. J. Appl. Microbiol. 105, 1521–1528. Liu, Y., Liao, W., Chen, S.L., 2008b. Study of pellet formation of filamentous fungi Rhizopus oryzae using a multiple logistic regression model. Biotechnol. Bioeng. 99, 117–128. Molina Grima, E., Belarbi, E.H., Acién Fernández, F.G., Robles Medina, A., Chisti, Y., 2003. Recovery of microalgal biomass and metabolites: process options and economics. Biotechnol. Adv. 20, 491–515. Olaizola, M., 2003. Commercial development of microalgal biotechnology: from the test tube to the marketplace. Biomol. Eng. 20, 459–466. Park, J.B.K., Craggs, R.J., Shilton, A.N., 2011. Wastewater treatment high rate algal ponds for biofuel production. Bioresour. Technol. 102, 35–42. Pieterse, A.J.H., Cloot, A., 1997. Algal cells and coagulation, flocculation and sedimentation processes. Water Sci. Technol. 36, 111–118. Rajab, A.H., 2007. Micro-Algae Removal in Domestic Wastewater Using Aspergillus Flavus Soft Pellets as A Bio-Coagulant. Universiti Putra Malaysia. Ryoo, D., Choi, C.-S., 1999. Surface thermodynamics of pellet formation in Aspergillus niger. Biotechnol. Lett. 21, 97–100. Sharma, B.R., Dhuldhoya, N.C., Merchant, U.C., 2006. Flocculants—an Ecofriendly Approach. J. Polym. Environ. 14, 195–202. Shelef, G., Sukenik, A., Green, M., 1984. Microalgae harvesting and processing: a literature review, p. Medium: ED; Size: P. 70. Tan, T.W., Hu, B., Su, H.J., 2004. Adsorption of Ni2+ on amine-modified mycelium of Penicillium chrysogenum. Enzyme Microb. Technol. 35, 508–513. Uduman, N., Qi, Y., Danquah, M.K., Forde, G.M., Hoadley, A., 2010a. Dewatering of microalgal cultures: A major bottleneck to algae-based fuels. Journal of Renewable and Sustainable Energy 2. Uduman, N., Qi, Y., Danquah, M.K., Hoadley, A.F.A., 2010b. Marine microalgae flocculation and focused beam reflectance measurement. Chem. Eng. J. 162, 935–940. Wilkie, A.C., Mulbry, W.W., 2002. Recovery of dairy manure nutrients by benthic freshwater algae. Bioresour. Technol. 84, 81–91. Xia, C.J., Zhang, J.G., Zhang, W.D., Hu, B., 2011. A new cultivation method for microbial oil production: cell pelletization and lipid accumulation by Mucor circinelloides. Biotechnology for Biofuels 4. Xiong, W., Li, X.F., Xiang, J.Y., Wu, Q.Y., 2008. High-density fermentation of microalga Chlorella protothecoides in bioreactor for microbio-diesel production. Appl. Microbiol. Biotechnol. 78, 29–36. Yin, L.J., Jiang, S.T., Pon, S.H., Lin, H.H., 2010. Hydrolysis of Chlorella by Cellulomonas sp YJ5 Cellulases and Its Biofunctional Properties. J. Food Sci. 75, H317–H323.
