Y. T. Shah

Abstract This chapter briefly reviews the state of art on the conversion of biomass to biofuels by gasification followed by the gas to liquid conversion of biosyngas via Fischer-Tropsch and related syntheses. An integrated process to produce heat, elec­tricity, or fuel by gasification and Fischer-Tropsch synthesis is analyzed. The present state of gasification reactor technology is outlined. The strategies for syn-gas cleanup are delineated. The catalysis and processes for methanol, Fischer-Tropsch, and isosynthesis are briefly evaluated. Finally, various approaches to an integrated process design depending on the desired end results (heat, electricity, or fuel) and the associated economics for each approach are outlined and briefly discussed.

1 Introduction

The world energy demand is increasing at a faster rate than its supply. This is particu­larly true for the transportation fuel. The sources for total world energy supply at the present time are graphically illustrated in Fig. 1. It is clear from this graph that coal and renewable sources combine to provide as much energy as oil. Heat, electricity, and transportation fuel are the major forms of energy use. While heat and electricity are derived from all sources of energy, transportation fuel has been largely obtained from the oil and natural gas. Although the major world reserve of crudes, heavy oils, and natural gas reside in regions like Middle East, Canada, Venezuela, Russia, etc., the United States is the largest consumer of these forms of energy. In fact, because of the supply and demand discrepancies for the United States, more than 60% of oil con­sumed by the United States is imported from the foreign countries. Furthermore, the global supply of oil is decreasing, while the demand is increasing. One of the reasons

Y. T. Shah (*)

Department of Engineering, Norfolk State University, Norfolk, VA 23504, USA e-mail: ytshah@nsu. edu

J. W. Lee (ed.), Advanced Biofuels and Bioproducts, DOI 10.1007/978-1-4614-3348-4_12, 185

© Springer Science+Business Media New York 2013

for this is the aggressive economic development and the resulting energy need by the countries such as China, India, Latin America, and Russia among others [1, 2].

It is imperative for the United States to be less dependent on foreign oil and develop alternate domestic sources to produce power for heat and electricity and synthetic oils. Besides crude oil and natural gas, the United States also uses considerable amount of coal, particularly for the power industries. The United States has the largest reserve of coal (which can last up to 200 years) [1, 2], which can also be a very good oil substi­tute for the production of transportation fuels. This can decrease the U. S. dependence on the foreign crude and heavy oil significantly. The technologies that have the most promise to convert coal to heat and electricity and/or liquid fuel include coal pyroly­sis, combustion, and gasification. Gasification converts coal into producer gas with different amounts of methane, depending upon the gasification technology. The pro­ducer gas can either be used for heat and electricity or it can be converted into syngas (which largely contains CO and H2) by reforming technology. The syngas can then be converted to a liquid fuel by means of Fischer-Tropsch (FT) synthesis. Both coal — based power production and CTL (coal-to-liquid) technology are commercially avail­able. Unfortunately, they suffer from unacceptable production of green house gases (GHG) such as carbon dioxide and other harmful volatile organic compounds. While other toxic compounds can be removed from power generation and CTL plants, the sequestration of carbon dioxide still remains a major issue.

While for the past several decades the use of biomass for the source of heat, electricity, and transportation fuel has been extensively examined in Europe, Brazil, and several other countries, in recent years the use of biomass as a raw material for heat and electricity and transportation fuels is also becoming increasingly important in the United States. The energy available from renewable biomass sources in the United States has been estimated to be about 20% of the U. S. energy consumption [1, 2]. There are basically three classes of feedstocks derived from biomass that are appropriate for the production of renewable fuels for heat, electricity, and transportation

fuel: (1) starchy and edible feedstocks such as corn, beets, sugar cane, (2) triglyceride feedstocks such as soybeans, algae, jatropha, and about 350 other types of crop oils, and (3) lignocellulosic feedstocks. While, bioethanol and biodiesel derived from first two sources are commercialized and continue to be examined, it is the lignocel­lulosic biomass that is the most abundant class of biomass. While starch and triglyc­erides are only present in some crops, lignocellulose contributes structural integrity to plants and thus always present. In general, most energy crops and waste biomass, such as switch grass, miscanthus, agricultural residues, municipal wastes, animal wastes, waste from wood processing, waste from paper and pulp industries, etc., are lignocellulose that can be used for the generation of heat, electricity, and transportation fuel. The analysis carried out by EPA [4] shows that the use of biomass fuel sources results in the generation of significantly lower quantities of anthropogenic CO2 emissions during power or fuel productions.

Biomass can be converted to biofuels in a number of different ways. For lignocel — lulosic biomass, thermochemical methods for the productions of biosyngas and bioliquids are very popular. These methods utilize gasification, reforming, pyrolysis, extraction, and liquefaction technologies to convert biomass into a variety of biogas and bioliquids. One method widely used is the gasification of biomass followed by reforming and gas cleaning to produce clean syngas. This syngas can be converted to a variety of bioliquids via well-known FT and related syntheses. This method is very versatile in that it can generate biofuels for heat, electricity, or transportation as well as for chemical feed stock. This brief chapter addresses three aspects of this method of biomass conversion: (a) basic chemistry and catalysis of FT and related processes for the conversion of biosyngas to bioliquids, (b) brief descriptions of various processes that currently exist for biomass to biosyngas and biosyngas to methanol and transportation fuels via FT and related syntheses, and (c) a brief assessment of an integrated process for the conversion of biomass to synthetic biodiesel fuel.

