Progress in energy from microalgae: A review

Renewable and Sustainable Energy Reviews 27 (2013) 128–148 Contents lists available at SciVerse ScienceDirect Renewable and Sustainable Energy Reviews journal homepage: www.elsevier.com/locate/rser Progress in energy from microalgae: A review Ali Bahadar a, M. Bilal Khan b,n a b School of Chemical and Materials Engineering, National University of Sciences and Technology, H-12 Sector, Islamabad, Pakistan Centre for Energy Systems, USAID Centre for Advanced Studies, National University of Sciences and Technology, Sector H-12, Islamabad, Pakistan art ic l e i nf o a b s t r a c t Article history: Received 1 January 2013 Received in revised form 22 June 2013 Accepted 24 June 2013 Microalgae have great potential as renewable fuel sources, but a dire need exists for high-level academic and industrial research into their growth and bioprocessing. New algae strains that efficiently use CO2 and wastes as nutrients, novel oil extraction methods, and industrial-scale designs for fuel production are imperative for long-term energy sustainability. A particular challenge to research in this field is the transition from pilot studies to industrial operations, which often exposes algae cells and their products to hostile environments, reducing yields. Hence, a need exists to integrate algae cell engineering with predictive bioprocess engineering to ensure economic and environmental feasibility and minimize the number of full-scale trials that fail. This review provides a brief overview of biofuel production from microalgal biomass. It highlights the most promising microalgae species for different types of fuel, the proper choice of photobioreactor and process parameters, product extraction techniques, and the main biofuel products. The main goal of this paper is to promote research into energetically- and environmentally-favorable technologies via the development of better algal strains and separation, extraction, and conversion methods. & 2013 Elsevier Ltd. All rights reserved. Keywords: Microalgae Photobioreactors Extraction Biofuels Biodiesel Liquefaction Contents 1. 2. 3. 4. 5. n Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biofuel algae strains and their culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Algae cultivation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Cultivation in ponds and raceways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Cultivation in photobioreactors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Extraction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Organic solvent extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Soxhlet solvent extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Ultrasonic-assisted organic solvent extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4. Microwave-assisted organic solvent extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5. Direct liquefaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6. Supercritical (CO2) fluid extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Microalgae biofuel products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. Bio-hydrogen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.1. Biophotolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.2. Direct photolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.3. Indirect photolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. Biodiesel. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.2. Transesterification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.3. Transesterification via liquefaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.4. Supercritical methanol transesterification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3. Biomethanol. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Corresponding author. Tel.: +925 190 855 100. E-mail address: [email protected] (M. Bilal Khan). 1364-0321/$ - see front matter & 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.rser.2013.06.029 129 129 132 132 132 136 137 137 137 137 137 137 138 138 138 138 139 139 139 140 140 140 142 A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 5.4. 5.5. Bioethanol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biohydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5.1. Pyrolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5.2. Direct liquefaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Production costs and life cycle assessment of algae derived fuels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Appendix A. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction Declining petroleum reserves, increasing fuel demands for commuting and power generation, and environmental problems have necessitated the search for sustainable sources of energy. Liquid biofuels have received much attention in this context. Numerous biomass feedstocks, including both terrestrial plants and aquatic algae, have been reported to produce renewable fuels to replace fossil fuels and mitigate carbon dioxide (CO2) emissions [1]. Marine biomass has the potential to provide both food and fuel at productivity levels per unit area that match or exceed those of terrestrial crops [2]. Among the available feedstocks, aquatic microalgae are ideal for producing liquid fuels [3]. Their rapid growth, high biomass yields, product diversity, and ease of harvest from ponds or closed photobioreactor systems give them excellent commercial potential as sustainable environmentally-friendly carbon-neutral fuel sources [4,5]. Microalgae are photosynthetic microorganisms from either marine or freshwater environments and can include bacteria (Cyanobacteria), diatoms (Chromalveolata), other protists (e.g., Chromista), and unicellular plants (e.g., Chlorophyta). They exist as individual cells or as chains of cells but do not form differentiated, multicellular organisms, as do macroalgae. Some species contain more than 70% lipids (dry weight basis) [6], and they grow exponentially under optimal conditions. They have been investigated because of their high photosynthetic rates (e.g., 6.9  104 cells/mL/h) [7] and efficiency, with biofuel yields up to 12,000 L/ha, which is much higher than terrestrial plants [8,9]. Microalgae currently cost more to cultivate than traditional crops and yield lower profits, but they remain a promising tool for future energy needs because they occupy less area than other energy crops, like Jatropha (Euphorbiaceae) or Pongamia (Fabaceae) [10,11]. The high CO2 and nutrient demands of microalgae can be met using flue gases and waste water from other industrial processes, providing ecological benefits while lowering the cost of biomass production. Some microalgae can also synthesize desirable compounds, like β-carotenoids, docosahexaenoic acid (DHA), and astaxanthin, with commercial or pharmaceutical applications [6,12,13]. Moreover, microalgae have shown great potential to produce a wide spectrum of fuel products in pilot studies: (1) hydrogen (H2) via direct and indirect biophotolysis, (2) biodiesel through transesterification, (3) biomethane via anaerobic digestion, (4) bioethanol by fermentation, (5) bio-oil via thermochemical conversion, and (6) green diesel and gasoline through direct catalytic hydrothermal liquefaction [14–17]. However, the transition from small-scale to industrial-scale operations often exposes algae cells to hostile conditions, with consequent declines in product yields. The recovery and concentration of algae from highly dilute suspensions, in particular, requires steps that can prematurely lyse cells and reduce extract yields. Therefore, an urgent need exists to integrate the best algae cell and bioprocessing engineering methods to ensure economic and environmental feasibility and to minimize the number of full-scale trials that fail. 129 142 142 142 143 143 145 145 145 Conditions for a technically and economically viable biofuel resource are that it should be competitive or cost less than petroleum fuels; should require low to no additional land use; should enable air quality improvement (e.g., CO2 sequesteration); and should require minimal water use [18]. Production cost associated with microalgae biofuel is the major barrier in its commercialization but still fuels from algae are promising, as they may already be viewed as competitive with petroleum fuels, if the full environmental impact of the latter types of fuels is taken into consideration. Issues of climate change may force us to move beyond petroleum long before it runs out [19,20]. A significant number of startup companies are making attempts to commercialize algae fuels. The table in Appendix A shows a list of 43 companies [19,21–23] which are actively participating in the development of algal fuels. Singh [21] presented the region wise percentage of companies around the world producing algae fuels. Most of the companies contributing to development of algal biofuel are based in America i.e. 78%, 13% are Europe based and 9% are from other regions. There are many research groups, companies and governments involved in the commercialization of algae based biofuels. The Department of Energy (DOE) currently spends about $85 million on 30 research projects to develop algal biofuels. Obama committed another $14 million to the idea. According to Piccolo [24] the biggest algae investment in the EU is the £26 million publically funded project by the UK Carbon Trust which planned to build a large algae farm in Northern Africa [25]. In another development, a Spanish renewable energy company Aurantia and Green Fuel Tech of Massachusetts (USA) formed a partnership through a $92 million project in 2007 to produce algae oil. In the long run, this project will target to scale up to 100 ha of algae greenhouses, producing 25,000 t of algae biomass per annum. The plant will obtain its CO2 from a cement plant near Jerez in Spain. In yet another endeavor, an Italian energy company, Eni, has installed a 1 ha pilot facility for algae oil production in Gela, Sicily. This project is testing the photobioreactor facility as well as open ponds [21,24]. This review provides an overview of recent progress in algal energy production, summarizing the most promising algal strains, cultivation methods, harvesting and extraction techniques, and conversion technologies for the commercial application of microalgae. We provide a critical analysis of the potential of selected microalgal strains and processes to yield biofuels. This review does not consider multicellular macroalgae, which are more difficult to grow in bioreactors, yield fewer products at lower concentrations, and have not yet been researched as thoroughly as microalgae [26,27]. Furthermore, microalgae have less complex structures, faster growth rates, and higher oil contents than do macroalgae. 2. Biofuel algae strains and their culture Like most other photosynthetic organisms, microalgae use light energy to convert CO2 into organic compounds. Microalgae are more photosynthetically efficient than higher plants [28]; thus 130 Table 1 Microalgae strain cultivation in optimized cultures. Light intensity Temp (lmol/m2/s) Medium Botryococcus braunii 765 (29–75%) 1507 10 BG11 medium (per liter): 1500 mg NaNO3, 40 mg K2HPO4
3H2O, 75 mg MgSO4
7H2O, 36 mg 20% CO2 CaCl2
2H2O, 6 mg C6H8O7
H2O, 6 mg Fe(NH4)3C18H10O14, 1 mg Na2
EDTA, 20 mg Na2CO3, 2.86 mg H3BO3, 1.81 mg MnCl2
H2O, 0.222 mg ZnSO4
7H2O, 0.079 mg CuSO4
5H2O, 0.39 mg 10% CO2 Na2MoO4
2H2O, 0.049 mg Co(NO3)2
6H2O 5% CO2 25 1C CO2 2% CO2 pH Yield 6.3 2.31 g/L biomass on day 25 with 12.71% (w/w) lipid content 6.5 1.61 g/L on day 24 with 12.44% lipid content 6.8 1.69 g/L on day 26 with 11.21% lipid content 7.0– 2.18 g/L on day 28 with 10.41% 7.5 lipid content Reference [32] [33,34] Schizochytrium limacinum SR21 (ATCC MYA-1381) (50–77%) – 25 1C Artificial seawater containing (per liter): 18 g NaCl, 2.6 g MgSO4
7H2O, 0.6 g KCl, 1.0 g NaNO3, 0.3 g CaCl2
2H2O, 0.05 g KH2PO4, 1.0 g Trizma base, 0.027 g/L NH4Cl, 1.35  10–4 g vitamin B12, 3 mL chelated iron solution, and 10 mL PII metal solution (boron, cobalt, manganese, zinc, and molybdenum) plus 10 g/L glucose, 1 g/L yeast extract, and 1 g/L peptone Nannochloropis sp. (31–68%) 300 26–27 1C (per liter): 1.45 g KNO3, 0.12 g KH2PO4, 0.04 g NaHCO3, 1 mL Fe-EDTA solution (240 mg 1.5% CO2 at aeration 7–8 Optimum cell density (for a 10-cm FeCl3
4H2O in 100 mL 0.05 M Na2-EDTA), 1 mL trace element stock solution (0.01 g CuSO4, rate of 0.31 min– light path reactor outdoors) is 1 –1 0.022 g ZnSO4, 0.01 g CoCl2
6H2O, 0.18 g MnCl2
H2O, and 0.006 g Na2MoO4
2H2O, in 1 L 5.5  108 cells/mL L distilled water) [35] 20 1C Batch-fed (based on glucose feedback) continuous supplemented LDMb medium (per liter): – 1 g tryptone, 892 mL artificial seawater, 100 mL Bristol solution, 6 mL PIV metal solution, 1 mL of 25.0  10–5 g/L biotin, 1 mL of 15.0  10–5 g/L vitamin B12, supplemented with 5 g/L glucose and 30 mg/L Na2SiO3
9H2O 8.2 3.14% dry wt EPAc content 695.2 mg/L EPA yield 49.7 mg/L/d EPA productivity [36] 6.0 Biphasic growth rate: 0.69 g/d for days 1–4, 0.12 g/d after day 4 7.0  106 cells/mL/d maximum 58% dry wt lipid content 28 kJ/g caloric value 24 mg/L/d biomass productivity [37] Nitzschia laevis UTEX 2047 (69.1%) – 7.5– 37.90 g/L maximum biomass yield 8.0 3.25 g/L/d maximum biomass productivity 1 73 mg/g dry wt DHAa content 6.56 g/L DHA yield Chlorella vulgaris (56%) 76 25 1C Low nitrogen medium: 203 mg/L (NH4)2HPO4, 2.236 g/L KCl, 2.465 g/L MgSO4, 1.361 g/L KH2PO4 and 10 mg/L FeSO4 Chlorella emersonii (63%) 76 25 1C Low nitrogen medium: 203 mg/L (NH4)2HPO4, 2.236 g/L KCl, 2.465 g/L MgSO4, 1.361 g/L KH2PO4, and 10 mg/L FeSO4 6.0 0.38 g/d growth rate 4.0  106 cells/mL/d maximum 34% dry wt lipid content 24 kJ/g caloric value 36 mg/L/d biomass productivity [31] Chlorella minutissima UTEX2341 (57%) 50 25 1C Basic Medium (BM ¼ N8Y medium, per liter distilled water): 1 g KNO3, 0.74 g KH2PO4, 0.207 g Atmospheric CO2 Na2HPO4, 0.013 g CaCl2
2H2O, 0.01 g FeNaEDTA, 0.025 g MgSO4, 0.1 g yeast extract, and 1 mL source was not micronutrient solution (3.58 g Al2(SO4)3
18H2O, 12.98 g MnCl2
4H2O, 1.83 g CuSO4 5H2O, used 3.2 g ZnSO4
7H2O per liter of distilled water). Added carbon sources (per liter): 1 g dextrose, 1.46 g oxalic acid, 0.88 g starch, 0.93 g sucrose, 1.22 g glycine, 1.33 g sodium acetate, and 1 g glycerin with 0.39 g carbon/L – 1.78 g/L/d biomass 16.11% lipid content 286.76 mg/L/d lipid productivity 180.68 mg/L/d FAMEd productivity [38] 28 1C Grown on crude glycerol. Basal culture medium (per liter): 0.7 g KH2PO4, 0.3 g K2HPO4, 0.3 g MgSO4
7H2O, 25 mg CaCl2
H2O, 25 mg NaCl, 3 mg FeSO4
7H2O, 0.01 mg vitamin B1, and – 1 mL A5e solution 6.8 3.9 g/L/d biomass productivity 0.62 g/g CDWf lipid content 2.4 g/L/d lipid productivity [39] – Chlorella protothecoides UTEX 256 (23/55%) 270 25 1C – 7.5 [40] A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 Algae strain (% lipid content) DHA, docosahexaenoic acid. LDM, Lewin's marine diatom medium. c EPA, eicosapentaenoic acid. d FAME, fatty acid methyl ester. e A5 solution: 286 mg H3BO3, 250 mg MnSO4
7H2O, 22.2 mg ZnSO4
7H2O, 7.9 mg CuSO4
5H2O, 2.1 mg Na2MoO4
2H2O and100 mL dH2O. f CDW, cell dry weight. b a 36.32 mg/L/d lipid productivity 523.19 mg/L volumetric yield Bold Basal Medium (BBM) (per liter): 1.5 g NaNO3, 0.075 g K2HPO4, 0.175 g KH2PO4, 0.075 g 1% CO2 (v/v) at rate – MgSO4
7H2O, 0.084 g CaCl2
2H2O, 0.00498 g FeSO2
7H2O, 0.05 g EDTA
2Na salt, 0.025 g of 1 L/min NaCl, 0.031 g KOH, 11.42 μg H3BO3, 1.44 μg MnCl2
4H2O, 8.82 μg ZnSO4
7H2O, 1.57 μg CuSO4
5H2O, 0.049 μg Co(NO3)2
6H2O, and 0.71 μg MoO3 Parietochloris incisa (62%) Neochloris oleoabundans strain 1185 (35–65%) – 26 1C 16.5 g/m2/d biomass area productivity 23% dry wt total lipid content 3.8 g/m2/d total lipid productivity Medium (per liter): 0.91 mM MgSO4
7H2O, 8.82 mM NaNO3, 0.43 mM KH2PO4, 1.29 mM Gas injection(CO2 K2HPO4, 0.43 mM NaCl, 0.17 mM CaCl2
2H2O, 30.7 mM ZnSO4
7H2O, 7.3 mM MnCl2
4H2O, and air), flow rate 4.9 mM MoO3, 6.3 mM CuSO4
5H2O, 1.7 mM CoNO3
6H2O, 0.185 mM H3BO3, 0.171 mM of 0.5 L/min EDTA, 0.553 mM KOH, 18 mM FeSO4
