Microalgal biodiesel production is relatively new and not very well explored. Some reports are available, where attempts have been made to produce biodiesel from algae (Table 11.3). Miao and Wu (2006) reported that lipid extracted from the heterotrophically grown microalga, C. protothecoides, transformed into biodiesel with a yield of 63% under 1:1 weight ratio of H2SO4 to oil, and 56:1 molar ratio of methanol to oil at 30 °C for a reaction time of 4 h. Xu et al. (2006) characterized the biodiesel obtained from the C. protothecoides oil by acid-catalyzed transesterification. The most abundant fatty acid methyl ester (FAME) in C. prothecoides bio­diesel was methyl oleate (61% of total FAME) followed by methyl linoleate (17%) and methyl palmitate (13%). Subsequently, Li et al. (2007) showed that it was feasible to grow C. protothecoides in a commercial-scale biore­actor. Using 75% immobilized lipase, these researchers claimed ~98% conversion could be obtained in 12 h when the reaction condition with respect to solvent type, water content and pH were optimized. Hossain and Salleh (2008) studied biodiesel production from Oedogonium and Spirogyra species using NaOH as cata­lyst. Algal oil and biodiesel production was higher in Oedogonium sp. than in Spirogyra sp. Umdu et al. (2009) studied the effects of Al2O3 supported CaO and MgO catalysts in the transesterification of lipid of N. oculata. These researchers found that pure CaO and MgO were not active, and CaO-Al2O3 catalyst showed the highest activity. Biodiesel yield was increased up to 98% from 23% under CaO-Al2O3 catalyzed reaction when methanol: lipid ratio was increased from 6:1 to 30:1.

Lipid extracted from N. oleoabundans was found to have an adequate fatty acid profile and iodine value according to the biodiesel specifications of European

Standards (EN, Gouveia et al. 2009). Converti et al.

(2009) analyzed the FAMEs in biodiesel produced from N. oculata and C. vulgaris. The most abundant composi­tion was methyl palmitate, which was 62% and 66%, respectively, in N. oculata and C. vulgaris biodiesel. However, the concentration of linolenic acid (18%) in N. oculata could not meet the requirement of European legislation for biodiesel. Johnson and Wen (2009) prepared biodiesel from the microalga Schizochytrium limacinum by direct transesterification of algal biomass. Parameters such as free glycerol, total glycerol, acid number, soap content, corrosiveness to copper, flash point and viscosity met the American Society for Testing and Materials (ASTM) and European standards, while the water and sediment content, as well as the sulfur content did not pass the standards. Damiani et al.

(2010) studied biodiesel production from H. pluvialis using potassium hydroxide as the catalyst. The major constituent of H. pluvialis biodiesel was palmitic acid fol­lowed by linoleic, oleic and linolenic acid methyl esters. The iodine value was within the limit established by European standards. Chinnasamy et al. (2010) produced biodiesel by a two-step transesterification process (acid-catalyzed followed by base-catalyzed) from a con­sortium of 15 native algae cultivated in carpet industry wastewater. Algal methyl esters were predominated by linolenic, linoleic, palmitic and oleic acids. The biodiesel was found to contain 0.0155% and 0.0001% bound and free glycerin, respectively, and met the ASTM and Euro­pean standard specifications.

Patil et al. (2012) optimized the direct conversion of wet Nannochlopsis sp. biomass to biodiesel under super­critical methanol treatment, without using any catalyst. In the supercritical state, at high pressure and tempera­ture, the methanol molecules enabled simultaneous extraction and transesterification of lipids in wet algal biomass. The abundant FAME in Nannochlopsis sp. biodiesel was methyl oleate (37%) followed by methyl palmitolate (32%) and methyl palmitate (8%). Velasquez-Orta et al. (2012) compared in situ transester­ification of C. vulgaris with acid as well as alkaline cat­alysts, in which the oil extraction step was eliminated. FAME yield reached a maximum of 77.6% after 45 min using a catalyst (NaOH) ratio of 0.15:1 and sol­vent ratio of 600:1 at 60 °C under constant stirring rate of 380 rpm. However, with sulfuric acid as catalyst FAME yield reached up to 96.9% with catalyst : oil ratio of 0.35:1 for a reaction time of 20 h. Recently, Mallick et al. (2012) characterized the biodiesel obtained from the C. vulgaris oil by acid-catalyzed transesterifica­tion. The fuel properties (density, viscosity, acid value, iodine value, calorific value, cetane index, ash and water contents) of C. vulgaris biodiesel are comparable with the international (ASTM and EN) and Indian standards (IS).

Slow Pyrolysis

Only a few percent of chlorine and potassium are car­ried in the pyrolysis gas and bio-oil at a pyrolysis temper­ature of less than 700 °C (Jensen et al., 1999). About 40% of the energy is at a temperature of 700 ° C in the gas frac­tion. Fifty percent of the energy remains in the biochar in slow pyrolysis at 400 °C. Total efficiency is 20% (Hal — wachs, 2010).

