The great influence of this gaseous hormone on different plant organs made it an interesting target for pathogens. Indeed, the mold Penicillium digitatum has been known to produce ethylene since the mid-1950s (Wang et al., 1962) and its production was shown for prokaryotic plant pathogens in the early 1960s (Freebairn and Buddenhagen, 1964). Not surprisingly, the pathways found in these microorganisms are not analogous to the one in plants. So far, two distinct routes for ethylene production have been described in microbes: a 2-oxoglutarate-dependent pathway and the 2-keto-

4- methyl-thiobutyric acid (KMBA) pathway (Nagahama et al., 1991,1992). The latter is the most common among microorganisms, composed of a series of chemical and enzymatic reactions, by which only trace amounts of ethylene are usually produced (Ogawa et al., 1990). The former pathway has been found to be more efficient, with 2-oxoglutarate being used as substrate in a single­step reaction by the ethylene-forming enzyme (EFE). This pathway has been found in several different microor­ganisms, including P. digitatum, Chaetomium globosum, Phycomyces nitens, Fusarium oxysporum, and in different pathovars of Pseudomonas syringae, where a comparison study found the pv. phaseolicola to be the most efficient ethylene-producing strain (Weingart et al., 1999). This enzyme catalyzes simultaneously two reactions (Fukuda et al., 1992b):

2 — oxoglutarate 4 ethylene + 3CO2 + H2O (22.2)

2 — oxoglutarate + L — arginine + O2 4 succinate +CO2 + guanidine +(S) — 1 — pyrroline — 5 —carboxylate + H2O (22.3)

These reactions are rather interesting as they keep the tricarboxylic acid (TCA) cycle closed through a shortcut, converting 2-oxoglutarate directly into succinate with the formation of ethylene "as a by-product" (Figure 22.2), and therefore substituting for the steps catalyzed by 2-oxoglutarate dehydrogenase and succinyl-CoA synthe­tase (Figure 22.2). The original two-step reaction between 2-oxoglutarate and succinate generates one NADH and one guanosine triphosphate, which are not produced by EFE. Thus, competition of the two pathways for substrate 2-oxoglutarate would lower the formation of NADH. Since NADH also has a role as an inhibitor for four enzymes associated with the TCA cycle, pyruvate dehy­drogenase, isocitrate dehydrogenase, 2-oxoglutarate de­hydrogenase, and citrate synthase (Figure 22.2), this could potentially upregulate the reactions performed by these enzymes, from the decarboxylation of pyruvate to 2-oxoglutarate. The last is a direct and indirect substrate to the EFE, directly to generate ethylene (Eqn (22.2)) and indirectly, as it is also a substrate for the synthesis of argi­nine, required for the simultaneous reaction of this enzyme (Eqn (22.3) and Figure 22.2).

Physical methods involve breakdown of biomass size by coarse size reduction, chipping, shredding, grinding, and milling in order to increase the available specific surface area and reduce the degree of polymerization, enhancing the digestibility of lignocellulosic biomass (Agbor et al., 2011; Brodeur et al., 2011). However, it has been shown that further reduction of biomass parti­cle size below 0.4 mm has little effect on the rates and yields of biomass hydrolysis (Agbor et al., 2011).

Chipping reduces heat and mass transfer limitations; grinding and milling are more effective at reducing the particle size and cellulose crystallinity than chipping, probably as result of the shear forces generated during milling. Vibratory ball milling has been used with more effective results in reducing cellulose crystallinity than ordinary ball milling. Also, disk milling, which pro­duces fibers, has been reported as more efficient in enhancing cellulose hydrolysis than hammer milling, which produces finer bundles (Agbor et al., 2011). Stir­ring ball milling also could significantly damage the structure of biomass, resulting in the variation of surface morphology, the increase in amorphous region ratio and hydrogen bond energy, and the decrease in crystallinity and crystalline size (Liao et al., 2011).

The energy requirements for physical pretreatments are dependent on the biomass characteristics, final par­ticle size and reduction in crystallinity, for example, hardwoods require more energy input than agricultural residues (Agbor et al., 2011). Taking into account the high-energy requirement of milling on an industrial scale and the rise in energy demands, this method is not economically feasible and likely will not be used in a full-scale process (Agbor et al., 2011; Saritha and Lata, 2011). In most cases where the only option avail­able for pretreatment is physical, the required energy is higher than the theoretical energy content available in the biomass (Brodeur et al., 2011).

