The raw syngas from the gasifier needs significant cleaning and conditioning and treating before it is suitable for subsequent catalytic synthesis. A typical composition of the raw biosyngas from an air-blown CFB gasifier is given in Table 9. The syngas contains H2/CO ratio of 0.9 and it makes up only 34 vol.% corresponding to 48.9% of chemical energy. The remainder of energy is mainly contained in CH4, C2H4, and benzene (total of 44.4%). As mentioned before, this can be converted to CO and H2 by appropriate reforming process. The major gas cleaning issue is the presence of 7 g/m3 of tars in the gas. Tars are condensable organic compounds with boiling points

between 80 and 350°C. When the temperature in the system decreases below 350°C, tars start condensing resulting in fouling and ultimately failing of the system. Typical inorganic biomass impurities are NH3, HCl, and H2S and in minor quantities COS, CS2, HCN, and volatile metals. Also, 2 g/m3 of solids are present in raw biosyngas.

A typical gas conditioning lineup comprises gas cooling, water gas shift (or other reforming), CO2 removal, and impurities removal (e. g., H2S, COS, HCN, volatile metals). Cooling can be achieved with a cooler or a water quench. While a cooler can utilize latent heat in the gas, for biomass there is an increased risk of fouling due to relative high alkaline and chloride concentrations compared with coal. In a gasifier using water quench, fouling problems are avoided. Except for gas cooling, there is a great deal of similarity in biosyngas conditioning and the treating of fossil fuel-based syngas. Thus, biosyngas can be cleaned using the available technologies for commercial coal to liquid operations. There are no biomass specific impurities that require a totally different gas cleaning approach [40] .

In commercial FT operation, catalysts are replaced or regenerated after a certain operational period. The level of gas cleaning is a matter of economics between the cost of catalyst regeneration and the cost of cleaning up front. This economics may depend on the nature of FT synthesis and the catalyst. As a rule of thumb, a maxi­mum value of 1 ppm V may be used for the sum of the nitrogen-containing (NH3 + HCN) and sulfur-containing (H2S + COS + CS2) compounds. For the halides (HCl + HBr+HF) and alkaline metals, a lower level of 10 ppb V should be targeted. There are no limits regarding the poisoning of the catalysts by the tars. For FT syn­thesis, since gas is compressed to 25-60 atm pressure, the concentration of organic compounds must be below the dew point at FT pressure so as to avoid condensation of tars which can poison the catalyst and foul up the entire system. For thiophene and pyridine, the concentrations should be below ppmV level since they poison the catalysts. Solids should be removed completely as they foul the system and obstruct the operation of the fixed bed reactor. For H2, CO, CO2, CH4, N2, paraffins such as ethane and propane, and olefins such as ethylene and propene there are no standards. While hydrocarbons can be further reduced to CO and H2 by reforming, their small concentrations do not affect the operation of the FT reactor. Often BTX (benzene, toluene, and xylene) are used as guidelines for gas cleaning.

It is well known that the hydrogenation of CO2 proceeds via the CO intermediate [9,11,18,22-26]. That is, during the hydrogenation of CO2 over Fe catalysts, some CO2 is firstly converted to CO through reverse WGS reaction and the formed CO is converted consecutively through FT reaction to hydrocarbons. In a previous study of CO2 hydrogenation [27], CO formation was found to be the essential step of producing C, + hydrocarbons with high conversion and selectivity. Pijolat et al. suggested that CO can be obtained in two different ways: by partial dissociation of CO2 and by the reverse WGS reaction [28].

H2 is adsorbed only on Fe [29, 30], but CO2 adsorbed on both Fe and K [29-31]. Higher K content is beneficial for CO2 conversion to FT product [16].

Iron carbides are responsible for the formation of olefins and long-chain hydro­carbons in CO2 hydrogenation [32]. However, Ando et al. [18] found that the major surface phases of the Fe-Cu catalysts were FeO and/or FeCO3 after CO2 hydrogena­tion. Suo et al. [33] studied CO2 hydrogenation on TiO2-, ZrO2-, and Al2O3-supported iron catalysts. The catalyst with the optimum ratio of iron cations vs. zero valent iron gave good catalytic activity and selectivity in the synthesis of C2 + hydrocarbons from CO2 and H2.

