- Email: [email protected]
Immobilization of microalgae on exogenous fungal mycelium: A promising separation method to harvest both marine and freshwater microalgae
Biochemical Engineering Journal 91 (2014) 53–57
Contents lists available at ScienceDirect
Biochemical Engineering Journal journal homepage: www.elsevier.com/locate/bej
Immobilization of microalgae on exogenous fungal mycelium: A promising separation method to harvest both marine and freshwater microalgae Md. Mahabubur Rahman Talukder ∗ , Probir Das, Jin Chuan Wu Institute of Chemical and Engineering Sciences, Singapore
a r t i c l e
i n f o
Article history: Received 8 March 2014 Received in revised form 20 June 2014 Accepted 1 July 2014 Available online 10 July 2014 Keywords: Microalgae Immobilization Separation Optimization Screening fungus Mycelium
a b s t r a c t Although various methods have been developed for microalgae harvesting, problems such as chemical toxicity, low efficacy, low flexibility, and high cost, still remain with the current methods. In the present study, an effective biological separation method has been developed to harvest microalgae via immobilization on exogenous fungal mycelium. Fungal mycelium was added into microalgae culture and air mixed for the immobilization. The immobilized microalgae with fungal mycelium were naturally precipitated within 10 min after stopping the mixing. Both marine (e.g. Nannochloropsis sp.) and freshwater (e.g. Chlorella vulguris) microalgae were almost completely (94–97%) precipitated using the mycelium of Aspergillus nomius CCK-PDA 7#6. The precipitated mixed biomass was separated by simple filtration using sieve. A 70% initial nutrients supplement to de-algated culture medium gave high growth yield (0.43 g/l) in subsequent microalgae cultivation. The mycelium can be obtained free or at low-cost as waste from a fungal fermentation process of producing valuable products. The developed method is, therefore, very promising for economical harvesting of microalgae. © 2014 Published by Elsevier B.V.
1. Introduction Microalgae biomass is a promising renewable source for fuels and chemicals production [1,2]. However, a lack of an economical and effective method for harvesting microalgae is one of the bottlenecks for commercialization of microalgae-based fuels and chemicals [3]. Harvesting via natural sedimentation or filtration has been attempted. However, sedimentation and simple filtration are not feasible except for large species such as Arthrospira due to the small size of microalgal cell (2–20 m) and their colloidal stability in suspension. Centrifugation is used for harvesting microalgae for high value products but it is expensive if microalgae biomass is used for low value products such as biofuel. Chemical precipitation or flocculation using inorganic coagulants such as alum and ferric chloride is widely used and has been reported to be most efficient for large-scale harvesting of microalgae [4]. However, the chemical flocculation contaminates the
∗ Corresponding author. Present address: Department of Industrial Biotechnology, Institute of Chemical and Engineering Sciences, 1 Pesek Road, Jurong Island, Singapore 627833, Singapore. Tel.: +65 67963826; fax: +65 63166182. E-mail addresses: [email protected], [email protected] (Md.M.R. Talukder). http://dx.doi.org/10.1016/j.bej.2014.07.001 1369-703X/© 2014 Published by Elsevier B.V.
harvested biomass and culture medium with metal salts, causing difficulties in downstream processing of the biomass, reuse of the culture medium and waste water treatment [5]. We have recently found that the fermentation of lipid depleted algal biomass to lactic acid was significantly inhibited at Fe+ concentration >17 ppm [6]. Chemical flocculation using alum has negatively affected the ferementative production of acetone, butanol and ethanol [7]. These drawbacks can be minimized by biological harvesting via bioflocculation [8] or co-culture of other microorganisms such as fungus [9,10] and bacteria [11]. Bioflocculants such as chitosan, starch and poly ␥-glutamic acid, although effective for freshwater microalgae, undergo coiling at high ionic strengths and consequently are less effective for marine microalgae [12]. The filamentous fungi [9,10] that used to harvest microalgae via co-culturing can only grow in freshwater, and are not applicable for harvesting marine microalgae. Furthermore, the carbon source needed for growing such fungi will evoke undesirable microbial contamination of microalgae culture in an open raceway pond [11]. Microalgae harvesting via immobilization on different carriers has also been studied [13,14]. However, there are no prior studies on immobilization of microalgae on exogenous fungal mycelium. The mycelium can be obtained free or at low-cost as waste from a fungal fermentation process of producing valuable products such as enzymes and soy sauce [15,16]. Furthermore, the addition of fungal mycelium will increase
54
Md.M.R. Talukder et al. / Biochemical Engineering Journal 91 (2014) 53–57
biomass yield. As both fungus and microalgae biomasses contain lipid, carbohydrate and protein, and can be processed together, the harvesting microalgae by immobilizing them on fungal mycelium may add value to the microalgae biorefinery. In this study, a simple but effective separation method to harvest both marine and freshwater microalgae via immobilization on exogenous fungal mycelium was developed. The effects of various parameters such as fungal mycelium to microalgae biomass ratio, pH and immobilization time on separation of microalgae from the culture were investigated. 2. Material and methods 2.1. Material Nannochloropsis sp. was locally isolated [17] while Chlorella vulgaris (ATCC 13482) and Dunaliella teriolecta (UTEX LB999) were from ATCC and UTEX, respectively. Waste cooking oil containing 18.5% free fatty acids and 0.9% water was provided by Vender Horst Biodiesel Pte Ltd. (Singapore). Polypeptone were from Nihon Seiyaku, Japan. Potato Dextrose Agar (PDA) was from Difco, USA. HPLC grade methanol and hexane were from J.T. Baker, USA. Dimethylacetamide (DMAC) was from Sigma-Aldrich. All chemicals unless mentioned otherwise were of analytical grade and used as received. 2.2. Methods 2.2.1. Cultivation of microalgae A locally isolated Nannochloropsis sp. [17] was cultured in Guillard f/2 (w/o Si) medium, which was prepared using synthetic sea salt (35 g/l) and deionized water. An inoculum of 200 ml (0.3 g/l) was added into 5 l capacity glass bottle filled with 3.8 l Guillard f/2 medium in advance. The bottle was continuously irradiated with two white fluorescent lights (TL5, 28 W) placed 20 cm away from the bottle. The light intensity received by the culture was not measured. The culture was mixed with sterile-filtered air through sparger at a flow rate of 2 l/min and maintained at room temperature (22–23 ◦ C) for 6–7 days. During this time the growth of microalgae was relying on atmospheric CO2 and microalgae density reached 0.4–0.45 g/l. Microalgae density was determined based on the calibration curve prepared by plotting microalgae dry weight against the value of optical density at 680 nm (OD680 ). C. vulgaris and D. teriolecta were cultured in BAR medium [18] and modified ATCC 1174DA medium (reduced NaCl concentration of 0.5 M), respectively under the same conditions mentioned above. 2.2.2. Screening fungi Fungi were isolated from soil as described previously [19]. All fungal isolates were grown on PDA agar at 30 ◦ C for 5–7 days. Spores or mycelium of a fungus from agar plate were transferred into a conical flask containing 50 ml of polypeptone medium: waste cooking oil 30 g/l, polypeptone 10 g/l, KH2 PO4 10 g/l, NaNO3 2 g/l, MgSO4 ·5H2 O 0.5 g/l and pH 6.0. The flask was incubated at 150 rpm and 30 ◦ C for 4 days. The whole fungal culture was mixed with 200 ml microalgae culture (0.4 g/l) in a glass bottle of capacity 500 ml by air through a sparger at a flow rate of 0.5 l/min for 5 h. The air flow was stopped to precipitate the immobilized microalgae with fungal mycelium for 10 min by natural sedimentation. The six isolates, which could precipitate ca. 90% of microalgae from culture, were primarily chosen as the promising candidates. The primarily selected candidates were further screened based on the amount of microalgae immobilized per gram fungal mycelium. The isolate PDA 7#6, which was identified as Aspergillus nomius CCK-PDA 7#6 [19] and deposited in the institute of DSMZ-German