Flash Pyrolysis

In flash pyrolysis the production of liquid bio-oil is maximized using high temperatures and high heating rates. The liquid fraction of fast pyrolysis products con­tains a high concentration of potassium and calcium and its use in diesel engines and gas turbines is not prudent.

DISCUSSION

Maximum Methane Yield

Jerger et al. (1982) demonstrated that a correct ratio of macronutrients and minimum values of micronutrients offer a way to increase the methane yield at low costs. Tests on the biodegradability should be made with optimal concentrations of macronutrients and sufficient micronutrients. Some companies offer the service to test for these conditions in biogas plants. It would be wise for Indian cattle manure plants to formulate packages with the required macro — and micronutrients.

Komatsu et al. (2007) showed that primary sewage sludge is an excellent medium of both macro and micro­nutrients. Secondary sludge not only has methanogenic bacteria but also contains enzymes that break down straws, bagasse and husks. They used 65% VS sewage sludge and demonstrated that high concentrations of micronutrients are not harmful. The codigestion of ligno — cellulosic materials with sewage sludge is attractive. The proteins in the sludge are difficult to digest. Codigestion reduces the VS more than digestion the sludge and ligno — cellulosic materials separately (Rashed et al., 2008).

Longer retention times will increase the methane yield further given optimal nutrient conditions. This requires more or larger digester tanks. The tanks form only 20% of the total investment in a biogas installation and retention time can be increased fivefold when the methane yield is doubled for the same investment per cubic meter methane. The size for concrete tanks is limited to around 3000 m3 and multiple tanks are nor­mally required.

Much work has been done on mechanical, chemical or biological pretreatment of straws, bagasse and husks. These are effective giving up to 50% more methane. No comparison has been made with longer retention times in terms of investments and operational costs. Enzyme addition when making silage and boiling water addition during loading of straw and husks are the next most cost-effective treatments. This involves no extra handling of the substrates.

Nutrient Recycling

Nutrient recycling back to the soil is possible with anaerobic digestion. Most of the nitrogen and phosphate is in the liquid fraction of the digestate, when the diges — tate is separated in a solid and liquid fraction. The liquid fraction could be spread at nearby fields. Lignocellulosic biomass should be codigested with sewage sludge, where nutrient recycling is not economic.

Soil Fertility

Soil fertility is enhanced by humus. Humus is in turn produced by the degradation of lignin. The lignin remains in the digestate in anaerobic digestion and can be used as "fertilizer". A similar effect is obtained by the biochar from the pyrolysis of lignocellulosic biomass (terra preta). No comparison has been made on invest­ment costs and operational costs between pyrolysis and anaerobic digestion.

Digesters

Solid biomasses need to be digested at optimum nutrient conditions and long retention times for maximum methane yields. The wet scum layer digester is one option. The maximum solids concentration in the systems without giving operational difficulties has not been established.

Batch systems with leachate recycling are also an option. Substrate handling is minimized. The disadvan­tage is their small size and the danger of explosions during opening of the digesters.

CONCLUSIONS

Anaerobic digestion of food processing and crop res­idues can contribute to reduce the dependency on fossil fuels. Energy recovery in methane is 70% with digestion periods between 100 and 150 days, depending on the lignin content of the substrates and temperature of the digestion. Digestion times can be reduced with opti­mum concentrations of macro and micronutrients.

Food processing residues are a cheaper substrate for anaerobic digestion than crop residues, as collection has already been paid for. Systems using an auger or paddles to transport the substrate inside the digester tank have a significant higher methane yield than batch systems with leachate recycling. Their investment war­rants only their use for kitchen and garden wastes, for which a gate fee is paid. Losses in batch systems may be reduced by longer retention times. Storage cum digester tanks with leachate recycling will reduce sub­strate handling to a minimum.

