Cyanobacteria are predicted to be the first microorganisms to develop the capability of oxygenic photosynthesis, some 2.7 billion years ago [112]. Similar to algae, cyanobacteria have a great range of diverse morphologies, cellular functions, and genetics, presumably due to their long evolutionary history and their diverse habitats. As discussed previously, the ASP initially deemed cyanobacteria unfit for fuel production due to their lack of natural TAG accumulation. Since they are amenable to genetic manipulation, however, cyanobacteria can be engineered to produce a range of biofuel products (Table 1). As prokaryotes, cyanobacteria are subject to the traditional methods employed for engineering other well-developed bacterial hosts like E. coli. Some strains of cyanobacteria are even naturally transformable, uptaking exogenous DNA from their environment without the use of cell permeablization techniques [113]. As progenitors of the algal chloroplast, cyanobacteria also integrate DNA into their chromosomes using homologous recombination. Moreover, cyanobacteria do not possess the cellular components for gene silencing. The genetic tools for engineering some model strains of cyanobacteria are well developed and have been used to genetically modify cyanobacteria for several decades [113]. Another advantage of using cyanobacteria as the microbial host for hydrocarbon-based fuel production is that they have been shown to excrete potential fuel precursors such as FFAs [73]. Fuel excretion enables a continuous production process, eliminating the cost associated with harvesting the algal biomass and the time and nutrients needed to repeatedly grow new batches of algae for fuel production. The advantages of straightforward genetic manipulation and fuel excretion make cyanobacteria contenders for large-scale biofuel production despite the disadvantage of low natural lipid yields.

After the initial demonstration of engineering cyanobacteria for ethanol production [97], the production of hydrocarbon-based fuels in engineered cyanobacteria has expanded to include isoprene, FFAs, FAEEs, fatty alcohols, and alkanes/alkenes (Table 1). Isoprene biosynthesis was established in the model cyanobacterium, Synechocystis sp. PCC 6803, through expression of the isoprene synthase (ispS) from kudzu [78]. Codon optimization of ispS and the use of a strong promoter (psbA2) increased isoprene production. Engineering strategies targeting the upstream MEP pathway for isoprenoid biosynthesis, as described in Section 2.2 of this chapter, will likely further improve isoprene productivity. The remaining four hydrocarbon-based fuels are all derived from the fatty acid biosynthesis pathway. Common strategies for im­proving FFA production (see Section 2.1) have proven successful in cyanobacteria [74—76]. Eliminating non-essential, competing pathways such as polyhydroxybutyrate (PHB), cyano — phycin, and acetate biosynthesis also improved FFA production [74]. Liu and colleagues engineered a more permeable peptidoglycan layer to improve FFA excretion in Synechocystis sp., yet this weakened cell membrane resulted in slower growth rates and may also make the engineered cyanobacterium more susceptible to external predators and toxins that may be present in large-scale cultivations. Initial engineering attempts for fatty alcohol and alkane/ alkene production entail expression of a heterologous FAR and overexpression of AAR and ADC, respectively [23, 26]. Alkane/alkene synthesis was also observed with ACC overexpres­sion and native AAR and ADC activities in cyanobacteria [23]. Despite being derived from fatty acids, the synthesis of fatty alcohols and alkanes/alkenes is up to 1000-fold lower than that observed with FFA production (Table 1), suggesting that the conversion of acyl-ACP to the final fuel product is rate limiting. These inaugural proof-of-concept reports illustrate the potential of cyanobacteria as hosts for autotrophic biofuel production, but additional metabolic engineering will be required to achieve the fuel titers necessary for large-scale synthesis.

As stated above, the use of acid or base as a co-catalyst gives 1,2-PDO as a main product. To obtain more valuable 1,3-PDO, the most effective approach has shown to be the use of noble metal (Ir, Rh or Pt) combined with oxophilic metals. Shinmi et al. [52] modified Rh/SiO2 cat­alyst with Re, W and Mo. Re addition showed the largest enhancing effect on catalytic activ­ity and also increased the selectivity to 1,3-PDO. The Rh-ReOx/SiO2 (Re/Rh = 0.5) exhibited 22 times higher glycerol conversion (79%) and 37 times higher 1,3-PD yield (11%) than

Rh/SiO2. In a more recent work, an Ir-ReOx/SiO2 (Re/Ir = 1) catalyst prepared by a similar method to that for Rh-ReOx/SiO2 catalyzed the hydrogenolysis of glycerol to 1,3-PDO in a more effectively way (1,3-PDO/1,2-PDO ratio = 11) [72]. Based on characterization results, the authors suggested that oxidized low-valence Re clusters are attached to the Ir or Rh met­al particles. Glycerol is adsorbed on the surface of MO, species (M = Mo, Re and W) at the OH group to form alkoxide. Hydrogen is activated on the noble-metal (Rh or Ir) surface. The alkoxide located on the interface between MO, and the noble-metal surface is attacked by the activated hydrogen species, and the C-O bond neighboring to the C-O-M group is dis­sociated. The hydrolysis of the resulting alkoxide releases the product (see Figure 11). One of the weak point of these catalytic systems is that they are also active in the further hydro — genolysis of both 1,2 and 1,3-PDO to 1-PO.

