Bioresource Technology 243 (2017) 163–168
Contents lists available at ScienceDirect
Bioresource Technology journal homepage: www.elsevier.com/locate/biortech
Application of agar liquid-gel transition in cultivation and harvesting of microalgae for biodiesel production Vinod Kumar a,⇑, Manisha Nanda b, Monu Verma a,c a
Dept. of Chemistry, Uttaranchal University, Dehradun, India Dept. of Biotechnology, Dolphin (PG) Institute of Biomedical and Natural Sciences, Dehradun, India c Dept. of Chemistry, Indian Institute of Technology, Roorkee, India b
h i g h l i g h t s
g r a p h i c a l a b s t r a c t
Microalgal cells are seeded in the agar
medium in four different concentrations. No difference in growth rate and biochemical contents of agar and control media. Microalgae grow within the agar gel in clusters rather than individual cells. Microalgal clusters gravimetrically settle at the bottom in 2 h. In this method agar can be reused.
a r t i c l e
i n f o
Article history: Received 27 April 2017 Received in revised form 13 June 2017 Accepted 14 June 2017 Available online 17 June 2017 Keywords: Chlorella sorokiniana Agar gel Cultivation Harvesting
a b s t r a c t In order to increase microalgal biomass productivity efficient cultivation and harvesting methods are needed against the available traditional methods. The present study focuses on the same by harvesting microalgae using agar gel. Agar medium containing bold’s basal medium (BBM) undergoes a thermoreversible gel transition. As compared to the traditional protocols, this gel is used to cultivate microalgae without even affecting the total productivity. To develop the gel for microalgae cultivation, agar was boiled in BBM. Then the agar was cooled to 35 °C and microalgae culture was added to it. After seeding the microalgae the temperature of the agar was further decreased by 10 °C to induce gelation. Instead of isolated cells microalgae were grown in clusters within the agar gel. Microalgal clusters gravimetrically settle at the bottom within 2 h. In this method agar can be reused. Ó 2017 Elsevier Ltd. All rights reserved.
1. Introduction Rapidly increasing fossil fuel demands across the globe is increasing fossil fuel depletion and carbon emissions (Liam et al., 2016). This has led to the discovery of the alternative fuels. Microalgae are currently attracting wide interests for alternative energy production. By photoautotrophic mechanism microalgae convert CO2 into biomass, lipid (fatty acid) and protein
⇑ Corresponding author. E-mail address: [email protected] (V. Kumar). http://dx.doi.org/10.1016/j.biortech.2017.06.080 0960-8524/Ó 2017 Elsevier Ltd. All rights reserved.
(Ravindran et al., 2016). The total lipid content in microalgae varies from 10 to 70% of dry algae biomass from species to species and has 20–40 times more productivity than oil crops (Mata et al., 2010; Li et al., 2011; Spolaore et al., 2006). The most tedious area of industrial algal biofuel production is harvesting of algal biomass (Greenwell et al., 2010). It is also a major factor that limits the commercial use of micro-algae (Olguı’n, 2003. Harvesting contributes to approximately 20–30% of the total cost of biofuel production (Mata et al., 2010; Molina et al., 2003; Verma et al., 2010). Thus, Industrial production of microalgae needs cost
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V. Kumar et al. / Bioresource Technology 243 (2017) 163–168
effective methods to cultivate, harvest, and separate biomass from growth media (Drexler et al., 2014). There are number of methods available for harvesting microalgae viz., sedimentation, flocculation, flotation, Tris-AcetatePhosphate-Pluronic (TAPP) gel, centrifugation and filtration. A combination of any of these can also be used (Ana et al., 2015; Estime et al., 2017). Recent studies have suggested various cost effective methods for the recovery of microalgae against the available conventional methods. Xu et al. (2011) harvested two microalgae species using nanoparticles. Vandamme et al. (2012) have used auto-flocculation, where increasing the pH value up to 11 caused flocculation due to magnesium precipitation. Fayad et al. (2017) harvested Chlorella vulgaris using electro-coagula tion-flocculation. Keeping in view the above mentioned facts, the present study focuses on the following: the first is to develop and grow microalgae culture on agar gel. Next is to characterize parameters for harvesting microalgal biomass from the agar gel. Finally, the potential of the harvested algal biomass to produce lipid and biodiesel was determined. 2. Materials and methods 2.1. Materials Chlorella singularis UUIND5 (GenBank accession number: KY745895) isolated earlier by our group from fresh water was used in this study. Chemicals used for the preparation of agar and solvents were acquired from Himedia, India. All solvents and reagents used in this study were of HPLC grade. Microalgae was grown in sterilized 250 ml conical flasks. 2.2. Preparation of agar media, culture conditions and harvesting For determining biomass productivity after every 2 days, 7 different sets of each of the four concentrations i.e., 0.2%, 0.3%, 0.5% and 1% of agar were prepared in Bold’s Basal Medium (BBM). BBM was prepared according to the composition given by Guarnieri et al. (2013). After adding the agar to BBM, boiled the solution for 5 min at 100 °C. 0.5 g L1 sodium bicarbonate as carbon source was added to agar and control medium. Further, it was allowed to cool up to 35–40 °C. Inoculated microalgae to the above agar solution and decreased the temperature by 10 °C to induce gelation. All cultures were incubated at photoperiod of 24 h: 0 h (light:dark cycle) with 200 lmol photons m2 s1 (illuminated externally) for a period of 14 days. Temperature was maintained at 25 °C. The control was maintained by inoculating microalgae into BBM alone (i.e., without agar) and incubated under same conditions. After 14 days of cultivation, the resulting agarmicroalgae matrix was used for biomass and biodiesel production analysis. Harvesting was done by gravimetrical and centrifugation methods. Microalgal clusters gravimetrically settle at the bottom within 2hrs on heating the gel for 5 min at 100 °C to bring it in liquid form (Fig. 1). Biomass samples were also hravested from agar gel by centrifugation at 1500 rpm for 5 min. The harvested cells were vacuum dried at 90 °C overnight. Dry biomass thus obtained was then measured. 2.3. Determination of cell size, biomass productivity, lipid content and lipid productivity Cell diameter (major axis recorded at 50 lm scale bar) of approximately 100 cells was estimated using ‘Image J 1.49 a’ software.