Acid pretreatment involves the use of concentrated and diluted acids to break the rigid structure of the lignocellulosic material. The most commonly used acids are sulfuric (H2SO4) and hydrochloric (HCl). Dilute sul­furic acid has traditionally been used to manufacture furfural (van Putten et al., 2013a, b) by hydrolyzing the hemicellulose of mainly corncobs and bagasse into sim­ple sugars. The pentose part (e. g. xylose) is subsequently converted into furfural. Dilute sulfuric acid has also been used commercially to pretreat a wide variety of biomass types to subsequently convert both the C6 and C5 part. Feedstocks evaluated include switchgrass, corn stover, spruce, and poplar (Brodeur et al., 2011 and references therein). Other acids have also been stud­ied, such as phosphoric acid (H3PO4), nitric acid (HNO3) and organic acids (Brodeur et al., 2011; Kootstra et al., 2009). Due to its ability to remove hemicellulose, acid pretreatments have also been integrated in other pro­cesses in fractionating the components of lignocellulosic biomass such as the production of dissolving cellulose. Acid pretreatment (removal of hemicellulose) followed by alkali pretreatment (removal of lignin) results in rela­tively pure cellulose. This chemical pretreatment usually consists of the addition of concentrated or diluted acids (usually between 0.2% and 2.5% w/w) to the biomass, followed by constant mixing at temperatures between 130 °C and 210 °C. Depending on the conditions of the pretreatment, the hydrolysis of the sugars could take from a few minutes to hours (Brodeur et al., 2011). A key advantage of acid pretreatment is that a subsequent enzymatic hydrolysis step is sometimes not required, as the acid itself hydrolyzes the biomass to yield fermentable sugars. This is especially true with concen­trated acid treatments. Hemicellulose and lignin are partly solubilized with relatively minor degradation [75], and the hemicellulose is converted to monomeric and oligomeric sugars with acid pretreatment. A poten­tial drawback is the production of fermentation inhibi­tors like furfural and HMF, which reduce the effectiveness of the further processes. Therefore, exten­sive washing and/or a detoxification step is sometimes required to remove these inhibitors before a fermenta­tion step. Most acids have a strong corrosive nature asking for special reactor requirements (material for the reactor) in order to withstand the required experi­mental conditions and corrosiveness of the acids.

The optimum conditions for the acid pretreatment depend highly on the targeted sugars and the purpose of the pretreatment. Up to now most times subsequent conversions were based on fermentative processes. These processes require low amounts of inhibitors (furfural, HMF, organic acids) but are relatively tolerant to inorganic components. But also for fermentative pro­cesses this is not clear cut. It was found that the optimal conditions for obtaining the maximum sugar yield depends on whether the goal is to maximize the yield after the pretreatment or after the enzymatic hydrolysis of the pretreated solids or if the goal is to obtain maximum yield after both steps (Lloyd and Wyman,

2005) . Delmas (2008) studied the use of formic acid/ acetic acid pretreatment at 105 °C and atmospheric pres­sure to fractionate wheat straw into high purity fractions of organosolv cellulose, lignin and a sugar syrup. The raw straw pulp was separated from the dissolved lignin and hemicelluloses. The pulp can be bleached with hydrogen peroxide and the commercial value of the raw pulp is close to that of eucalyptus chemical pulp. A high-purity lignin, with linear structures, was recovered from the organic medium.

Methionine is a limiting amino acid in the monogas­tric’s feed. Its isolation was first reported from casein (Mueller, 1923) and since then methionine was commer­cially produced either by chemical synthesis or enzymatic methods. Unlike other amino acids, methionine has an advantage that it can be supplied to animal feed as a race­mate or a racemic mixture as the mammals are able to convert it to utilizable form with a methionine racemase enzyme. Even so, the microbial production has added ad­vantages over the racemate that it helps optimal nutrient utilization. With the discovery of fermentative production of amino acids by Kinoshita, attempts were being made to commercialize methionine production by submerged fermentation. The initial attempts on microbial fermenta­tion were done in the 1970s (Kase and Nakayama, 1974). As amino acid synthesis is an energy expensive process and is feedback inhibited, the wild-type bacteria were not reported to overproduce methionine. Hence, overpro­duction was achieved either by classical mutagenesis or deregulation of the biosynthetic pathway. Species of

Corynebacterium and Brevibacterium have much simpler regulatory mechanisms than E. coli for methionine biosyn­thesis and are the preferred microbes for overproduction. Corynebacterium is also able to switch between the transsulfuration pathway and the direct sulfhydrylation for methionine production (Hwang et al., 2002). Even though attempts are still being made to tailor the methio­nine producers to utilize raw sugars from complex sources, glucose and maltose are most extensively used. Substrates for fermentation varied over coconut water, banana, cassava, molasses, sugarcane juice, etc. (Pham et al., 1992). Methanol and n-alkanes were also used (Morinaga et al., 1982; Ghosh and Banerjee, 1986).