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Bioresource Technology journal homepage: www.elsevier.com/locate/biortech
A novel method to harvest microalgae via co-culture of filamentous fungi to form cell pellets Jianguo Zhang, Bo Hu ⇑ Department of Bioproducts and Biosystems Engineering, University of Minnesota, USA
a r t i c l e
i n f o
Article history: Received 1 November 2011 Received in revised form 13 March 2012 Accepted 17 March 2012 Available online 28 March 2012 Keywords: Microalgae cultivation Chlorella vulgaris Filamentous fungus Cell pelletization Microalgae cell harvest
a b s t r a c t While current approaches have limitations for efficient and cost-effective microalgal biofuel production, new processes, which are financially economic, environmentally sustainable, and ecologically stable, are needed. Typically, microalgae cells are small and grow individually. Harvest of these cells is technically difficult and it contributes to 20–30% of the total cost of biomass production. A new process of pelletized cell cultivation is described in this study to co-culture a filamentous fungal species with microalgae so that microalgae cells can be co-pelletized into fungal pellets for easier harvest. This new process can be applied to microalgae cultures in both autotrophic and heterotrophic conditions to allow microalgae cells attach to each other. The cell pellets, due to their large size, can be harvested through sieve, much easier than individual cells. This method has the potential to significantly decrease the processing cost for generating microagal biofuel or other products. Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction Producing biofuels and bioproducts via microalgae is promising; however, new technical processes must be developed to capitalize on the economically feasible potential of accumulating bioproducts and biofuel inside microalgae biomass. For instance, many microalgae (e.g., Chlorella 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 et al., 2009; HerediaArroyo et al., 2011). 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 cost. Open-pond cultures are only commercialized to produce some value-added health food supplements such as feed and reagents (Chisti, 2007). Photobioreactors are developed to achieve higher productivity and to maintain monoculture of algae;
⇑ Corresponding author. Address: Department of Bioproducts and Biosystems Engineering, University of Minnesota, 316 BAE, 1390 Eckles Ave., Saint Paul, MN 55108-6005, USA. Tel.: +1 612 625 4215; fax: +1 612 624 3005. E-mail address: [email protected] (B. Hu). URL: http://bohu.cfans.umn.edu/ (B. Hu). 0960-8524/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2012.03.054
however, the unit cost of microalgae production in these enclosed photobioreactors are actually much higher than those achievable in open-pond cultures despite photoreactors’ higher biomass concentration and better control of culture parameters (Lee, 2001). The algae cell harvest from cultivation broth has always been one of the major obstacles for the algae-to-fuel approach. Microalgae cell harvest is technically challenging, especially considering the low concentration (typically in the range of 0.3–5 g/L), the small size of the oleaginous algal cells (typically in the range of 2–40 lm), and their similar density to water (Li et al., 2008). 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 usually accounts for 20–30% of the total costs of production (Uduman et al., 2010a). How to harvest microalgae cells from cultivation broth is dependent on the characteristics of the microalgae, such as size and density (Olaizola, 2003); and harvesting usually requires a separate step after the cell cultivation. All of the available harvest approaches, which include flocculation, flotation, centrifugal sedimentation, and filtration, have limitations for efficient, costeffective production of biofuel (Shelef et al., 1984). 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 filtration and gravitational sedimentation are widely applied in wastewater treatment facilities to harvest relatively large (>70 lm) microalgae such as Coelastrum and Spirulina. However,
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they cannot be used to harvest algae species approaching bacterial dimensions (<30 lm) like Scenedesmus, Dunaliella, and Chlorella (Brennan and Owende, 2010), to which most oleaginous microalgae species belong. Centrifugation is widely used to recover microalgae biomass, especially small-sized algae cells; however, its application is restricted to algae cultures for high-value metabolites due to intensive energy needs and high equipment maintenance requirements. While flocculation is used to harvest small-sized microalgae cells, it is a preparatory step to aggregate the microalgae cells and increase the particle size so that other harvesting methods such as filtration, centrifugation, or gravity sedimentation can be applied (Molina Grima et al., 2003). Several flocculants have been developed to facilitate the aggregation of microalgae cells, including multivalent metal salts like ferric chloride (FeCl3), aluminum sulphate (Al2(SO4)3) and ferric sulphate (Fe2(SO4)3), and organic polymers such as chitosan and modified starch (Li et al., 2008). Chemical flocculation can be reliably used to remove small algae cells from pond water by forming large (1–5 mm) sized flocs (Sharma et al., 2006). However, besides the high cost of chemical flocculants and possible pollution effects that may generate, the chemical reactions are highly sensitive to pH and the high doses of flocculants required