Continuous fermentation is a preferred operational mode to decrease cost of production and increase efficiency. It can easily be performed with using cascade reactors with suppressing butanol concentration below the inhibition limit. The butanol concentration supressing can be performed by dilution or with various methods of recovery with adsorption, extraction, stripping, membrane techniques or with combination of these methods. Increase of the active biomass amount in the mash by cell recycling plays key role in continuous ABE fermentation processes, as well.

Dyr et al. [100] observed formation of neutral solvents in continuous ABE fermentation process by means of C. acetobutylicum without morphological adaptation due to the altered way of cul­tivation. The results obtained leave no doubt as to the possibility of employing the continuous method for acetone-butanol fermentation [101]. A cascade type continuous ABE fermentation method was developed from soluble starch by building an equipment consistsing a battery of 11 fermenting tanks [102]. The first tank is used as an incubator and an activator for the culture. In the remaining tanks, the actual fermentation is carried out. The feed liquor is continuously sup­plied. The continuous fermentation process for Me2CO-BuOH production is a lst-order reac­tion. A continuous ABE fermentation process was developed and adopted in plants using starch raw materials by Yarovenko [103]. The basis for a continuous process is knowledge of laws of continuous mixing of liquids in batteries of connected vessels which are discussed by Yarovenko [104]. The length of fermentation considerably influences the acidity of the ferment­ed mixture at the end of the process. Owing to differences in mash composition and duration of process, acid level is mostly higher in continuous fermentation than in a discontinuous one. With continuous acetone-butanol process fermentation, speed could be raised 1.58 times com­pared to the semicontinuous method. In the continuous fermentation, it is useful to operate with 2-5 parallel batteries and to cultivate bacteria in separated vessels. The carbohydrates produced by saccharification under different conditions were studied as they were of great importance on length and course of fermentation. Operation of the battery’s head fermentor has a great influ­ence on the whole process, the amount of inoculum, acid production, and fermentation speed. To provide an adequate microorganism concentration and to reduce the risk of infection in the battery’s head fermentor, mash from the 2nd vessel is recycled. The acidity increase was evident primarily in the last tank. Optimum concentration of the cells to be inoculated at the start of the fermentation 7*109/ml for C. acetobutylicum and physiologically mature cells should comprise about 80% of the total inoculum. The flow rate into the main fermentor should be harmonized with the utilization rate of carbohydrate in the battery. Bacteria in the main vessel must be main­tained at their respective stationary phase of growth. The continuous ABE fermentation in­creased productivity efficiency 20%. The carbohydrate utilization was improved by 2.4%, along with the characteristics of the beer [103,104]. The Japanese K. F. Engineering [105] described an apparatus for production of Me2CO and BuOH by immobilized ABE-producing microorgan­isms, where the immobilized microorganisms are first exposed to a batch process until active gas formation is observed, and then, a continuous production process was performed.

The availability and demand of biosynthetic energy (ATP) is an important factor in the regula­tion of solvent production in steady state continuous cultures of C. acetobutylicum. The effect of biomass recycle at a variety of dilution rates and recycle ratios on product yields and selectivi — ties was determined. Under conditions of non-glucose limitation, when the ATP supply is not growth-limiting, a lower growth rate imposed by biomass recycle leads to a reduced demand for ATP and substantially higher acetone and butanol yields. When the culture is glucose limit­ed, however, biomass recycle results in lower solvent and higher acid yields [106]. Wijjeswara — pu et al. studied continuous BuOH fermentation by C. acetobutylicum in a stirred tank reactor. The results of glucose fermentation with cell recycling revealed the formation of small amounts of EtOH, moderate amounts of Me2CO and BuOH, and large amounts of AcOH and butyric acid. Without cell recycling overall BuOH production was decreased by a factor of 3.5 [107]. Af — shar et al. used a cascade system and cell recycling. At a dry cell mass concentration of 8 g/L and a dilution rate of D=0.64 h-1, a solvent productivity of 5.4 g/L-1 h-1 could be attained. To avoid de­generation of the culture which occurs with high concentrations of ABE solvents a 2-stage cas­cade with cell recycling and turbidostatic cell concentration control was used as optimal solution, the 1st stage of which was kept at relatively low cell and product concentrations. A sol­vent productivity of 3 and 2.3 g L-1 h-1, respectively, was achieved at solvent concentrations of 12 and 15 g L-1 [108]. Huang and Ramey [109] determined the influence of dilution rate and pH in continuous cultures of Clostridium acetobutylicum in a fibrous bed bioreactor with high cell density and butyrate concentrations at pH 5.4 and 35°C. By feeding glucose and butyrate as co­substrates, the fermentation was maintained in the solventogenesis phase, and the optimal bu­tanol productivity of 4.6 g L-1 h-1 and a yield of 0.42 g g-1 were obtained at a dilution rate of 0.9 h-1 and pH 4.3. Eight Clostridium acetobutylicum strains were examined for a-amylase and strains B-591, B-594 and P-262 had the highest activities. Defibered-sweet-potato-slurry containing starch supplemented with potassium phosphate, cysteine-HCl, and polypropylene glycol was used as continuous feedstock to a multistage bioreactor system. The system consisted of four columns (three vertical and one near horizontal) packed with beads containing immobilized cells of C. acetobutylicum P-262. The effluent contained 7.73 g solvents L-1 (1.56 g acetone; 0.65 ethanol; 5.52 g butanol) and no starch. Productivity of total solvents synthesized during contin­uous operation was 1.0 g L-1 h-1 and 19.5% yield compared to 0.12 g L-1 h-1 with 29% yield in the batch system [110]. Pierrot et al. introduced a hollow-fiber ultrafiltration to separate and recycle cells in continuous ABE fermentation. Under partial cell recycling and at a dilution rate of 0.5 h-1, a cellular concentration of 20 g L-1 and a solvent productivity of 6.5 g L-1 h-1 is maintained for sev­eral days at a total solvent concentration of 13 g L-1 [111]. The device developed was sterilizable by steam and permitted drastic cleaning of the ultrafiltration membrane without interrupting continuous fermentation. With total recycle of biomass, a dry weight concentration of 125 g L-1 was attained, which greatly enhanced the volumetric solvent productivity averaging 4.5 g L-1 h-1 for significant periods of time (>70 h) and maintaining solvent concentration and yield at accept­able levels [112].

