Microalgae

To protect the rights of the author(s) and publisher we inform you that this PDF is an uncorrected proof for internal business use only by the author(s), editor(s), reviewer(s), Elsevier and typesetter TNQ Books and Journals Pvt Ltd. It is not allowed to publish this proof online or in print. This proof copy is the copyright property of the publisher and is confidential until formal publication.

C H A P T E R

11 Microalgae: The Tiny Microbes with a Big Impact

c0011

Shovon Mandal 1, Nirupama Mallick 2,* 1

Section of Ecology, Behavior and Evolution, University of California, San Diego, CA, USA, Agricultural and Food Engineering Department, Indian Institute of Technology, Kharagpur, West Bengal, India *Corresponding author email: [email protected]

2

O U T L I N E

s0005 p0010

Renewable Energy

1

Fatty Acid Methyl Esters and Fuel Properties

Petroleum Fuel Scenario in India

2

9

Biodiesel

2 3

Waste Utilization for Biodiesel Production: A Case Study with Scenedesmus obliquus in a Recirculatory Aquaculture System

Microalgae: Viable Feedstocks for Biodiesel

9

Selection of Potent Strains

3

Concluding Remarks

11

Genetic Engineering Approach

5

References

11

Microalgal Biodiesel Production

7

RENEWABLE ENERGY

planet (Shay, 1993). The use of renewable energy is largely motivated from the standpoint of global energy crisis and environmental issues. Renewable energy is a form of energy that is produced from natural sources like sunlight, wind, hydropower, geothermal and biomass, which can be naturally replenished. Currently, renewable energy supplies only w18% of the world’s energy consumption (Kumar et al., 2010). Most of these renewable energy sources (hydropower, wind, solar and geothermal) target the electricity market, while the majority of world energy consumption (about two-thirds) is derived from liquid fuels (Campbell, 2008; Hankamer et al., 2007). This has stimulated recent interest to explore alternative sources for petroleumbased fuels and much of the attention has been focused on biomass-derived liquid fuels or biofuels (Haag, 2007; Schneider, 2006).

Energy is an important currency for human society. The world population growth and rapid economic progresses are expected to result in considerable increase in the demand for energy. In the reference scenario, the International Energy Agency has projected an increase in energy need by 55%, between 2005 and 2030, at an average annual rate of 1.8% (IEA, 2007). Driven by such increasing demand, and the dwindling fuel production, the cost of petroleum fuel has gone up sky high in recent times, which can jeopardize the economic progresses of a nation. Despite the fuel crisis, increasing concentrations of CO2 and other heattrapping greenhouse gases (GHGs) in the atmosphere, primarily due to the combustion of fossil fuels, is clearly the prime reason for rapid warming of the

Bioenergy Research: Advances and Applications http://dx.doi.org/10.1016/B978-0-444-59561-4.00011-5

1 Copyright Ó 2014 Elsevier B.V. All rights reserved.

10011-GUPTA-9780444595614


2

11. MICROALGAE: THE TINY MICROBES WITH A BIG IMPACT

s0010 PETROLEUM FUEL SCENARIO IN INDIA p0015

s0015 p0020

India ranks seventh as the world’s energy producer accounting for about 2.5% of the world’s total annual energy production, and world’s fifth largest energy consumer with about 3.5% of the global primary energy demand (IEA, 2007; Planning Commission, Govt. of India, 2007). Despite being among the largest energy producer, India is a net importer of energy, largely due to huge imbalance between energy consumption and production. About 30% of India’s total primary energy need is being met by petroleum oil, of which 76% is imported. India’s transportation fuel requirements are unique in the world. India consumes almost five times more diesel fuel than gasoline, whereas all other countries in the world use more gasoline than diesel fuel (Khan et al., 2009). Thus, search for alternatives to diesel fuel is of special importance in India. Bioalcohols are unsuitable substitutes for diesel engines, because of their low cetane numbers (CNs) along with poor energy content per unit biomass (Bhattacharyya and Reddy, 1994; Rao and Gopalkrishnan, 1991). Therefore, biodiesel is the only option to fulfill the requirements in future.

BIODIESEL

Biodiesel is chemically monoalkyl esters of longchain fatty acids derived from vegetable oils or animal fats. The history of using vegetable oil as an alternative fuel dates back to 1900, when Rudolph Diesel used peanut oil as fuel in the World Exhibition in Paris. It was found that vegetable oils, in general, have acceptable CNs and calorific values comparable with the conventional diesel. However, the major problem with the direct use of vegetable oils as fuel of compression ignition engine is their high viscosity, which interferes with the fuel injection and atomization contributing to incomplete combustion, nozzle clogging, injector coking, severe engine deposits, ring sticking and gum formation leading to engine failure (Knothe, 2005; Meher et al., 2006; Singh and Rastogi, 2009). Therefore, vegetable oils need to be modified to bring their combustion-related properties closer to those of diesel fuel. One possible method to overcome the problem of high viscosity of vegetable oils is their chemical modification to esters, what is nowadays called as “biodiesel”. p0025 Biodiesel has emerged as the most suitable alternative to petroleum diesel fuel owing to its ecofriendly characteristics and renewability (Krawczyk, 1996). It burns in conventional diesel engines with or without any modifications while reducing pollution (100% less sulfur dioxide, 37% less unburned hydrocarbons,

46% less carbon monoxide, and 84% less particulate matter) in comparison to the conventional diesel fuel (McMillen et al., 2005). The basic feedstocks for the production of first-generation biodiesel were mainly edible vegetable oils like soybean, rapeseed, sunflower and safflower. The use of first-generation biodiesel has generated a lot of controversy, mainly due to their impact on global food markets and food security for diverting food away from the human food chain. The second-generation biodiesel was produced by using nonedible oil sources like used frying oil, grease, tallow, lard, karanja, jatropha and mahua oils (Alcantara et al., 2000; Francis and Becker 2002; Canakci and Gerpen, 1999; Dorado et al., 2002; Ghadge and Raheman, 2006; Mittelbach, 1990). Nevertheless, the cost of biodiesel production is still a major obstacle for large-scale commercial exploitation, mainly due to the high feed cost of vegetable oils (Lang et al., 2001). Moreover, the first- as well as the second-generation biodiesel based on terrestrial plants initiate land clearing and potentially compete with net food production (Chisti, 2008; Marsh, 2009). The focus of researchers has now been shifted to the next generation biodiesel. The third-generation biodiesel is both promising and different; it is based on simple microscopic organisms that live in water and grow hydroponically, i.e. microalgae. The possibilities of biodiesel production from edible p0030 oil resources in India is almost impossible, as primary need is to first meet the demand of edible oil that is being imported. India accounts for 9.3% of the world’s total oil seed production and contributes as the fourth largest edible oil producing country. Even then, about 46% of edible oil is imported for catering the domestic needs (Jain and Sharma, 2010). So the nonedible oil resources like Jatropha, pongamia, mahua, etc. seem to be the only possibility for biodiesel production in the country. The Government of India has duly realized the importance of biodiesel and introduced a nationwide program under the National Biodiesel Mission in 2003 with the aim of achieving a target of meeting 13.4 Mt of biodiesel (@ 20% blending) from Jatropha curcas by 2012, and to achieve the target about 27 billion of planting materials are required to be planted over 11.2 million hectares of land (Planning Commission, Govt. of India, 2003). At the current rate of consumption, if all petroleumderived transport fuel is to be replaced with biodiesel from Jatropha oil, Jatropha would need to be grown over an area of 384 million hectares, which is more than 100% of the geographic area of India (Khan et al., 2009). Therefore, India must find additional, reliable, cost-effective and sustainable feedstock for biodiesel production. In this context, biodiesel from microalgae seems to be a suitable substitute for diesel fuel in the long run.