In summary, Cu based catalysts are active and selective for the production of 1,2-PDO from glycerol. However, if the aim is to produce the more valuable 1,3-PDO, different approaches are required. The used of noble metals combined with low-valence metal oxide seems to be a promising alternative. Nonetheless, there is still room for improvement; both in catalyst design and in process engineering, as PDOs further hydrogenolysis significantly affect the final yields to target products.

Electrical energy is added to the "TONADO-LE" plasma-liquid system in the form of plasma power. Plasma acts as a catalyst and thus this power should be controlled. In addition to electric energy for plasma we incorporate hydrocarbon (ethanol or bioglycerol) as an input to the system. These hydrocarbons are raw material for syn-gas generation but they are also a fuel which has some energy associated with it. So, we input some energy to the system (hydrocar­bon + electricity) and we get syn-gas, which is potentially a source of energy as well.

Carbon dioxide adding leads to a significant increase the percentage of H2 + CO (syn-gas) and CH4 components in the exhaust. This may indicate that the CO2 addition under the ethanol reforming increases the conversion efficiency, because CO2 plays a role of the retarder in the system by reducing the intensity of the conversion components combustion.

The transmission spectra of infrared radiation indicate that the exhaust gas obtained by ethanol solution conversion, contains such components as CO, CO2, CH4, C2H2. It was found that CO2 adding reduces the CH4 and C2H2 amount, but does not affect the amount of producted CO.

The possibility of hydrocarbons reforming, which have considerable viscosity (bioglycerol) in the "TORNADO-LE" is shown. This gives a possibility to avoid environmental problems due to the bioglycerol accumulation during biodiesel production.

The a coefficient [see (2)] in bioglycerol reforming is higher than ethanol reforming at the same ratios of CO2/Air in the input gas. This may be connected with the lower power consumption on the plasma generation in case of bioglycerol reforming. Bioglycerol contains alkaline dash, which increases the bioglycerol conductivity. Bioglycerol reforming products contain mainly CO and hydrocarbons CH4, C2H2 which also gives some contribution to energy yield.

The genomes of several solventogenic Clostridia, including gas fermenting species, have been sequenced since 2001 [54, 62, 109, 119, 123, 130], and an array of transcriptomic [100, 116, 121, 131, 132], proteomic [132] and systems analysis [133, 134] are being made increas­ingly available. However, the generation of stable recombinant Clostridia has been severely hindered by the difficulties encountered introducing foreign DNA into cells and a lack of established genetic tools for this genera of bacteria. In comparison to starch-utilizing Clostri­dia, very little information is available for metabolic engineering of acetogens. Although this section describes recent advances in the development of genetic tools for mostly sugar-uti­lizing Clostridia, these techniques are highly relevant and applicable to the closely related acetogenic Clostridia for biofuels or chemical production via gas fermentation.

The ideal microbial catalyst for industrial scale gas fermentation might exhibit the following traits: high product yield and selectivity, low product inhibition, no strain degeneration, as — porogenous, prolonged cell viability, strong aero-tolerance, high biomass density and effi­cient utilization of gas substrates. These can be achieved by directed evolution, random mutagenesis and/or targeted genetic engineering. Traditionally, chemical mutagenesis [135—137] and adaption strategies [138, 139] have been deployed to select for these traits. However, these strategies are limited and often come with the expense of unwanted events.

First attempts of targeted genetic modification of Clostridia were made in the early 1990s by the laboratory of Prof. Terry Papoutsakis [140—142]. While these pioneering efforts relied on use of plasmids for (over)expression of genes in C. acetobutylicum, more sophisticated tools were later developed for a range of solventogenic and pathogenic Clostridia.

Antisense RNA (asRNA) has been employed to down-regulate genes. Here, single stranded RNA binds to a complementary target mRNA and prevents translation by hindering ribo­some-binding site interactions [143]. For instance, this method has been used to knockdown ctfB resulting in production of 30 g/l solvents with significantly suppressed acetone yield in C. acetobutylicum ATCC 824 [144, 145].