The Cell Dry Weight (CDW) (mg/L/day) was calculated according to the Eq.
P¼
ðCDW x CDW 1 Þ tx t1
where CDWx and CDW1 are the cell dry weight at time tx and t1 (the time recorded after lag phase). Accumulation of lipid droplets was monitored by rapidly using Nile red method (Arora et al., 2016) by staining the cells at regular intervals and visualizing under a fluorescent microscope (EVOS-FL, AMG, USA). Lipids were extracted from fresh microalgal biomass using a modified method of Bligh and Dyer (1959). Cell disruption was done using liquid nitrogen. The disrupted cells were treated with chloroform:methanol (2:1; v/v) and stirred at 180 rpm at room temperature for 3 h. After incubation, the suspension was centrifuged. The supernatant was then collected in a screw cap glass tube. Further 0.034% MgCl2 was added to the supernatant followed by centrifugation at 5000 rpm for 10 min. The lower phase was then separated in a glass tube. To the lower phase 2 N KCl was added followed by centrifugation at 5000 rpm for 10 min. Lastly, lower phase thus obtained was mixed with artificial organic layer (chloroform:methanol:water; 3:47:48; v/v/v). The mixture was again centrifuged at 5000 rpm for 5 min. The lower phase so obtained was vacuum dried to obtain total lipids (Patel et al., 2015). The weight of oily extract was weighed and the total lipids obtained were measured gravimetrically and then lipid content (%) was calculated by the following equations:
Lipid contentð%Þ ¼ total lipidsðgÞ=dry biomassðgÞ 2.4. Determination of biochemical composition Total protein isolation and estimation was done by the method given by Slocombe et al. (2013) with some modification. In this method 5 mg of freeze-dried micro-algae material was taken. The Sample was vortexed in either 10 ml of 24% (w/v) TCA. The homogenate was incubated in a water bath at 95 °C, for 15 min, in screw-capped micro-centrifuge tube. After incubation the sample was allowed to cool to room temperature. Then added 1 ml water to it and further centrifuged 15,000 rpm for 20 min at 4 °C. The supernatant was discarded. The precipitate was resuspended in phosphate buffer pH 7.4. Protein quantification followed the method of Lowry et al. (1951). Total carbohydrate content in lipid extracted algal biomass (10 mg) was hydrolyzed by 5 ml of 5% H2SO4 then autoclaved at 121 °C for 1 h. Supernatant obtained was used for total sugar estimation by phenol sulphuric acid method (Dubois et al., 1956) taking D-glucose as standard. 2.5. Biodiesel production – acid catalyzed transesterification The total extracted lipids were transesterified into fatty acid methyl esters (FAMEs) by methanolic sulphuric acid (6%) (Arora et al., 2016). Treating extracted lipids with methanolic sulphuric acid (6%) at 90 °C for 1 h. FAMEs were recovered by mixing with hexane and then washing with distilled water (2:1; v/v). The suspension was then centrifuged at 5000 rpm for 5 min. Hexane containing FAMEs were then transferred into new beaker. The FAMEs were analyzed using gas chromatography–mass spectroscopy (GC– MS; Agilent technologies, USA). 1 ll of sample was injected and GC–MS analysis was done as described by Patel et al. (2015). 2.6. Statistical analysis The statistical analysis was carried out by analyzing the triplicate (n = 3) results for each culture. These results have been
V. Kumar et al. / Bioresource Technology 243 (2017) 163–168
165
Fig. 1. Microalgae cells are added in the agar medium in liquid phase at 35 °C. Then, the temperature is decreased to 25 °C for gel formation and microalgal cultivation. After the cultivation gel is boiled to 2 min and gel comes in liquid form. Allowing microalgal clusters to gravimetrically settle at the bottom in 2 h. Agar can be used reused for new cultivation.
reported as mean ± SD. The data was further validated by One-way ANOVA using Graph Pad Prism software (version 6.0f) with p < 0.05.