10011-GUPTA-9780444595614


SELECTION OF POTENT STRAINS

s0020 p0035

MICROALGAE: VIABLE FEEDSTOCKS FOR BIODIESEL Microalgae are a diverse group of photosynthetic organisms whose systematics is based on the kinds and combinations of photosynthetic pigments present in different species. They can grow in diverse environmental conditions, and are able to produce a wide range of chemical products with applications in feed, food, nutritional, cosmetic and pharmaceutical industries. These are primitive organisms with a simple cellular structure and a large surface to volume body ratio, which gives them the ability to take up a large amount of nutrients. While the mechanism of photosynthesis in microalgae is similar to that of higher plants, they have the ability to capture solar energy with an efficiency of 10e50 times higher than that of terrestrial plants (Li et al., 2008). Moreover, because the cells grow in aqueous suspension, they have more efficient access to water, CO2 and other nutrients. For these reasons, microalgae are capable of producing more amount of oil per unit area of land in comparison to that of all other known oil-producing crops (Chisti, 2007; Haag, 2007). The per hectare yield of microalgal oil has been projected to be 58,700e136,900 l/year depending upon the oil content of algae, which is about 10e20 times higher than the best oil producing crop, i.e. palm (5950 l/ha year, Chisti, 2007). The most acclaimed energy crop, i.e. Jatropha has been estimated to produce only 1892 l/ha year. More importantly, due to being aquatic in nature, algae do not compete for arable land for their cultivation; they can be grown in freshwater or saline, and salt concentrations up to twice that of seawater can be used effectively for few species (Aresta et al., 2005; Brown and Zeiler, 1993). The utilization of wastewaters that are rich in nitrogen and phosphorus may bring about remarkable advantages by providing N and P nutrients for growing microalgae, while removing N and P from the wastewaters (Mallick, 2002). This implies that algae need not compete with other users for freshwater (Campbell, 2008). On top of these advantages, microalgae grow even better when fed with extra carbon dioxide, the main GHG. If so, these tiny organisms can fix CO2 from power stations and other industrial plants, thereby cleaning up the greenhouse problem. Each ton of algae produced consumes about 1.8 ton of CO2 (Chisti, 2007). Thus, the integrated efforts to cleanup industrial flue gas with microalgal culture by combining it with wastewater treatment will significantly enhance the environmental and economical benefits of the technology for biodiesel production by minimizing the additional cost of nutrients and saving the precious freshwater resources.

SELECTION OF POTENT STRAINS

3

s0025

Realizing the oil-yielding potentialities with much p0040 faster growth rate and efficient CO2 fixation, microalgae appear to be the best option as a renewable source of biodiesel that has the potentiality to completely replace the petroleum diesel fuel. However, the lipid content in the selected microalga/strain is required to be high; otherwise the economic performance would be hard to achieve. Each species of microalga produces different ratios of p0045 lipids, carbohydrates and proteins. Nevertheless, these tiny organisms have the ability to manipulate their metabolism by simple manipulations of the chemical composition of the culture medium (Behrens and Kyle, 1996); thus, high lipid productivity can be achieved. Physiological stresses such as nutrient limitation/deficiency, salt stress and high light intensity have been employed for directing metabolic fluxes to lipid biosynthesis of microalgae. Many reports are available, where attempts have been made to raise the lipid pool of various microalgal species. Table 11.1 summarizes those studies. Exceptionally, an oil content of 86% of dry cell weight p0050 (dcw) was reported in the brown resting state colonies of Botryococcus braunii, while the green active state colonies were found to account for 17% only (Brown et al., 1969). However, the major obstacle in focusing B. braunii as an industrial organism for biodiesel production is its poor growth rate (Dayananda et al., 2007). Nitrogen limitation/deficiency has been found to raise the lipid content of a number of microalgal species profoundly. For instance, Piorreck and Pohl (1984) reported an increased lipid pool from 12% to 53% (dcw) in Chlorella vulgaris under nitrogen-limited condition. Unlike the green algae, the blue-green algae viz. Anacystis nidulans and Oscillatoria rubescens contained the same quantities of lipid at different nitrogen concentrations. It was observed by Illman et al. (2000) that four species of Chlorella (Chlorella emersonii, Chlorella minutissima, C. vulgaris and Chlorella pyrenoidosa) could accumulate lipid up to 63, 57, 40 and 23% (dcw), respectively, in low N-medium. These values in control vessels were, respectively, 29%, 31%, 18% and 11% in the above order. In the same year, Takagi et al. (2000) observed an increase in intracellular lipid pool up to 51% (dcw) against 31% control in 3% CO2-purged cultures of Nannochloris sp. UTEX LB1999 grown in continuous low nitrate (0.9 mM)-fed medium. Chlorella protothecoides also showed a rise in lipid pool from 15% to 55% (dcw), when grown heterotrophically with glucose (1%) under reduced nitrogen concentration (Miao and Wu, 2004). Similarly, C. protothecoides depicted a lipid pool of 55% (dcw) when grown heterotrophically with corn powder hydrolysate under nitrogen limitation (Xu et al., 2006).

10011-GUPTA-9780444595614


4 t0010


TABLE 11.1

A List of Studies on Increased Lipid Accumulation in Microalgae under Various Specific Conditions

Microalga

Growth Condition

Lipid Content as Percent of Dry Cell Weight

Botryococcus braunii

Brown resting state

86 (17*)

Brown et al. (1969)

Chlorella vulgaris

Nitrogen limitation

53 (12*)

Piorreck and Pohl (1984)

Chlorella emersonii

Nitrogen limitation

63 (29*)

Illman et al. (2000)

Chlorella minutissima

57 (31*)

Chlorella vulgaris

40 (18*)

Chlorella pyrenoidosa

23 (11*)

References

Nannochloris sp. UTEX LB1999

Nitrogen limitation

51 (31*)

Takagi et al. (2000)

Chlorella protothecoides

Heterotrophy with 0.1% glucose under nitrogen limitation

55 (15*)

Miao and Wu (2004)

Heterotrophy with corn powder hydrolysate under nitrogen limitation

55 (15*)

Xu et al. (2006)

Dunaliella sp.

1 M NaCl

71 (64*)

Takagi et al. (2006)

Chlorella sp.

Heterotrophy with 1% sucrose

33 (15*)

Rattanapoltee et al. (2008)

Scenedesmus obliquus

Nitrogen and phosphorus limitations in presence of thiosulphate

58 (13*)

Mandal and Mallick (2009)

Neochloris oleoabundans

Nitrogen deficiency

56 (29*)

Gouveia and Oliveira (2009)

Nannochloropsis oculata NCTU-3

2% CO2

50 (31*)

Chiu et al. (2009)

Nannochloropsis sp. F&M-M24

Nitrogen deficiency

60 (31*)

Rodolfi et al. (2009)

Phosphorus deficiency

50 (31*)

Nannochloropsis oculata

Nitrogen limitation

15 (8*)

Chlorella vulgaris

Converti et al. (2009)

16 (6*)

Choricystis minor

Nitrogen and phosphorus deficiencies

60 (27*)

Sobczuk and Chisti (2010)

Haematococcus pluvialis

High light intensity

35 (15*)

Damiani et al. (2010)

High light intensity under nitrogen deficiency

33 (15*)


Heterotrophy with sweet sorghum hydrolysate under nitrogen limitation

50 (15*)

Gao et al. (2010)

Chlorella zofingiensis

Nitrogen limitation

55 (27*)

Feng et al. (2011)a

Isochrysis zhangjiangensis

High nitrogen (0.9%) supplementation

53 (41*)

Feng et al. (2011)b

Dunaliella tertiolecta

Nitrogen deficiency

26 (12*)

Jiang et al. (2012)

Thalassiosira pseudonana Chlorella vulgaris

20 (13*) Nitrogen, phosphorus and iron limitations

57 (8*)

* Lipid content of control culture.

10011-GUPTA-9780444595614

Mallick et al. (2012)


GENETIC ENGINEERING APPROACH

p0055

Gao et al. (2010) used sweet sorghum hydrolysate instead of corn powder for C. protothecoides culture, and lipid yield of 50% (dcw) was recorded. Nitrogen limitation/starvation also enhanced the lipid content in Neochloris oleoabundans, Nannochloropsis oculata, C. vulgaris, Chlorella zofingiensis, Dunaliella tertiolecta and Thalassiosira pseudonana (Converti et al., 2009; Feng et al., 2011a; Gouveia and Oliveira, 2009; Jiang et al., 2012). However, the marine microalga Isochrysis zhangjiangensis was found to accumulate lipid under high nitrate concentration, rather than limitation or depletion (Feng et al., 2011b). p0060 Limitation of phosphate was also found to enhance lipid accumulation in Ankistrodesmus falcutus and Monodus subterraneus (Kilham et al., 1997; Khozin-Goldberg and Cohen, 2006). Rodolfi et al. (2009) screened 30 microalgal strains for lipid production, among which the marine genus Nannochloropsis sp. F&M-M24 emerged as the best candidate for oil production (50% under phosphorus deficiency against 31% control). Sobczuk and Chisti (2010) observed an increase in intracellular lipid content up to 60% (dcw) against 27% control in Choricystis minor under simultaneous nitrate and phosphate deficiencies. In Scenedemus obliquus, lipid accumulation up to 58% (dcw) was recorded when subjected to simultaneous nitrate and phosphate limitations in presence of sodium thiosulphate (against 13% under control condition, Mandal and Mallick, 2009). Simultaneous nitrate, phosphate and iron limitations have also been reported to stimulate lipid accumulation in a microalga C. vulgaris (57% against 8% control, Mallick et al., 2012). p0065 In addition to nutrient limitations/deficiencies, other stress conditions may also enhance lipid accumulation in microalgae. Takagi et al. (2006) studied the effect of NaCl on accumulation of lipids and triacylglycerides in the marine microalga Dunaliella sp. Increase in initial NaCl concentration from 0.5 M (seawater) to 1.0 M resulted in a higher intracellular lipid accumulation (71% dcw). Damiani et al. (2010) studied the effects of continuous high light intensity (300 mmol photons/ m2 s) on lipid accumulation in Haematococcus pluvialis grown under nitrogen-sufficient and nitrogen-deprived conditions. A lipid yield of 33e35% was recorded under the high light intensity as compared to 15% yield in control cultures. Nitrogen deprivation was, however, not found to raise the lipid content of H. pluvialis cultures. p0070 Nutrient limitations/deficiencies or physiological stresses required for accumulation of lipids in microalgal cells is associated with reduced cell division (Ratledge, 2002). The overall lipid productivity is therefore compromised due to the low biomass productivity. For instance, Scragg et al. (2002) studied the energy recovery from C. vulgaris and C. emersonii grown in complete Watanabe medium and also in low nitrogen

