Although the cost of agricultural residues is much lower than the cost of other conventional substrates such as corn, the process of butanol production from residues requires additional process steps. One such step is the hydrolysis of residues to simple sugars prior to their conversion to butanol. Unless these residues are converted to simple sugars, they cannot be used by butanol producing cultures. The hydrolysis first requires pretreatment using dilute sulfuric acid or dilute alkali at 121°C or higher. This is done to make cellulosic fibers accessible to enzymes. For our studies dilute sulfuric acid was used in order to make the process simple [57]. Following pretreatment, the biomass was hydrolyzed with enzymes which were then fermented to butanol. Butanol or ABE was then recovered using the gas stripping technique [57] . It is suggested that any of the product recovery techniques described by

Recovery (Step III)

An integrated/combined process of hydrolysis (Step I), fermentation
(Step II), and product recovery (Step III)

Fig. 1 A schematic diagram of production of butanol/ABE from lignocellulosic biomass employ­ing Clostridium beijerinckii P260. (a) SHFR (separate hydrolysis fermentation and recovery pro­cess); (b) SSFR (simultaneous saccharification, fermentation, and recovery process; also known as an integrated process)

Maddox [37] or Qureshi [48] can be used for ABE removal. The overall process of production of butanol from cellulosic biomass by this process requires three sepa­rate steps (hydrolysis, fermentation, and recovery) and is called “Separate Hydrolysis, Fermentation, and Recovery (SHFR).” Figure 1a shows a schematic diagram of butanol production by this process.

2.1 Primary Wall Modification: Yield, Growth, and Porosity

The primary cell wall allows the plant to expand and elongate while maintaining mechanical strength and support. Elongation rates and cell size and shape are largely governed by primary wall plasticity and cell-cell adhesions. Thus, the primary wall has a direct impact on plant yield. Hence, growth zones are rich in modifying enzymes such as cellulases, xylanases, xyloglucanases and pectin modifying enzymes that allow cell expansion [81].

2.1.1 Cell Wall Modulation and Increased Plant Biomass via Overexpression of Glycoside Hydrolases

Plants synthesize an extensive family of glycoside hydrolases termed endogluca — nases. Some are secreted to the extracellular matrix during growth and some form part of the cellulose synthase complex. Endoglucanases are naturally involved in the process of plant cell wall development and degradability. These enzymes are capa­ble of hydrolyzing noncrystalline cellulose and xyloglucans, enabling xyloglucan — cellulose matrix remodeling and cell wall plasticity during growth and development via wall loosening [18, 113]. The Arabidopsis endo-(1-4)-b-glucanase protein (Cell) accumulates in young, expanding tissues, playing a key role in cell elonga­tion of rapidly growing tissues [109-112]. Heterologous overexpression of cell in poplar trees or of poplar endoglocanase (PaPopCell) in Arabidopsis resulted in longer internodes, increased cell elongation, and subsequent biomass accumulation [89, 111]. Mechanical analysis, studying leafblade extension at constant load and breakage at changing load was conducted. An elongation vs. load curve demonstrated higher elongation rates in transgenic Arabidopsis leaf blades when compared to wild type [89 ] . As highly crosslinked materials are more resistant to elongation under specific load, it is speculated that the cell wall of these transgenic plants con­tained less crosslinked material [89]. It is proposed that noncrystalline glucan chains intercalated with hemicelluloses are unraveled by endoglucanases, lowering the amount of tethered xyloglucans, ultimately increasing cell wall plasticity and allow­ing for continued cell wall deposition ] 56 ] . Similar results were obtained upon expression of Aspergillus niger xyloglucanase in poplar trees. Both stem length and cellulose content increased, but at the same time, the growth zone had reduced Young’s elastic modulus [88]. Furthermore, when plants were placed horizontally, the basal regions of stems of these transgenic poplars failed to bend upward due to lower tensile strength in newly formed wood [6] .

Overexpression of poplar endoglucanase PaPopCell in the leguminous tropical tree Paraserianthes falcataria. resulted in increased biomass. Disturbance of the biological clock by altering the closing movements of the leaves was also detected [55]. Thus in summary, overexpression of endoglucanases can accelerate growth, but may also result in undesirable effects. Maximal benefits may be achieved by using specific promoters for targeting the gene product to specific organs or for restricting expression to specific developmental stages.