Threonine

Threonine is the third limiting amino acid after lysine and methionine in the animal feeds and was discovered by W. C. Rose (Rose, 1931). Presently, the major share of commercially produced threonine employs fermentative production. The microbial strains employed in the pro­duction process were genetically manipulated for threo­nine overproduction including Serratia marcescens (Komatsubara et al., 1978), C. glutamicum and E. coli (Dong et al., 2011). Escherichia coli dominates as the thre­onine producer due to their dynamic growth pattern and better substrate utilization, but cannot be used in the synthesis of pharmaceutical grade amino acid owing to their endotoxin production. The generally recognized as safe status, the highly defined genome database and scope for further genetic manipulations make C. glutami — cum dominate the pharmaceutical grade threonine pro­duction. Generally the strains employed in methionine fermentation utilize directly available monosaccharide sugars. The broadening of substrate utilization range to polysaccharides and biomass-derived sugars including pentoses is highly desired, as this helps in cost reduction and flexibility of the overall industrial production pro­cess. Most importantly it will help utilize the renewable sugar sources that will otherwise go underutilized. But this will require manipulations in the sugar uptake sys­tems of the microorganisms under concern, as both the industrial strains—E. coli and C. glutamicum—are unable to directly utilize polysaccharides. Escherichia coli strains are reported to have pentose utilization systems, but this capability has to be incorporated in the strains with amino acid production ability and resistance to biomass pretreatment-derived inhibitors. Thus, the extended sub­strate utilization spectrum will be a step toward produc­tion of threonine from biorefinery.

Light-Dependent Hydrogen Production Pathways

Like some cyanobacteria, many species of eukaryotic green algae are capable of direct water biophotolysis (Boichenko and Hoffmann, 1994). In green algae, water biophotolysis typically proceeds in two steps:

H2O + 2Fdox / 2H+ + 1 /2O2 + 2Fdred (21.5)

2H+ + 2Fdred / H2 + 2Fdox (21.6)

The first reaction is common to all oxygenic phototrophs, and was explained above (Section Oxygenic Photosyn­thesis). The second step, however, occurs only under anaerobic or microaerobic conditions, since it is extremely oxygen sensitive (Ghirardi et al., 1997). The reaction is catalyzed by the [Fe—Fe]-hydrogenase enzyme that accepts electrons from photosynthetically reduced Fd and reduces protons of water to molecular hydrogen (Happe and Naber, 1993; Happe and Kamin­ski, 2002; Foriestier et al., 2003). Although six different Fds are known in green algae, it is most likely that only PetF serves as the physiological electron donor to [Fe—Fe]-hydrogenase and, thus, links the photosynthetic electron-transport chain to the hydrogenase-driven reaction in vivo (Happe and Naber, 1993; Winkler et al., 2009; Long et al., 2008).

In photosynthetically active algal cells, the direct water biophotolysis process usually occurs when cul­tures are exposed to the light after a period of dark, anaerobic adaptation (Gaffron and Rubin, 1942; Green — baum, 1982; Appel and Schulz, 1998). It proceeds at very high initial rates (up to 300 mmol H2 mg/Chlh) that are comparable to the rates of O2 evolution in green algae under optimal light conditions (Boichenko and Hoffmann, 1994; Boichenko et al., 2004). It should be noted, however, that such high rates of H2 photoproduc­tion in healthy algal cells occur only for a very short period of time and that the duration of the process depends directly on the light intensity. For example, in dark-adapted Chlorella vulgaris cultures the H2 photo­production rate reaches the maximum in 2.5 s after exposing cells to about 0.03 W/m2 light and the kinetics is linear for at least 1 min (Boichenko et al., 1983). Under 2 W/m2 illumination, cells show the maximum rate after 0.6 s, which starts declining soon after 1 s. The pro­cess can also be extended in the presence of 3-(30,4′- dichlorophenyl)-1,1-dimethylurea (DCMU), a specific inhibitor of electron transport from PSII to the PQ-pool (Happe and Naber, 1993; Florin et al., 2001). In this case, however, H2 photoproduction is driven through the photofermentation pathway (see below) and thus depends on the level of stored carbohydrates and pro­teins. Nevertheless, the extension of the H2 photopro­duction period in DCMU-treated cultures indicates that inhibition of H2 evolution in dark-adapted and DCMU-untreated algae in light is mainly due to a fast accumulation of O2 inside algal chloroplasts. This inhibits hydrogenase-driven reaction and switches the physiological state of cells from anaerobic to aerobic. Indeed, using the Clark-type O2 and H2 sensors, Boichenko and et al. (1983) showed that the decrease in the rate of H2 photoproduction in algae is always followed by accumulation of O2 in the culture. The sensitivity to O2 occurs at four levels: (1) gene transcrip­tion, (2) [Fe—Fe]-hydrogenase maturation, (3) activity of the hydrogenase catalytic site, and (4) competition for photosynthetic reductants (Ghirardi, 2006).

Although H2 photoproduction in green algae is a short-term phenomenon, its theoretical sunlight to hydrogen conversion efficiency (STHE) is higher than in N2-fixing cyanobacteria. In addition, algae split water and, contrasting many anoxygenic photosynthetic bacte­ria, they do not require any organic substrates for pro­duction of H2 gas. The maximum efficiency for the direct water biophotolysis process has been estimated at around 10% (Bolton, 1996; Akkerman et al., 2002). Since this value is comparable to the power conversion efficiency of present commercial silicon solar cell mod­ules, which are rated at around 10—11% with water­splitting process considered (Blankenship et al., 2011), the high efficiency of algal H2 photoproduction raises the possibility of industrial application of the process in the future. At the current state, however, the direct water biophotolysis systems are not cost-effective and will require significant biochemical and engineering improvements to achieve commercial viability.