5

medium. The results showed that the low nitrogen medium, although induced higher lipid accumulation in both the test algae with high calorific values, the overall energy recovery was lower in comparison to Watanabe’s medium. A commonly suggested counter measure is to use a two-stage cultivation strategy, dedicating the first stage for cell growth/division in nutrient sufficient medium, and the second stage for lipid accumulation under nutrient starvation or other physiological stresses. To get maximal biomass and lipid yield, CO2 can also be utilized. Chiu et al. (2009) reported an increased accumulation of lipid (from 31% to 50% dcw) in the stationary phase cultures of N. oculata NCTU-3 grown under 2% CO2 aeration.

GENETIC ENGINEERING APPROACH

s0030

High oil-yielding transgenic microalgae could be a p0075 promising source for biodiesel production. However, the biotechnological processes based on transgenic microalgae are still in infancy. In manipulation of genetically modified algae for high oil content, acetyl-CoA carboxylase (ACCase) was first isolated from the diatom Cyclotella cryptica by Roessler (1990), and then successfully transformed into the diatoms C. cryptica and Navicula saprophila (Dunahay et al., 1995, 1996; Sheehan et al., 1998). A plasmid was constructed that contained acc1 gene driven by the cauliflower mosaic virus 35S ribosomal gene promoter (CaMV35S) and the selectable marker nptII from Escherichia coli. Introduction of plasmids into the diatoms was mediated by microprojectile bombardment. The acc1 was overexpressed with the enzyme activity enhanced by threefold. These experiments demonstrated that ACCase could be transformed efficiently into microalgae, although no significant increase in lipid accumulation was observed in the transgenic diatoms (Dunahay et al., 1995, 1996). Recently, diacylglycerol acyltransferases (DGATs) homologous genes have been identified in the genome of Chlamydomonas reinhardtii and were overexpressed in the same microalga (Russa et al., 2012). This resulted in an enhanced mRNA expression level of DGAT genes, but did not boost the intracellular triacylglycerol (TAG) synthesis. Thus, till date, there is no success story with respect to lipid overproduction in microalgae using the genetic engineering approach. Extensive studies have also been carried out on p0080 enhancement of lipid production using genetic engineering approaches in different bacterial and plant species, which may provide valuable background for future studies with microalgae. Some of these studies are summarized in Table 11.2. The cytosolic ACCase from Arabidopsis sp. was overexpressed in Brassica napus (rapeseed) plastids. The fatty acid content of the

10011-GUPTA-9780444595614


6 t0015 TABLE 11.2


A List on Trials to Enhance Lipid Biosynthesis in Transgenic Organisms

Gene (Enzyme)

Source Species

Receiver Species

Result

References

acc1 (ACCase)

Cyclotella cryptica

Cyclotella cryptica

3-fold rise in ACCase activity, no change in lipid content

Dunahay et al. (1995, 1996)

Navicula saprophila

3-fold rise in ACCase activity, no change in lipid content

acc1 (ACCase)

Arabidopsis sp.

Brassica napus

2-fold rise in plastid ACCase, 6% rise in fatty acid content

Roesler et al. (1997)

LPAT

Saccharomyces cerevisiae

Brassica napus

6-fold rise in oil content

Zou et al. (1997)

accA, accB, accC, accD (ACCase)

E. coli

E. coli

6-fold rise in fatty acid synthesis

Davis et al. (2000)

are1 and are2 (DGAT)

Arabidopsis thaliana

Saccharomyces cerevisiae

9-fold rise in TAG content

Bouvier-Nave et al. (2000)

DGAT

Arabidopsis sp.

Arabidopsis sp.

70% rise in lipid content

Jako et al. (2001)

acc1 (ACCase)

Arabidopsis sp.

Solanum tuberosum

5-fold rise in TAG content

Klaus et al. (2004)

acs (ACS)

E. coil

E. coli

9-fold rise in ACS activity

Lin et al. (2006)

malEMt and malEMc (malic enzyme, ME)

Mortierella alpina and Mucor circinelloides

M. circinelloides

2.5-fold rise in lipid accumulation

Zhang et al. (2007)

fadD, ACCase, thioesterase (TE)

E. coil

E. coli

20-fold rise in fatty acid synthesis

Lu et al. (2008)

wri1

Brassica napus

Arabidopsis thaliana

40% rise in oil content

Liu et al. (2010)

Acyl-ACP thioesterase

Diploknema butyracea, Ricinus communis and Jatropha curcas

E. coli

0.2e2.0 g/l free fatty acid yield

Zhang et al. (2011)

DGAT

Chlamydomonas reinhardtii

C. reinhardtii

29-Fold rise in mRNA level, no change in TAG

Russa et al. (2012)

Source: Modified from Courchesne et al., 2009

recombinant was 6% higher than that of the control (Roesler et al., 1997). In prokaryotes like E. coli, overexpression of four ACCase subunits resulted in sixfold rise in the rate of fatty acid synthesis (Davis et al., 2000), confirming that the ACCase-catalyzed committing step was indeed the rate-limiting step for fatty acid biosynthesis in this strain. Nevertheless, Klaus et al. (2004) achieved an increase in fatty acid synthesis and a more than fivefold rise in the amount of TAG in Solanum tuberosum (potato) by overexpressing the ACCase from Arabidopsis in the amyloplasts of potato tubers. p0085 Transformation of rape seed with a putative sn-2-acyltransferase gene from Saccharomyces cerevisiae was carried out by Zou et al. (1997), leading to overexpression of seed lysophosphatidate acid acyl-transferase (LPAT) activity. This enzyme is involved in TAG formation and its overexpression led to profound rise in oil content from 8% to 48% on seed dry weight basis. However, it was cautioned that the steady state level of diacylglycerol could be perturbed by an increase in LPAT activity in the developing seeds. Transformations of

S. cerevisiae with the Arabidopsis DGAT were performed by Bouvier-Nave et al. (2000). About 600-fold rise in DGAT activity in the transformed S. cerevisiae was observed, which led to a ninefold increase in TAG accumulation. DGAT gene has also been overexpressed in the plant Arabidopsis and it was shown that the oil content was enhanced in correlation with the DGAT activity (Jako et al., 2001). All these results suggest that the reaction catalyzed by ACCase, LPAT and DGAT are important rate-limiting steps in lipid biosynthesis. A few enzymes that are not directly involved in lipid p0090 metabolism have also been demonstrated to influence the rate of lipid accumulation. For instance, it was observed by Lin et al. (2006) that by overexpressing the acs gene in E. coli, the acetyl-CoA synthase activity was increased by ninefold, leading to a significant increase in the assimilation of acetate from the medium, which can contribute to lipid biosynthesis. The genes coding for malic enzyme from Mucor circinelloides (malEMt) and from Mortierella alpina (malEMc), respectively, were overexpressed in M. circinelloides which led to a 2.5-fold increase in lipid accumulation (Zhang et al.,

10011-GUPTA-9780444595614


MICROALGAL BIODIESEL PRODUCTION

2007). Lu et al. (2008) reported a 20-fold enhancement of fatty acid productivity of E. coli by combining four targeted genotypic changes: deletion of the fadD gene encoding the first enzyme in fatty acid degradation, overexpression of the genes encoding the endogenous ACCase, and overexpression of both an endogenous thioesterase (TE) as well as a heterologous plant TE. Overexpression of wri1 gene from B. napus in transgenic Arabidopsis thaliana resulted into 40% increased seed oil content (Liu et al., 2010). Zhang et al. (2011) studied the effects of the overexpression of different acyl-ACP TE genes from Diploknema butyracea, Ricinus communis and J. curcas on free fatty acid contents of E. coli. The strain carrying the acyl-ACP TE gene from D. butyracea produced approximately 0.2 g/l of free fatty acid while the strains carrying acyl-ACP TE genes from R. communis and J. curcas produced the free fatty acid at a high level of more than 2.0 g/l.

