Depending on the composition and properties of raw materials, the selection and conditioning of the appropriate bacterium strain are essential. In order to improve the economic efficacy of ABE fermentation, the butanol ratio is to be increased by eliminating the production of other byproducts such as acetone and specific mutants are to be developed which show high butanol tolerance, high productivity or other advantageous properties.

Harada [28, 29] isolated a new strain of Clostridium (Cl. Madisonii) which produced BuOH amounting to 28.7% of the initial total sugar and the fermented broth included 1.38% BuOH. The age of the culture also plays important role in the productivity. By using older inoculated bacteria, the production of acetone increased and the ratio of BuOH to Me2CO decreased from 2.24 to 1.88 [30]. Harada [31] concluded that the seed culture at the last stage of the acid- decreasing phase gave the best yield as inoculum in the main fermentation. Butanol-resistant mutants have been isolated by Hermann from soil which produced significantly higher solvent concentrations (about 30%) than the wild-type strain [32]. The sporulation-deficient (spo) early-sporulation Clostridium acetobutylicum P262 mutants produced higher solvent yields than did the spoB mutant which was a late-sporulation one. In conventional batch fermenta­tion, the wild-type strain produced 15.44 g L-1 of solvents after 50 h at a productivity of 7.41 g L-1 d-1 of solvents. The spoA2 mutant produced 15.42 g L-1 of solvents at a productivity of 72.4 g L-1 d-1 of solvents with a retention time of 2.4 h in a continuous immobilized cell system employing a fluidized bed reactor [33].

Using two different types of Clostridia to improve the productivity of each (acidogenic and solventogenic) phase is also known. Bergstroem and Foutch [34] improved the BuOH pro­duction from sugars by combining two cultures of Clostridium: one that produces butyric acid, and another that converts butyrate to BuOH. Thus, C. butylicum NRRL B592 and C. pasteur — ianum NRRL B598 were cultured together in thioglycolate medium containing 2.5% added glucose and a CaCO3 chip to maintain pH, at 37 °C under anaerobic conditions. The yield of BuOH was 20 % more as compared to the value when C. butylicum was cultured alone.

Initiation of gene-structure changes by destructive methods such as irradiations or chemicals followed by selection is a well known method in the production of highly effective Cl. Acetobutylicum strains. Yasuda [35] heated ABE producing microorganisms at 100 °C to destroy all vegetative forms except spores which were kept at -10 °C, then treated with electric discharge in vacuum by using 50,000 V and 0.002 A DC for stimulation. High-yield butanol producing Clostridium strain was prepared through irradiation of the wild strain with 60Co y-rays at an irradiation dosage of 100-1,000 Gy and a dosage rate of 3-5 Gy/min [36].

Chemical mutation with N-methyl-N’-nitrosoguanidine is one of the most frequently used method to produce excellent ABE fermenting strains. Hermann et al [37] prepared a strain of C. acetobutylicum that hyperproduces acetone and BuOH by mutation of C. acetobutylicum IFP903. A new mutant (CA101) of C. pasteurianum prepared in this way could produce 2.1 g BuOH/L in 2 days. By using the parental strain, the production of BuOH was only 0.6 g/L [38]. The C. acetobutylicum strain 77 was isolated from the parent strain ATCC 824 with the abovementioned method in the presence of butanol. The mutant grew more rapidly (|j = 0 69 h-1) than the parent strain (p = 0 27 h-1) and, at the stationary phase, the cell dry weight of mutant strain was about 50% higher than that of the parent strain. Strain 77 metabolised glucose faster than wild strain and solvent production started earlier with higher specific production rates than the parent strain. From 65 g of glucose, 20 g L-1 of solvents (butanol, 14 5 g; acetone, 3 5 g; ethanol, 2 g) were formed by the wild strain in 53 h, whereas the mutant used 75 g of glucose and excreted nearly 24 g L-1 of solvents (butanol, 15.6 g; acetone, 4.5 g; ethanol, 3.7 g) in 44 h [39]. A frequently used chemical to inititate mutation in C. Acetobuty — licum strains is methanesulfonic acid ethyl ester (EMS). EMS is effective in inducing mutants resistant to ampicillin, erythromycin, and butanol (15 g/l). Optimal mutagenesis occurs at 85­90% kill corresponding to a 15 minute exposure to 1.0% (v/v) EMS at 35 C. At optimal condi­tions, the frequency of resistant mutant CFU/ total CFU plated increases 100-200 fold [40].