produce large amounts of sludge and may leave a residue in the treated effluent. In summary, most technologies including chemical and mechanical methods greatly increase operational costs for algal production and are only economically feasible for production of high-value products (Park et al., 2011). Besides traditional methods mentioned above, there are several new technology developments in this field. DOE-ARPA-E recently funded a research project for Algae Venture Systems (AVS) to develop a Harvesting, Dewatering and Drying (AVS-HDD) technology using the principles of liquid adhesion and capillary action to extract water from dilute microalgae solutions. Attached algal culture systems have been developed for growing microalgae on the surface of polystyrene foam to simplify the cell harvest (Johnson and Wen, 2010; Wilkie and Mulbry, 2002). New bioflocculants, which are more environmentally friendly, are also proposed to address the cost and environmental concerns for current flocculation methods (Uduman et al., 2010a). These methods are innovative and will decrease the harvest cost to some extent if developed successfully, but require heavy investments on equipment and chemical supplies. Enhancing natural algae aggregation to encourage simple gravity settling or filtration appears to be the most 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 use (Uduman et al., 2010a). It will also 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 (50–200 lm), and their cell aggregation can be achieved through nitrogen limitation and CO2 addition (Park et al., 2011); however, most of these microalgae species are not oleaginous species. Methods to enable oleaginous microalgae aggregate during their cultivation are strategically and urgently needed to develop efficient and economic means of biofuel production. In submerged cultures, filamentous microorganisms, including some species of molds and bacteria, tend to aggregate and grow as pellets/granules. These fungal cell pellets are 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, 2008; Hu et al., 2009). Fungal cell pelletization can significantly decrease the viscosity of the fermentation broth; therefore it has been widely researched to increase the cultural performance on the mixing and mass transfer properties. Other advantages of cell pelletization to the micro-oil production process include the ease of
harvesting cells and of re-using pond water (Johnson and Wen, 2010; Xia et al., 2011). Conditions for cell pelletization seem to be strain-specific and that not all the filamentous fungal strains can form pellets during their growth. A recent study at University of Minnesota showed that Mucor circillenous is relatively difficult to form pellets during their cell growth; however, with induction of CaCO3 powder in the early stage of its cultivation, cell pellets can be formed homogeneously, lasting for the entire cultivation cycle. Changing operational conditions during cell cultivation was found to be able to induce fungal cells to aggregate and form pellets (Xia et al., 2011). This method avoids traditional approaches that use CaCO3 powder or other nuclei to induce the fungal pelletization (Liao et al., 2007; Liu et al., 2008a) which are costly and cause solid waste disposal issues. Pelletization is more widely seen in the fungal fermentation process where the microorganisms are filamentous. However, most oleaginous microalgae are not filamentous and oleaginous microalgae pelletization has not been seen in a complete review of current literature. In this study, a preliminary study was conducted to inoculate filamentous fungal spores when culturing mixotrophic green algae C. vulguris and it was found that pellets clearly formed within two days of culture. The microalgae cells, aggregated together with fungal cells, were immobilized in the pellets. This paper described this new technology which uniquely addresses the cell harvest of microalgae and has the potential to greatly reduce the algae biofuel cost. While pelletization and granulation have already been developed for commercial production of numerous products to increase culture conditions (e.g., mass transfer and oxygen transfer), this paper is the first study to investigate co-culturing microalgae with filamentous fungi for possible direct pelletization of microalgae to lower the overall cost of the process. 2. Materials 2.1. Microbial species Filamentous fungi Aspergillus niger Ted S-OSU was tested in this study for its co-cultivation with microalgae, as well as other filamentous fungal strains, which were all purchased from ATCC. C. vulguris was the model microalgae species to test its co-pelletization with filamentous fungi. Previous research has revealed that C. vulguris 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). 2.2. Cell cultivation 2.2.1. Cultivation medium Autotrophic medium A (per L): KNO3 1 g, KH2PO4 0.075 g, K2HPO4 0.1 g, MgSO42H2O 0.5 g, Ca(NO3)24H2O 0.0625 g, FeSO47H2O 0.01 g, Yeast extract 0.5 g, A5 1 ml. A5 (L): H3BO3 2.86 g, Na2Mo42H2O 0.039 g, ZnSO47H2O 0.222 g, MnCl24H2O 1.81 g, CuSO45H2O 0.074 g, CoCl2 0.03 g. Heterotrophic medium B (per L): Potato dextrose broth 12 g, Glucose 15 g. 2.2.2. Cultivation of seed C. vulgaris An autotrophic flask culture of C. vulgaris was maintained in the lab to provide algae seeds for the co-cultural experiments. A 4 L flask was filled with 3 L culture medium A with a magnetic stir (100 rpm) and four fluorescent lights were provided outside the flask to culture the algae cells at 25–27 °C. Fresh cultural medium was routinely added into the flask to maintain the active cell growth.