3. Results and discussion 3.1. Biomass and biochemical productivity analysis Four different concentrations of agar were used to cultivate the microalgae. To correlate the change in cell morphology and size on exposure to agar medium, cell diameter (major axis for ellipsoid) was measured using both FE-SEM and light microscopic images. No statistical difference in cell size was recorded when grown in 0.2% agar and control. Cell size decreases with increasing concentration of agar in media (Fig. 2A). Amongst these 0.2% agar displayed the maximum biomass productivity followed by 0.3, 0.4 and 0.5%. Microalgal biomass concentrations, after 14 days cultivation period, were found to be 3.2 ± 0.1 g/l, 2.9 ± 0.3 g/l, 2.7 ± 0.2 g/l, 2.3 ± 0.1 g/l with 0.2%, 0.3%, 0.4% and 0.5% agar concentrations respectively. Dry Cell Mass (DCM) were recorded to be approximately equal in control 3.3 ± 0.1 g/l and 0.2% agar (3.2 ± 0.1 g/l) (Fig. 2B). No statistical difference was recorded in the lipid, carbohydrates and protein productivity of microalgae grown in control and 0.2% agar (Fig. 3A). Estime et al. (2017) have also reported the growth of microalgae Chlamydomonas reinhardtii with similar
biomass productivity, lipid and carbohydrate composition in TrisAcetate-Phosphate-Pluronic (TAPP) gel, as obtained from cultivation in a traditional method. In this study continuous supply of CO2 was not possible in agar gel medium, thus 0.5 g L1 sodium bicarbonate as carbon source was added to both agar and control medium. Microalgae utilize bicarbonate as the external source of carbon for photosynthesis. It derives CO2 via the action of carbonic anhydrase (Bozzo et al., 2000). NaHCO3 can be used as an alternative inorganic carbon source (Hsueh et al., 2007) for microalgal cultivation as it has greater solubility than CO2. Addition of bicarbonate increases inorganic carbon uptake which increases the productivity of algal biomass (Keyuri et al., 2016). Bicarbonate salts can easily be stored and transported to algal facilities as compared to gaseous CO2 which is costly to store and transport. FAME profile (Fig. 3B) showed that palmitic acid (C16:0), 7,10hexadecadienoic acid methyl ester (C16:2), stearic acid (C18:0), oleic acid (C18:1) and linoleic acid (C18:2) were present in maximum proportions. Myristic acid (C14:0), linoleic acid (C18:2) and C19:2 were present in small amount. 3.2. Characterization of microalgae on the basis of cellular morphology The effect of agar on the microalgae morphology was determined by FE-SEM. The cell surfaces of microalgae grown in agar medium appeared smooth and contained some traces of agar gel.
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A Cell size (µm)
6 5
4 3 2 1 0 Control 0.20%
0.30%
0.40%
0.50%
B Microalgal biomass (g/l)
3.3 3.2 3.1 3 CDW
2.9 2.8 2.7 2.6
Control 0.20%
0.30%
0.40%
0.50%
Fig. 2. (A) Average diameter of microalgal cells growing in control, 0.2%, 0.3%, 4% and 0.5%. (B) Percentage of microalgal biomass recovery through gravimetrically settling. The data are mean ± S.D. for triplicate (n = 3) results (p < 0.05).
3.3. Harvesting, energy usage, and recultivation of microalgae using thermoreversible liquid-gel transition The main reasons for high harvesting costs of microalgae are its small size and its concentration in growth media (Danquah et al., 2009; Molina et al., 2003). One of the major advantages of using agar medium is the potential for a time reducing and efficient microalgae harvesting process than the traditional method. Growth of microalgae cells within the agar gel is economical method for harvesting the algal biomass. Microalgae cell grow in clusters in agar medium. Estime et al. (2017) also reported the growth of microalgae in clusters using Tris-Acetate-PhosphatePluronic (TAPP) medium. The morphology of the algae cells affects the settling speed during the gravimetric separation. Settling velocity of a particle depends on fluid density, particle density, size, shape and concentration (Liyanage et al., 2016; Jang and Choi, 2007). Grijspeerdt and Verstraete (1997) reported that spherical shaped clusters settle faster than anisotropic shape. Smith and Friedrichs (2011) reported the correlation between the size and the settling velocity. According to Stokes’s law, settling velocity depends upon the size of algae cell/cluster in medium (Bai et al., 2006). Microalgae clusters were harvested by gravimetric separation method in this study. On boiling agar gel at 100 °C for 2 min it turned into solution phase. This allows the gravimetric separation
of microalgae which settles down at the bottom within 2 h at 40 °C. The agar used for cultivating microalgae was reused thrice by adding BBM after the harvesting process. Microalgal clusters were also harvested by centrifugation method. The growth of microalgae in clusters made the harvesting approximately ten times faster than centrifugal harvesting of microalgae grown as isolated cells. Microalgae clusters were separated in 5 min at 1500 rpm as compared to microalgae grown as isolated cells which were separated in 50 min at 1500 rpm. Commercial harvesting of microalgae using centrifugation is time consuming and costly. Combination of agar gel method with centrifugation can significantly reduce process costs of microalgae (Girma et al., 2003; Chen et al., 2014). Schlesinger et al. (2012) reported that the combination of floccula tion–sedimentation with centrifugation can reduce harvesting costs of microalgae. Comparative study of different harvesting methods, effectiveness and energy requirements given in Table 1. Electricity required to increases the temperature of 1 L of agar medium from 25 °C to 100 °C during the harvesting of microalgae.
Q ¼ m:cðDTÞ; Where Q = the heat energy transferred (joule, J), m = the mass of the liquid being heated (grams, g), c = the specific heat capacity of the liquid (joule per gram degree Celsius, J/g °C), DT = the change in temperature of the liquid (degree Celsius, °C)
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Relative (%) of proteins, carbohydrate and lipids
A 80 70 60 50
Carbohydrate
40
Protein
30
Lipid
20 10 0
Control 0.20%
0.30%
0.40%
0.50%
B
Fay Acid Composion (%)
70 C16:2
60
C19:2
50
C18:1
40
C16:1
30
C18
20
C16 C14
10
0
Control
0.20%
Fig. 3. (A) Weight percentage of protein, lipid and carbohydrate in microalgal biomass. The data are mean ± S.D. for triplicate (n = 3) results (p < 0.05). (B) Fatty acid methyl ester (FAME) profile of microalgae control and 0.2 agar.