Genetical engineering opened unlimited perspectives in the preparation of ABE ferment­ing microorganisms. Genetically modified C. Acetobutylicum, E. Coli and S. Cereviase and other microorganisms play important role in the future production of ABE solvents under more convenient conditions than in classical ABE fermentation. The acetoacetate decarboxylase gene (adc) in the hyperbutanol-producing industrial strain Clostridium acetobutylicum EA 2018 was disrupted when the butanol ratio was increased from 70 to 80.05%, while acetone production decreased to approx. 0.21 g/L in the adc-disrupted mu­tant (2018adc). Regulation of the electron flow by addition of methylviologen altered the carbon flux from acetic acid production to butanol production in strain 2018adc, which resulted in an increased butanol ratio of 82% and a corresponding improvement in the overall yield of butanol from 57 to 70.8% [41].

Larossa and Smulski found genes involved in a complex that is a three-component proton motive force-dependent multidrug efflux system to be involved in E. coli cell response to butanol by screening of transposon random insertion mutants. Reduced production of the AcrA and/or AcrB proteins of the complex confers increased butanol tolerance [42]. Green and Bennett subcloned the genes coding for enzymes involved in butanol or butyrate formation into a novel Escherichia coli-Clostridium acetobutylicum shuttle vector constructed from pIMPI and a chloramphenicol acetyl transferase gene [43]. The resulting replicative plasmids, referred to as pTHAAD (aldehyde/alcohol dehydrogenase) and pTHBUT (butyrate operon), were used to complement C. acetobutylicum mutant strains, in which genes encoding aldehyde/alcohol dehydrogenase (aad) or butyrate kinase (buk) had been inactivated by recombination with Emr constructs. Complementation of strain PJC4BK (buk mutant) with pTHBUT restored butyrate kinase activity and butyrate production during exponential growth. Complementation of strain PJC4AAD (aad mutant) with pTHAAD restored NAD(H)- dependent butanol dehydrogenase activity, NAD(H)-dependent butyraldehyde dehydrogen­ase activity and butanol production during solventogenic growth [43]. Shen and Liao constructed an Escherichia coli strain that produces 1-butanol and 1-propanol from glucose [44]. First, the strain converts glucose to 2-ketobutyrate, a common keto-acid intermediate for isoleucine biosynthesis. Then, 2-ketobutyrate is converted to 1-butanol via chemicals involved in the synthesis of the unnatural amino acid norvaline. The synthesis of 1-butanol is improved through deregulation of amino-acid biosynthesis and elimination of competing pathways. The final strain demonstrated a production titre of 2 g/L with nearly 1:1 ratio of butanol and propanol [44]. Green et al [45] made recombinant thermophilic bacteria of the family Bacilla — ceae which have been engineered to produce butanol and/or butyrate. The Bacillaceae is preferably of the genus Geobacillus or Ureibacillus [45]. Young et al described a method of modifying prokaryotic and eukaryotic hosts for the fermentation production of aliphatic alcohols. Elements of the gene for a CAAX proteinase (prenylated protein-processing C- terminal proteinase) are used to increase alcohol tolerance. This can be used in combination with other changes to increase alcohol tolerance [46]. Fermenting with modified eukaryotic cells in a suitable fermentation broth, wherein butanol and ethanol are produced at a ratio between 1:2 to 1:100, is described by Dijk et al. [47]. Since fermentations with yeasts do not require sterile environment, genetically modified yeasts are very prosperous microrganisms in ABE fermentation. Yeast cells capable of producing butanol and comprising a nucleotide sequence encoding a butyryl-CoA dehydrogenase and at least one nucleotide sequence encoding an electron transfer flavoprotein were described by Mueller et al. [48].

Catalytic hydroprocessing of liquid biomass is a technology currently under developed and there is a lot of room for optimization. For example there are not many commercial catalysts specifically designed and developed for such applications, while conventional commercial cat­alysts, employed for catalytic hydroprocessing of refinery streams, are used instead. Common
hydrotreating catalysts employed contain active metals on alumina substrate with increased surface area. The most known commercial catalysts employ Cobalt and Molybdenum (CoMo) or Nikel and Molybdenum (NiMo) in alumina substrate (Al2O3) as shown in Figure 4.