Wensheng Ning and Muneyoshi Yamada

Abstract Fischer-Tropsch (FT) synthesis is an effective method to produce liquid fuels from biomass. This chapter reports the study on precipitated Fe catalysts for conversion of CO2-containing syngas to liquid fuels. The influences of promoter Zn, K, and Cu on CO2 activation were analyzed by CO2 temperature-programmed — desorption (CO2-TPD). Cu has no strong effect to activate CO2. K increases mainly CO2 adsorption and is inferior to Zn in producing CO. The catalysts with high Zn/K ratio or low K content possess desorbed CO peak around 930 K in CO2-TPD and decreased CO2 selectivity resulted from CO2 addition in FT synthesis. The Fe catalyst with high Zn/K ratio shows high C2+ hydrocarbon selectivity for CO2 hydrogenation, too. It indicates that the CO2 contained in syngas is able to be activated by suitable promoter(s) for hydrocarbon synthesis at low temperature. The correlation between promoter composition and catalyst reactivity found for SiO2-free Fe catalysts is effective for SiO2-added Fe catalysts.

1 Introduction

The combustion-engine-based transportation system demands liquid fuels worldwide. Crude oil is currently the main source for liquid fuels; however, its reserve is limited. At the same time, the consumption of fuels generates significant pollutants that are damaging the environment. New technologies which can supply environmental — friendly fuels are required to overcome the above problems.

W. Ning (*)

College of Chemical Engineering and Materials Science, Zhejiang University of Technology, Chaowang Road 18#, Hangzhou 310032, China e-mail: wenshning@sohu. com

M. Yamada

Department of Applied Chemistry, Graduate School of Engineering, Tohoku University, Aoba 6-6-07, Aramaki, Aoba-ku, Sendai 980-8579, Japan

J. W. Lee (ed.), Advanced Biofuels and Bioproducts, DOI 10.1007/978-1-4614-3348-4_14, 225

© Springer Science+Business Media New York 2013

Production of butanol is adversely affected by the high costs of traditional substrates such as glucose, corn, sugarcane molasses, and whey permeate. To reduce the cost of production, this biofuel could be produced from economically available renewable feedstocks such as corn stover, wheat, barley, and rice straws, corn fiber, switchgrass, alfalfa, reed canary grass, sugarcane bagasse, miscanthus, waste paper, distillers dry grains and solubles (DDGS), and soy molasses. Currently, costs of corn stover, grasses, and straws are in the range of $24-60/ton as opposed to corn which has ranged from $153-218/ton during recent months. It should be noted that while prices of these residue feedstocks are low, they are associated with additional process steps such as pretreatment, and hydrolysis prior to fermentation. Additionally, fermentation inhibitors are generated during the pretreatment process which either halt or slow down reaction rates or fermentation. This section describes production of butanol from wheat straw, barley straw, corn stover, switchgrass, corn fiber, and DDGS and challenges that are faced when handling these feedstocks for the production of this biofuel.

Lignin is a complex aromatic heteropolymer deposited within the SCWs of all vas­cular plants, and accounts for approximately 30% of the terrestrial organic carbon fixed annually in the biosphere, placing it second to cellulose as the most abundant biopolymer on earth [12]. Lignification aids the plant by providing added strength to xylem fibers that give support for upright growth, by waterproofing tracheary elements that make up the vascular system and by helping increase the resistance of plants to pathogen attack [12] (Fig. 1). Lignin content can vary with environmental factors, but in general comprises around 13-19% of the biomass in switchgrass (Panicum virgatum) [108, 173], 22-25% in Miscanthus (Miscanthusxgiganteus) [18, 169], and around 20% in big bluestem (Andropogon gerardii) and eastern gamagrass (Tripsacum dactyloides) [173], all of which are C4 grass species that have potential as bioenergy feedstocks. In addition, lignin accounts for approxi­mately 25-30% of the dry weight of potential hardwood bioenergy tree crops like poplar and can be even higher in softwood species [134]. The prominence of lignin in a majority of plant tissues has been recognized by reference to the nonstarch or nonsugar components of the plant body as simply “lignocellulosic” biomass. Traditional research attention was given to lignin with respect to chemical pulping and forage digestibility, but recently interest has intensified concerning conversion processes to biofuels and biochemicals. Much of the focus has centered on the fact that the cellulose microfibrils of the SCWs are embedded in a meshwork of HCs and lignin that create a barrier for cellulase enzymes and decrease saccharification efficiency. However, from a thermochemical conversion prospective, the associa­tion of lignin with cellulose is not a major issue and more important is the fact that lignin contains structural units that are more chemically reduced and energy dense than any of the cell wall carbohydrates and thus could serve as a source of hydrocar­bon fuels and high-value chemicals, if means can be found to free those structural units from the polymer.