Taking into account a high sensitivity of algal hydrog — enases to O2, much attention in the past was focused on the development of the efficient methods of keeping cul­tures anaerobic throughout the H2 production period. These methods included the addition of O2 scavengers (Healey, 1970; Randt and Senger, 1985), the use of added reductants (Randt and Senger, 1985), sparging cultures with inert gases (e. g. argon or helium) (Greenbaum, 1982; Greenbaum et al., 2001) and the addition of PSII in­hibitors (Gfeller and Gibbs, 1984; Fouchard et al., 2005). Although some of these methods indeed prolong H2 production in algae, none of them resulted in bulk pro­duction of H2 gas. The expense of such approach also limits their application on larger scales. Many early research efforts also concentrated on screening for natu­rally better H2 producers. As a result, a few dozen algal species were tested for their ability to photoproduce H2 gas after the period of dark anaerobic adaptation. Many of them, but not all, were capable of direct water biophotolysis only for a very short period of time (Ben-Amotz et al., 1975; Boichenko and Hoffmann, 1994).

According to Eqns (21.5) and (21.6), direct water biophotolysis gives a maximum theoretical H2 to O2 (mol:mol) ratio of 2 to 1. Since both PSII and PSI are involved in the process, the minimum number of quanta required for generating one H2 molecule is equal to four. In many short-term experiments performed with dark — adapted algae, this value was above 4, but in some cases only 2—2.5 quanta were required (Greenbaum, 1988; Boichenko et al., 1989, 2004). The latter value indicates that H2 photoproduction in green algae may occur through a mechanism independent of water oxidation. The existence of this pathway was also confirmed by the experiments with inhibitors of the photosynthetic electron transport chain. It was found that DCMU, a PSII inhibitor (see above), does not completely block H2 photoproduction in algal cells, while 2,5-dibromo-

3- methyl-6-iso-propyl-p-benzoquinone, which blocks PQ oxidation by the Cyt b6f complex, inhibits the process almost completely (Ben-Amotz et al., 1975; Kosourov et al., 2003; Antal et al., 2009). In contrast to water biophotolysis, this pathway depends on the meta­bolic oxidation of organic substrates that are coupled to PSI and the [Fe—Fe]-hydrogenases through the PQ-pool (Stuart and Gaffron, 1972; Gfeller and Gibbs, 1984; Gibbs et al., 1986). According to Gibbs and coworkers, starch, acetate and proteins could be the main substrates for photofermentation in algae, which results in production of CO2 and H2 gases. For example, the full degradation of 1 mol (in glucose equivalents) starch will give up to 6 mol CO2 and up to 12 mol H2:

CbHi2O6 + 6H2O / 6CO2 + 12H2 (21.7)

In C. reinhardtii and other green algae, degradation of substrates through photofermentation is not complete. The major by-products are formate, acetate, ethanol, and, in some rare cases, lactate and glycerol (Gfeller and Gibbs, 1984; Kreuzberg, 1984; Zhang et al., 2002; Kosourov et al., 2003). According to a number of studies, the final composition varies significantly on the strain and the environmental condition. Since accumulation of organic substrates, mainly starch, also depends on PSII activity, the photofermentation pathway in green algae proceeds in two stages and, thus, can be considered as an indirect water biophotolysis. Starch accumulated through photosynthetic activity later undergoes degra­dation through glycolysis that yields pyruvate, ATP and reductants. In green algae, oxidation of reductants may occur via Nda2, a monomeric class-II type NAD(P) de­hydrogenase, which feeds electrons into the photosyn­thetic electron transport chain at the point of the PQ-pool (Jans et al., 2008; Desplats et al., 2008). From the PQ-pool, electrons follow to [Fe—Fe]-hydrogenase through PSI and Fd. Since both direct and indirect water biophotolysis pathways are linked to [Fe—Fe]- hydrogenase via PSI, it becomes clear that in algae they operate simultaneously most of the time. Nevertheless, their contribution into the overall H2 photoproduction process varies depending on the organism (Meuser et al., 2009) and the physiological state of algal cells (Laurinavichene et al., 2004).