(b)

Hydrotreating catalysts are dual action catalytic material, triggering both hydrogenation and cracking/isomerization reactions. On one hand hydrogenation takes place on the active metals (Mo, Ni, Co, Pd, Pt) which catalyze the feedstock molecules rendering them more ac­tive when subject to cracking and heteroatom removal, while limiting coke formation on the catalyst. Furthermore hydrogenation supports cracking by forming an active olefinic inter­mediate molecule via dehydrogenation. On the other hand both cracking and isomerization reactions take place in acidic environment such as amorphous oxides (SiO2 — Al2O3) or crys­talline zeolites (mainly z-zeolites) or mixtures of zeolites with amorphous oxides.

During the first contact of the feedstock molecules with the catalyst, a temperature increase is likely to develop due to the exothermic reactions that occur. However, during the continu­ous utilization of the catalyst and coke deposition, the catalyst activity eventually reduces from 1/3 to 1/2 of its initial one. The catalyst deactivation rate mainly depends on tempera­ture and hydrogen partial pressure. Increased temperatures accelerate catalyst deactivation while high hydrogen partial pressure tends to mitigate catalyst deactivation rate. Most of the catalyst activity can be recovered by catalyst regeneration.

The selection of a suitable hydroprocessing catalyst is a critical step defining the hydro­processing product yield and quality as well as the operating cycle time of the process in petroleum industry [5]. However the hydrotreating catalyst selection for biomass applica­tions is particularly crucial and challenging for two reasons: a) catalyst activity varies sig­nificantly, as commercial catalysts are designed for different feedstocks, i. e. feedstocks with high sulfur concentration, heavy feedstocks (containing large molecules), feedstocks with high oxygen concentration etc, and b) there are currently no commercial hydropro­cessing catalysts available for lipid feedstocks and other intermediate products of biomass conversion processes (e. g. pyrolysis biooil), and thus commercial hydrotreating catalysts need to be explored and evaluated as different catalyst have different yields (Figure 5) and different degradation rate [8]. Nevertheless, significant efforts have been directed to-

wards developing special hydrotreating catalysts for converting/upgrading liquid biomass to biofuels [9—12].

The CDTECH process utilizes C4s and alcohol as feed to a fixed-bed downflow adiabatic re­actor. The equilibrium-converted reactor effluent is introduced to the reactive distillation column where the reaction continues. Concurrently, ether is separated from unreacted C4s as the bottom product.

In case of MTBE, the reactive distillation column overhead is washed in an extractor with a countercurrent H2O stream to extract MeOH. The H2O extract stream is sent to a MeOH re­covery column to recover both MeOH and H2O for recycle.

This scheme can provide overall IB conversions of up to 99.99% for MTBE process. Conver­sion is slightly less for ETBE than MTBE. For TAME and TAEE, isoamylene conversions of 95%+ are achievable.

Due to the various conditions that microorganisms grow and the constant flux of nu­trients that can persist in nature, there are numerous types of lipids found that can change in concentration as the local environments evolve through typical ebbs and flows of materials. In response to these changes, microorganisms will change their cellular struc­tures (i. e., lipid accumulation) by storing energy in various forms in order to utilize exist­ing nutrients and energy to prepare for leaner conditions that may occur. In practical terms, this concept can be leveraged in order to produce high concentrations of intracellu­lar lipids in marine and aquatic biomass in order to maximize the amount of lipids that can be harvested. Several studies have been conducted to determine what conditions af­fect the lipid composition and concentration of microorganisms. The more common tech­niques applied to increase the production of lipids from algae have through genetic manipulation [16], where genetic markers are manipulated that allow for increased lipid production to occur in the cell under normal conditions, by alteration of the cultivation conditions[17, 18], or by addition and manipulation of nutrients and chemicals added to the media [19]. By utilizing methods such as these, algal lipids can be increased by a sub­stantial amount without increasing the footprint of required reactor space, nor greatly in­creasing the amount of time between harvests.

Like other organisms, acetogens have a limited range of pH for optimal growth so the pH of the fermentation medium needs to be closely controlled. The extracellular pH directly influ­ences the intracellular pH, membrane potential, proton motive force, and consequently sub­strate utilization and product profile [208, 209]. In most studies, lowering pH medium divert carbon and electron flow from cell and acid formation towards alcohol production [113, 209—211]. By applying this knowledge, Gaddy and Clausen performed a two-stage CSTR syngas fermentation systems using C. ljungdahlii where they set the first reactor at pH 5 to promote cell growth, and pH 4 — 4.5 in the second reactor to induce ethanol production [212]. One recent study with C. Ijungdahlii showed conflicting results in which cell density and ethanol production were both higher at pH 6.8 when compared to pH 5.5 [213].