Culture Collection of Microorganism and cell cultures (DSM 26642), was finally selected. 2.2.3. Cultivation of fungus in a fermenter and preparation of mycelium A flask (250 ml) containing 100 ml polypeptone medium was inoculated by 1 ml spores suspension (106 –107 /ml) and incubated for 1 day at 150 rpm and 30 ◦ C. The culture was transferred into a 2 l fermenter (BIOSTAT, B plus, Sartorius, Germany) containing 1000 ml polypeptone medium. The cultivation conditions: pH 6.0, temperature 30 ◦ C, stirring speed 300 rpm and aeration (0.5 l/min) were kept constant. The mycelium after 4–7 days of cultivation was washed with deionized water and vacuum filtered through a filter paper (Whatman 125). The wet mycelium (filter cake) containing ca. 80% water was used for immobilising microalgae. The wet mycelium was freeze dried for 48 h to measure the water content. 2.2.4. Immobilization of microalgae on fungal mycelium The wet fungal mycelium was mixed with 1000 ml microalgae culture (density 0.4 g/l) at room temperature (22–23 ◦ C) by air through a sparger at a flow rate of 1 l/min. After a specified mixing or immobilization time, air flow was stopped to precipitate immobilized microalgae with fungal mycelium for 10 min. The immobilization was optimized by varying fungal mycelium load (0.2–1.6 g dry weight), immobilization time (0.5–10 h) and microalgae culture pH (5.5–8.5). The original pH of microalgae culture was adjusted to different pH (5.5–8.5) by supplying 100% CO2 (1 l/min) prior to the mixing with fungal mycelium. The precipitate (immobilized microalgae with fungal mycelium) was separated by simple filtration using sieve (mesh no. 18, nominal sieve opening 1 mm). The immobilization efficiency (I.E) was calculated using the formula I.E = (1 − ODb /ODa )×100, where ODb and ODa are OD680 of the culture before and after precipitating microalgae, respectively. All data are averaged of at least three replicate experiments and varied within ±3–8%. 2.2.5. Extraction of chitin from fungal mycelia Chitin in different fungal mycelia was extracted by using the method described in the literature [20]. Freeze dried mycelia were placed in 5% LiCl/DMAC solvent at a ratio of 0.5 g/75 ml with constant stirring at room temperature (ca. 23 ◦ C) for 48 h. The viscous suspension was centrifuged at 4000 rpm for 5 min, the supernatant containing dissolved chitin was collected, and distilled water (1:1) was added to the supernatant to precipitate the chitin over 24 h. The precipitate was recovered as chitin pellets by centrifugation. The chitin pellets were washed three times with distilled water to remove remaining DMAC, freeze-dried and weighed. 2.2.6. Measurement of microalgal zeta potential The zeta potential of the microalgal cells were measured at different pHs adjusted by adding acid (0.1 M HCl) or alkali (0.1 M NaOH) by using Nano Zetasizer 3600 (Malvern Instruments, Worchestershire, UK) using the electrophoretic light scattering technique. Microalgae culture (1.0 ml) was pipetted into a cuvette and inserted into the units for zeta potential measurements. Zeta potential measurements were performed in triplicate. 3. Results and discussion 3.1. Screening fungi for immobilising microalgae About 300 fungal isolates were screened to immobilize microalgae. After preliminary screening (described in Section 2.2.2), five best fungal isolates was chosen to determine the immobilization capacities of their mycelia at a fungal mycelium to microalgae biomass ratio of 1:2 (w/w). Fig. 1 shows that the immobilization
Md.M.R. Talukder et al. / Biochemical Engineering Journal 91 (2014) 53–57
16
14
14
12
12
10
10 8
8
6
6 4
4
2
2
0
0
-10 -12
80
-14 -16
60
Immobilizaon efficiency
-18
Zeta potenal
40
-20 -22
20
Zeta potential (mV)
16
Chitin content
Immobilisation efficiency (%)
Immobilisation efficiency (%)
18
100
18
Immobilization efficiency
Chitin in dry fungal biomass (%)
20
55
-24 0
5
6
7
8
9
-26
PDA 5#198 PDA 6#39 PDA 6#139 PDA 7#6 PDA 7#129
pH
Fig. 1. Microalgae (Nannochloropsis sp.) immobilization capacities of the mycelium of five best fungal isolates and their chitin content. Immobilization conditions: 1000 ml microalgae culture (0.4 g/l), fungal mycelium to microalgae biomass ratio 1:2 (w/w), pH 6, mixing time 2 h and air flow rate 1 l/min. The numbers 7 and 6 in the name of “PDA 7#6” indicate the pH of PDA agar and strain number, respectively. The same principle was applied to name the other isolates.
Fig. 3. Effect of pH on microalgae (Nannochloropsis sp.) immobilization and zeta potential of microalgae cells. The immobilization conditions were the same as those mentioned in Fig. 1 except fungal mycelium to microalgae biomass ratio (w/w) was 4:1.
efficiency (I.E.) varied with fungal isolates and the mycelium of PDA 7#6 identified as A. nomius CCK-PDA 7#6 (DSM 26642) was able to immobilize highest amount of microalgae. The immobilization could be attributed to the ionic attraction between fungal mycelium (positively charged) and microalgae cell (negatively charged). The fungal cell wall material chitin has strong positive charge aiding immobilization of microalgae. It can be seen that PDA 7#6 mycelium contained highest amount of chitin and microalgae I.E. correlated well with the chitin content in different fungal mycelia (Fig. 1). In addition to the ionic attraction, microalgae could be trapped inside the porous structure of fungal mycelium. As the net surface charge and structure of the mycelium differ with the fungal species, the amount of microalgae immobilized varied with the isolates. 3.2. Effect of carbon and nitrogen sources used for growing PDA 7#6 on microalgae immobilization
18
20
16
18
14
16 14
12
Growth Yield
10
Immobilisation efficiency
12 10
8
8
6
6
4
4
2
2
0
Waste cooking oil
Glucose
Xylose
Glycerol
Immobilisation efficiency (%)
PDA 7#6 growth yield (g/l)
The cultivation of PDA 7#6 and the preparation of mycelium were described in Section 2.2.3. Fig. 2 shows that the maximum growth yield (dry biomass per liter) reached 15.62 g/l using waste cooking oil as carbon source. The growth yield on waste cooking oil
0
Fig. 2. Effect of different carbon sources on PDA 7#6 growth yield and the immobilization capacity of its mycelium. PDA 7#6 was separately grown in a fermenter as described in Section 2.2.3. The immobilization conditions were the same as those mentioned in Fig. 1.