The ultimate source of lignin in the plant is the amino acid phenylalanine (Phe) which is derived from the shikimate biosynthesis pathway in the plastid ] 154]. Current evidence suggests that through the general phenylpropanoid and monolignol — specific pathways located on or near the cytosolic side of the ER membrane, Phe is deaminated to form cinnamic acid, followed by a series of ring hydroxylations,

O-methylations and side-chain modifications culminating in the production of the p-hydroxycinnamyl alcohol monomers (monolignols) coniferyl and sinapyl alcohol, and to a lesser extent p-coumaryl alcohol [105]. Upon incorporation into the lignin polymer, these monomers are referred to as guaiacyl (G), syringyl (S), or p-hydroxy — phenyl (H) units, respectively ] 149] , In general, angiosperm dicot lignins are composed of G — and S-units, while gymnosperms, with a few notable exceptions, are composed almost entirely of G-units with minor amounts of H-units [188]. Most, if not all, of the enzymes required for monolignol biosynthesis are known and include: phenylalanine ammonia lyase (PAL), the three ER membrane bound cytochrome P450 monooxygenases cinnamate 4 hydoxylase (C4H), coumarate 3-hydroxylase (C3’H) and ferulate 5-hydroxylase (F5H), the two methyltransferases caffeoyl-CoA

3- O-methyltransferase (CCoAOMT) and caffeic acid/5-hydroxyferulic acid

O-methyltransferase (COMT), the two oxidoreductases cinnamoyl-CoA reductase (CCR) and cinnamyl alcohol reductase (CAD) as well as two enzymes 4-coumarate — CoA ligase (4CL) and shikimate hydroxycinnamoyl transferase (HCT) that are involved in the generation of pathway intermediates [37, 38, 70, 93, 109, 144, 194].

Although it is uncertain how the newly synthesized monolignols are translocated to the apoplast (cell wall), once there most evidence suggests that the single electron oxidation of the monolignol phenol by wall-bound peroxidases and/or laccases followed by combinatorial radical coupling commences formal lignin polymerization [12]. Presumably, the coupling of two monolignols with one another initiates polymer­ization. Most likely due to the lack of steric hindrance, coupling between monoli — gnols is favored at the central b carbon of their side chain, resulting in the most common b-b dimer, however b-O-4 and b-5 linked dimers can and do occur [188]. In order for polymerization to continue, the lignin dimer must be dehydrogenated once more to a phenolic radical before it can couple with the next monomer radical. Bond formation is again favored at the central b carbon of the monolignol side chain and depending on the bond configuration and the subunit composition of the dimer, the end-wise coupling process can produce either more b-O-4, when a mono­mer adds to S — and G-units (most common), or b-5 bonds that occur only when adding to G-units [13]. If only the three previously mentioned bonds contributed to lignin polymerization, then the lignin polymer would form a relatively straight lin­ear chain. However, two oligolignol radicals with G-unit ends can also react to form

4- O-5 or 5-5 couplings that generate a branch-like quality to the polymer structure. In fact, lignin containing a high proportion of G-units is more highly cross-linked than lignin rich in S-units, which may contribute to the more rigid and hydrophobic character of G-unit lignin [13]. Therefore, the relative proportion of a given lignin monomer dictates the relative abundance of the inter-unit linkage present in the lignin polymer. Interestingly, the b-O-4 linkage is not only the most common linkage found in plants [62], but it is also the easiest of all the linkages to chemically cleave and increasing this linkage could potentially enhance the efficiency of conversion processes [75]. It should also be noted that an alternative hypothesis with regards to lignin polymerization suggests that lignin monomers are coupled with absolute structural control by proteins in the cell wall bearing arrays of dirigent sites, how­ever there has been no genetic data yet produced to support this claim [ 188] . Additional evidence supporting the predominant radical coupling model of lignification has shown that all phenolic compounds, monolignols or otherwise, that enter into the region of the cell wall where oxidation and radical coupling occur have the potential to be radicalized and incorporated into the lignin polymer, sug­gesting a very flexible process not likely mediated by ligand-specific enzymes [188].

This phenomenon may also allow for a strategy of designing lignins for industrial applications, specifically by regulating the influx and species of monolignol or other phenolic compound into the cell wall [68]. And finally, regarding the global control of lignification, several transcription factors belonging to the MYB and NAC gene families, similar to those responsible for SCW biogenesis, have been shown to play a key role in regulating the expression of many of the genes in the monolignol biosynthesis pathway [204, 211].

There are a large number of cellulosic substrates that are used in the laboratory to assay cellulase reactivity (Table 2). Cellulose is synthesized by plants, and some algae, bacteria, and fungi. In plants, cellulose microfibrils may be covered by hemicellulose polymers and lignin matrices [69]. Cellulose chains from plant fibers typically have a degree of polymerization (DP) of 300-10,000 glucosyl subunits [37]. Although direct assay of deconstruction of native lignocellulosic materials such as corn stover and sugarcane bagasse would be ideal for large-scale biofuel production, cellulase activity is more easily studied using simpler substrates that are enriched in the cellulosic fraction. Moreover, it is often preferable to perform cellulase assays with substrates that have been modified (either noncovalently or covalently) to enhance accessibility, uniformity, or solubility. The substrates listed in Table 2 are commercially available or can be prepared from commercial material by simple pretreatment steps.