1.5. Temperature

The optimum temperature for mesophilic acetogens are between 30-40°C, while thermophil­ic acetogens grow best between 55 and 58°C. The fermentation temperature not only affects substrate utilization, growth rate and membrane lipid composition of the acetogens, but also gas substrate availability because gas solubility increases with decreasing temperature [24, 211]. "C. ragsdalef’was reported to produce more ethanol at 32°C than at the optimum growth temperature of 37°C [211].

Solvent extraction techniques have the potential for tremendous energy savings in the recovery of fermentation products such as ABE solvents. Such savings will have a direct impact on the economics for the entire fermentation. In order to find the optimal conditions of extractive butanol recovery, however, numerous conditions and factors have to be taken into consider­ation. A special case of the extractive recovery is the so-called in-situ extractive fermentation (see in chapter 6.5), where extraction is performed during, and together with fermentation. In this case, not only the separation and recovering characteristics play key role in the process, but also the toxicity of the non-miscible solvents basically determines the applicability of the method. In presence of an extractant solvent, however, due to distribution equilibriums, the concentration of each component of the fermentation broth (acids used in decreasing the pH to initiate the solventogenic stage, substrates or intermediates such as glucose, acetate, butyrate) changes. Being aware of these concentration relationships is essential to be able to control the process. Mass and energy balances of side-stream and countercurrent extraction were compared with the appropriate parameters of a classic distillation procedure in recovery of ABE solvents from fermentation broth [136]

A general mathematical model for performance evaluation of acetone-butanol continuous flash extractive fermentation system was formulated in terms of productivity, energy require­ment (energy utilization efficiency) and product purity. Simulation results based on experi­mental data showed that the most pronounced performance improvement could be achieved by using a highly concentrated substrate as feed and the increase in solvent dilution rate could only improve the total productivity at the expense of energy utilization efficiency. A two-vessel partial flash system, with the first vessel of two to three plates and the second vessel as a complete flash vessel, is required to ensure high product purity [137]. Extraction with solvents having distribution coefficients above one appears to have a more favourable energy balance than in case of distillation [136].

Distribution coefficients of ABE solvents between water and the selected extractant and biocompatibility of the extractant are crucial parameters. A solvent screening criterion was developed based on the maximum product concentartion attainable for the assessment of batch and semicontinuous multicomponent extractive fermentations [138]. Dadgar and Foutsch evaluated 47 solvents for the ability to recover Clostridium fermentatiuon products. Equili­brium distribution coefficients and separation factors from water for ethanol, butanol, and acetone were determined [139]. Griffith et al. [140] measured the organic/aqueous distribution coefficients of numerous potential BuOH extractants and simultaneously tested several in bacterial culture. The most effective appeared to be polyoxyalkylene ethers which had distribution coefficients in the range of 1.5-3 and showed little or no toxicity toward the fermentation. The esters and alcohols tested generally had better distribution coefficients but higher biotoxicity. Barton and Daugulis performed biocompatibility tests on 63 organic solvents, including alkanes, alcohols, aldehydes, acids, and esters. Thirty-one of these solvents were further tested to determine their partition coefficients for butanol in fermentation medium of C. acetobutylicum. The biocompatible solvent with the highest partition coefficient for BuOH (4.8) was poly(propylene glycol) 1200 which was selected for fermentation experi­ments F141G [141]. Thirty-six chemicals were tested for the distribution coefficients for BuOH, the selectivity of alcohol/water separation and the toxicity towards Clostridia. Convenient extractants were found in the group of esters with high molar mass. Liquid-liquid extraction was carried out in a stirred fermentor and a spray column. Formation of emulsions and fouling of the solvent in fermentation broth causes problems with the operation of this type of equipment [142].

Based on the solvent screening criterion and practical experience, one of the best solvents proved to be oleyl alcohol [143]. Oleyl alcohol was used in 40% that of the culture medium to extract BuOH and acetone from the fermentation broth produced from glucose by C. Aceto­butylicum and fermentation of the raffinate was continued after the extraction [144]. With a known biocompatibility of extractants such as oleyl alcohol, 1-decanol, 1-octanol, 1-heptanol and ethyl acetate, considering economic viewpoints as well, a mixed extractant of oleyl-alkohol and decanol was chosen for extraction at phase rate of 1:5 [145].