7H2O, 10.2 mM H2SO4 [41] A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 131 they are important potential sources of biodiesel and can be used to mitigate the industrial production of CO2. Under normal growing conditions, microalgae yield a caloric value of 18–21 kJ/g/d. However, the optimal CO2 concentrations, nutrient sources, and biofuel yields must be investigated for each strain of microalga [29]. For example, lipid production in Chlorella emersonii could be as high as 58–63% dry weight when grown with low nitrogen levels, and C. protothecoides contained 55% lipid (dry weight) when grown heterotrophically with corn powder hydrolysate under nitrogen limitation [30]. Furthermore, the high production costs of microalgal biofuels can be mitigated by selecting the best strains and by developing process technologies that are economically viable at industrial scales [31]. In this section, we discuss some of the most economically-promising microalgal strains, their cultivation conditions, and their yields. All example strains are summarized in Table 1. Botryococcus braunii 765 (Plantae: Chlorophyta) is a green colonial microalga that can yield biodiesel, hydrocarbons, and biocrude oil. Typically, these algal cells are incubated in laboratory bioreactors at 25 1C under 150710 mmol/m2/s light for 2 weeks before being transferred to an enclosed photobioreactor (see Section 3) containing sterile modified BG11 medium. The effects of CO2 concentrations (from 2% to 20%) on the growth rate of B. braunii were studied. Maximum biomass production occurred with 20% CO2, giving a biomass yield of 2.31 g/L that was 12.71% (w/w) lipids on day 25 [32]. This shows that hydrocarbon content increases as the concentration of CO2 increases and by adding 2% sodium hypochlorite in photobioreactors. Strains of Schizochytrium (Chromalveolata: Heterokontophyta) have excellent production rates on the order of 7.3–9.4/day. Biomass densities of some strains reach 200 g/L within fermentation cycles of 90–100 h under nitrogen- and glucose-fed cultures [33]. Schizochytrium can also ferment glycerol to produce DHA. Crude glycerol is a major byproduct of biodiesel production. DHA yield significantly increases to 4.91 g/L by adding 1 g/L ammonium acetate at 19.2 1C. However, glycerol concentrations above 25– 35 g/L decrease cell growth. At 35 g/L glycerol, cellular lipid content reached 73.3% [34]. Algal growth and productivity are highly affected by the light path (LP), which can range from 1.3 to 17 cm in vertical glass photobioreactors. Shorter LPs (e.g., 10 cm) resulted in higher mass productivity of Nannochloropsis sp. (Chromista: Ochrophyta) [35]. The two main factors that affect reactor efficiency are the total illuminated surface area and culture volume. The lower these values, the more efficient and cost effective the reactor. Open raceway ponds are more productive per unit illuminated area than flat-plate reactors, but they require six times more volume than closed photobioreactors to produce the same amount of algae. To increase the productivity of the diatom Nitzschia laevis (Chromista: Ochrophyta), a continuous fed-batch process was developed [36]. Glucose, tryptone, nitrates, and yeast extract were considered essential media components to increase productivity. The optimum ratio of 31:1 glucose: nitrate increased cell growth yield to 22.1 g/L; these conditions also increased eicosapentaenoic acid (EPA) production. For fuel use, microalgae must have high caloric value and biomass productivity. The microalgae C. vulgaris and C. emersonii (Plantae: Chlorophyta) were cultivated in a pumped tubular photobioreactor to compare low-nitrogen and Watanabe's media [37]. Using the low-nitrogen medium increased both lipid content and caloric value for both species, but the biomass productivity of C. vulgaris was higher in Watanabe's medium (40 mg dry wt/L/d) than in the low-nitrogen medium (24 mg dry wt/L/d). C. minutissima UTEX2341 is a promising source of biodiesel, because it contains C16 and C18 lipids, which are components of diesel oil. When cultivated in a flask receiving 50 mmol m2/s light 132 A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 in organic carbon medium, its biomass productivity was 1.78 g/L/d. To increase its biomass productivity, glycerin can be used as an organic carbon source and casein as a nitrogen source [38]. Crude glycerol has been considered as an important alternative carbon source to glucose for the cultivation of microalgae. It is a byproduct during biodiesel production, so using it as a carbon source makes the cultivation process more cost-effective and eliminates its disposal cost. Strains of C. protothecoides cultivated in continuous fed-batch mode with glycerol as the carbon source had higher maximum lipid productivity (3.9 g/L) than in batch mode. Another advantage of fed-batch cultivation is that it can use glycerol with 62% purity with similar productivity to the use of pure glycerin in batch mode [39]. Neochloris oleoabundans (strain 1185) (Plantae: Chlorophyta), a freshwater microalga, is known for its ability to store lipids and triacylglycerides (TAG). However, the choice of nutritional medium did not affect TAG productivity [40]. Parietochloris incise (Plantae: Chlorophyta) is a potential source of polysaturated fatty acids (PAC) and arachidonic acid (AA). It was cultivated in a vertical tubular photobioreactor to optimize its nutrient proportions and productivity to improve its cost effectiveness. Nitrogen starvation decreased AA productivity; the critical concentration was 0.5 g–1 sodium nitrate, giving 36.32 mg/L/d lipid productivity and 523.19 mg/L volumetric yield. In general, phosphorus and nitrogen starvation seem to enhance AA productivity more than nitrogen starvation alone [41]. A primary strategy for most algal biofuel producers is to identify the algal species that have a high oil content, that will also grow quickly to produce biodiesel, biocrude and drop-in fuels. Algae with high oil content such as B. braunii (Bb) grow slowly and can be harvested only a few times a week, whereas algae with lower oil content such as Dunaliella or Nannochloropsis (in the 20– 40% range) will grow more quickly and can be harvested daily or a few times a day. For this reason, most algal R&D projects and precommercial projects are using algal strains with 20–40% content [42]. 3. Algae cultivation Microalgae can be cultivated in different types of systems, mainly in open ponds or raceways and in enclosed photobioreactors. Open cultures are usually located outdoors and rely on direct sunlight, while closed photobioreactors can be either indoors or, preferably, outdoors to use free sunlight [43]. Most species of Fig. 1. Schematics of a raceway pond. microalgae can be grown in photobioreactors, while open systems are more limited. 3.1. Cultivation in ponds and raceways Open ponds are simple cultivation tanks that are largely obsolete, having been replaced by more efficient raceway ponds. Raceway ponds as shown in Fig. 1 form closed circuits approximately 0.25 m wide and 0.4 m deep through which the water is circulated using a paddle wheel [44]. They are shallow to maximize light penetration. Where land and water are inexpensive, raceways are extremely cost effective to construct and operate. They require no cooling and do not experience oversaturation with oxygen, which can threaten biofuel production in closed systems. However, they have lower productivity per unit area and volume than closed photobioreactor systems because of the low light-tovolume ratio [45]. Additionally, these systems can be easily contaminated by other microorganisms that can compete with the cultivated algal strain [46,47]. They must be kept highly alkaline to prevent contamination; this high pH limits their suitability to only a few species. The possibility of contamination is often cited as a serious limitation of open systems [5] and it is true that most of the species cultured in such systems currently do grow in selective environments, i.e. Arthrospira (Spirulina) [high alkalinity], Dunaliella salina [high salinity], and Chlorella [high nutrients] [48,49]. However, other species with ‘normal' growth requirements have also been grown successfully in open ponds [5], either in batch mode [e.g., Haematococcus pluvialis] [50], or continuously for very long periods [e.g., Phaeodactylum tricornutum, Nannochloropsis and Pleurochrysis carterae] without significant contamination problems [5]. Odlare [51] studied the cultivation of algae in open pond (Lake Mälaren) in Sweden. The idea behind this research was to enhance indigenous algae production rather than inoculate new species into the system. The production rate of biodiesel from algae was estimated using data from Weyer [52], Amin [53] and Chisti [54] for the estimation of potential for using algae as energy source in the Mälardalen region. The biomass productivity was enhanced by nutrient addition by using Jaworskis Medium (JM). The estimate of the potential of algae to replace vehicle fuels in the Mälardalen region shows that an area corresponding to at least 40% of the Mälaren would be required to satisfy current demand [51]. 3.2. Cultivation in photobioreactors Closed photobioreactors are designed to overcome the limitations of open pond systems [55]. They have higher efficiency and biomass productivity, shorter harvest times, high surface-to-volume ratios, reduced contamination risks, and can be used to cultivate a greater range of algal species than open systems [29,54]. Additionally, they can use wastewater or flue gases from power plants, providing additional environmental benefits [56]. However, they are much more expensive to construct than open systems. Closed photobioreactors involve a thin panel of transparent tubes or plates placed horizontally or vertically and provided with CO2 cylinders. There are several types, including tubular, flat plate, column, and biofilm photobioreactors. They can be airlift, flat inclined, bubble column, column aeration, solar penthouse-roof, and multistage continuous flow photobioreactors. Table 2 summarizes some important process variables and specifications for photobioreactor designs used for different microalgae species. Tubular photobioreactors as shown in Fig. 2 are the most widely used and considered the most promising, because they yield high biomass and have short harvest times [41]. They comprise parallel tubes (0.2 m in diameter or less) that are positioned horizontally or vertically to maximize sun exposure. Table 2 Cultivation of microalgae species in closed photobioreactor systems. Biomass concentration during production Reference 25 1C In cabinet (25 1C) 2.31 g/L max [32] 25 1C 257 2 1C 0.21 g/L/d [63] Possible 30 1C _ 0.89–0.28 g/L/d [68] Cultivates algae as a biofilm with reduced energy and water requirement 25 1C Transparent film capable of blocking infrared radiation 0.71 g/m2/day [64] 4.25–6.50 kW/ m2/day _ 35 1C _ 0.770 g/L/d [61] 8.2 430 730  10–6 mol/m2/s Difficult _ 22.0 7 0.1 1C _ [70] SA¼ 0.1096 m2 _ 120 μE/m2/s1 _ 25 1C Transparent jacket with thermostat unit (25 1C) _ [65] SA¼ 0.084 m2, capacity ¼ 1.5 L, 3.0 L 6.8 7 0.1 980 780  10–6 E/m2/s Possible 30 1C Cooling jacket located at front of reactor (29 1C) 0.027–0.045 g/ L/h [71] Polycarbonate: 0.01 m thick; Capacity ¼30 L plates 0.61  0.61  0.1 m3 7.6 Optical density was measured at 806 nm Commercial 22.5 1C Acrylic water basin (40 L, 3.42 7 0.39 g/d ≈10 cm high) with type K thermocouples (26–35 1C, accurate to 7 0.3 1C) [72] Spirulina platensis θ ¼301 (summer), 601 Glass tank: 90 cm length, 2.6 cm internal width, 70 cm (winter) high 9.5 10% Of incident irradiance _ 35 1C Water sprinklers along top of front panel _ [69] Flat plate Synechocystis aquatilis SA¼ 3.5 m2 Three plates (1000 m, 1000 m, and 360 m) separated by 0.5 m in a glass green house _ 1–12 MJ/m2/d _ _ _ 30 g/m2/d [67] Multistage continuous flow Scenedesmus sp. Standard window glass: 0.25 Capacity ¼900 L in. thick, six flat plate 2.1 m long, 0.5 m high, 0.15 m deep _ 750 nm light Commercial 35 1C Evaporative cooling (simple, economical) (25–35 1C) _ [73] Stirred tank Spirulina maxima Acrylic glass _ 10.3–11.4 _ _ 30 1C Roux bottles (31 1C) 10.8 g/m2/d [74] Tubular Haematococcus pluvialis Parallel plastic tubes, diam. ¼0.18–0.41 m SA¼ 100 m2; capacity ¼ 25,000 L _ _ Difficult 20 1C PBR flooded with cold sea 13 g/m2/d water (5–25 1C) [75] Tubular Phaeodactylum tricornutum Acrylic glass tubes; 4-m tall airlift section with degasser zone; riser/downcomers 0.053 m diam., 0.007 m thick Tube length ¼ 80 m; surface area (SA) ¼ 12/m2; degasser zone length ¼0.22 m; θ ¼601; reactor volume ¼ 0.2 m3. 7.7 Saturation Difficult irradiance ¼ 185  10–6 E/m2/s 35 1C PBR immersed in pond water (207 2 1C) [59] Specifications Size and capacity pH Solar irradiance Airlift Botryococcus braunii Chlorella sp. Tubes: 10 cm diam., 50 cm long Acrylic glass tubes Capacity ¼3 L 6.0–8.0 SA¼ 1.337 m2 _ 120  10–6 mol/ Difficult m2 /s 350 μmol/m2/s at Commercial surface of reactor from 16 cool white 40 W lamps Airlift Chlorella vulgaris Acrylic glass columns: 10 cm Capacity ¼2 L diam., 25 cm high Biofilm photobioreactor Botryococcus braunii A biofilm growth surface (8 mm thick concrete layer) SA¼ 0.275 m2 Capacity ¼0.60 L 8.3 Bubble column Aphanothece microscopica Glass tube: 7.5 cm diam., 4 mm thick, 75 cm high Capacity ¼3 L; 1.5-cm diam. air diffuser in center of column _ Bubble column Isochrysis aff. Galbana _ Capacity ¼5 L Bubble column Porphyridium purpureum Glass bubble-column tubes, 0.4 m high, 0.1 m diam. Flat plate Chlorella vulgaris Acrylic glass Flat plate Dunaliella tertiolecta Flat plate Airlift 8.0–10.0 _ Standardization 1.5 g/L/d. 133 Temperature control Algae taxon A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 Limiting temp. Type of photobioreactor (PBR) 134 [60] Shading with dark plastic 0.01 g/L/g (air)/m2 sheets; overlapping of tubes; spraying with water 36 1C Commercial 9.4–9.8 Spirulina sp. Tubular Acrylic glass tubes, 0.13 m diam., 0.004 m thick SA¼100 m2; capacity ¼ 100 L /m2 _ [77] 32.5 g/m2/d Two double-jacket heat exchangers in lower degasser and counterflow cooler (357 1 1C) 35 1C Lab Scale 7  10 6 mol photon/m2/s 9.4 SA¼1.2 m2, capacity ¼ 65 L Spirulina platensis Tubular Glass tubes: 24 m long, 48 mm i.d., in six parallel horizontal rows connected by glass U-bends [43] 35 g/m2/d _ _ Commercial _ _ Porphyridium cruentum Tubular Tubes 0.064 m diam., 1500 m Capacity ¼6 m3 total length [76] _ _ _ Commercial _ 7.7 _ Phaeodactylum tricornutum Tubular Tubes: 0.03–0.06 m diam. [47] 1.5 g/L/d Heat exchanger (28 1C) 35 1C Saturation Difficult irradiance ¼ 200  10–6 E/m2/ s; average irradiance ¼ 250  10–6 E/m2/s 7.7 Capacity ¼75 L; height of degasser above helical loop¼ 2 m Plastic tubes: 106 m length, 0.03 m diam.; arranged around a circular frame, 1.2 m diam., 0.8 m high Phaeodactylum tricornutum Tubular Solar irradiance pH Size and capacity Specifications Algae taxon Type of photobioreactor (PBR) Table 2 (continued ) Standardization Limiting temp. Temperature control Biomass concentration during production Reference A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 The growth medium is circulated from a reservoir through the reactor using an airlift device without moving parts, reducing contamination and preventing the cell damage caused by mechanical pumping. Airlift devices also remove excess oxygen, which would otherwise inhibit photosynthesis [54,57,58] Culture efficiency is highly dependent on optimizing flow and gas exchange, and photobioreactor geometry should also maximize the illumination area. Volumetric productivity decreases as tube diameter increases, while areal productivity increases with volume. For maximum production in a given area, the suggested tube diameter is 0.06 m. The annual areal productivity of P. tricornutum using 0.06-m diameter tubes was 35 g/m2/d and 1.5 g/L/d [59]. Tubular photobioreactor can also be designed helically to provide a larger surface area to volume ratio. This design maximizes light penetration, limits contamination, allows for easy temperature control, and provides maximum CO2 transfer in the culture medium. P. tricornutum grown in a helical photobioreactor had 1.5 g/L/d biomass productivity at 30 1C [47]. Spirulina was cultured in a flat inclined photobioreactor made of flat glass tank with the temperature maintained below 35 1C. The inclined face was set at angles of 301 in the summer and 601 in the winter in the northern hemisphere [60]. The cyanobacterium Aphanothece microscopica was cultured in a bubble column photobioreactor, and the effects of photoperiod on biomass productivity and CO2 fixation rates were studied at 35 1C. The duration of light directly impacted biomass productivity, and CO2 fixation rates reduce by 99.69% when deprived of light [61]. Airlift photobioreactors are