Several homologous recombination methods have been developed for integration or knock-out of genes in a range of sugar-utilizing Clostridia. In early stage, knockout mu­tants were almost exclusively generated from single crossover events that could revert back to wild-type [146—152], with stable double crossovers only observed in rare cases [153, 154]. For C. acetobutylicum [155] and cellulolytic C. thermocellum [156] counter selecta­ble markers have been developed to allow more efficient screening for the rare second re­combination event.

ClosTron utilizes the specificity of mobile group II intron Ll. ltrB from Lactoccocus lactis to propagate into a specified site in the genome via a RNA-mediated, retro-homing mecha­nism which can be used to disrupt genes [157]. This technique has initially been devel­oped by InGex and Sigma-Aldrich under the name ‘TargeTron™’ and successfully adapted to a range of solventogenic and pathogenic Clostridia including C. acetobutyli­cum, C. difficile, C. sporogenes, C. perfringens, and C. botulinum [158—160] by the laboratory of Prof. Nigel Minton.

The same laboratory recently also developed another method for integration of DNA into the genome. Termed Allele-Coupled Exchange (ACE), this approach does not employ a counter selective marker to select for the rare second recombination event. Rather, it utilizes the activation or inactivation of gene(s) that result in a selectable phenotype, and asymmetri­cal homology arms to direct the order of recombination events [161]. Remarkably, the whole genome of phage lambda (48.5kb minus a 6kb region) was successfully inserted into the ge­nome of C. acetobutylicum ATCC 824 in three successive steps using this genetic tool. This technique was also demonstrated in C. difficile and C. sporogenes [161].

For reverse engineering, mainly transposon mutagenesis has been utilized. Earlier efforts of transposon mutagenesis were demonstrated in C. acetobutylicum P262 (now: C. saccharobuty — licum [162]), C. acetobutylicum DSM792, C. acetobutylicum DSM1732, and C. beijerinckii NCIM 8052, but issues with multiple transposon insertions per mutant, and non-random distribu­tion of insertion were reported [163, 164]. Recent developments have seen the successful generation of mono-copy random insertion of transposon Tn1545 into cellulolytic C. cellulo — lyticum [165] and mariner transposon Himarl into pathogenic C. difficile [166].

While there is still a lack of some other essential metabolic engineering tools such as efficient inducible promoters, the array of available tools that enabled significant improvements to the ABE process and cellulolytic Clostridia fermentations as summarized in Table 3.

In contrast, to date only a limited number of acetogenic Clostridia have been successfully modified. Pioneering work in this area has been undertaken in the laboratory of Prof. Peter Durre. C. Ijungdahlii, a species that does not naturally produce butanol, was modified with butanol biosynthetic genes (thlA, hbd, crt, bcd, adhE and bdhA) from C. acetobutylicum ATCC 824 resulting in production of up to 2 mM of butanol using synthesis gas as sole energy and carbon source [62]. By delivering a plasmid with acetone biosynthesis genes ctfA, ctfB, adc, and thlA in C. aceticum, production of up to 140 |oM acetone was demonstrated from various gas mixes (80% H/20% CO2 and 67% H2/33% CO2) [169, 170]. Recent patent filings by Lanza — Tech describe the production of butanol as main fermentation product and increased alcohol tolerance in genetically engineered acetogens. Up to 26 mM butanol were produced with ge­netically modified C. ljungdahlii and C. autoethanogenum using steel mill gas (composition 44% CO, 32% N2, 22% CO2, and 2% H2) as the only source of carbon and energy when the butanol biosynthetic genes thlA, hbd, crt, bcd, etfA, and etfB were heterologously expressed [167]. Overexpression of native groESL operon in C. autoethanogenum resulted in a strain that displayed higher alcohol tolerance relative to wild-type when challenged with ethanol [168].