The different views about the active phase in Fe catalysts for CO2 hydrogenation indicate that much study is needed to make it clear.

The purpose of process integration is to combine more than one unit operations into a single unit to reduce both capital and operational cost. In butanol fermentation, early reports on process integration were published in the late 1980s and the early 1990s [13, 52] where butanol fermentation was integrated with product separation. In the case of butanol production by fermentation, following process integrations can occur depending upon the feedstock used:

1. Fermentation and recovery of butanol.

2. Hydrolysis of feedstock and fermentation to butanol.

3. Hydrolysis of feedstock, fermentation, and separation of butanol.

In the case of fermentation and recovery, feedstock does not require hydrolysis such as glucose or if it requires hydrolysis, the later can be performed by the butanol producing culture while fermentation occurs. An example of this is the production of butanol from whey permeate (a byproduct of cheese making industry) and simul­taneous recovery by gas stripping. Whey permeate contains lactose (a disaccharide of glucose and galactose) which can be hydrolyzed by the culture into monomeric sugar units. Then both of these sugars can be converted to butanol. Butanol that is produced by the culture can be recovered simultaneously by one of the product recovery techniques mentioned in Sect. 4.

The second group is where a feedstock requires hydrolysis using exogenous enzymes. In this case, the butanol producing culture is not capable of hydrolyzing the substrate and hence either enzymes are added to the reactor or hydrolytic enzyme producing culture is propagated with the butanol producing culture. The conditions under which enzymes perform hydrolysis need to be close to the cultivation condi­tions of butanol producing culture. In this system, hydrolysis and fermentation are all combined. This group is called SSF (Simultaneous Saccharification and Fermentation). The third group is where hydrolysis, fermentation, and recovery are combined. In this case, enzymes or a hydrolytic enzyme producing culture hydroly­ses the feedstock such as cellulosic biomass, the butanol producing culture performs fermentation, and application of product (butanol) recovery technique recovers the product simultaneously from the reactor. The third process can be abbreviated as SSFR (simultaneous saccharification, fermentation, and recovery).

Large portions of crop biomass is wasted during harvest, mostly in the form of straw that carries the grain [50]. This straw—from corn, barley, oat, rice, wheat, sorghum, and sugarcane are currently the major feedstocks used for second-generation bio­fuel production and yield nearly 1.5 billion tons of lignocellulosic biomass which can potentially provide up to 442 billion liters of bioethanol per year [67]. Seventeen years of research conducted by Gelfand et al. [45] potentially redefined the “food versus fuel” debate by concluding that growing grain for food and use of the residue stalks for biofuel is more energetically efficient than the use of grain for bioethanol. The cellulose, hemicelluloses, lignin, and ash content of some common straw feed­stocks are presented in Table 1.

Relatively high percentages of lignin limit digestibility of residual biomass, while ash contains large amount of silica microparticles released at burning. Reduction of silica content in straw can enhance enzymatic digestibility, but may simultaneously reduce plant resistance to pathogens [50]. Thus, feedstock should be engineered to contain less silica while programming in alternative modes of disease resistance, such as genetic engineering or increased pesticide use.

Arabinoxylan is the major noncellulosic polysaccharide in cereals. It is composed of a xylose backbone with arabinose residues and contributes to the structural rigidity

of grass cell walls. Arabinoxylans can covalently bind ferulic acids and other phe — nolics [63], which in turn crosslink neighboring arabinoxylans and form ester bridges with lignin, hindering enzymatic degradation of polysaccharides.

Soluble cellulose chains (CMC and HEC) are long polymers, and thus in solution they act as strong viscogens. Upon endocellulase cleavage of these polymers into shorter fragments, the viscosities of CMC and HEC solutions decrease as digestion progresses [14, 21]. Viscosity changes can easily be measured by using a vertical capillary tube (such as an Ostwald viscometer) to measure the time it takes for the solution level to fall a fixed distance into a reservoir. By repeating the measurement at different times, a time-resolved progress curve can be determined. Alternatively, viscosity can be measured with a rotating disc rheometer, which can be interfaced to a computer to give real-time measurements. Because exocellulases shorten cel­lulose chains by only two glucoside units per enzyme cycle, and because they are likely to be blocked after only a few steps by chemical modification, exocellulases produce very little change in viscosity. Thus, by comparing activity viscometrically and colorimetrically, exo — and endocellulolytic activity can be resolved [68].