simple and cost-effective reactors for the mass culture of various types of algae. They are made of acrylic glass, which is inexpensive and easily obtained, and meet the desired criteria for new generation photobioreactors of high light penetration and biomass production, ease of maintenance, and minimal contamination [62]. They have three major parts, a draft tube, an outer tube, and an air duct. They are well suited for cultivating Chlorella sp. Volumetric productivity obtained at a superficial velocity of 4 mm/s was 0.21 g/L/d [63]. Ozkan [64] studied the use of performance of an algae biofilm photobioreactor that offers a significant reduction of the energy and water requirements of cultivation. The green alga B. braunii was cultivated as a biofilm. The system achieved a direct biomass harvest concentration of 96.4 kg/m3 with a total lipid content 26.8% by dry weight and a productivity of 0.71 g/m2 day, representing a light to biomass energy conversion efficiency of 2.02%. Moreover, it reduced the volume of water required to cultivate a kilogram of algal biomass by 45% and reduced the dewatering energy requirement by 99.7% compared to open ponds. Porphyridium purpureum was cultivated in a bubble column photobioreactor with a diameter of 0.4 m and height of 0.1 m to maximize volumetric productivity. Air was supplied with flow rate of 2.5 L gas per L liquid per hour along with 2% CO2 from below to provide agitation and fixation [65]. Flat plate photobioreactors as shown in Fig. 3 are very effective for biomass cultivation of microalgae. They provide a high surface area to volume ratio for illumination and have easy design features [66]. Biomass productivity of microalgal cultures rapidly increases with mixing rate, which provides an adequate supply of CO2 to the culture while removing excess oxygen and increasing the flashing effect. However, because of higher running costs, high aeration rates are not recommended for large-scale production. Synechocystis (Cyanobacteria) was cultivated in vertical flat plate reactors and aeration along with 5–10% CO2 introduced from the bottom of the column to increase biomass productivity [67]. The effects of CO2 aeration rates from 2% to 20% on the biomass productivity of B. braunii were investigated in a closed airlift photobioreactor. All strains could grow at all CO2 concentrations A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 with no obvious inhibition at an aeration rate of 0.02 volume gas per volume liquid per minute. Maximum biomass productivity was 2.31 g/L on the 25th day at a 20% CO2 aeration rate. Moreover, cultivation and harvest could be made more economical by using sodium hypochlorite to sterilize the reactor, by not adjusting the pH, and by using flocculation to harvest the algae [32]. Studies have been done on the production of lipids using C. vulgaris and wastewater in a batch or semi-continuous manner in a column aeration photobioreactor. This is a promising means for lipid production. Maximum lipid production was 42% with biomass productivity of 147 mg/L/d using a semi-continuous process with daily replacement rate of one of the two cultures, making it competitive with petroleum at US $63.97/barrel [68]. A penthouse-roof photobioreactor was used to cultivate Spirulina. These reactors comprise both indoor and outdoor units and Fig. 2. Working of a horizontal tubular photobioreactor. 135 are most efficient in temperate climate zones. They use collectors to concentrate light and all cultivation parameters (temperature, flow rate and oxygen concentration) can be easily controlled. The reactor in this study had a tilt angle of 401 to provide maximum light. Spirulina maximum biomass concentration was 1.2–2.2 g/L and productivity was 0.5 g/L/d in September, with a biomass yield of 32.5 g/m2/d [69]. In Arizona, USA, groundwater provides about 40% of the drinking water, so it is a precious resource. Ten percent of Arizona groundwater exceeds the maximum allowable concentration of nitrates (10 mg/L), making it unfit for drinking. Growing algal cultures using such water could make cultivation more economical and also purify water. To test this potential, Scenedesmus (Planta: Chlorophyta) was grown in an outdoor multistage photobioreactor consisting of six flat glass plates arranged in ascending levels (total capacity, 900 L) [73]. The culture was provided with air along with 0.05–1% CO2 through submerged tubing, which also mixed it. The reactor was designed to overflow from one plate to another under gravity, making it very effective at maximizing biomass productivity and also removing nitrates. C. vulgaris was studied in flat plate airlift (FPA) and bubble column photobioreactors provided with 3 cm LPs, 980 E/m2/s of light, and caloric values of 25 kJ/g [71]. The volumetric productivity in the FPA (0.045 g/L/h) was 1.7-fold higher than in the bubble column photobioreactor (0.027 g/L/h), and the photosynthetic efficiency was also higher (4.7% vs. 2.9%). Thus, FPA was considered to be more effective. The advantages and disadvantages of different types of photobioreactors used to cultivate microalgae are listed in Table 3. Fig. 3. Front and side view of the flat panel photobioreactor. 136 A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 Table 3 Advantages and disadvantages of different photobioreactors used to cultivate microalgae. Type of photobioreactor Advantages Disadvantages Reference Tubular Very effective light use; excellent temperature control; reasonable scale-up Very high productivities and cell densities Absorbs more oblique light from light source Fouling with some growth along walls [78] Scale-up requires many compartments and support materials Difficulty controlling temperature; possible hydrodynamic stress to some algal strains High heating and illumination costs Complexity; difficult to scale up High production cost [79] [80] May be dark regions away from the center, depending on reactor depth [83] Vertical Flat panel tubular with fresnel coating Helical tubular Air lift tubular Multiple airlifting membrane Cuboidal Stirred draught tube Stirred tank Air lift Flat plate Bubble column tubular Modified cascade High surface area Capable of large scale production; easy CO2 supply Control over overall gas holdup and liquid circulation; production of metabolites High cell concentration; effective total light incidence Combines stirred tank and plate reactors; constant light conditions Largely uniform mixing; excellent temperature control Good light use; high temperature control; high mass transfer coefficient Excellent light use and temperature control; high gas transfer coefficient Scalable; homogeneous culture environment; low cooling requirement; effective light use Effective light use; high mass transfer coefficient; economical [81] [62] [82] [84] Higher cultivation cost Difficult to scale up; hydrodynamic stress on algae Low hydrodynamic stress on algae; difficult to scale up [78] [85] Difficult to scale up [67] Low surface to volume ratio [85] Increased shear stress by pumps may limit biomass productivity [86] Table 4 Methods used to extract lipids and oils from microalgae. Product Extraction method Species Solvent Efficiency/yield (wt%) Time (min) T (oC) P (MPa) Reference Lipids Lipids Lipids Lipids Lipids Lipids Lipids Lipids Lipids Lipids Lipids Lipids Lipids Lipids Lipids Lipids Lipids Oils Oils Oils Oils Oils Oils Organic solvent Organic solvent Organic solvent Organic solvent Cold pressing Bead beater+solvent Bead beater+solvent Soxhlet Soxhlet Soxhlet Soxhlet Supercritical fluid Supercritical fluid Subcritical ethanol Supercritical fluid Supercritical fluid Supercritical fluid Organic solvent Bead beater+solvent Soxhlet Soxhlet Supercritical fluid Supercritical fluid Chaetoceros muelleri Chlorococcum sp. Chlorococcum sp. Phaeodactylum tricornutum Scenedesmus obliquus Botryococcus braunii Botryococcus sp. Scenedesmus obliquus Botryococcus braunii Chlorella vulgaris Chlorococcum sp. Crypthecodinium cohnii Chlorococcum sp. Nannochloropsis sp. Nannochloropsis sp. Spirulina maxima Spirulina platensis Oedogonium sp. Chlorella vulgaris Isochrysis galbana Isochrysis galbana Spirulina platensis Tetraselmis chui 1-Butanol Isopropanol/hexane Hexane system Ethanol, 5 mL/g dried microalgae Ethanol Chloroform/methanol Chloroform/methanol Hexane DBUa/octanol Hexane Hexane system CO2, 10 g/min CO2 Ethanol CO2 CO2 CO2 Hexane+ether CO2 CO2, 2.0 mL/min CO2/ethanol CO2 DCMb/MeOH in a ratio of 9:1 94 6.8 1.5 29 62.047 2.42 28.6 28.1 40.717 4.46 81 1. 77 3.2 9 5.8 90.21 25 3.1 8.6 9.20 13.30 4–10 5–11 90 15 60 450 450 1440 – 50 – – 240 140 330 180 80 – 70 25 25 – 73–75 – – 63–65 60 70 – 49.85 60 – 40 35 40 80 – 40 50 55 100 – – – – – – – – – – – 30 10–50 – 55 60 40 – – 69 6.89 70 10.34 [113] [90] [90] [114] [106] [96] [96] [106] [115] [102] [90] [116] [90] [117] [118] [119] [120] [112] [112] [121] [121] [112] [112] a b 60 20 – – – 15 – DBU, 1,8-diazabicyclo-[5.4.0]-undec-7-ene. DCM, Dichloromethane. 4. Extraction To extract lipids from microalgae, the algae must first be harvested from photobioreactors or open pond cultures and concentrated by filtration, centrifugation, flocculation, or agglomeration to remove the water [87]. Dewatered algae is then dried, milled into a fine powder, and pretreated by bead milling, microwaving, chemical lysis, or high-pressure homogenization to increase the mass transfer of lipids during extraction. Pretreatment greatly improves the extraction efficiency by disrupting the cellular structure, releasing lipids into the solvent mixture, and enhancing overall yield. Oil expellers and presses along with hexane solvent extraction can increase lipid yield up to 75 wt% [88,89]. Lipids can be extracted from the dried algal biomass using different chemical and physical means. Chemical extraction uses organic solvents like hexane or methanol. Other techniques include expeller presses, electromagnetic methods, direct liquefaction, Soxhlet extraction, supercritical fluids (CO2), ultrasonic waves, and microwave-assisted organic solvent extraction [90]. Several methods are summarized below and in Table 4. After lipid extraction, the remaining constituents (solvent, water, cell debris, and unextracted lipids) are sent to a solid–liquid separation system to remove cell debris [91]. In organic solvent extraction, water and solvent are removed using liquid–liquid separation methods, such as evaporation, vacuum distillation, or solvent adsorption. In supercritical fluid extraction, the mixture is A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 137 pressure decomposed, converting the solvent and residual water into gases and precipitating the lipids. Extracted lipids are then transesterified into biodiesel [90]. Technology for the production of biodiesel from microalgae must have high specificity for lipids to minimize contaminants such as carbohydrates and proteins [92]. Purification technology should also favor the production of acylglycerols over other lipids, such as ketones, chlorophylls, sterols, polar lipids, and carotenes, that are not readily converted to biodiesel [93]. Additionally, the technology should have low operating and capital costs, require little energy and time, be safe, and should not react with lipids [89]. was obtained from C. vulgaris using ultrasonic-assisted solvent extraction in 2.33 h, while Soxhlet extraction yielded 1.58 wt% oil after 18 h [102]. Thus, a higher yield was obtained in less time using ultrasonic waves versus Soxhlet extraction. To avoid filtration and other chemical extraction techniques, microalgal suspensions can also be placed in an electromagnetic field and the pH varied using CO2; this process destroys the cell wall and allows the oils to float to the surface. Lipids can be extracted from live algal cultures using mesoporous nanoparticles [103–105]. 4.1. Organic solvent extraction Microwaves combined with organic solvent can also be used to extract lipids from microalgae. The process uses electromagnetic radiation within a specific frequency range to heat the cells and increase the internal pressure. The cells rupture, forcing their constituents out. The rapid explosion rapidly diffuses the lipids into the organic solvent. Microwave-assisted hexane extraction yielded more lipids than other conventional heating methods and was more rapid [106]. The process is economical and environmentally friendly. The microalga Scenedesmus obliquus was heated to 80–95 1C using microwaves (1.2 kW, 2450 MHz frequency) for 20–30 min. Maximum oil yield obtained was 76–77 wt% at 95 1C with a holding time of 30 min, whereas solvent extraction yielded 52 wt% oil [106,107]. Lipid–solvent systems are governed by the principle that like dissolves like, so lipids are extracted using non-polar organic solvents like chloroform or hexane [94]. The extraction can be divided into five steps. (1) Microalgae are exposed to the solvents, which penetrate the cell membrane and enter the cytoplasm. (2) The solvents interact with neutral lipids via Van der Waal's forces to form a solvent–lipid complex. (3) This complex diffuses across the cell membrane, such that neutral lipids enter the organic phase while water and solvent–contaminant complexes (with carbohydrates or proteins) enter the aqueous phase. (4) The organic phase is then separated, and (5) crude lipids are transesterified to produce biodiesel [7,88,95]. To increase lipid yield, two or more organic solvents can be used simultaneously in polar/non-polar combinations, e.g., methanol/ chloroform or hexane/isopropanol. One study found that using isopropanol as a co-solvent increased lipid yield from Chlorococcum sp. up to 300% more than using pure hexane; yields were 0.068 and 0.015 g lipid/g algal biomass, respectively [90]. In another study, beadbeaten B. braunii was exposed to five different organic solvents; chloroform/methanol yielded the highest lipid content (0.29 g/g algal biomass) [96]. 4.2. Soxhlet solvent extraction Organic solvent extraction is usually carried out in a noncontinuous batch process, which limits lipid mass transfer equilibrium [97]. To solve this problem, a continuous solvent extraction process is required, which requires large amounts of solvent [98]. Soxhlet solvent extraction continuously evaporates and condenses the solvent, avoiding the lipid mass transfer limitation and reducing solvent consumption [99]. The Soxhlet apparatus comprises a round-bottom flask holding solvent, a Soxhlet extractor containing algal biomass, and a condenser. Heated solvent enters the condenser, which channels it into the extractor. A thimble filter can be used to prevent algal biomass from being carried out of the extractor with the solvent [100]. When the organic solvent reaches its maximum volume in the extractor, it is siphoned into the flask, where it is again heated and evaporated, leaving crude lipids behind [99]. Soxhlet lipid extraction is more effective than batch extraction. Yields from Chlorococcum sp. were 0.015 g lipid/g dried microalgal biomass using Soxhlet extraction and 0.057 g lipid/g biomass using a batch process [90]. However, continuous distillation is energyconsuming. 4.3. Ultrasonic-assisted organic solvent extraction Ultrasonic waves can be used to increase lipid yield as compared with other extraction methods. Ultrasonic methods can also extract other biochemical compounds, such as carotenoids and chlorophyll. In the reactor, ultrasonic waves create solvent bubbles that explode and rupture the cell walls, forcing constituents out of the cells into the solvent mixture [91,101]. An oil yield of 1.77 wt% 4.4. Microwave-assisted organic solvent extraction 4.5. Direct liquefaction Oils can be directly obtained from dried or wet microalgae by liquefaction. Algae have high moisture content (up to 78.4%), requiring a lot of energy to dewater. Liquefaction directly converts the biomass into oil. Maximum oil yield was 25–44.8 wt% at 300–360 1C and 10 MPa. In experiments with high-moisture B. braunii treated with or without a catalyst (5% Na2CO3) at 300 1C, more than 95% of hydrocarbons were recovered [96]. The origin oil company introduced a new method that can extract oil from microalgae at a rate of 5 gallons/min with an efficiency of 94–97 wt% [53] without requiring dewatering, making the process economical for commercial use. The method combines an electromagnetic field to rupture the cells, pH adjustments, and tank settling and gravity clarification [108]. 