In addition to the assays above (colorimetry, viscometry, and chromatography), there are several other physical methods that have been used to quantify breakdown. While most of these methods have seen limited use, they have the advantage that they can directly monitor breakdown in real time. One particularly interesting approach is time-resolved isothermal batch calorimetry [51, 54]. Slurried cellulosic substrates are prepared, and enzymatic degradation is monitored by heat release as a function of time. In addition to following hydrolysis in real time, this method is well-suited for complex substrates (e. g., corn stover). Another real-time method for monitoring cellulose degradation is direct determination of mass-decrease from immobilized cellulose using a quartz crystal microbalance [35] .

3.1 Plate Assays and Live-Cell Assays

In protein engineering applications, it is often desirable to use a screen that is qualitative, so that a small number of active enzymes (or enzyme combinations) can be resolved from a large background of inactive enzymes. Several methods can be used to screen colonies for cellulose degradation activity (reviewed in [60]). One of the most common plate screening methods uses celluloses with tightly bound dyes and fluorophores. Congo red dye interacts tightly with b-1,4-D-glucans including chemically modified carboxymethyl cellulose (CMC). Because CMC is large and does not diffuse in agar, CMC-bound dye will remain uniformly immobilized in the absence of cellulase activity. Cellulose hydrolysis releases the dye; subsequent diffusion results in a dye-free halo around colonies expressing active cellulase. The halo radius gives a semiquantitative indication of activity. In a similar approach, unlabelled soluble cellulose (e. g., CMC) can be included in the agar, and after incu­bation to allow for colony growth and cellulase activity, the remaining CMC is precipitated on the plate using ethanol, acetone, or cetylammonium bromide (CTAB). Precipitated CMC renders the plate opaque; colonies expressing active cellulase can be identified as having a clear halo [60] .

Alternatively, cellulase activity can be selected for using agar plates in which the only source of carbon is cellulose. This approach requires both high purity cellulose (organic contaminants may provide an alternative carbon source) and a uniformly high DP (contaminating cellodextrins would also short-circuit the selection). This approach has also been used in liquid culture to select for yeast strains producing active cellulases on their surface. An advantage of this method is that saccharification products can be directly fermented by the yeast to produce ethanol, a process referred to as “consolidated bioprocessing” [18, 40, 72].

According to one of the various embodiments, a designer Calvin-cycle-channeled pathway is created that takes the Calvin-cycle intermediate product, 3-phospho — glycerate, and converts it into isobutanol by using, for example, a set of enzymes consisting of (as shown with numerical labels 34, 35, 03-05, 53-55, 42, 43 (or 44) in Fig. 6): NADPH-dependent glyceraldehyde-3-phosphate dehydrogenase 34, NAD-dependent glyceraldehyde-3-phosphate dehydrogenase 35, phosphoglycerate mutase 03 , enolase 04 , pyruvate kinase 05, acetolactate synthase 53, ketol-acid reductoisomerase 54, dihydroxy-acid dehydratase 55, 2-keto acid decarboxylase 42, and NAD-dependent alcohol dehydrogenase 43 (or NADPH-dependent alcohol dehydrogenase 44). The net result of this pathway in working with the Calvin cycle is photobiological production of isobutanol ((CH3)2CHCH2OH) from carbon diox­ide (CO,) and water (H, O) using photosynthetically generated ATP and NADPH according to the following process reaction:

4CO2 + 5H2O ^ (CH3)2CHCH2OH + 6O2 (7)