A stable continuous production system with nongrowing cells of C. acetobutylicum adsorbed to beechwood shavings was obtained by different types of adsorption procedures for produc­tion of ABE solvents by Foerberg and Haegsstroem [113]. The system was started with continuous flow of a complete nutrient medium. A thick cell layer was formed on the wood shavings during the 1st day but it disappeared rapidly. Under glucose limitation, a new cell layer developed during the following period (2-5 days). After this phase, a continuous flow of nongrowth medium with nutrient dosing (8 h dosing interval) was started. This led to a washout of most adsorbed cells and ~85% of suspended cells. Another cell layer was formed during this period and the system was controlled by the nutrient dosing technique. The system was stable with no cell leakage for weeks. The maximal productivity of butanol, acetone, and EtOH was 36 g L-1 d-1 with a product ratio of 6:3:1 [113].

A continuous ABE production system with high cell density obtained by cell-recycling of Clos­tridium Saccharoperbutylacetonicum N1-4 was also studied. In a conventional continuous ABE culture without cell-recycling, the cell concentration was below 5.2 g L-1 and the maximal ABE productivity was only 1.85 g L-1 h-1 at a dilution rate of 0.20 h-1. To obtain a high cell density at a faster rate, we concentrated the solventogenic cells of the broth 10 times by membrane filtra­tion and were able to obtain ~20 g L-1 of active cells after only 12 h of cultivation. Continuous cul­ture with cell recycling was then started, and the cell concentration increased gradually through cultivation to a value greater than 100 g L-1. The maximum ABE productivity of 11.0 g L-1 h-1 was obtained at a dilution rate of 0.85 h-1. However, a cell concentration >100 g L-1 resulted in heavy bubbling and broth outflow, which made it impossible to carry out continuous culture. There­fore, to maintain a stable cell concentration, cell bleeding and cell recycling were performed. At dilution rates of 0.11 h-1 and above for cell bleeding, continuous culture with cell recycling could be operated for more than 200 h without strain degeneration and an overall volumetric ABE productivity of 7.55 g L-1 h-1 was achieved at an ABE concentration of 8.58 g L-1 [114].

Biofuels production via the Fischer-Tropsch technology is a conversion process of solid bio­mass into liquid fuels (Biomass-To-Liquid or BTL) as it is depicted in Figure 2. More specifi­cally the solid biomass is gasified in the presence of air and the produced biogas rich in CO and H2 (synthesis gas), after being pretreated to remove coke residues and sulfur com­pounds, enters the Fischer-Tropsch reactor. The Fischer-Tropsch reactions allow the catalytic conversion of the synthesis gas into a mixture of paraffinic hydrocarbons consisting of light (Q-C4), naphtha (C5-Cu), diesel (C12-C20) and heavier hydrocarbons (>C20). Even though the Fischer-Tropsch reactions yields depend on the catalyst and operating parameters employed [43—45], the liquid product (naphtha, diesel and heavier hydrocarbons) yield is high (~95%). The produced synthetic naphtha and diesel fuels can be used similarly to their fossil coun­terparts. The heavier product however, which is called as Fischer-Tropsch wax, due to its waxy/paraffinic nature should get upgraded via catalytic hydrocracking to get converted to mid-distillate fuels (naphtha and diesel).

The conversion of Fischer-Tropsch wax into mainly diesel was studied in virtue of the Euro­pean Project RENEW [46]. During this project Fischer-Tropsch wax with high paraffinic con­tent of C20-C45 was converted into a total liquid product consisting of naphtha, kerosene and diesel fractions via catalytic hydrocracking. However the total liquid product content of die­sel molecules was the highest and the diesel fraction was further separated and character­ized having density of 0.78gr/ml and cetane index of 76 [47]. The schematic of the BTL process with actual images of the feedstock, Fischer-Tropsch wax and synthetic diesel are given in Figure 8.

As a replacement for IB, TBA was first used in the synthesis of ETBE over 80 years ago uti­lizing concentrated H2SO4 as a catalyst as shown in reaction scheme in Figure 2 [11]. Habe- nicht et al. [12] suggested that TBA was preferred over IB as a reactant for ETBE synthesis at elevated pressures and temperatures. The reason for this is that the protonated IB (the key component in ETBE formation) forms only from TBA (not IB) under the conditions em­ployed. Yin et al. [13] also studied liquid-phase synthesis of ETBE from TBA and EtOH cata­lyzed by ion-exchange resin and heteropoly acid (HPA) at mild pressures and temperatures. Knifton et al. [7] also investigated different types of zeolites catalysts for direct synthesis of ETBE from TBA and EtOH. At temperature ranges of 40-140 oC and pressure ranges of 0.1-7 MPa, liquid-phase synthesis of ETBE resulted to a 40-70 % yield and 65-95 % selectivity.