End endocellulase; Exo exocellulase; fi-Gluc b-glucosidase activity a DP, degree of polymerization b CR, crystallinity (percentage)

cAssay, type of activity most easily measured with a given substrate d Data from Zhang et al. [81] e Data from Park et al. [55] fData from Valjamae et al. [74]

8 Data from Stalbrand et al. [67] h Data from Zhang et al. [82]

The present invention [2, 3] is directed to a photobiological butanol and related high alcohols production technology based on designer photosynthetic organisms such as designer transgenic plants (e. g., algae and oxyphotobacteria) or plant cells. In this context throughout this specification, a “higher alcohol” or “related higher alco­hol” refers to an alcohol that comprises at least four carbon atoms, which includes both straight and branched alcohols such as 1-butanol and 2-methyl-1-butanol. The Calvin-cycle-channeled and photosynthetic-NADPH-enhanced pathways are con­structed with designer enzymes expressed through use of designer genes in host photosynthetic organisms such as algae and oxyphotobacteria (including cyanobac­teria and oxychlorobacteria) organisms for photobiological production of butanol and related higher alcohols. The said butanol and related higher alcohols are selected from the group consisting of: 1-butanol, 2-methyl-1-butanol, isobutanol, 3-methyl-

1- butanol, 1-hexanol, 1-octanol, 1-pentanol, 1-heptanol, 3-methyl-1-pentanol,

4- methyl-1-hexanol, 5-methyl-1-heptanol, 4-methyl-1-pentanol, 5-methyl-1- hexanol, and 6-methyl-1-heptanol. The designer plants and plant cells are created using genetic engineering techniques such that the endogenous photosynthesis reg­ulation mechanism is tamed, and the reducing power (NADPH) and energy (ATP) acquired from the photosynthetic water splitting and proton gradient-coupled elec­tron transport process can be used for immediate synthesis of higher alcohols, such as 1-butanol (CH3CH2CH2CH2OH) and 2-methyl-1-butanol (CH3CH2CH(CH3)

CH2OH), from carbon dioxide (CO2) and water (H2O) according to the following generalized process reaction (where m, n, x, and y are its molar coefficients) in accordance with the present invention:

m(CO2) + n(H2O) ^ x(higher alcohols) + y(O2) (2)

The photobiological higher alcohols production methods of the present invention completely eliminate the problem of recalcitrant lignocellulosics by bypassing the bottleneck problem of the biomass technology. As shown in Fig. 1, for example, the photosynthetic process in a designer organism effectively uses the reducing power (NADPH) and energy (ATP) from the photosynthetic water splitting and proton gradient-coupled electron transport process for immediate synthesis of butanol (CH3CH2CH2CH2OH) directly from carbon dioxide (CO2) and water (H2O) without being drained into the other pathways for synthesis of the undesirable lignocellu — losic materials that are very hard and often inefficient for the biorefinery industry to use. This approach is also different from the existing “cornstarch butanol produc­tion” process. In accordance with this invention, butanol can be produced directly from carbon dioxide (CO2) and water (H2O) without having to go through many of the energy consuming steps that the cornstarch butanol-production process has to go through, including corn crop cultivation, corn-grain harvesting, corn-grain corn­starch processing, and starch-to-sugar-to-butanol fermentation. As a result, the pho­tosynthetic butanol-production technology of the present invention is expected to have a much (more than ten times) higher solar-to-butanol energy-conversion efficiency than the current technology. Assuming a 10% solar energy conversion efficiency for the envisioned photosynthetic butanol production process, the maxi­mal theoretical productivity (yield) could be about 72,700 kg of butanol per acre per year, which could support about 70 cars (per year per acre). Therefore, this inven­tion could bring a significant capability to the society in helping to ensure energy security. The present invention could also help protect the Earth’s environment from the dangerous accumulation of CO2 in the atmosphere, because the present methods convert CO2 directly into clean advanced biofuels (e. g., butanol) energy.

A fundamental feature of the present methodology is utilizing a plant (e. g., an alga or oxyphotobacterium) or plant cells, introducing into the plant or plant cells nucleic acid molecules encoding for a set of enzymes that can act on an intermediate product of the Calvin cycle and convert the intermediate product into butanol as illustrated in Fig. 1, instead of making starch and other complicated cellular (bio­mass) materials as the end products by the wild-type photosynthetic pathways. Accordingly, the present invention provides, inter alia, methods for producing butanol and/or related higher alcohols based on a designer plant (such as a designer alga and a designer oxyphotobacterium), designer plant tissue, or designer plant cells, DNA constructs encoding genes of a designer butanol — and/or related higher alcohols-production pathway(s), as well as the designer algae, designer oxyphoto — bacteria (including designer cyanobacteria), designer plants, designer plant tissues, and designer plant cells created. The various aspects of the present Host Photosynthetic Organisms

According to the present invention, a designer organism or cell for the photosyn­thetic butanol and/or related higher alcohols production of the invention can be created utilizing as host, any plant (including alga and oxyphotobacterium), plant tissue, or plant cells that have a photosynthetic capability, i. e., an active photosyn­thetic apparatus and enzymatic pathway that captures light energy through photo­synthesis, using this energy to convert inorganic substances into organic matter. Preferably, the host organism should have an adequate photosynthetic CO2 fixation rate, for example, to support photosynthetic butanol (and/or related higher alcohols) production from CO2 and H2 O at least about 1,450 kg butanol per acre per year, more preferably, 7,270 kg butanol per acre per year, or even more preferably,

72,700 kg butanol per acre per year.