Table 1 Harvesting methods, effectiveness and energy requirements. Harvesting process
Advantages
Disadvantages
Energy requirement (kWh)
References
Centrifugation Gravity sedimentation Filtration (natural) Filtration (pressurized) Polymer flocculation Electro-flotation Agar liquid-gel transition
Rapid, easy, efficient Low cost, potential for water recycling Less shear stress Less shear stress High efficiency, no damage to cells High efficiency Rapid, easy, efficient, potential for water recycling, no pumping cost
Very high energy input, extra pumping cost Slow process Slow process, scale up potentially has problems Scale up potentially has problems No water reuse, higher energy input High energy input Energy input
8 0.1 0.4 0.88 14 5.0 0.9 (for 100 L)
Girma et al. (2003) Shelef et al. (1984) Semerjian and Ayoub (2003) Semerjian and Ayoub (2003) Danquah et al. (2009) Azarian et al. (2007) Current method
Q ¼ 1000 4:2 ð100 25 CÞ 315 KJ ¼ 298:56 BTU ¼ 0:09 kWh
4. Conclusions In this study, cultivation and harvesting of microalgae using agar gel was investigated. The growth of microalgae in agar gel was observed as large clusters as compared to control media where it was grown as isolated cells. Clusters increase the settling veloc-
ity as compared to that of individual cells. This allowed for an energy efficient gravimetric separation of the algae biomass from the culture medium. Microalgal clusters gravimetrically settle at the bottom within 2 h. Moreover, experimental data showed that agar medium did not affect biomass and biochemical contents. In this method agar can be reused. Acknowledgements This research was supported by a grant from Uttaranchal University under Project UU-0005-2016.
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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.2017.06. 080. References Ana, I., Barros, A., Gonçalves, L., Simões, M., José, C.M., Pires, N., 2015. Harvesting techniques applied to microalgae: a review. Renewable Sustainable Energy Rev. 41, 1489–1500. Arora, N., Patel, A., Pruthi, P.A., Pruthi, V., 2016. Synergistic dynamics of nitrogen and phosphorous influences lipid productivity in Chlorella minutissima for biodiesel production. Bioresour. Technol. 213, 79–87. Azarian, G.H., Mesdaghinia, A.R., Vaezi, F., Nabizadeh, R., Nematollahi, D., 2007. Algae removal by electro-coagulation process, application for treatment of the effluent from an industrial wastewater treatment plant. Iran. J. Public Health 36, 57–64. Bai, G., Jiang, W., Chen, L., 2006. Effect of interfacial thermal resistance on effective thermal conductivity of MoSi2/SiC composites. Mater. Trans. 47 (4), 1247–1249. Bligh, B.G., Dyer, W.J., 1959. A rapid method for total lipid extraction and purification. Can. J. Biochem. Phys. 37, 911–917. Bozzo, G.G., Colman, B., Matsuda, Y., 2000. Active transport of CO2 and bicarbonate is induced in response to external CO2 concentration in the green alga Chlorella kessleri. J. Exp. Bot. 51, 1341–1348. Chen, G., Zhao, L., Qi, Y., Cui, Y.U., 2014. Chitosan and its derivatives applied in harvesting microalgae for biodiesel production: an outlook. J. Nanomater. http://dx.doi.org/10.1155/2014/217537. Danquah, M.K., Ang, L., Uduman, N., Moheimani, N., Forde, G.M., 2009. Dewatering of microalgal culture for biodiesel production: exploring polymer flocculation and tangential flow filtration. J. Chem. Technol. Biotechnol. 84, 1078–1083. Drexler Ivy, L.C., Yeh, H.D., 2014. Membrane applications for microalgae cultivation and harvesting: a review. Rev. Environ. Sci. Biotechnol. 13, 487–504. Dubois, M., Gilles, K.A., Hamilton, J.K., Rebers, P.A., Smith, F., 1956. Colorimetric method for determination of sugars and related substances. Anal. Chem. 28, 350–356. Estime, B., Ren, D., Sureshkumar, R., 2017. Cultivation and energy efficient harvesting of microalgae using thermoreversible sol-gel transition. Sci. Rep. 10.1038/srep40725. Fayad, N., Yehya, T., Audonnet, F., Vial, C., 2017. Harvesting of microalgae Chlorella vulgaris using electro-coagulation-flocculation in the batch mode. Algal Res. 25, 1–11. Girma, E., Belarbi, E.H., Fernandez, G.A., Medina, A.R., Chisti, Y., 2003. Recovery of microalgal biomass and metabolites: process options and economics. Biotechnol. Adv. 20, 491–515. Greenwell, H.C., Laurens, L.M.L., Shields, R.J., Lovitt, R.W., Flynn, K.J., 2010. Placing microalgae on the biofuels priority list: a review of the technological challenges. J. R. Soc. Interface 7, 703–726. Grijspeerdt, K., Verstraete, W., 1997. Image analysis to estimate the settleability and concentration of activated sludge. Water Res. 31, 1126–1134. Guarnieri, M.T., Nag, A., Yang, S., Pienkos, P.T., 2013. Proteomic analysis of Chlorella vulgaris: potential targets for enhanced lipid accumulation. J. Proteome 93, 245– 253. Hsueh, H.T., Chu, H., Yu, S.T., 2007. A batch study on the biofixation of carbon dioxide in the absorbed solution from a chemical wet scrubber by hot spring and marine algae. Chemosphere 66, 878–886.