The present invention further discloses designer Calvin-cycle-channeled and photo — synthetic-NADPH (reduced nicotinamide adenine dinucleotide phosphate)- enhanced pathways, associated designer DNA constructs (designer genes), and designer transgenic photosynthetic organisms for photobiological production of butanol and related higher alcohols from carbon dioxide and water. In this context throughout this specification as mentioned before, a “higher alcohol” or “related higher alcohol” refers to an alcohol that comprises at least four carbon atoms, including both straight and branched higher 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 photosynthetic organisms such as designer transgenic algae and oxyphotobacteria (including cyanobacteria and oxychlorobacteria) comprise designer Calvin-cycle-channeled and photosynthetic NADPH-enhanced pathway gene(s) and biosafety-guarding technology for enhanced photobiological production of butanol and related higher alcohols from carbon dioxide and water.

Photosynthetic water splitting and its associated proton gradient-coupled elec­tron transport process generates chemical energy intermediate in the form of ade­nosine triphosphate (ATP) and reducing power in the form of reduced nicotinamide adenine dinucleotide phosphate (NADPH). However, certain butanol-related meta­bolic pathway enzymes such as the NADH-dependent butanol dehydrogenase (GenBank accession numbers: YP_148778, NP_561774, AAG23613, ZP_05082669, ADO12118, ADC48983) can use only reduced nicotinamide adenine dinucleotide (NADH) but not NADPH. Therefore, to achieve a true coupling of a designer path­way with the Calvin cycle for photosynthetic production of butanol and related higher alcohols, it is a preferred practice to use an effective NADPH/NADH conver­sion mechanism and/or NADPH-using enzyme(s) (such as NADPH-dependent enzymes) in construction of a compatible designer pathway(s) to couple with the photosynthesis/Calvin-cycle process in accordance with the present invention.

According to one of the various embodiments, a number of various designer Calvin-cycle-channeled pathways can be created by use of an NADPH/NADH con­version mechanism in combination with certain amino-acids-metabolic pathways for production of butanol and higher alcohols from carbon dioxide and water. The Calvin-cycle-channeled and photosynthetic-NADPH-enhanced pathways are con­structed typically with designer enzymes that are selectively expressed through use of designer genes in a host photosynthetic organism such as a host alga or oxypho- tobacterium for production of butanol and higher alcohols. A list of exemplary enzymes that can be selected for use in construction of the Calvin-cycle-channeled and photosynthetic-NADPH-enhanced pathways are presented in Table 1. As shown in Figs. 4-10, the net results of the designer Calvin-cycle-channeled and photosyn­thetic NADPH-enhanced pathways in working with the Calvin cycle are production of butanol and related higher alcohols from carbon dioxide (CO2) and water (H2O) using photosynthetically generated ATP (adenosine triphosphate) and NADPH (reduced nicotinamide adenine dinucleotide phosphate). A significant feature is the innovative utilization of an NADPH-dependent glyceraldehyde-3-phosphate dehy­drogenase 34 and a nicotinamide adenine dinucleotide (NAD)-dependent glyceral- dehyde-3-phosphate dehydrogenase 35 to serve as a NADPH/NADH conversion mechanism that can convert certain amount of photosynthetically generated NADPH to NADH which can then be used by NADH-requiring pathway enzymes such as an NADH-dependent alcohol dehydrogenase 43 (examples of its encoding gene with GenBank accession numbers are: BAB59540, CAA89136, NP_148480) for pro­duction of butanol and higher alcohols.

