The metals Fe, Ni, Co, and Ru have the required FT activity for commercial appli­cation. Under practical operating conditions Ni produces too much CH4, while the available supply of Ru is insufficient for large scale application. This leaves only Fe and Co as viable catalysts [11].

Fe and Co are commercial catalysts for FT synthesis with CO2 — free syngas. Riedel et al. [16] compared the performance of Fe and Co catalysts in the mixtures of CO, CO2, and H2. With increasing CO2 and decreasing CO content in the feedgas, the product composition shifts from a mixture of mainly higher hydrocarbons to almost exclusively methane for Co catalyst, while Fe-based catalyst synthesizes the same hydrocarbon products from CO2/H2 as from CO/H2 syngas. Zhang et al. [17] also found that the CO2 hydrogenation products contained about 70% or more methane for supported cobalt.

These distinctions are partly attributed to that Fe catalyst is active for WGS reaction, but Co catalyst has no such activity. On Fe catalyst, CO2 can be hydrogenated into FT products by two steps as shown in (1) and (2): CO is converted from CO2 by reverse WGS reaction, then produced CO is further hydrogenated to hydrocar­bons [9, 11, 18]. In contrast, Co catalyst cannot convert CO2 into CO.

Another factor is the different prerequisites to achieve the kinetic regime of FT synthesis for Fe and Co catalysts. With Fe catalyst, the FT kinetic regime is gener­ated through the formation of stable FT sites via carbiding, and their selectivity is invariant against changes of reactant concentrations [19] . On the contrary, the FT regime can exist only at a sufficiently high CO partial pressure for Co catalyst [20]. Therefore, Co is not fit to hydrogenate CO2 in nature, while Fe is promiseful to FT synthesis from CO2-containing syngas.

As indicated above, conversion of cellulosic biomass to butanol requires pretreatment employing dilute sulfuric acid or dilute sodium hydroxide, or alkaline peroxide. During the pretreatment and neutralization process, inhibitors such as salts (sodium acetate, sodium chloride, and sodium sulfate), and chemicals including furfural, hydroxymethyl furfural (HMF), syringealdehyde, and acids (acetic, glucuronic, ferulic, and p-coumaric) are produced. Some of these chemicals are toxic to the culture. In a recent study it was observed that sodium sulfate [18], sodium chloride [59], glucuronic, ferulic, and p-coumaric acids, phenol, and syringealdehyde [18] were toxic to a butanol producing culture. Ferulic acid, at a concentration as low as 0.3 g/L, was a strong inhibitor to both cell growth and fermentation. On the contrary, syringealdehyde (0.3-1.0 g/L) was not so toxic to the cell growth; how­ever, it resulted in complete arrest of ABE production [18]. As expected, phenol was found to be inhibitory to both cell growth and ABE production.

In order to produce ABE from agricultural residues successfully, inhibitors present in the hydrolyzates must be removed prior to fermentation. In recent studies, Qureshi et al. [61, 62] attempted to produce butanol from untreated and treated BSH and CSH. ABE was successfully produced from the treated hydrolyzates, however, strong inhibition of cell growth was still observed. In case of BSH and CSH, 0.80 and 0.77 g/L cell mass was obtained as compared to 2.66 g/L in the control experiment in which glucose was used. It is likely that chemical inhibitors were removed from the hydrolyzates by overliming leaving behind salts that were generated during neutralization. In order to reduce or eliminate cell growth inhibition, it is recom­mended that salts also should be removed from the medium by electrodialysis [59] followed by fermentation. Possibly, removal of salts could improve both cell growth and productivity.

M. Abramson, O. Shoseyov, S. Hirsch, and Z. Shani

Abstract To date, most ethanolic fuel is generated from “first generation” crop feedstocks by conversion of soluble sugars and starch to bioethanol. However, these crops exploit land resources required for production of food. On the other hand, utilization of “second generation” lignocellulosic biofuels derived from the inedible parts of plants remains problematic as high energy inputs and harsh condi­tions are required to break down the composite cell walls into fermentable sugars. This chapter reviews and discusses genetic engineering approaches for the generation of plants modified to increase cellulose synthesis, enhance plant growth rates, cell wall porosity and solubility, as well as improve cell wall sugar yields following enzymatic hydrolysis. Strategies focusing on increased accessibility of cellulose­degrading enzymes to their substrates have been developed. These approaches reduce cell wall crystallinity or alter the hemicellulose-lignin complexes. A novel approach to cell wall modification involving the introduction of noncrystalline, soluble polysaccharides into cell walls is also presented. The use of such approaches may promote and accelerate the future use of lignocellulosic feedstocks for the bioethanol industry.