In our previous work, reactive distillation, a configuration in which the reactive section was located inside the column, was employed to continuously synthesize ETBE from bioe­thanol and TBA using Amberlyst 15 in pellet form as a catalyst. Results under standard operating conditions indicated that ETBE at about 60 mol% could be obtained in the distil­late, and almost pure water in the residue. The conversion of TBA and the selectivity of ETBE were 99.9 and 35.9 %, respectively. The effects of operating conditions on conver­sion and selectivity were also investigated. Further purification of the distillate using the residue results in 95 mol% ETBE. Simulation of the process was also carried out using AS­PEN PLUS simulator, and results showed good agreement with the obtained experimental results as shown in Figure 3 [14].

Biogasification (or anaerobic digestion) is a biochemical process that converts organic matter to biogas (a mixture of methane, 50-70%, and balance carbon dioxide) under anaerobic con­ditions. Biogas can be used as a replacement for natural gas or it can be converted to electric­ity. The process is mediated by a mixed, undefined culture of microorganisms at near ambient conditions. Several terrestrial biomass feedstocks (agricultural residues, urban or­ganic wastes, animal wastes and biofuel crops) have been anaerobically digested and com­mercial scale digesters exist for the biogasification of such feedstocks.

Anaerobic digestion offers several advantages over other biofuel production processes like ethanol fermentation or thermochemical conversion. The microbial consortia in an anaerobic digester are able to naturally secrete hydrolytic enzymes for the solubilization of macromo­lecules like carbohydrates, proteins and fats. Therefore, unlike in ethanol fermentation proc­ess there is no need to incorporate a pretreatment step to solubilize the macromolecules prior to fermentation. In addition, since the process is mediated by a mixed undefined cul­ture, issues of maintaining inoculum (or culture) purity does not arise. Being a microbial process, there is no need to dewater the feedstock prior to processing unlike in thermochem­ical conversion where the feedstock is dried, to improve net energy yield. This is advanta­geous when it comes to processing aquatic biomass as these can be processed without dewatering. The anaerobic digestion process will also mineralize organic nitrogen and phos­phorous, and these nutrients can be recycled for algae growth [37].

The process primarily takes place in four steps. A mixed undefined culture of mciroorgan — isms mediates hydrolysis, fermentation, acetogenensis and methanogenesis of the organic substrates as shown in Figure 3. During hydrolysis, the complex organic compounds are broken down into simpler, soluble compounds like sugars, amino acids and fatty acids. These soluble compounds are fermented to a mixture of volatile organic acids (VOA). The higher chain VOAs like propionic, butyric, and valeric acids are then converted to acetic acid in the acetogenesis step. Acetic acid is converted to methane during methanogenesis. Hydrogen and carbon dioxide are also liberated during fermentation and acetogenesis. A different group of methanogens converts hydrogen and carbon dioxide to methane. This mixed microbial culture thrives in the pH range of 6-8. Digestion can be performed either at mesophilic conditions (30 — 38°C) or thermophilic conditions (49 — 57°C).

Aquatic biomass — macrophytes [38], micro and macro algae, have all been tested as feed­stock for biogasification. Microalgae have proportions of proteins (6-52%), lipids (7-23%)

and carbohydrates (5-23%) that are strongly dependent on the species and environmental conditions [39—41]. Compared with terrestrial plants microalgae have a higher proportion of proteins, which is characterized by a low carbon to nitrogen (C/N) ratio. The average C/N for freshwater microalgae is around 10.2 while it is 36 for terrestrial plants [40]. Usually the digestion of terrestrial plants is limited by nitrogen availability; however for microalgae this situation does not arise. Besides carbon, nitrogen and phosphorus, which are major compo­nents in microalgae composition, oligo nutrients such as iron, cobalt, zinc are also found [42]. These characteristics of microalgae make it a good feedstock for anaerobic digestion.

Previous studies have shown that macro algae like Ulva lactuca, Gracillaria vermiculophylla, Saccharina latissima etc. can be anaerobically digested producing methane at yields ranging from 0.1-0.3 LCH/g volatile solids (VS) [43]. Methane yields of microalgae like Spirulina pla — tensis (fresh water), and Scenedesmus spp. and Chlorella spp. (fresh water) ranged between 0.2 and 0.3 L CH4/g VS [44, 45] when these were codigested with other feedstocks like dairy manure and waste paper sludge, whereas other microalgae like Tetraselmis sp (marine), Chlorella vulgaris (fresh water), Scendesmus obliquess (fresh water) and Phaeodactylum tricornu — tum (fresh water) produced an average methane yield ranging from 0.17 to 0.28 L CHyg VS [45—47] when digested as sole feedstock. Table 3 summarizes microalgae digestion studies reported in the literature. The Table also lists the methane yield of cellulose powder as a benchmark to compare the methane potentials of microalgae feedstocks. Depending on the

type of microalgae, the methane potentials range from 5 to 78% of methane potential of cel­lulose. Choice of microalgae has an impact on the methane yield.

