While T. saccharolyticum cannot be considered as a strict CBP candidate since it is noncellulolytic, it does have important properties that give it some warranted atten­tion potentially as a member of a CBP consortium. It is an anaerobic thermophilic bacterium but unlike C. thermocellum, T. saccharolyticum can ferment xylan and all other biomass-derived sugars making the engineering of this organism somewhat less complicated in that regard. Furthermore, high frequency gene transfer has been achieved in this organism [28, 29] increasing the capacity for strain engineering. Importantly, the catabolic pathways have been mapped out with key metabolic enzymes identified [30] providing the necessary knowledge base to address the relevant metabolic pathways. In fact, highly effective metabolic engineering has been performed on T. saccharolyticum allowing it to produce ethanol with a maximum titer of 37 g/L, which while greatly lower than many ethanologens, is the highest reported for a thermophilic anaerobe [31]. While the use of this organism alone is unlikely to achieve the endpoints required for a robust CBP host, we feel that given its great promise as a biocatalyst for the conversion of hemicellulose, it is an organism worthy of future research endeavors.

The classification above resolves whether catalytic domains of cellulases hydrolyze glycosidic bonds at the end of cellulose chains (exocellulases) or in the middle (endocellulases). An important additional distinction among endocellulases is whether they are processive (binding to a cellulase chain and then hydrolyzing at multiple sites, perhaps in sequence on the chain) or nonprocessive (binding, hydro­lyzing a single bond, and releasing). Comparison of multiple crystal structures of cellulases has revealed some of the basis of processivity, along with the basis for endo — vs. exocellulolytic activity (see [84] for a review). It should be emphasized that the type of processivity discussed above relates to individual catalytic domains, since the attachment of CBDs in cis is likely to significantly enhance processivity by tethering the enzyme to the substrate, as is the presence of multiple catalytic domains in a single polypeptide chain.

An important subdivision of the exocellulases is whether they attack the reduc­ing end or nonreducing end of the cellulose chain [2]. In many cases, exocellulases produce the disaccharide cellobiose and are appropriately designated “cellobiohy — drolases,” although many exocellulases also produce longer oligosaccharides such as cellotriose and cellotetraose.

A final mechanistic subdivision that can be made is whether a cellulase cleaves the b-glucosidic bond with retention of chirality at the C1 position (producing the b-anomer) or with inversion (producing the a-anomer). For both mechanisms, cel — lulases use a pair of acidic (asp/glu) residues in a general acid/general base scheme. Other common features include water attack at the C1 position on the nonreducing side of the glycosidic bond to be cleaved and displacement of the O4-sugar. In the inverting mechanism, the water attacks C1 directly, with one of the acidic residues acting as a general base to abstract a proton from the attacking water, and the other acting as a general acid to protonate the O4 leaving group. In the retaining mecha­nism, the general base forms a covalent adduct (an ester linkage between the side — chain carboxyl and C1), displacing the O4, with protonation from the general acid. In a second step, water is activated by the general acid (now in its deprotonated conjugate base form), which attacks the C1 and displaces the C1-asp/glu linkage.

2.2.1 Selecting Appropriate Designer Enzymes

One of the key features in the present invention is the creation of a designer ethanol — production pathway to tame and work with the natural photosynthetic mechanisms to achieve the desirable synthesis of ethanol directly from CO2 and H2O. The natural photosynthetic mechanisms (illustrated in Fig. 1) include (1) the process of photo­synthetic water splitting and proton gradient-coupled electron transport through the
thylakoid membrane of the chloroplast, which produces the reducing power (NADPH) and energy (ATP), and (2) the Calvin cycle, which reduces CO2 by consumption of the reducing power (NADPH) and energy (ATP).

In accordance with the present invention, a series of enzymes are used to create a designer ethanol-production pathway that takes an intermediate product of the Calvin cycle and converts the intermediate product into ethanol. A “designer etha­nol-production-pathway enzyme” is hereby defined as an enzyme that serves as a catalyst for at least one of the steps in a designer ethanol-production pathway. The intermediate products of the Calvin cycle are shown in Fig. 4a. According to the present invention, a number of intermediate products of the Calvin cycle can be utilized to create designer ethanol-production pathway(s); and the enzymes required for a designer ethanol-production pathway are selected depending on from which intermediate product of the Calvin cycle the designer ethanol-production pathway branches off.