In the indirect water biophotolysis process, O2 evolu­tion and H2 production stages can be separated from each other either temporally or spatially, thus circum­venting the apparently inherent O2 sensitivity of H2 photoproduction. Despite lower efficiency (as compared to the direct water splitting), separation of the process into two distinct stages gives a significant advantage to the biotechnological applications (Benemann, 1994, 1996). According to the early Benemann’s concept, during the first aerobic stage algal cultures accumulate biomass enriched in carbohydrates as a result of photo­synthetic CO2 fixation. During the second, anaerobic stage carbohydrates and other materials stored in biomass are processed to H2 gas. The two stages are sepa­rated either physically in two different photobioreactors or temporally through additional dark adaptation and fermentation periods. In this approach, O2-evolving activity inside the cells is totally inactivated during the H2 production stage without application of any inhibi­tors, and algae evolve H2 gas through the photofermenta­tion pathway linked to PSI. The second stage can also be driven in the dark by fermentative bacteria. Thus, the discovery of the indirect water biophotolysis pathway was the first step toward development of the protocol for long-term H2 photoproduction in green algae.

The major components of lignocellulosic feedstocks are cellulose and hemicellulose that can be converted to sugars through a series of thermochemical and bio­logical processes and eventually fermented to bio­ethanol, other solvent biofuel or biogas. Therefore, lignocellulosic feedstocks are mainly categorized as agricultural residues (e. g. crop residues and sugarcane bagasse), forest residues, herbaceous and woody energy
crops. There are three principal technological end points for bioenergy crops: (1) conversion to liquid fuels, (2) combustion (alone or in combination with fossil fuels) to produce heat, steam, or electricity, and (3) gasification to simpler gaseous products that can have various uses. Agricultural residues can differ significantly in their chemical composition, which can lead to different bio­energy and biofuel yields per unit feedstock (Carriquiry et al., 2010). Commonly available agricultural residues for bioenergy production are barley straw, corn stover, rice straw, sorghum straw, wheat straw and sugarcane bagasse and having available carbohydrate contents of 70%, 58.3%, 49.3%, 61%, 54% and 67.2%, respectively. If the available carbohydrate of these feedstocks is fully converted to bioenergy, they can potentially yield approximately 367, 503, 392, 199, 1413 and 3133 liters biofuel/hectare, respectively (Carriquiry et al., 2010; US-DOE, 2008a, b; Kim and Dale, 2004). Forest residues include logging residues produced from harvest opera­tions, fuel wood extracted from forestlands, and pri­mary and secondary wood-processing mill residues (Perlack et al., 2005). Some important forest residues include hardwoods (black locust and hybrid poplar), softwoods (eucalyptus and pine) and switchgrass, which comprise approximately 57.15—66.45% carbohy­drate, depending on the residue (Menon and Rao, 2012; van Dyck and Pletschke, 2012; Alves et al., 2010; Carriquiry et al., 2010; US-DOE, 2008a, b; Merino and Cherry, 2007; Hamelinck et al., 2005 Howard et al.,

2003) . However, several factors restrict the potential use of forest residues for biofuel production (Perlack et al., 2005). The first factor is the economic cost of transportation as limited accessibility largely increases the operational costs of logging/collection activities. Another factor is a potential reduction in recoverability of harvest areas due to environmental considerations (Richardson, 2008). Therefore a shift in research efforts to dedicated biofuel crops is taking place in order to have a continuous supply of feedstock for second — generation bioenergy requirements.

Strategies for Microorganism Application in Biomass

Most naturally occurring microorganisms cannot uti­lize untreated lignocellulose efficiently for the produc­tion of biofuel or bioproducts due to the inaccessibility of the carbohydrate polymers, even though many of them secrete a variety of hydrolytic enzymes. For effi­cient utilization, biomass must first be pretreated to open up the cell wall and then hydrolyzed by acidic or enzymatic processes to fermentable sugar monomers. In addition to monomeric sugars, the pretreatment and acidic hydrolysis processes may also produce low mo­lecular weight organic acids like acetic acid, furfural, hydroxymethylfurfural and various lignin-degradation products that are potent inhibitors of microbial meta­bolism (Larsson et al., 1999; Palmqvist and Hahn — Hagerdal, 2000).

For an economically viable manufacturing process from lignocellulosic biomass, both hexose and pentose sugars produced during hydrolysis of both cellulose and hemicelluloses need to be utilized efficiently. In the course of cellulosic biomass conversion into biofuels and bioproducts, four biologically mediated processes are involved: (1) saccharolytic enzyme production, (2) enzymatic hydrolysis of biomass, (3) fermentation of hex — ose sugars, and (4) fermentation of pentose sugars (Lynd et al., 2005,2002). For an industrially viable process, each of the four steps must be rapid and efficient. As suggested by a recent calculation, an economically competitive fermentation process for industrial application needs to approach an anaerobic yield of ~ 95% of the theoretical yield, produce around 100 g/l of end product with a pro­ductivity of more than 2 g/l/h (Sheridan, 2009).