The pretreated biomasses by physical methods are subjected to heating, mixing and shearing resulting in physical and chemical modifications (Karunanithy and Muthukumarappan, 2011; Lamsal et al., 2010; Saritha and Lata, 2011). Also, Agbor et al., in 2011, suggested that the materials can be milled after chemical pretreat­ment with significantly reduction of (1) milling energy consumption, (2) reduce cost of solid liquid separation because the pretreated chips can be easily separated,

(3) eliminate energy-intensive mixing of pretreatment slurries, (4) liquid to solid ratio and (5) did not result in the production of fermentation inhibitors.

Another physical method is extrusion that disrupts the lignocellulose structure and increases the accessi­bility of carbohydrates to enzyme attack. This method has reported the improvement on sugar recovery up to 63.5% (Karunanithy and Muthukumarappan, 2011). Other physical pretreatments involve the use of gamma rays that cleave the b-1,4 glycosidic bonds, thus giving a larger surface area and lower crystallinity. This method will undoubtedly be very expensive on a large scale with huge environmental and safety concerns (Agbor et al., 2011). Proton beam irradiation has also been tested reporting a glucose conversion of 68% of the theoretical maximum at 72 h (Kim et al., 2011b).

The effect of microwave and microwave-chemical pre­treatments on densification characteristics and physical quality of pellets has also been investigated showing that microwave pretreatment was significantly able to disintegrate the lignocellulosic structure of wheat and barley straw grinds (Kashaninejad and Tabil, 2011). These pretreatments also have been tested on barley husks, sweet sorghum bagasse, bamboo, coconut husk and gar­den biomass (Choudhary et al., 2012; Ding et al., 2012; Gabhane et al., 2011; Jackowiak et al., 2011; Janker — Obermeier et al., 2012; Roos et al., 2009; Wu et al., 2012). However, some restrictions on energy consumption must be accomplished to obtain a positive energy balance with these pretreatments. Ultrasounds and ultrasound — assisted alkaline pretreatment also has been reported (Sun et al., 2002; Velmurugan and Muthukumar, 2012b). On the other hand, some continuous system includes a wet disk milling of rice straw (Hideno et al., 2012) and pulsed electric field of wood chip and switchgrass (Kumar et al., 2011).

In an effort to metabolically engineer microorganisms for efficient isobutanol production, various researchers sought to understand isobutanol production at both the molecular and protein levels. A few years before 2013, a number of studies directed at finding molecular and biochemical bases for isobutanol synthesis have been conducted (Table 7.2). Using a prokaryote (E. coli) as host, Atsumi et al. (2008) demonstrated that KIV can serve as a precursor for efficient isobutanol produc­tion from glucose in a biosynthetic pathway consisting of 2-KDC from L. lactis and ADH2 from S. cerevisiae with broad-range substrate specificity in combination with the expression of the alsS gene (encoding AHAS (acetohydroxy acid synthase)) from B. subtilis and the ilvCD gene from E. coli. A follow-up study on the role of different ADHs on isobutanol production with E. coli showed that chromosomally encoded YqhD is the major isobutyraldehyde-converting enzyme, and ADH2 from S. cerevisiae contributes only to a minor extent to isobutanol production in E. coli (Atsumi et al., 2010). This supposition was made after yqhD gene was deleted from the genome of E. coli; the generated recombinant E. coli strain (yqhD deficient) accumulated isobutyraldehyde during fermen­tation and experienced 80% reduction in isobutanol production. Using an a eukaryote (S. cerevisiae) and work­ing on the assumption that inefficient production of isobu­tanol from glucose may be due to limited supply of KIV, a precursor for the valine biosynthesis pathway (Figure 7.2), Lee et al. (2012) added exogenous KIV (0.5 g/l) into the growth medium. As expected, supplementation of the medium with KIV improved the production of isobutanol, which suggests that the endogenous pathway for produc­ing KIV in S. cerevisiae (and potentially, other producing microorganisms) is the limiting step in the isobutanol pro­duction pathway. Consequently, this finding made KIV biosynthesis a rational target for metabolic engineering toward designing more robust isobutanol-producing strains.