In one example, a designer pathway is created that takes glyceraldehydes-3- phosphate and converts it into ethanol by using, for example, a set of enzymes consisting of glyceraldehyde-3-phosphate dehydrogenase, phosphoglycerate kinase, phosphoglycerate mutase, enolase, pyruvate kinase, pyruvate decarboxylase, and alcohol dehydrogenase, as shown in Fig. 4b. In this designer pathway, for conversion of one molecule of glyceraldehyde-3-phosphate to ethanol, an NADH molecule is generated from NAD+ at the step from glyceraldehyde-3-phosphate to 1,3-diphos — phoglycerate catalyzed by glyceraldehyde-3-phosphate dehydrogenase; meanwhile an NADH molecule is converted to NAD+ at the terminal step catalyzed by alcohol dehydrogenase to reduce acetaldehyde to ethanol. Consequently, in this designer pathway (Fig. 4b), the number of NADH molecules consumed is balanced with the number of NADH molecules generated. Therefore, this designer ethanol-production pathway can operate continuously.

In another example, as shown in Fig. 4c, a designer pathway is created that takes the intermediate product, 3-phosphoglycerate, and converts it into ethanol by using, for example, a set of enzymes consisting of phosphoglycerate mutase, enolase, pyruvate kinase, pyruvate decarboxylase, and alcohol dehydrogenase. It can be seen that the last five enzymes of the designer pathway shown in Fig. 4b are identical with those utilized in the designer pathway shown in Fig. 4c. In other words, the designer enzymes depicted in Fig. 4b permit ethanol production from both the point of 3-phosphoglycerate and the point glyceraldehydes 3-phosphate in the Calvin cycle. These two pathways (Fig. 4b. c), however, have different characteristics. Unlike the glyceraldehyde-3-phosphate-branched ethanol-production pathway (Fig. 4b), the 3-phosphoglycerate-branched pathway which consists of the activities of only five enzymes as shown in Fig. 4c could not itself generate any NADH for use in the terminal step to reduce acetaldehyde to ethanol. That is, if (or when) an alcohol dehydrogenase that strictly uses only NADH but not NADPH is employed, it would require a supply of NADH for the 3-phosphoglycerate-branched pathway to operate. Consequently, in order for the 3-phosphoglycerate-branched ethanol — production pathway (Fig. 4c) to operate, it is important to use an alcohol dehydro­genase that can use NADPH which can be supplied by the photo-driven electron

this designer ethanol-production pathway (Fig. 4c). Alternatively, when an alcohol dehydrogenase that can use only NADH is employed, it is prefer­ably here to use an additional embodiment that can confer an NADPH/NADH conversion mechanism (to supply NADH by converting NADPH to NADH, see more detail later in the text) in the designer organism’s chloroplast to facilitate photosynthetic production of ethanol through the 3-phosphoglycerate-branched designer pathway.

In still another example, a designer pathway is created that takes fructose-1,6- diphosphate and converts it into ethanol by using, for example, a set of enzymes consisting of aldolase, triose phosphate isomerase, glyceraldehyde-3-phosphate dehydrogenase, phosphoglycerate kinase, phosphoglycerate mutase, enolase, pyru­vate kinase, pyruvate decarboxylase, and alcohol dehydrogenase, as shown in Fig. 4d, with aldolase and triose phosphate isomerase being the only two additional enzymes relative to the designer pathway depicted in Fig. 4b. The addition of yet one more enzyme in the designer organism, phosphofructose kinase, permits the creation of another designer pathway which branches off from the point of fructose — 6-phosphate for the production of ethanol (Fig. 4e) . Like the glyceraldehyde-3- phosphate-branched ethanol-production pathway (Fig. 4b), both the fructose-1, 6-diphosphate-branched pathway (Fig. 4d) and the fructose-6-phosphate-branched pathway (Fig. 4e) can themselves generate NADH for use in their terminal step to reduce acetaldehyde to ethanol. In each of these designer ethanol-production pathways, the numbers of NADH molecules consumed are balanced with the numbers of NADH molecules generated. Therefore, these designer ethanol-production pathways can operate continuously.