Both butyric acid and butanol could readily be extracted from microbial fermentation broth with vinyl bromide. The vinyl bromide fraction was separated from the aqueous broth and evaporated to give substantially pure butyric acid and (or) BuOH. Three passes of broth through separation columns of vinyl bromide at 4° enabled to isolate ~65% of total butyric acid and ~60% of BuOH in the broth [146]. The methyl, ethyl, propyl and butyl esters of vegetable oils are effective extractants for butanol from aqueous solutions. The effect of four salts, three alcohols and a ketone could be expected to affect the extraction of BuOH from industrial fermentation systems were evaluated. Variations in NaCl, Na2SO4, Na2SO3 and KH2PO4 from 0 to 0.15 M on the extraction of 0.1-4.1% BuOH from aqueous solutions at 25, 40, and 55° gave small changes in distribution coefficients. Mild increases occurred with increasing temperature and increasing NaCl, Na2SO4, and KH2PO4. Mild decreases in BuOH extraction occurred with increasing Na2SO3. Variations in acetone, EtOH, and 2-PrOH concentration ranging between 0 and 4% at 25, 40, and 55° gave small changes in distribution coefficients at BuOH concen­trations of 0.1-4.1%. A slight increase in BuOH extraction was observed with increasing 1- pentanol under similar conditions [147]. Extraction of ABE solvents with long-chain fatty acid esters/using the extracts without separation as diesel fuel is discussed in chapter 8.

Ionic liquids are novel green solvents that have the potential to be employed as extraction agents to remove butanol from aqueous fermentation media. An extraction procedure used 1- butyl-3-methylimidazolium-bis(trifluoromethylsulfonyl)imide or 1-butyl-3-methylimidazoli — um hexafluoro-phosphate ionic liquids was developed by Eom et al. [148]. Knowledge of phase behaviour of ionic-liquid-butanol-water systems is essential in selection the appropriate solvent [149,150]. The 1-hexyl-3-methylimidazolium bis(trifluoromethylsulfonyl)imide exhibits Type 2 liquid-liquid equilibrium behavior toward butanol-water system, thus this ionic liquid can easily separate 1-butanol from water [150].

Inaki Gandarias and Pedro Luis Arias

Additional information is available at the end of the chapter http://dx. doi. org/10.5772/52581

1. Introduction

In a future sustainable scenario a progressive transition by the chemical and energy indus­tries towards renewable feedstock will become compulsory. Energy demand is expected to grow by more than 50% by 2035 [1], with most of this increase in demand emerging from developing nations. Clearly, increasing demand from finite petroleum resources cannot be a satisfactory policy for the long term. The transition to a more renewable production system is now underway; however, this transition needs more research and investment in new tech­nologies to be feasible.

Biomass appears as the only renewable source for liquid fuels and most commodity chemi­cals [2]. This is the reason why, in the near future, bio-refineries in which biomass is catalyti — cally converted to pharmaceuticals, agricultural chemicals, plastics and transportation fuels will take the place of petrochemical plants [3]. Indeed, biomass represents 77.4% of global renewable energy supply [4]. Current technologies to produce liquid fuels from biomass are typically multistep and energy-intensive processes, including the production of ethanol by fermentation of biomass derived glucose [5],bio-oils by fast pyrolysis or high pressure lique­faction of biomass [6,7], polyols and alkanes from hydrogenolysis of biomass derived sorbi­tol [8],and biodiesel from vegetable oils [9].Biomass can also be gasified to produce CO and H2(synthesis gas), which can be further processed to produce methanol or liquid alkanes through Fischer-Tropsch synthesis [10].

The so-called "First Generation" biofuels, such as sugarcane ethanol in Brazil, corn ethanol in US, oilseed rape biodiesel in Germany, and palm oil biodiesel in Malaysia, already present mature commercial markets and well developed technologies. Nonetheless, there is a world­wide increasing awareness against the use of edible oils and seeds to generate transporta­tion fuels, and critical voices have aroused questioning the actual sustainability of these

"First Generation" biofuels. In fact, nowadays 95 % of biodiesel is made from edible oil [9]. This means that possible food resources are being used as automotive fuels when some part of the World’s population is suffering from hunger. Therefore, large-scale production of bio­diesel from edible oils may bring about a global imbalance in the food supply market. An­other significant concern of using "First Generation" technologies is the deforestation and the destruction of ecosystems. Indeed, the expansion of oil-crop plantations for biofuel pro­duction on a large scale has caused deforestation in countries such as Malaysia, Indonesia and Brazil because more and more forest has been cleared for plantation purposes. In addi­tion to this, in developing countries energy crops are powerful competitors for scarce water resources [11].

