Biofuels derived from microalgae are currently considered to be the most economi­cal and technically viable route for producing biofuels to compete with petroleum — based fuels [40] . This is due to a number of intrinsic advantages they could have, when compared to biofuels extracted from terrestrial bioenergy crops. These include: (1) higher annual growth rates, e. g. rates of up to 37 tonnes ha-1 per annum have been recorded, primarily due to higher photosynthetic efficiencies when compared with terrestrial plants [221]; (2) higher lipid productivity (up to 75% dry weight for some algae species), with higher proportion of triacylglycerol (TAG) that is essential for efficient biodiesel production [185]; (3) microalgae production can effect biofixation of CO2 (production utilises about 1.83 kg of CO2 per kg of dry algal biomass yield) thus making a contribution to air quality improvement [38]; (4) capability of growing in wastewater, which offers the duel potential for integrating the treatment of organic effluent with biofuel production [28]; and (5) inherent yield of valuable co-products such as proteins and polyunsaturated fatty acids (PUFAs) may be used to enhance the economics of production systems [197]. Figure 2 summarises the distinct stages in the production and processing of microalgae to biofuel.

Despite the outlined advantages, there are still a number of significant obstacles to realisation of the full potential of microalgae-derived biofuels. They include: (1) energy-intensive downstream processes including pumping and dewatering of bio­mass, and conversion processes can result in negative energy balance [ 88] ; (2) trade-off requirements in species selection and genetic enhancement for biofuel production vs. extraction of co-products [ 157 ] ; and (3) need for techniques for enabling real-time control of pH, photoinhibition, evaporation and CO. diffusion losses in cultivation systems [213].

Previous research has shown that a 400°C pyrolysis process produced a preferred biochar for agriculture and ammonia adsorption [3]. A 400°C biochar sample was produced by Day using pyrolysis of pelletized peanut hulls at 400°C. Specifically, the cross-draft reactor was brought up to 400°C empty with the exhaust from natural gas burner. Then, pelletized peanut hulls (biomass) were slowly fed in maintaining that internal reactor temperature. Once the biochar discharge sensor reached 400°C, the rotary discharge valve released biochar, the burner was switched off and the

system operated without any air or outside heat. This point coincides near the end of the exothermic zone for peanut shells. It also corresponds with the temperature range at which the resulting biochar had an increased fertilizer binding capacity as previously reported [3, 4]. The feed rate was controlled by the automated discharge of the biochar reaching 400°C and averaged 5-7 kg h-1 . The exothermic reaction allowed continuous feed to the pyrolysis reactor; however, the feed rate was 20% of what occurs when combustion of part of the pyrolysis vapors was used to augment the natural exothermic reaction.

In addition to the above chemicals, many other chemicals can be produced by selective fast pyrolysis of biomass. For example, Chen et al. [18] reported that fast pyrolysis of biomass impregnated with Na2 CO3 produced HA with high purity. Badri [10] revealed that catalytic pyrolysis of cotton with reactive dyes favored the formation of 5-hydroxymethyl-furfural (HMF) and 3-(hydroxymethyl)-furan. Lu et al. [51] found that fast pyrolysis of cellulose followed with catalytic cracking of the vapors by sulfated metal oxides could obtain high yields of furan and 5-methyl furfural. In another study, Lu et al. [52] confirmed that catalytic cracking of the biomass fast pyrolysis vapors using ZrO2 and TiO2 increased the formation of three light carbo­nyl products (acetaldehyde, acetone, and 2-butanone).

2 Conclusion

Most of the selective fast pyrolysis techniques are only in their early stage of devel­opment, and none of the techniques is commercially practical to produce specific chemicals in marketable quantities at present. Three aspects should be considered for the commercialization of the selective fast pyrolysis techniques, including (1) the technique to produce specific chemicals in high yield and purity, (2) the method to recover the target chemicals from pyrolysis liquids, and (3) the ready markets for the chemicals.

Among the above indicated chemicals, the LG, HAA, and AA can be produced without catalyst utilization, and thus, their large-scale production might be rela­tively easy to achieve through slight modification of the conventional fast pyrolysis technique. The production of other chemicals requires catalysts, which will add difficulty to their scale up. Various methods have been proposed for the chemicals recovery, and further studies should be conducted to reduce the purification cost. Finally, it is important to note that currently there are no existing markets for the LG, LGO, LAC, anhydro-oligosaccharides, and some other chemicals. Corresponding markets should be developed by manufacturers who would incorporate these chemi­cals into various products.