was 1.3 fold higher than that on glucose. This is not surprising as oil gives almost twice the usable chemical energy that glucose can deliver. The use of xylose and glycerol as a carbon source decreased PDA 7#6 growth yield. The similar result has been reported by Gulari et al., 1999 [21]. It is obvious that microalgae immobilization capacity of the mycelium was highest with glucose followed by waste cooking oil, xylose and glycerol (Fig. 2). To better understand, chitin content in the mycelium of PDA 7#6 grown on glucose and waste cooking oil were determined. The chitin content in PDA 7#6 mycelium increased from 14.8% to 18.5% (dry weight) when waste cooking was replaced by glucose. This result suggests that glucose induced the production of polysaccharide chitin in fungal cell wall, enhancing the adsorption of microalgae. Hence, the combination of waste cooking oil and glucose could be a better carbon source than either one alone to immobilize microalgae more efficiently. However, waste cooking oil yielded higher biomass and is inexpensive than glucose. Waste cooking oil was, therefore, chosen for subsequent studies. Effects of four different nitrogen sources including polypeptone, yeast extract, (NH4 )2 SO4 , and NaNO3 on PDA 7#6 growth yield and microalgae immobilization were also investigated. The biomass yield was highest with polypeptone (15.61 g/l) followed by yeast extract (14.21 g/l), NaNO3 (6.13 g/l) and (NH4 )2 SO4 (8.33 g/l) while microalgae I.E. remained almost the same (ca. 16–17%) regardless of the type of nitrogen source used. 3.3. Optimization of the immobilization of marine microalgae (Nannochloropsis sp.) Microalgae cells contain amino and carboxylic groups as well as polysaccharides and acidic compounds such as uronic acid. The contributions of each of these groups to the overall surface charge of the microalgae depend on the culture pH. The original pH (ca. 8.5) of microalgae culture was, therefore, adjusted to 5.5–8.5 by supplying 100% CO2 prior to the addition of fungal mycelium and the effect of pH on I.E. was investigated at a fungal mycelium to microalgae biomass ratio of 4:1 (w/w). Fig. 3 shows that lowering the culture pH favored the immobilization of microalgae and the maximum I.E. was ca. 85% at a pH of 6.0–6.5 after 2 h mixing. It is also obvious that immobilization efficiency correlated well with the zeta potential, which was found to lie in the range of −19.8 to −24.1 mV for Nannochloropsis sp. used in this studies for the pH range of 5.5–8.5. The pH affects zeta potential, which represents the surface charges of microalgae cells. The higher negative zeta potential (ca. −24 mV) at pH 6.0–6.5 thus suggests that the ionic attraction
Md.M.R. Talukder et al. / Biochemical Engineering Journal 91 (2014) 53–57
0.6 Microalgae dry cell biomass (g/l)
Immobilisation Efficiency (%)
100
80
60 4:1
40
2:1 1:1 20
1:2
0.5 0.4 0.3 0.2 0.1 0
2
4 6 Mixing time (h)
8
0
1
2
10
Fig. 4. Effect of mixing time on the immobilization of Nannochloropsis sp. at different ratios (w/w) of fungal mycelium to microalgae biomass. The ratio (w/w) was varied by increasing the mycelium load while the other immobilization conditions were the same as those mentioned in Fig. 1.
between microalgae (negatively charged) and fungal mycelium (positively charged) could reach to a maximum at a pH 6.0–6.5. Fig. 4 shows the effect of mixing time on the immobilization of microalgae at different ratios of fungal mycelium to microalgae biomass. The ratio was varied by adding different amounts of fungal mycelium into 1000 ml microalgae culture (0.4 g/l). It is evident from Fig. 4 that the immobilization rate increased with the increase in fungal mycelium load and the maximum immobilization efficiency (I.E.) reached 97.2% at a ratio of 4:1 after 3 h mixing. A longer mixing time was needed at a lower mycelium load. The I.E. was improved at a higher ratio because of the larger amount of mycelium available for immobilizing microalgae. At a lower ratio (e.g. 1:2), the mycelium was insufficient to yield higher I.E. To prove the robustness of the developed method, another seawater microalgae species (D. teriolecta, UTEX LB999) was immobilized at a fungal mycelium to microalgae biomass ratio of 4:1 under the same conditions used for Nannochloropsis sp. The maximum I.E. reached 92.7% after 9 h. This result suggests that the developed method could be used for other microalgae species. 3.4. Reuse of de-algated culture medium After separating the precipitate (the immobilize microalgae with fungal mycelium), 500 ml de-algated culture medium was supplemented with fresh nutrients, inoculated with 25 ml microalgae (Nannochloropsis sp.) seed culture (0.3 g/l), and air mixed. Microalgae growth in the de-algated culture medium with and without supplement (30% and 70%) and fresh f/2 medium were shown in Fig. 5. The microalgae growth yield in the de-algated medium without supplement was low (0.23 g/l after 7 days) due to the insufficient nutrients. The growth yield increased to 0.43 g/l after 7 days by supplementing 70% initial nutrients and was comparable with that in the fresh f/2 medium. This result suggests that the harvesting microalgae via immobilization on fungal mycelium allowed to reuse the de-algated culture medium and can contribute to economical microalgae cultivation. 3.5. Flexibility of the developed method to harvest freshwater microalgae The freshwater microalgae C. vulgaris (ATCC 13482) were also immobilized on PDA 7#6 fungal mycelium to examine the flexibility of the developed method. The experiment was performed using the microalgae culture of density 0.4 g/l at a fungal mycelium to microalgae biomass ratio of 4:1, pH 6.0 and air flow rate 1 l/min.
3
4
5
6
7
8
Cultivation time (day) Fig. 5. Nannochloropsis sp. growth in de-algated culture and fresh f/2 medium.
100
Immobilisation Efficiency (%)
0
80
60
40
20
0
0
1
2
3 4 Mixing time (day)
5
6
Fig. 6. Effectiveness of PDA 7#6 fungal mycelium to immobilize freshwater microalgae (C. vulgaris). Immobilization conditions were the same as those mentioned in Fig. 1 except fungal mycelium to microalgae biomass ratio was 4:1.
Fig. 6 shows that the maximum I.E. with C. vulgaris reached 89.7% after 4 h. To further improve I.E., the optimal pH for the immobilization was investigated. Fig. 7 shows that I.E. increased with the increase in pH and was maximum (94.5%) at pH 7. This result suggests that the developed method can also be applied to harvest freshwater microalgae. I.E. also correlated well with zeta potential, which represent the surface charges of the microalgae cells. The similar phenomenon was also observed in the case of 100 Immobilisation efficiency (%)
0
70% Supplement Fresh f/2 medium 30% Supplement De-algated culture medium
-22 -23
80
-24
60
Immobilisaon efficiency
-25
Zeta potenal
-26 -27
40
-28 -29
20
Zeta potential (mV)
56
-30 -31
0
-32 5
6
7 pH
8
9
Fig. 7. Immobilization of C. vulgaris at different pHs. Immobilization conditions were the same as those mentioned in Fig. 6 except mixing time was kept constant at 4 h.