Flavins

In addition to the mediator-less form of electrogenesis discussed above (TFP and soluble/insoluble cyto­chromes), S. oneidensis is an electrogenic bacterium that can also secrete extracellular, soluble, and redox-active mediators. Mediators can accept electrons in the anaer­obic extracellular milieu or directly from the bacterial cell surface, and, due to their lower redox potential (Eo0 in V), they donate electrons directly to the anodic surface. One group of the S. oneidensis mediators is fla­vins. In addition to pili and cytochromes, S. oneidensis produces extracellular flavins that contribute to the elec­trogenic process and can actually be reduced, in part, via the Mtr/Omc system. Such flavins include riboflavin (vitamin B2) (Figure 9.10), flavin (isoalloxazine from which flavins are derived), and flavin mononuclotide (FMN). Thus, cells that are unbound to the metal oxide surface are still capable of reducing it, although the reduction process also requires reduced members of MtrC and OmcA (Figure 9.7).

FIGURE 9.8 DET and MET electron transfer pathways utilized by S. oneidensis and selected mutant strains. (a) Electron transfer via the cytochrome pool. Transmembrane pilus electron transfer via (b) pil-type pilus and via (c) msh-type pilus, and (d) biofilm formation behaviour. OM: Outer membrane and IM: Inner membrane. Source: Figure from Carmona et al. (2011) with permission from Elsevier

Phenazines

Phenazines are tricyclic, redox-active compounds that are produced by a number of species of the genus Pseudo­monas. Pseudomonas aeruginosa, depending upon the mutations acquired in a specific microniche, can produce, or in the case of mutations within negative regulators or modulators such as RpoS, actually overproduce redox-active 1-hydroxyphenazine, pyorubrin, or pyo — cyanin (Figure 9.11). The process of phenazine biosyn­thesis in these organisms was highlighted by Mavrodi et al. (2001) where the entire pathway is based upon anthranilate synthesis and genes beginning with the acronym phn (for phenazine). A classic demonstration of the electrogenic contribution of P. aeruginosa phena — zines to the electrogenic properties of this organism was shown using the power of classical bacterial genetics. Using a mutant approach, Rabaey et al. (2005a) showed that pyocyanin, pyorubrin, and 1-hydroxyphenazine could act as excellent mediators in MFCs. Bacteria that could not produce these mediators possessed reduced
electrogenic properties relative to those genetically inca­pable of producing them, or organisms whose media were amended with known quantities of each phenazine. Given the known electrochemical potential of each of the aforementioned mediators, it is not surprising that power density was greatest in MFCs containing pyocya — nin > pyorubrin (aeruginosin A; Rabaey et al., 2005a) > 1-hydroxyphenazine. Supportive of these results were those of Luo et al. (2009b) in the isolation of strain RE7.

Independent of the biomass source, biolipids can be used in various ways as a source of bioenergy. There are a number of basic steps involved in processing bio­lipids to biofuel. These can include some or all of lipid extraction, degumming, neutralization, winterization, bleaching and transesterification. The sources of biomass and how they are produced have been described previously in this chapter and the first pro­cessing step will usually involve efficient extraction of the biolipid from the biomass. Following extraction, some biolipids can be used in their pure form as pure plant oils (PPOs). Other biolipids are further processed, usually into biodiesel. Here the extraction step is followed by purification and stabilization of the biolipid and the conversion to biodiesel. The various steps involved in processing biolipids are described below, beginning with extraction, along with the fuel properties of both PPO and biodiesel.

Extraction

Extraction is a process consisting of the separation of a specific substance from a complex matrix. In the context of extraction lipids from biomass, the purpose is to use standardized extraction procedures to isolate the biomolecules of interest, i. e. lipids, concurrent to rejection of the remaining inert biomass. This is most commonly achieved by using a selective solvent known as menstruum (Handa, 2008), or by solventless physical extraction means. The resultant lipid may be ready for use in the form of fluid extracts, it may be further pro­cessed into a variety of biofuel and nutraceutical prod­ucts, or it may be fractionated to isolate individual chemical entities or a combination of the above as proposed by the "biorefinery" concept discussed previ­ously. The most common biolipid extraction procedures are summarized below.

Steam Distillation

Steam distillation is a process that is commonly applied to the extraction of essential oils (Gutierrez et al., 2009). Plant material is placed into a still where pressurized steam penetrates the plant material causing internal lipid vacuoles to rupture. Upon exposure to the surrounding environment, the lipid evaporates to form a mixture of easily separable vapors (essential oil and water). The vapors condense and the distillate (sepa­rated into two immiscible layers) is collected in a gradu­ated tube connected to a condenser. The aqueous phase is recirculated into the flask, while the volatile oil is collected separately. The main disadvantage associated with steam distillation is that thermolabile components risk being degraded (Sarker et al., 2005). A combination of solvents and steam distillation is often used to improve the final product of a biodiesel production process.