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2.2.3. Preparation for seed fungal spores Fungal spore solution was stored at 70 °C with 25% of glycerol. The stored spores were activated by cultivation on the petri dish with medium B and 2% agar at 27 °C for 7 days. Sterilized DI (10 ml) water was applied on the culture to harvest the spores and the spore solution was used as the inoculation for the co-culture after the number of spores in the solution was counted. 2.2.4. Flask co-culture of filamentous fungi with autotrophic microalgae Prepare 100 ml culture medium A in 250 mL flasks, and then adjust pH by adding 2 mol/L HCl or 1 mol/L NaOH to pH 4.0. Transfer 20 ml microalgae seed broth into these 250 ml flasks. Finally, inoculate fungal spores into medium A to reach certain spore concentration (7.6 10E6/L). These flasks were cultured at 27 °C, 150 rpm for 3 days. 2.2.5. Cellulase degradation of C. vulgaris cell wall Inoculate with 7.6 10E6/L spores of A. niger in a flask with 10 times dilution of medium B and culture at 27 °C, 100 rpm for 3 days to obtain the fungal cultivation broth. The broth was filtered with sterilized filters to remove the fungal cells. C. vulgaris cell broth (1.76 10E10/L, 10 ml) was harvested by centrifuge, then inoculated into 4.5 ml of fungal supernatant solution together with 0.5 ml 500 mmol/L phosphate buffer and 100 ll cellulase (C8546, Sigma, St. L MO. USA), and reacted at 50 °C, for 1 h in water bath (Model 25, Precision scientific Inc. Chicago, IL, USA). 2.2.6. Flask co-culture of filamentous fungi with heterotrophic microalgae Prepare 100 ml culture medium B in 250 mL flasks, and then adjust pH to 5.0. Transfer 20 ml microalgae seed broth and inoculate fungal spores (7.6 10E6/L) into these 250 ml flasks. These flasks were cultured at 27 °C, 150 rpm for 3 days. 2.2.7. Co-culture of C. vulgaris and A. niger in a bioreactor Prepare 1 L of medium B together with 1 L water in the Bioreactor (Bioflo 110, New Brunswick, MA USA), and pH was adjusted to 5.0 after sterilization. Inoculate with C. vulgaris cultivation broth and A. niger spores (7.6 10E6/L) into the bioreactor. The culture condition was 27 °C, 200 rpm. The pH value was maintained by alkaline (1 mmol/L NaOH) addition automatically and the air feeding rate is 1 vvm. 2.3. Analysis At the end of each flask culture, pour all the pellets of the cocultivation of microalgae and filamentous fungi from each flask into the Petri dish (discard extra liquid if necessary) 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. The microalgae cell numbers in the supernatant were measured after diluting the supernatant multiple times until the cell numbers can be counted under microscope. Total microalgae algae cells were measured after the cultivation broth was blended completely to break the structure of pellets in order to release microalgae cells from pellets and then diluting the cell concentrations until the cell numbers can be counted under microscope. 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. Glucose concentration was estimated by using DNS method (Xiong et al., 2008). The total biomass concentration of the flask cultures were determined by the dry cell biomass. Dry biomass was then processed with 10 ml mixture of chloroform and metha-
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nol (2:1 v/v) to extract the microbial oil. Finally fatty acid methyl ester (FAME) was generated via transesterification reaction at 90 °C water bath (Model 25, Precision Scientific, Chicago, IL, USA) for 90 min with concentrated sulfuric acid as the catalyst. FAME in chloroform was carefully collected and subjected to GC after filtration (0.22 lm). The GC was equipped with a flame ionization detector and a HP-5 capillary column. The oven temperature was 80 °C, held for 5 min, raised to 290 °C at a rate of 4 °C/min, and held at 290 °C for 5 min, 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/min. 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, MO). All values were means of triplicate determinations. 3. Results and discussion 3.1. Description of pellets formed in the co-cultivation of C. vulgaris and A. niger Microalgae cells were suspended in the cultivation broth and the solution can be relatively stable and homogeneous. With the inoculation of fungal spores and addition of organic nutrients, the algae solution lost most of its green color within two days of cultivation, and turned transparent, indicating the majority of microalgae cells were pelletized. Pellets were formed in the cultivation broth with almost similar density as the cultivation solution. The pellets are spherical, packed, relatively homogeneous in size, and will settle down once the shaking stops. Significantly different from the color of pellets formed by fungi itself; the pellets in the co-cultivation are green, instead of white or grey. The microscopic image showed that the skeleton structure of the pellets is still filamentous fungal cells; while green algae cells are entrapped inside the fungal mycelia and attached on the fungal cell surface (Supplementary Figure). Since these microalgae cells are not filamentous, natural algae aggregation, simple gravity settling or filtration cannot be applied to these microalgae species for harvesting their cell biomass. Having a separate harvesting step, no matter by using flocculation, centrifugation or other methods, is costly and environmental non-friendly (Olaizola, 2003). Co-culturing filamentous fungi with microalgae cells enabled these microalgae cells to pelletize, and these cell pellets (size at 2–5 mm) can be harvested by a simple filtration with sieves, significantly easier than harvesting individual cells. The process is a totally different method compared to chemical flocculation. Instead of adding a chemical flocculent, which may be expensive, needs large quantity and/or leaves toxic residue to the harvested microalgae pat, fungal spores or hyphae were inoculated into the microalgae cultivation broth and were cultured together with microalgae cells for the harvest purpose. The pellets formed in this co-culture are more packed than ‘‘fluffy’’ floc, which gives more advantages to pellets for cell harvest (Olaizola, 2003). 