Development of hyper-isobutanol-producing strains has typically followed one of the three approaches: (1) identification of rate-limiting steps in Ehrlich/2-ketoacid biosynthetic pathways and overexpression of KIV biosyn­thetic genes; (2) improvement of heterologous expression of enzymes of the isobutanol pathway by codon optimiza­tion; and (3) removal of feedback inhibition and deletion of other competitive pathways, especially competition for pyruvate. Pursuant to the first strategy, Lee et al. (2012) screened and identified a 2-KDC exhibiting a relatively higher activity on KIV through in vitro activity assays of KDC using crude extracts of transformants overexpress­ing KDCs from various microorganisms. The highest KDC activity with KIV was observed from the transform­ant expressing kivd from L. lactis subsp. lactis KACC13877. Subsequently, Chen et al. (2011) evaluated the effect of overexpressing the genes, ILV2, ILV3, ILV5, ILV6, and BAT2, involved in valine metabolism, in different combi­nations in S. cerevisiae, on isobutanol production. Following cultivation of the ILV2, ILV3, and ILV5 overex­pressing strain (ILV235_XCY561) and the reference strain (CEN. PK113-5D) in mineral glucose medium supple­mented with uracil in fermentors under anaerobic condi­tions, the recombinant strain ILV235_XCY561 produced

0. 97 ± 0.14 mg isobutanol/g glucose, which was sixfold higher than the control strain (Chen et al., 2011), hence attesting to the fact that overexpression of the genes ILV2, ILV3, and ILV5 may have led to a higher concentra­tion of KIV, which resulted in higher isobutanol produc­tion. In parallel, Atsumi et al. (2009) engineered a cyanobacterium, Synechococcus elongatus, by expressing a KDC gene (kivd) from L. lactis in this cyanobacterium using an expression cassette under the control of the iso- propyl-b-D-thiogalactoside-inducible promoter Ptrc and integration into neutral site I (Bustos and Golden, 1992) by homologous recombination (Golden et al., 1987), and strain SA578 was generated. To improve flux toward KIV, alsS gene from B. subtilis and the ilvC and ilvD genes from E. coli were integrated into neutral site II (Andersson et al., 2000) in the genome of strain SA578 to generate strain SA590, which produced high levels of isobutyraldehyde upon cultivation in a Roux culture bottle at 30 °C (Atsumi et al., 2009, Figure 7.2). Given the fact that isobutyralde — hyde can easily undergo a reduction reaction to produce isobutanol, Atsumi et al. (2009) evaluated feasibility of using ADHs (ADH2 from S. cerevisiae, YqhD from E. coli, and AdhA from L. lactis) and Kivd from L. lactis (strain SA590) to achieve this reduction reaction. These genes (ADH2, YqhD, and AdhA) were integrated downstream of kivd individually, hence, strains SA413, SA561 and SA562 were generated, respectively. Whereas YqhD, an NADPH-dependent enzyme, was the most active one in S. elongatus, AdhA and ADH2 were nicotinamide adenine

dinucleotide-dependent, and generated strain SA579, which produced 450 mg/l isobutanol.

The second strategy derives from differences in codon bias among different microorganisms, based on their individual transfer RNA content and requisite expression levels of specific proteins in each microor­ganism (Ikemura, 1985; Percudani et al., 1997). Given the fact that codons at the beginning of an open reading frame play a critical role in protein expression (Vervoort et al., 2000), codon bias influences heterologous expres­sion of foreign proteins to a great extent. For instance, heterologously expressed protein levels in E. coli (Atsumi et al., 2010) and S. cerevisiae (Brat and Boles,

2013) were improved by codon optimization, especially at the 50 end of the coding sequence.