4.6. Supercritical (CO2) fluid extraction Supercritical CO2 (SC-CO2) extraction is a green technology that promises to replace organic solvent extraction. When the temperature and pressure of a fluid increase above its critical point, the fluid behaves as both a liquid and a gas. This method is very efficient for extracting lipids for several reasons. (1) The crude lipid products are solvent free. (2) The solvent rapidly penetrates the algal cells, giving a higher lipid yield. (3) Solvent power is a function of fluid density, which can be tuned by adjusting the temperature and pressure to get neutral lipids (acylglycerols). (4) Supercritical fluids are non-corrosive, non-toxic, non-flammable, and inert. (5) No degumming is required because SC-CO2 does not solubilize polar phospholipids [109–111]. Supercritical organic solvent extractions use a decompressed CO2 flow rate of 400 mL/min, with a holding time of 4.9–14.1 min depending upon fluid density. The temperature is varied from 60 to 80 1C and the pressure from 10 to 50 MPa for 80–120 min. Under optimum conditions, i.e., 30 MPa and 49.85 1C, almost 50 wt% of the oil is extracted from the dried algal biomass after 2 h [112]. Lipid yields using SC-CO2 increase with decreasing temperature and increasing pressure and are much higher in less time than with ordinary solvent extraction. A yield of 0.058 g lipids/g microalgae was obtained from Chlorococcum sp. using SC-CO2 138 A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 with a residence time of 80 min, while Soxhlet solvent extraction yielded 0.032 g lipids/g microalgae after 5.5 h [90]. In one study, 90% of oils were recovered from Spirulina platensis in less than 15 min at 70 MPa and 55 1C using SC-CO2, while it took 6 h to extract the same amount by Soxhlet hexane extraction [65]. 5. Microalgae biofuel products 5.1. Bio-hydrogen The need for energy, global warming, and pollution combined to make research on alternative energy sources vital. Hydrogen has excellent potential to provide energy while alleviating global warming and pollution concerns [122]. Renewable biohydrogen production is of increasing interest as fossil fuel supplies are being depleted [123]. A number of methods can generate renewable hydrogen, such as biomass gasification, electrolysis, and photovoltaic generation, which all produce hydrogen for less than US $20/GJ, which is quite reasonable [124,125]. Hydrogen has the highest energy content per unit weight (142 kJ/g or 16,000 BTU/lb) of the known fuels, although it requires special handling because little energy is needed for ignition and it leaks easily [125]. Hydrogen gas has excellent potential as a renewable energy source because it produces only water when combusted, unlike the carbon pollution of fossil fuels. The only carbon released by hydrogen gas is derived from CO2fixation and from microbial fermentation [126]. Thus, biohydrogen can be considered carbon-free. Hydrogen is also very good for internal combustion engines and maintains long-term engine efficiency [127,128] Most hydrogen is used in the fertilizer (≈50%) and petroleum (≈30%) industries. Sales have increased by 6% per year in the last 5 years, indicating the increased production in refineries. Hydrogen produced today comes from natural gas (40%), heavy hydrocarbons like naphtha (30%), coal (18%), and electrolysis (4%). Biological hydrogen has become a viable source given the current energy demand and environmental issues. The main objective is now to improve hydrogen yield to make it more economically viable [129]. A number of methods are currently being used to produce biohydrogen, for example, hybrid bioreactors, electrochemical-assisted bioreactors, and metabolic and genetic engineering techniques [127]. The main problem for commercializing biohydrogen production is the low yield and rate of production. Using cheaper raw materials, efficient production techniques, and pilot tests of photofermentation plants should make biohydrogen a commercially viable source of energy in the near future [130]. 5.1.1. Biophotolysis Blue-green and green algae produce biological hydrogen by photolyzing water using solar energy and hydrogenase and/or nitrogenase enzymes. Microalgae use solar energy to transfer electrons to NADPH and ferredoxin, which in turn generates hydrogen. This process was studied by Gaffron and Rubin [131]. To offer an economically-competitive source of hydrogen, these organisms must achieve at least 10% solar conversion efficiency; the green alga Chlamydomonas reinhardtii achieved the maximum theoretical light-conversion efficiency of 22% under controlled laboratory conditions, as summarized in Table 5 [125]. Photolysis can be further classified into several subcategories, as discussed below. 5.1.2. Direct photolysis Direct photolysis involves splitting water into hydrogen and oxygen using sunlight energy, as follows: 2H2 O þ light energy-2H2 þ O2 C. reinhardtii under anaerobic conditions is widely used to produce hydrogen. These hydrogen generated ions are then used for the production of hydrogen in the medium of ferredoxin which are catalyzed by hydrogenase enzyme present in the algal cells. Hydrogen is produced in two steps. First, photosystem II absorbs light and generates electrons that are transferred to ferredoxin using light energy absorbed by the photosystem. Hydrogenase Table 5 Hydrogen production from algae. Species Process Catalyst Yield Time (h) Reference Chlamydomonas reinhardtii Chlamydomonas reinhardtii Biophotolysis Direct biophotolysis Indirect biophotolysis Direct biophotolysis Biophotolysis Direct biophotolysis Biophotolysis Hydrogenase – 40 mol% more with blockage of DCMUa – 96 – [125] [138] Hydrogenase and nitrogenase – – [122] Hydrogenase enzyme – – [122] Hydrogenase and nitrogenase In vitro chloroplast–ferredoxin– hydrogenase system Pigment–protein antennae complexes 0–5 mol% – – – [141] [141] – [135] Bacterial hydrogenases/ferredoxin 2.2 mol% increase with two-compartment flat plate reactor 10 mol% 0.25 [142] Hydrogenase and nitrogenase 20 mol% – [142] Bacterial hydrogenases/ferredoxin Hydrogenase enzyme 10–13 mol% – – – [143] [136] Bacterial hydrogenases/ferredoxin – – [136] Hydrogenase enzyme Fe hydrogenase 15–20 mol% 10–20 mol% more with methylamine hydrochloride 23.6 mL/h 112.7 mL/h – – [144] [137] 182 210 [145] [145] Anabaena variabilis (blue algae) Chlamydomonas reinhardtii Blue-green algae Green algae Rhodopseudomonas sphaeroides Chlamydomonas reinhardtii Chlamydomonas reinhardtii Green algae Chlamydomonas reinhardtii Anabaena variabilis Scenedesmus obliquus Chlamydomonas reinhardtii Chlamydomonas reinhardtii Chlamydomonas reinhardtii a Direct biophotolysis Dark fermentation Biophotolysis Direct biophotolysis Indirect biophotolysis Biophotolysis Direct biophotolysis Biophotolysis Biophotolysis Dilution S-deprivation Nutrient control of S-deprivation DCMU, 3-(3,4-dichlorophenyl)-1,1-dimethylurea. A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 then accepts the electron from ferredoxin to generate hydrogen H2 O-Photo system II-Photo system I-Fd Hydrogenase-H2 ↓ O2 Hydrogenase activity is also found in S. obliquus, Chlorococcum littorale, Platymonas subcordiformis, and C. fusca but not in D. salina or C. vulgaris [122,132,133]. C. reinhardtii in the presence of 2–7 mM methylamine hydrochloride produces 10–20% more hydrogen (cf. Table 5). 5.1.3. Indirect photolysis Some cyanobacteria, such as non-marine Anabaena sp., marine Calothrix sp., Oscillatoria sp., Synechococcus sp., and Gloeobacter sp., can fix nitrogen and produce hydrogen via both hydrogenase and nitrogenase [122,126,134]. Indirect biophotolysis is well suited for nitrogenase-based systems. The reaction is 12H2O+6CO2+light energy-C6H12O6+O2 C6H12O6+12H2O+light energy-12H2+6CO2 However, hydrogen production is 1000-fold lower in nitrogenasebased systems than in reversible hydrogenase-based ones. When Rhodopseudomonas sphaeroides [135] was cultured in a flat plate reactor with two compartments, it showed a 1.4-fold increase in hydrogen conversion efficiency (to 2.2%) relative to wild type with both compartments, as shown in Table 5. Anabaena is now being considered for hydrogen production. It has a solar 139 conversion efficiency of about 1–2%, with a maximum efficiency of 16% [136]. Nitrogenase is inhibited by oxygen, so oxygen must be excluded from these systems. Also, CO2 concentrations between 4 and 18% w/v are optimal, resulting in higher cell densities and more hydrogen production [137,138]. Researchers believed that one-third of the hydrogen stored in carbohydrates can be recovered by dark fermentation, for a maximum yield of 20% [135,139]. Current photolysis yields under laboratory conditions are 15–20% [140]. However, hydrogen is presently produced mostly by dark or photofermentation. Photosynthetic hydrogen production is best for converting solar energy into hydrogen, so the current focus of research is to increase light-use efficiency and to design better reactors for hydrogen production [132]. 5.2. Biodiesel 5.2.1. Introduction Rudolph diesel first produced methyl esters (diesel) from crops in 1900 [146], and biodiesel has since received much attention as a renewable, biodegradable, and nontoxic source of diesel [147,148]. The United Kingdom consumes nearly 25 billion L of diesel annually; to produce this much biodiesel using oil seeds would require more than half of the land in the UK. In recent years, microalgae have been increasingly considered promising sources of biodiesel because of their high reproduction rates and lipid contents (50–70%); the lipids are transesterified into methyl esters. Algal biodiesel can contain 39–41 MJ/kg caloric value, close to that of petrodiesel (46 MJ/kg) [149]. A recent survey indicated that to fulfill Table 6 Advantages and disadvantages of biofuel derived from microalgae. Advantages Disadvantages High growth rate More cost effective farming Less water demand than land crops Low biomass concentration Difficult to harvest due to microscopic size of most planktonic microalgae Algae can grow on brackish water from saline aquifers or in sea water. While this may solve some of the water availability problems, it will result in other undesirable side effects: salt precipitation on the bioreactor walls; precipitates on pumps and valves leading to reduced lifecycle; presence of salts in the final biomass, which will likely have to be purged with steam. There is a need to develop techniques for growing a single species, reducing evaporation losses and increasing the utilization of CO2. Drying and extraction is difficult. In dry extraction (drying the algae by using the sun or artificially), they receive a much lower yield. When using artificial dryers (such as using electricity) it takes more energy to extract than the energy you can get from the yield. Natural algal stands are not favoured probably due to their low productivity for target organisms. Most of microalgae species are unadapted to local climates and outdoor cultivation. Drying and extraction is difficult. In dry extraction (drying the algae by using the sun or artificially), they receive a much lower yield. When using artificial dryers (such as using electricity) it takes more energy to extract than the energy you can get from the yield. Biodiesel performs poorly compared to its mainstream alternative. High-efficiency CO2 mitigation Algae biofuel contains no sulfur Algae produce nontoxic and highly biodegradable biofuels. Growing algae do not require the use of herbicides or pesticides. High levels of polyunsaturates in algae biodiesel is suitable for cold weather climates A high per-acre yield (7–31 times greater than the next best crop—palm oil) Easy to provide optimal nutrient levels due to the well-mixed aqueous environment as compared to soil Continuous production avoids establishment periods of conventional plants Produces unstable biodiesel with many polyunsaturated Limited genomic data for algal species Large scale extraction procedures for microalgal lipids are complex and still in development stage. Microalgae grown in open pond systems are prone to contamination. Ability to adjust harvest rates to keep culture densities at optimal levels at all times. Especially with the continuous culture systems, such as raceway ponds and bioreactors, harvesting efforts can be controlled to match productivity. Algae oil extracts can be used as livestock feed and even processed into ethanol There exist few commercial cultivating “farms”, so there is a lack of data on largescale cultivation. Capability of performing the photobiological production of biohydrogen. Higher capital costs Algae-based fuel properties allow use in jet fuels. Large-scale production could present many other drawbacks compared to those found in laboratory experiments. 140 A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 50% of the US fuel needs using algae would require only 1–3% of the total US cropping area, while using palm oil would require 24% of that area [54]. Table 6 lists the advantages and disadvantages of biofuel derived from microalgae [42,54,147,149,150]. There are about 300,000 species of algae, some of which, like C. reinhardtii, D. salina, and various Chlorella species, can contain 60% lipids. For algae, concerns are whether to use closed or open photobioreactors, measures to prevent contamination, and how to supply nutrients and CO2 to the cultures. Biodiesel can be extracted from algae using expellers, hexane extraction, or supercritical CO2 extract, with up to 70–75 wt% oil [151]. Biodiesel does not require engine modifications and also reduces CO (by 50%) and CO2 (by 78%) emissions [54]. Algal fuel currently costs about US $52–91/barrel (based on the inflation of 2008) [152]. A main problem of algal biofuels is their high viscosity, which can be 10–20 times more than that of no. 2 diesel fuel. Such high viscosity fuels are difficult to combust and leave deposits on the fuel injector of diesel engines. Therefore, biodiesel is usually blended with conventional diesel. Four techniques can address this problem: pyrolysis, micro-emulsification, dilution, and transesterification [105,153]. Transesterification offers the most promise for lowering viscosity. 5.2.2. Transesterification Transesterification is the chemical conversion of triglycerides into methyl esters using a solvent and catalyst. The reaction converts three moles of alcohol and one mole of triglyceride into one mole of glycerol and three moles of methyl esters, although excess methanol is used to promote biodiesel formation [148,154], giving methyl ester yields greater than 98% w/w [54]. Catalysts include acids (e.g., sulfuric acid), bases (e.g., sodium hydroxide, potassium hydroxide), supercritical fluids, and enzymes such as lipase. Experimentally, base-catalyzed reactions were 4000 times faster than acid-catalyzed ones. Commercially, alkoxides of sodium and potassium are used at 1% per weight of oil formed because of their better catalytic activity compared with simple alcohols. Basecatalyzed reactions are optimized at 60 1C under atmospheric pressure for 90 min. At higher temperatures and pressures, the reaction proceeds faster but is more expensive. The reaction yields two layers, because the excess methanol is insoluble in oil. The biodiesel is then separated from contaminants, e.g., glycerol and solids in a flask separator [155]. Biodiesel may contain free fatty acids which can cause saponification, so after separation the oil is washed with 5% water to prevent yield loss. Transesterification consumes 4.3 MJ/L of biodiesel [148]. The optimum conditions for acidic transesterification are 100% catalyst (e.g., sulfuric acid), a methanol-to-oil ratio of 56:1, and a temperature of 30 1C. The specific gravity of oil reduces from 0.912 to 0.8637 in 4 h [148]. Transesterification by using lipases at low temperatures yields more biodiesel than other transesterification methods. The amount of lipase affects the reaction rate; using 75% immobilized lipase with 10% water gives a 98.15% conversion in 12 h. Unfortunately, lipases are too expensive for commercial production of biodiesel and are deactivated by impurities [156–158]. Nannochloropsis oculata was transesterified in the presence of CaO and Al2O3 catalysts at 50 1C to give an oil conversion of 97.5% [159] (Table 7). In another experiment, algae were transesterified in the presence of zirconia, titania, and alumina catalysts at 350– 400 1C and 2500 psi (17.23 MPa), giving an oil conversion of 90.2% [160]. C. protothecoides was transesterified in the presence of 75% lipase (from Candida sp.) and methanol at 38 1C, with an oil conversion of 98.15% after 12 h [161]. Biodiesel produced from various microalgae species is listed in Table 7. 