A process for converting fossil fuels such as crude oil, heavy oil, coal, shale oil, tar sand, bitumen, etc., or biomass to liquid fuels such as methanol, biogasoline, biod­iesel, or biojet fuel, etc., generally contain five steps: pretreatment, gasification, gas cleaning and conditioning, FT process, and final product separation and possible upgrading. This section outlines some details of each of these steps with special attention to biomass feedstock.

2.1 Pretreatment

The first step is the pretreatment of fossil fuel or biomass. Here undesired impurities such as mineral matters, inorganic impurities (particularly in biomass), sulfur, etc., are removed and particle size of the feed stock is adjusted to satisfy the need of the subsequent gasification process. Pretreatment of coal requires removal of ash and sulfur along with adjustment of the particle size. The inorganic sulfur and ash are often removed by the floatation technique supported by the fine particle technology. Pretreatment step is particularly important for biomass. Unlike coal, biomass is a low density (both mass and energy) product and easily degradable in the natural environment. The storage and transportation of biomass are very problematic because of its low density and easy degradation. The reduction of biomass particle size to the desired level is also problematic because it requires high grinding energy. Biomass is generally wet and contains a large amount of water. The removal of this water is also important for high energy efficiency of the gasifier.

Numerous pretreatment processes for biomass have been examined. The advan­tages and disadvantages of these processes are outlined in Table 6.

For combustion and gasification, biomass particle size, water content, decay resistance, and inorganic impurities are important parameters. For these reasons, leaching and torrefaction are considered to be the two most important pretreatment steps. Leaching process will remove salts and other inorganic impurities from the biomass and can be easily carried out using hot water or steam. Torrefaction process makes biomass hydrophobic, easy to crush, and improve its resistance for decay. A typical comparison of grinding energy required for untreated and torrefied wood is illustrated in Fig. 2. This comparison clearly indicates the important role torrefac­tion can play in reducing the grinding energy of biomass. Torrefied biomass also has higher mass density and it is resistant to water and biological decay. These param­eters help the transportation and storage of biomass. The positive features of torrified biomass for gasification are given in Table 7.

In Part 1.2, it is suggested that to use CO. — containing syngas directly for FT synthesis can solve the shortcoming of current FT technology. The studies of CO2 hydrogenation and CO. + CO hydrogenation on Fe catalysts are reviewed in the following.

In view of engineering, to use CO2-containing syngas is easy to control temperature of synthesis reactor because of the lower exothermicity of the overall reaction of CO2 as compared to CO [ 16] . Furthermore, a perceived disadvantage of using Fe-based catalysts for FT is that a large proportion of the CO in the syngas is con­verted to CO. rather than the desired hydrocarbons [21]. The addition of CO2 to syngas can inhibit CO2 formation from CO and may increase the ratio of CO to hydrocarbons rather than CO2.

In an interesting investigation, it was observed that some of the cellulosic hydrolyzate chemicals that are generated during the pretreatment or the neutralization process stimulate both cell growth and ABE production. These chemicals include sodium acetate, furfural, and HMF. In the presence of 8.9 g/L sodium acetate cell growth was slightly improved. In the control experiment 17.8 g/L ABE was produced while in the presence of 8.9 g/L acetate 20.3 g/L ABE was produced, showing an increase of 14%. An improvement in fermentation performance on supplementation of acetate has previously been documented [11,24]. Inclusion of furfural and HMF (0.3-2.0 g/L) in the fermentation medium improved both cell concentration and ABE production [18]. Itis suggested that acetate, furfural, and HMF are beneficial to this fermentation within a certain concentration range.

Plant cell walls are composed of several distinct chemical polymers arranged as a composite network. Their structural rigidity minimizes water loss and protects against various biotic and abiotic stresses. The plasticity of the plant cell wall allows cell elongation, plant growth and enables reorganization that accommodates environmental changes [18, 107, 120]. The construction and breakdown of the cell wall is complex, involving a large number of glycosyl synthases, hydrolytic and disruptive enzymes that act at different developmental stages in various cellular organelles. A thorough understanding of the synthesis and breakdown of the cell wall will enable the development of techniques for effective manipulation of cell wall polymers for the production of biofuel. Recent reviews cover synthesis, composition, and remodeling of plant cell walls [18, 53, 105]; the fundamentals are outlined below.