Table 1 lists examples of the enzymes including those identified above for con­struction of the designer ethanol-production pathways. Throughout this specification, when reference is made to an enzyme, such as, for example, any of the enzymes listed in Table 1, it include their isozymes, functional analogs, designer modified enzymes, and combinations thereof. These enzymes can be selected for use in con­struction of the designer ethanol-production pathways. The “isozymes or functional analogs” refer to certain enzymes that have the same catalytic function but may or may not have exactly the same protein structures. For example, in Saccharomyces bayanus, there are four different genes (accession numbers: AY216992, AY216993, AY216994, and AY216995) encoding four alcohol dehydrogenases. These alcohol dehydrogenases essentially have the same function as an alcohol dehydrogenase, although there are some variations in their protein sequences. Therefore, the isozymes or functional analogs can also be selected and/or modified for use in con­struction of the designer ethanol-production pathway. The most essential feature of an enzyme is its active site that catalyzes the enzymatic reaction. Therefore, certain enzyme-protein fragment(s) or subunit(s) that contains such an active catalytic site may also be selected for use in this invention. For various reasons, some of the natu­ral enzymes contain not only the essential catalytic structure but also other structure components that may or may not be desirable for a given application. With techniques

Chlamydomonas reinhardtii cytoplasm; Aspergillus fumigatus; Coccidioides immitis; Leishmania braziliensis; Ajellomyces capsulatus; Monocercomonoicles sp.; Aspergillus clavatus; Arabidopsis thaliana; Zea mays C. reinhardtii cytoplasm; A. thaliana’, Leishmania Mexicana; Lodderomyces elongisporus; Babesia bovis; Sclerotinia sclerotiorum; Pichia guilliennon- clii; Spirotrichonympha leidyi; Otyza sativa; T. pyrifonnis; Leuconostoc mesenteroides; Davidiella tassiana; Aspergillus oryzjae; Schizosaccharomyces pombe; Brassica napus; Z. mays C. reinhardtii cytoplasm; A. thaliana; Saccharomyces cerevisiae; B. bovis; S. sclerotiorum; Trichomonas vaginalis; P. guillietmondii; Pichia stipitis; L. elongisporus; C. immitis; T. pyrifonnis; Glycine max (soybean)

C. reinhardtii cytoplasm; P. stipitis; L. elongisporus; A. thaliana; Lycoris aurea; Chaetomium globosum; Citrus sinensis; Petunia x hybrida; Candida glabrata; Saccharomyces kluyveri; Z. mays; Rhizopus oryzae; Lotus comiculatus; Zymomonas mobiles; Lachancea kluyveri; O. sativa C. reinhardtii mitochondria; Kluyveromyces lactis; Kluyveromyces marxianus; S. cerevisiae; Saccharomyces bayanus; P. stipitis; Entamoeba histolytica; T. vaginalis; L. braziliensis; Botryotinia fuckeliana; A. fumigatus; Dianthus caiyophyllus; Saccharomyces pastorianus; L. kluyveri

JGI Chlre2 protein ID 161689, GenBank: AF268078; XMJ747847; XM_749597; XM 001248115; XM_001569263; XM_001539892; DQ665859; XM_001270940; NM_117020; M80912

GenBank: X66412, P31683; AK222035;DQ221745; XM_001528071; XM_001611873; XMOO1594215; XM_001483612; AB221057; EF122486, U09450; DQ845796; AB088633; U82438; D64113; U13799; AY307449; U17973