More recently when Nannochloropsis oculata was biogasified [48] in laboratory scale digesters at thermophilic temperature, the methane yield obtained was 0.20 L at STP/g VS. N. oculata was chosen because it can be grown easily in brackish or seawater, has a satisfactory growth rate and can tolerate a wide range of pH (7-10) and temperature (17 — 27° C). N. oculata is not rich in lipids but contains predominantly cellulose and other carbohydrates, which makes it a good feedstock for anaerobic digestion instead of biodiesel production. On a % (w/w dry matter) basis, the composition of N. oculata is: 7.8% carbohydrate, 35% protein and 18% lip­id. Rest of the components are amino acids, fatty acids, omega-3, unsaturated alcohols, as­corbic acid [49]. About 88% of the carbohydrate is polysaccharide. Of the polysaccharides, 68.2% is glucose, and the rest are fucose, galactose, mannose, rhamnose, ribose and xylose.

Based on N. oculata growth observed in the pilot raceways and the methane yield from di­gestion of this alga, an analysis was carried out to estimate energy production and land re­quirements. Currently the algae harvesting rate from the raceways are 9.64 g ash free dry weight (afdw)/m2/d. Note that afdw (ash free dry weight) is the same as volatile solids con­tent. An often cited study for algae growth has yielded a much higher productivity of 50 g afdw/m2/d for Platyomonas sp [50]. The algae biomass yield obtained in this study was only about 20% of the productivity potentially attainable. Optimization of growth conditions for N. oculata may improve its productivity. Using the methane yield value of 204 L/kg VS for anaerobic digestion of N. oculata, the annual energy output from a facility that grows the al­gae and subsequently digests it would be 27 MJ/m2/year. The area occupied (or footprint) of the digester(s) would be far less than the land area required for growing the algae. If the methane produced from this facility is converted to electricity, the electrical energy output would be 2.25 kWHe/m2/year assuming that the efficiency of converting thermal energy to electrical energy is 30%. The household electrical energy and natural gas consumption in the US for the year 2010 was 11,496 kWH/year and 2070 m3/year respectively. If the algae bioga­sification facility were to supply the entire electrical energy requirements for a household, the land area required would be 5108 m2 (1.26 acres). If in addition, the facility were to sup­ply the natural gas needs, then an additional 2900 m2 (0.77 acres) would be needed. In other words ~2 acres of land could supply all the energy needs of a household in US. If the algae productivities were improved then land requirement could be further reduced. At 50 g afdw/m2/d algae productivity, the land requirement would only be about 0.4 acres.

Despite useful methane production potential from biogasification and the ability to process dilute algal slurries in a digester, there are challenges to be overcome to commercialize this approach for producing bioenergy from microalgae. One bottleneck is that some feedstock characteristics can adversely affect anaerobic digestion. Unlike defined cultures used for production of biofuels like ethanol or butanol, the microbial consortia in an anaerobic di­gester is capable of secreting extracellular enzymes to hydrolyze and solubilize macromole­cules like cellulose, hemicellulose, proteins and fats. This characteristic has enabled several terrestrial biomass feedstocks like sugarbeets, sugarbeet tailings, napier grass, sorghum and aquatic biomass like water hyacinth and giant kelp to be successfully digested using practi­cal retention times. However, degradability of feedstocks containing high fraction of lignin (for example sugarcane bagasse, switchgrass, miscanthus and woody biomass like pine, eu­calyptus) is poor in an anaerobic digester. The refractoriness of these feedstocks has been at­tributed to low moisture, crystalline nature of the cellulose, and complex association of the component carbohydrates within lignin [51]. As seen from Table 3, the digestibility of micro­algae varies. Species with no cell wall or cell encapsulation composed of proteins like Chlor — ella vulgaris and Phaeodactylum tricomutum, has a higher yield of methane. Dunaliella tertiolecta has very low methane yield of 0.018 L/kg VS due to the presence of a cell wall con­sisting of cellulose fibers distributed within an organic matrix. So depending on the type of microalgae used it may be necessary to carry out some form of pretreatment of algae to im­prove methane yield and rate of methane production. The type of pretreatment may depend on algae type.

4. Conclusion

Aqueous and marine biomass can be processed into a variety of sources of energy. Due to the extreme dilution in water, non-thermal processes such as anaerobic digestion, fermenta­tion to bioalcohols, and lipid extraction are logical and useful methods to utilize key compo­nents of microorganisms to produce biofuels for the replacement or supplementing of traditional fossil fuels. However, thermal methods such as gasification of wet biomass may play a role in producing specialty fuels such as jet fuel that require a specific ratio of higher hydrocarbons that would prove otherwise difficult to manufacture, even given the require­ment of intense drying.

In order for biofuels sourced from aqueous and marine biomass to secure a market share in the world, research and development needs to further nature’s ability to produce higher concentrations of biomass with targeted characteristics and reduced footprints, while better utilizing available nutrients. This will allow for an ample supply of biomass to be produced without competition with the human food chain, that can be used renewably produce fuel that can power the world’s mobile fleet.