In a preferred embodiment, an aquatic plant is utilized to create a designer plant. Aquatic plants, also called hydrophytic plants, are plants that live in or on aquatic environments, such as in water (including on or under the water surface) or perma­nently saturated soil. As used herein, aquatic plants include, for example, algae, blue-green algae (cyanobacteria and oxychlorobacteria), submersed aquatic herbs (Hydrilla verticillata, Elodea densa, Hippuris vulgaris, Aponogeton boivinianus, Aponogeton rigidifolius, Aponogeton longiplumulosus, Didiplis diandra, Vesicularia dubyana, Hygrophilia augustifolia, Micranthemum umbrosum, Eichhornia azurea, Saururus cernuus, Cryptocoryne lingua, Hydrotriche hottoniiflora, Eustralis stel — lata, Vallisneria rubra, Hygrophila salicifolia, Cyperus helferi, Cryptocoryne petchii, Vallisneria americana, Vallisneria torta, H. hottoniiflora, Crassula helmsii, Limnophila sessiliflora, Potamogetonperfoliatus, Rotala wallichii, Cryptocoryne becketii, Blyxa aubertii, Hygrophila difformmis), duckweeds (Spirodela polyrrhiza, Wolffia globosa, Lemna trisulca, Lemna gibba, Lemna minor, Landoltiapunctata), water cabbage (Pistia stratiotes), buttercups (Ranunculus), water caltrop (Trapa natans and Trapa bicornis), water lily (Nymphaea lotus, Nymphaeaceae and Nelumbonaceae), water hyacinth (Eichhornia crassipes), Bolbitis heudelotii, Cabomba sp., seagrasses (Heteranthera zosterifolia, Posidoniaceae, Zosteraceae, Hydrocharitaceae, and Cymodoceaceae). Butanol (and/or related higher alcohols) produced from an aquatic plant can diffuse into water, permitting normal growth of the plants and more robust production of butanol from the plants. Liquid cultures of aquatic plant tissues (including, but not limited to, multicellular algae) or cells (including, but not limited to, unicellular algae) are also highly preferred for use, since the butanol (and/or related higher alcohols) molecules produced from a designer butanol (and/or related higher alcohols) production pathway(s) can readily diffuse out of the cells or tissues into the liquid water medium, which can serve as a large pool to store the product butanol (and/or related higher alcohols) that can be subsequently harvested by filtration and/or distillation/evaporation techniques.

Although aquatic plants or cells are preferred host organisms for use in the meth­ods of the present invention, tissue and cells of nonaquatic plants, which are photo­synthetic and can be cultured in a liquid culture medium, can also be used to create designer tissue or cells for photosynthetic butanol (and/or related higher alcohols) production. For example, the following tissue or cells of nonaquatic plants can also be selected for use as a host organism in this invention: the photoautotrophic shoot tissue culture of wood apple tree Feronia limonia, the chlorophyllous callus-cultures of corn plant Zea mays, the green root cultures of Asteraceae and Solanaceae spe­cies, the tissue culture of sugarcane stalk parenchyma, the tissue culture of bryo — phyte Physcomitrella patens, the photosynthetic cell suspension cultures of soybean plant (Glycine max), the photoautotrophic and photomixotrophic culture of green Tobacco (Nicofiana tabacum L.) cells, the cell suspension culture of Gisekiapharna — ceoides (a C4 plant), the photosynthetic suspension cultured lines of Amaranthuspow — ellii Wats.,Datura innoxia Mill., Gossypium hirsutum L., and N. tabacumx Nicotiana glutinosa L. fusion hybrid.

By “liquid medium” is meant liquid water plus relatively small amounts of inor­ganic nutrients (e. g., N, P, K, etc., commonly in their salt forms) for photoautotrophic cultures; and sometimes also including certain organic substrates (e. g., sucrose, glu­cose, or acetate) for photomixotrophic and/or photoheterotrophic cultures.

In an especially preferred embodiment, the plant utilized in the butanol (and/or related higher alcohols) production method of the present invention is an alga or a blue-green alga. The use of algae and/or blue-green algae has several advantages. They can be grown in an open pond at large amounts and low costs. Harvest and purification of butanol (and/or related higher alcohols) from the water phase is also easily accomplished by distillation/evaporation or membrane separation.