of bioinformatics-assisted molecular design, it is possible to select the essential catalytic structure(s) for use in construction of a designer DNA construct encoding a desirable designer enzyme. Therefore, in one of the various embodiments, a designer enzyme gene is created by artificial synthesis of a DNA construct according to bioinformatics-assisted molecular sequence design. With the computer-assisted synthetic biology approach, any DNA sequence (thus its protein structure) of a designer enzyme may be selectively modified to achieve more desirable results by design. Therefore, the terms “designer modified sequences” and “designer modified enzymes” are hereby defined as the DNA sequences and the enzyme proteins that are modified with bioinformatics-assisted molecular design. For example, when a DNA construct for a designer chloroplast-targeted enzyme is designed from the sequence of a mitochondrial enzyme, it is a preferred practice to modify some of the protein structures, for example, by selectively cutting out certain structure component(s) such as its mitochondrial transit-peptide sequence that is not suitable for the given application, and/or by adding certain peptide structures such as an exogenous chloroplast transit-peptide sequence (e. g., a 135-bp Rubisco small — subunit transit peptide (RbcS2)) that is needed to confer the ability in the chloroplast — targeted insertion of the designer protein. Therefore, one of the various embodiments flexibly employs the enzymes, their isozymes, functional analogs, designer modified enzymes, and/or the combinations thereof in construction of the designer ethanol — production pathway(s).

As shown in Table 1, many genes of the enzymes identified above have been cloned and/or sequenced from various organisms. Both genomic DNA and/or mRNA sequence data can be used in designing and synthesizing the designer DNA constructs for transformation of a host alga, plant, plant tissue or cells to create a designer organism for photobiological ethanol production (Fig. 5). However, because of possible variations often associated with various source organisms and cellular compartments with respect to a specific host organism and its chloroplast environment where the ethanol-production pathway(s) is designed to work with the Calvin cycle, certain molecular engineering artwork in DNA construct design including codon-usage optimization and sequence modification is often necessary for a designer DNA construct (Fig. 6) to work well. For example, if the source sequences are from cytosolic enzymes (sequences), a functional chloroplast-target — ing sequence must be added to provide the capability for a designer unclear gene- encoded enzyme to insert into a host chloroplast to confer its function for a designer ethanol-production pathway. Furthermore, to provide the switchability for a designer ethanol-production pathway, it is also important to include a functional inducible promoter sequence such as the promoter of a hydrogenase (Hydl) or nitrate reductase (Nial) gene in certain designer DNA construct(s) as illustrated in Fig. 6a to control the expression of the designer gene(s). In addition, as mentioned before, certain functional derivatives or fragments of these enzymes (sequences), chloroplast-tar — geting transit peptide sequences, and inducible promoter sequences can also be selected for use in full, in part or in combinations thereof, to create the designer organisms according to various embodiments of this invention. The arts in creating and using the designer organisms are further described herein below.

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The growing demand for lower-cost, nonfood, nonrainforest-based feedstock for biodiesel provides new opportunities and stimulate fresh investment in the produc­tion of lower-cost, alternative feedstock such as Jatropha. The governments in South Asia and Africa have identified between 20 and 50 million ha of suitable land for Jatropha cultivation. Indonesia has identified nearly 23 million ha of Jatropha land potential. One hectare of Jatropha can produce between 1.5 and 2.5 tons of seed oil. Jatropha is now becoming one of the prime contenders for biodiesel feed­stock supply in the near future [87] . This is due to the expansion of commercial — scale Jatropha production from India to Africa, Southeast Asia, and Latin America and to pilot programs and larger-scale ventures in China, Central Asia, South/ Central America, and southern parts of the USA.

A variety of equipment is available to obtain oil from the seeds. The oil can be extracted mechanically using a press (ram, hydraulic, or screw) or chemically using organic solvents or water [12, 26, 65] . three phase partitioning (TPP) extraction method [76], and supercritical extraction method [97].

The objective of oil preparation is to find an efficient and effective method in extracting oil from Jatropha seed. Technique of TPP with enzyme pretreatment and sonication constitutes an efficient procedure to obtain oil from Jatropha seed kernels. This technique can extract 97% oil within 2 h [77]. Extraction using ethyl acetate and methyl acetate is better than using hexane [83]. It was found that Gas-Assisted Mechanical Expression (GAME) process in Jatropha oil extraction is capable of reaching yields up to 30 wt.% higher than conventional expression under the same conditions [96] .