3.2. Test of co-cultivation of C. vulguris with different filamentous fungal strains Aspergillus oryzae, a commercial fungal strain for cellulase production, can easily form relatively large cell pellets, which are stable and homogenous, during the cell growth without any inducing approaches (Table 1). Same phenomena also showed on many other fungal strains such as ATCC 11730, RLG 9902 and Aspergillus flavus. Mortierella isabellina, on the other hand, is nearly incapable of forming stable pellets (Table 1). During a cell culture in flasks,
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Table 1 Photos of co-pellets of fungal species and Chlorella vulgaris. Fungal species
M. isabellina
M. circinelloides
A. flavus
Fungal Pellets
Co-pellets
RLG 9902
Ted S-OSU
ATCC 9642
No photo
No pellets
pellets were formed in the first day of culture of M. isabellina; while all the pellets disappeared after 4 days of cultivation. M. circillenous is also relatively difficult to form pellets during their cell growth; however, with induction of CaCO3 powder in the early stage of its cultivation, cell pellets can be formed homogeneously, lasting for the entire cultivation cycle. Key operational conditions were revealed in the previous research to induce the pelletization of filamentous fungi M. circillenous (Xia et al., 2011). It was discovered that by only changing conditions during cell cultivation can force fungal cells to aggregate and form pellets. This method avoids traditional approaches that use CaCO3 powder or other nuclei to induce the fungal pelletization, which are costly and cause solid waste disposal issues (Liao et al., 2007; Liu et al., 2008a). Conditions for cell pelletization seem to be strain-specific and that not all the filamentous fungal strains can form pellets during their growth. Different species of fungal spores were tested for their co-pelletization capability with the green algae C. vulguris, as shown in Table 1 and Table 2. Several fungal species, such as ATCC 11730, RLG 9902 and A. flavus, can form pellets that can entrap almost all of the individual microalgae cells, resulting in a clear co-culture broth of those strains (Table 2). As for Mucor circinelloides (Table 2), there are still some C. vulgaris cells in supernatant, although green pellets also formed in their cultures. For Phanerochaete chrysosporium and A. oryzae, even though pellets were still formed in the coculture with microalgae, these pellets were either grey or milky while with almost no algae cells entrapped inside the pellets. There were no harvest effects on these cultures because microalgae cells were predominantly in the supernatant. Strains such as M. isabellina could form unstable pellets on its own; but it could not form any pellets when co-cultured with C. vulguris. The formation of fungal pellets and co-pellets were depending on the strains as well
as the cultivation conditions. Numerous researches have showed some specific methods on inducing the fungal pelletization (Liao et al., 2007; Liu et al., 2008a; Xia et al., 2011), however, there is no generally applicable strategy, especially for the co-pelletization of fungal and algae strains. Detailed study on the co-cultivation conditions is needed in each strain combination in order to induce the pellets formation of these cells. Depending on the application of microalgae cells, different filamentous fungal species can be co-cultured to pelletize microalgae cells for easier harvest. For instance, if the microalgae cells can generate cellulose on their cell wall, co-cultivation of a cellulaseproducing fungus with this microalgae will be beneficial because fungal cells can support themselves by partially degrading the cell wall of microalgae to generate sugars as well as pelletize microalgae cells into pellets for easer harvest. In this case, no external sugars are needed for this process. If the primary purpose of culturing microalgae is to generate microbial lipids, co-culturing an oleaginous fungus can be similarly beneficial. This is 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 high content of lipids that can be extracted for biodiesel production, co-culturing these oleaginous fungi cells will not only help with the oil-producing purpose, but will also facilitate the pelletization of microalgae cells for easier harvest. 3.3. Test of co-cultivation of A. niger with C. vulguris on different growth mode C. vulguris can grow on both autotrophic and heterotrophic growth modes and spores of A. niger were inoculated to both
Table 2 Co-pelletization of fungal species and C. vulgaris after 24 h heterotrophic condition. Fungal species
Pellets
Initial cell number (10 E9/L)