s0035 MICROALGAL BIODIESEL PRODUCTION p0095

Microalgal biodiesel production is relatively new and not very well explored. Some reports are available, where attempts have been made to produce biodiesel from algae (Table 11.3). Miao and Wu (2006) reported that lipid extracted from the heterotrophically grown microalga, C. protothecoides, transformed into biodiesel with a yield of 63% under 1:1 weight ratio of H2SO4 to oil, and 56:1 molar ratio of methanol to oil at 30 C for a reaction time of 4 h. Xu et al. (2006) characterized the biodiesel obtained from the C. protothecoides oil by acid-catalyzed transesterification. The most abundant fatty acid methyl ester (FAME) in C. prothecoides biodiesel was methyl oleate (61% of total FAME) followed by methyl linoleate (17%) and methyl palmitate (13%). Subsequently, Li et al. (2007) showed that it was feasible to grow C. protothecoides in a commercial-scale bioreactor. Using 75% immobilized lipase, these researchers claimed w98% conversion could be obtained in 12 h when the reaction condition with respect to solvent type, water content and pH were optimized. Hossain and Salleh (2008) studied biodiesel production from Oedogonium and Spirogyra species using NaOH as catalyst. Algal oil and biodiesel production was higher in Oedogonium sp. than in Spirogyra sp. Umdu et al. (2009) studied the effects of Al2O3 supported CaO and MgO catalysts in the transesterification of lipid of N. oculata. These researchers found that pure CaO and MgO were not active, and CaO-Al2O3 catalyst showed the highest activity. Biodiesel yield was increased up to 98% from 23% under CaO-Al2O3 catalyzed reaction. p0100 Lipid extracted from N. oleoabundans was found to have an adequate fatty acid profile and iodine value according to the biodiesel specifications of European

7

Standards (EN, Gouveia et al. 2009). Converti et al. (2009) analyzed the FAMEs in biodiesel produced from N. oculata and C. vulgaris. The most abundant composition was methyl palmitate, which was 62% and 66%, respectively, in N. oculata and C. vulgaris biodiesel. However, the concentration of linolenic acid (18%) in N. oculata could not meet the requirement of European legislation for biodiesel. Johnson and Wen (2009) prepared biodiesel from the microalga Schizochytrium limacinum by direct transesterification of algal biomass. Parameters such as free glycerol, total glycerol, acid number, soap content, corrosiveness to copper, flash point and viscosity met the American Society for Testing and Materials (ASTM) and European standards, while the water and sediment content, as well as the sulfur content did not pass the standards. Damiani et al. (2010) studied biodiesel production from H. pluvialis using potassium hydroxide as the catalyst. The major constituent of H. pluvialis biodiesel was palmitic acid followed by linoleic, oleic and linolenic acid methyl esters. The iodine value was within the limit established by European standards. Chinnasamy et al. (2010) produced biodiesel by a two-step transesterification process (acid-catalyzed followed by base-catalyzed) from a consortium of 15 native algae cultivated in carpet industry wastewater. Algal methyl esters were predominated by linolenic, linoleic, palmitic and oleic acids. The biodiesel was found to contain 0.0155% and 0.0001% bound and free glycerin, respectively, and met the ASTM and European standard specifications. Patil et al. (2012) optimized the direct conversion of p0105 wet Nannochlopsis sp. biomass to biodiesel under supercritical methanol treatment, without using any catalyst. In the supercritical state, at high pressure and temperature, the methanol molecules enabled simultaneous extraction and transesterification of lipids in wet algal biomass. The abundant FAME in Nannochlopsis sp. biodiesel was methyl oleate (37%) followed by methyl palmitolate (32%) and methyl palmitate (8%). Velasquez-Orta et al. (2012) compared in situ transesterification of C. vulgaris with acid as well as alkaline catalysts, in which the oil extraction step was also eliminated. FAME yield reached a maximum of 77.6% after 45 min using a catalyst (NaOH) ratio of 0.15:1 and solvent ratio of 600:1 at 60 C under constant stirring rate of 380 rpm. However, with sulfuric acid as catalyst FAME yield reached up to 96.9% with catalyst : oil ratio of 0.35:1 for a reaction time of 20 h. Recently, Mallick et al. (2012) characterized the biodiesel obtained from the C. vulgaris oil by acid-catalyzed transesterification. The fuel properties (density, viscosity, acid value, iodine value, calorific value, cetane index, ash and water contents) of C. vulgaris biodiesel are comparable with the international (ASTM and EN) and Indian standards (IS).

10011-GUPTA-9780444595614

Attempts on Biodiesel Production from Microalgae

8

Properties of Biodiesel Major Ester

Physical Property

References


H2SO4-catalyzed (63%)

NC

Density: 0.86 kg/l, viscosity: 5.2 cSt, flash point: 115 C, acid value: 0.37 mg KOH/g, heating value: 41 MJ/kg

Miao and Wu (2006)


Methyl oleate: 61%, methyl linoleate: 17%, methyl palmitate: 13%

Density: 0.86 kg/ l, viscosity: 5.2 cSt, flash point: 115 C, solidifying point: 12 C, acid value: 0.37 mg KOH/g

Xu et al. (2006)

Lipase-catalyzed (98%)

Methyl oleate: 65%, methyl linoleate: 18%, methyl palmitate: 10%

NC

Li et al. (2007)

Oedogonium sp.

NaOH-catalyzed (95%)

NC

NC

Hossain and Salleh (2008)

Spirogyra sp.

NaOH-catalyzed (93%)


Heterogeneous catalyst (Al2O3supported CaO & MgO) (98%)

NC

NC

Umdu et al. (2009)

Neochloris oleoabundans

BF3-catalyzed (NR)

Methyl oleate: 38%, methyl palmitate: 17%, methyl stearate: 14%, methyl linolenate: 8%

Iodine value: 72 g I2/100 g

Gouveia et al. (2009)


Acid-catalyzed (NR)

Methyl palmitate: 62%, methyl linolenate: 18%, methyl linoleate: 12%, methyl oleate: 6%

NC

Converti et al. (2009)

Chlorella vulgaris

Methyl palmitate: 66%, methyl linolenate: 12%, methyl linoleate: 11%, methyl oleate: 7%

Schizochytrium limacinum


Methyl palmitate: 57%, methyl ester of C22: 6:30%

Viscosity: 3.87 cSt, flash point: 204 C, moisture content: 0.11%, acid value: 0.11 mg KOH/g, total glycerin: 0.097%, free glycerin: 0.003%,

Johnson and Wen (2009)

Haematococcus pluvialis

KOH-catalyzed (NR)

Methyl palmitate: 23%, methyl linoleate: 20%, methyl oleate: 19%, methyl linolenate: 16%

Iodine value: 111 g I2/100 g

Damiani et al. (2010)

A consortium of 15 native microalgae

Acid-catalyzed followed by base-catalyzed (64%)

Methyl linolenate: 28%, methyl linoleate: 20%, methyl palmitate: 16%, methyl oleate: 12%

Bound glycerin: 0.0155%, free glycerin: 0.0001%

Chinnasamy et al. (2010)

Nannochloropsis sp.

Supercritical methanol

Methyl oleate: 37%, methyl palmitoleate: 23%, methyl palmitate: 8%

NC

Patil et al. (2012)

Chlorella vulgaris

Alkaline in situ (78%)

Methyl linolenate: 22%, methyl oleate: 21%, methyl stearate: 11%

NC

Velasquez-Orta et al. (2012)

Chlorella vulgaris

HCl-catalyzed (NR)

Methyl palmitate: 62%, methyl oleate: 20%, methyl linoleate: 10%

Density: 0.88 kg/l, viscosity: 4.5 cSt, calorific value: 38.4 MJ/kg, iodine value: 56.2 g I2/ 100 g, acid value: 0.6 mg KOH/g, cetane index: 54.7, ash content: 0.01%, water content: 0.03%

Mallick et al. (2012)

NR, not reported; NC, not characterized.