Strategies for enhancing security of energy supply and mitigation of energy-related GHG emissions, include, inter alia, the need for: increased energy efficiency (i. e. decreasing energy use per unit of product, process or service); increased use of clean fossil energy (i. e. use of fossil fuels coupled with CO2 sequestration and storage); and increased use of carbon neutral renewable energy, mainly biofuels [23, 196]. Conditions for technically and economically viable biofuel resources are that, they should be [100, 195]: cost-competitive against petroleum-based fuels, require low to no additional land use, enable air quality improvements (e. g. CO2 sequestration), and require minimal water usage. Biofuels are technically carbon neutral, i. e. the release of CO2 during conversion is equivalent to what is captured during growth of the parent material via the photosynthesis process [ 1] . However, they may not always be carbon neutral where energy from fossil fuels are used in the cultivation, harvesting, manufacture of fertiliser and herbicides, processing, and direct conver­sion to energy or specific energy carriers as depicted in the illustration in Fig.1 for biomass from willow short rotation coppice.

The deployment of first — and second-generation biofuels has generated a lot of controversy, mainly due to the negative impact on global food markets [144] and

energy-intensive processes for conversion to fuels [226], respectively. Any effort towards increased production of biofuel-directed biomass must therefore consider the overall sustainability. Microalgae-derived biofuels are devoid of the major drawbacks associated with first — and second-generation biofuels. Their judicious exploitation could therefore make a significant contribution to meeting the primary energy demand, while simultaneously providing environmental benefits [218]

Table 5 provides a preliminary comparison between organic solvent extraction and SCCO2 extraction for lipid extraction from microalgae. A more thorough under­standing of mass transfer mechanism and kinetic parameters involved in lipid extraction from microalgae is needed for an extensive comparison. Large use of toxic solvents and energy-intensive solvent-lipid separation represent the main dis­advantages with commercial use of organic solvent extraction, while expensive fluid compression and high installation cost of an extraction pressure vessel remain the primary obstacles for large-scale SCCO2 extraction [50]. The theoretical advan­tages of SCCO2 extraction, such as tunable selectivity for specific lipid fractions and enhanced kinetics due to the fluid intermediate physicochemical properties, still need to be verified for microalgal lipid extraction.

CEC analysis was performed using the following method: The ground char sample was thoroughly mixed and 2 g was placed in a 250-mL Erlenmeyer flask. Hundred milliliters of 0.5 N HCl was added, the flask was covered with parafilm and shaken vigorously periodically for 2 h. Sample was filtered using a glass fiber fi. ter in a Buchner funnel, washing with 100 mL portions of H2O until wash shows no precipitate with AgNO3. Filtrate was discarded. Moist char was immediately transferred to a clean 250-mL Erlenmeyer flask and a total of 100 mL 0.5 N Ba(OAc)2 was added and a stopper placed on the flask. The mixture was shaken vigorously periodically for 1 h and was filtered, washing with three 100 mL portions H2O. The char was discarded, and the filtrate was titrated with 0.0714 N NaOH using phenolphthalein to first pink. The following equation was used to calculate the CEC value:

milliequivalent mL x normality NaOH x 100 (1)

100g air — dried char (g) sample

Biochar is attracting increasing scientific, political, and industry attention for its potential benefits as a soil amendment. Issues such as food security, declining soil fertility, climate change adaptation, and profitability, are all drivers for implement­ing new technologies or new farming systems. Application of biochar to soil has been shown to have effects ranging from very positive, through to neutral and even negative impacts for crop production. It is therefore essential that the mechanisms for action of biochar in soil be understood before it is applied.

The application of biochar to soil can influence a wide range of soil constraints, including low pH and high available Al [51], soil structure and nutrient availability [13], bioavailability of organic [56] and inorganic contaminants [25], cation exchange capacity (CEC) and nutrient retention [33, 46], and organic matter decline [32]. Biochars have a highly porous structure with surface areas sometimes exceed­ing 1,000 m2/g [18]. Like activated charcoal, they adsorb organics, nutrients and gases, and are likely to provide habitats for bacteria, actinomycetes, and fungi [49]. Increases in water holding capacity following biochar application to soil have been well established [11, 41] . and this may influence crop production, soil microbial populations, and population flux during wetting/drying cycles.