Mortierella isabellina Aspergillus flavus Aspergillus niger Aspergillus versicolor (ATCC 11730) Corynebacterium nephridii (Bacteria, ATCC 11425) Pycnoporus sanguineus (ATCC 24598) Aerococcus viridans (ATCC 9642) Phanerochaete chrysosporium (ATCC 24725) Trametes lilacinogilva (ATCC 46156) Pycnoporus sanguineus (ATCC 18395) Leucogyrophana arizonica (RLG 9902) Mucor circinelloides Aspergillus oryzae Control (No fungi)
p p p
1.80 2.65 3.30 1.85 2.60 2.40 3.95 2.40 2.75 2.35 2.80 2.75 1.75 2.45
p p⁄ p p p⁄
Algae in supernatant (10 E9/L) in 24 h 0 0.65 0 (48 h)
0.25 White pellets
0 6.95 White pellets 20.80
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70 60
Harvest ratio (%)
growth conditions to study the co-pelletization of these strains as shown in Table 3. The initial microalgae cell concentration both started at 6.9 10E9/L with medium A for the autotrophic cultivations (no sugar addition) and medium B for the heterotrophic cultivations. The microalgae final cell concentration reached at (10.5 ± 3.16) 10E9/L for the control tests with no fungal spores inoculated. For the co-culture at autotrophic conditions, the suspended algae concentration in the supernatant was about same as the control ((10.7 ± 0.78) 10E9/L), while a large amount of the microalgae cells were co-pelletized with the fungi, which enlarged the total microalgae cells in the co-culture almost three times more the control. The maximum microalgae cell concentration in autotrophic cultivations is usually limited by the mutual shading effects, where microalgae cells in the pond water cannot access to the sun light if the final algae cells reach to the maximum concentration (Lee and Palsson, 1994). The experiments showed that the co-cultivation of fungal cells actually stimulated the overall microalgae cultivation and dramatically increased the total algae production. Pelletization creates an inhomogeneous cultural system that large amount of algae cells were immobilized inside the pellets while the suspended algae cell density in the culture broth could remain at low level, which allows the penetration of light. For the co-culture with autotrophic microalgae, around 60% of algae cells were pelletized with fungal cells (Table 3 and Fig. 1). No sugar was added to the autotrophic co-cultivation to support the germination and growth of A. niger. Due to the low concentration of fungal cells, there was not any measurable cellulase activity in the co-cultivation broth of autotrophic microalgae and fungi (Data not shown). However, it was confirmed by many researchers that A. niger can produce cellulase during their cell growth (Acharya et al., 2008) and the cellulase is excreted in the fermentation broth to degrade the cellulose to support the cell growth. Test of degradation of C. vulgaris cell wall by external cellulase (Table 4) showed that sugar concentration increased from 1.74 to 2.36 g/L. It was also confirmed by other researchers 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) and also to facilitate the lipid extraction (Fu et al., 2010). The carbohydrates composition of microalgae is mainly polysaccharide, entrapped in the cell walls with up to 70% dry weight. The composition varies among microalgae strains, and cellulose is reported as the main structural component of the cell wall for most microalgae species. 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 the oil extraction (Fu et al., 2010; Yin et al., 2010). This experiment may provide some supports that fungal cells grew on the microalgae cell degradation, but further research is needed in the future to provide direct evidence. For the heterotrophic co-culture, the supernatant microalgae cell concentration reached to (72.50 ± 21.21) 10E9/L and the total microalgae cells reached to (95.75 ± 5.30) 10E9/L. Since algae
50 40 30 20 10 0 Autotrophic condition
Heterotrophic condition
Fig. 1. Harvest ratio of C. vulgaris at autotrophic and heterotrophic growth condition.
Table 4 Test of cellulase degradation of C.vulgaris cell wall. Cellulase addition
Initial
Fungal supernatant
Microalgae solution
Sugar concentration (g/L)
2.5
1.74 ± 0.03
2.36 ± 0.07
growth in heterotrophic conditions relied on the available external sugar, instead of light, microalgae cell concentration increased dramatically, with relatively smaller portion being pelletized (Fig. 1). Total cell biomass in the heterotrophic co-cultivation was significantly higher than other conditions because sugars were provided and consumed both by the fungus and microalga. A. niger, which only generated 17.69% of its biomass as oil (measured based on Fatty Acid Methyl Ester FAME), is not an oleaginous strain (Table 3). This explained that the oil content (FAME) of total biomass from co-cultivations was lower than the purely cultured C. vulguris. Although the fungus’s fatty acid profile was significantly different from C. vulguris (with relatively higher content of C18:2, less content of C16:1), the fatty acid profile of total biomass from co-cultivations were not much different than pure culture of C. vulguris (Fig. 2). The predominant fatty acid components of C. vulguris include C16:0, C16:1, C18:0, C18:1 and C18:2. Since A. niger cells did not contribute significantly to the total oil extracted from cocultivation biomass, the fatty acid composition was primarily determined by the algae. When co-culturing an oleaginous filamentous fungus, the fungal cells will contribute to the total oil production and the fatty acid profile of total biomass may change more significantly.