To realize the full potential of heterologously overex­pressed genes in producing microorganisms with respect to efficient isobutanol production, six genes including adhE (ADH), ldhA (lactate dehydrogenase), frd (fumarate reductase), fnr (encodes redox-sensing transcription regulator, which partakes in the regulation of lactate synthesis), pta (phosphate acetyltransferase), and pflB (pyruvate formate lyase) that are involved in byproduct formation in E. coli were deleted following overexpres­sion of AlsS (B. subtilis), IlvC (E. coli), and IlvD (E. coli) (Atsumi et al., 2008). These deletions may have increased the level of pyruvate available for the valine biosynthesis pathway. As a consequence, the isobutanol strain (JCL260/pSA55/pSA69) produced more than 22g/l in 112 h (Atsumi et al., 2008). In a similar study, Kondo
et al. (2012) overexpressed 2-KDC and ADH in S. cerevi- siae to enhance the endogenous activity of the Ehrlich pathway followed by overexpression of Ilv2, which cata­lyzes the first step in the valine synthetic pathway and deletion of the PDC1 gene encoding a major PDC with the intent of reducing ethanol flux via pyruvate. As a result, S. cerevisiae YTD306 was generated. Upon cultiva­tion of S. cerevisiae YTD306 along with modification of culture conditions, a 13-fold increase in isobutanol titer was produced (from 11 mg/l to 143 mg/l) when compared with the control (Table 7.2, Kondo et al., 2012).

The strategy described here, in which amino acid bio­synthetic and 2-ketoacid degradation pathways were exploited for isobutanol production, represents a new paradigm for biofuel production. Indeed, this paradigm employs non-CoA-mediated chemistry and uses only pyruvate as a precursor, unlike ethanol and butanol production by native alcohols producing microorgan­isms that are CoA-dependent (Ezeji et al., 2010; Atsumi et al., 2008).

Hydrogen is seen as one of the most promising fuels for the future owing to the fact that it is renewable and liberates large amounts of energy per unit weight without evolving CO2 when combusted. Biohydrogen production has several advantages over hydrogen pro­duction by photoelectrochemical or thermochemical processes. For example, whereas electrochemical hydrogen production requires the use of solar batteries with high energy requirements to split water and form the hydrogen product, biohydrogen production by photosynthetic microorganisms only requires simple PBRs with low energy requirements. A select group of green algae (including Chlamydomonas reinhardtii) and cyanobacteria offer an alternative route to renewable H2 production (Levin et al., 2004; Sakurai and Masukawa,

2007) . Cyanobacteria are able to diverge the electrons emerging from the two primary reactions of oxygenic photosynthesis directly into the production of H2, making them attractive for the production of renewable H2 from solar energy and water.

Cyanobacteria utilize two enzymatic pathways for H2 production, either nitrogenases or bidirectional hydroge — nases (Angermayr et al., 2009). Nitrogenases require ATP, whereas bidirectional hydrogenases do not require ATP for H2 production, hence making them more efficient and favorable for H2 production with a much higher turnover. The fundamental aspects of cyanobacterial hy — drogenases, and their more applied potential use as future producers of renewable H2 from sun and water, are receiving increased international attention. At the same time, significant progress is being made in the un­derstanding of the molecular regulation of the genes encoding both the enzymes and the accessory proteins needed for the correct assembly of an active hydrogenase. With the increasing interest of both scientific and public communities in clean and renewable energy sources, and consequent funding opportunities, rapid progress will likely be made in the fundamental understanding of the regulation of cyanobacterial hydrogenases at both genetic and proteomic levels. Bandyopadhyay et al.

(2010) have described Cyanothece sp. ATCC 51142, a uni­cellular, diazotrophic cyanobacterium with capacity to generate high levels of hydrogen under aerobic condi­tions. Wild-type Cyanothece sp. 51142 can produce hydrogen at rates as high as 465 mmol/mg of chloro — phyll/h in the presence of glycerol. Authors also report that hydrogen production in this strain is mediated by an efficient nitrogenase system, which can be manipu­lated to convert solar energy into hydrogen at rates that are several fold higher, compared to other previously described wild-type hydrogen-producing photosynthetic microbes. These strains have evolved the ability to use so­lar energy to produce H2 from water (Esquivel, 2011; Levin et al., 1961). The theoretical conversion efficiency from light to H2 is calculated to be as high as ~10% (Levin et al., 1961).

Photosystem II (PSII) drives the first stage of the pro­cess (Figure 10.10), by splitting H2O into protons (H2), electrons (e~), and O2.