10011-GUPTA-9780444595614

Name of the Alga

Transesterification Process with % Conversion


t0020 TABLE 11.3


WASTE UTILIZATION FOR BIODIESEL PRODUCTION: A CASE STUDY WITH SCENEDESMUS OBLIQUUS

s0040 p0110

p0115

p0120

p0125

p0130

FATTY ACID METHYL ESTERS AND FUEL PROPERTIES As stated before, biodiesel is the best substitute for petrodiesel due to its fuel properties, which are very close to those of diesel. Diesel is a mixture of C15 to C18 hydrocarbons obtained from crude oil in the distillation range of 250e350 C. It contains only carbon and hydrogen atoms, which are arranged in straight or branched chain structures, as well as aromatic configurations. Diesel may contain both saturated and unsaturated hydrocarbons. Biodiesel rather has a different chemical structure than the conventional diesel fuel. It is monoalkyl esters of long-chain fatty acids derived from various types of vegetable oils. The fatty acids are of C12 to C24, with over 90% of them being between C16 and C18. Fuel properties of biodiesel that are influenced by the fatty acid profile and, in turn, by the structural features of various fatty acid esters is CN, which ultimately affects the exhaust emission, heat of combustion, cold flow, oxidative stability, viscosity, and lubricity. Structural features of a fatty acid ester molecule that influence the physical and fuel properties are chain length and degree of unsaturation (Knothe, 2005). Since biodiesel is produced in quite differently scaled plants from vegetable oils of varying origin and quality, it is necessary to install a standardization of fuel quality to guarantee engine performance without any difficulties. CN is widely used as diesel fuel quality parameter related to the ignition delay time and combustion quality. The higher the CN is the better are the ignition properties (Meher et al., 2006). High CNs ensure good cold start properties and minimize the formation of white smoke. The longer the fatty acid carbon chains and the more saturated the molecules are, the higher are the CNs (Bajpai and Tyagi, 2006). According to Knothe and Dunn. (2003), high CNs are observed for esters of saturated fatty acids such as palmitic and stearic acids. The oxidation stability decreased with increase in the contents of polyunsaturated FAMEs (Ramos et al., 2009). The limitation of unsaturated fatty acids is also necessary due to the fact that heating of higher unsaturated fatty acids results in polymerization of glycerides. This can lead to the formation of heavy deposits in the machines (Mittelbach, 1996). One of the major problems associated with the use of biodiesel is its poor cold flow property, indicated by relatively high cloud point and pour point. Saturated fatty acids have significantly higher melting points and crystallize even at room temperature. Thus biodiesel produced from the sources with high amounts of saturated fats would show higher cloud points and pour points. Viscosity also increases with the increasing degree of saturation and chain length (Knothe, 2005). Unsaturated fatty acids exhibit better lubricity than

9

saturated ones (Kenesey and Ecker, 2003). Heat of combustion increases with the chain length and decreases with unsaturation (Goering et al., 1982). The increase in heat content results from a gross increase in number of carbon and hydrogen as well as increase in the ratio of these elements relative to oxygen. Therefore no single fatty acid could fulfill every fuel properties. Rather, a very good compromise can be reached by considering a fuel rich in the monounsaturated fatty acids, such as oleate or palmitoleate, and low in both saturated and polyunsaturated fatty acids (Durrett et al., 2008).

WASTE UTILIZATION FOR BIODIESEL PRODUCTION: A CASE STUDY WITH SCENEDESMUS OBLIQUUS IN A RECIRCULATORY AQUACULTURE SYSTEM

s0045

Nowadays, waste disposal is a worldwide problem. In p0135 agricultural countries like India, waste discharges from agriculture, agrobased industries and city sewages are the main sources of water pollution. Conventional wastewater treatment systems do not seem to be the definitive solution to pollution and eutrophication problems. The major drawbacks are cost and lack of nutrient recycling (Eisenberg et al., 1981). Secondary sewage treatment plants are specifically designed to control the quantity of organic compounds in wastewaters. Other pollutants including nitrogen and phosphorus are only slightly affected by this type of treatment (Gates and Borchardt, 1964). Owing to the ability to use nitrogen and phosphorus for growth, algae can successfully be cultivated in such type of wastewaters (Mallick, 2002). This has been evolved from the early work of Oswald (Oswald et al., 1953) using microalgae in tertiary treatment of municipal wastewaters. The widely used microalgae cultures for nutrient removal are Chlorella (Gonza´lez et al., 1997; Lee and Lee, 2001), Scenedesmus (Martinez et al., 1999, 2000) and Spirulina (Olguıń et al., 2003). Nutrient removal efficiency of Nannochloris sp. (Jimenez-Perez et al., 2004), B. brauinii (An et al., 2003), and Phormidium sp. (Dumas et al., 1998; Laliberte et al., 1997) has also been investigated. One of the well-known algae-based bioprocesses for wastewater treatment is high-rate algal ponds (Cromar et al., 1996; Deviller et al., 2004). Recently, corrugated raceways (Craggs et al., 1997; Olguıń et al., 2003), triangular photobioreactors (Dumas et al., 1998), and tubular photobioreactors (Briassoulis et al., 2010; Molina et al., 2000) have been developed for nutrient removal. Among agroindustries, a large quantity of waste- p0140 water is generated from intensified aquaculture practices. The main source of potentially polluting waste in fish culture is feed derived, mainly un.consumed and undigested feed and fish excreta. Discharging these

10011-GUPTA-9780444595614


10


effluents directly into water resources causes eutrophication of the receiving waters. Qian et al. (1996) reported the collapse of a prawn industry in China due to outbreak of pathogenic bacteria caused by high nutrient load. A few studies have shown the efficiency of algae biofilters in removing nitrogen from fish effluents (Cohen and Neori, 1991; Jimenez del Rio et al., 1996; Schuenhoff et al., 2003). These works are based on the use of seaweeds of the genera Ulva and Gracilaria to treat effluent water from aquaculture. p0145 Recently, we intend to explore an integrated approach to produce biodiesel with simultaneous waste recycling by a green microalga S. obliquus with three types of wastes, viz. poultry litter (PL), fish pond discharge (FPD), and municipal secondary settling tank discharge. Our initial trial under laboratory batch culture conditions (Mandal and Mallick, 2011) encouraged us to conduct a small-scale field experiment in a recirculatory aquaculture system (RAS) using FPD and PL with the same microalga (Mandal and Mallick, 2012). Figure 11.1 presents a schematic diagram of RAS, developed at Agricultural and Food Engineering Department, Indian Institute of Technology Kharagpur, West Bengal, India. The effluent from a fish pond was pumped into a settling tank for removal of large solids. After 24 h, the supernatant was siphoned to an inclined plate settler for removal of fine solids. To have a clear picture of the

inclined plate settler readers are requested to refer Sarkar et al. (2007). The effluent was then entered into fiber-reinforced plastic tanks (length 125 cm, breadth 60 cm, depth 45 cm) for culturing the test microalga. FPD has a very high load of solid particles in suspen- p0150 sion, which contributes to increase in turbidity. Experiments carried on with sedimented (after passing through inclined plate settler) and nonsedimented FPD showed that the nutrient removal efficiency of S. obliquus was higher in the sedimented one. Further experiments with sedimented FPD demonstrated that biomass and lipid yield was maximum at 15 cm culture depth with stirring. In seasonal variation study, the maximum algal biomass and lipid productivity was recorded during summer when sunshine hour was relatively large. During the summer season, when S. obliquus cultures pregrown in FPD supplemented with 5 g PL/l were transferred to the optimized conditions to maximize the lipid accumulation (to have details on optimized condition readers are requested to refer Mandal and Mallick, 2009), lipid yield was raised by more than sixfold (up to 780 mg/l, Mandal and Mallick, 2012). During rainy and winter seasons, comparable lipid yield was recorded by providing artificial lights for few hours. Thus an areal lipid productivity of 14,000 l/ha year (approximately) has been projected assuming 11 cultivation cycles per year, leaving the rest of the period for

FP : fish pond CP : centrifugal pump ST : settling tank IPS : inclined plate settler ACT : algae culture tank RWT : remediated water tank ST

RWT

f0010 FIGURE 11.1

Diagrammatic representation of recirculatory aquaculture system (RAS).

10011-GUPTA-9780444595614


REFERENCES

cleaning and maintenance of the system (Mandal and Mallick, 2012). Nevertheless, this value is w8 times higher than that of Jatropha, one of the most acclaimed energy crops (Khan et al., 2009).

s0050

CONCLUDING REMARKS

p0155

Chisti (2007) envisioned a lipid productivity of 58,700e136,900 l/ha year, considering lipid content of 30e70% of dry biomass. However, to attain this, increasing the volumetric and areal production rates should be the focus (Grobbelaar, 2012). Grobbelaar (2009) projected the upper limits of biomass productivity of about 200 g (dcw)/m2 day. Six decades of worldwide research on outdoor mass cultivation of microalgae, however, have demonstrated only a diminutive fraction of this, where the highest value being recorded was 30 g (dcw)/m2 day (Lee, 2001). As opined by Grobbelaar (2012), with the available strains, an average long-term rate close to 50 g (dcw)/m2 day could be attainable by optimizing various conditions, such as culture depth, mixing, nutrients and CO2 supply, temperature and light, and controlling predators, pathogens and alien microalgal invasion in open raceways. This equates to an annual productivity of about 150 tons of algal biomass per hectare. Thus, maximum lipid productivity would vary between 50,000 and 84,000 l/ha year, considering lipid content of 30e50% of dry biomass. p0160 For the last few years, there has been a worldwide impetus to achieve commercial-scale production of biodiesel from microalgae. In October 28, 2010, US Naval base in Norfolk, Virginia, completed a successful test by running a 15 m gunboat with 50:50 mix of algaebased fuel and diesel. However, the cost of this mix was $112 for liter. In March 2012, the US Navy put another milestone by sailing a fleet ship w1200 miles on “Soladiesel”, an algae-based fuel blend. Recently, researchers at Brookhaven National Laboratory, USA, have announced the development of a new process that could significantly lower the cost. Nevertheless, cost of producing microalgal biodiesel can be reduced further by using a biorefinery-based production strategy, like a petroleum refinery, where each and every component is used to produce valuable products. There is much to be researched in this exciting and upcoming field, and the only thing we can say for certain is that the best method/technology of biofuel production will survive and rise above the others. It is our job as researchers to find this, and be proud to be a part of this endeavor. c0011

Acknowledgments Nirupama Mallick is thankful to NFBSFARA, Indian Council of Agricultural Research, New Delhi, India, for financial support in the form of

11

a project (Studies on Microalgal Triacylglycerols as a Source of Biodiesel) to continue research efforts in this exciting and imminent field.