Author details

Robert Diltz1 and Pratap Pullammanappallil2 *Address all correspondence to: Robert. diltz@us. af. mil

1 Air Force Research Laboratory, Tyndall AFB, FL, USA

2 Department of Agricultural and Biological Engineering, University of Florida, Gainesville, FL, USA

Except the use of solvent, chemical intermediate and extract agent, butanol also can be used as fuel, which attracted people’s attention in recent years. Because of the good properties of high heat value, high viscosity, low volatility, high hydrophobicity, less corrosive, butanol has the potential to be a good fuel in the future. The properities of butanol and other fuels or homologues are compared as Table 2. (Freeman et al., 1988; Dean, 1992)

Butanol appeared the good properties compared with it’s homologues such as 2-butanol, iso-butanol and tert-butanol and other fuels such as Gasoline and ethanol. Actually, when ethanol is mixed with gasoline (less than 10%), there exists some disadvantages. Firstly, the heating value of ethanol is one sixth of gasoline. The fuel consumption will increase 5% if the engine is not retrofitted. Secondly, acetic acid will be produced during the burning proc­ess of ethanol, which is corrosive to the materials of vehicle. The preservative must be added when the ethanol proportion upper than 15%. Thirdly, ethanol is hydroscopic and the liquid phase separation may be occurring with high water proportion. Furthermore, ethanol as fuel cannot be preserved easily and it is more difficult in the process of allocation, storage, transi­tion than that of gasoline.

Compared with ethanol, butanol overcomes above disadvantages and it shows potential ad­vantages. For example, Butanol has higher energy content and higher burning efficiency, which can be used for longer distance. The air to fuel ratio and the energy content of butanol are closer to gasoline. So, butanol can be easily mixed with gasoline in any proportion. Buta­nol is less volatile and explosive, has higher flash point, and lower vapor pressure, which makes it safer to handle and can be shipped through existing fuel pipelines. In addition, Bu­tanol can be used directly or blended with gasoline or diesel without any vehicle retrofit (Durre, 2007; Pfromm et al., 2010).

Actually, the first-time synthesis of biobutanol at laboratory level was reported by Pasteur in 1861 (Durre, 1998) and the industrial synthesis of biobutanol was started during 1912­1914 by fermentation (Jones and Woods, 1986). However, before 2005, butanol was mainly used as solvent and precursor of other chemicals due to the product inhibition and low bu­tanol productivity. To bring awareness to butanol’s potential as a renewable fuel, David Ra­mey drove his family car from Ohio to California on 100% butanol (http:// www. consumerenergyreport. com /2011/02/09/reintroducing-butanol/). And then, two giant companies DuPont and BP have declared to finance development of a modernize produc­tion plant supported by research and development. (http://biomassmagazine. com/articles/ 2994 /eu-approves-bp-dupont-biobutanol-venture) The economy of biobutanol production also was revaluated. The research of a continuous fermentation pilot plant operating in Aus­tria in the 1990s introduced new technologies and proved economic feasibility with agricul­tural waste potatoes. (Nimcevic and Gapes, 2000).

Anne M. Ruffing

Additional information is available at the end of the chapter http://dx. doi. org/10.5772/52050

1. Introduction

The world’s supply of petroleum hydrocarbons, which serve as feedstock for the fuel and chemical industries, is rapidly diminishing to satisfy the global demand for energy and consumer goods. In response to this increasing demand and limited supply, the cost of crude oil has risen to over $100 per barrel in 2012, a 10-fold increase compared to prices in the late 1990s [1]. As fossil fuels are nonrenewable resources, the price of oil is only expected to increase in the future. This unavoidable reality necessitates the development of renewable energy sources in order to maintain the current standard of living. Among the alternative energy options under development, biofuels are anticipated to supplement and eventually replace the petroleum-based fuels that supply the transportation and chemical industries. Currently, first generation biofuels like corn-based ethanol are blended into conventional petroleum fuels, with biofuels supplying 2.7% of the world’s transportation fuel in 2010 [2]. It appears that biofuels are on their way to becoming a viable renewable energy source, yet technological and biological advancements are necessary for sustainable and economical biofuel production at the scales necessary to support the world’s energy needs.

The current practice of using food crops, like corn or soybean, as feedstocks for biofuel production is not a viable, long-term solution to the energy crisis. In fact, to replace our current petroleum usage with crop-based ethanol production, the entire surface area of land on Earth would be needed for corn production [3]. In addition to this shortcoming, first generation biofuels compete with food production for arable land, require significant nutrient resources (fertilizer and fresh water), and typically have low net energy yields due to the low energy density of the product fuel (i. e. ethanol) and the energy input required to harvest the feedstock and convert it into fuel [4]. Second and third generation biofuels address these limitations. Second generation biofuels use lignocellulosic biomass as the feedstock for fuel production.

Lignocellulose, the main component of plant biomass, is the most abundant form of renewable carbon on the Earth, making it an ideal feedstock for renewable hydrocarbon production. The cellulose and hemicellulose components of lignocellulose can be degraded into fermentable sugars to serve as the carbon source for microbial-based fuel production. The carbon feedstocks for both first and second generation biofuels are ultimately derived from carbon dioxide (CO2) fixation through the process of photosynthesis. Third generation biofuels use photo­synthetic microorganisms (i. e. microalgae) to directly convert CO2 into fuel molecules or fuel precursors, eliminating the biomass intermediate (Figure 1). While both second and third generation biofuels require land, nutrients, and energy investment for harvesting and fuel production, the fuel production yields from these processes are predicted to be capable of meeting energy needs. However, these technologies have yet to be demonstrated at scale and still require further improvement before they can be economically competitive with fossil fuels.