5.2.3. Transesterification via liquefaction Liquefaction is a process by which wet algal mass is decomposed into liquid fuel in an autoclave at moderate temperatures (300–350 1C) and pressures (5–20 MPa). Fresh algae (thalli) are first autoclaved for 1 h in a nitrogen atmosphere at 3 MPa to prevent water evaporation. Water heated to subcritical conditions decomposes biomass to smaller high-energy molecules. The autoclave is then cooled, the gas fraction is transferred to a cylinder, and the reaction mixture recovered is treated with dichloromethane to form two phases. The solvent is evaporated from the organic mixture to yield an amber-colored oil. Liquefaction requires less input energy (300–350 1C) than either pyrolysis (500 1C) or gasification (2000 1C). Liquefaction can also be used for algae with high moisture contents (4 80– 90 wt%), but reactors are expensive and complex. Supercritical CO2 extraction and liquefaction differ in that the former extracts more long-chain compounds and polysaturates, while liquefaction gives more oils. Quantitatively, liquefaction seems more effective than supercritical CO2 extraction [147]. B. braunii was thermochemically liquefied at 300 1C and 10 MPa in the presence of sodium carbonate to yield 57–64 wt% oil. The caloric value was 45.9 MJ/kg, very close to that of petrodiesel [162]. Similarly, Dunaliella tertiolecta with 78.4 wt% moisture was liquefied at 340 1C and 10 MPa in a hydrogen environment for 60 min to yield 34.9–37% oil of 34.9–36 MJ/kg [163]. 5.2.4. Supercritical methanol transesterification Supercritical methanol transesterification is now being tested for the production of biodiesel. The process is carried out in 100 mL cylinders containing algae, to which methanol is Table 7 Biodiesel production from microalgae. Species Method Catalyst T (1C) P (MPa) Time (min) Yield (wt%) Reference Chlorella protothecoides Botryococcus braunii Dunaliella tertiolecta Nannochloropsis oculata Algae oil Chlorella protothecoides Chlorella protothecoides Chlorella minutissima Transesterification Liquefaction Liquefaction Transesterification Transesterification Transesterification Transesterification Transesterification Transesterification Transesterification H2SO4 Sodium carbonate – CaO/Al2O3 catalyst Zirconia, titania and alumina 75% Candida sp. lipase 30% Immobilized lipase Sodium methylate NaOH KOH 30 300 340 50 300–450 38 38 110 60 90 – – – – 17.23 – – – 240 120 60 240 – 720 720 300 120 90 80 37 97.5 90.20 98.15 98.15 82 93 88 [30] [162] [163] [159] [160] [161] [161] [167] [168] [169] 120 30 [170] Chlorella emersonii Schizochytrium sp Neochloris oleoabundans In Situ Transesterification 70 141 A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 Table 8 Biohydrocarbon production from microalgae. Product Species Process Catalyst/raw material Yield/conversion T (1C) P HHV MJ/kg Reference (MPa) Bio-crude Nannochloropsis/ algae species Dunaliella parva Emiliania huxleyi Botryococcus braunii Microalgae oil Hydrothermal conversion – 30 wt% 300 8.10 – [176] Hydrothermal conversion Pyrolysis Hydrocracking – – – 15.70 wt% 72 wt% methane relative scale 40 wt% (67% jet, 15% diesel) 350 400 – – – – 36.7 – – [177] [181] [196] 75 wt% 270 4 – [196] 37 wt% 300 10 36 [163] 300 – 30–35 [197] Liquefaction in water 5%sodium 64% dry wt basis of oil carbonate Pd/C-catalyzed 57 wt% 3501 – 38 [187] Hydrothermal liquefaction Na2CO3 23.0 wt%. 220–320 – 28–30 [198] Hydro treated 10 wt% Ni/HB with Si/Al ratio of 75 Na2CO3 H+ZSM-5 (1:9) catalyst – 260 78 wt% (60 wt% yield of C18 octadecane, propane (3.6 wt%) and methane (0.6 wt%)) 40 wt% 400 52.7 wt% 500 4 – [179] – – 21.2 18.6 [180] [180] 34.7–39.9 [188] 72.00 wt% 200 2.0 30.11 [189] 65 wt% – – 22 [175] 70–75 wt% (CARBON CONTENT) 300–350 – 23.2/21.2 [191] C20–C30 CH4/alkanes Bio-crude Alkanes (C15 C18) Bio-crude Hydrogenation decarbonylation Zeolitesupported Ni catalysts/ZrO2 Direct liquefaction – Alkanes Dunaliella tertiolecta Botryococcus braunii Nannochloropsis sp. Enteromorpha prolifera Algae species Liquid fuel Bio-crude Chlorella Catalytic pyrolysis Chlorella vulgaris Catalytic pyrolysis Bio-crude/ light fraction Bio-crude Spirulina platensis Thermochemical liquefaction Dunaliella salina Hydrothermal liquefaction Ethanol Chlorella sp. Fermentation Bio-crude Hydrothermal liquefaction Hydrothermal liquefaction Na2CO3 57–65 wt% oil – – – [162] Hydrothermal liquefaction Na2CO3 37 wt% – – – [163] Hydrothermal liquefaction – – – [190] 20.0 wt% – – – [191] Bio-crude Chlorella Catalyzed liquefaction 27.3 wt% – – – [191] Bio-crude Bio-crude Bio-crude Bio-crude C10–C30 Spirulina Chlorella Spirulina Chlorella vulgaris Chllorella protothecoides Microcystis aeruginosa Spirulina Spirulina Chlorella vulgaris Catalyzed liquefaction Catalyzed Liquefaction hydrothermal liquefaction hydrothermal liquefaction pyrolysis Sodium carbonate Sodium carbonate Sodium carbonate KOH KOH Acetic acid Acetic acid – 33–40 wt% Bio-crude Chlorella vulgaris/ Spirulina Botryococcus braunii Dunaliella tertrolecta Microcystis viridis Spirulina Ni/REHY catalyst Yeast Saccharomyces cerevisiae Acetic acid 9 wt% 13.6 wt%. 19.5 wt% 15.7 wt% 17.50 wt% 350 350 300–350 300–350 500 – – – – – – – 21.2 23.2 30 [191] [191] [191] [191] [183] pyrolysis – 23.70 wt% – – 29 [183] Liquefaction in toluene Liquefaction in water Hydrothermal liquefaction Fe(CO)5–S – Ni/Al2O3 catalyst Ni/Al2O3 catalyst – 52.3–66.9 wt% 78.3 wt% 30 wt% 350 350 350 5.0 32–33 – 26 15–20 23.2 [193] [193] [192] 18.10 wt% 350 15–20 17.9 [192] 37 wt% 302 10 36 [104] Sodium carbonate – 64 wt% basis 302 – – [149] 57–64 wt%/( 495%) recovery 302 – – [149] 5% Sodium carbonate Ethanol solvent Methanol solvent Sodium carbonate – – 25.80 wt% 360 – 30.74 [199] 22.7 wt% (C18H36O2) 380 – – [200] 35.53 wt% (C18H36O2) 380 – 39.83 [200] 42 wt% 350 20 17.9 [194] 32.60 wt% 64 wt% 300 1000 10–12 32.0–34.7 – 16.9 Bio-crude CH4/Alkanes Bio-crude Bio-crude Bio-crude Bio-crude C17–C19 Bio-oil Bio-crude Bio-crude Bio-crude Nannochloropsis occulta Bio-oil Dunaliella tertiolecta Bio-oil Botryococcus braunii petroleum oil Botryococcus braunii Bio-oil Dunaliella tertiolecta Bio-crude Spirulina Direct liquefaction Catalyzed liquefaction Hydrothermal liquefaction Direct liquefaction Catalyzed Liquefaction Liquefaction Hydrothermal liquefaction Thermochemical liquefaction Bio-crude Spirulina Thermochemical liquefaction Bio-crude Nannochloropsis sp. Spirulina Spirulina Hydrothermal liquefaction Bio-crude Methanol Hydrothermal liquefaction Gasification 39.9 wt%/50–63 wt% light biocrude 350 [195] [172] 142 A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 Table 8 (continued ) Product Species Process Catalyst/raw material Yield/conversion T (1C) P HHV MJ/kg Reference (MPa) Bio-oil Polytrichum commune Dicranum scoparium Thuidium tamarascinum Sphagnum palustre Drepanocladus revolvens Cladophora fracta Chlorella protothecoides Chlorococcum littorale Spirogyra Pyrolysis – 39.10 wt% 750–775 – 17.4 [184] Pyrolysis – 34.30 wt% 750–775 – 16 [184] Pyrolysis – 33.60 wt% 750–775 – 15.8 [184] Pyrolysis – 37 wt% 750–775 – 16.6 [184] Pyrolysis – 35.40 wt% 750–775 – 15.9 [184] Pyrolysis – 48.20 wt% 750–775 – 19.8 [184] Pyrolysis – 55.30 wt% 750–775 – 23.6 [184] Dark Fermentation – 450 μmol ethanol g 30 – – [175] Fermentation – 14–17 wt% – – – [201] Bio-oil Bio-oil Bio-oil Bio-oil Bio-oil Bio-oil Bio-ethanol Bio-ethanol introduced at supercritical conditions (350–400 1C, 10–25 MPa) [164]. The newly-developed McGyan process yields considerably more biodiesel using a fixed-plate metal oxide catalyst instead of supercritical alcohol and does not lose efficiency over an extended period of time [165]. Biodiesel production using supercritical methanol is economical and gives higher yields than other processes. To produce 1 L of biodiesel, transesterification requires 4.3 MJ of energy, while supercritical methanol requires only 3.3 MJ [166]. 5.3. Biomethanol Microalgae can also be used to produce biomethanol as a renewable fuel, although bioethanol has received more attention than biomethanol, which is corrosive, toxic, and has a high cold point, which causes engine start problems in cold weather [171]. Spirulina is converted into methanol by gasification. Microalgae are first concentrated (1–2% w/v) in settling ponds and centrifuges (2–21% w/v). Spirulina oxidizes into gas, the composition of which determines the methanol yield. Temperatures of 850–1000 1C increase the methanol yield; the maximum yield was 0.64 g of methanol per gram of algal biomass at 1008 1C. Oxygen during gasification is supplied by adsorption. Hot steam helps to reform hydrocarbons, and ash, tar and particulates are removed with a scrubber. Carbon dioxide produced by the reaction is removed by absorption using mono-ethanol–amine. Spirulina yields a heat value of 40–50 kcal/kg by gasification [172]. 1 C. vulgaris and Chlorococcum sp. are widely used for bioethanol production because of their high starch contents. On average, 3.83 g/L of ethanol is produced from 10 g/L of lipid extracted from algae [173,174]. C. littorale produced 450 mmol of ethanol per gram of algae during fermentation at 30 1C [175]. Fermentation of Spirogyra yielded 14–17% bioethanol [123]. In one study, C. vulgaris achieved a conversion efficiency of about 65%. Fermentation enzymes are more active at 25 1C than at 35 1C, and ethanol production ceases at 45 1C, so lower temperatures are better [175]. See Table 8 for additional examples. Bioethanol fermentation from microalgae requires less energy than biodiesel production. The undesired CO2 byproduct can be recycled to cultivate additional algae. However, the commercial production of bioethanol from microalgae is still in the research stages [91]. 5.5. Biohydrocarbons C6H12O6+2ADP+2Pi-2C2H5OH+2CO2+2ATP+2H2O In an experiment oil obtained by hydrothermal conversion of Nannochloropsis water at supercritical conditions was used for hydrolysis. Upon hydrolysis, an organic layer and a solid layer form. The organic layer contains proteins, carbohydrates and fatty acids from the hydrolysis of lipids. Oil is then separated from the aqueous layer via distillation or decanting. Nannochloropsis yielded 30% hydrocarbons at 300 1C and 8.16 MPa, while D. parva yielded 15.7% at 350 1C [176,177]. Other examples are shown in Table 8. Alkanes are also produced by hydrogenation and decarbonylation of microalgae. The reaction is catalyzed by synthetic nickel catalyst and ZrO2 support to produce ketone intermediates, which are then hydrogenated to aldehydes in the presence of Ni catalysts. The reaction occurs in either an autoclave or a continuous flow trickle bed reactor, which stabilizes the catalyst. Algae are placed in the reactor at 260 1C with a 40 MPa hydrogen environment. The total liquid yield contained 70–75 wt% of C17 and n-heptadecane [178]. Alkanes in the range of C15–C18 were obtained by hydro treatment of microalgae in the presence of 10 wt% Ni/Hbeta (Si/Al ¼180) catalyst in batch mode. Conversion took place at 260 1C in a hydrogen environment at 4 MPa and yielded 78 wt% liquid alkanes (60 wt% C18 octadecane, 3.6 wt% propane, and 0.6 wt% methane). A Ni/HBeta catalyst increases the hydrogenation rate and produces propane and fatty acids from saturated triglycerides [179]. See Table 8 for additional examples. The process occurs in two steps. First, enzymatic saccharification hydrolyzes large carbohydrates and cell walls to produce starches. Then, yeasts such as Saccharomyces cerevisiae are added to convert sugars into ethanol. The ethanol is then purified by distillation. 5.5.1. Pyrolysis The pyrolysis of microalgae gives up to 40% oil recovery. This process has a high fuel-to-feed ratio and is the most effective way 5.4. Bioethanol In recent years, ethanol has become an important alternative fuel or fuel supplement. Bioethanol fuel reduces lead, sulfur, CO, and particulate emissions. Ethanol used as a fuel in Brazil reduced carbon emissions by 9.56–106 t, contributing to a reduction of 15% of Brazil's total emissions. Bioethanol can be produced using microalgae as a feedstock for fermentation; bacteria or yeast ferment the carbohydrates, such as glucose and starch, in the microalgae. The overall reaction for ethanol production from biomass is A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 to convert algal biomass to biofuels to eventually replace nonrenewable fossil fuels. Bio-oil production can be considerably improved by using proper catalysts that are effectively preactivated. Chlorella when pretreated with Na2CO3 was less acidic and produced more aromatics with higher caloric value, suggesting that the appropriate catalysts and pretreatment techniques could yield more hydrocarbons than simple pyrolysis alone. After pretreatment, the Chlorella was placed in a fixed-bed reactor and heated from above for 30 min to ensure complete conversion of the microalgae. Hydrocarbons were then condensed as they arose from the reactor. The energy conversion efficiency was 40%, with a caloric value of 21.2 MJ/kg [180]. Emiliania huxleyi, a marine coccolithophore, is a candidate for hydrocarbon production. Methane was obtained when it was subjected to vacuum pyrolysis at 100–500 1C. The highest hydrocarbon yield obtained was 129 mL at 400 1C, 10 times more than at 300 1C, while fewer liquid saturates and aromatics formed at 400 1C, showing that low-temperature pyrolysis is a direct source of hydrocarbon gases [181]. In an experiment, C. vulgaris was pyrolyzed in a fixed-bed reactor in the presence of a 1:9 ZSM-5 catalyst-to-biomass ratio. The yield was 52.7 wt% (Table 6); 25% of the hydrocarbons were alkanes, alkenes, or aromatic compounds. Recent studies show that aromatic yield can be increased to 40 wt% of the bio-oil obtained from pyrolysis by adding appropriate catalysts. Thus, catalysts can both increase the amount of aromatics in bio-oil and also reduce negative outcomes, e.g., oxygen from 30.1 to 19.5 wt%, with an increased caloric value of 32.2 MJ/kg from 24.4 MJ/kg [182]. Recently, fast pyrolysis was performed using C. protothecoides and M. aeruginosa in a fluid bed reactor. High quality bio-oils were produced when the reactor was heated to 500 1C with an average algal feed rate of 4 g/min, a method that could be used for largescale commercial production of bio-oils. Bio-oils obtained by fast pyrolysis have reduced oxygen content with higher caloric values of 29 MJ/kg. Bio-oil yields obtained from C. protothecoides and M. aeruginosa were 17.5 and 23.7 wt% with higher asphaltene and organic content of 35.9 and 29.99 wt% respectively [183]. The bio-oil yield increases considerably as temperature increases from 476.85 to 501.85 1C. The maximum oil yields from Cladophora fracta and C. protothecoides were 48.2% and 55.3 wt%, respectively (Table 6). Bio-oil yields from Chlorella rose from 5.7 to 55.3 wt% as the temperature increased from 202 to 502 1C and decreased to 51.8% at 602 1C [149,184]. 