Md.M.R. Talukder et al. / Biochemical Engineering Journal 91 (2014) 53–57
Nannochloropsis sp. (Fig. 3). The experiment results thus suggest that the ionic attraction between microalgae cells and fungal mycelium due to the difference in surface charge was the main reason for the immobilization of microalgae on fungal mycelium. Harvesting C. vulguris via co-cultivation with PDA 7#6 under heterotrophic condition was also investigated by following the protocol reported by Zhang and Hu [9] and the result was compared with that obtained with the method developed in this study. Small portion of the microalgae was immobilized on fungal mycelium via co-cultivation, and the supernatant remained green color even after 6 days of co-cultivation. The maximum harvesting efficiency reached 30.04%. This harvesting efficiency was slightly higher than that (ca. 25%) reported by Zhang et al. [9]. This result suggests that the method developed in the study is more promising as it is almost completely (94.3%) immobilized and precipitates the same microalgae species from the culture medium. It should be noted that PDA 7#6 does not grow in autotrophic condition. Hence, harvesting C. vulguris via co-cultivation with PDA 7#6 under autotrophic condition was not possible. 4. Conclusions Several fungi were screened to immobilize microalgae and a separation method using the mycelium of A. nomius CCK-PDA 7# 6 was developed to harvest both marine and freshwater microalgae. The immobilization capacity of the mycelium was affected by the carbon sources used for growing PDA 7#6. Zeta potential measurement revealed that the charge difference between microalgae and fungal cells affects the capability of PDA 7#6 to immobilize microalgae. The maximum immobilization efficiency for Nannochloropsis sp. reached ca. 97% at a fungal mycelium to microalgae biomass ratio 4:1, pH 6.0 and mixing time 3 h. It has been demonstrated that the immobilization of microalgae on fungal mycelium can be regarded as a simple, effective and flexible separation method to harvest microalgae. The method also contributes in economical cultivation of microalgae by the reuse of de-algated culture medium. Acknowledgments Financial support from the Agency for Science Technology and Research (A*STAR) project number (ICES/10-273A01) of Singapore is gratefully acknowledged. Special thanks to Dr. Yvonne Chow for providing us with C. vulgaris (ATCC 13482) and D. teriolecta (UTEX LB999). Thanks are extended to Mr. Ng Jun Wei for helping us to measure microalgal zeta potential.
57
References [1] S.H. Ho, S.W. Huang, C.Y. Chen, T. Hasunuma, A. Kondo, J.S. Chang, Bioethanol production using carbohydrate-rich microalgae biomass as feedstock, Bioresour. Technol. 135 (2013) 1891–2198. [2] C.Y. Chen, X.Q. Zhao, H.W. Yen, S.H. Ho, C.L. Cheng, D.J. Lee, F.W. Bai, J.S. Chang, Microalgae-based carbohydrates for biofuel production, Biochem. Eng. J. 78 (2013) 1–10. [3] M.K. Lam, K.T. Lee, Immobilisation as a feasible method to simplify the separation of microalgae water for biodiesel production, Chem. Eng. J. 191 (2012) 263–268. [4] L. Kim, K. Kwon, Y. Oh, Effect of harvesting method and growth stage on the flocculation of the green algae Botryococcus braunii, Lett. Appl. Microbiol. 27 (1998) 14–18. [5] I. Rawat, R.R. Kumar, T. Mutanda, F. Bux, Biodiesel from microalgae: a critical evaluation from laboratory to large scale production, Appl. Energy 103 (2013) 444–467. [6] M.M.R. Talukder, D. Probir, J.C. Wu, Microalgae (Nannochlorpsis salina) biomass to lactic acid and lipid, Biochem. Eng. J. 68 (2012) 109–113. [7] R.J. Anthony, J.T. Ellis, A. Sathish, A. Rahman, C.D. Miller, R.C. Sims, Effect of coagulant/flocculants on bioproducts from microalgae, Bioresour. Technol. 149 (2013) 65–70. [8] D.G. Kim, H.J. La, C.Y. Ahn, Y.H. Park, H.M. Oh, Harvest of Scenedesmus sp. with bioflocculant and reuse of culture medium for subsequent high density culture, Bioresour. Technol. 102 (2011) 3163–3168. [9] J. Zhang, B. Hu, A novel method to harvest microalgae via co-culture of filamentous fungi to form cell pellets, Bioresour. Technol. 114 (2012) 529–535. [10] S. Xie, S. Sun, S. Dai, J.S. Yuan, Efficient coagulation of microalgae in cultures with filamentous fungus, Algal Res. 2 (2013) 28–33. [11] S. Salim, R. Bosma, M.H. Vermue, R.H. Wiiffels, Harvesting of microalgae via bio-flocculation, J. Appl. Phycol. 23 (2011) 849–855. [12] D. Bilanovic, G. Shelef, Flocculation of microalgae with cationic polymers—effects of medium salinity, Biomass 17 (1988) 65–76. [13] S. Nayar, B.P.L. Goh, L.M. Chou, Settlement of marine periphytic algae in a tropical estuary, Estuar. Coast. Shelf Sci. 64 (2005) 241–248. [14] N. Akhtar, J. Iqbal, M. Iqbal, Removal and recovery of nickel (II) from aqueous solution by loofa sponge immobilised biomass of Chlorella sorokiniana: characterization studies, J. Hazard. Mater. B 108 (2004) 85–94. [15] D. Teng, M. Gao, Y. Yang, B. Liu, Z. Tian, J. Wang, Biomodification of soybean meal with Bacillus subtilis or Aspergillus oryzae, Biocatal. Agric. Biotechnol. 1 (2012) 32–38. [16] X. Meng, J. Yang, X. Xu, L. Zhang, Q. Nie, M. Xian, Biodiesel production from oleaginous microorganisms, Renew. Energy 34 (2009) 1–15. [17] P. Das, W. Lei, S.S. Aziz, J.P. Obbard, Enhanced algae growth in both phototrophic and mixotrophic culture under blue light, Bioresour. Technol. 102 (2011) 3383–3387. [18] A.R.D. Bocanegra, I.G. Legarreta, F.M. Jeronimo, A.T. Campocosio, Influence of environmental and nutritional factors in the production of astaxanthin from Haematococcus pluvialis, Bioresour. Technol. 92 (2004) 209–214. [19] M.M.R. Talukder, H.Z.S. Lee, R.F. Low, L.C. Pei-Lyn, D. Warzecha, J.C. Wu, Potential use of whole cell lipase from a newly isolated Aspergillus nomius for methanolysis of palm oil to biodiesel, J. Mol. Catal. B—Enzym. 89 (2013) 108–113. [20] W.L. Teng, E. Khor, T.K. Tan, L.Y. Lim, S.C. Tan, Concurrent production of chitin from shrimp shells and fungi, Carbohydr. Res. 332 (2001) 305–316. [21] R. Gulari, R.K. Saxena, R. Gupta, R.P. Yadav, W.S. Davidson, Parametric optimization of Aspergillus terreus lipase production and its potential in ester synthesis, Process Biochem. 35 (1999) 459–464.