Algae suitable for use in the present invention include both unicellular algae and multiunicellular algae. Multicellular algae that can be selected for use in this inven­tion include, but are not limited to, seaweeds such as Ulva latissima (sea lettuce), Ascophyllum nodosum, Codium fragile, Fucus vesiculosus, Eucheuma denticula — tum, Gracilaria gracilis, Hydrodictyon reticulatum, Laminaria japonica, Undaria pinntifida, Saccharinajaponica, Porphyra yezoensis, and Porphyra tenera. Suitable algae can also be chosen from the following divisions of algae: green algae (Chlorophyta), red algae (Rhodophyta), brown algae (Phaeophyta), diatoms (Bacillariophyta), and blue-green algae (Oxyphotobacteria including Cyanophyta and Prochlorophytes). Suitable orders of green algae include Ulvales, Ulotrichales, Volvocales, Chlorellales, Schizogoniales, Oedogoniales, Zygnematales, Cladophorales, Siphonales, and Dasycladales. Suitable genera of Rhodophyta are Porphyra, Chondrus, Cyanidioschyzon, Porphyridium, Gracilaria, Kappaphycus, Gelidium, and Agardhiella. Suitable genera of Phaeophyta are Laminaria, Undaria, Macrocystis, Sargassum, and Dictyosiphon. Suitable genera of Cyanophyta (also known as Cyanobacteria) include (but not limited to) Phoridium, Synechocystis, Syncechococcus, Oscillatoria, and Anabaena. Suitable genera of Prochlorophytes (also known as oxychlorobacteria) include (but not limited to) Prochloron, Prochlorothrix, and Prochlorococcus. Suitable genera of Bacillariophyta are Cyclotella, Cylindrotheca, Navicula, Thalassiosira, and Phaeodactylum. Preferred species of algae for use in the present invention include: C. reinhardtii, P. subcordi — formis, C. fusca, Chlorella sorokiniana, Chlorella vulgaris, “Chlorella” ellipsoidea, Chlorella spp., D. salina, Dunaliella viridis, Dunaliella bardowil, Haematococcus pluvialis; Parachlorella kessleri, Betaphycus gelatinum, Chondrus crispus, Cyanidioschyzon merolae, Cyanidium caldarium, Galdieria sulphuraria, Gelidiella acerosa, Gracilaria changii, Kappaphycus alvarezii, Porphyra miniata, Ostreococcus tauri, P. yezoensis, Porphyridium sp., Palmariapalmata, Gracilaria spp., Isochrysis galbana, Kappaphycus spp., L. japonica, Laminaria spp., Monostroma spp., Nannochloropsis oculata, Porphyra spp., Porphyridium spp., Undaria pinnatifida, Ulva lactuca, Ulva spp., Undaria spp., Phaeodactylum tricornutum, Navicula sap — rophila, Crypthecodinium cohnii, Cylindrotheca fusiformis, Cyclotella cryptica, Euglena gracilis, Amphidinium sp., Symbiodinium microadriaticum, Macrocystis pyrifera, A. braunii, and S. obliquus.

Preferred species of blue-green algae (oxyphotobacteria including cyanobacteria and oxychlorobacteria) for use in the present invention include: Thermosynechococcus elongatus BP-1, Nostoc sp. PCC 7120, Synechococcus elongatus PCC 6301, Syncechococcus sp. strain PCC 7942, Syncechococcus sp. strain PCC 7002, Syncechocystis sp. strain PCC 6803, Prochlorococcus marinus MED4, P. marinus MIT 9313, P. marinus NATL1A, Prochlorococcus SS120, Spirulina platensis (Arthrospira platensis), Spirulina pacifica, Lyngbya majuscule, Anabaena sp., Synechocystis sp., Synechococcus elongates, Synechococcus (MC-A),Trichodesmium sp., Richelia intracellularis, Synechococcus WH7803, Synechococcus WH8102, Nostocpunctiforme, Syncechococcus sp. strain PCC 7943, Synechocyitis PCC 6714 phycocyanin-deficient mutant PD-1, Cyanothece strain 51142, Cyanothece sp. CCY0110, Oscillatoria limosa, Lyngbya majuscula, Symploca muscorum, Gloeobacter violaceus, Prochloron didemni, Prochlorothrix hollandica, Synechococcus (MC-A), Trichodesmium sp., R. intracellularis, P. marinus, Prochlorococcus SS120, Synechococcus WH8102, L. majuscula, S. muscorum, Synechococcus bigranulatus, cryophilic Oscillatoria sp., Phormidium sp., Nostoc sp.-1, Calothrixparietina, ther­mophilic S. bigranulatus, Synechococcus lividus, thermophilic Mastigocladus laminosus, Chlorogloeopsis fritschii PCC 6912, Synechococcus vulcanus, Synechococcus sp. strain MA4, Synechococcus sp. strain MA19, and T. elongatus.

Proper selection of host photosynthetic organisms for their genetic backgrounds and certain special features is also beneficial. For example, a photosynthetic — butanol-producing designer alga created from cryophilic algae (psychrophiles) that can grow in snow and ice, and/or from cold-tolerant host strains such as Chlamydomonas cold strain CCMG1619, which has been characterized as capable of performing photosynthetic water splitting as cold as 4°C [4], permits photobio­logical butanol production even in cold seasons or regions such as Canada. Meanwhile, a designer alga created from a thermophilic/thermotolerant photosyn­thetic organism such as thermophilic algae C. caldarium and G. sulphuraria and/or thermophilic cyanobacteria (blue-green algae) such as T. elongatus BP-1 and