Table 3 Co-culture of A. niger with C. vulguris at different growth. C. vulgaris Volume (ml) Initial microalgae (10E9/L) Initial spores (10E6/L) Microalgae in supernatant (10E9/L) Total microalgae cell (10E9/L) Biomass (g/L) Fatty acid (%) Fatty acid (mg/L)
100 6.9 10.5 ± 3.16 10.5 ± 3.16 0.142 ± 0.01 37.40 ± 1.50 53.11 ± 1.51
A. niger
Autotrophic
Heterotrophic
100 0 7.6 0 0 0.215 ± 0.02 17.69 ± 3.08 38.03 ± 10.05
100 6.9 7.6 10.7 ± 0.78 29 ± 3.50 0.304 ± 0.02 32.58 ± 3.26 99.04 ± 16.30
100 6.9 7.6 72.50 ± 21.21 95.75 ± 5.30 0.968 ± 0.129 28.50 ± 1.93 275.88 ± 52.40
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Chrolella vulgaris Aspergillus niger Autotrophic condition Heterotrophic condition
40
Percentage (%)
30
20
10
0 C14
C15 C16:1 C16 C18:3 C18:2 C18:2 C18:1 C18
C20
C22
Fatty acid Fig. 2. Fatty acid profiles of co-cultivated C. vulguris and A. niger.
Experiments in the bioreactor containing C. vulgaris and A. niger showed typical filamentous growth, which pure culture of microalgae C. vulgaris should not have, although some cells form pellet like aggregates while other cells attached on the mixer or other surface inside the bioreactor. With the co-culture of fungal cells, the supernatant microalgae cells dropped to 0.2E9/L from 7.8E9/L at initial inoculations while the total microalgae reached to 10.4 10E9/L, which means that 98.1% microalgae cells were entrapped in the fungal clumps (Fig. 3). Factors to affect the pelletization process and how to scale up the co-cultivation should be investigated in details in the future.
3.4. Mechanism of co-pelletization of fungal and microalgae cells
12
100
10
DO (%)
80
8 60 6 40 4 DO Glucose Microalgae
20
2
0
0 0
10
20
30
40
50
Time (h) Fig. 3. Co-culture of C. vulgaris and A. niger in the bioreactor.
Microalgae supernatant (10E9/L) Glucose (g/L)
Fungal cell pelletization has been studied for decades and widely applied in the industrial processes to produce organic acid, pharmaceuticals, enzymes and other high-valued fermentation products, and wastewater treatment to remove pollutants. The research results showed conditions for cell pelletization seem to be strain-specific and that not all the filamentous fungal strains can form pellets during their growth. The capability of cell aggregation of filamentous fungal cells is predominately rooted to the production of hydrophobic proteins (referred to as hydrophobins), a family of low molecular weight amphipathic proteins. For those dimorphic fungal species, hydrophobin were detected on the hyphal surface while missing when the cells were grown in the yeast
form (Linder, 2009). One of the biological functions of these hydrophobic proteins is to coordinate the adherence of hyphae to solid substrates, which facilitates biofilm formation (attaching on a solid surface) and cell pelletization/granulation/aggregation (attaching on each other) (Feofilova, 2010). Numerous theories have been proposed to explain the fungal pelletization process, which can be categorized into two types of mechanisms (Liu et al., 2008b; Ryoo and Choi, 1999). The first one is coagulative mechanism, where spores coagulate in the early stage of cultivation and develop into pellets through their intertwining hyphae. The second one is non-coagulative mechanism, where the spores germinate into hyphae, and then intertwine into pellets. Many genus of fungus such as Aspergillus, basidiomycete, Phanerochaete chrysosporium follows the coagulative mechanism where fungal spores conglomerate at an early stage of development and then each pellet may arise from each spore aggregate. While other genus of fungus, such as Rhizopus spp, Mucor spp. Penicillium chrysogenum, follows the non-coagulative mechanism. The experiment results showed that co-pelletization of fungal and microalgae cells only occurred on certain filamentous fungal strains (Table 2) and the detailed mechanisms are still not clear. The cell interaction information could have significant potential impacts in inducing the co-pelletization so that co-pelletization conditions can be refined for the large-scale production of microalgae-fungal pellets as a biofuel feedstock. One microscopic photo taken on the co-pellets of A. niger hyphae and C. vulgaris grinded by a blender clearly showed that the microalgae cells were attracted on the filamentous hyphae (Supplementary Figure). The mutual attraction of microalgae and filamentous cells may be attributed to several possible reasons, but the primary reason might be due to the surface charge. Most microalgae cells carry negative charges on their cellular surfaces