Besides the classical Clostridial butanol pathway (which constitutes genes thlA, crt, hbd, bcd, etfA and etfB; see earlier section), a non-fermentative approach has been described and dem­onstrated in E. coli for branched chain higher alcohol production [182]. This alternative ap­proach requires a combination of highly active amino acid biosynthetic pathway and artificial diversion of 2-keto acid intermediates into alcohols by introduction of two addi­tional genes: broad substrate range 2-keto-acid decarboxylase (kdc) which converts 2-keto acids into aldehydes, followed by Adh to form alcohols [182]. Engineered strains of E. coli have been shown to produce alcohols such as isobutanol, n-butanol, 2-methyl-1-butanol, 3- methyl-1-butanol and 2-phenylethanol via this strategy [182]. For instance, the overexpres­sion of kivD (KDC from Lactococcus lactis), adh2, ilvA, and leuABCD operon, coupled with deletion of ilvD gene and supplementation of L-threonine, increased n-butanol yield to 9 mM while producing 10 mM of 1-propanol [182].An even more remarkable yield of 300 mM isobutanol was achieved through introduction of kivD, adh2, alsS (from B. subtilis), and ilvCD into E. coli [182]. Like butanol, isobutanol exhibits superior properties as a transporta­tion fuel when compared to ethanol [177]. By applying similar strategy into C. cellulolyticum, 8.9 mM isobutanol was produced from cellulose when kivD, yqhD, alsS, ilvC, and ilvD were overexpressed [177]. This result suggests that such non-fermentative pathway is suitable tar­get for metabolic engineering of acetogens for the biosynthesis of branched chain higher al­cohols. Via synthetic biology and metabolic engineering, production of additional potential liquid transportation fuels like farnesese or fatty acid based fuels has successfully been dem­onstrated in E. coli or yeast from sugar [183, 184]. Given the unsolved energetics in aceto — gens, it is unclear if production of such energy dense liquid fuels could be viable via gas fermentation.

Wyne [79] studied the inhibition of ABE fermentation of maize mash by C. acetobutylicum influenced by 30 representative inorganic and organic acids. Several acids caused complete inhibition when the initial reaction was between pH 3.90 and 3.65, the following being included: HCl, HNO3, H2SO4 H3PO4, succinic, maleic, malonic, levulinic, crotonic, glycolic, p — hydroxybutyric, formic, acetic, propionic, butyric and isobutyric. The toxic effects are probably associated with a critical CH+ in the cell interior, closely approximating the observed extracel­lular CH+ associated with an inhibitory effect. All three chloroacetic acids are much more toxic than acetic acid, but hydroxy derivatives of the lower fatty acids are not more toxic than the corresponding normal acids. Pyruvic, lactic and glyceric acids are tolerated at higher CH+ levels. In the lower fatty acids the inhibiting CH+ was appreciably lower with each successive higher homolog. On the basis of molar concentration the order of effectiveness of inhibition was as follows: nonylic > caprylic > heptylic > formic > caproic = isocaproic > valeric = isovaleric > isobutyric = butyric > propionic = acetic. Capillary activity has relatively little effect with formic, acetic, propionic and butric acids, but was very marked with higher homologs [79]. Inhibitory effect of these acids can easily be removed by neutralization [80]. When the ABE fermentation is over, the culture medium may be treated by blowing NH3 to neutralize most of organic acids and, after distilling out the solvents, the residue can be treated with non-N-containing nutrients, e. g. dried sweet potatoes, and the fermentation may be repeated in the same way in order to save the quantity of nutrient and to increase the yield [81].

The effect of agitation speed and pressure was studied by Doremus et al [82]. Batch fermen­tations were run at varying agitation rates and were either pressurized to 1 bar or nonpres — surized. Agitation and pressure both affect the level of dissolved H2 in the media which, in turn, influence solvent production. In nonpressurized fermentations volumetric productivity of BuOH increased as the agitation rate decreased. While agitation had no significant effect on BuOH productivity under pressurized conditions, overall BuOH productivity increased over that obtained in nonpressurized runs. Maximal butyric acid productivity, however, occurred earlier and increased as agitation increased. Peak H2 productivity occurred simultaneously with peak butyric acid productivity. The proportion of reducing equivalents used in forming the above products was determined using a redox balance based on the fermentation stoichi­ometry. An inverse relationship between the final concentrations of acetone and acetoin was found in all fermentations studied [82]. Using shear activation of C. acetobutylicum by pumping the cells through capillaries, the cell growth, glucose consumption and product formation rates are considerably increased. Shear-activated continuous cell culture can be used as an inoculum with a well-defined fermentation activity for batch cultures. Different runs of such batch cultivation yield well-reproducible results which could not be obtained from inocula of other cultures or even of heat-shocked spores. The cells can attain a growth rate higher than 1.6 h-1. The shear-activated continous culture growth is affected already at a butanol concentration lower than 1.6 g L-1[83], Afschar et al (1986) [80]. The effect of viscosity on the ABE fermentation was studied by Korneeva et al. [84]. Viscosity of the medium was a limiting factor in ABE production by C. acetobutylicum during fermentation with starch and grains such as wheat and rye flour. Various concentrations of agar-agar (0.1, 0.5 and 0.8%) were added to the medium which showed that elevation of viscosity reduces saccharification, increases the concentration of nonfermented sugars, and decreases the yield of solvents. Prior treatment of the substrate with a-amylase reduced the viscosity of the medium and improved fermentation and solvent yields [84].