According to another embodiment, a designer Calvin-cycle-channeled pathway is created that takes the intermediate product, 3-phosphoglycerate, and converts it into 3-methyl-1-butanol by using, for example, a set of enzymes consisting of (as shown with the numerical labels 34, 35, 03-05, 53-55, 40, 38, 39, 42, 43 (or 44/57) in Fig. 6): NADPH-dependent glyceraldehyde-3-phosphate dehydrogenase 34, NAD-dependent glyceraldehyde-3-phosphate dehydrogenase 35, phosphoglycerate mutase 03 , enolase 04 , pyruvate kinase 05, acetolactate synthase 53, ketol-acid reductoisomerase 54, dihydroxy-acid dehydratase 55, 2-isopropylmalate synthase 40, 3-isopropylmalate dehydratase 38, 3-isopropylmalate dehydrogenase 39,2-keto acid decarboxylase 42, and NAD-dependent alcohol dehydrogenase 43 (or NADPH- dependent alcohol dehydrogenase 44; or more preferably, 3-methylbutanal reductase 57). The net result of this pathway in working with the Calvin cycle is photobiologi­cal production of 3-methyl-1-butanol (CH3CH(CH3)CH2CH2OH) from carbon diox­ide (CO2 ) and water (H2 O) using photosynthetically generated ATP and NADPH according to the following process reaction:

10CO2 + 12H2O ^ 4CH3CH(CH3)CH2CH2OH + 15O2 (8)

These designer pathways (Fig. 6) share a number of designer pathway enzymes with those of Figs. 4 and 5, except that a 3-methylbutanal reductase 57 is preferably used for production of 3-methyl-1-butanol; they all have a common feature of using an NADPH-dependent glyceraldehyde-3-phosphate dehydrogenase 34 and an NAD-dependent glyceraldehyde-3-phosphate dehydrogenase 35 as an NADPH/ NADH conversion mechanism to convert certain amount of photosynthetically gen­erated NADPH to NADH which can be used by NADH-requiring pathway enzymes such as an NADH-requiring alcohol dehydrogenase 43 , The net results of the designer photosynthetic NADPH-enhanced pathways (Fig. 6) in working with the

Calvin cycle are also production of isobutanol ((CH3)2CHCH2OH) and/or 3-methyl — 1-butanol (CH3CH(CH3)CH2CH2OH) from carbon dioxide (CO2) and water (H2O) using photosynthetically generated ATP and NADPH.

In situ transesterification [32, 33, 39, 78], a biodiesel production method that uti­lizes the original agricultural products instead of purified oil as the source of trig­lycerides for direct transesterification, eliminates the costly hexane extraction process and works with virtually any lipid-bearing material. It could reduce the long production system associated with preextracted oil and maximize alkyl ester yield. The use of reagents and solvents is reduced, and the concern about waste disposal is avoided.

The experimental results showed that the amount of J. curcas seed oil dis­solved in methanol was approximately 83% of the total oil and the conversion of this oil could achieve 98% under the following conditions: less than 2% moisture content in J. curcas seed flours, 0.3-0.335 mm particle size, 0.08 mol/L NaOH concentration in methanol, 171:1 methanol/oil molar ratio, 45.66°C reaction tem­perature, and 3.02 h reaction time. The use of alkaline methanol as extraction and reaction solvent, which would be useful for extraction oil and phorbol esters, would reduce the phorbol esters content in the Jatropha seedcake. The seedcakes after in situ transesterification is rich in protein and is a potential source of live­stock feed [58].

Life cycle analysis (LCA) is a systems approach that is aimed at evaluating the environmental impact of all processes that contribute to the entire life cycle of the product of interest, carried out in accordance with the ISO 14040 standard [187]. To objectively determine if microalgae biofuel production chains are environmentally sustainable it is deemed important to carry out a full LCA of the relevant process chains for rational boundary conditions. LCA can then be used to identify the “hotspots” in the process chains, i. e. where potential environmental burdens may be dominant, and to identify downstream processes requiring technological improvement and innovation. Most LCAs incorporate impacts assessment such as GHG reduction potential and energy balance; additional impacts of processes such as the nitrogen cycle changes and recycling of spent biomass and nutrients (see biorefinery concept) provide more rational representation of the production process for algae-derived biofuels [106]. Currently, there are no validated life cycle inventories pertaining to microalgae-derived biofuel production process, e. g. water use and elemental and nutrient recycling, that could form bases for realistic scenarios [170].