5.5.2. Direct liquefaction Bio-hydrocarbons were produced when D. tertiolecta and B. braunii were subjected to direct liquefaction at 300 1C and 10 MPa; oil yields were 37% and 64% (dry wt basis of oil), respectively (Table 8). Caloric values ranged from 30 to 36 MJ/kg [163,185,186]. Bio-oil was produced from Nannochloropsis sp. by direct liquefaction at 350 1C using different catalysts, e.g., Pd/C, Pt/C, Ru/C, Ni/ SiO2–Al2O3, CoMo/γ-Al2O3, and zeolite, under a hydrogen environment at high pressure. The gaseous products were mainly CO2, H2, and CH4 with minor quantities of C2H2 and C2H6. The highest methane yield was obtained using Ru and Ni catalysts. Higher pressures of hydrogen suppress the gaseous fraction during liquefaction. Bio-oil produced by using Ni catalyst had sulfur content below detection levels. Bio-hydrocarbon yield was 35% from non-catalyzed liquefaction reactions, while a Pd/C catalyst in an H2 atmosphere increased yield up to 57%, with lesser amount of nitrogen [187]. During an investigation of the production of bio-oil from S. platensis, maximum hydrocarbon yield was 39.9% with a carbon conversion efficiency of 98.3% at 350 1C with a holding time of 143 60 min. Biocrude obtained above 300 1C had 71–77% elemental carbon and 0.6–11.6% oxygen with a caloric value of 34.7–39.9 MJ/ kg (Table 8). Gas chromatography–mass spectroscopy (GC/MS) analysis identified hydrocarbons in the range of C16–C17 [188]. Hydro-liquefaction of D. salina was performed in a hydrogen atmosphere with a bifunctional Ni/REHY catalyst at 200 1C and 2 MPa for 60 min. The maximum yield was 72%, with a conversion efficiency of 87.6% (Table 8) [189]. Yang et al. [190] obtained an oil yield of 33–40 wt% from Microcystis viridis subjected to liquefaction in the presence of a Na2CO3 catalyst. Maximum bio-oil yields obtained from Spirulina and Chlorella using an alkali KOH catalyst at 350 1C were 9 wt% and 13.6 wt%, while yields obtained using acetic acid were 19.5 wt% and 15.7 wt%, respectively [191]. However, when yields were expressed on organic basis, Na2CO3 produced higher bio-oil yields of 20.0 wt% and 27.3 wt%, correspondingly (Table 8). The different catalysts were ranked by yield in the order Na2CO3 4 CH3COOH 4 KOH4 HCOOH [110]. Spirulina was liquefied in tetralin at 350 1C for 60 min and yielded 52.3–66.9 wt% oil in the presence of Fe (CO)5–S catalyst in a hydrogen environment at 5 MPa (Table 8). Liquefaction in water gave an oil yield of 78.3 wt% at 350 1C in the presence of nitrogen. GC/MS analysis showed that liquefaction in toluene resulted in higher carbon content with little oxygen and a caloric value of 32–33 MJ/kg [191]. Hydrothermal liquefaction of C. vulgaris and N. occulta gave maximum oil yields of 30 wt% and 18.1 wt%, respectively, in the presence of a Ni/Al2O3 catalyst at 350 1C and 15–20 MPa (Table 6) [113]. Hydrothermal liquefaction converts algal biomass to bio-oil thermochemically. Recent work investigated the hydrothermal conversion of D. tertiolecta at different temperatures and using various catalysts. A maximum oil yield of 25.8% was obtained at 360 1C with a holding time of 50 min using a 5% Na2CO3 catalyst (Table 8). The bio-oil obtained was a mixture of ketones, aldehydes, methyl esters, and fatty acids having a caloric value of 30.74 MJ/kg [192]. The organic solvent can greatly impact the bio-oil product in thermochemical liquefaction. In one study, methanol, ethanol, and 1,4-dioxane were tested with Spirulina. The bio-oil produced using methanol contained more carbon and hydrogen compounds with little oxygen content and a caloric value of 39.83 MJ/kg. The major component obtained using methanol and ethanol was hexadecanoic acid methyl ester (35.53% and 26.27%, respectively; Table 8), while 1,4-dioxane favored the formation of C16H31N (22.7%) [193] Thermochemical liquefaction is a process by which algal biomass is converted into bio-oil using extremely hot (300– 340 1C), high-pressure (up to 20 MPa) water with or without a catalyst. A maximum oil yield of 33 wt% (organic basis) was obtained when M. viridis was subjected to liquefaction in the presence of 5% Na2CO3 at 340 1C for 60 min (Table 8). The bio-oil composition was 62% carbon, 28% hydrogen, 8% N2, and 2% sulfur. Liquefied oil mainly contained alkanes in the range of C17–C18 [190]. Bio-oil yields obtained by hydrothermal liquefaction of Nannochloropsis sp. and Spirulina in the presence of 5% Na2CO3 were 42% and 32.60%, respectively, at 300–350 1C and 10–12 MPa (Table 8) [194,195]. 6. Production costs and life cycle assessment of algae derived fuels Liquid fuels production from microalgae is proven technically, but is still expensive compared to petroleum fuels. Algae fuel is economically viable only in a scenario with crude petroleum selling for ≥$100 per barrel [19,202]. Producing microalgal biomass is generally more expensive than growing crops. Photosynthetic growth requires light, carbon dioxide, water and inorganic 144 A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 salts. Temperature must remain generally within 19–23 1C. To minimize expense, biodiesel production must rely on freely available sunlight, despite daily and seasonal variations in light levels [54]. Microalgae production in closed photobioreactors (PBRs) is highly expensive. Closed systems are much more expensive than ponds. However, the closed systems require much less light and agricultural land to grow the algae. High oil species of microalgae cultured in growth optimized conditions of PBRs have the potential to yield 19,000–57,000 L of microalgal oil per acre per year. The yield of oil from algae is over 200 times the yield from the best-performing plant/vegetable oils [54]. According to Singh et al. [21] more thrust is being made to cultivate algae in closed systems or using photobioreactors i.e. 52% of the world wide technologies being used for algae biofuel production companies, 25% from open ponds and 22% from natural settings. Recovery of oil from microalgal biomass and conversion of oil to biodiesel are not affected by whether the biomass is produced in raceways or photobioreactors. Hence, the cost of producing the biomass is the only relevant factor for a comparative assessment of photobioreactors and raceways for producing microalgal biodiesel. If the annual biomass production capacity is increased to 10,000 t, the cost of production per kilogram reduces to roughly $0.47 and $0.60 for photobioreactors and raceways, respectively, because of economy of scale. Assuming that the biomass contains 30% oil by weight, the cost of biomass for providing a liter of oil would be something like $1.40 and $1.81 for photobioreactors and raceways, respectively [54]. There have been many attempts to estimate this for large scale micro-algae biofuels production using life cycle assessment (LCA) methods to describe and quantify inputs and emissions from the production process. Attempts have been hampered, however, by the fact that no industrial scale process designed specifically for biofuel production yet exists [44]. Slade [44] reviewed the seven recent LCA studies done by Kadam [203], Jorquera [204], Campbell [205], Sander [206], Stephenson [207], Lardon [208], and Clarens [209]. Production systems were compared in terms of the net energy ratio (NER) of biomass production. NER is defined here as the sum of the energy used for cultivation, harvesting and drying, divided by the energy content of the dry biomass. According to the results of this comparison he found NER less than 1 for six raceway ponds out of eight. Hence a positive energy balance may be achievable for these systems, although this benefit is marginal in the normalized case. The NER of the PBR systems are all greater than 1. The best performing PBR is the flat-plate system which outperforms the tubular PBRs as it benefits from a large illumination surface area and low oxygen build-up [44]. He also found that for raceway ponds the base case biomass production cost is ∼1.6–1.8 € kg 1 and the projected case cost is ∼0.3–0.4 € kg 1, and for idealized tubular PBR the base case cost is ∼9–10 € kg 1 and the projected case cost is ∼3.8 € kg 1 [44]. Raceway pond systems Table A1 List of startup companies attempting to commercialize algal fuels. S.N. Company/location Country Website 01 02 03 04 05 06 07 09 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 Algenol Biofuels, Bonita Springs, FL, Fort Meyers Aquaflow Bionomics, Nelson Aurora Algae Inc., Hayward CA Algae Link Roosendaal Aquatic Energy, LLC, Lake Charles Louisiana ALG Western Oil Alga Fuel, S.A., Sines A2BE Carbon Capture, Boulder Colorado Bioalgene, Seatle, WA BFS Biopetróleo, San Vicente del Raaspeig Blue Marble Energy, Seattle, Washington Bionavitas, Inc, Redmond, WA. Bodega Algae, LLC, Boston, MA Cellana, Hawaii Circle Biodiesel and Ethanol Corp., San Marcos, CA Community Fuels, Encinitas, CA Diversified Energy, Gilbert, Arizona Eni Galp Energia, Lisbon Global Energy Solutions, Vancouver Green Fuel Technologies, Cambridge, Massachusetts Green Shift Corp, New York HR Biopetroleum, Hawaii Ingrepo B.V, Zutphen International Energy, Vancouver Inventure Chemical, Seattle LiveFuels, Inc., San Carlos, CA Mighty Algae Biofuels, CA Neste Oil, Helsinki Origin Oil Inc., Los-Angeles, California OilFox S.A Parabel Inc. formerly PetroAlgae Inc., Melbourne, FL PhycoBiosciences, Chandler, AZ PetroSun, Scottsdale, Arizona Sapphire Energy, San Diego SolenaFuels, WA Solix Biofuels, Fort Collins, Colorado Solazyme, Inc., San Francisco Seambiotic, Ashkelon Sartec Anoka, Minnesota Solarvest BioEnergy XL Renewables, Phoenix, Arizona USA New Zealand USA Netherlands USA South Africa Portugal USA USA Spain USA USA USA USA USA USA USA Italy Portugal Canada USA USA USA Netherlands Canada USA USA USA Finland USA Argentina USA USA USA USA USA USA USA Israel USA Canada USA www.algenolbiofuels.comhtpp www.aquaflowgroup.com www.aurorainc.com www.algaelink.com www.aquaticenergy.com www.algbf.co.za www.a4f.pt www.algaeatwork.com www.bioalgene.com www.biopetroleo.com www.bluemarblebio.com www.bionavitas.com www.bodegaalgae.com www.cellana.com www.circlebio.com www.communityfuels.com www.diversified-energy.com www.eni.com www.galpenergia.com www.globalgreensolutionsinc.com 〈www.greenfuelonline.com〉 www.greenshift.com www.hrbiopetroleum.com www.ingrepro.nl www.internationalenergyinc.com www.inventurechem.com www.livefuels.com www.nesteoil.com www.originoil.com www.oilfox.com.ar www.parabel.com www.phyco.net www.sapphireenergy.com www.solenafuels.com www.solixbiofuels.com www.solazyme.com www.seambiotic.com www.xlrenewables.com/ www.xlrenewables.com www.xlrenewables.com A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 demonstrate a lower cost of algal biomass production than photobioreactor systems. Most of the production costs in raceway system are associated with operation (labor, utilities and raw materials). The cost of production in PBRs, in contrast, is dominated by the capital cost of the PBRs [44]. Another recent study done by Jonker [210], regarding the energy consumption ratio and overall bio-energy production costs of micro-algae cultivation, harvesting and its conversion to energy products, gives an insight for future perspectives of micro-algae production for energy purposes in three different climate profiles of Spain. Jonker [210] covered three phases of total chain (cultivation, harvesting and conversion) in the economic evaluation which resulted in total production costs of heat, fuels and electricity derived from micro-algae. He found that the lower end of fuel production cost calculated for raceway ponds is 136 € 2010/GJ and 153 €2010/GJ for horizontal tubular systems which is very much greater than gasoline–diesel 5–20€ 2010/GJ [210]. He found low end of cultivation costs that is 31€ 2010/GJ and 59€ 2010/GJ for raceway and horizontal tubular PBR respectively. And the overall production costs with cost reduction measures are 65, 105, 111 € /GJ for heat, fuel and electricity respectively with raceway ponds, and for horizontal tubular they are 72, 116 and 122 €/GJ [210]. Almost all the LCA studies recommend more technology advancements to make algae fuel commercially viable. 145 Microalgae are an important source of biomass. It has been reviewed that algae biomass can be used to produce biodiesel, biohydrogen, bio-oil, bio-ethanol and biomethanol using thermochemical/hydrothermal liquefaction and gasification processes. Most of the literature reports the advantage of the use of catalysts in the bio-crude production from microalgae. Both homogeneous and heterogeneous catalysts systems have been used in hydrothermal liquefaction of microalgae. Current developments in biofuel catalyst systems for pyrolysis and liquefaction are promising; new catalytic processes may allow the production of gasoline, diesel, and jet fuel that is more economical and competitive than existing fuels. However, many issues are still unclear and require further research to be used on industrial scale. Biohydrogen from microalgae as a clean future energy carrier has also been reported in this study and this also needs more innovative research to improve photo-conversion efficiencies by improving culture conditions and effectiveness of photobioreactors. Appendix A See Table A1. References 7. Conclusion In this work we have reviewed and presented the progresses in the production technologies for making microalgae based liquid fuels. Outcomes of different technologies used for cultivation, extraction and biofuel production via diverse routes from promising microalgae strains were investigated to see the evolution in this field for future perspective. The immense potential of microalgae for producing environmentally sustainable transport fuels is the main motivation behind their development and is receiving support from R&D and investors around the world. This is due to their high biomass productivity, readiness for harvest and high lipid contents (20–75 wt%) than other terrestrial crops. In this review we have covered the best suitable biofuel microalgae strains; among those few are B. braunii, Nannochloropis sp, C. vulgaris, C.a minutissima, C. protothecoides, C. emersonii, S. platensis, S. maxima, D. tertiolecta, P. tricornutum, S. obliquus, Chlorococcum sp, Crypthecodinium cohnii, C. reinhardtii, Schizochytrium sp, D. salina, and Microcystis aeruginosa. These strains are investigated heavily in the literature for fuel purpose due to their higher lipid contents and ability to yield biodiesel, biohydrogen, biohydrocarbons and biomethane. However, these strains still need more ideal culture and growth conditions to obtain algal biomass cheaply. This review also reveals the associated cultivation systems for these strains. Closed photobioreactor configurations seem more promising than open ponds or raceways to meet the need of biofuel industries. Different types of PBRs have been developed over the past decades and T-PBR, FP-PBR and bubble column have shown optimum biomass concentrations. Photobioreactor designs are evolving rapidly and efforts are being made to improve light distribution, mass transfer, shear stress and other PBR operations to make them highly efficient for commercial use. This study also underlines the various methods to extract the algal oil. The solvent based route for different microalgae species has been used extensively and is believed to be most suitable on a large scale. An environmentally friendly technique like SC-CO2 and non-solvent based technologies like microwaves, pulse electric filed and ultrasonic are still in the developmental stage and require detailed R&D for industrial scale processes. [1] Williams PJlB, Laurens LM. Microalgae as biodiesel & biomass feedstocks: review & analysis of the biochemistry, energetics & economics. Energy and Environmental Science 2010;3:554–90. [2] Kheshgi HS, Prince RC, Marland G. The potential of biomass fuels in the context of global climate change: focus on transportation fuels 1. Annual Review of Energy and the Environment 2000;25:199–244. [3] Gallagher BJ. The economics of producing biodiesel from algae. Renewable Energy 2011;36:158–62. [4] Gao Y, Gregor C, Liang Y, Tang D, Tweed C. Algae biodiesel—a feasibility report. Chemistry Central Journal 2012;6:1–16. [5] Sing SF, Isdepsky A, Borowitzka MA, Moheimani NR. Production of biofuels from microalgae. Mitigation and Adaptation Strategies for Global Change 2013;18:47–72. [6] Spolaore P, Joannis-Cassan C, Duran E, Isambert A. Commercial applications of microalgae. Journal of Bioscience and Bioengineering 2006;101:87–96. [7] Suali E, Sarbatly R. Conversion of microalgae to biofuel. Renewable and Sustainable Energy Reviews 2012;16:4316–42. [8] Ratlege C, Dawson PSS, Rattray JB. Biotechnology for the oils & fats industry. Champaign, IL, USA: The American Oil Chemists Society; 1984. [9] Schenk PM, Thomas-Hall SR, Stephens E, Marx UC, Mussgnug JH, Posten C, et al. Second generation biofuels: high-efficiency microalgae for biodiesel production. Bioenergy Research 2008;1:20–43. [10] Singh A, Nigam PS, Murphy JD. Renewable fuels from algae: an answer to debatable land based fuels. Bioresource Technology 2011;102:10–6. [11] Posten C, Schaub G. Microalgae and terrestrial biomass as source for fuels—a process view. Journal of Biotechnology 2009;142:64–9. [12] C. Alasalvar, T. Taylor (Eds.), Food and health applications of marine nutraceuticals: a review. Seafoods—Quality, Technology and Nutraceutical Applications. Springer-Verlag, Berlin (2002), pp. 175–204. [13] Milledge JJ. Commercial application of microalgae other than as biofuels: a brief review. Reviews in Environmental Science and Bio/Technology 2011;10:31–41. [14] Fatih Demirbas M. Biorefineries for biofuel upgrading: a critical review. Applied Energy 2009;86:S151–61. [15] Nigam PS, Singh A. Production of liquid biofuels from renewable resources. Progress in Energy and Combustion Science 2011;37:52–68. [16] Damartzis T, Zabaniotou A. Thermochemical conversion of biomass to second generation biofuels through integrated process design—a review. Renewable and Sustainable Energy Reviews 2011;15:366–78. [17] Demirbas A. Biofuels securing the planet's future energy needs. Energy Conversion and Management 2009;50:2239–49. [18] Khosla V. Where will biofuels and biomass feedstocks come from? White Paper, AfhwkcpWd. [19] Chisti Y, Yan J. Energy from algae: current status and future trends: algal biofuels—a status report. Applied Energy 2011;88:3277–9. [20] Chisti Y. Biodiesel from microalgae beats bioethanol. Trends in Biotechnology 2008;26:126–31. [21] Singh J, Gu S. Commercialization potential of microalgae for biofuels production. Renewable and Sustainable Energy Reviews 2010;14:2596–610. [22] Waltz E. Biotech's green gold? Nature Biotechnology 2009;27:15–8. [23] Torrey M. Algae in the tank. International News on Fats, Oils and Related Materials 2008;19(7):432–7. 