Contents lists available at ScienceDirect
Biochemical Engineering Journal journal homepage: www.elsevier.com/locate/bej
Immobilization of microalgae on exogenous fungal mycelium: A promising separation method to harvest both marine and freshwater microalgae Md. Mahabubur Rahman Talukder ∗ , Probir Das, Jin Chuan Wu Institute of Chemical and Engineering Sciences, Singapore
a r t i c l e
i n f o
Article history: Received 8 March 2014 Received in revised form 20 June 2014 Accepted 1 July 2014 Available online 10 July 2014 Keywords: Microalgae Immobilization Separation Optimization Screening fungus Mycelium
a b s t r a c t Although various methods have been developed for microalgae harvesting, problems such as chemical toxicity, low efficacy, low flexibility, and high cost, still remain with the current methods. In the present study, an effective biological separation method has been developed to harvest microalgae via immobilization on exogenous fungal mycelium. Fungal mycelium was added into microalgae culture and air mixed for the immobilization. The immobilized microalgae with fungal mycelium were naturally precipitated within 10 min after stopping the mixing. Both marine (e.g. Nannochloropsis sp.) and freshwater (e.g. Chlorella vulguris) microalgae were almost completely (94–97%) precipitated using the mycelium of Aspergillus nomius CCK-PDA 7#6. The precipitated mixed biomass was separated by simple filtration using sieve. A 70% initial nutrients supplement to de-algated culture medium gave high growth yield (0.43 g/l) in subsequent microalgae cultivation. The mycelium can be obtained free or at low-cost as waste from a fungal fermentation process of producing valuable products. The developed method is, therefore, very promising for economical harvesting of microalgae. © 2014 Published by Elsevier B.V.
1. Introduction Microalgae biomass is a promising renewable source for fuels and chemicals production [1,2]. However, a lack of an economical and effective method for harvesting microalgae is one of the bottlenecks for commercialization of microalgae-based fuels and chemicals [3]. Harvesting via natural sedimentation or filtration has been attempted. However, sedimentation and simple filtration are not feasible except for large species such as Arthrospira due to the small size of microalgal cell (2–20 m) and their colloidal stability in suspension. Centrifugation is used for harvesting microalgae for high value products but it is expensive if microalgae biomass is used for low value products such as biofuel. Chemical precipitation or flocculation using inorganic coagulants such as alum and ferric chloride is widely used and has been reported to be most efficient for large-scale harvesting of microalgae [4]. However, the chemical flocculation contaminates the
∗ Corresponding author. Present address: Department of Industrial Biotechnology, Institute of Chemical and Engineering Sciences, 1 Pesek Road, Jurong Island, Singapore 627833, Singapore. Tel.: +65 67963826; fax: +65 63166182. E-mail addresses: [email protected], [email protected] (Md.M.R. Talukder). http://dx.doi.org/10.1016/j.bej.2014.07.001 1369-703X/© 2014 Published by Elsevier B.V.
harvested biomass and culture medium with metal salts, causing difficulties in downstream processing of the biomass, reuse of the culture medium and waste water treatment [5]. We have recently found that the fermentation of lipid depleted algal biomass to lactic acid was significantly inhibited at Fe+ concentration >17 ppm [6]. Chemical flocculation using alum has negatively affected the ferementative production of acetone, butanol and ethanol [7]. These drawbacks can be minimized by biological harvesting via bioflocculation [8] or co-culture of other microorganisms such as fungus [9,10] and bacteria [11]. Bioflocculants such as chitosan, starch and poly ␥-glutamic acid, although effective for freshwater microalgae, undergo coiling at high ionic strengths and consequently are less effective for marine microalgae [12]. The filamentous fungi [9,10] that used to harvest microalgae via co-culturing can only grow in freshwater, and are not applicable for harvesting marine microalgae. Furthermore, the carbon source needed for growing such fungi will evoke undesirable microbial contamination of microalgae culture in an open raceway pond [11]. Microalgae harvesting via immobilization on different carriers has also been studied [13,14]. However, there are no prior studies on immobilization of microalgae on exogenous fungal mycelium. The mycelium can be obtained free or at low-cost as waste from a fungal fermentation process of producing valuable products such as enzymes and soy sauce [15,16]. Furthermore, the addition of fungal mycelium will increase
54
Md.M.R. Talukder et al. / Biochemical Engineering Journal 91 (2014) 53–57
biomass yield. As both fungus and microalgae biomasses contain lipid, carbohydrate and protein, and can be processed together, the harvesting microalgae by immobilizing them on fungal mycelium may add value to the microalgae biorefinery. In this study, a simple but effective separation method to harvest both marine and freshwater microalgae via immobilization on exogenous fungal mycelium was developed. The effects of various parameters such as fungal mycelium to microalgae biomass ratio, pH and immobilization time on separation of microalgae from the culture were investigated. 2. Material and methods 2.1. Material Nannochloropsis sp. was locally isolated [17] while Chlorella vulgaris (ATCC 13482) and Dunaliella teriolecta (UTEX LB999) were from ATCC and UTEX, respectively. Waste cooking oil containing 18.5% free fatty acids and 0.9% water was provided by Vender Horst Biodiesel Pte Ltd. (Singapore). Polypeptone were from Nihon Seiyaku, Japan. Potato Dextrose Agar (PDA) was from Difco, USA. HPLC grade methanol and hexane were from J.T. Baker, USA. Dimethylacetamide (DMAC) was from Sigma-Aldrich. All chemicals unless mentioned otherwise were of analytical grade and used as received. 2.2. Methods 2.2.1. Cultivation of microalgae A locally isolated Nannochloropsis sp. [17] was cultured in Guillard f/2 (w/o Si) medium, which was prepared using synthetic sea salt (35 g/l) and deionized water. An inoculum of 200 ml (0.3 g/l) was added into 5 l capacity glass bottle filled with 3.8 l Guillard f/2 medium in advance. The bottle was continuously irradiated with two white fluorescent lights (TL5, 28 W) placed 20 cm away from the bottle. The light intensity received by the culture was not measured. The culture was mixed with sterile-filtered air through sparger at a flow rate of 2 l/min and maintained at room temperature (22–23 ◦ C) for 6–7 days. During this time the growth of microalgae was relying on atmospheric CO2 and microalgae density reached 0.4–0.45 g/l. Microalgae density was determined based on the calibration curve prepared by plotting microalgae dry weight against the value of optical density at 680 nm (OD680 ). C. vulgaris and D. teriolecta were cultured in BAR medium [18] and modified ATCC 1174DA medium (reduced NaCl concentration of 0.5 M), respectively under the same conditions mentioned above. 2.2.2. Screening fungi Fungi were isolated from soil as described previously [19]. All fungal isolates were grown on PDA agar at 30 ◦ C for 5–7 days. Spores or mycelium of a fungus from agar plate were transferred into a conical flask containing 50 ml of polypeptone medium: waste cooking oil 30 g/l, polypeptone 10 g/l, KH2 PO4 10 g/l, NaNO3 2 g/l, MgSO4 ·5H2 O 0.5 g/l and pH 6.0. The flask was incubated at 150 rpm and 30 ◦ C for 4 days. The whole fungal culture was mixed with 200 ml microalgae culture (0.4 g/l) in a glass bottle of capacity 500 ml by air through a sparger at a flow rate of 0.5 l/min for 5 h. The air flow was stopped to precipitate the immobilized microalgae with fungal mycelium for 10 min by natural sedimentation. The six isolates, which could precipitate ca. 90% of microalgae from culture, were primarily chosen as the promising candidates. The primarily selected candidates were further screened based on the amount of microalgae immobilized per gram fungal mycelium. The isolate PDA 7#6, which was identified as Aspergillus nomius CCK-PDA 7#6 [19] and deposited in the institute of DSMZ-German