Soil constraints where biochar may provide benefits to productivity include:

• Low pH and high Al availability.

• Low CEC and nutrient holding capacity.

• Low water holding capacity, poor infiltration.

• Poor soil aeration, root development.

• Hard setting soils.

• Residual herbicide or heavy metal phytotoxicity.

• Presence of certain soil-borne diseases.

In some cases, biochar application to soil may influence nutrient availability and nutrient use efficiency [54]. The application of a low nutrient biochar derived from timber increased the retention of N in soil and uptake of N into crop biomass [48]. Lehmann et al. [30] showed that biochar reduced leaching of NH4+, maintaining it in the surface soil where it is available for plant uptake. Similarly, the application of charcoal derived from bamboo into a sludge composting system was shown to provide significant increases in N retention in the compost [25] . Increased fertility of soil resulting from biochar application is likely to increase crop vigor, and thus may enhance disease tolerance.

Biochar is also likely to influence a range of soil physical properties. For example, [11, 13] demonstrated significant declines in soil tensile strength following addition of biochar derived from green waste or pecan shells. These declines in soil tensile strength may allow for better crop root penetration (especially during dry periods), and will also reduce costs associated with soil preparation (such as tillage).

Biochar has been shown to increase biological N2 fixation (BNF) of Phaseolus vulgaris [44], largely due to greater availability of plant micronutrients following biochar application. By increasing potential for BNF, and increasing N use efficiency, lower rates of synthetic N fertilizers may be acceptable for maintaining productivity. Synthetic N fertilizers have a significant C footprint, with over 4t CO, emission required per t N fertilizer produced [55].

Although there is a paucity of published data on the effects of biochar on soil-borne pathogens, evidence is mounting that control of certain pathogens may be possible. The addition of biochar (0.32, 1.60, and 3.20% (w/w)) to asparagus soils infested with Fusarium root rot pathogens increased asparagus plant weights and reduced Fusarium root rot disease [19]. Further, Matsubara et al. (2002) (cited in [49]) have shown that biochar inoculated with mycorrhizal fungi are effective in reducing Fusarium root rot disease in asparagus. A study of bacterial wilt suppression in tomatoes found that biochar derived from municipal organic waste reduced the incidence of disease in Ralstonia solanacearum infested soil [38]. The mechanism of disease suppression was attributed to the presence of calcium compounds, as well as improvements in the physical, chemical, and biological characteristics of the soil. Likewise, Ogawa [57] describes the use of biochars and biochar amended composts in reducing bacterial and fungal soil-borne diseases.

The economic value of biochar as an agricultural commodity is largely untested. Although the benefit of biochar in many systems has been described to increase crop yield, the cost-benefit ratio of applying the technology has not been completed. Van Zwieten et al. [52] discusses several mechanisms for valuing biochar as a commodity. Simply, it could be valued based on its nutrient or liming value, replacing commodities such as fertilizer or lime, alternatively, it could be valued according to benefits to productivity or projected productivity. A recent study [6] using biochar derived from Eucalyptus banded at a low rate of 1 ton/ha was shown to have a breakeven valuation of around Aus$170 per ton of biochar in broadacre wheat, assuming yield benefits for 12 years. In the cost-benefit outcome described by Van Zwieten et al. [52], biochar derived from poultry litter waste was valued at $300 per ton, based on the performance enhancement of three crops following the single
application of biochar. Clearly, the economic value of biochar will depend on its properties, but will also be driven by supply and demand, inherent value of the target enterprise, and demonstrated benefits.

Hydrocarbons are usually formed in very low yields during fast pyrolysis of bio­mass, but can be greatly increased by using proper cracking catalysts with deoxy­genation capability [16 ] . Zeolite catalysts (such as HZSM-5, HY, etc.) are very effective to convert the highly oxygenated crude bio-oils or pyrolysis vapors to hydrocarbons which are dominated by several light aromatic hydrocarbons (ben­zene, toluene, xylene, and naphthalene) [43, 67, 70, 95, 97] . For example, in the studies performed by Adjaye et al. [4, 5], catalytic cracking of the crude bio-oil by HZSM-5 catalysts obtained a organic liquid product with up to 90 wt% of aromatic hydrocarbons, and the aromatic hydrocarbons contained abundant toluene (31.8 wt%) and xylene (33.1 wt%).