S. bigranulatus may permit the practice of this invention to be well extended into the hot seasons or areas such as Mexico and the Southwestern region of the United States including Nevada, California, Arizona, New Mexico, and Texas, where the weather can often be hot. Furthermore, a photosynthetic-butanol-producing designer alga created from a marine alga, such as P. subcordiformis, permits the practice of this invention using seawater, while the designer alga created from a freshwater alga such as C. reinhardtii can use freshwater. Additional optional features of a photosynthetic butanol (and/or related higher alcohols) producing designer alga include the benefits of reduced chlorophyll antenna size, which has been demon­strated to provide higher photosynthetic productivity [5] and butanol tolerance (and/ or related higher alcohol tolerance) that allows for more robust and efficient photo­synthetic production of butanol (and/or related higher alcohols) from CO2 and H2O. By use of a phycocyanin-deficient mutant of Synechocystis PCC 6714, it has been experimentally demonstrated that photoinhibition can be reduced also by reducing the content of light-harvesting pigments [6]. These optional features can be incor­porated into a designer alga, for example, by use of a butanol-tolerant and/or chlo­rophyll antenna-deficient mutant (e. g., C. reinhardtii strain DS521) as a host organism, for genetic transformation with the designer butanol-production-pathway genes. Therefore, in one of the various embodiments, a host alga is selected from the group consisting of green algae, red algae, brown algae, blue-green algae (oxy- photobacteria including cyanobacteria and prochlorophytes), diatoms, marine algae, freshwater algae, unicellular algae, multicellular algae, seaweeds, cold-tolerant algal strains, heat-tolerant algal strains, light-harvesting-antenna-pigment-deficient mutants, butanol-tolerant algal strains, higher alcohols-tolerant algal strains, and combinations thereof.

GAME is another potential alternative process for the production of oil with high yields which do not use organic solvents. In this process, CO2 is dissolved in the oil contained in the seeds before pressing the seeds [93]. It was found that at the same effective mechanical pressure (absolute mechanical pressure minus the actual CO2- pressure), the liquid content was the same in both conventional and GAME press cakes. The liquid in the GAME press cake was saturated with CO2 (typically 20-50 wt.%), reducing the oil content compared to the conventional cake by the same amount. The contribution of this effect increased with increasing solubility of the CO2 in the oil. Furthermore, the dissolved CO2 reduced the viscosity of oil by about an order of magnitude [93], which could increase oil extraction. Some addi­tional oil was removed by entrainment in the gas flow during depressurization of the cake.

GAME has some advantages compared with conventional pressing. The first advantage of GAME is the increased yield at lower mechanical pressure. Compared with supercritical extraction, the amount of CO2 that has to be recycled is reduced by two orders of magnitude from typically 1 kg of CO2/kg of seeds [93] to 100 kg of CO2/kg of seeds [69]. Therefore, the energy and equipment cost for the solvent recycle can be reduced. Compared with SCE, the second advantage of GAME is CO2-pressure required is low, which is approximately 10 MPa. In contrast, for SCE extraction, pressures of 40-70 MPa are not unusual ([69]; Rosa et al. 2005). These two effects provide a significant reduction in the energy requirements for recycling and repressurising the CO2. Additionally, some reports in literature suggest that the use of CO2 at 7-20 MPa has a sterilizing effect on the substrates [80, 94]; this may be a beneficial side-effect of the GAME process.

The general applicability of the GAME process to enhance the oil recovery from oilseeds was shown by pressing experiments for sesame, linseed, rapeseed, Jatropha, and palm kernel by Willems et al. [95]. It was proved that GAME was capable of reaching yields that were up to 30 wt.% higher than conventional expression under the same conditions. Despite the lower yields for hulled seeds in conventional expression, GAME yields for hulled and dehulled seeds were very similar. The oil yields obtained for GAME increased with increasing effective mechanical pressure; the yields were the highest at a temperature of 100°C. These effects were similar to conventional expression. With CO2-pressure up to 10 MPa, the oil yield increased significantly. However, increasing the CO2-pressure above 10 MPa did not significantly increase the oil yield.

Genetic engineering of microalgae can provide important and significant improve­ments for algal-biofuel production, by increasing the yields of TAGs to facilitate more efficient biodiesel conversion [187]. In the 1960s, the genome of Anabaena PCC7120 (Chloroxybacteria) was successfully cloned to produce a model organism for academic research [92] . However, for eukaryotic algae there is still a lack of understanding of the detailed molecular biological and regulation of lipid body

TRENDS in Biotechnology

Fig. 5 Basic design of an enclosed horizontal tubular photobioreactor (Adapted from Chisti [39]). Two main sections, airlift system and solar receiver array. The degassing column allows for the transfer of O2 out of the systems and transfer of CO2 into the system as well as providing a means to harvest the biomass. The solar receiver provides a platform for growth enhancement by giving a high surface area to volume ratio (With permission from Elsevier Ltd.)

metabolism [187]. Therefore, genetic manipulation (transgenics) remains limited to a few selected algal laboratory models, i. e. Chlamydomonas reinhardtii, Volvox car — teri, Cyanidioschyzon merolae, Emiliania huxleyi, and the diatoms Phaeodactactylum tricornutum, and Thalassiosira pseudonana. The expanding research interest in biofuel-directed microalgae has led to genetic engineered model organisms [13, 217] and general advances in algal transgenics. Additional genome sequencing for strains with suitable characteristics for biofuels and more universal genetic transformation tools that could enable further development of trangenetic based microalgae-derived biofuel production are considered to be necessary for further advancement [13]