146 A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 [24] Piccolo A. Algae oil production and its potential in the Mediterranean region; 2009. [25] Jha A. UK announces world's largest algal biofuel project. The Guardian; 2008. p. 23. [26] Mc Gill R. Algae as a feedstock for transportation fuels–the future of biofuels. A white paper prepared for the IEA advanced motor fuels implementing agreement Presented to the 35th Executive Committee Meeting; 2008. [27] McHugh DJ. A guide to the seaweed industry. Rome: Food and Agriculture Organization of the United Nations; 2003. [28] Anders RMo, David Clayton. Micro- and macro-algae:utility, for industrial applications. CPL Press; 2007. [29] Wang B, Li Y, Wu N, Lan CQ. CO2 bio-mitigation using microalgae. Applied Microbiology and Biotechnology 2008;79:707–18. [30] Xu H, Miao X, Wu Q. High quality biodiesel production from a microalga Chlorella protothecoides by heterotrophic growth in fermenters. Journal of Biotechnology 2006;126:499–507. [31] Scragg A, Illman A, Carden A, Shales S. Growth of microalgae with increased calorific values in a tubular bioreactor. Biomass and Bioenergy 2002;23: 67–73. [32] Ge Y, Liu J, Tian G. Growth characteristics of Botryococcus braunii 765 under high CO2 concentration in photobioreactor. Bioresource Technology 2011;102:130–4. [33] Ethier S, Woisard K, Vaughan D, Wen Z. Continuous culture of the microalgae Schizochytrium limacinum on biodiesel-derived crude glycerol for producing docosahexaenoic acid. Bioresource Technology 2011;102:88–93. [34] Chi Z, Liu Y, Frear C, Chen S. Study of a two-stage growth of DHA-producing marine algae Schizochytrium limacinum SR21 with shifting dissolved oxygen level. Applied Microbiology and Biotechnology 2009;81:1141–8. [35] Richmond A, Cheng-Wu Z. Optimization of a flat plate glass reactor for mass production of Nannochloropsis sp. outdoors. Journal of Biotechnology 2001;85:259–69. [36] Wen Z-Y, Jiang Y, Chen F. High cell density culture of the diatom Nitzschia laevis for eicosapentaenoic acid production: fed-batch development. Process Biochemistry 2002;37:1447–53. [37] Illman A, Scragg A, Shales S. Increase in chlorella strains calorific values when grown in low nitrogen medium. Enzyme and Microbial Technology 2000;27:631–5. [38] Li Z, Yuan H, Yang J, Li B. Optimization of the biomass production of oil algae Chlorella minutissima UTEX2341. Bioresource Technology 2011;102:9128–34. [39] Liang Y, Sarkany N, Cui Y. Biomass and lipid productivities of Chlorella vulgaris under autotrophic, heterotrophic and mixotrophic growth conditions. Biotechnology Letters 2009;31:1043–9. [40] Pruvost J, Van Vooren G, Cogne G, Legrand J. Investigation of biomass and lipids production with Neochloris oleoabundans in photobioreactor. Bioresource Technology 2009;100:5988–95. [41] Tababa HG, Hirabayashi S, Inubushi K. Media optimization of Parietochloris incisa for arachidonic acid accumulation in an outdoor vertical tubular photobioreactor. Journal of Applied Phycology 2012;24:887–95. [42] Gendy T.S., El-Temtamy S.A. Commercialization potential aspects of microalgae for biofuel production: an overview. Egyptian Journal of Petroleum. 2013. [43] Gudin C, Chaumont D. Cell fragility—the key problem of microalgae mass production in closed photobioreactors. Bioresource Technology 1991;38: 145–51. [44] Slade R., Bauen A. Micro-algae cultivation for biofuels: cost, energy balance, environmental impacts and future prospects. Biomass and Bioenergy. 2013. [45] Ugwu C, Aoyagi H, Uchiyama H. Photobioreactors for mass cultivation of algae. Bioresource Technology 2008;99:4021–8. [46] Morita M, Watanabe Y, Saiki H. Investigation of photobioreactor design for enhancing the photosynthetic productivity of microalgae. Biotechnology and Bioengineering 2000;69:693–8. [47] Acien Fernandez F, Hall DO, Canizares Guerrero E, Krishna Rao K, Molina Grima E. Outdoor production of Phaeodactylum tricornutum biomass in a helical reactor. Journal of Biotechnology 2003;103:137–52. [48] Belay A. Mass culture of Spirulina outdoors: the Earthrise farms experience. In: Vonshak A, editor. Spirulina platensis (Arthrospira): physiology, cellbiology and biotechnology. London: Taylor and Francis; 1997. p. 131–158. [49] Soong P. Production and development of Chlorella and Spirulina in Taiwan. Algae biomass. Amsterdam: Elsevier/North Holland Biomedical Press; 97–113. [50] Cysewski G, Lorenz RT. 14 Industrial production of microalgal cell-mass and secondary products–species of high potential. In: Richmond A, editor. Handbook of microalgal culture: biotechnology and applied phycology. Oxford: Blackwell; (2004), p. 281–88. [51] Odlare M, Nehrenheim E, Ribé V, Thorin E, Gavare M, Grube M. Cultivation of algae with indigenous species–potentials for regional biofuel production. Applied Energy 2011;88:3280–5. [52] Weyer KM, Bush DR, Darzins A, Willson BD. Theoretical maximum algal oil production. BioEnergy Research 2010;3:204–13. [53] Amin S. Review on biofuel oil and gas production processes from microalgae. Energy Conversion and Management 2009;50:1834–40. [54] Chisti Y. Biodiesel from microalgae. Biotechnology Advances 2007;25: 294–306. [55] Richmond A. Principles for attaining maximal microalgal productivity in photobioreactors: an overview. Hydrobiologia 2004;512:33–7. [56] Carlsson AS, Bowles DJ. Micro-and macro-algae: utility for industrial applications: outputs from the EPOBIO project. Tall Gables, The Sydings, Speen, Newbury, Berks RG14 1RZ, UK: CPL Press; 2007. [57] Campbell MN. Biodiesel: algae as a renewable source for liquid fuel. Guelph Engineering Journal 2008;1:2–7. [58] Patil V, Reitan K, Knutsen G, Mortensen L, Källqvist T, Olsen E, et al. Microalgae as source of polyunsaturated fatty acids for aquaculture. Current Topics in Plant Biology 2005;6:57–65. [59] Molina E, Fernández J, Acién F, Chisti Y. Tubular photobioreactor design for algal cultures. Journal of Biotechnology 2001;92:113–31. [60] Torzillo G, Pushparaj B, Bocci F, Balloni W, Materassi R, Florenzano G. Production of Spirulina biomass in closed photobioreactors. Biomass 1986;11:61–74. [61] Jacob-Lopes E, Scoparo CHG, LMCF Lacerda, Franco TT. Effect of light cycles (night/day) on CO2 fixation and biomass production by microalgae in photobioreactors. Chemical Engineering and Processing: Process Intensification 2009;48:306–10. [62] Barbosa MJ, Janssen M, Ham N, Tramper J, Wijffels RH. Microalgae cultivation in air‐lift reactors: modeling biomass yield and growth rate as a function of mixing frequency. Biotechnology and Bioengineering 2003;82:170–9. [63] Xu Z, Baicheng Z, Yiping Z, Zhaoling C, Wei C, Fan O. A simple and low-cost airlift photobioreactor for microalgal mass culture. Biotechnology Letters 2002;24:1767–71. [64] Ozkan A., Kinney K., Katz L., Berberoglu H. Reduction of water and energy requirement of algae cultivation using an algae biofilm photobioreactor. Bioresource Technology. 2012. [65] Becerra-Celis G, Tebbani S, Joannis-Cassan C, Isambert A, Boucher P. Estimation of microalgal photobioreactor production based on total inorganic carbon in the medium. In: Proceedings of the 17th IFAC World Congress; 2008. p. 6–11. [66] Barbosa MJ, Zijffers JW, Nisworo A, Vaes W, van Schoonhoven J, Wijffels RH. Optimization of biomass, vitamins, and carotenoid yield on light energy in a flat‐panel reactor using the A‐stat technique. Biotechnology and Bioengineering 2005;89:233–42. [67] Zhang K, Kurano N, Miyachi S. Optimized aeration by carbon dioxide gas for microalgal production and mass transfer characterization in a vertical flat-plate photobioreactor. Bioprocess and Biosystems Engineering 2002;25:97–101. [68] Feng Y, Li C, Zhang D. Lipid production of Chlorella vulgaris cultured in artificial wastewater medium. Bioresource Technology 2011;102:101–5. [69] Gitelson A, Qiuang H, Richmond A. Photic volume in photobioreactors supporting ultrahigh population densities of the photoautotroph Spirulina platensis. Applied and Environmental Microbiology 1996;62:1570–3. [70] Mairet F, Bernard O, Masci P, Lacour T, Sciandra A. Modelling neutral lipid production by the microalga Isochrysis aff. galbana under nitrogen limitation. Bioresource Technology 2011;102:142–9. [71] Degen J, Uebele A, Retze A, Schmid-Staiger U, Trösch W. A novel airlift photobioreactor with baffles for improved light utilization through the flashing light effect. Journal of Biotechnology 2001;92:89–94. [72] Zemke P.E. Mathematical modeling of light utilization and the effects of temperature cycles on productivity in a steady-state algal photobioreactor; 2010. [73] Report for 2005 AZ90B: an outdoor multi-stage C-FPfBoN-CG. [74] Belkacemi K, Hamoudi S. Photobioreactor stability by phase plane technique applied to Spirulina maxima growth. International Journal of Chemical and Reactor Engineering 2003;1:1041. [75] Olaizola M. Commercial development of microalgal biotechnology: from the test tube to the marketplace. Biomolecular Engineering 2003;20:459–66. [76] Molina E, Fernández FA, Camacho FG, Rubio FC, Chisti Y. Scale-up of tubular photobioreactors. Journal of Applied Phycology 2000;12:355–68. [77] Masojídek J, Papáček Š, Sergejevova M, Jirka V, Červený J, Kunc J, et al. A closed solar photobioreactor for cultivation of microalgae under supra-high irradiance: basic design and performance. Journal of Applied Phycology 2003;15:239–48. [78] Kumar K, Dasgupta CN, Nayak B, Lindblad P, Das D. Development of suitable photobioreactors for CO2 sequestration addressing global warming using green algae and cyanobacteria. Bioresource Technology 2011;102:4945–53. [79] Tredici MR, Zittelli GC. Efficiency of sunlight utilization: tubular versus flat photobioreactors. Biotechnology and Bioengineering 1998;57:187–97. [80] Zijffers J-WF, Janssen M, Tramper J, Wijffels RH. Design process of an areaefficient photobioreactor. Marine Biotechnology 2008;10:404–15. [81] Morita M, Watanabe Y, Okawa T, Saiki H. Photosynthetic productivity of conical helical tubular photobioreactors incorporating Chlorella sp. under various culture medium flow conditions. Biotechnology and Bioengineering 2001;74:136–44. [82] Xu Z, Yu J. Hydrodynamics and mass transfer in a novel multi-airlifting membrane bioreactor. Chemical Engineering Science 2008;63:1941–9. [83] Ogbonna JC, Yada H, Tanaka H. Light supply coefficient: a new engineering parameter for photobioreactor design. Journal of Fermentation and Bioengineering 1995;80:369–76. [84] Csőgör Z, Herrenbauer M, Schmidt K, Posten C. Light distribution in a novel photobioreactor–modelling for optimization. Journal of Applied Phycology 2001;13:325–33. [85] Janssen M, Tramper J, Mur LR, Wijffels RH. Enclosed outdoor photobioreactors: light regime, photosynthetic efficiency, scale‐up, and future prospects. Biotechnology and Bioengineering 2003;81:193–210. A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 [86] Scarsella M, Bravi M. Microalgae culturing in thin-layer photobioreactors. Chemical Engineering 2010:20. [87] Mercer P, Armenta RE. Developments in oil extraction from microalgae. European Journal of Lipid Science and Technology 2011;113:539–47. [88] Halim R, Danquah MK, Webley PA. Extraction of oil from microalgae for biodiesel production: a review. Biotechnology Advances 2012;30:709–32. [89] Harun R, Danquah MK. Influence of acid pre-treatment on microalgal biomass for bioethanol production. Process Biochemistry 2011;46:304–9. [90] Halim R, Gladman B, Danquah MK, Webley PA. Oil extraction from microalgae for biodiesel production. Bioresource Technology 2011;102:178–85. [91] Harun R, Singh M, Forde GM, Danquah MK. Bioprocess engineering of microalgae to produce a variety of consumer products. Renewable and Sustainable Energy Reviews 2010;14:1037–47. [92] Banerjee A, Sharma R, Chisti Y, Banerjee U. Botryococcus braunii: a renewable source of hydrocarbons and other chemicals. Critical Reviews in Biotechnology 2002;22:245–79. [93] Mojaat M, Foucault A, Pruvost J, Legrand J. Optimal selection of organic solvents for biocompatible extraction of β-carotene from Dunaliella salina. Journal of Biotechnology 2008;133:433–41. [94] Geciova J, Bury D, Jelen P. Methods for disruption of microbial cells for potential use in the dairy industry—a review. International Dairy Journal 2002;12:541–53. [95] Lewis T, Nichols PD, McMeekin TA. Evaluation of extraction methods for recovery of fatty acids from lipid-producing microheterotrophs. Journal of Microbiological Methods 2000;43:107–16. [96] Lee SJ, Yoon B-D, Oh H-M. Rapid method for the determination of lipid from the green alga Botryococcus braunii. Biotechnology Techniques 1998;12:553–6. [97] Medina AR, Grima EM, Giménez AG, Gonzalez M. Downstream processing of algal polyunsaturated fatty acids. Biotechnology Advances 1998;16:517–80. [98] Kates M. Lipid extraction procedures. Amsterdam, The Netherlands: Techniques of lipidology Elsevier; 100–11. [99] Wang L, Weller CL. Recent advances in extraction of nutraceuticals from plants. Trends in Food Science and Technology 2006;17:300–12. [100] Luque de Castro M, Garcıa-Ayuso L. Soxhlet extraction of solid materials: an outdated technique with a promising innovative future. Analytica Chimica Acta 1998;369:1–10. [101] Cravotto G, Boffa L, Mantegna S, Perego P, Avogadro M, Cintas P. Improved extraction of vegetable oils under high-intensity ultrasound and/or microwaves. Ultrasonics Sonochemistry 2008;15:898–902. [102] Suarsini E, Subandi S. Utilization ultrasonic to increase the efficiency of oil extraction for microalgae indigenous isolates from pond gresik, east java. In: Proceedings of the 2011 IEEE first conference on clean energy and technology (CET); 2011. p. 275–9. [103] /http:// ANPSfAB, www.technologyreview.com/energy/22572/page1/S; 2009. [104] Lee J-Y, Yoo C, Jun S-Y, Ahn C-Y, Oh H-M. Comparison of several methods for effective lipid extraction from microalgae. Bioresource Technology 2010;101: S75–S77. [105] Awasthia M, Singh RK. Development of algae for the production of bioethanol, biomethane, biohydrogen and biodiesel. Trees 2011; 0:50. [106] Balasubramanian S, Allen JD, Kanitkar A, Boldor D. Oil extraction from Scenedesmus obliquus using a continuous microwave system-design, optimization, and quality characterization. Bioresource Technology 2011;102: 3396–403. [107] Patil PD, Gude VG, Mannarswamy A, Cooke P, Munson-McGee S, Nirmalakhandan N, et al. Optimization of microwave-assisted transesterification of dry algal biomass using response surface methodology. Bioresource Technology 2011;102:1399–405. [108] Heger M. A new processing scheme for algae biofuels. Technology review Afhwtcec. [109] Sahena F, Zaidul I, Jinap S, Karim A, Abbas K, Norulaini N, et al. Application of supercritical CO2 in lipid extraction–a review. Journal of Food Engineering 2009;95:240–53. [110] Mendes RL, Nobre BP, Cardoso MT, Pereira AP, Palavra AF. Supercritical carbon dioxide extraction of compounds with pharmaceutical importance from microalgae. Inorganica Chimica Acta 2003;356:328–34. [111] Herrero M, Cifuentes A, Ibanez E. Sub-and supercritical fluid extraction of functional ingredients from different natural sources: plants, foodby-products, algae and microalgae: a review. Food Chemistry 2006;98: 136–48. [112] Grierson S, Strezov V, Bray S, Mummacari R, Danh LT, Foster N. Assessment of biooil extraction from Tetraselmis chui microalgae comparing supercritical CO2, solvent extraction, and thermal processing. Energy and Fuels 2011;26:248–55. [113] Nagle N, Lemke P. Production of methyl ester fuel from microalgae. Applied Biochemistry and Biotechnology 1990;24:355–61. [114] Fajardo AR, Cerdan LE, Medina AR, Fernández FGA, Moreno PAG, Grima EM. Lipid extraction from the microalga Phaeodactylum tricornutum. European Journal of Lipid Science and Technology 2007;109:120–6. [115] Samorì C, Torri C, Samorì G, Fabbri D, Galletti P, Guerrini F, et al. Extraction of hydrocarbons from microalga Botryococcus braunii with switchable solvents. Bioresource Technology 2010;101:3274–9. [116] Couto RM, Simões PC, Reis A, Da Silva TL, Martins VH, Sánchez‐Vicente Y. Supercritical fluid extraction of lipids from the heterotrophic microalga Crypthecodinium cohnii. Engineering in Life Sciences 2010;10:158–64. [117] Chen M, Chen X, Liu T, Zhang W. Subcritical ethanol extraction of lipid from wet microalgae paste of Nannochloropsis sp. Journal of Biobased Materials and Bioenergy 2011;5:385–9. 