Culture Collection of Microorganism and cell cultures (DSM 26642), was finally selected. 2.2.3. Cultivation of fungus in a fermenter and preparation of mycelium A flask (250 ml) containing 100 ml polypeptone medium was inoculated by 1 ml spores suspension (106 –107 /ml) and incubated for 1 day at 150 rpm and 30 ◦ C. The culture was transferred into a 2 l fermenter (BIOSTAT, B plus, Sartorius, Germany) containing 1000 ml polypeptone medium. The cultivation conditions: pH 6.0, temperature 30 ◦ C, stirring speed 300 rpm and aeration (0.5 l/min) were kept constant. The mycelium after 4–7 days of cultivation was washed with deionized water and vacuum filtered through a filter paper (Whatman 125). The wet mycelium (filter cake) containing ca. 80% water was used for immobilising microalgae. The wet mycelium was freeze dried for 48 h to measure the water content. 2.2.4. Immobilization of microalgae on fungal mycelium The wet fungal mycelium was mixed with 1000 ml microalgae culture (density 0.4 g/l) at room temperature (22–23 ◦ C) by air through a sparger at a flow rate of 1 l/min. After a specified mixing or immobilization time, air flow was stopped to precipitate immobilized microalgae with fungal mycelium for 10 min. The immobilization was optimized by varying fungal mycelium load (0.2–1.6 g dry weight), immobilization time (0.5–10 h) and microalgae culture pH (5.5–8.5). The original pH of microalgae culture was adjusted to different pH (5.5–8.5) by supplying 100% CO2 (1 l/min) prior to the mixing with fungal mycelium. The precipitate (immobilized microalgae with fungal mycelium) was separated by simple filtration using sieve (mesh no. 18, nominal sieve opening 1 mm). The immobilization efficiency (I.E) was calculated using the formula I.E = (1 − ODb /ODa )×100, where ODb and ODa are OD680 of the culture before and after precipitating microalgae, respectively. All data are averaged of at least three replicate experiments and varied within ±3–8%. 2.2.5. Extraction of chitin from fungal mycelia Chitin in different fungal mycelia was extracted by using the method described in the literature [20]. Freeze dried mycelia were placed in 5% LiCl/DMAC solvent at a ratio of 0.5 g/75 ml with constant stirring at room temperature (ca. 23 ◦ C) for 48 h. The viscous suspension was centrifuged at 4000 rpm for 5 min, the supernatant containing dissolved chitin was collected, and distilled water (1:1) was added to the supernatant to precipitate the chitin over 24 h. The precipitate was recovered as chitin pellets by centrifugation. The chitin pellets were washed three times with distilled water to remove remaining DMAC, freeze-dried and weighed. 2.2.6. Measurement of microalgal zeta potential The zeta potential of the microalgal cells were measured at different pHs adjusted by adding acid (0.1 M HCl) or alkali (0.1 M NaOH) by using Nano Zetasizer 3600 (Malvern Instruments, Worchestershire, UK) using the electrophoretic light scattering technique. Microalgae culture (1.0 ml) was pipetted into a cuvette and inserted into the units for zeta potential measurements. Zeta potential measurements were performed in triplicate. 3. Results and discussion 3.1. Screening fungi for immobilising microalgae About 300 fungal isolates were screened to immobilize microalgae. After preliminary screening (described in Section 2.2.2), five best fungal isolates was chosen to determine the immobilization capacities of their mycelia at a fungal mycelium to microalgae biomass ratio of 1:2 (w/w). Fig. 1 shows that the immobilization
Md.M.R. Talukder et al. / Biochemical Engineering Journal 91 (2014) 53–57
16
14
14
12
12
10
10 8
8
6
6 4
4
2
2
0
0
-10 -12
80
-14 -16
60
Immobilizaon efficiency
-18
Zeta potenal
40
-20 -22
20
Zeta potential (mV)
16
Chitin content
Immobilisation efficiency (%)
Immobilisation efficiency (%)
18
100
18
Immobilization efficiency
Chitin in dry fungal biomass (%)
20
55
-24 0
5
6
7
8
9
-26
PDA 5#198 PDA 6#39 PDA 6#139 PDA 7#6 PDA 7#129
pH
Fig. 1. Microalgae (Nannochloropsis sp.) immobilization capacities of the mycelium of five best fungal isolates and their chitin content. Immobilization conditions: 1000 ml microalgae culture (0.4 g/l), fungal mycelium to microalgae biomass ratio 1:2 (w/w), pH 6, mixing time 2 h and air flow rate 1 l/min. The numbers 7 and 6 in the name of “PDA 7#6” indicate the pH of PDA agar and strain number, respectively. The same principle was applied to name the other isolates.
Fig. 3. Effect of pH on microalgae (Nannochloropsis sp.) immobilization and zeta potential of microalgae cells. The immobilization conditions were the same as those mentioned in Fig. 1 except fungal mycelium to microalgae biomass ratio (w/w) was 4:1.
efficiency (I.E.) varied with fungal isolates and the mycelium of PDA 7#6 identified as A. nomius CCK-PDA 7#6 (DSM 26642) was able to immobilize highest amount of microalgae. The immobilization could be attributed to the ionic attraction between fungal mycelium (positively charged) and microalgae cell (negatively charged). The fungal cell wall material chitin has strong positive charge aiding immobilization of microalgae. It can be seen that PDA 7#6 mycelium contained highest amount of chitin and microalgae I.E. correlated well with the chitin content in different fungal mycelia (Fig. 1). In addition to the ionic attraction, microalgae could be trapped inside the porous structure of fungal mycelium. As the net surface charge and structure of the mycelium differ with the fungal species, the amount of microalgae immobilized varied with the isolates. 3.2. Effect of carbon and nitrogen sources used for growing PDA 7#6 on microalgae immobilization
18
20
16
18
14
16 14
12
Growth Yield
10
Immobilisation efficiency
12 10
8
8
6
6
4
4
2
2
0
Waste cooking oil
Glucose
Xylose
Glycerol
Immobilisation efficiency (%)
PDA 7#6 growth yield (g/l)
The cultivation of PDA 7#6 and the preparation of mycelium were described in Section 2.2.3. Fig. 2 shows that the maximum growth yield (dry biomass per liter) reached 15.62 g/l using waste cooking oil as carbon source. The growth yield on waste cooking oil
0
Fig. 2. Effect of different carbon sources on PDA 7#6 growth yield and the immobilization capacity of its mycelium. PDA 7#6 was separately grown in a fermenter as described in Section 2.2.3. The immobilization conditions were the same as those mentioned in Fig. 1.