and therefore are capable of forming stable suspensions in the cultivation pond water. The stability of these microalgal suspensions depends on the forces that interact between the cells themselves and between the cells and water. Hence they are considered as hydrophilic bio-colloids (Uduman et al., 2010b). Current flocculation method to harvest microalgae is to add flocculants such as metals ions or organic polymers that have counterions to charge neutralisation and polymer bridging, and then allow cells to aggregate to form floc (Pieterse and Cloot, 1997). Fungal hyphae and mycelia contain polysaccharide with active sites that may enable their surface capability of bioadsorption and also enable the fungal cells to be charged (Tan et al., 2004). In a similar published research study to search for a biological flocculation agent for microalgae cell harvest (Rajab, 2007), A. flavus were grown to form cell pellets, and then applied to the microalgae cultural broth to flocculate the microalgae cells. The fungal cell pellets did absorb the microalgae cells, and their zeta potential measurement revealed the average number for microalgae as 23.7 millivolt while A. flavus as +46.1 mV. The charge difference between the microalgae and fugal cells indicated the capability of A. flavus to capture microalgae, and it provides some clue that surface charge might be the reason of co-pelletization of microalgae and fungal cell cultures.
4. Conclusions This paper studied the possibility to co-culture non-filamentous microalgae cells with filamentous fungi to form cell pellets for easier harvest. The co-culture of filamentous fungi to enable filamentous features on non-filamentous microalgae cultures will potentially provide an innovative process with direct commercial application potential on microalgae production of biofuel and other products. Compared to other algae harvesting methods, this method will potentially decrease harvest costs, avoid second pollu-
J. Zhang, B. Hu / Bioresource Technology 114 (2012) 529–535
tion from chemical flocculants and stimulate the algae production, although further research is needed to study the detailed pelletization conditions in the larger scale industrial applications. Acknowledgements Dr. Jianguo Zhang’s research was supported by Dr. Bo Hu’s faculty seed money program at University of Minnesota. We appreciate Dr. Jonathan Schilling at University of Minnesota for providing some fungal strains. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.biortech.2012. 03.054. References Acharya, P.B., Acharya, D.K., Modi, H.A., 2008. Optimization for cellulase production by Aspergillus niger using saw dust as substrate. Afr. J. Biotechnol. 7, 4147–4152. Brennan, L., Owende, P., 2010. Biofuels from microalgae – A review of technologies for production, processing, and extractions of biofuels and co-products. Renew. Sustain. Energy Rev. 14, 557–577. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294–306. Fakas, S., Papanikolaou, S., Batsos, A., Galiotou-Panayotou, M., Mallouchos, A., Aggelis, G., 2009. Evaluating renewable carbon sources as substrates for single cell oil production by Cunninghamella echinulata and Mortierella isabellina. Biomass Bioenergy 33, 573–580. Feofilova, E.P., 2010. The fungal cell wall: modern concepts of its composition and biological function. Microbiology 79, 711–720. Fu, C.-C., Hung, T.-C., Chen, J.-Y., Su, C.-H., Wu, W.-T., 2010. Hydrolysis of microalgae cell walls for production of reducing sugar and lipid extraction. Bioresour. Technol. 101, 8750–8754. Harun, R., Danquah, M.K., 2011. Enzymatic hydrolysis of microalgal biomass for bioethanol production. Chem. Eng. J. 168, 1079–1084. Heredia-Arroyo, T., Wei, W., Ruan, R., Hu, B., 2011. Mixotrophic cultivation of Chlorella vulgaris and its potential application for the oil accumulation from non-sugar materials. Biomass Bioenergy 35, 2245–2253. Hu, B., Chen, S.L., 2007. Pretreatment of methanogenic granules for immobilized hydrogen fermentation. Int. J. Hydrogen Energy 32, 3266–3273. Hu, B., Chen, S.L., 2008. Biological hydrogen production using chloroform-treated methanogenic granules. Appl. Biochem. Biotechnol. 148, 83–95. Hu, B., Zhou, X., Forney, L., Chen, S.L., 2009. Changes in Microbial Community Composition Following Treatment of Methanogenic Granules with Chloroform. Environ. Prog. Sustainable Energy 28, 60–71. Johnson, M.B., Wen, Z.Y., 2010. Development of an attached microalgal growth system for biofuel production. Appl. Microbiol. Biotechnol. 85, 525–534.
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