M. Abramson

The Robert H. Smith Institute of Plant Sciences and Genetics in Agriculture, and The Otto Warburg Minerva Center for Agricultural Biotechnology, Faculty of Agricultural, Food and Environmental Quality Sciences,

The Hebrew University of Jerusalem, P. O. Box 12, Rehovot 76100, Israel

FuturaGene Ltd., 2 Pekeris Street, P. O. Box 199, Rehovot 76100, Israel

O. Shoseyov

The Robert H. Smith Institute of Plant Sciences and Genetics in Agriculture, and The Otto Warburg Minerva Center for Agricultural Biotechnology, Faculty of Agricultural, Food and Environmental Quality Sciences,

The Hebrew University of Jerusalem, P. O. Box 12, Rehovot 76100, Israel

S. Hirsch • Z. Shani (H)

FuturaGene Ltd., 2 Pekeris Street, P. O. Box 199, Rehovot 76100, Israel e-mail: ziv@futuragene. com

J. W. Lee (ed.), Advanced Biofuels and Bioproducts, DOI 10.1007/978-1-4614-3348-4_18, 315

© Springer Science+Business Media New York 2013

In general, cellulase activities measured using insoluble substrates are more likely to be relevant to large-scale biomass deconstruction than chemically modified, soluble chains. A variety of different insoluble substrates are used to measure cellulase activ­ities. These substrates differ in their biological sources and pretreatment methods, and as a result, they have different structural and physical properties (Table 2). As a result, different substrates are more appropriate for different types of enzymes.

By pretreatment with acid (typically phosphoric acid), insoluble cellulose preparations can be obtained that have decreased crystallinity, and thus are typically more susceptible to enzymatic digestion than crystalline substrates [22] . Although published protocols differ in subtle but important ways, high concentrations of phosphoric acid can produce PASC (for phosphoric acid swollen cellulose, [78]). By prehydrating Avicel prior to acid treatment, Zhang et al. prepared an amorphous cellulose of even higher reactivity, which they term regenerated amorphous cellu­lose (RAC, [82] ).

Cellulose substrates with high crystallinity include bacterial cellulose (BC), which has high DP values of 2,000-8,000, and a high 60-90% crystallinity index [37]. Microcrystalline cellulose, or “Avicel” PH, which is prepared by acid hydro­lysis of wood pulp, has a lower DP value (150-300) [37], although it retains a high level of crystallinity. Because it has a high ratio of free ends to accessible b-gluco — sidic bonds due to its lower DP, Avicel is especially suited for the measurement of exoglucanase activity from a crystalline substrate [ 82]. Finally, Whatman No. 1 filter paper, manufactured from cotton linters, is highly heterogeneous and is often used for measuring total cellulase activity.

DNA constructs are generated in order to introduce designer butanol-production­pathway genes to a host alga, plant, plant tissue, or plant cells. That is, a nucleotide sequence encoding a designer butanol-production-pathway enzyme is placed in a vector, in an operable linkage to a promoter, preferably an inducible promoter, and in an operable linkage to a nucleotide sequence coding for an appropriate chloro- plast-targeting transit-peptide sequence. In a preferred embodiment, nucleic acid constructs are made to have the elements placed in the following 5′ (upstream) to 3′ (downstream) orientation: an externally inducible promoter, a transit targeting sequence, and a nucleic acid encoding a designer butanol-production-pathway enzyme, and preferably an appropriate transcription termination sequence. One or more designer genes (DNA constructs) can be placed into one genetic vector. An example of such a construct is depicted in Fig. 2a. As shown in the embodiment illustrated in Fig. 2a, a designer butanol-production-pathway transgene is a nucleic acid construct comprising: (a) a PCR forward primer; (b) an externally inducible promoter; (c) a transit targeting sequence; (d) a designer butanol-production-path­way-enzyme-encoding sequence with an appropriate transcription termination sequence; and (e) a PCR reverse primer.

In accordance with various embodiments, any of the components (a)-(e) of this DNA construct are adjusted to suit for certain specific conditions. In practice, any of the components (a)-(e) of this DNA construct are applied in full or in part, and/or in any adjusted combination to achieve more desirable results. For example, when an algal hydrogenase promoter is used as an inducible promoter in the designer butanol — production-pathway DNA construct, a transgenic designer alga that contains this DNA construct will be able to perform autotrophic photosynthesis using ambient air CO2 as the carbon source and grows normally under aerobic conditions, such as in an open pond. When the algal culture is grown and ready for butanol production, the designer transgene(s) can then be expressed by induction under anaerobic condi­tions because of the use of the hydrogenase promoter. The expression of designer gene(s) produces a set of designer butanol-production-pathway enzymes to work with the Calvin cycle for photobiological butanol production (Fig. 1).