was 1.3 fold higher than that on glucose. This is not surprising as oil gives almost twice the usable chemical energy that glucose can deliver. The use of xylose and glycerol as a carbon source decreased PDA 7#6 growth yield. The similar result has been reported by Gulari et al., 1999 [21]. It is obvious that microalgae immobilization capacity of the mycelium was highest with glucose followed by waste cooking oil, xylose and glycerol (Fig. 2). To better understand, chitin content in the mycelium of PDA 7#6 grown on glucose and waste cooking oil were determined. The chitin content in PDA 7#6 mycelium increased from 14.8% to 18.5% (dry weight) when waste cooking was replaced by glucose. This result suggests that glucose induced the production of polysaccharide chitin in fungal cell wall, enhancing the adsorption of microalgae. Hence, the combination of waste cooking oil and glucose could be a better carbon source than either one alone to immobilize microalgae more efficiently. However, waste cooking oil yielded higher biomass and is inexpensive than glucose. Waste cooking oil was, therefore, chosen for subsequent studies. Effects of four different nitrogen sources including polypeptone, yeast extract, (NH4 )2 SO4 , and NaNO3 on PDA 7#6 growth yield and microalgae immobilization were also investigated. The biomass yield was highest with polypeptone (15.61 g/l) followed by yeast extract (14.21 g/l), NaNO3 (6.13 g/l) and (NH4 )2 SO4 (8.33 g/l) while microalgae I.E. remained almost the same (ca. 16–17%) regardless of the type of nitrogen source used. 3.3. Optimization of the immobilization of marine microalgae (Nannochloropsis sp.) Microalgae cells contain amino and carboxylic groups as well as polysaccharides and acidic compounds such as uronic acid. The contributions of each of these groups to the overall surface charge of the microalgae depend on the culture pH. The original pH (ca. 8.5) of microalgae culture was, therefore, adjusted to 5.5–8.5 by supplying 100% CO2 prior to the addition of fungal mycelium and the effect of pH on I.E. was investigated at a fungal mycelium to microalgae biomass ratio of 4:1 (w/w). Fig. 3 shows that lowering the culture pH favored the immobilization of microalgae and the maximum I.E. was ca. 85% at a pH of 6.0–6.5 after 2 h mixing. It is also obvious that immobilization efficiency correlated well with the zeta potential, which was found to lie in the range of −19.8 to −24.1 mV for Nannochloropsis sp. used in this studies for the pH range of 5.5–8.5. The pH affects zeta potential, which represents the surface charges of microalgae cells. The higher negative zeta potential (ca. −24 mV) at pH 6.0–6.5 thus suggests that the ionic attraction
Md.M.R. Talukder et al. / Biochemical Engineering Journal 91 (2014) 53–57
0.6 Microalgae dry cell biomass (g/l)
Immobilisation Efficiency (%)
100
80
60 4:1
40
2:1 1:1 20
1:2
0.5 0.4 0.3 0.2 0.1 0
2
4 6 Mixing time (h)
8
0
1
2
10
Fig. 4. Effect of mixing time on the immobilization of Nannochloropsis sp. at different ratios (w/w) of fungal mycelium to microalgae biomass. The ratio (w/w) was varied by increasing the mycelium load while the other immobilization conditions were the same as those mentioned in Fig. 1.
between microalgae (negatively charged) and fungal mycelium (positively charged) could reach to a maximum at a pH 6.0–6.5. Fig. 4 shows the effect of mixing time on the immobilization of microalgae at different ratios of fungal mycelium to microalgae biomass. The ratio was varied by adding different amounts of fungal mycelium into 1000 ml microalgae culture (0.4 g/l). It is evident from Fig. 4 that the immobilization rate increased with the increase in fungal mycelium load and the maximum immobilization efficiency (I.E.) reached 97.2% at a ratio of 4:1 after 3 h mixing. A longer mixing time was needed at a lower mycelium load. The I.E. was improved at a higher ratio because of the larger amount of mycelium available for immobilizing microalgae. At a lower ratio (e.g. 1:2), the mycelium was insufficient to yield higher I.E. To prove the robustness of the developed method, another seawater microalgae species (D. teriolecta, UTEX LB999) was immobilized at a fungal mycelium to microalgae biomass ratio of 4:1 under the same conditions used for Nannochloropsis sp. The maximum I.E. reached 92.7% after 9 h. This result suggests that the developed method could be used for other microalgae species. 3.4. Reuse of de-algated culture medium After separating the precipitate (the immobilize microalgae with fungal mycelium), 500 ml de-algated culture medium was supplemented with fresh nutrients, inoculated with 25 ml microalgae (Nannochloropsis sp.) seed culture (0.3 g/l), and air mixed. Microalgae growth in the de-algated culture medium with and without supplement (30% and 70%) and fresh f/2 medium were shown in Fig. 5. The microalgae growth yield in the de-algated medium without supplement was low (0.23 g/l after 7 days) due to the insufficient nutrients. The growth yield increased to 0.43 g/l after 7 days by supplementing 70% initial nutrients and was comparable with that in the fresh f/2 medium. This result suggests that the harvesting microalgae via immobilization on fungal mycelium allowed to reuse the de-algated culture medium and can contribute to economical microalgae cultivation. 3.5. Flexibility of the developed method to harvest freshwater microalgae The freshwater microalgae C. vulgaris (ATCC 13482) were also immobilized on PDA 7#6 fungal mycelium to examine the flexibility of the developed method. The experiment was performed using the microalgae culture of density 0.4 g/l at a fungal mycelium to microalgae biomass ratio of 4:1, pH 6.0 and air flow rate 1 l/min.
3
4
5
6
7
8
Cultivation time (day) Fig. 5. Nannochloropsis sp. growth in de-algated culture and fresh f/2 medium.
100
Immobilisation Efficiency (%)
0
80
60
40
20
0
0
1
2
3 4 Mixing time (day)
5
6
Fig. 6. Effectiveness of PDA 7#6 fungal mycelium to immobilize freshwater microalgae (C. vulgaris). Immobilization conditions were the same as those mentioned in Fig. 1 except fungal mycelium to microalgae biomass ratio was 4:1.
Fig. 6 shows that the maximum I.E. with C. vulgaris reached 89.7% after 4 h. To further improve I.E., the optimal pH for the immobilization was investigated. Fig. 7 shows that I.E. increased with the increase in pH and was maximum (94.5%) at pH 7. This result suggests that the developed method can also be applied to harvest freshwater microalgae. I.E. also correlated well with zeta potential, which represent the surface charges of the microalgae cells. The similar phenomenon was also observed in the case of 100 Immobilisation efficiency (%)
0
70% Supplement Fresh f/2 medium 30% Supplement De-algated culture medium
-22 -23
80
-24
60
Immobilisaon efficiency
-25
Zeta potenal
-26 -27
40
-28 -29
20
Zeta potential (mV)
56
-30 -31
0
-32 5
6
7 pH
8
9
Fig. 7. Immobilization of C. vulgaris at different pHs. Immobilization conditions were the same as those mentioned in Fig. 6 except mixing time was kept constant at 4 h.