References Alcantara, R., Amores, J., Canoira, L., Fidalgo, E., Franco, M.J., Navarro, A., 2000. Catalytic production of biodiesel from soybean oil, used frying oil and tallow. Biomass Bioenergy 18, 515e527. An, J.Y., Sim, S.J., Lee, J.S., Kim, B.K., 2003. Hydrocarbon production from secondarily treated piggery wastewater by the green algae Botryococcus braunii. J. Appl. Phycol. 15, 185e191. Aresta, M., Dibenedetto, A., Carone, M., Colonna, T., Fagale, C., 2005. Production of biodiesel from macroalgae by supercritical CO2 extraction and thermochemical liquifaction. Environ. Chem. Lett. 3, 136e139. Bajpai, D., Tyagi, V.K., 2006. Biodiesel: source, production, composition, properties and its benefits. J. Oleo Sci. 55, 487e502. Behrens, P.W., Kyle, D.J., 1996. Microalgae as a source of fatty acids. J. Food Lipids 3, 259e272. Bhattacharyya, S., Reddy, C.S., 1994. Vegetable oils as fuel for internal combustion engine: a review. J. Agric. Eng. Res. 57, 157e166. Bouvier-Nave, P., Benveniste, P., Oelkers, P., Sturley, S.L., Schaller, H., 2000. Expression in yeast and tobacco of plant cDNAs encoding acyl CoA: diacylglycerol acyltransferase. Eur. J. Biochem. 267, 85e96. Briassoulis, D., Panagakis, P., Chionidis, M., Tzenos, D., Lalos, A., Tsinos, C., Berberidis, K., Jacobsen, A., 2010. An experimental helical-tubular photobioreactor for continuous production of Nannochloropsis sp. Bioresour. Technol. 101, 6768e6777. Brown, A.C., Knights, B.A., Conway, E., 1969. Hydrocarbon content and its relationship to physiological state in the green alga Botryococcus braunii. Phytochemicals 8, 543e547. Brown, L., Zeiler, K.G., 1993. Aquatic biomass and carbon dioxide trapping. Energy Convers. Manage. 34, 1005e1013. Campbell, M.N., 2008. Biodiesel: algae as a renewable source for liquid fuel. Guelph Eng. J. 1, 1916-1107. Canakci, M., Gerpan, J.V., 1999. Biodiesel production via acid catalysis. Trans. ASAE 42, 1203e1210. Chinnasamy, S., Bhatnagar, A., Hunt, R.W., Das, D.C., 2010. Microalgae cultivation in a wastewater dominated by carpet mill effluents for biofuel applications. Bioresour. Technol. 101, 3097e3105. Chisti, Y., 2008. Biodiesel from microalgae beats bioethanol. Trends Biotechnol. 26, 126e131. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294e306. Chiu, S.Y., Kao, C.Y., Tsai, M.T., Ong, S.C., Chen, C.H., Lin, C.S., 2009. Lipid accumulation and CO2 utilization of Nannochloropsis oculata in response to CO2 aeration. Bioresour. Technol. 100, 833e838. Cohen, I., Neori, A., 1991. Ulva lactuca biofilters for marine fishpond effluents: 1. ammonia uptake kinetics and nitrogen content. Bot. Mar. 34, 475e482. Converti, A., Casazza, A.A., Ortiz, E.Y., Perego, P., Del Borghi, M., 2009. Effect of temperature and nitrogen concentration on the growth and lipid content of Nannochloropsis oculata and Chlorella vulgaris for biodiesel production. Chem. Eng. Process 48, 1146e1151. Courchesne, N.M.D., Parisien, A., Wang, B., Lan, C.Q., 2009. Enhancement of lipid production using biochemical, genetic and transcription factor engineering approaches. J. Biotechnol. 141, 31e41. Craggs, R.J., McAuley, P.J., Smith, V.J., 1997. Wastewater nutrient removal by marine microalgae grown on corrugated raceway. Water Res. 31, 1701e1707. Cromar, N.J., Fallowfield, H.J., Martin, N.J., 1996. Influence of environmental parameters on biomass production and nutrient

10011-GUPTA-9780444595614


12


removal in a high rate algal pond operated by continuous culture. Water Sci. Technol. 34, 133e140. Damiani, M.C., Cecilia, A.P., Diana, C., Patricia, I.L., 2010. Lipid analysis in Haematococcus pluvialis to assess its potential use as a biodiesel feedstock. Bioresour. Technol. 101, 3801e3807. Davis, M.S., Solbiati, J., Cronan, J.E., 2000. Overproduction of acetylCoA carboxylase activity increases the rate of fatty acid biosynthesis in Escherichia coli. J. Biol. Chem. 275, 28593e28598. Dayananda, C., Sarada, R., Usha Rani, M., Shamala, T.R., Ravishankar, G.A., 2007. Autotrophic cultivation of Botryococcus braunii for the production of hydrocarbons and exopolysaccharides in various media. Biomass Bioenergy 31, 87e93. Deviller, G., Aliaume, C., Nava, M.A.F., Casellas, C., Blancheton, J.P., 2004. High-rate algal pond treatment for water reuse in an integrated marine fish recirculating system: effect on water quality and sea bass growth. Aquaculture 235, 331e344. Dorado, M.P., Ballesteros, E., Almeida, J.A., Schellert, C., Lohrlein, H.P., Krause, R., 2002. An alkali-catalyzed transesterification process for high free fatty acid waste oils. Trans. ASAE 45, 525e529. Dumas, A., Laliberte´, G., Lessard, P., de la Nouë, J., 1998. Biotreatment of fish farm effluents using the cyanobacterium Phormidium bohneri. Aqua. Eng. 17, 57e68. Dunahay, T.G., Jarvis, E.E., Roessler, P.G., 1995. Genetic transformation of the diatoms Cyclotella cryptica and Navicula saprophila. J. Phycol. 31, 1004e1012. Dunahay, T.G., Jarvis, E.E., Dais, S.S., Roessler, P.G., 1996. Manipulation of microalgal lipid production using genetic engineering. Appl. Biochem. Biotechnol. 57, 223e231. Durrett, T., Benning, C., Ohlrogge, J., 2008. Plant triacylglycerols as feedstocks for the production of biofuels. Plant J. 54, 593e607. Eisenberg, M., Koopman, B., Benemann, J.R., Oswald, W.L., 1981. Algal bioflocculation and energy conservation in microalgal sewage ponds. Biotechnol. Bioeng. Symp. 11, 429e448. Feng, D., Chen, Z., Xue, S., Zhang, W., 2011b. Increased lipid production of the marine oleaginous microalgae Isochrysis zhangjiangensis (Chrysophyta) by nitrogen supplement. Bioresour. Technol. 102, 6710e6716. Feng, P., Deng, Z., Hu, Z., Fan, L., 2011a. Lipid accumulation and growth of Chlorella zofingiensis in flat plate photobioreactors outdoors. Bioresour. Technol. 102, 10577e10584. Francis, G., Becker, K., 2002. Biodiesel from Jatropha plantations on degraded land. University of Hohenheim, Stuttgart, Germany p 9. http://www.youmanitas.nl/pdf/Bio-diesel.pdf. Gao, C., Zhai, Y., Ding, Y., Wu, Q., 2010. Application of sweet sorghum for biodiesel production by heterotrophic microalga Chlorella protothecoides. Appl. Energy 87, 756e761. Gates, W.E., Borchardt, J.A., 1964. Nitrogen and phosphorus extraction from domestic waste water treatment plant effluents by controlled algal culture. Res. J. Water Pollut. Control Fed. 36, 443e462. Ghadge, S.V., Raheman, H., 2006. Process optimization for biodiesel production from mahua (Madhuca indica) oil using response surface methodology. Bioresour. Technol. 97, 379e384. Goering, C.E., Schwab, A.W., Daugherty, M.J., Pryde, E.H., Heakin, A.J., 1982. Fuel properties of eleven vegetable oils. Trans. ASAE 25, 1472e1477. Gonza´lez, L.E., Can˜izaresb, R.O., Baenaa, S., 1997. Efficiency of ammonia and phosphorus removal from a colombian agroindustrial wastewater by the microalgae Chlorella vulgaris and Scenedesmus dimorphus. Bioresour. Technol. 60, 259e262. Gouveia, L., Oliveira, A.C., 2009. Microalgae as a raw material for biofuels production. J. Ind. Microbiol. Biotechnol. 36, 269e274. Gouveia, L., Marques, A.E., da Silva, T.L., Reis, A., 2009. Neochloris oleoabundans UTEX #1185: a suitable renewable lipid source for biofuel production. J. Ind. Microbiol. Biotechnol. 36, 821e826.