147 [118] Andrich G, Nesti U, Venturi F, Zinnai A, Fiorentini R. Supercritical fluid extraction of bioactive lipids from the microalga Nannochloropsis sp. European Journal of Lipid Science and Technology 2005;107:381–6. [119] Mendes RL, Coelho JP, Fernandes HL, Marrucho IJ, Cabral J, Novais JM, et al. Applications of supercritical CO2 extraction to microalgae and plants. Journal of Chemical Technology and Biotechnology 1995;62:53–9. [120] Sajilata M, Singhal RS, Kamat MY. Supercritical CO2 extraction of γ-linolenic acid (GLA) from Spirulina platensis ARM 740 using response surface methodology. Journal of Food Engineering 2008;84:321–6. [121] Perretti G, Bravi E, Montanari L, Fantozzi P. Extraction of PUFAs rich oil from algae with supercritical carbon dioxide. In: Proceedings of the 6th international symposium on supercritical fluids; 2003. p. 29–34. [122] Boboescu I, Mirel I, Gherman V. Some considerations on fermentative biohydrogen production. [123] Davila-Vazquez G, Arriaga S, Alatriste-Mondragón F, de León-Rodríguez A, Rosales-Colunga LM, Razo-Flores E. Fermentative biohydrogen production: trends and perspectives. Reviews in Environmental Science and Bio/Technology 2008;7:27–45. [124] Xiong W, Li X, Xiang J, Wu Q. High-density fermentation of microalga Chlorella protothecoides in bioreactor for microbio-diesel production. Applied Microbiology and Biotechnology 2008;78:29–36. [125] Hemschemeier A, Fouchard S, Cournac L, Peltier G, Happe T. Hydrogen production by Chlamydomonas reinhardtii: an elaborate interplay of electron sources and sinks. Planta 2008;227:397–407. [126] Hallenbeck PC. Fermentative hydrogen production: principles, progress, and prognosis. International Journal of Hydrogen Energy. 2009;34:7379–89. [127] Meher Kotay S, Das D. Biohydrogen as a renewable energy resource—prospects and potentials. International Journal of Hydrogen Energy 2008;33:258–63. [128] Das L. Hydrogen engine: research and development (R&D) programmes in Indian Institute of Technology (IIT), Delhi. International Journal of Hydrogen Energy 2002;27:953–65. [129] Das D, Veziro|glu TN. Hydrogen production by biological processes: a survey of literature. International Journal of Hydrogen Energy 2001;26:13–28. [130] Levin DB, Pitt L, Love M. Biohydrogen production: prospects and limitations to practical application. International Journal of Hydrogen Energy 2004;29:173–85. [131] Gaffron H, Rubin J. Fermentative and photochemical production of hydrogen in algae. The Journal of General Physiology 1942;26:219–40. [132] Kruse O, Hankamer B. Microalgal hydrogen production. Current Opinion in Biotechnology 2010;21:238–43. [133] Ni M, Leung MK, Sumathy K, Leung DY. Potential of renewable hydrogen production for energy supply in Hong Kong. International Journal of Hydrogen Energy 2006;31:1401–12. [134] Abed R, Dobretsov S, Sudesh K. Applications of cyanobacteria in biotechnology. Journal of Applied Microbiology 2009;106:1–12. [135] McKinlay JB, Harwood CS. Photobiological production of hydrogen gas as a biofuel. Current Opinion in Biotechnology 2010;21:244–51. [136] Sen U, Shakdwipee M, Banerjee R. Status of biological hydrogen production. Journal of Scientific and Industrial Research 2008;67:980. [137] Zhang L, Melis A. Probing green algal hydrogen production. Philosophical Transactions of the Royal Society of London Series B: Biological Sciences 2002;357:1499–509. [138] Das D, Veziroglu TN. Advances in biological hydrogen production processes. International Journal of Hydrogen Energy 2008;33:6046–57. [139] Hallenbeck PC, Benemann JR. Biological hydrogen production; fundamentals and limiting processes. International Journal of Hydrogen Energy 2002;27:1185–93. [140] Matsumi Y, Comes F., Hancock G, Hofzumahaus A, Hynes A, Kawasaki M, et al. Quantum yields for production of O (1D) in the ultraviolet photolysis of ozone: Recommendation based on evaluation of laboratory data. Journal of Geophysical Research: Atmospheres (1984–2012). 2002;107:ACH 1–12. [141] Benemann J. Feasibility analysis of photobiological hydrogen production. International Journal of Hydrogen Energy 1997;22:979–87. [142] Benemann JR. Hydrogen production by microalgae. Journal of Applied Phycology 2000;12:291–300. [143] Turner J, Sverdrup G, Mann MK, Maness PC, Kroposki B, Ghirardi M, et al. Renewable hydrogen production. International Journal of Energy Research 2008;32:379–407. [144] Melis A, Happe T. Hydrogen production. Green algae as a source of energy. Plant Physiology 2001;127:740–8. [145] Tamburic B., Zemichael F.W., Maitland G.C., Hellgardt K. A novel nutrient control method to deprive green algae of sulphur and initiate spontaneous hydrogen production. International Journal of Hydrogen Energy. 2012. [146] Ramadhas A, Muraleedharan C, Jayaraj S. Performance and emission evaluation of a diesel engine fueled with methyl esters of rubber seed oil. Renewable Energy 2005;30:1789–800. [147] Demirbas A, Fatih Demirbas M. Importance of algae oil as a source of biodiesel. Energy Conversion and Management 2011;52:163–70. [148] Demirbas A. Importance of biodiesel as transportation fuel. Energy Policy 2007;35:4661–70. [149] Demirbas A. Use of algae as biofuel sources. Energy Conversion and Management 2010;51:2738–49. [150] Cheng JJ, Timilsina GR. Status and barriers of advanced biofuel technologies: a review. Renewable Energy 2011;36:3541–9. [151] Sheehan J, Dunahay T, Benemann J, Roessler PA. Look back at the US Department of Energy's Aquatic Species. Golden, Colorado, USA: Close-Out 148 [152] [153] [154] [155] [156] [157] [158] [159] [160] [161] [162] [163] [164] [165] [166] [167] [168] [169] [170] [171] [172] [173] [174] [175] [176] [177] [178] [179] [180] A. Bahadar, M. Bilal Khan / Renewable and Sustainable Energy Reviews 27 (2013) 128–148 Report National Renewable Energy Laboratory, US Department of Energy's Office of Fuels Development; 1998. Demirbas MF. Biofuels from algae for sustainable development. Applied Energy 2011;88:3473–80. Marchetti J, Miguel V, Errazu A. Possible methods for biodiesel production. Renewable and Sustainable Energy Reviews 2007;11:1300–11. Guan G, Kusakabe K, Sakurai N, Moriyama K. Transesterification of vegetable oil to biodiesel fuel using acid catalysts in the presence of dimethyl ether. Fuel 2009;88:81–6. Hossain A, Salleh A, Boyce AN, Chowdhury P, Naqiuddin M. Biodiesel fuel production from algae as renewable energy. American Journal of Biochemistry and Biotechnology 2008;4:250–4. Fjerbaek L, Christensen KV, Norddahl B. A review of the current state of biodiesel production using enzymatic transesterification. Biotechnology and Bioengineering 2009;102:1298–315. Akoh CC, Chang S-W, Lee G-C, Shaw J-F. Enzymatic approach to biodiesel production. Journal of Agricultural and Food Chemistry 2007;55:8995–9005. Du W, Li W, Sun T, Chen X, Liu D. Perspectives for biotechnological production of biodiesel and impacts. Applied Microbiology and Biotechnology 2008;79:331–7. Umdu ES, Tuncer M, Seker E. Transesterification of Nannochloropsis oculata microalga's lipid to biodiesel on Al2O3 supported CaO and MgO catalysts. Bioresource Technology 2009;100:2828–31. McNeff CV, McNeff LC, Yan B, Nowlan DT, Rasmussen M, Gyberg AE, et al. A continuous catalytic system for biodiesel production. Applied Catalysis A: General 2008;343:39–48. Cheng Y, Zhou W, Gao C, Lan K, Gao Y, Wu Q. Biodiesel production from Jerusalem artichoke (Helianthus Tuberosus L.) tuber by heterotrophic microalgae Chlorella protothecoides. Journal of Chemical Technology and Biotechnology 2009;84:777–81. Dote Y, Sawayama S, Inoue S, Minowa T, Yokoyama S-Y. Recovery of liquid fuel from hydrocarbon-rich microalgae by thermochemical liquefaction. Fuel 1994;73:1855–7. Minowa T, Yokoyama S-Y, Kishimoto M, Okakura T. Oil production from algal cells of Dunaliella tertiolecta by direct thermochemical liquefaction. Fuel 1995;74:1735–8. Demirbaş A. Production of biodiesel from algae oils. Energy Sources, Part A: Recovery, Utilization, and Environmental Effects 2008;31:163–8. Krohn BJ, McNeff CV, Yan B, Nowlan D. Production of algae-based biodiesel using the continuous catalytic Mcgyans process. Bioresource Technology 2011;102:94–100. Saka S, Kusdiana D. Biodiesel fuel from rapeseed oil as prepared in supercritical methanol. Fuel 2001;80:225–31. Tang H, Chen M, Garcia M, Abunasser N, Ng K, Salley SO. Culture of microalgae Chlorella minutissima for biodiesel feedstock production. Biotechnology and Bioengineering 2011;108:2280–7. Carvalho J, Ribeiro A, Castro JDF, Vilarinho C, Castro F. Biodiesel production by microalgae and macroalgae from north littoral portuguese coast; 2011. Veillette M, Chamoumi M, Faucheux N, Heitz M. Microalgae-based oil for biodiesel production; 2011. Levine RB, Costanza-Robinson MS, Spatafora GA. Neochloris oleoabundans grown on anaerobically digested dairy manure for concomitant nutrient removal and biodiesel feedstock production. Biomass and Bioenergy 2011;35:40–9. Yeole SD, Aglave B, Lokhande M. Algaeoleum—a third generation biofuel. Asian Journal of Bio Science 2009;4:344–7. Hirano A, Hon-Nami K, Kunito S, Hada M, Ogushi Y. Temperature effect on continuous gasification of microalgal biomass: theoretical yield of methanol production and its energy balance. Catalysis Today 1998;45:399–404. Hirano A, Ueda R, Hirayama S, Ogushi Y. CO2 fixation and ethanol production with microalgal photosynthesis and intracellular anaerobic fermentation. Energy 1997;22:137–42. Harun R, Danquah MK, Forde GM. Microalgal biomass as a fermentation feedstock for bioethanol production. Journal of Chemical Technology and Biotechnology 2010;85:199–203. Ueno Y, Kurano N, Miyachi S. Ethanol production by dark fermentation in the marine green alga, Chlorococcum littorale. Journal of Fermentation and Bioengineering 1998;86:38–43. WU BDeluca C Payne E. Systems and methods for hydrothermal conversion of algae into biofuel. WO patent 2,010,021,753; 2010. Bloch MR, Sasson J., Ginzburg ME, Goldman Z, Ginzburg BZ, Garti N, et al. Oil products from algae. Google patents; 1982. Peng B, Yuan X, Zhao C, Lercher JA. Stabilizing catalytic pathways via redundancy: selective reduction of microalgae oil to alkanes. Journal of the American Chemical Society 2012;134:9400–5. Peng B, Yao Y, Zhao C, Lercher JA. Towards quantitative conversion of microalgae oil to diesel—range alkanes with bifunctional catalysts. Angewandte Chemie 2012;124:2114–7. Babich I, van der Hulst M, Lefferts L, Moulijn J, O'Connor P, Seshan K. Catalytic pyrolysis of microalgae to high-quality liquid bio-fuels. Biomass and Bioenergy 2011;35:3199–207. [181] Wu Q, Dai J, Shiraiwa Y, Sheng G, Fu J. A renewable energy source— hydrocarbon gases resulting from pyrolysis of the marine nanoplanktonic alga Emiliania huxleyi. Journal of Applied Phycology 1999;11:137–42. [182] Thangalazhy-Gopakumar S, Adhikari S, Chattanathan SA, Gupta RB. Catalytic pyrolysis of green algae for hydrocarbon production using H+ ZSM-5 catalyst. Bioresource Technology 2012. [183] Miao X, Wu Q, Yang C. Fast pyrolysis of microalgae to produce renewable fuels. Journal of Analytical and Applied Pyrolysis 2004;71:855–63. [184] Demirbaş A. Oily products from mosses and algae via pyrolysis. Energy Sources Part A 2006;28:933–40. [185] Patil V, Tran K-Q, Giselrød HR. Towards sustainable production of biofuels from microalgae. International Journal of Molecular Sciences 2008;9:1188–95. [186] Sawayama S, Inoue S, Dote Y, Yokoyama S-Y. CO2 fixation and oil production through microalga. Energy Conversion and Management 1995;36:729–31. [187] Duan P, Savage PE. Hydrothermal liquefaction of a microalga with heterogeneous catalysts. Industrial and Engineering Chemistry Research 2010;50: 52–61. [188] Jena U, Das K, Kastner J. Effect of operating conditions of thermochemical liquefaction on biocrude production from Spirulina platensis. Bioresource Technology 2011;102:6221–9. [189] Yang C, Jia L, Chen C, Liu G, Fang W. Bio-oil from hydro-liquefaction of Dunaliella salina over Ni/REHY catalyst. Bioresource Technology 2011;102:4580–4. [190] Yang Y, Feng C, Inamori Y, Maekawa T. Analysis of energy conversion characteristics in liquefaction of algae. Resources, Conservation and Recycling 2004;43:21–33. [191] Ross A, Biller P, Kubacki M, Li H, Lea-Langton A, Jones J. Hydrothermal processing of microalgae using alkali and organic acids. Fuel 2010;89: 2234–43. [192] Biller P, Riley R, Ross A. Catalytic hydrothermal processing of microalgae: decomposition and upgrading of lipids. Bioresource Technology 2011;102: 4841–8. [193] Matsui T-o Nishihara A, Ueda C, Ohtsuki M, Ikenaga N-O, Suzuki T. Liquefaction of micro-algae with iron catalyst. Fuel 1997;76:1043–8. [194] Biller P, Ross A. Potential yields and properties of oil from the hydrothermal liquefaction of microalgae with different biochemical content. Bioresource Technology 2011;102:215–25. [195] Vardon DR, Sharma B, Scott J, Yu G, Wang Z, Schideman L, et al. Chemical properties of biocrude oil from the hydrothermal liquefaction of Spirulina algae, swine manure, and digested anaerobic sludge. Bioresource Technology 2011;102:8295–303. [196] Hillen L, Pollard G, Wake L, White N. Hydrocracking of the oils of Botryococcus braunii to transport fuels. Biotechnology and Bioengineering 1982;24: 193–205. [197] Sawayama S, Inoue S, Dote Y, Yokoyama S-Y. CO2 fixation and oil production through microalgae. Energy Conversion and Management 1995;36(6-9):729–31. [198] Zhou D, Zhang L, Zhang S, Fu H, Chen J. Hydrothermal liquefaction of macroalgae Enteromorpha prolifera to bio-oil. Energy and Fuels 2010;24: 4054–61. [199] Shuping Z, Yulong W, Mingde Y, Kaleem I, Chun L, Tong J. Production and characterization of bio-oil from hydrothermal liquefaction of microalgae Dunaliella tertiolecta cake. Energy 2010;35:5406–11. [200] Yuan X, Wang J, Zeng G, Huang H, Pei X, Li H, et al. Comparative studies of thermochemical liquefaction characteristics of microalgae using different organic solvents. Energy 2011;36:6406–12. [201] Bush RA, Hall KM. Process for the production of ethanol from algae. Google Patents; 2006. [202] Stephens E, Ross IL, King Z, Mussgnug JH, Kruse O, Posten C, et al. An economic and technical evaluation of microalgal biofuels. Nature Biotechnology 2010;28:126–8. [203] Kadam K. Environmental implications of power generation via coalmicroalgae cofiring. Energy 2002;27:905–22. [204] Jorquera O, Kiperstok A, Sales EA, Embiruçu M, Ghirardi ML. Comparative energy life-cycle analyses of microalgal biomass production in open ponds and photobioreactors. Bioresource Technology 2010;101:1406–13. [205] Campbell PK, Beer T, Batten D. Life cycle assessment of biodiesel production from microalgae in ponds. Bioresource Technology 2011;102:50–6. [206] Sander K, Murthy GS. Life cycle analysis of algae biodiesel. International Journal of Life Cycle Assessment 2010;15:704–14. [207] Stephenson AL, Kazamia E, Dennis JS, Howe CJ, Scott SA, Smith AG. Life-cycle assessment of potential algal biodiesel production in the United Kingdom: a comparison of raceways and air-lift tubular bioreactors. Energy and Fuels 2010;24:4062–77. [208] Lardon L, Hélias A, Sialve B, Steyer J-P, Bernard O. Life-cycle assessment of biodiesel production from microalgae. Environmental Science and Technology 2009;43:6475–81. [209] Clarens AF, Resurreccion EP, White MA, Colosi LM. Environmental life cycle comparison of algae to other bioenergy feedstocks. Environmental Science and Technology 2010;44:1813–9. [210] Jonker J, Faaij A. Techno-economic assessment of micro-algae as feedstock for renewable bio-energy production. Applied Energy 2012;102:461–75 February 2013.