The two PCR primers are a PCR forward primer (PCR FD primer) located at the beginning (the 5′ end) of the DNA construct and a PCR reverse primer (PCR RE primer) located at the other end (the 3′ end) as shown in Fig. 2a. This pair of PCR primers is designed to provide certain convenience when needed for relatively easy PCR amplification of the designer DNA construct, which is helpful not only during and after the designer DNA construct is synthesized in preparation for gene trans­formation, but also after the designer DNA construct is delivered into the genome of a host alga for verification of the designer gene in the transformants. For example, after the transformation of the designer gene is accomplished in a C. reinhardtii — arg7 host cell using the techniques of electroporation and argininosuccinate lyase (arg7) complementation screening, the resulted transformants can be then analyzed by a PCR DNA assay of their nuclear DNA using this pair of PCR primers to verify whether the entire designer butanol-production-pathway gene (the DNA construct) is successfully incorporated into the genome of a given transformant. When the nuclear DNA PCR assay of a transformant can generate a PCR product that matches with the predicted DNA size and sequence according to the designer DNA construct, the successful incorporation of the designer gene(s) into the genome of the transfor­mant is verified.

Therefore, the various embodiments also teach the associated method to effec­tively create the designer transgenic algae, plants, or plant cells for photobiological butanol production. This method, in one of embodiments, includes the following steps: (a) selecting an appropriate host alga, plant, plant tissue, or plant cells with respect to their genetic backgrounds and special features in relation to butanol produc­tion; (b) introducing the nucleic acid constructs of the designer genes into the genome of said host alga, plant, plant tissue, or plant cells; (c) verifying the incorporation of the designer genes in the transformed alga, plant, plant tissue, or plant cells with DNA PCR assays using the said PCR primers of the designer DNA construct; (d) measuring and verifying the designer organism features such as the inducible expression of the designer butanol-pathway genes for photosynthetic butanol production from carbon dioxide and water by assays of mRNA, protein, and butanol-production characteris­tics according to the specific designer features of the DNA construct(s) (Fig. 2a).

The above embodiment of the method for creating the designer transgenic organ­ism for photobiological butanol production can also be repeatedly applied for a plurality of operational cycles to achieve more desirable results. In various embodiments, any of the steps (a)-(d) of this method described earlier are adjusted to suit for certain specific conditions. In various embodiments, any of the steps

(a) -(d) of the method are applied in full or in part, and/or in any adjusted combina­tion. Many examples of designer butanol-production-pathway genes (DNA con­structs) are shown in the sequence listings: SEQ ID NOS:1-57 of the PCT Patent Application Publication No. WO 09105733 and SEQ ID NOS:1-165 of the US Patent Application Publication No. 2011/0177571 A1.

The nucleic acid constructs, such as those presented in the examples earlier, may include additional appropriate sequences, for example, a selection marker gene, and an optional biomolecular tag sequence (such as the Lumio tag). Selectable markers that can be selected for use in the constructs include markers conferring resistances to kanamycin, hygromycin, spectinomycin, streptomycin, sulfonyl urea, gentamy — cin, chloramphenicol, among others, all of which have been cloned and are avail­able to those skilled in the art. Alternatively, the selective marker is a nutrition marker gene that can complement a deficiency in the host organism. For example, the gene encoding argininosuccinate lyase (arg7) can be used as a selection marker gene in the designer construct, which permits identification of transformants when C. reinhardtii arg7- (minus) cells are used as host cells.

Nucleic acid constructs carrying designer genes can be delivered into a host alga, blue-green alga, plant, or plant tissue or cells using the available genetic transforma­tion techniques, such as electroporation, PEG-induced uptake, and ballistic delivery of DNA, and Agrobacterium-mediated transformation. For the purpose of deliver­ing a designer construct into algal cells, the techniques of electroporation, glass bead, and biolistic gene gun can be selected for use as preferred methods, and an alga with single cells or simple thallus structure is preferred for use in transforma­tion. Transformants can be identified and tested based on routine techniques.

An environmentally kind process was developed for the production of biodiesel from Jatropha oil using a heterogeneous solid super base catalyst and calcium oxide. The results revealed that under the optimum conditions of catalyst calcina­tions, temperature of 900°C, reaction temperature of 70°C, reaction time of 2.5 h, catalyst dosage of 1.5%, and methanol/oil molar ratio of 9:1, the oil conversion was 93% [34]. Nazir [58] found that the yield of JCO FAME could reach up to 94.35% using the following reaction conditions: 79.33 min reaction time, 10.41:1 methanol:oil molar ratio, and 0.99% of CaO catalyst at reaction temperature 65°C.