H2 Production

H2O/2H++ 2e~ + У2 O2
2H+ + 2e~ /H2

H2 Combustion

H2 + У2 O2 / H2O + 285.8 kJ/mol

Normally, the photosynthetic light reactions and the Cal­vin cycle produce carbohydrates that fuel mitochondrial respiration and cell growth. Under anaerobic conditions, however, mitochondrial oxidative phosphorylation is largely inhibited, which leads some organisms (e. g. Chla- mydomonas reinhardtii) to reroute the energy stored in

carbohydrates to a chloroplast hydrogenase (HydA), likely using an NAD(P)H~PQ e~ transfer mechanism, to facilitate ATP production via photophosphorylation. Thus, hydrogenase reacts with H+ (from the medium) and e~ (from reduced ferredoxin) to produce H2 gas that is subsequently excreted from the cell. The combus­tion of the recovered H2 yields only heat and H2O and thus is a model green technology.

Several renewable energy laboratories have concluded that production efficiencies must be improved from 0.2% photon to H2 conversion efficiency at 20 W/m2 illumination to ~ 7—10% at 230 W/m2 illumination (day light) to make the process economically viable. Through extensive preliminary work, the efficiency of this process has been enhanced to ~1.0% from light to H2 and 2% to biomass. The H2 gas produced in such mutants has a purity of ~90—95% and typical yields are 500 ml H2 for a 11 culture (10 days; 110 W illumination). Without further purification, the H2 gas can used to power a small-scale fuel cell car.

In addition to work with Chlamydomonas, a large number of unicellular, filamentous, freshwater, and ma­rine cyanobacterial species have been reported to pro­duce large quantities of biohydrogen. Among other species, Anabaena azollae, Anabaena cylindrica, Anabaena variabilis, Arthrospira (Spirulina) platensis, Cyanothece, Gloeocapsa alpicola, and Nostoc muscorum have been re­ported to produce high levels of hydrogen gas (Jeffries et al., 1978; Aoyama et al., 1997; Antal and Lindblad, 2005). In particular, Anabaena sp. is reported to produce relatively large quantities of biohydrogen. Among these species, nitrogen-starved A. cylindrica cells produce the highest concentration of biohydrogen (30 ml H2/l/h) (Margheri et al., 1990).

These cyanobacterial strains use two sets of enzymes to generate hydrogen gas. The first enzyme is nitroge — nase, and it is found in the heterocysts of filamentous cyanobacteria when grown under nitrogen-limiting con­ditions. Hydrogen is produced as a by-product of fixa­tion of nitrogen into ammonia. The reaction consumes 16 ATP for fixation of 1 mol of N2, and results in forma­tion of 1 mol of H2. The other hydrogen-metabolizing or hydrogen-producing enzymes in cyanobacteria are hy- drogenases, which occur as two distinct types in different cyanobacterial species. The first type is uptake hydrogenase (encoded by hupSL), which has the ability to oxidize hydrogen via oxyhydrogenation or the Knall — gas reaction. The other type of hydrogenase is reversible or bidirectional hydrogenase (encoded by hoxFUYH), and it is capable of uptake and production of hydrogen (Schmitz et al., 1995; Tamagnini et al., 2002). Hydrogen is an important fuel source and is widely applied in fuel cells, coal liquefaction, upgrading of heavy oils, and several other operations. Hydrogen can be produced biologically by various means, including the steam reformation of bio-oils, dark — and photofermentation of organic materials, and photolysis of water catalyzed by special microalgal species (Kapdan and Kargi, 2006; Ran et al., 2006; Wang et al., 2008).

BIOCRUDE

In addition to direct combustion, there is growing attention to conversion of biomass into liquid energy carriers. Applying more traditional biofuel production processes (e. g. lipid extraction followed by transesterifi­cation, fast pyrolysis and gasification) to algal biomass requires that the algae be dried prior to use. Unless ac­cess to waste heat is available, the energy required to first concentrate the biomass to a paste followed by com­plete drying far exceeds to energy value of the produced biocrude. An alternative production pathway called hy­drothermal liquefaction (HTL) bypasses the drying step and converts the algal biomass into a hydrocarbon — based biocrude fuel in the aqueous phase. A simple comparison of the enthalpies of liquid water at 350 °C and water vapor at 50 °C (i. e. drying the biomass) indi­cates that processing in liquid water saves 921 kJ/kg.