The outlined limitations notwithstanding several LCA studies have been used to evaluate the potential environmental impacts of microalgae biofuel production systems. Liu and Ma [121] carried out an LCA of methanol production from microalgae and recorded a positive energy balance and reduction of environmental loading compared to equivalent fossil fuels. Campbell et al. [27] also recorded lower cost for producing algae-derived biodiesel with a substantial GHG reduction and a positive energy balance over petroleum diesel. However, they also noted that, for full-scale systems, the economic cost could exceed those for diesel due to inapplicable economies of scale, and the process being highly dependent upon the selection of algae species with high oil yields. For example, Lardon et al. [110] performed an LCA on biodiesel produced from Chlorella vulgaris grown in raceway ponds and recorded a negative energy balance (1.66 MJ of energy input to 1 MJ output).

Kadam [98] established the potential benefits to utilising recycled CO2 towards microalgae production from LCA of algae co-firing scenario. He recorded lowered net fossil energy consumption, SOx and NOx, particulates, CO2 and methane. Reijnders [173] has argued that, after considering the total fuel inputs during the biofuel life cycle (e. g. fossils fuel used for building the facilities and for operational activities such as supplying nutrients, mixing, harvesting and processing), microal — gae-derived biodiesel is inferior to petroleum diesel. He illustrated that both ethanol from sugarcane (161-175 GJ ha-1 year-1) and palm oil from oil palm (142­180 GJ ha-1 year-1) returned higher net energy yield compared to methanol from microalgae (127 GJ ha-1 year-1).

Greenhouse gas emissions as CO2, CH4, and N2O as a result of fossil fuel usage and agricultural activity within the USA have increased 14% between 1990 and 2008 [40]. The agricultural sector was estimated by the US-EPA to contribute approximately 6% of the total GHG emissions. Animal and crop production may account for as much as 70% of the annual global anthropogenic N2O emitted [65]. Globally, N2O is a significant contributor to the emission total (»8%, [35]) and has a global warming potential of 298 times greater than CO2 [41]. The large difference in N2O radiative force with CO2 causes it to have a larger destructive potential to the stratospheric ozone layer [26].

N2O fluxes have been measured in agricultural field, but estimates of their overall contributions to the global GHG budget is difficult to estimate because fluxes have been linked to differences in soil N application, N form, soil pH, soil wetness, and tillage practices [33, 54, 76] . Nevertheless, the sizable hazard that N. O poses for climate change relative to CO2, suggests that it is important to have management strategies available to curtail N2 O production from agricultural soils. This will require both field and laboratory evaluations between feedstock, pyrolysis conditions, and biochar chemical properties on N2O dynamics.

Both field and laboratory studies reported that biochar additions to soil can reduce N2O emissions [42, 67, 83 , 90, 96, 104]. In the field, biochar applications at 20 Mg ha-1 to soybean plots were found to cause a 50% curtailment in N2O emissions [83]. While in the lab, N2O production was suppressed by a variety ofbiochars pro­duced from nut shell wastes and hardwoods [96] and from poultry litter and wood [90]. In fact, both studies employed biochars produced at different temperatures and reported difference in N2O reduction. Neither study reported an over-arching biochar chemical/structural characteristic as responsible for reducing N2O emissions.

Not all biochars will suppress N2O emissions when added to soils, in fact, Spokas and Reicosky [96] reported that two out of 16 biochars stimulated N2O production relative to the control. Similarly, Singh et al. [90] reported that a biochar produced from pyrolyzed poultry manure at 400°C stimulated N. O production. Based on these reports, it is difficult to make suggestions (Fig. 11) to create a biochar tailored to effectively suppress N2O production.

What we do know, however, is that biochars may suppress N2 O production if they have properties that influence N availability [ 90] , decrease soil microbial activity [96] , and improve soil physical properties that promotes aeration [ 58] . Whereas, others have reported soil N2O production can be stimulated with biochar. These conflicting conclusions suggest that additional laboratory and field evalua­tions involving biochars produced from multi-feedstocks and under different pyrolysis conditions are needed.

1.2 Levoglucosan

LG (1,6-anhydro-P-D-glucopyranose) is the most important pyrolytic product of pure cellulose, formed through the depolymerization reaction. The chemistry of LG has long been known, and it can be used as a chiral synthon for the synthesis of

stereoregular polysaccharides possessing biological activities [11, 56]. Moreover, LG can also be hydrolyzed to glucose, providing a potentially rapid route to produce bio-ethanol [14].