The cell wall is generally divided into primary wall and secondary wall. Following cell division, the middle lamella and primary wall are formed. The primary wall determines the cell’s shape and size and it basically consists of cellulose, hemicel — luloses, pectin, structural proteins, and phenolic compounds. The amount and ratio between these components are highly depended on the primary wall type. While cellulose, or (1-4)-b-linked glucose, is the most abundant polysaccharide on earth, control of its biosynthesis is not completely understood. Cellulose synthesis occurs in the plasma membrane by hexagonal-shaped complexes termed rosettes, each containing 36 cellulose synthase modules (CesA) [3, 32]. A highly compact lattice of microfibrils is assembled and linked via hydrogen bonds that form crystalline cellulose. These microfibrils serve as a scaffold for the deposition of other wall components, such as hemicellulose and pectin [84, 119] . In contrast to cellulose, hemicellulose and pectin synthesis occurs in the Golgi bodies [94]. Hemicellulosic polysaccharides are complex and heterogeneous molecules which crosslink neigh­boring cellulose microfibrils. Cell walls of dicots or type I primary walls consist of equal amounts of xyloglucans and cellulose that are embedded in a rich pectin matrix and are further crosslinked with structural proteins. On the other hand, type II walls or cell walls of poaceae (grasses) and gymnosperms contain arabinoxylan instead of xyloglucan. Moreover, this kind of wall does not contain large amounts of pectin but contains phenolics that are crosslinked with arabinoxylan, thus strengthening the wall [18, 70, 90, 126].

Secondary wall deposition occurs in most plant cells and accounts for most of the cell wall mass. Secondary cell walls are composed of cellulose, hemicelluloses (mostly xylans), and lignin and include several layers that vary between species in the composition and ratio of components and in the arrangement of the cellulose microfibrils. The key differences between the secondary cell walls of distinct plant families lie in the quantity and nature of their hemicelluloses and Lignins. Lignins are the product of an oxidative combinatorial coupling of 4-hydroxyphenylpropanoids, principally involving the hydroxycinnamyl alcohols, coniferyl alcohol, sinapyl alcohol, and small amounts of p-coumaryl alcohol. These monolignols differ in their degree of aromatic methoxylation. Following coupling, the resulting units in the lignin polymer are the guaiacyl (G), syringyl (S), and p-hydroxyphenyl (H) units, respectively [100, 123, 124]. S lignin is unique to flowering plants while H and G lignins are fundamental to all vascular plants [131].

The hemicelluloses of poaceae and gymnosperms mainly comprises large amounts of glucuronoarabinoxylan (GAX) and galactoglucomannan in their sec­ondary walls while woody angiosperms mostly contain glucuronoxylan (GX). GAX contains many arabinose residues which can be further esterified with ferulic acid. The hemicelluloses in the secondary wall are crosslinked with lignin and cellulose. The interactions of hemicelluloses with other components in the secondary cell wall is stronger than in the primary cell wall due to the presence of fewer side chains in secondary wall hemicellulose [126]. In summary, cellulose crystallinity, degree of polymerization, hemicellulose species, and lignin content all influence cell wall resistance to enzymatic degradation and these constraints differ between different plant varieties.

One of the most common approaches to measuring cellulase activity involves monitoring spectroscopic changes resulting from product formation or substrate depletion. Some of these assays involve covalent attachment of a probe that changes its absorbance (or fluorescence) when released from cellulose. Other assays involve the addition of colorimetric reagents that react with product to produce an absorbance change. The easiest of these assays takes advantage of the reactivity of the reducing end of cellulose. The reducing end is in equilibrium with an aldehyde, which can reduce a chromogenic substrate. Each time a glycosidic bond is hydrolyzed, a new reducing end is produced, regardless of enzyme type (endo — vs. exocellulase vs. b-glucosidase).

Two methods to detect reducing end formation are the dinitrosalicylic acid (DNS) and the Nelson-Somogyi assays. In the DNS assay, oxidation of the reducing-end aldehyde to a carboxylic acid is coupled to the reduction of DNS to 3-amino-5- nitrosalicylic acid [45] [19]. Because this redox reaction requires elevated tempera­tures (typically 100°C) and high pH, the color must be developed as an end-point to the reaction, limiting the degree to which time-course of the reaction can be sam­pled. Another drawback in the DNS assays is a rather low sensitivity, due in part to the high absorbance of unreacted DNS. Sensitivity can also be adversely affected by buffer conditions and the presence of metals, and in some cases, the degree of cel­lulose polymerization [59] .