Maceration (Solvent Extraction)

Maceration is used for creating extracts and resins in a simple yet well-established procedure. Whole or coarsely powdered biomass is placed in intimate contact with a suitable extractant in a closed vessel. The mixture is allowed to stand at room temperature for a defined period of time, typically at least 3 days, with frequent agitation (using mechanical shakers or mixers) to ensure homogeneity (Sarker et al., 2005). The organic phase is separated from the solids by either filtration, decanta­tion or in some cases centrifugation and the remaining solid material is pressed to ensure efficient solvent recovery. The recovered liquid phases are combined and clarified for further processing. This process can be repeated several times to achieve maximum lipid recovery. The main disadvantage associated with macer­ation is that the process can be quite onerous, potentially taking from a few days up to several weeks (Takahashi et al., 2001).

The heteropolyacid (HPA) solids, related to the class of polyoxometalates, are often remembered when there is a need for catalysts that are tolerant to the large amounts of water. As already discussed above, such conditions are usually found in the catalytic conversion of low-cost raw materials to liquid biofuels such as biodiesel.

Besides their inherent superacidity (pKH+ > 12) (Mizuno and Misono, 1998), which already ensures the achievement of relatively high yields, these compounds can be devised in such a way to produce a pore architec­ture and a chemical composition that meets the structure and size of the molecules that are involved in both ester­ification and transesterification reactions.

Polyoxometalates, frequently named as POMs, are anionic metal-oxo clusters whose chemical properties can be modulated by the presence of one or more different transition metal ions and the cation used to generate the salt form.

The presence of two different metal atoms per poly — oxometalate molecule generates compounds with mixed metals, vanadium and molybdenum being the most commonly used. Furthermore, the presence of other atoms in the structure, besides the metal and the oxygen atoms, leads to heteropolyoxometalate compounds with the general formula (X”+Mo12O40)(8~”)~, where X can often be as W (V), Si (IV), Ge (IV) and Ti (IV). These an­ions can be arranged in typical structures such as Keggin and Dawson structures.

The protonated form of heteropolyoxometalate an­ions is referred to as heteropolyacids, which may be defined as a condensed structure of different types of oxyacids. In water, it is expected that all HPA protons are dissociated. The strength of these acids in acetoni­trile was estimated. For instance, the acidity of H3PW12O40 in acetonitrile is greater than that observed for the p-toluenesulfonic acid and H2SO4, two acids usu­ally used as catalysts for (trans)esterification in homoge­neous catalytic systems (Drago et al., 1997).

HPAs are generally soluble in water and other polar media. Therefore, they are usually unsuitable for bio­diesel production by heterogeneous catalysis. However, these anionic compounds are water insoluble when pre­sented as salts with large cations such as Cs+. Because of this characteristic, there is a great interest in the applica­tion of this family of solid compounds in heterogeneous process, acting as acid, redox and bifunctional catalysts (Li et al., 2007).

The (C16TA)H4TiPWnO40 solid, resulting from the combination of a surfactant (C16TA, cetyltrimethylam — monium) with an HPA, was recently reported as a water-tolerant solid for the heterogeneous catalytic esterification of palmitic acid (Zhao et al., 2012). The observed high conversion (94.7 wt%) and high efficiency (91.8 wt% yield) were attributed to the presence of Bronsted and Lewis acid sites, its amphiphilic property and high water tolerance. The authors claimed that sub­strate molecules concentrate around the catalyst through hydrophobic interactions with its lipophilic tail while methanol molecules are absorbed by HPA through hydrogen bonding. The hydrophobic surroundings also promote the separation and/or repulsion of water

molecules from the surface of the catalyst. Also, the solid catalyst showed a good recyclability and its heteroge­neous character was proved through several cycles of re­covery and reuse.

In order to heterogenize the HPAs and improve their recovery and reuse, their impregnation on zirconia was also investigated (Oliveira et al., 2010). The H3PW12O40 was immobilized on zirconia at different ratios and calcined at 200 °C for 4 h. No decomposition of the Keg — gin anion structure was observed under these conditions. The resulting solids were used in the esterification of oleic acid with ethanol at a 20 wt% loading, 100 °C and 4 h with an ethanol:acid molar ratio of 6:1, conditions under which an 88 wt% conversion of oleic acid to ethyl oleate was obtained. A small leaching (8 wt%) of the catalyst was observed at the end of the reaction, therefore affecting the reaction kinetics. The recyclability study indicated that, after being recovered, washed and ther­mally treated, the solid presented conversion values as high as 70 wt%, that is, 80% of the original value of 88 wt%.

These examples and many others have shown that HPA and POWs represent promising catalysts for both esterification and transesterification (Giri et al., 2005; Caetano et al., 2008; Wee et al., 2010; Leng et al., 2009) but their heterogenization in different supports still needs to be improved in order to keep the process totally heterogeneous.