Being the non-edible portion of the plant and the most abundant source of biomass, ligno — cellulosic biomass materials are attracting growing attention as sustainable and renewable energy sources. The so-called "Second Generation" technologies for the production of fuels and chemicals can use a wide range of lignocellulosic biomass residues such as agricultural, industrial, and forest wastes, and also energy crops (willow, switchgrass) that do not com­pete with food crops for available land. The average composition of lignocellulosic material is as follows: 50% cellulose, 25% hemicellulose, and 20% lignin [12]. Cellulose is a linear pol­ysaccharide with p-1,4 linkages of D-glucopyranose monomers (Figure 1). Hemicellulose is a more complex polymer containing five different sugar monomers: five carbon sugars (xy­lose and arabinose) and six carbon sugars (galactose, glucose, and mannose). Lignin is a highly branched aromatic polymer, that consists of an irregular array of variously bonded "hydroxy-" and "methoxy-" substitutedphenylpropane units. Lignin is mainly found in woody biomass. Lignocellulosic materials can be converted into liquid fuels by three pri­mary routes, including (i) syngas production by gasification, (ii) bio-oil production by pyrol­ysis or liquefaction, and (iii) acid hydrolysis reactions [13].

In the pyrolysis process, biomass feedstock is heated in the absence of oxygen, forming a gaseous product, which after cooling condenses. Depending on the operating conditions that are used, pyrolysis processes are known as slow or fast pyrolysis. Fast pyrolysis proc­esses are characterized by high rates of particle heating (heating rate > 1000°C/min) to tem­peratures around 500°C, and rapid cooling of the produced vapors to condense them (vapor residence time 0.5-5s). In order to obtain that fast heating rates, it is essential to use reactors that provide high external heat transfer (such as fluidized bed reactors) and to guarantee an efficient heat transfer through the biomass particle, using biomass particle size of less than 5 mm [7]. Fast pyrolysis produce 60-75 wt% of liquid bio-oil, 15-25 wt% of solid char, and 10-20 wt% of non condensable gases, depending on the feedstock. In slow pyrolysis biomass is heated to around 500°C at much lower heating rates than those used in fast pyrolysis. The vapor residence times are much longer; they vary from 5 min to 30 min. As a consequence of the lower heating rate and of the longer vapor residence time, lower yields to pyrolysis oils and higher yields to char and gas products are obtained (Figure 2). As a result of all this, for bio-oil production from biomass, fast pyrolysis processes are preferred.

Bio-oils are dark-red brown color liquids. They are also known as pyrolysis oils, bio-crude oil, wood oil or liquid wood. Bio-oils usually have higher density, viscosity and oxygen con­tent compared to fuel-oil. While the sulfur and nitrogen content is usually smaller (Table 1). The high oxygen content of bio-oils generates some negative characteristics like low heating value (HV), immiscibility with conventional fuels and high viscosity. A serious problem of bio-oils is their instability during storage, as their viscosity, HV and density are affected. This is because some of the organic compounds present in bio-oils are highly reactive. For instance, ketones, aldehydes and organic acids react to form ethers, acetals and hemiacetals respectively [15]. Therefore, bio-oils need to be upgraded to reduce their oxygen content in order to increase their stability, to be miscible with conventional oil, and to increase their H/C ratio. This upgrading can be carried out through three different routes: (i) catalytic hy­drotreating, usually known as hydrodeoxygenation (HDO), which consists mainly on decar­boxylation, hydrocracking, hydrogenolysis and hydrogenation reactions, (ii) zeolite upgrading or (iii) through esterification reactions. Zeolite upgrading is carried out without external hydrogen sources, and therefore the resulting oil has lower HV and H/C than con­ventional fuels. Esterification can significantly increase the chemical and physical properties of bio-oil, however it requires using high amounts of alcohols, which are highly demanded. Catalytic hydrotreating appears to have the greatest potential to obtain high grade oils which are compatible with the already available infrastructure for fossil fuels.