In the Nelson-Somogyi assay, reducing sugar oxidation is coupled to a Cu2+ reduc­tion to Cu1+. In a subsequent step, Cu1+ is oxidized back to Cu2+ by an arsenomolyb — date complex that becomes colored upon reduction [66]. Unlike the DNS method, the Nelson-Somogyi method depends neither on the substrate concentration nor on the degree of polymerization. However the Nelson-Somogyi assay is also sensitive to composition and to the nature of the substrate. The Nelson-Somogyi assay has higher sensitivity than the DNS assay, but is more cumbersome [ 6] . Both the DNS and Nelson-Somogyi assays can be performed on a variety of cellulosic substrates, includ­ing soluble substrates and insoluble amorphous and crystalline material. A number of additional colorimetric assays have been used to monitor cellulase activity [81].

Direct labeling methods to monitor cellulase activity include labeling of long — chain insoluble cellulose polymers with dyes such as azure [38] and fl uorescein; hydrolysis is monitored by determining the amount of dye released into the soluble phase. By casting dyed cellulose films on the bottom of microtiter plates, this approach can be used to screen activity in a high-throughput format [25] . Short, water-soluble cellodextrins can also be labeled with spectroscopic probes (e. g., p-nitrophenyl [78]; 4-methylumbelliferyl [43]). These short-chain derivatives are useful for measuring b — glucosidase activity. b — glucosidase activity can also be assayed by monitoring the formation of free glucose, using coupled enzyme detec­tion [83]; colorimetric assay kits are commercially available.

In prokaryotic photosynthetic organisms such as blue-green algae (oxyphotobacte — ria including cyanobacteria and oxychlorobacteria), which typically contain photosynthetic thylakoid membrane but no chloroplast structure, the Calvin cycle is located in the cytoplasm. In this special case, the entire designer butanol-production pathway(s) (Fig. 1) including (but not limited to) the glyceraldehyde-3-phosphate — branched butanol-production pathway (01-12) , the 3-phosphpgly cerate-branched butanol-production pathway (03-12), the fructose-1,6-diphosphate-branched path­way (20-33), the fructose-6-phosphate-branched pathway (19-33), and the starch (or glycogen)-to-butanol pathways (17-33) are adjusted in design to operate with the Calvin cycle in the cytoplasm of a blue-green alga. The construction of the cyto­plasm designer butanol-production pathways can be accomplished by use of designer butanol-production pathway genes (DNA construct of Fig. 2a) with their chloroplast-targeting sequence all omitted. When the chloroplast-targeting sequence is omitted in the designer DNA construct(s) as illustrated in Fig. 2e, the designer gene(s) is transcribed and translated into designer enzymes in the cytoplasm whereby conferring the designer butanol-production pathway(s). The designer gene(s) can be incorporated into the chromosomal and/or plasmid DNA in host blue-green algae (oxyphotobacteria including cyanobacteria and oxychlorobacteria) by using the techniques of genetic transformation known to those skilled in the art. It is a preferred practice to integrate the designer genes through an integrative transformation into the chromosomal DNA that can usually provide better genetic stability for the designer genes. In oxyphotobacteria such as cyanobacteria, integrative transforma­tion can be achieved through a process of homologous DNA double recombination into the host’s chromosomal DNA using a designer DNA construct as illustrated in Fig. 2f, which typically, from the 5′ upstream to the 3′ downstream, consists of: recombination site 1, a designer butanol-production-pathway gene(s), and recombi­nation site 2. This type of DNA constructs (Fig. 2f) can be delivered into oxyphoto — bacteria (blue-green algae) with a number of available genetic transformation techniques including electroporation, natural transformation, and/or conjugation. The transgenic designer organisms created from blue-green algae are also called designer blue-green algae (designer oxyphotobacteria including designer cyanobac­teria and designer oxychlorobacteria). Examples of designer oxyphotobacterial butanol-production-pathway genes are shown in SEQ ID NO: 34-45 listed in PCT Patent Application Publication No. WO 09105733. Recently, certain independent studies [27, 28] have also applied synthetic biology in certain model cyanobacteria such as S. elongatus PCC7942 for photobiological production of isobutanol and 1-butanol. According to another independent study [29 ] for this type of direct photosynthesis-to-biofuel process, the practical maximum solar-to-biofuel energy — conversion efficiency could be about 7.2% while the theoretical maximum solar-to — biofuel energy-conversion efficiency is calculated to be 12%.