Jang, S.P., Choi, S.U.S., 2007. Effects of various parameters on nanofluid thermal conductivity. J. Heat Transfer 129 (5), 617–623. Keyuri, M., Vishaka, S., Sangeetha, A.G., Sibi, G., 2016. Sodium bicarbonate as inorganic carbon source for higher biomass and lipid production integrated carbon capture in Chlorella vulgaris. Achiev. Life Sci. 10, 111–117. Li, Y., Chen, Y.F., Chen, P., Min, M., Zhou, W., Martinez, B., Zhu, J., Ruan, R., 2011. Characterization of a microalga Chlorella sp. well adapted to highly concentrated municipal wastewater for nutrient removal and biodiesel production. Bioresour. Technol. 102, 5138–5144. Liam, W., Ian, R., John, F., Hankamer, B., 2016. Trading off global fuel supply, CO2 emissions and sustainable development. PLoS One 11 (3), e0149406. Liyanage, D.D., Rajika, Thamali, J.K.A., Kumbalatara, A.A.K., Weliwita, J.A., Witharana, S., 2016. An analysis of nanoparticle settling times in liquids. J. Nanomater. http://dx.doi.org/10.1155/2016/7061838. Lowry, O.H., Rosebrough, N.J., Farr, A.L., Randall, R.J., 1951. Protein measurement with the folin phenol reagent. J. Biol. Chem. 193, 265–275. Mata, T.M., Martins, A.A., Caetano, N.S., 2010. Microalgae for biodiesel production and other applications: a review. Renewable Sustainable Energy Rev. 14, 217– 232. Molina, G.E., Belarbi, E.H., Acien, F.F.G., Robles, M.A., Yusuf, C., 2003. Recovery of microalgal biomass and metabolites: process options and economics. Biotechnol. Adv. 20, 491–515. Olguı’n, E.J., 2003. Phycoremediation: key issues for cost-effective nutrient removal processes. Biotechnol. Adv. 22, 81–91. Patel, A., Sindhu, D.K., Arora, N., Singh, R.P., Pruthi, V., Pruthi, P.A., 2015. Biodiesel production from non-edible lignocellulosic biomass of Cassia fistula L. fruit pulp using oleaginous yeast Rhodosporidium kratochvilovae HIMPA1. Bioresour. Technol. 197, 91–98. Ravindran, B., Gupta, S.K., Cho, W.M., Kim, J.K., Lee, S.R., Jeong, K.H., Lee, D.J., Choi, H. C., 2016. Microalgae potential and multiple roles—current progress and future prospects—an overview. Sustainability 8, 1215. Schlesinger, A., Eisenstadt, D., Bar-Gil, A., Carmely, H., Einbinder, S., Gressel, J., 2012. Inexpensive non-toxic flocculation of microalgae contradicts theories; overcoming a major hurdle to bulk algal production. Biotechnol. Adv. 30, 1023–1030. Semerjian, L., Ayoub, G.M., 2003. High-pH–magnesium coagulation–flocculation in wastewater treatment. Adv. Environ. Res. 7, 389–403. Shelef, G., Sukenik, A., Green, M., 1984. Microalgae Harvesting and Processing: A Literature Review. Report. Solar Energy Research Institute, Golden, CO. SERI Report No. 231-2396. Slocombe, P.S., Ross, M., Mc.Neill Thomas, S., Stanley, S.M., 2013. A rapid and general method for measurement of protein in micro-algal biomass. Bioresour. Technol. 129, 51–57. Smith, S.J., Friedrichs, C.T., 2011. Size and settling velocities of cohesive flocs and suspended sediment aggregates in a trailing suction hopper dredge plume. Cont. Shelf Res. 31, S50–S63. Spolaore, P., Joannis-Cassan, C., Duran, E., Isambert, A., 2006. Commercial applications of microalgae. J. Biosci. Bioeng. 101, 87–96. Vandamme, D., Foubert, I., Fraeye, I., Meesschaert, B., Muylaert, K., 2012. Flocculation of Chlorella vulgaris induced by high pH: role of magnesium and calcium and practical implications. Bioresour. Technol. 105, 114–119. Verma, N.M., Mehrotra, S., Shukla, A., Mishra, B.N., 2010. Prospective of biodiesel production utilizing microalgae as the cell factories: a comprehensive discussion. Afr. J. Biotechnol. 9, 1402–1411. Xu, L., Guo, C., Wang, F., Zheng, S., Liu, C., Z., 2011. A simple and rapid harvesting method for microalgae by in situ magnetic separation. Bioresour. Technol. 102, 10047–10051.
Contents lists available at ScienceDirect
Bioresource Technology journal homepage: www.elsevier.com/locate/biortech
Application of agar liquid-gel transition in cultivation and harvesting of microalgae for biodiesel production Vinod Kumar a,⇑, Manisha Nanda b, Monu Verma a,c a
Dept. of Chemistry, Uttaranchal University, Dehradun, India Dept. of Biotechnology, Dolphin (PG) Institute of Biomedical and Natural Sciences, Dehradun, India c Dept. of Chemistry, Indian Institute of Technology, Roorkee, India b
h i g h l i g h t s
g r a p h i c a l a b s t r a c t
Microalgal cells are seeded in the agar
medium in four different concentrations. No difference in growth rate and biochemical contents of agar and control media. Microalgae grow within the agar gel in clusters rather than individual cells. Microalgal clusters gravimetrically settle at the bottom in 2 h. In this method agar can be reused.
a r t i c l e
i n f o
Article history: Received 27 April 2017 Received in revised form 13 June 2017 Accepted 14 June 2017 Available online 17 June 2017 Keywords: Chlorella sorokiniana Agar gel Cultivation Harvesting
a b s t r a c t In order to increase microalgal biomass productivity efficient cultivation and harvesting methods are needed against the available traditional methods. The present study focuses on the same by harvesting microalgae using agar gel. Agar medium containing bold’s basal medium (BBM) undergoes a thermoreversible gel transition. As compared to the traditional protocols, this gel is used to cultivate microalgae without even affecting the total productivity. To develop the gel for microalgae cultivation, agar was boiled in BBM. Then the agar was cooled to 35 °C and microalgae culture was added to it. After seeding the microalgae the temperature of the agar was further decreased by 10 °C to induce gelation. Instead of isolated cells microalgae were grown in clusters within the agar gel. Microalgal clusters gravimetrically settle at the bottom within 2 h. In this method agar can be reused. Ó 2017 Elsevier Ltd. All rights reserved.