Fast pyrolysis of pure cellulose can produce LG with the yield up to 40 wt%, but fast pyrolysis of raw biomass materials would produce much lower LG yield due to the presence of inorganic impurities. Even the minor amounts of alkaline cations would shift the pyrolytic pathways of cellulose, to promote the formation of ring — scission products (such as HAA, HA, etc.) and char on the expense of LG [61, 68]. Hence, it is necessary to use pure cellulose or demineralized biomass for the LG production. Furthermore, some studies pointed out that when small amounts of acids or acidic salts were added to demineralized biomass, the LG yield could be increased. For example, it was reported in an analytical pyrolysis study that, the LG content in the pyrolytic products was 5.3% from raw birch wood, 17.0% from decationized wood, 33.6% from wood impregnated with 1.0% phosphoric acid, and 27.3% from decationized wood adsorbed with iron ions [28].

The main difficulty in LG preparation is not the pyrolysis process, but its isolation from pyrolysis liquids. Due to its high boiling point (386°C), LG could not be simply recovered by distillation. Currently, several methods have been proposed or patented for the purification of LG [44, 57, 81].

Lignocellulosic biomass has emerged as a potential renewable biomass resource for the bioethanol production [92, 117]. The concept is to hydrolyze the cellulose and hemicelluloses fraction of biomass (holocellulose) to recover C5 and C6 sugars and then ferment the sugars to bioethanol [93] . The recovered lignin in the process which has relatively higher heating value in the range of 24-26 MJ/kg is typically used for generating steam or providing the process heat. The biochemical pathways which can be realized at a very moderate process conditions using cellulase and accessory enzymes to convert holocelluloses to fermentable sugars are the most promising ones for large-scale bioethanol production. But the efficiency of this technology is limited due to the complex chemical structure of lignocellulose biomass and the inaccessibility of b-glycosidic linkages to cellulase enzymes because of the low surface area and small size of pores in multicomponent structure. Hence, pre­treatment is nowadays viewed as a critical step in lignocellulose processing [66] .

Pretreatment alters both the structural barrier (removal of lignin and hemicellu — loses) and physical barrier (surface area, crystallinity, pore size distribution, degree of polymerization) which help in improving the accessibility of enzyme for hydro­lysis [38, 67, 83]. It enhances the rate of production and the yield of monomeric sugars from biomass. Pretreatment is among the most costly step in the bioethanol conversion process as it may account for up to 40% of the processing cost. Moreover, it also affects the cost of upstream and downstream processes [69, 91, 130, 135]. Hence, an efficient, less energy intensive and cost-effective pretreatment method is a necessity for producing ethanol at an economically viable cost. Different pretreatment methods are broadly classified into physical, chemical, physicochemical, and biologi­cal processes. The conventional pretreatment, by using acids or alkalis, is associated with the serious economic and environmental constraints due to the heavy use of chemicals and chemical resistant materials [21, 43, 115].

Hydrothermal pretreatment employing subcritical water has attracted much attention because of its suitability as a nontoxic, environmentally benign and inex­pensive media for chemical reactions [63, 76]. One of the most important benefits of using water instead of acid as pretreatment media is that there is no need of acid recovery process and related solid disposal and handling cost [21, 51]. Below the critical point, the ionization constant of water increases with temperature and is about three orders of magnitude higher than that of ambient water. Also, the dielectric constant of water decreases with temperature. A low dielectric constant allows liquid water to dissolve organic compounds, while a high ionization constant provides an acidic medium for the hydrolysis of biomass components via the cleavage of ether and ester bonds and favor the hydrolysis of hemicelluloses [31, 65, 79, 103]. The structural alterations due to the removal of hemicelluloses increase the acces­sibility and enzymatic hydrolysis of cellulose. Enzyme accessibility is increased as a result of the increase in mean pore size of the substrate which enhances the prob­ability of the hydrolysis of glycosidic linkage [42].