Rice Husks

The production of rice husks is about 100 million tons per year. Only a fraction of it is used as animal bedding or as fuel for energy production. In Asia bri­quettes are produced from rice husks. These are expen­sive to produce, due to the silicon content of the husks. Hill et al. (1981) obtained 110 l/kg VS at a retention time of 17 days. Pretreatment with 8% NaOH gave a methane yield of 200 l/kg VS (Vevekanandan et al.,

2011) .

Bagasse

Bagasse is the pressed stalks from sugarcane. It is washed to remove nearly all the sugar in the stalks and leaves the factory at 50% humidity with 5% sugar remaining. World production is 140 million tons (dry weight). Most of the bagasse is used as fuel in the sugar

factory and some is made into paper and fiberboard. The factories have an excess of bagasse and this is stored in the open air. The stacks produce methane and open fires are common giving off soot and polluting the air. Bagasse has a low biodegradability of 120 l/kg VS (Table 13.8).

Coffee Husks and Mucilage

The production of coffee husks and mucilage is 5 million tons per year. They are dumped near the fac­tories causing methane emissions and degrading the environment. They have a high lignin content of 21%

Often the biological energy storage systems are also based on carbohydrates like sucrose (saccharose), starch and inulin, which will not be discussed here. In this overview we will only focus on the structural carbohy­drates of terrestrial biomass.

Structural Carbohydrates

Cellulose and hemicellulose can be found in the cell wall of all terrestrial plant cells. In terrestrial biomass the combined cellulose and hemicellulose fraction represents almost always more than 50% of the total biomass based on dry weight. Cellulose is a linear poly­mer composed of b-D-glucopyranose (glucose) units forming microfibrils that give strength and resistance to the cell wall. The hemicellulose consists of a wide variety of polysaccharides (composed of pentoses, hexoses, hexuronic acids), which are interspersed with the microfibrils of cellulose, conferring consistency and flexibility to the structure of the cell wall (Spiridon and Popa, 2008).

CELLULOSE

Cellulose is the basic structural component of plant cell walls and comprises about a third of all vegetable materials. Cellulose is a complex polysaccharide, con­sisting of 3000 or more b-(1 / 4) linked D-glucose units (Table 17.2). It is present in wood in quantities between 40% and 50% on dry matter basis (Table 17.1). It is the most abundant of all naturally occurring organic compounds, comprising over 50% of all the carbon in vegetation. Cellulose is a straight-chain polymer where no coiling or branching occurs, and the molecule adopts an extended and rather stiff rodlike conformation. Cellulose consists of crystalline parts together with some amorphous regions. The chains can stack together to form larger microfibrils, which make cellulose highly insoluble in water. Cellulose microfibrils may also associate with water and matrix polysaccharides, such as the (1 / 3, 1 / 4)-b-D-glucans, heteroxylans (arabinoxylans (AXs)) and glucomannans (GMs) (Sinha et al., 2011; Fengel and Wegener, 1984).

HEMICELLULOSES

Hemicelluloses are the world’s second most abun­dant renewable polymers after cellulose in lignocellu — losic materials. Hemicelluloses are a heterogeneous class of polymers representing, in general, 15—35% of plant biomass and which may contain pentoses (b-D-xylose and a-L-arabinose), hexoses (b-D-mannose, b-D-glucose, and a-D-galactose) and/or uronic acids (a-D-glucuronic, a-D-4-O-methylgalacturonic and

a-D-galacturonic acids). Other sugars such as a-L-rham — nose and a-L-fucose may also be present in small amounts and the hydroxyl groups of sugars can be partially substituted with acetyl groups (Ebringerova et al., 2005; Peng et al., 2012; Girio et al., 2010). Compo­sition and amounts strongly depend on plant source, plant tissue and geographical location. Hemicelluloses are usually bonded to other cell wall components such as cellulose, cell wall proteins, lignin, and phenolic com­pounds by covalent and hydrogen bonds, and by ionic and hydrophobic interactions. The most relevant hemi — celluloses are the xylans and the GMs, with xylans being the most abundant. Xylans are the main hemicellulose components of secondary cell walls constituting about 20—30% of the biomass of hardwoods (angiosperms) and herbaceous plants. In some tissues of grasses and cereals xylans can account up to 50% (Ebringerova et al., 2005). Xylans are usually available in large amounts as by-products of forest, agriculture, agroin­dustries, wood and pulp and paper industries. Mannan-type hemicelluloses such as GMs and galacto — glucomannans (GGMs) are the major hemicellulosic components of the secondary wall of softwoods (gymnosperms), whereas in hardwoods they occur in minor mounts. Depending on their biological origin, different hemicellulose structures can be found (Table 17.2). Upon hydrolysis, the hemicelluloses are converted into the corresponding monosaccharides (Table 17.1). The major hemicelluloses are discussed below.