More specifically, an NADPH-dependent glyceraldehyde-3-phosphate dehy­drogenase 34 (e. g., GenBank accession numbers: ADC37857, ADC87332, YP_003471459, ZP_04395517, YP_003287699, ZP_07004478, ZP_04399616) catalyzes the following reaction that uses NADPH in reducing 1,3- Diphosphoglycerate (1,3-DiPGA) to 3-Phosphoglyaldehyde (3-PGAld) and inor­ganic phosphate (Pi):

1,3 — DiPGA + NADPH + H + ^ 3 — PGAld + NADP+ + Pi (3)

Meanwhile, an NAD-dependent glyceraldehyde-3-phosphate dehydrogenase 35 (e. g., GenBank: ADM41489, YP_003095198, ADC36961, ZP_07003925, ACQ61431, YP_002285269, ADN80469, ACI60574) catalyzes the oxidation of 3-PGAld by oxidized nicotinamide adenine dinucleotide (NAD+) back to1,3- DiPGA:

3 — PGAld + NAD+ + Pi ^ 1,3- DiPGA + NADH + H+ (4)

The net result of the enzymatic reactions (3) and (4) is the conversion of photo­synthetically generated NADPH to NADH, which various NADH-requiring designer pathway enzymes such as NADH-dependent alcohol dehydrogenase 43 can

butanol and related higher alcohols. When there is too much NADH, this NADPH/NADH conversion system can run also reversely to balance the supply of NADH and NADPH. Therefore, it is a preferred practice to innovatively utilize this NADPH/NADH conversion system under control of a designer switchable promoter such as nirA (or Nial for eukaryotic system) promoter when/if needed to achieve robust production of butanol and related higher alcohols. Various designer Calvin — cycle-channeled pathways in combination of a NADPH/NADH conversion mecha­nism with certain amino-acids-metabolism-related pathways for photobiological production of butanol and related higher alcohols are further described hereinbelow.

The synthesis of biodiesel from Jatropha oil has been investigated in supercritical methanol and ethanol, without using any catalysts, from 200 to 400°C at 200 bar

[68] . It is found that for the synthesis of biodiesel in supercritical alcohols with an optimum molar methanol oil ratio of 50:1, very high conversions (>80%) were obtained within 10 min and nearly complete conversions within 40 min. The con­version into ethyl esters is higher than that of methyl esters [68] .

Tang et al. [86] studied transesterification of the crude Jatropha oil which was catalyzed by micro-NaOH (0.2 to 0.5 to 0.8 wt-%o) in supercritical methanol. When the catalyst content, reaction temperature, and molar ratio of methanol to oil were developed at 0.8 wt-%o NaOH, 534 K, 7.0 MPa, and 24:1, respectively, the methyl ester yield reached 90.5% within 28 min.

The life cycle energy analysis of Jatropha biodiesel production was conducted by evaluating direct energy input (such as electricity, diesel, gasoline, fuel oil, palm fiber, palm shell, etc.) and indirect energy input (energy accumulated in fertilizers, agrochemicals, and chemical production, excluding equipment and machinery used in the processes). The net energy value (NEV) and the net energy ratio (NER) can be estimated. The NEV is a measure of the energy gain or loss from the biodiesel used, which is defined as the energy content of the biodiesel minus the nonrenew­able energy used in the life cycle of the biodiesel production [63]. The NER is a ratio of energy output to total energy input for the life cycle of the product [64].

Prueksakorn and Gheewala [64] calculated the energy consumption in every pro­cess in producing 1 kg of Jatropha biodiesel. The energy analysis results of the present situation of Jatropha biodiesel production compared to palm oil methyl ester is shown in Table 5. The results show that the selected biodiesel production process determines energy efficiency and environmental impacts.

9 Conclusions

High cost of biodiesel production is the major impediment to its large-scale com­mercialization. Methods to reduce the production cost of Jatropha biodiesel must be developed. One way to reduce production costs is to increase the added values of protein-rich Jatropha seedcakes, by-product of oil extraction, through detoxification process. The development of integrated biodiesel production process and the detoxification process results in two products, namely biodiesel and protein-rich seedcakes that can be used for animal feed. Assuming that an average seed yield on land of 5 tons/ha/year (2 tons/acre/year) could be achieved, the estimated theoretical maximum yield of biodiesel would be 750 kg/acre/year and seedcake products would be 500 kg/acre/year.