Both second and third generation biofuels rely on microbes to convert the carbon feedstock into the desired hydrocarbon fuels. Microorganisms have been identified that are capable of producing a range of fuel molecules and fuel precursors, yet the natural rates of microbial fuel synthesis are typically too low to support industrial-scale production. Metabolic engineering is a powerful tool to improve microbial fuel production, either through engineering the metabolic pathways within the native microorganism to encourage high fuel synthesis or though transferring the fuel production pathway into a model organism for optimization. This chapter will focus on the application of metabolic engineering to increase hydrocarbon fuel production. Within this chapter, hydrocarbon-based fuels are defined to include oxygen — containing fuel molecules with long hydrocarbon chains, such as fatty alcohols and fatty acid ethyl esters (FAEE), in addition to pure hydrocarbons like alkanes, alkenes, and isoprenoid — based molecules: hemiterpene (C5), monoterpenes (C10), and sesquiterpenes (C15). Hydro­carbon-based fuel precursors will also be considered, including free fatty acids (FFAs) and triacylglycerol (TAG). The structures of these hydrocarbon-based fuels and precursors are illustrated in Figure 2. Hydrocarbon-based fuels and precursors can be produced by both second and third generation biofuel processes. Therefore, the first section in this chapter will

discuss the metabolic pathways for hydrocarbon fuel production and common metabolic engineering strategies for improving fuel synthesis. Because second and third generation biofuel processes rely on different carbon sources, sugars and CO2 respectively, the remaining sections will focus on the use of organic carbon (heterotrophy) and inorganic carbon (auto­trophy) as feedstocks for biofuel production. This division, based on carbon source, is impor­tant from both the biofuel production and metabolic engineering perspectives. The chapter will conclude with a discussion of the future outlook for microbial-based, hydrocarbon fuel synthesis.

Figure 2. Chemical structures of hydrocarbon-based biofuels and fuel precursors. (A) Fuels derived from fatty acid bio­synthesis and (B) fuels derived from isoprenoid biosynthesis, including (1) hemiterpene, (2) monoterpenes, and (3) ses­quiterpenes.

Biodiesel is currently obtained from the transesterification reaction of vegetable oils. A pos­sible drawback of this technology is that large investment is required to build up new bio­diesel plants. An interesting alternative is to directly feed the vegetable oil into the hydrotreating unit of a petroleum refinery, for instance, vegetable oil can be co-fed with heavy vacuum oil HVO. Under typical hydrotreating conditions (300-450°C, 50 bar H2 pres­sure, sulfidedNiMo/Al2O3 catalyst), vegetable oils are transformed into alkanes through three different pathways: decarboxylation, decarbonylation and HDO. The straight chain al­kanes can undergo isomerization and cracking to produce lighter and isomerized alkanes (Figure 8) [37]. It was reported that mixing the sunflower oil with HVO does not decrease the rate of desulfurization. Moreover, the rate of vegetable oil hydrotreating is faster that the rate of HVO desulfurization. For industrial application, corrosion problems should be taken into account and the formation of waxes should be minimized, as they can plug the reactor.

Isomerization lso"^i5

Propane

Cracking lighter alkanes

HsO

Figure 8. Reaction pathway for conversion of tri-glycerides to alkanes [37].

Fatty alcohols can be obtained by catalytic hydrogenolysis of fatty acid methyl esters. Small- chain fatty alcohols are used in cosmetics and food and as industrial solvents or plasticizers, while the large-chain fatty alcohols are important as biofuels and as nonionic surfactants or
emulsifiers. Fatty alcohols are produced by hydrogenolysis, in the presence of Cu based het­erogeneous hydrogenation catalysts, operating under H2 pressures between 20 and 30 bar and temperatures in the range of 97-197°C [38]. High hydrogen pressures are required to in­crease the solubility of hydrogen in the reaction mixture, in order to boost the availability of H2 at the catalyst surface and to reduce mass transport limitations [39].The stoichiometry of the reaction is presented below:

R-COOCH3+ 2H2 — R-CH2OH + CH3OH

A research in order to determine the initial feasibility to generate biohydrogen from urban organics wastes and then established some conditions to operate a batch bioreactor was developed in Colombia. This section presents the results of this research and analysis the potential use of urban wastes as sources to generate hydrogen.

1.2. Localization

The research was performance between the years 2009 and 2012, at the Laboratory of Agri­cultural Mechanization of the National University of Colombia in Medellm, localized in 6°13’55"N and 75°34’05"W, with average annual temperature of 24°C, relative humidity of 88% and average annual precipitation of 1571mm.

1.3. Methods

Two stages were established to develop the research, the first had five phases.

1.3.1. First stage

Phase 1. Identification of organic wastes generated at the Central Wholesaler of Antioquia

The Central Wholesaler of Antioquia is the main company dedicated to trade food in the city of Medellin (fruits, vegetable and some grains). At the first phase historical information related to organic wastes production during two year was supplied by Central Wholesaler of Antio- quia and was made a photographic register of solids wastes generated. The photographs were taken twice per day at the morning and afternoon.

Phase 2. Selection of wastes with greater production

According to the information collected and the photographic register from the first phase, the wastes with greater production were selected to be introduced into a batch bioreactor.