Hydrothermal pretreatment is typically conducted in the range of 150-220°C. The temperature range, aiming for the fractionation of hemicelluloses, is decided based on the fact that at temperature below 100°C less/small extent of hydrolytic reaction is observed whereas cellulose hydrolysis and degradation become significant above 210°C [32, 41]. The severity factor (R0) has been used by several researchers to measure the combined effect of temperature and residence time in hot water treatment of biomass processes [1, 88,96]. The severity index is defined as

„ Г t -1001

R =’x exp 1 14T7T}

where t is the residence time in minutes and T is temperature in °C.

Ether bonds of the hemicelluloses are most susceptible to breakage by the hydronium ions. Depending on the operational conditions, hemicelluloses are depolymerized to oligosaccharides and monomers, and the xylose recovery from biomass can be as high as 88-98%. For example, Suryawati et al. have reported 90% removal of hemi­celluloses from Kanlow switchgrass at 200°C [116] . Acetic acid is also generated from the splitting of thermally labile acetyl groups of hemicelluloses. In further reactions, the hydronium ions generated from the autoionization of acetic acid also acts as catalyst and promotes the degradation of solubilized sugars. In fact, the forma­tion of hydronium ions from acetic acid is much more than from water [32, 41] .

The low pH (< 3) of the medium causes the precipitation of solubilized lignin and also catalyzes the degradation of hemicelluloses. To avoid the formation of inhibitors, the pH should be kept between 4 and 7 during the pretreatment. This pH range minimizes the formation of monosaccharides, and therefore the formations of degradation products that can further catalyze hydrolysis of the cellulosic material during pretreatment [10, 41, 58, 67, 82, 110, 128]. Maintaining the pH near neutral (5-7) helps in avoiding the formation of fermentation inhibitors during the pretreatment. The addition of small amount of K2CO3 increases the glucan digestibility even at low pretreatment temperatures (150-175°C) [66]. Figure 11 shows the SEM images

of untreated and hydrothermally pretreated switchgrass in a flow-through reactor where additional pores created after the pretreatment can be seen [62].

In general, the concentrations of solubilized products are lower in hydrothermal pretreatment compared to the steam pretreatment [ 9 ] . Since the hot compressed water is used instead of steam, the latent heat of evaporation is saved which makes it easier to apply for a continuous process [57]. Yang and Wyman have reported that flow-through process fractionated more hemicelluloses and lignin from corn stover as compared to batch system under the conditions of similar severity [132]. In a flow-through system, the product is continuously removed from the reactor which reduces the risk on condensation and precipitation of lignin components, making the biomass less digestible. The soluble lignin compounds are very reactive at the pretreatment temperature and if not removed rapidly part of these compounds recondense and precipitate on the biomass [68, 96].

In a subcritical water pretreatment study, the microcrystalline cellulose (MCC) pretreated at 315°C in a continuous flow reactor for about 4 s of residence time showed nearly threefold increase in the initial enzymatic reactivity as compared to the untreated MCC at 3.5 FPU/g of glucan enzyme loading [63]. The percentage crystallinity of MCC slightly increased after the subcritical water pretreatment and remained high (>81%) throughout the treatment range (200-315°C). Increase in percentage crystallinity is generally attributed to the hydrolysis and removal of the amorphous part of cellulose during pretreatment [63]. The DPv of cellulose reduced with the pretreatment temperature and sharp decline was observed in cellulose samples pretreated at 315°C (Fig. 12). The DPv of untreated MCC was 327. It decreased with temperature as expected, but reduced rapidly for treatment above 300°C.

N. Escalona, R. Garcia, and P. Reyes

Abstract The gasification of biomass followed by a Fischer-Tropsch Synthesis (FTS) is a good alternative for synthesis of gasoline and/or diesel. However, this process may be considered as a high-cost technology, depending on crude oil and biomass raw material prices. The viability may be increased depending on the value of biomass, cost of transportation of biomass and the separation (conditioning) of gases produced in the gasification (elimination of CO2, CH4, N2 and others). Nevertheless, this gas mixture “called biosyngas” may be used in the FTS without pre-conditioned for producing gasoline and/or diesel. The main focus of this chapter will be on the latest investigations in the FTS carried out in a microreactor from a simulated biosyngas (without conditioning), as an alternative to decrease the cost of this process. This chapter reports results of catalytic activity and characterization of Fe/SiO2 and Co/SiO2 catalysts and Cu, Re, Ru and Zn promoted Co/SiO2 catalysts.