Although the ABE fermentation is a strictly anaerobic process, [2] Nakhmanovich and Kochkina [85] could increase the BuOH yield by 3.4-9.1% by short periodical aeration of the medium. Redox potential was measured before and after bubbling and decreased sharply by aeration. In batch and continuous cultivations of C. acetobutylicum ATCC 824 on lactose, a strong relationship was observed between redox potential of broth and cellular metabolism [86]. The specific productivity of BuOH and of butyric acid was maximal at a redox potential of -250 mV. The specific production rate of butyric acid decreased rapidly at higher and lower redox potentials. For BuOH, however, it achieved a lower but stable value. This was true for both dynamic and steady states. Continuous fermentations involving lactose exhibited sustained oscillation at low dilution rates. Such oscillation appears to be related to BuOH toxicity to the growth of cells. At higher dilution rates, where BuOH concentrations were relatively low, no such oscillation was observed. Broth redox potential apparently is an excellent indicator of the resulting fermentation product partitioning [86]. Some selected examples are given in Table 2 and 3.

Vegetable oils are the main feedstock for the production of first generation biofuels, which can offer several CO2 benefits and limit the consumption of fossil fuels. Raw vegetable oils consist of fatty acid triglycerides, the consistency of which depends on their origin (i. e. plant type) as shown in Table 2. Their production, however, is competing for the cultivated areas that were originally dedicated for the production of food and feed crops. As a result the pro­duction and utilization of vegetable oils for biofuels production has instigated the "food vs. fuel" debate. For this reason traditional energy crops (soy, cotton, etc) with low oil yield per hectare are being substituted by new energy crops (eg. jatropha, palm, castor etc).

Catalytic hydrotreatment was explored for conversion of vegetable oils in the early 90’s. The investigation of the hydrogenolysis of various vegetable oils such as maracuja, buritimtucha and babassu oils over a Ni-Mo/y-Al2O3 catalyst as well as the effect of temperature and pressure on its effectiveness was firstly investigated [16][17]. The reaction products included a gas product rich in the excess hydrogen, carbon monoxide, carbon dioxide and light hy­drocarbons as well as a liquid organic product of paraffinic nature. In more detail these studies showed the conversion of triglycerides into carboxyl oxides and then to high quality hydrocarbons via decarboxylation and decarbonylation reactions. Rapeseed oil hydropro­cessing was also studied in lab-scale reactor for temperatures 310° and 360°C and hydrogen pressures of 7 and 15 MPa using three different Ni-Mo/alumina catalysts [18]. These prod­ucts contained mostly n-heptadecane and n-octadecane accompanied by low concentrations of other n-alkanes and i-alkanes [19].

3.2. Waste cooking oils conversion to paraffinic biofuels

Even though vegetable oils are the main feedstock for the production of first generation bio­fuels, soon their production has troubled the public opinion due to their abated sustainabili­ty and to their association with the food vs. fuel debate. As a result the technology has shifted towards the exploitation of both solid and liquid residual biomass. Waste Cooking Oils (WCOs) is a type of residual biomass resulting from frying with typical vegetable fry­ing oils (e. g. soybean-oil, corn-oil, olive-oil, sesame-oil etc). WCOs have particular problems regarding their disposal. In particular grease may result in coating of pipelines within the residential sewage system and is one of the most common causes of clogs and sewage spills. Furthermore, in the cases that sewage leaks into the environment, WCOs can cause human and environmental health problems because of the pathogens contained. It has been estimat­ed that by disposing 1 lit of WCO, over 1,000,000 of liters of water can be contaminated, which is estimated as the average demand of a single person for 14 years.

Catalytic hydroprocessing of WCO was studied as an alternative approach of producing 2nd generation biofuels [20—24]. Initially catalytic hydrocracking was investigated over commer­cial hydrocracking catalysts leading not only to biodiesel but also to lighter products such as biogasoline [20], employing a continuous-flow catalytic hydroprocessing pilot-plant with a fixed-bed reactor. During this study several parameters were considered including hydro­cracking temperature (350-390°C) and liquid hourly space velocity or LHSV (0.5-2.5 hr-1) un­der high pressure (140 bar), revealing that the conversion is favoured by high reaction temperature and low LHSV. Lower and medium temperatures, however, were more suita­ble for biodiesel production while higher temperatures offered better selectivity for biogaso­line production. Furthermore, heteroatom removal (S, N and particularly O) was increased while saturation of double bonds was decreased with increasing hydrocracking tempera­ture, indicating the necessity of a pre-treatment step.