Grobbelaar, J.U., 2009. Upper limits of photosynthetic productivity and problems of scaling. J. Appl. Phycol. 21, 519e522. Grobbelaar, J.U., 2012. Microalgae mass culture: the constraints of scaling-up. J. Appl. Phycol. 24, 315e318. Haag, A.L., 2007. Algae bloom again. Nature 447, 520e521. Hankamer, B., Lehr, F., Rupprecht, J., Mussgnug, J.H., Posten, C., Kruse, O., 2007. Photosynthetic biomass and H2 production by green algae: from bioengineering to bioreactor scale-up. Physiol. Plant. 131, 10e21. Hossain, A.B.M.S., Salleh, A., 2008. Biodiesel fuel production from algae as renewable energy. Am. J. Biochem. Biotechnol. 4, 250e254. Illman, A.M., Scragg, A.H., Shales, S.W., 2000. Increase in Chlorella strains calorific values when grown in low nitrogen medium. Enzyme Microb. Technol. 27, 631e635. International Energy Agency, 2007. World Energy Outlook. Executive Summary, China and India insight. Paris, France, pp.14. Jain, S., Sharma, M.P., 2010. Prospects of biodiesel from Jatropha in India: a review. Renewable Sustainable Energy Rev. 14, 763e771. Jako, C., Kumar, A., Wei, Y., Zou, J., Barton, D.L., Giblin, E.M., Covello, P.S., Taylor, D.C., 2001. Seed-specific over-expression of an Arabidopsis cDNA encoding a diacylglycerol acyltransferase enhances seed oil content and seed weight. Plant Physiol. 126, 861e874. Jiang, Y., Yoshida, T., Quigg, A., 2012. Photosynthetic performance, lipid production and biomass composition in response to nitrogen limitation in marine microalgae. Plant Physiol. Biochem. 54, 70e77. Jimenez del Rio, M., Ramazanov, Z., Garcia-Reina, G., 1996. Ulva rigida (Ulvales, Chlorophyta) tank culture as biofilters for dissolved inorganic nitrogen from fishpond effluents. Hydrobiologia 326, 61e66. Jimenez-Perez, M.V., Sanches-Castillo, P., Romera, O., FernandezMoreno, D., Perez-Martinez, C., 2004. Growth and nutrient removal in free and immobilized planktonic green algae isolated from pig manure. Enzyme Microb. Technol. 34, 392e398. Johnson, M.B., Wen, Z., 2009. Production of biodiesel fuel from the microalga Schizochytrium limacinum by direct transesterification of algal biomass. Energy Fuels 23, 5179e5183. Kenesey, E., Ecker, A., 2003. Oxygen bond to improve the lubricity of fuel. Tribol. Schmierungstech. 50, 21e26. Khan, S.A., Rashmi, M.Z., Hussain, S., Prasad, Banerjee, U.C., 2009. Prospects of biodiesel production from microalgae in India. Renewable Sustainable Energy Rev. 13, 2361e2372. Khozin-Goldberg, I., Cohen, Z., 2006. The effect of phosphate starvation on the lipid and fatty acid composition of the fresh water eustigmatophyte Monodus subterraneus. Phytochemicals 67, 696e701. Kilham, S.S., Kreeger, D.A., Gulden, C.E., Lynn, S.G., 1997. Effects of nutrients limitation on biochemical constituents of Ankistrodesmus falcatus. Freshwater Biol. 38, 591e596. Klaus, D., Ohlrogge, J.B., Neuhaus, H.E., Dormann, P., 2004. Increased fatty acid production in potato by engineering of acetyl-CoA carboxylase. Planta 219, 389e396. Knothe, G., 2005. Dependence of biodiesel fuel properties on the structure of fatty acid alkyl esters. Fuel Process. Technol. 86, 1059e1070. Knothe, G., Dunn, R.O., 2003. Oxidative stability of biodiesel/jet fuel blends by oil stability index (OSI) analysis. J. Am. Oil Chem. Soc. 80, 1047e1048. Krawczyk, T., 1996. Biodiesel alternative fuel makes inroads but hurdles remain. Inform 7, 801e829. Kumar, A., Kumar, K., Kaushik, N., Sharma, S., Mishra, S., 2010. Renewable energy in India: current status and future potentials. Renewable Sustainable Energy Rev. 14, 2434e2442. Laliberte, G., Lessard, P., de la Nouë, J., Sylvestre, S., 1997. Effect of phosphorus addition on nutrient removal from wastewater with the cyanobacterium Phormidium bohneri. Bioresour. Technol. 59, 227e233.

10011-GUPTA-9780444595614


REFERENCES

Lang, X., Dalai, A.K., Bakhshi, N.N., Reany, M.J., Hertz, P.B., 2001. Preparation and characterization of biodiesel from various bio-oils. Bioresour. Technol. 80, 53e62. Lee, Y.-K., 2001. Microalgal mass culture systems and methods: their limitation and potential. J. Appl. Phycol. 13, 307e315. Lee, K., Lee, C.G., 2001. Effect of light/dark cycles on wastewater treatments by microalgae. Biotechnol. Bioprocess Eng. 6, 194e199. Li, X., Xu, H., Wu, Q., 2007. Large-scale biodiesel production from microalga Chlorella protothecoides through heterotrophic cultivation in bioreactors. Biotechnol. Bioeng. 98, 764e771. Li, Y., Horsman, M., Wu, N., Lan, C.Q., Dubois-Calero, N., 2008. Biofuels from microalgae. Biotechnol. Prog. 24, 815e820. Lin, H., Castro, N.M., Bennett, G.N., San, S.Y., 2006. Acetyl-CoA synthetase overexpression in Escherichia coli demonstrates more efficient acetate assimilation and lower acetate accumulation: a potential tool in metabolic engineering. Appl. Microbiol. Biotechnol. 71, 870e874. Liu, J., Hua, W., Zhan, G., Wei, F., Wang, X., Liu, G., Wang, H., 2010. Increasing seed mass and oil content in transgenic Arabidopsis by the overexpression of wri1-like gene from Brassica napus. Plant Physiol. Biochem. 48, 9e15. Lu, X., Vora, H., Khosla, S., 2008. Over production of free fatty acids in E. coli: implications for biodiesel production. Metab. Eng. 10, 333e339. Mallick, N., 2002. Biotechnological potential of immobilized algae for wastewater N, P and metal removal: a review. BioMetals 15, 377e390. Mallick, N., Mandal, S., Singh, A.K., Bishai, M., Dash, A., 2012. Green microalga Chlorella vulgaris as a potential feedstock for biodiesel. J. Chem. Technol. Biotechnol. 87, 137e145. Mandal, S., Mallick, N., 2009. Microalga Scenedesmus obliquus as a potential source for biodiesel production. Appl. Microbiol. Biotechnol. 84, 281e291. Mandal, S., Mallick, N., 2011. Waste utilization and biodiesel production by the green microalga Scenedesmus obliquus. Appl. Environ. Microbiol. 77, 374e377. Mandal, S., Mallick, N., 2012. Biodiesel production by the green microalga Scenedesmus obliquus in a recirculatory aquaculture system. Appl. Environ. Microbiol. 78, 5929e5934. Marsh, G., 2009. Small wonders: biomass from algae. Renewable Energy Focus 9, 74e78. Martinez, M.E., Castillo, J.M., El Yousfi, F., 1999. Photoautotrophic consumption of phosphorus by Scenedesmus obliquus in a continuous culture. Influence of light intensity. Process Biochem. 34, 811e818. Martinez, M.E., Sanchez, S., Jimenez, J.M., El Yousfi, F., Munoz, L., 2000. Nitrogen and phosphorus removal from urban wastewater by the microalga Scenedesmus obliquus. Bioresour. Technol. 73, 263e272. McMillen, S., Shaw, P., Jolly, N., Goulding, B., Finkle, V., 2005. Biodiesel: Fuel for Thought, Fuel for Connecticut’s Future. Connecticut Center for Economic Analysis Report. University of Connecticut, pp. 56. Meher, L.C., Vidya Sagar, D., Naik, S.N., 2006. Technical aspects of biodiesel production by transesterification - a review. Renewable Sustainable Energy Rev. 10, 248e268. Miao, X., Wu, Q., 2004. High yield bio-oil production from fast pyrolysis by metabolic controlling of Chlorella protothecoides. J. Biotechnol. 110, 85e93. Miao, X., Wu, Q., 2006. Biodiesel production from heterotrophic microalgal oil. Bioresour. Technol. 97, 841e846. Mittelbach, M., 1990. Lipase-catalyzed alcoholysis of sunflower oil. J. Am. Oil Chem. Soc. 67, 168e170. Mittelbach, M., 1996. Diesel fuel derived from vegetable oils, VI: specifications and quality control of biodiesel. Bioresour. Technol. 27, 435e437.