1 Introduction

Global warming has pushed the necessity to reduce the emissions of gases responsible for the Greenhouse effect, mainly CO2. Public transportation produces approximately a 22% of the total CO2 , because of the use of fossil fuels. Nowadays there is great interest in the search of environmentally friendly fuels, in order to decrease the green­house gas emission. Biomass appears as an attractive alternative since it is a clean, renewable and sustainable energy resource. Different transformation routes have been used to convert biomass to liquid or gaseous fuels [1-5]. The first step of the overall process (biomass-to-liquid fuels, BTL) includes biomass gasification to yield a gas mixture mainly containing H2, CO, CO2 and CH4, called biosyngas, which can then be

N. Escalona (H) • R. Garcia • P. Reyes

Universidad de Concepcion, Facultad de Ciencias Quimicas,

Casilla 160, Concepcion, Chile

e-mail: nescalona@udec. cl

J. W. Lee (ed.), Advanced Biofuels and Bioproducts, DOI 10.1007/978-1-4614-3348-4_13, 209

© Springer Science+Business Media New York 2013

adjusted to the desired H2/CO ratio prior to being converted by the Fischer-Tropsch synthesis (FTS) [4, 5]. The FTS is the most important catalytic process in the synthesis of gasoline and/or diesel from syngas (H2/CO) mixture [6]. The first commercial FTS plants operated with syngas mixtures produced from coal gasification, whereas the modern FTS units use CO/H2 mixtures mainly obtained from the methane steam reforming, where the molar H2/CO ratio is around of 2, which is much higher than the H2/CO molar ratio close to 1 derived from biomass gasification [7, 8]. The metals used as catalysts in the FTS traditional are mainly Fe and Co promoted by K, Re, Cu or Zn [9, 10]. The hydrocarbons produced according to the FT synthesis using biosyngas mixtures are free of S and N, and in principle can be considered neutral in the CO2 greenhouse effect. Significant efforts have been devoted to hydrocarbon synthesis using syngas mixtures produced according to the steam methane reforming technol­ogy [6-13]; however, the literature describing the production ofhydrocarbons from biosyngas is quite limited [2, 14-19]. In this context, Tijmensen et al. [2] explore the technical and economic feasibility using an integrated biomass gasification and Fischer-Tropsch synthesis (BIG-FT). The results show that an integrated plant (BIG-FT) can only be profitable if the price of a barrel of oil exceeds US $50 per bar­rel, in agreement with Hamelinck et al. [5]. Additionally, the later author found that the cost of producing diesel from biomass by integrated gasification and FTS is 16 €/ GJ. This value may decrease in the future to 9 €/GJ by further development and research in integrated technologies for the gasification and FTS. In these studies the biosyngas mixture used is pre-conditioned before being introduced to the FT reactor. In other words, the CO2, CH4, N2 and other gases are removed from the mixture of feed. The cost of the gas removal processes has increased significantly. An alternative to lower the cost of production is to use biosyngas directly, without being pre-condi­tioned, in the FT synthesis. In this aspect, Jun et al. [14] studied the synthesis of liquid hydrocarbons over Fe/Cu/Al/K (100/6/16/4) catalysts using the biosyngas as feed. They found that this catalyst has a high fuel yield and selectivity to olefins. Recently, Co/SiO2, CoM/SiO2 (M=Cu, Re, Ru and Cu) and Fe/SiO2 catalysts were studied in the SFT from simulated biosyngas feed [15, 16, 19] . These catalysts display a high CO conversion activity and the formation of hydrocarbon with Fe/SiO2 and Co/SiO2 catalysts was centred around of C8-15 chain length, while the formation of hydrocarbon by Co/SiO2 promoted was centred around of C8-9 chain length. Therefore, the focus of this work is on the latest research in the FTS over Fe/SiO. , Co/SiO2 and Co/SiO2- promoted using a model of biosyngas as feed.