Md.M.R. Talukder et al. / Biochemical Engineering Journal 91 (2014) 53–57
Nannochloropsis sp. (Fig. 3). The experiment results thus suggest that the ionic attraction between microalgae cells and fungal mycelium due to the difference in surface charge was the main reason for the immobilization of microalgae on fungal mycelium. Harvesting C. vulguris via co-cultivation with PDA 7#6 under heterotrophic condition was also investigated by following the protocol reported by Zhang and Hu [9] and the result was compared with that obtained with the method developed in this study. Small portion of the microalgae was immobilized on fungal mycelium via co-cultivation, and the supernatant remained green color even after 6 days of co-cultivation. The maximum harvesting efficiency reached 30.04%. This harvesting efficiency was slightly higher than that (ca. 25%) reported by Zhang et al. [9]. This result suggests that the method developed in the study is more promising as it is almost completely (94.3%) immobilized and precipitates the same microalgae species from the culture medium. It should be noted that PDA 7#6 does not grow in autotrophic condition. Hence, harvesting C. vulguris via co-cultivation with PDA 7#6 under autotrophic condition was not possible. 4. Conclusions Several fungi were screened to immobilize microalgae and a separation method using the mycelium of A. nomius CCK-PDA 7# 6 was developed to harvest both marine and freshwater microalgae. The immobilization capacity of the mycelium was affected by the carbon sources used for growing PDA 7#6. Zeta potential measurement revealed that the charge difference between microalgae and fungal cells affects the capability of PDA 7#6 to immobilize microalgae. The maximum immobilization efficiency for Nannochloropsis sp. reached ca. 97% at a fungal mycelium to microalgae biomass ratio 4:1, pH 6.0 and mixing time 3 h. It has been demonstrated that the immobilization of microalgae on fungal mycelium can be regarded as a simple, effective and flexible separation method to harvest microalgae. The method also contributes in economical cultivation of microalgae by the reuse of de-algated culture medium. Acknowledgments Financial support from the Agency for Science Technology and Research (A*STAR) project number (ICES/10-273A01) of Singapore is gratefully acknowledged. Special thanks to Dr. Yvonne Chow for providing us with C. vulgaris (ATCC 13482) and D. teriolecta (UTEX LB999). Thanks are extended to Mr. Ng Jun Wei for helping us to measure microalgal zeta potential.
57
References [1] S.H. Ho, S.W. Huang, C.Y. Chen, T. Hasunuma, A. Kondo, J.S. Chang, Bioethanol production using carbohydrate-rich microalgae biomass as feedstock, Bioresour. Technol. 135 (2013) 1891–2198. [2] C.Y. Chen, X.Q. Zhao, H.W. Yen, S.H. Ho, C.L. Cheng, D.J. Lee, F.W. Bai, J.S. Chang, Microalgae-based carbohydrates for biofuel production, Biochem. Eng. J. 78 (2013) 1–10. [3] M.K. Lam, K.T. Lee, Immobilisation as a feasible method to simplify the separation of microalgae water for biodiesel production, Chem. Eng. J. 191 (2012) 263–268. [4] L. Kim, K. Kwon, Y. Oh, Effect of harvesting method and growth stage on the flocculation of the green algae Botryococcus braunii, Lett. Appl. Microbiol. 27 (1998) 14–18. [5] I. Rawat, R.R. Kumar, T. Mutanda, F. Bux, Biodiesel from microalgae: a critical evaluation from laboratory to large scale production, Appl. Energy 103 (2013) 444–467. [6] M.M.R. Talukder, D. Probir, J.C. Wu, Microalgae (Nannochlorpsis salina) biomass to lactic acid and lipid, Biochem. Eng. J. 68 (2012) 109–113. [7] R.J. Anthony, J.T. Ellis, A. Sathish, A. Rahman, C.D. Miller, R.C. Sims, Effect of coagulant/flocculants on bioproducts from microalgae, Bioresour. Technol. 149 (2013) 65–70. [8] D.G. Kim, H.J. La, C.Y. Ahn, Y.H. Park, H.M. Oh, Harvest of Scenedesmus sp. with bioflocculant and reuse of culture medium for subsequent high density culture, Bioresour. Technol. 102 (2011) 3163–3168. [9] J. Zhang, B. Hu, A novel method to harvest microalgae via co-culture of filamentous fungi to form cell pellets, Bioresour. Technol. 114 (2012) 529–535. [10] S. Xie, S. Sun, S. Dai, J.S. Yuan, Efficient coagulation of microalgae in cultures with filamentous fungus, Algal Res. 2 (2013) 28–33. [11] S. Salim, R. Bosma, M.H. Vermue, R.H. Wiiffels, Harvesting of microalgae via bio-flocculation, J. Appl. Phycol. 23 (2011) 849–855. [12] D. Bilanovic, G. Shelef, Flocculation of microalgae with cationic polymers—effects of medium salinity, Biomass 17 (1988) 65–76. [13] S. Nayar, B.P.L. Goh, L.M. Chou, Settlement of marine periphytic algae in a tropical estuary, Estuar. Coast. Shelf Sci. 64 (2005) 241–248. [14] N. Akhtar, J. Iqbal, M. Iqbal, Removal and recovery of nickel (II) from aqueous solution by loofa sponge immobilised biomass of Chlorella sorokiniana: characterization studies, J. Hazard. Mater. B 108 (2004) 85–94. [15] D. Teng, M. Gao, Y. Yang, B. Liu, Z. Tian, J. Wang, Biomodification of soybean meal with Bacillus subtilis or Aspergillus oryzae, Biocatal. Agric. Biotechnol. 1 (2012) 32–38. [16] X. Meng, J. Yang, X. Xu, L. Zhang, Q. Nie, M. Xian, Biodiesel production from oleaginous microorganisms, Renew. Energy 34 (2009) 1–15. [17] P. Das, W. Lei, S.S. Aziz, J.P. Obbard, Enhanced algae growth in both phototrophic and mixotrophic culture under blue light, Bioresour. Technol. 102 (2011) 3383–3387. [18] A.R.D. Bocanegra, I.G. Legarreta, F.M. Jeronimo, A.T. Campocosio, Influence of environmental and nutritional factors in the production of astaxanthin from Haematococcus pluvialis, Bioresour. Technol. 92 (2004) 209–214. [19] M.M.R. Talukder, H.Z.S. Lee, R.F. Low, L.C. Pei-Lyn, D. Warzecha, J.C. Wu, Potential use of whole cell lipase from a newly isolated Aspergillus nomius for methanolysis of palm oil to biodiesel, J. Mol. Catal. B—Enzym. 89 (2013) 108–113. [20] W.L. Teng, E. Khor, T.K. Tan, L.Y. Lim, S.C. Tan, Concurrent production of chitin from shrimp shells and fungi, Carbohydr. Res. 332 (2001) 305–316. [21] R. Gulari, R.K. Saxena, R. Gupta, R.P. Yadav, W.S. Davidson, Parametric optimization of Aspergillus terreus lipase production and its potential in ester synthesis, Process Biochem. 35 (1999) 459–464.