13

Molina, E., Fernańdez, A.F.G., Camacho, G.F., Rubio, C.F., Chisti, Y., 2000. Scale-up of tubular photobioreactors. J. Appl. Phycol. 12, 355e368. Olguıń, E.J., Galicia, S., Mercado, G., Perez, T., 2003. Annual productivity of Spirulina (Arthrospira) and nutrient removal in a pig wastewater recycle process under tropical conditions. J. Appl. Phycol. 15, 249e257. Oswald, W.J., Gotaas, H.B., Ludwig, H.F., Lynch, W., 1953. Algae symbiosis in oxidation ponds: III. phototosynthetic oxygenation. Sewage Ind. Wastes 25, 692e704. Patil, P.D., Gude, V.G., Mannarswamy, A., Deng, S., Cooke, P., Munson-McGee, S., Rhodes, I., Lammers, P., Nirmalakhandan, N., 2012. Optimization of direct conversion of wet algae to biodiesel under supercritical methanol conditions. Bioresour. Technol. 101, 118e122. Piorreck, M., Pohl, P., 1984. Formation of biomass, total protein, chlorophylls, lipids and fatty acids in green and blue-green algae during one growth phase. Phytochemicals 23, 217e223. Planning Commission, 2003. Report of the Committee on Development of Biofuel. Government of India, New Delhi, India, pp. 140. Planning Commission, 2007. Eleventh Five Year Plan 2007e2012, Energy Sector. Government of India, New Delhi, India, pp. 65. Qian, P., Wu, C.Y., Wu, M., Xie, Y.K., 1996. Integrated cultivation of the red alga Kappaphycus alvarezii and the pearl oyster Pinctada martensi. Aquaculture 147, 21e35. ´ ., Ramos, M.J., Fernańdez, C.M., Casas, A., Rodrı´guez, L., Pe´rez, A 2009. Influence of fatty acid composition of raw materials on biodiesel properties. Bioresour. Technol. 100, 261e268. Rao, P.P., Gopalkrishnan, K.V., 1991. Vegetable oils and their methyl esters as fuels diesel engines. Indian J. Technol. 29, 292e297. Ratledge, C., 2002. Regulation of lipid accumulation in oleaginous micro-organisms. Biochem. Soc. Trans. 30, 1047e1050. Rattanapoltee, P., Chulalaksananukul, W., James, A.E., Kaewkannetra, P., 2008. Comparison of autotrophic and heterotrophic cultivations of microalgae as a raw material for biodiesel production. J. Biotechnol. 136, S412. Rodolfi, L., Zittelli, G., Bassi, N., Padovani, G., Bonini, N., Bonini, G., Tredici, M.R., 2009. Microalgae for oil: strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnol. Bioeng. 102, 100e112. Roesler, K., Shintani, D., Savage, L., Boddupalli, S., Ohlrogge, J., 1997. Targeting of the Arabidopsis homomeric acetyl-coenzyme A carboxylase to plastids of rapeseeds. Plant Physiol. 113, 75e81. Roessler, P.G., 1990. Purification and characterization of acetyl-CoA carboxylase from the diatom Cyclotella cryptica. Plant Physiol. 92, 73e78. Russa, L.M., Bogen, C., Uhmeyer, A., Doebbe, A., Filippone, E., Kruse, O., Mussgnug, J.H., 2012. Functional analysis of three type2 DGAT homologue genes for triacylglycerol production in the green microalga Chlamydomonas reinhardtii. J. Biotechnol. 162, 13e20. Sarkar, S., Kamilya, D., Mal, B.C., 2007. Effect of geometric and process variables on the performance of inclined plate settlers in treating aquacultural waste. Water Res. 41, 993e1000. Schneider, D., 2006. Grow your own? Would the widespread adoption of biomass-derived transportation fuels really help the environment. Am. Sci. 94, 408e409. Schuenhoff, A., Shpigel, M., Lupatsch, I., Ashkenazi, A., Msuya, F.E., Neori, A., 2003. A semi-recirculating, integrated system for the culture of fish and seaweed. Aquaculture 221, 167e181. Scragg, A.H., Illman, A.M., Carden, A., Shales, S.W., 2002. Growth of microalgae with increased calorific values in a tubular bioreactor. Biomass Bioenergy 23, 67e73. Shay, E., 1993. Diesel fuel from vegetable oils: status and opportunities. Biomass Bioenergy 4, 227e242.

10011-GUPTA-9780444595614


14


Sheehan, J., Dunahay, T., Benemann, J., Roessler, P., 1998. A Look Back at the U.S. Department of Energy’s Aquatic Species Program Biodiesel from Algae. National Renewable Energy Laboratory Report. U.S. Department of Energy, pp. 323. Singh, I., Rastogi, V., 2009. Performance analysis of a modified 4-stroke engine using biodiesel fuel for irrigation purpose. Int. J. Environ. Sci. 4, 229e242. Sobczuk, T.M., Chisti, Y., 2010. Potential fuel oils from the microalga Choricystis minor. J. Chem. Technol. Biotechnol. 85, 100e108. Takagi, M., Karseno, Yoshida, T., 2006. Effect of salt concentration on intracellular accumulation of lipids and triacylglyceride in marine microalgae Dunaliella cells. J. Biosci. Bioeng. 101, 223e226. Takagi, M., Watanabe, K., Yamaberi, K., Yoshida, T., 2000. Limited feeding of potassium nitrate for intracellular lipid and triglyceride accumulation of Nannochloris sp. UTEX LB1999. Appl. Microbiol. Biotechnol. 54, 112e117. Umdu, E.S., Tuncer, M., Seker, E., 2009. Transesterification of Nannochloropsis oculata microalga’s lipid to biodiesel on Al2O3

supported CaO and MgO catalysts. Bioresour. Technol. 100, 2828e2831. Velasquez-Orta, S.B., Lee, J.G.M., Harvey, A., 2012. Alkaline in situ transesterification of Chlorella vulgaris. Fuel 94, 544e550. Xu, H., Miao, X., Wu, Q., 2006. High quality biodiesel production from a microalgae Chlorella protothecoides by heterotrophic growth in fermenters. J. Biotechnol. 126, 499e507. Zhang, Y., Adams, I.P., Ratledge, C., 2007. Malic enzyme: the controlling activity for lipid production? Overexpression of malic enzyme in Mucor circinelloides leads to a 2.5-fold increase in lipid accumulation. Microbiology 153, 2013e2025. Zhang, Z., Li, M., Agarwal, A., San, K.-Y., 2011. Efficient free fatty acid production in Escherichia coli using plant acyl-ACP thioesterases. Metab. Eng. 13, 713e722. Zou, J., Katavic, V., Giblin, E.M., Barton, D.L., MacKenzie, S.L., Keller, W.A., Hu, X., Taylor, D.C., 1997. Modification of seed oil content and acyl composition in the Brassicaceae by expression of a yeast sn-2 acyltransferase gene. Plant Cell 9, 909e923.

10011-GUPTA-9780444595614

GUPTA: 11 Non-Print Items

Abstract: Biodiesel production from microalgae is potentially feasible because these tiny photosynthetic microbes offer a combination of high biomass productivity and lipid content. Over the past few decades, several microalgal species have been screened for high lipid content. Nutrient starvation has so far been the most commonly employed strategy for improving lipid accumulation in microalgae. The properties of biodiesel are largely determined by the structure of its component fatty acid esters. Depending on the growth conditions, microalgae, even the same strain, can exhibit large variations in fatty acid composition. Biodiesel containing saturated and monounsaturated fatty acids exhibits improved fuel properties. The major bottleneck for producing microalgal biodiesel is the huge consumption of water resources in addition to inorganic nutrients for large-scale cultivation. One promising approach is to couple biodiesel production with wastewater treatment as algae can successfully be cultivated in wastewaters. Here, a case study with Scenedesmus obliquus in a recirculatory aquaculture system using fish pond discharge and poultry litter for biodiesel production has been briefly included. Keywords: Biodiesel, Lipids, Microalgae, Recirculatory aquaculture system (RAS), Waste discharges.