However catalytic hydrotreatment was later examined in more detail as a more promising technology particularly for paraffinic biodiesel production (Figure 1). The same team has studied the effect of temperature (330-398°C) on the product yields and heteroatom removal [21]. The study was conducted in the same pilot plant utilizing a commercial NiMo/Al2O3 hydrotreating catalyst over lower pressure (80 bar). According to this study, the hydrotreat­ing temperature is the key operating parameter which defines the catalyst effectiveness and life. In fact lower temperatures (330°C) favour diesel production and selectivity. Sulfur and nitrogen removal were equally effective at all temperatures, while oxygen removal and satu­ration of double bonds were favoured by hydrotreating temperature. The same team also studied the effect of the other three operating parameters i. e. pressure, LHSV and H2/WCO ratio [22]. Moreover they also studied the hydrocarbon content of the products [23] qualita­tively via two-dimensional chromatography and quantitatively via Gas Chromatography with Flame Ionization Detector (GC-FID), which indicated the presence of C15-C18 paraf­fins. Interestingly this study showed that as hydrotreating temperature increases, the con­tent of normal paraffins decreases while of iso-paraffins increases, revealing that isomerization reactions are favoured by temperature.

Figure 7. Catalytic hydrotreatment of WCO to 2nd generation biodiesel

The total liquid product of WCO catalytic hydrotreatment was further investigated in terms of its percentage that contains paraffins within the diesel boiling point range (220-360°C) [24]. The properties of WCO, hydrotreated WCO (total liquid product) and the diesel frac­tion of the hydrotreated WCO are presented in Table 3. Based on this study the overall yield of the WCO catalytic hydrotreatment technology was estimated over 92%v/v. The properties of the new 2nd generation paraffinic diesel product indicated a high-quality diesel with high heating value (49MJ/kg) and high cetane index (77) which is double of the one of fossil die­sel. An additional advantage of the new biodiesel is its oxidation stability (exceeding 22hrs) and negligible acidity, rendering it as a safe biofuel, suitable for use in all engines. The prop­erties and potential of the new biodiesel were further studied [25], for evaluating different fractions of the total liquid product and their suitability as an alternative diesel fuel.

Similar to Philips Etherification Process, ethers are produced by the addition of alcohol to reactive olefins in the presence of an ion exchange resin at mild temperature and pressure.

The feed passes through two reactors in series — an isothermal tubular reactor and an adia­batic drum reactor. The second reactor effluent goes to the product fractionation tower where the ether product leaves the bottom stream and hydrocarbon is recovered overhead.

In the MTBE process, MeOH in the overhead stream is extracted with H2O in the MeOH re­moval tower. The extract from the bottom enters the MeOH-H2O fractionator, while the MeOH overhead is recycled to reactor feed.

Pyrolysis is a process in which organic matter is exposed to heat and pressure in the absence of oxygen. The primary components of this process are syngas molecules like those found in gasification, as well as bio-oils and charred solid residues [26]. Pyrolysis methods are de­fined by the rate of heating, which directly affects the residence time of the reaction [27]. In slow pyrolysis, for example, the material is exposed to reactor conditions for five minutes; in fast pyrolysis, residence time is reduced to one to two minutes and in flash pyrolysis to less than five seconds. The residence time of the pyrolysis reaction greatly influences the compo­sition of oils, gases and chars that are formed [28—30]. Several studies have been performed to identify the effect of operational variables— reactor conditions and variations in feed­stock material —on the quality of the pyrolysis oils, gases, and chars [27, 30]. The oils typi­cally produced during pyrolysis reactions are high in moisture content, and corrosive due to low pH. Pyrolysis of biomass is typically constrained by the high water content of the raw material, and current pyrolysis methods for biomass conversion have not reached the stage of commercial development. Ongoing research, however, aims at maximizing energy poten­tial from biomass and optimizing conversion methods to achieve commercialization at mar­ketable levels [31, 32].

The growing commercial interests in using gas fermentation as a platform for biofuels pro­duction is evident in the recent spike in patent fillings within the field [105]. A 2009 report compared mass and energy conversion efficiencies from a process engineering standpoint between enzymatic hydrolysis fermentation of lignocellulose, syngas fermentation and FTP [227]. The authors concluded that while syngas fermentation offers a range of advantages such as low pretreatment requirement and low energy requirement for bioconversion, the technology is severely limited by low ethanol productivity [227]. Another report document­ed the techno-economic analysis of gas fermentation and concluded that the selling price of ethanol using this technology would still be significantly higher than gasoline in 2009 [228]. In contrast, Griffin and Schultz recently compared the production of ethanol from CO-rich gas using thermo-chemical route and biological gas fermentation route [22]. The authors concluded that gas fermentation offers superior fuel yield per volume of biomass feed, car­bon conversion to fuel, energy efficiency and lower carbon emissions relative to the thermo­chemical approach to bioethanol production.