Not only fuels, but also commodity chemicals are nowadays derived from petroleum-based resources. Commodity chemicals are involved in the production of a wide variety of prod­ucts and thus are an essential and integral part of the modern societies. Hence, in the search for a sustainable scenario, it is crucial to also look towards alternative biorenewable sources for these chemicals. In the case of platform chemicals coming from biomass, such as glucose, levulinic acid, 5-(hydroxyl-methyl furfural), sorbitol, or glycerol, they usually have higher O/C ratio than most commodity chemicals. Therefore, the conversion of these platform chemicals into value-added chemicals usually requires O removal reactions.

This book chapter summarizes the main aspects involved in the catalytic hydrotreating processes for the oxygen removal from bio-oils and from biomass based platform chemicals.

In an equilibrium reaction, simultaneous separation of obtained products will shift the re­action forward, thus further increasing the yield. In case of ETBE synthesis, simultaneous removal of the product (ETBE) and byproduct (H2O) could increase the yield of the target compound. Preliminary studies were carried out using the apparatus shown in Figure 12. In this set-up, the catalysts were placed at the middle of the column. Initially, an equimo­lar mixture of TBA and EtOH was placed in a round bottom flask in B, then microwave irradiated. The products (distillates) were condensed and collected outside the cavity. Af­ter 25 min, the collected products in the distillates (D) and bottoms (B) were analyzed of its composition.

Results in Figure 13, show that the bottom consisted of mostly H2O and unreacted EtOH, while the distillates consisted of ETBE, unreacted TBA, unreacted EtOH and H2O indicating possibility of simultaneous separation of the products. The future will look at the control and optimization of its operation to obtain better yield of the target compound.

Figure 12. Microwave apparatus for preliminary studies on reactive separation for ETBE synthesis

1.3. Biodiesel

1.3.1. Biodiesel extraction

Biodiesel is a clean-burning diesel fuel produced from vegetable oils, animal fats, or grease. Its chemical structure is that of fatty acid alkyl esters (FAAE). Biodiesel as a fuel gives much lower toxic air emissions than fossil diesel. In addition, it gives cleaner burning and has less sulfur content, and thus reducing emissions. Because of its origin from renewable resources, it is more likely that it competes with petroleum products in the future. To use biodiesel as a fuel, it should be mixedwith petroleum diesel fuel to create a biodiesel-blended fuel. Biodie­sel refers to the pure fuel before blending. Commercially, biodiesel is produced by transes­terification of triglycerides which are the main ingredients of biological origin oils in the presence of an alcohol (e. g. methanol, ethanol) and a catalyst (e. g. alkali, acid, enzyme) with glycerine as a major by-product [Ma and Hanna, 1999 ; Dube et al., 2007 ]. After the reaction, the glycerine is separated by settling or centrifuging and the layer obtained is purified prior to using it for its traditional applications (pharmaceutical, cosmetics and food industries) or for the recently developed applications (animal feed, carbon feedstock in fermentations, pol­ymers, surfactants, intermediates and lubricants) [Vicente et al., 2007].

However, one of the most serious obstacles to use biodiesel as an alternative fuel is the com­plicated and costly purification processes involved in its production. Therefore, biodiesel must be purified before being used as a fuel in order to fulfil the EN 14214 and ASTM D6751 standard specifications listed in Table 2; otherwise the methyl esters formed cannot be clas­sified as biodiesel. Removing glycerine from biodiesel is important since the glycerine con­tent is one of the most significant precursors for the biodiesel quality. Biodiesel content of glycerine can be in the form of free glycerine or bound glycerine in the form of glycerides. In this work we refer to the total glycerine, which is the sum of free glycerine and bound glyc­erine. Severe consequences may result due to the high content of free and total glycerine, such as buildup in fuel tanks, clogged fuel systems, injector fouling and valve deposits (Hayyan et al., 2010).

The ABE producing strains can hydrolyze starch to glucose or other hexose by amylases. Glucose was firstly converted to pyruvate through the Embden-Meyerhoff pathway (EMP, or glycolysis). Pyruvate was then cleaved to acetyl-CoA by pyruvate ferredoxin oxidoreduc — tase. Acetyl-CoA is the common precursor of all the fermentation intermediate and end products. The enzyme activity and the coding genes have been widely assayed and descri­bed in butanol-producing strains (Durre et al., 1995; Gheshlaghi et al., 2009).