1. Introduction Rapidly increasing fossil fuel demands across the globe is increasing fossil fuel depletion and carbon emissions (Liam et al., 2016). This has led to the discovery of the alternative fuels. Microalgae are currently attracting wide interests for alternative energy production. By photoautotrophic mechanism microalgae convert CO2 into biomass, lipid (fatty acid) and protein
⇑ Corresponding author. E-mail address: [email protected] (V. Kumar). http://dx.doi.org/10.1016/j.biortech.2017.06.080 0960-8524/Ó 2017 Elsevier Ltd. All rights reserved.
(Ravindran et al., 2016). The total lipid content in microalgae varies from 10 to 70% of dry algae biomass from species to species and has 20–40 times more productivity than oil crops (Mata et al., 2010; Li et al., 2011; Spolaore et al., 2006). The most tedious area of industrial algal biofuel production is harvesting of algal biomass (Greenwell et al., 2010). It is also a major factor that limits the commercial use of micro-algae (Olguı’n, 2003. Harvesting contributes to approximately 20–30% of the total cost of biofuel production (Mata et al., 2010; Molina et al., 2003; Verma et al., 2010). Thus, Industrial production of microalgae needs cost
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effective methods to cultivate, harvest, and separate biomass from growth media (Drexler et al., 2014). There are number of methods available for harvesting microalgae viz., sedimentation, flocculation, flotation, Tris-AcetatePhosphate-Pluronic (TAPP) gel, centrifugation and filtration. A combination of any of these can also be used (Ana et al., 2015; Estime et al., 2017). Recent studies have suggested various cost effective methods for the recovery of microalgae against the available conventional methods. Xu et al. (2011) harvested two microalgae species using nanoparticles. Vandamme et al. (2012) have used auto-flocculation, where increasing the pH value up to 11 caused flocculation due to magnesium precipitation. Fayad et al. (2017) harvested Chlorella vulgaris using electro-coagula tion-flocculation. Keeping in view the above mentioned facts, the present study focuses on the following: the first is to develop and grow microalgae culture on agar gel. Next is to characterize parameters for harvesting microalgal biomass from the agar gel. Finally, the potential of the harvested algal biomass to produce lipid and biodiesel was determined. 2. Materials and methods 2.1. Materials Chlorella singularis UUIND5 (GenBank accession number: KY745895) isolated earlier by our group from fresh water was used in this study. Chemicals used for the preparation of agar and solvents were acquired from Himedia, India. All solvents and reagents used in this study were of HPLC grade. Microalgae was grown in sterilized 250 ml conical flasks. 2.2. Preparation of agar media, culture conditions and harvesting For determining biomass productivity after every 2 days, 7 different sets of each of the four concentrations i.e., 0.2%, 0.3%, 0.5% and 1% of agar were prepared in Bold’s Basal Medium (BBM). BBM was prepared according to the composition given by Guarnieri et al. (2013). After adding the agar to BBM, boiled the solution for 5 min at 100 °C. 0.5 g L1 sodium bicarbonate as carbon source was added to agar and control medium. Further, it was allowed to cool up to 35–40 °C. Inoculated microalgae to the above agar solution and decreased the temperature by 10 °C to induce gelation. All cultures were incubated at photoperiod of 24 h: 0 h (light:dark cycle) with 200 lmol photons m2 s1 (illuminated externally) for a period of 14 days. Temperature was maintained at 25 °C. The control was maintained by inoculating microalgae into BBM alone (i.e., without agar) and incubated under same conditions. After 14 days of cultivation, the resulting agarmicroalgae matrix was used for biomass and biodiesel production analysis. Harvesting was done by gravimetrical and centrifugation methods. Microalgal clusters gravimetrically settle at the bottom within 2hrs on heating the gel for 5 min at 100 °C to bring it in liquid form (Fig. 1). Biomass samples were also hravested from agar gel by centrifugation at 1500 rpm for 5 min. The harvested cells were vacuum dried at 90 °C overnight. Dry biomass thus obtained was then measured. 2.3. Determination of cell size, biomass productivity, lipid content and lipid productivity Cell diameter (major axis recorded at 50 lm scale bar) of approximately 100 cells was estimated using ‘Image J 1.49 a’ software.
The Cell Dry Weight (CDW) (mg/L/day) was calculated according to the Eq.