Ethanol and butanol are the most attractive fuel products from current gas fermentation but other by-products such as 2,3-butanediol, acetic acid and butyric acid are also valuable com­modities that have the potential to provide significant additional revenue streams, setting off costs for biofuel production. 2,3-butanediol is a high value commodity which can be used to synthesize chemical products such as 1,3-butanediane, methyl ethyl ketone, and gamma butyrolactone, with a combined potential market value of $43 billion [104]. Acetic acid is an important precursor for synthesis of polymers while butyric acid can be used as a flavouring agent in the food industry [229, 230]. With the development of advanced genetic

tools for expansion of product range, the industry might witness an increasing emphasis on the production of high-value commodities in addition to biofuels.

Several companies are actively engaged in the development of the gas fermentation technol­ogy and some are approaching commercialization. Bioengineering Resources Inc (BRI) founded by Prof. James Gaddy of University of Arkensas, Fayetteville, an early pioneer in the investigation of gas fermentation at scale, was the first company to explore the potential of gas fermentation for industrial bioethanol production. BRI was acquired by chemical company INEOS and rebranded as INEOS Bio (www. ineosbio. com). A pilot-scale facility in Arkansas has been operated since 2003 using several isolates of C. Ijungdahlii [231] and is building a US$130 million commercial facility in Florida with its joint venture partner New Planet Energy Florida [232]. The commercial facility is expected to start operation in the sec­ond quarter of 2012 and is aiming to generate 8 million gallon of cellulosic ethanol per an­num and 6 MW of power to the local communities [232]. INEOS Bio also announced design of a second plant, the Seal Sands Biorefinery in Teeside, UK [233].

Founded in 2006, Coskata Inc. (www. coskata. com) is a US-based company that has reported achieving ethanol yields of 100 gallons per dry ton of wood biomass in a semi-commercial facility in Pennsylvania [234]. The company licensed several microbial strains from the Uni­versity of Oklahoma [235], which has filed patents and journal publications for acetogens such as "C. ragsdalei" [211, 236, 237] and C. carboxidivorans [55, 112]. A patent documenting a new ethanologenic species, "C. coskatii" was also recently filed by Coskata [238]. Backed by a conditional US$250 million loan guarantee from the US Department of Agriculture (USDA), Coskata has announced that it is planning to build a commercial plant with the capacity to produce 55 million gallon fuel grade ethanol per annum in Alabama [234, 239]. While the initial strategy saw biomass as feedstock, the company recently announced its first commer­cial plant will be switched to 100% natural gas as feedstock [240]. A planned IPO with the aim to tap into private investors to finance the plant was put on hold [241]. In 2012, Coskata and INEOS Bio were involved in a trade secret dispute which culminated in a settlement that see INEOS Bio receiving US$2.5 million cash payment, shares and right to receive 2.5% of future ethanol royalties from Coskata [242].

LanzaTech is a NZ/US based company that has developed a gas fermentation technology to utilize industrial off-gases from steel making and other sources, as well as syngas pro­duced from biomass as feedstocks. The company has reported the development of a pro­prietary Clostridial biocatalyst that is able to convert the CO-rich waste gas with minimal gas conditioning into bioethanol and the platform chemical 2,3-butanediol. The use of in­dustrial off-gases as feedstock not only helps to reduce the carbon footprint of the steel­making operations but also allows the production of valuable commodities without the costs associated with feedstock gasification. The company has estimated that up to 30 bil­lion gallon of bioethanol per year can be produced from the CO-rich off gases produced through steel manufacturers globally [243]. Founded in 2005, LanzaTech has successfully demonstrated bioethanol production at a pilot plant at BlueScope Steel in Glenbrook, NZ, since 2008 and the company has recently started operating its 100,000 gallon bioethanol per year demonstration facility in Shanghai, China, using waste gas collected from an ad­jacent steel mill plant owned by its partner Baosteel Group [243, 244]. LanzaTech is plan­ning to build a commercial facility with the capacity to produce 50 million gallon of bioethanol per annum in China by 2013 [243]. The recent acquisition of a biorefinery fa­cility developed by the US-based gasification technology company Range Fuels in Geor­gia, and a milestone signing of its first commercial customer, Concord Enviro Systems (India), highlighted LanzaTech’s intention to utilize MSW and lignocellulosic waste as feedstocks for biofuel and chemical production [243, 244].