Wastewater sparged with CO2 provides a conducive growth medium for microalgae, enabling faster production rates and reduced nutrient levels in treated wastewater (Table 6), decreased harvesting costs, and increased lipid production [123]. Therefore, coupling of the production of biofuel-directed microalgae with bioreme­diation of wastewaters is considered an important strategy for successful deploy­ment [16, 123,215].

Microalgae are efficient removers of chemical and organic contaminants, heavy metals, and pathogens from wastewater [150], which provide a pathway for combating eutrophication in conjunction with the production of microalgae as energy resource [228]. This characteristic enhances the sustainability potential of the production process, through potential savings on requirements for chemical remediation of waste­water [218], and minimises the need for freshwater for algae production [118,195].

Microalgae have also exhibited significant potential for biological removal of hazardous or toxic compounds due to their negatively charged surfaces [106]. For example, they have demonstrated a strong absorption of polyvalent cations; an ion exchange capacity that is the basis for removal of heavy metals from wastewaters [149]. Another advantage is in the production of photosynthetic oxygen in wastewa­ter, which reduces the need for external aeration. The oxygen is also required by bacteria for biodegradation of pollutants such as polycyclic aromatic hydrocarbons (PAHs), phenolics and organic solvents [149]. However, some of these pollutants are potent inhibitors of photosynthesis in microalgae because they can induce mor­phological changes in the cells that lead to physiological incompatibility [106].

In principle, therefore the problem of increasing ethanol production was solved. The remaining problem was to make sure that the ethanol produced was consumed.

The Government solved the problem using two instruments [1]:

• Adopting mandates for mixing ethanol to gasoline. Up to 1979, the mixture of ethanol in the gasoline increased gradually to approximately 10% which required small changes in the existing motors. In 1981, ethanol consumption reached 2.5 billion liters.

• Setting the price of ethanol paid to producers at 59% of the selling price of gasoline (which was more than twice the cost of imported gasoline). The high price of gasoline has been used for a long time by the Government as a method of collecting resources to subsidize diesel oil. Parts of such resources were then used to subsidize ethanol.

Subsidies of approximately one billion dollars per year on the average over the 30 years were needed to sustain the program. These subsidies were removed gradu­ally and in 2004 the price paid to ethanol producers was similar to the cost of gaso­line in the international market as seen in Fig. 3.

Various solid-state 1 3C NMR techniques, such as CP/MAS and occasionally DP technique, have been employed to identify carbon functionality and aromaticity of biochars [1, 3, 7, 8, 10, 11, 13, 19, 23, 26]. In these studies, chemical structure of biochars to a greater extent depends upon the pyrolysis temperature, but is not much affected by heating rate and the nature of biomass [1, 13]. Biochars prepared at rela­tively low temperatures up to ~350°C retain spectral features of the original ligno — cellulosic composition of biomass [25]. For example, characteristic peaks of cellulose (0-alkyl carbons around 62, 72, 84 ppm, and di-O-alkyl carbons around 103 ppm), as well as those of lignin (methoxyl carbons ~57 ppm, aromatic carbons ~130 ppm, and aromatic C-O ~150 ppm) can still be observed in spectra of biochars within this temperature range (<350°C). For higher heat treatment temperature (HTT) biochars, a well-defined aromatic resonance evolves simultaneously with the decrease of the lignocellulosic resonances and signals of aliphatic, carboxyl, and carbonyl carbons. Lignin structures are more thermally stable than cellulose struc­tures because characteristic peaks of lignin such as the phenolic shoulders around 150 ppm can survive even at temperatures about ~550°C [25]. For HTT about 600°C upwards, the general shapes of the 13C NMR spectra of biochars are very alike, with a strong broad resonance line near 128 ppm in the aromatic region [1, 8, 22, 25].

Chemical transformation of lignocellulosic materials into graphitic structures with the temperatures between 800 and 1,000°C is well documented [1, 13, 25]. Aromatic cluster typical of chars is also reported to grow with increasing HTT [21].

To assess a new energy technology before considering its implementation, it is essential to perform a life-cycle analysis on its total energy efficiency and environ­ment impact, including both its potential benefits and risks. A viable energy tech­nology should have a significant net energy gain or a carbon footprint reduction based on its objective life-cycle analysis. Chapter 30 reports the process economics and greenhouse gas audit for microalgal biodiesel production. Chapter 31 discusses the sustainability considerations about microalgae for biodiesel production while Chap. 32 reports a life-cycle assessment for algae-to-energy systems.