The ABE fermentation process can be divided into two successive and distinct phase as acidogenesis phase and solvetogenesis phase. The acidogenesis phase is accompanied with cell exponential growth and pH drop, accumulation of acetate and butyrate. Solventogenesis phase begins with endospore forming and the cells entering stationary state. The products of acidogenesis phase include acetate and butyrate. Acetate forms from Acetyl-CoA, which is catalyzed by two enzymes, phosphotransacetylase (PTA, or phosphate acetyltransferase, endoced by pta gene) and acetate kinase (AK, encoded by ak gene). The butyrate synthesis is a little complicated with more steps. At first, two molecular of acetyl-CoA is catalyzed by thiolase (thl, or acetyl-CoA acetyltransferase, encoded by thl gene) and transforms into one molecular C4 unit acetoacetyl-CoA, which is another important node and precursor of buty­rate, acetone, and butanol synthesis. The acetoacetyl-CoA is subjected to three enzymes in turn and another C4 unit butyryl-CoA is the intermediate product. The three enzymes are hydroxybutyryl-CoA dehydrogenase (encoded by hbd gene) (Youngleson et al., 1995), croto — nase (CRT, or hydroxybutyryl-CoA dehydrolase, encoded by crt gene), and butyryl-CoA de­hydrogenase (BCD, encoded by bcd gene). Accordingly, three encoded genes coexist in the BCS operon with additional two genes coding for the a and p subunit of electron transfer protein (Bennett and Rudolph, 1995). Butyryl-CoA was then catalyzed by phosphotransbu- tylase (PTB, or phosphate butyltransferase, encoded by ptb gene) and butyrate kinase (BK, encoded by bk gene) to form butyrate during acidogenesis phase.

As the organic acid accumulation, pH drop to the lowest point during the fermentation. This leads to the switch of acidogenesis phase to solventogenesis phase. Acetate and butyrate are reassimilated and participate in the solvent formation. Under the catalyzing of CoA transfer­ase (CoAT, two unit encoded by ctfa and ctffi), acetate and butyrate was transformed into acetyl-CoA and butyryl-CoA respectively again. The alcohols formation share the same key enzymes, NAD(P)H dependent aldehyde/alcohol dehydrogenases (encoded by adhl and adh2 gene) (Chen, 1995). In addition, Butanol owns its unique butanol dehydrogenase (en­coded by bdh gene) (Welch et al., 1989). The formation of acetone from acetoacetyl-CoA is a two-step reaction. Acetoacetyl-CoA is catalyzed to acetoacetate by CoA transferase. Acetone is produced after a molecular CO2 released from acetoacetate by decarboxylase (AADC, en­coded by aadc gene) (Janati-Idrissi et al., 1988; Cary et al., 1993). Both acid reassimilation and acetone formation utilize CoA transferase, however, the butyrate uptake was not concomi­tant with the production of acetone (Desai et al., 1999). The metabolic pathway accompanied by electron transfer and reduction force forming. The main ABE fermentation pathway was illustrated in Fig.3.

Solventogenic genes aad, ctfA, ctfB and adc constitute the sol operon (Durre et al., 1995). In some conditions, butanol producing strains lose the ability to produce solvents after repeat­ed subculturing, called as degenerated (DGN) strain. In C. acetobutylicum ATCC 824, the plasmid pSOL1 carrying the sol operon was found missing during degenerating process (Cornillot et al., 1997). For C. saccharoperbutyl acetonicum strain N1-4, the sol genes main­tained in degenerated DGN3-4 strain, while the sol operon was hardly induced during sol- ventogenesis. Extract from the culture supernatants of wild-type N1-4 is enough to induce the transcription of the sol operon in DGN3-4 (Kosaka et al., 2007). It suggested that the de­generation maybe caused by the incompetence of the induction mechanism of the sol oper — on. The transcription of sol operon may be under the control of the quorum-sensing mechanism in C. saccharoperbutyl acetonicum.

Though the metabolic pathway is clear, the underlying regulation mechanism is poorly un­derstood, such as the phase switch of fermentation, the relationship between solventogene — sis and sporulation. Answering these questions is critical to improve the efficiency of butanol producing fundamentally. Proteomics and transcriptomics can provide more un­known details, which will be helpful for solving these problems (Sivagnanam et al., 2011; Sivagnanam et al., 2012).