Other catalysts were also investigated for the production of light aromatic hydro­carbons. For example, Wang et al. [94] reported that catalytic pyrolysis of biomass using CoMo-S/Al2O3 catalyst produced the four light aromatic hydrocarbons (ben­zene, toluene, xylene, and naphthalene) with the yield reaching 6.3 wt% at 590°C.

The FT synthesis process can be made generally selective by proper choice of oper­ating conditions and catalysts. Most of the studies over last several decades have been focused in this area. The status of the several FT processes through 1950 is given in Table 4. The yield of C3+ product per cubic meter of synthesis gas given in this table is very important because it directly relates to the purity of the synthesis gas. The cost of the production of purified synthesis gas can be as high as 70% of the total cost of the FT process [16]. Earlier work focused on Fe catalysts and the improvements in these catalysts that would increase their operability and selectivity and decrease the operating costs of FT process [16]. The efforts were made to pre­vent the reaction 2CO = CO2 + C and improve the steady-state life of the catalyst. Significant developmental efforts were also made [17, 18] for a more active and mechanically stable catalyst to further reduce the yields of C1 and C2 [19]. The pro­cess improvement efforts were also focused on developing a catalyst which mini­mized the shift reaction.

It was suggested [8] that further selectivity in FT reactions can be attained by either poisoning acceptable catalysts with sulfur compounds or by selecting sulfides of less frequently used catalysts. Storch et al. [20] reported that the initial effect of small amounts of H2S is the increase in nickel-manganese catalyst activity. This was also confirmed by Herrington and Woodward [21] for cobalt-thoria-kisselguhr

“Kilogram of total product, excluding water, carbon dioxide, methane, ethane, and ethylene per volume of reactor per unit time

bWeight percent of oil plus water-soluble product

‘In converter and its accessories only

dBauxite treated, but no T. F.L. added

‘Very small, less than 1%

(100:18:100) catalysts. In their experiments, H2S is mixed with the synthesis gas in small batches and no H2S was eliminated in the off-gas during the course of sulfur poisoning experiments. The first addition of H2S increased the yield of liquid hydrocarbons at constant temperature. As sulfur addition continued, there was a decrease in the yield of gaseous hydrocarbons. Total hydrocarbon yield increased with sulfur addition to the catalyst until 8 mg. of sulfur was added to each gram of catalyst. This work suggested the advantage of stopping sulfidization at low level (1-4 mg of sulfur/g of catalyst) to obtain the benefits of increased liquid hydrocar­bons yield. The results also suggested that the catalyst might show the same behav­ior if presulfided to the same degree before introducing the synthesis gas. The patent of Storch et al. [22] showed that 69% of CO conversion can be obtained for a molybde­num disulfide catalyst alkalized with 2-3% KOH in a feed of 2H2 + CO at 530°F and

13.6 atm. Products from this synthesis were low boiling with 30% of the product C3+ hydrocarbons and organic oxygenated compounds. Laynes [23] indicated that by allowing the sulfur content of the iron catalyst to build up to an optimum ratio and maintain at that level will minimize CO2 formation during hydrogenation of CO to form hydrocarbons.

Over last several decades, a continuous effort to improve catalyst activity, selec­tivity, and stability has been carried out. Besides looking at different forms of iron catalysts, nickel, cobalt, and ruthenium catalysts have been extensively studied. Both cobalt and ruthenium catalysts with different types of promoters have been extensively examined by Exxon and other oil companies. Their studies indicate that while these catalysts give higher initial activity, they also tend to decay rapidly. Numerous patents on these catalysts have been reported by Exxon and other oil companies. In recent years, MOF (metal organic framework) have been tested to improve the selectivity of FT reactions. This work is being carried out at NETL in Pittsburgh and it is still at the development stage. More work in this area (perhaps using recent developments in nanotechnology) is needed.