P¼
ðCDW x CDW 1 Þ tx t1
where CDWx and CDW1 are the cell dry weight at time tx and t1 (the time recorded after lag phase). Accumulation of lipid droplets was monitored by rapidly using Nile red method (Arora et al., 2016) by staining the cells at regular intervals and visualizing under a fluorescent microscope (EVOS-FL, AMG, USA). Lipids were extracted from fresh microalgal biomass using a modified method of Bligh and Dyer (1959). Cell disruption was done using liquid nitrogen. The disrupted cells were treated with chloroform:methanol (2:1; v/v) and stirred at 180 rpm at room temperature for 3 h. After incubation, the suspension was centrifuged. The supernatant was then collected in a screw cap glass tube. Further 0.034% MgCl2 was added to the supernatant followed by centrifugation at 5000 rpm for 10 min. The lower phase was then separated in a glass tube. To the lower phase 2 N KCl was added followed by centrifugation at 5000 rpm for 10 min. Lastly, lower phase thus obtained was mixed with artificial organic layer (chloroform:methanol:water; 3:47:48; v/v/v). The mixture was again centrifuged at 5000 rpm for 5 min. The lower phase so obtained was vacuum dried to obtain total lipids (Patel et al., 2015). The weight of oily extract was weighed and the total lipids obtained were measured gravimetrically and then lipid content (%) was calculated by the following equations:
Lipid contentð%Þ ¼ total lipidsðgÞ=dry biomassðgÞ 2.4. Determination of biochemical composition Total protein isolation and estimation was done by the method given by Slocombe et al. (2013) with some modification. In this method 5 mg of freeze-dried micro-algae material was taken. The Sample was vortexed in either 10 ml of 24% (w/v) TCA. The homogenate was incubated in a water bath at 95 °C, for 15 min, in screw-capped micro-centrifuge tube. After incubation the sample was allowed to cool to room temperature. Then added 1 ml water to it and further centrifuged 15,000 rpm for 20 min at 4 °C. The supernatant was discarded. The precipitate was resuspended in phosphate buffer pH 7.4. Protein quantification followed the method of Lowry et al. (1951). Total carbohydrate content in lipid extracted algal biomass (10 mg) was hydrolyzed by 5 ml of 5% H2SO4 then autoclaved at 121 °C for 1 h. Supernatant obtained was used for total sugar estimation by phenol sulphuric acid method (Dubois et al., 1956) taking D-glucose as standard. 2.5. Biodiesel production – acid catalyzed transesterification The total extracted lipids were transesterified into fatty acid methyl esters (FAMEs) by methanolic sulphuric acid (6%) (Arora et al., 2016). Treating extracted lipids with methanolic sulphuric acid (6%) at 90 °C for 1 h. FAMEs were recovered by mixing with hexane and then washing with distilled water (2:1; v/v). The suspension was then centrifuged at 5000 rpm for 5 min. Hexane containing FAMEs were then transferred into new beaker. The FAMEs were analyzed using gas chromatography–mass spectroscopy (GC– MS; Agilent technologies, USA). 1 ll of sample was injected and GC–MS analysis was done as described by Patel et al. (2015). 2.6. Statistical analysis The statistical analysis was carried out by analyzing the triplicate (n = 3) results for each culture. These results have been
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Fig. 1. Microalgae cells are added in the agar medium in liquid phase at 35 °C. Then, the temperature is decreased to 25 °C for gel formation and microalgal cultivation. After the cultivation gel is boiled to 2 min and gel comes in liquid form. Allowing microalgal clusters to gravimetrically settle at the bottom in 2 h. Agar can be used reused for new cultivation.
reported as mean ± SD. The data was further validated by One-way ANOVA using Graph Pad Prism software (version 6.0f) with p < 0.05.
3. Results and discussion 3.1. Biomass and biochemical productivity analysis Four different concentrations of agar were used to cultivate the microalgae. To correlate the change in cell morphology and size on exposure to agar medium, cell diameter (major axis for ellipsoid) was measured using both FE-SEM and light microscopic images. No statistical difference in cell size was recorded when grown in 0.2% agar and control. Cell size decreases with increasing concentration of agar in media (Fig. 2A). Amongst these 0.2% agar displayed the maximum biomass productivity followed by 0.3, 0.4 and 0.5%. Microalgal biomass concentrations, after 14 days cultivation period, were found to be 3.2 ± 0.1 g/l, 2.9 ± 0.3 g/l, 2.7 ± 0.2 g/l, 2.3 ± 0.1 g/l with 0.2%, 0.3%, 0.4% and 0.5% agar concentrations respectively. Dry Cell Mass (DCM) were recorded to be approximately equal in control 3.3 ± 0.1 g/l and 0.2% agar (3.2 ± 0.1 g/l) (Fig. 2B). No statistical difference was recorded in the lipid, carbohydrates and protein productivity of microalgae grown in control and 0.2% agar (Fig. 3A). Estime et al. (2017) have also reported the growth of microalgae Chlamydomonas reinhardtii with similar
biomass productivity, lipid and carbohydrate composition in TrisAcetate-Phosphate-Pluronic (TAPP) gel, as obtained from cultivation in a traditional method. In this study continuous supply of CO2 was not possible in agar gel medium, thus 0.5 g L1 sodium bicarbonate as carbon source was added to both agar and control medium. Microalgae utilize bicarbonate as the external source of carbon for photosynthesis. It derives CO2 via the action of carbonic anhydrase (Bozzo et al., 2000). NaHCO3 can be used as an alternative inorganic carbon source (Hsueh et al., 2007) for microalgal cultivation as it has greater solubility than CO2. Addition of bicarbonate increases inorganic carbon uptake which increases the productivity of algal biomass (Keyuri et al., 2016). Bicarbonate salts can easily be stored and transported to algal facilities as compared to gaseous CO2 which is costly to store and transport. FAME profile (Fig. 3B) showed that palmitic acid (C16:0), 7,10hexadecadienoic acid methyl ester (C16:2), stearic acid (C18:0), oleic acid (C18:1) and linoleic acid (C18:2) were present in maximum proportions. Myristic acid (C14:0), linoleic acid (C18:2) and C19:2 were present in small amount. 3.2. Characterization of microalgae on the basis of cellular morphology The effect of agar on the microalgae morphology was determined by FE-SEM. The cell surfaces of microalgae grown in agar medium appeared smooth and contained some traces of agar gel.