Phase 3. Elemental Composition and chemical composition analysis

The quantity of volatile solids, total solids and elemental composition on both wet and dry basis (coal, nitrogen and hydrogen) were obtained for each wastes. Were taken samples of 5 grams and the analysis method applied was the Wendee method (the analysis was made at the chemical analysis laboratory of National University in Medellin). With that information was calculated the quantity of wastes to use. Six samples of 3 grams in each wastes were taken in order to obtain the elemental analysis, in this case the method applied was burn of sample and the equipment employed was an elemental analyzer CE — 440 (Figure 2a). The samples were triturated with a precision crusher — IKA WERNE with sieve of 0,5 mm (Figure 2b) and then were dried in a lyophilizer LABCONCO Freezone 12L (Figure 2c). In order to determine the quantity of wastes and water to be employed, 6 grams of volatile solids per liter-day were used as organics load [19], additionally was employed a concentration on volatile solids of 5% [20].

A batch bioreactor of 2000 liters was installed, the wastes were triturated to facilitate its access into bioreactor and its process by the bacteria. The quantity of gas generated was registered with a gas flow meter Metrex G 2,5 with accurate of 0,040 m3/h; maximum pressure of 40 kPa, additionally was employed a gel of silica to remove the wet of gas. The load of bioreactor was made during four days, each day was used the same quantity until to complete the total load.

Phase 5. Principal variables to register

The relativity humidity and environment temperature were registered daily, was used a thermohygrometer with rank in temperature until 120°C and 100% in relativity humidity (Figure 4). The pH into the bioreactor was registered daily too, in this case was employed a digital pH-meter Hanna Instruments, with accurate of ± 0,2 (reference temperature of 20°C).

Figure 4. Thermohygrometer and pH-meter

The organics load was determined at the beginning and end of bioprocess; in this case the total suspended solids (TSS), total solids (TS), volatile fatty acids (VFAs), chemical oxygen demand (COD) and biochemical oxygen demand (BOD) were determined. The analytics method employed were Standard Method by water and residual water of the APHA-AWWA-WPCF, edition 19 of 1995.

The production of gas was registered daily, samples were collected in Tedlar bags (with capacity of 1 liter, Figure 5) and then were analyzed in a chromatographic gas (Perkin Elmer) to establish its composition (percentage of CO2 O2 H2 CH4 and N2). During the tests, the wastes were subjected to an acid pretreatment to eliminate the methanogenic bacteria, after several days, agricultural lime was added to increase the pH until to obtain a value most adequate to the acidogenic bacteria.

Due to the excessive exploitation, the fossil fuels are facing scarce and they cannot be generated. On the other hand, most of the carbon emissions result from fossil fuel combustion. Reducing the use of fossil fuels will considerably reduce the amount of carbon dioxide and other pollutants produced. Renewable energy has the potential to provide energy services with low emissions of both air pollutants and greenhouse gases. Currently, renewable energy sources supply over 14% of the total world energy demand. Biofuels as the important renewable energy are generally considered as sustainability, reduction of greenhouse gas emissions, regional development, so­cial structure and agriculture, and security of supply (Reijnders, 2006). Biodiesel and bioethanol are presently produced as a fuel on an industrial scale, including ETBE partially made with bioe­thanol, these fuels make up most of the biofuel market (Antoni et al., 2007).

Biobutanol also has a promising future for the excellent fuel properties. It has been demon­strated that n-butanol can be used either 100% in unmodified 4-cycle ignition engines or blended up with diesel to at least 30% in a diesel compression engine or blended up with kerosene to 20% in a jet turbine engine in 2006 (Schwarz et al., 2006). The production of bio­butanol from lignocellulosic biomass is promising and has been paid attention by many companies. Dupont and BP announced a partnership to develop the next generation of bio­fuels, with biobutanol as first product (Cascone, 2007). In 2011, Cobalt Technologies Compa­ny and American Process Inc. (API) have been partnering to build an industrial-scale cellulosic biorefinery to produce biobutanol. Additionally, the companies agreed to jointly market a GreenPower+ biobutanol solution to biomass power facilities and other customers worldwide. The facility is expected to start ethanol production in early 2012 and switch to biobutanol in mid-2012. The annual production of biobutanol is estimated to 470, 000 gal­lons. (http://www. greencarcongress. com/2011/04/cobalt-20110419.html, http://www. renewa- bleenergyfocususa. com/view/17558/cobalt-and-api-cooperate-on-biobutanol/) Gevo, Inc. signed a Joint Development Agreement with Beta Renewables, a joint venture between Chemtex and TPG, to develop an integrated process for the production of bio-based isobuta­nol from cellulosic, non-food biomass, such as switch grass, miscanthus, agriculture resi­dues and other biomass will be readily available. (http://www. greencarcongress. com/ biobutanol/). Syntec company also is currently developing catalysts to produce bio-butanol from a range of waste biomass, including Municiple Solid Waste, agricultural and forestry wastes. (http://www. syntecbiofuel. com/butanol. php). Utilization the waste materials im­prove the economy of butanol production that makes biobutanol great potential to be the next new type of biofuel in spite of the existing drawbacks.

7. Conclusions

Biobutanol production has only recent years booming again after long time of silence. Quite a lot of progress has been made with the technology development of metabolic engineering in enhancing solvent production, increasing the solvent tolerance of bacteria, improving the selectivity for butanol. Fortunately, Clostridia have been tested being able to consume ligno — cellulosic biomass for ABE fermentation. The complex regulation mechanism of butanol synthesis is still need to be further study. For the strain improvement, for example, con­structing better butanol tolerance strains, more suitable hosts and genetic methods are re­quired to be set up. Furthermore, more efficient techniques for removing the inhibitors in the lignocellulosic hydrolysate need to be developed. In addition, from the economic point of view, the integrated system of hydrolysis, fermentation, and recovery process also are im­portant to be further developed to reduce the operation cost of butanol synthesis.