Interest in using lipases as enzymatic catalyst for the production of alkyl fatty acid esters continuously grows. Some people work on the triglyceride by converting them to methyl esters, while some work on fatty acids. One of the main obstacles to the biocatalytic production of biodiesel is high cost of the enzyme; enzyme recy­cling might be the solution to this problem.

It was found that Pseudomonas fluorescens lipase immobilized on kaolinite lost one third of its activity when it is used for the second time, but no further decrease was observed in successive applications. The initial decrease in activity was put down to enzyme desorption from the solid support that was not observed after repeated (ten times) use [36 ] . Repeated batch reactions revealed that Rhizomucor miehei lipase had high stability, which retained about 70% of its initial conversion after 8 cycles (24 h each cycle). Meanwhile, under the same experimental conditions, Thermomyces lanuginosa retained only 35% of the initial conversion. This differ­ence was credited to factors such as inactivation of the biocatalyst in the oil phase, the type of carrier which is used for the immobilization, or enzyme sensitivity to long-term methanol exposure [79].

It is observed that various substances can slow down lipase activity (methanol, glycerol, phospholipids); however, a number of ways have been proposed to over­come these problems [71]. Commonly, biocatalytic process does not produce soaps or other by-products. If the reaction completes, only esters and glycerol are pro­duced. This makes purification steps much simpler and consequentially lowers plant costs. Once immobilized, lipases can be used many times or even in continuous processes. This resolves the major disadvantage related to high cost. In conclusion, if obtained by biocatalysis, biodiesel is an environmentally friendly fuel which will contribute to reducing negative impacts to the environment.

Modi et al. [56] used Propan-2-ol as an acyl acceptor for immobilized lipase- catalyzed preparation of biodiesel. The optimum conditions set for transesterification of crude Jatropha oil were 10% Novozym 435 (immobilized Candida antarctica lipase B) based on oil weight and alcohol to oil molar ratio of 4:1 at 50°C for 8 h. The maximum conversion reached using propan-2-ol was 92.8% from crude Jatropha oil. Reusability of the lipase was preserved over 12 repeated cycles with propan-2-ol as it reached to zero by the seventh cycle when methanol was utilized as an acyl acceptor, under standard reaction condition [55] .

Modi et al. [55, 57] explored ethyl acetate as an acyl acceptor for immobilized lipase-catalyzed preparation of biodiesel from the crude oil of J. curcas. The opti­mum reaction conditions set for interesterification of the oils with ethyl acetate were 10% of Novozym 435 (immobilized C. antarctica lipase B) which is based on oil weight, ethylacetate to oil molar ratio of 11:1, and the reaction period of 12 h at 50°C. Under the above optimum conditions, the maximum result of ethyl esters was 91.3% of purity. Reusability of the lipase over repeated cycles in interesterification and ethanolysis was also examined under standard reaction conditions. The relative activity of lipase could be well preserved over 12 repeated cycles with ethyl acetate as it reached to zero by the sixth cycle when ethanol was used as an acyl acceptor.

Shah and Gupta [73] conducted a process of optimization of monoethyl esters of the long chain fatty acids (biodiesel) by alcoholysis of Jatropha oil using lipase. The process includes (a) screening of various commercial lipase preparations, (b) pH tuning, (c) immobilization, (d) varying water content in the reaction media, (e) vary­ing amount of enzyme used, and (f) varying temperature of the reaction. The best yield 98% (w/w) was obtained by using Pseudomonas cepacia lipase which was immobilized on celite at 50°C in the presence of 4-5% (w/w) water in 8 h. The yield was not affected if analytical grade alcohol was replaced by commercial grade alco­hol. This biocatalyst could be applied four times without loss of any activity.

In order to lower the cost of biodiesel fuel production from Jatropha, Tamalampudi et al. [ 85] used lipase producing whole cells of Rhizopus oryzae which was immobilized onto biomass support particles. The activity of R. oryzae was compared to Novozym 435, the most effective lipase. Methanolysis of Jatropha oil progresses faster than other alcoholysis regardless of any lipase used. The maxi­mum methyl ester content in the reaction mixture reached 80 wt.% after 60 h using R. oryzae, but 76% after 90 h using Novozym 435. Both lipases could be used for repeated batches. They also exhibited more than 90% of their initial activities after 5 cycles. Whole-cell immobilized R. oryzae is a promising biocatalyst for produc­ing biodiesel from oil [85] .