Since the cost and efficiency of the selected process will be closely correlated with the production for a long time and affect the capital and operating costs and the environmental load of the product, selecting an appropriate process for the biodiesel production is a critical decision. There are still many future potential improvement of biodiesel production of J. curcas. These include (1) development of better and cheaper oil extraction and postreaction processing methods; (2) development of better and cheaper catalysts; (3) improvements in current technology for producing high-quality biodiesel with cheaper cost production; (4) development of technology to use methanol/ethanol in in situ extraction and transesterification; (5) develop­ment of technique to improve fuel stability of Jatropha biodiesel; (6) conversion of by-products, such as glycerol and seedcake to useful and value-added products, such as methanol and ethanol or glycerol tert-butyl ether (GTBE); and (7) develop­ment of integrated Jatropha biodiesel processing and detoxification process.

LCA has become an important decision-making tool for promoting alternative fuels because it can systematically analyze the fuel life cycle in terms of energy efficiency and environmental impacts. LCA analysis shows that the selected biodie­sel production process determines energy efficiency and environmental impacts of Jatropha biodiesel production.

Depending on the degree of drying during the pre-treatment step, the state of the microalgal cells during lipid extraction can be either a wet paste (approximately 10-30 wt.% solid) or dried powder. During lipid extraction, lipid molecules are extracted out of the microalgal cells and separated from the cellular matrix.

The isolated lipids are often referred to as extracts or analytes. The selected lipid extraction method has to be lipid-specific in order to minimize the co-extraction of non-lipid contaminants (proteins, carbohydrates) and be able to exhibit some degree of selectivity towards biodiesel-convertible neutral lipid fractions (acylglycerols) in order to minimize the downstream removal of unusable lipid components (polar lipids and non-saponifiable neutral lipids) [39]. Additionally, the method should be efficient (both in terms of time and energy), non-reactive with the lipids, and rela­tively safe [30] . The practice of completely drying microalgal wet paste prior to lipid extraction is energetically prohibitive and needs to be avoided. As a conse­quence, the selected lipid extraction method needs to be effective when applied directly to microalgal wet paste. Two of the most commonly used microalgal lipid extraction technologies are reviewed: traditional organic solvent extraction and emerging supercritical fluid extraction.

Because of the coastal plain soils advanced age, the single most important soil quality issue to improve, arguably, is the low soil SOC contents (Fig. 4). While most of the OC from crop residue is lost within a few months [72]; the logical remedy would be to increase long-term SOC by applying a biochar that has recalcitrant properties (Fig. 11). Biochars suited to long-term C storage in soils have highly aromatic composition [45, 87] and black carbon with low O/C ratios (0.2-0.4, [68, 97]).

To design biochar with these properties, feedstocks should be pyrolyzed at high temperature (500-700°C) leading to biochar composed of poly-condensed aromatic structures [7,48] and O/C ratio similar to charcoal (0.2-0.4, [49]). A good example of an appropriate feedstock choice is pecan shells, which after high temperature pyrolysis at 700°C, had 58% C in aromatic structures and an atomic O/C ratio of

0. 02 [73]. After 67 days of laboratory incubation in a Norfolk E horizon, pecan shell biochar (700°C) had the lowest CO2 evolved when compared to the control and several raw crop residues (Table 3). In fact, its CO2 mass evolved was similar to soil treated with hardwood shavings. These are laboratory results that were obtained only after few months of biochar incubation in the sandy Norfolk soil. But, the rela­tive difference in CO2 evolution suggests that if pecan shells were pyrolyzed at a high temperature (700°C), they would serve as a suitable designer biochar to increase C sequestration in the sandy Norfolk soil. Other feedstocks (i. e., hard­woods, shells from other nut crops, etc.) may also be suitable, but should also have high aromaticity and atomic O/C ratios of <0.4.

Another characteristic for biochar stability in soils is its volatile matter/fixed carbon (VM/FC) ratio [3] . Biochar with VM/FC of 0.5-1.0 are speculated to be stable in soils [3]. As an acceptable index of biochars longevity in soil, the actual relationship between its VM/FC ratio with CO2 evolution from soils/culture media needs further evaluation.