Author details

Hongjuan Liu*, Genyu Wang and Jianan Zhang

*Address all correspondence to: liuhongjuan@tsinghua. edu. cn; zhangja@tsinghua. edu. cn Institute of Nuclear and New Energy Technology, Tsinghua University, Beijing, P. R., China

The microbial production of drop-in replacement fuels faces unprecedented challenges. The sheer quantity of hydrocarbon product required to meet the world’s ever increasing demand for energy dwarfs the supply of any current microbially synthesized product. Moreover, both second (lignocellulosic feedstock) and third (microalgal feedstock) generation bio­fuels ultimately rely on sunlight and photosynthesis to supply the energy and carbon feedstocks necessary for production. This requires the development of new technology and infrastructure to facilitate the construction of this new supply chain. Finally, the low value of the final fuel product places additional financial restrictions on the development of large — scale biofuel production processes. For example, previous reports include the addition of exogenous metabolic precursors like mevalonate for isoprenoid production or FFA for FAEE biosynthesis [18, 50]. While these exogenous metabolites boost production of the desired hydrocarbon-based product, this practice is too expensive for large-scale biofuel applica­tions. These challenges currently limit the industrial production of second and third generation biofuels.

Fortunately, new biological and technological tools are rapidly being developed and applied to overcome the obstacles in biofuel production. In addition to the metabolic engineering strategies previously described in this chapter, new global strategies are being applied to engineer microbes for biofuel production. With the affordability of next-generation DNA sequencing technologies, new microbial genomes are being reported at an unprecedented rate, and this information can be used to generate metabolic models for biofuel-producing hosts. In turn, these models can be leveraged to analyze proposed metabolic engineering strategies in silico, reducing the number of costly and time-intensive strain constructions and experi­ments. This technique was shown to be successful at increasing lycopene production, an isoprenoid derivative, in E. coli [69, 138]. The advancement of synthetic DNA technology enables new engineering approaches such as multiplex automated genome engineering (MAGE) [139]. In MAGE, synthetic oligomers, consisting of degenerate DNA sequences flanked by regions homologous to the target sequences, are simultaneously transformed into E. coli, and the modified strains are screened for improvements. MAGE was used to target ribosome binding sites, for optimization of protein translation, and to inactivate genes by inserting nonsense mutations; this technique can also be applied to target promoters for improved gene transcription and enzyme active sites for enhanced activities. The technique does have some limitations, however. MAGE will likely require modification of the host organism to allow for efficient integration of the single-stranded oligonucleotides, and a high — throughput screening method is essential for screening the billions of genetic variants that are generated with MAGE. Global or systems-level technologies can also be applied to advance our fundamental understanding of genetic and regulatory mechanisms within a microbial host; this is vital to host development of non-model organisms and newly isolated strains. Omics technologies including genomics, transcriptomics, metabolomics, and proteomics provide global insight at the cellular level, which can be compared across different conditions or time points to identify the native mechanisms that control the cell metabolism. Integration of omics data can identify bottlenecks at the transcriptional, translational, and protein levels, and as such, can be applied to inform the metabolic engineering strategy for biofuel production [34]. Systems-level tools for engineering microbial hosts, including metabolic modeling, MAGE, and omics technologies, will be integral to the successful development of hosts for biofuel production.

Commercial interest in the production of second and third generation biofuels has developed rapidly in the past decade. As evidence of this, there has been a flurry of activity in patent applications regarding microbial hydrocarbon production. Companies invested in heterotro­phic hydrocarbon-based fuel production include LS9 [27, 59, 65, 66, 140, 141] and Amyris Biotechnologies [72, 142], which focus mainly on E. coli as the host, and Solazyme [143, 144], which initially focused on fuels derived from algae but has since moved toward more high — value markets, such as cosmetics and nutraceuticals. Most companies interested in algae and cyanobacteria are focused on autotrophically-produced hydrocarbon fuels. Notable compa­nies in this industry include Sapphire Energy [145, 146], Joule Unlimited [26, 77, 147], and Synthetic Genomics [68, 75]. The hydrocarbon-based fuels targeted by these companies span the entire gamut of fatty acid and isoprenoid derived fuel products. Despite this commercial interest, hydrocarbon biofuel production still remains to be demonstrated at scale and in a sustainable manner.

This chapter has described the challenges in microbial hydrocarbon production and presented metabolic engineering strategies to resolve these issues. As is evident from this discussion, microbial-based fuel production is only in the initial stages of exploration, and additional research and innovation is necessary to enable large-scale biofuel production. New metabolic engineering tools and techniques are currently being developed for engineering untraditional hosts like eukaryotic algae and cyanobacteria, and as our understanding of these new hosts matures, significant improvement in hydrocarbon yields is anticipated.

Abbreviations

Acknowledgements

This work was supported by the Harry S. Truman Fellowship in National Security Science and Engineering and the Laboratory Directed Research and Development program at Sandia National Laboratories. Sandia National Laboratories is a multi-program laboratory managed and operated by Sandia Corporation, a wholly owned subsidiary of Lockheed Martin Corpo­ration, for the U. S. Department of Energy’s National Nuclear Security Administration under contract DE-AC04-94AL85000.

Author details

Anne M. Ruffing

Sandia National Laboratories, Department of Bioenergy and Defense Technologies, Albu­querque, NM, USA