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A Cell size (µm)
6 5
4 3 2 1 0 Control 0.20%
0.30%
0.40%
0.50%
B Microalgal biomass (g/l)
3.3 3.2 3.1 3 CDW
2.9 2.8 2.7 2.6
Control 0.20%
0.30%
0.40%
0.50%
Fig. 2. (A) Average diameter of microalgal cells growing in control, 0.2%, 0.3%, 4% and 0.5%. (B) Percentage of microalgal biomass recovery through gravimetrically settling. The data are mean ± S.D. for triplicate (n = 3) results (p < 0.05).
3.3. Harvesting, energy usage, and recultivation of microalgae using thermoreversible liquid-gel transition The main reasons for high harvesting costs of microalgae are its small size and its concentration in growth media (Danquah et al., 2009; Molina et al., 2003). One of the major advantages of using agar medium is the potential for a time reducing and efficient microalgae harvesting process than the traditional method. Growth of microalgae cells within the agar gel is economical method for harvesting the algal biomass. Microalgae cell grow in clusters in agar medium. Estime et al. (2017) also reported the growth of microalgae in clusters using Tris-Acetate-PhosphatePluronic (TAPP) medium. The morphology of the algae cells affects the settling speed during the gravimetric separation. Settling velocity of a particle depends on fluid density, particle density, size, shape and concentration (Liyanage et al., 2016; Jang and Choi, 2007). Grijspeerdt and Verstraete (1997) reported that spherical shaped clusters settle faster than anisotropic shape. Smith and Friedrichs (2011) reported the correlation between the size and the settling velocity. According to Stokes’s law, settling velocity depends upon the size of algae cell/cluster in medium (Bai et al., 2006). Microalgae clusters were harvested by gravimetric separation method in this study. On boiling agar gel at 100 °C for 2 min it turned into solution phase. This allows the gravimetric separation
of microalgae which settles down at the bottom within 2 h at 40 °C. The agar used for cultivating microalgae was reused thrice by adding BBM after the harvesting process. Microalgal clusters were also harvested by centrifugation method. The growth of microalgae in clusters made the harvesting approximately ten times faster than centrifugal harvesting of microalgae grown as isolated cells. Microalgae clusters were separated in 5 min at 1500 rpm as compared to microalgae grown as isolated cells which were separated in 50 min at 1500 rpm. Commercial harvesting of microalgae using centrifugation is time consuming and costly. Combination of agar gel method with centrifugation can significantly reduce process costs of microalgae (Girma et al., 2003; Chen et al., 2014). Schlesinger et al. (2012) reported that the combination of floccula tion–sedimentation with centrifugation can reduce harvesting costs of microalgae. Comparative study of different harvesting methods, effectiveness and energy requirements given in Table 1. Electricity required to increases the temperature of 1 L of agar medium from 25 °C to 100 °C during the harvesting of microalgae.
Q ¼ m:cðDTÞ; Where Q = the heat energy transferred (joule, J), m = the mass of the liquid being heated (grams, g), c = the specific heat capacity of the liquid (joule per gram degree Celsius, J/g °C), DT = the change in temperature of the liquid (degree Celsius, °C)
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Relative (%) of proteins, carbohydrate and lipids
A 80 70 60 50
Carbohydrate
40
Protein
30
Lipid
20 10 0
Control 0.20%
0.30%
0.40%
0.50%
B
Fay Acid Composion (%)
70 C16:2
60
C19:2
50
C18:1
40
C16:1
30
C18
20
C16 C14
10
0
Control
0.20%
Fig. 3. (A) Weight percentage of protein, lipid and carbohydrate in microalgal biomass. The data are mean ± S.D. for triplicate (n = 3) results (p < 0.05). (B) Fatty acid methyl ester (FAME) profile of microalgae control and 0.2 agar.
Table 1 Harvesting methods, effectiveness and energy requirements. Harvesting process
Advantages
Disadvantages
Energy requirement (kWh)
References
Centrifugation Gravity sedimentation Filtration (natural) Filtration (pressurized) Polymer flocculation Electro-flotation Agar liquid-gel transition
Rapid, easy, efficient Low cost, potential for water recycling Less shear stress Less shear stress High efficiency, no damage to cells High efficiency Rapid, easy, efficient, potential for water recycling, no pumping cost
Very high energy input, extra pumping cost Slow process Slow process, scale up potentially has problems Scale up potentially has problems No water reuse, higher energy input High energy input Energy input
8 0.1 0.4 0.88 14 5.0 0.9 (for 100 L)
Girma et al. (2003) Shelef et al. (1984) Semerjian and Ayoub (2003) Semerjian and Ayoub (2003) Danquah et al. (2009) Azarian et al. (2007) Current method
Q ¼ 1000 4:2 ð100 25 CÞ 315 KJ ¼ 298:56 BTU ¼ 0:09 kWh
4. Conclusions In this study, cultivation and harvesting of microalgae using agar gel was investigated. The growth of microalgae in agar gel was observed as large clusters as compared to control media where it was grown as isolated cells. Clusters increase the settling veloc-
ity as compared to that of individual cells. This allowed for an energy efficient gravimetric separation of the algae biomass from the culture medium. Microalgal clusters gravimetrically settle at the bottom within 2 h. Moreover, experimental data showed that agar medium did not affect biomass and biochemical contents. In this method agar can be reused. Acknowledgements This research was supported by a grant from Uttaranchal University under Project UU-0005-2016.
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