14. Conclusion

One of the fundamental factors that govern the environmental and economical sustaina­bility of biofuel production is feedstock. Through gasification, a spectrum of renewable non-food feedstock such as agricultural wastes, dedicated energy crops, forest residues, and MSW can be converted into syngas. This article presents a detailed examination of gas fermentation technology in capturing the carbon and energy from syngas and pro­duce biofuels and chemicals. In comparison to indirect fermentation of lignocellulose via enzymatic hydrolysis, and thermo-chemical FTP, gas fermentation offers several advan­tages such as good product yield and selectivity, operation in ambient conditions, high tolerance to gas impurities, and elimination of expensive pre-treatment steps and costly enzymes. Furthermore, some industries such as steel mill, natural gas steam reforming, oil refining and chemical production generate large volumes of CO-rich off-gas. Gas fer­mentation can access these existing feedstocks and generate valuable products from these while reducing carbon emissions. Pivotal to gas fermentation is acetogens such as C. Ijungdahlii, C. carboxidivorans, "C. ragsdalei" and C. autoethanogenum, which are able to me­tabolize CO, and CO2/H2 into a range of products such as ethanol, butanol, isopropanol, acetone, 2,3-butanediol, acetic acid and butyric acid. Sustained effort in studying the physiology and biochemistry using advanced molecular techniques such as genomics, transcriptomics, proteomics, metabolomics and systems biology are essential to further the understanding of these microbes. Furthermore, recent advances in Clostridial genetic tools offer endless opportunities to engineer strains that have improved product yield, sub­strate utilization, no strain degeneration, and synthesis of new products.

The main challenges associated with commercialization of gas fermentation have been identified as gas-to-liquid mass transfer limitation, product yield, substrate utilization effi­ciency, low biomass density and product recovery. Further development of bioreactor is necessary to improve the availability of gas substrates and maintain high cell density for higher productivity. Improvement in integrated product recovery technology is also es­sential to lower the costs of product recovery and alleviate product inhibition. Gas fer­mentation appears to be mature enough for commercialization since several companies have already demonstrated their technologies at pilot scale and are moving towards com­mercialization in the near future.

Author details

Fung Min Liew, Michael Kopke and Sean Dennis Simpson LanzaTech NZ Ltd., Parnell, Auckland, New Zealand

The Promising Fuel-Biobutanol

Hongjuan Liu, Genyu Wang and Jianan Zhang

Additional information is available at the end of the chapter http://dx. doi. org/10. 5772/52535

1. Introduction

In recent years, two problems roused peoples’ concern. One is energy crisis caused by the depleting of petroleum fuel. The other is environmental issues such as greenhouse effect, global warming, etc. Therefore, renewable sources utilization technology and bioenergy pro­duction technology developed fast for solving such two problems. Bioethanol as one of the biofuel has been applied in automobiles with gasoline in different blending proportions (Zhou and Thomson, 2009; Yan and Lin, 2009). Biobutanol is one of the new types of biofuel. It continuously attracted the attention of researchers and industrialists because of its several distinct advantages.

Biodiesel is typically synthesized from triacylglycerides derived from vegetable oils and an alcohol with base catalysis, yielding the fatty acid ester type biodiesel. Wahlen et al. determined conditions that allowed rapid and high yield conversion of oil feedstocks containing significant concentrations of free fatty acids into biodiesel using an acid-cata­lyzed reaction with longer chain alcohols such as n-butanol at a slight molar excess. Bio­diesel yields >98% were achieved in <40 min. Key properties of the resulting butyl-diesel were determined, including cetane number, pour point, and viscosity [229]. The batch and continuous-flow preparation of biodiesel derived from vegetable oil and 1-butanol using a microwave apparatus has been reported. The methodology allows for the reac­tion to be run under atmospheric conditions and in continuous-flow mode. It can be uti­lized with new or used vegetable oil with 1-butanol and a 1:6 molar ratio of oil to alcohol. Sulfuric acid or potassium hydroxide can be used as catalyst [230]. High conver­sion could be reached when the transesterification of triglycerides with 1-butanol was performed under near-critical or supercritical conditions with microwave heating [231].

Biodiesel synthesis by butanolysis of vegetable oils (soybean, sunflower, and rice bran) catalyzed by Lipozyme RM-IM), and the optimization of the enzyme stability over repeated batches has been described. The enzyme showed the highest activity at a 9:1 BuOH:oil molar ratio and in the 30-35 °C temperature range [232]. Transesterification reaction using sunflower oil and butanol catalyzed by immobilized lipases can be carried out without auxiliary solvent. Immobilized porcine pancreatic lipase (PPL) and Candida rugosa lipase (CRL) showed satisfactory activity in these reactions. Activities of immobilized lipases were highly increased in comparison with free lipases because its activity sites became more effective. Immobilized enzyme could be repeatedly used without difficult method of separation and the decrease in its activity was not largely observed [233].