The application of genetic engineering has progressed in several paths, with most recently, the direct manipulation of the microalgal lipid synthesis through gene expres­sion and shunting of photosynthetic carbon partitioning to TAG synthesis [119]. The high expression of acetyl-coA carboxylase gene, which has a role in controlling the level of lipid accumulation, has lead to an improvement of lipid content in the engi­neered microalgae cells [92]. Another genetic engineering process for enhancing the lipid yields that has been tested is the shunting of pathways from starch production to lipid synthesis, by freeing precursor metabolites for desired biofuel products [187]. Both Li et al. [119] and Wang et al. [219] successfully induced inactivation of ADP — glucose pyrophosphorylase in a C. reinhardtii starchless mutant (sta6). Li et al. [119] reported a tenfold increase in lipid synthesis, while Wang et al. [219] reported a 30-fold increase in lipid synthesis, ultimately leading to a high concentration of TAG per cell. Therefore, future use of transgenics to improve TAG content of biofuel-directed microalgae is potentially an important bearing in the quest for economic production.

1.1 Geomorphic Properties

The coastal plain is an expansive geomorphic region of the Southeastern USA that extends from southern New Jersey along the Atlantic coast through the coast of the Gulf of Mexico to South Texas. It comprises nearly 2/3 of the land area of South Carolina (Fig. 1); most of which is either in agriculture or forestry. The coastal plain was initially deposited during a series of sea level rises and recessions; it has been subject to depositional and erosional forces moving and relocating sediments from the Pliocene Epoch (1.8-5 million years ago, [91]) to today. Below Pliocene age sediments are geologic strata consisting of beds of multicolored sands, intermixed with gravel and clay beds laid down during the Tertiary Epoch from 5 to 38 million years ago [91].

Terraces and scarps commonly occur across the coastal plain that are reflective of glacioeustatic changes in ocean level, deposition of sediments, and river dissec­tion during the last 5 million years [30]. The terraces are gently eastward-sloping on the surface, which are bounded by seaward-facing scarps [25] . These scarps are a

few meters in height and demark a time when sea levels were higher. Some of the scarps are definitive on the landscape [32] and are used to divide the area into phys­iographic divisions consisting of (1) lower, (2) middle, and (3) upper coastal plain, based on topography, sediments, elevations above mean sea level, and soils [30]. Their elevations range from sea level to about 150 m.

During 5-6-h long experiments the product gas composition did not change significantly; CO and CO2 were the major gas components with Ct-C4 hydrocar­bons accounting for approximately 20 vol% of the product gas. The product col­lected in the ESP that operated at the exit temperature of 80°C was a dark brown viscous liquid while that collected in the condensers was mostly water (90-95 wt%) with a thin layer of organics floating on the top. In every test significant amounts of carbon (coke and char) were produced which were collected in the reactor, the cyclone, and the filter. The carbon yields were determined by weight difference of the collected solids and of the initial catalysts. The yields of liquids corresponded to the weight of the fractions collected in the ESP and condensers, and the amount of gaseous product was calculated based on its volume and composition (excluding the carrier gas). Table 1 shows the results of two experiments carried out using the

Albemarle catalyst. The first experiment was run for 6 h and the final ratio of cata­lyst to biomass was 0.38 and the second experiment was run from 2 h with a final catalyst to biomass ratio of 1.2.

Mass balance closures were in the range 83-85% with 15-17% not accounted for. These losses most likely comprised of noncondensed water and volatile organic compounds that were not analyzed by gas chromatography. The amounts of carbon solids (char and coke) were in the range of 15-23% of the weight of the feedstock while the yields of total liquid were 35-40%, and those of gas were 25-28%. Similar product distribution was reported by Zhang et al. [32], Aho et al. [33], and Agblevor et al. [28] in bubbling fluidized beds. However, Lappas et al. [27] achieved significantly higher yields of liquids using a fluidized catalytic cracking system (circulating fluidized bed) with ZSM-5 catalyst. The most likely reason for that was maintaining the catalyst activity by continuous regeneration.

The liquid product included three phases: a very thin top organic layer, the middle most abundant water-rich fraction, and the bottom dark brown organic fraction. The yield of the heavy organic fraction was about 12% based on the feedstock. The light liquid hydrocarbon phase was 3-5% of the total condensate. The analysis of the organic liquid from the first test showed the following elemental composition: C 67-68 wt%, H 6.2-6.6 wt%, O 25-26 wt%. The oxygen content of this liquid was significantly less than for crude bio-oil from a non-catalytic process that contains 45-50% oxygen (37­40% on water-free basis). The organic liquid from the second test was even much more deoxygenated and included 83% C, 7.3% H, and 9.6% O, which is the lowest known to us oxygen content from bench-scale catalytic pyrolysis reported in the litera­ture ([27,28, 32, 33], all reporthigher oxygen contents of 13.5-22%).

In both tests, the biomass carbon conversion to organic liquid was 20-22% with most of carbon converted to gas and char/coke. It seems that in our experiments the lignin part of the feedstock was partly depolymerized (Mw 330-350) while the car­bohydrates were mostly converted to solids (char and coke), gas, and water. Agblevor et al. [29] had also concluded that the majority of the organic oil from catalytic pyrolysis in their experiments originated from lignin. The results showed that pyrol­ysis oil with highly reduced oxygen content can be produced by catalytic pyrolysis; however, the biomass carbon to liquid conversion was still low and more research is needed to improve the process performance.