Routine analytical methods, even comprehensive 13C NMR analysis, have failed to reveal the key lignin structures originated by different biomass processes, for example, differences between Alcell OS and kraft Indulin lignins (Tables 18.3 and 18.6). Therefore, this methodology is not sufficient to distinguish typical structural features of different lignins neither in qualitative nor in quantitative analytical mode. This indicates the necessity of advanced analytical methods to describe the key characteristics in the structure of technical lignins.

Two-dimensional NMR methods, specifically the HSQC technique, allow to distinguish specific structural characteristics of various technical lignins (Capanema et al., 2001; Balakshin et al., 2003; Liitia et al., 2003). The most advanced structural characterization has been achieved so far for kraft lignins. The HSQC analysis of OS lignins showed significant amounts of specific struc­tures (Balakshin et al., 2000; Capanema et al., 2001; Berlin et al., 2006), however, their exact signal assignment was not possible due to limited NMR data for specific model compounds. Further studies are required in order to perform the proper assignments in this type of lignins. The first attempts to quantify specific lignin functional­ities in different technical lignins have already been un­dertaken (Capanema et al., 2008). The 2D NMR approach pursued by the authors of the latter article focused on the quantification of lignin moieties, which were not possible to quantify with 13C NMR alone (and other 1D NMR techniques), especially of those structures formed during pulping. The study provided important quantitative information on various structural lignin units, such as condensed lignin moieties, products of b­O-4 bond cleavage, vinyl, and alkyl-aryl structures, saturated aliphatic moieties and others, as well as lignin-carbohydrate linkages (Table 18.7).

Another useful advanced NMR analytical method is DEPT 13C NMR that allows for quantification of specific lignin functionalities overlapped in routine 13C NMR spectra (Gellerstedt and Robert, 1987). These advanced NMR methods showed the possibility of expanding our understanding of the structure of technical lignins. How­ever, more comprehensive studies and cross-validation of the advanced methodologies with independent methods are needed before these methods can be routinely used.

Many cyanobacteria are able to fix atmospheric N2 into ammonia (NH3) and produce H2 as a by-product. The reaction of nitrogen fixation is catalyzed by nitroge — nase, a complex metalloenzyme and results in the formation of 1 mol of H2 per 1 mol of fixed N2 (Phelps and Wilson, 1942):

N2 + 8H+ + 8e~ + 16ATP / 2NH3 2 , 3 (21.2) + 16(ADP + PO+H2,

where Pi is inorganic phosphate.

Nitrogenases have relatively low turnover numbers. N2 fixation is an energy-expensive process that requires two ATP molecules per electron transfer. It has, howev­er, an advantage of catalyzing an irreversible reaction and not being inhibited by H2 accumulation.

The best studied type of nitrogenase is conventional molybdenum (Mo)-nitrogenase, which is encoded by the structural genes nifHDKl. Very little is known about the regulation of nif genes. Nitrogenase consists of two components: dinitrogenase, or the MoFe protein

composed of NifD and NifK subunits, and dinitrogenase reductase, or the Fe protein, consisting of two subunits of NifH. The substrate-binding and reducing active site is located in the MoFe protein. The Fe protein con­taining a [4Fe—4S] cluster and a Mg-ATP binding site acts as electron donor to a MoFe protein. This cluster accepts electrons from Fd or flavodoxin. The fixation of N2 is always accompanied by H2 evolution (Hadfield and Bulen, 1969). The reason for production of H2 as a by-product is not yet clear. It could be a result of unavoidable leakage of reducing potential, or formation of H2 could be a prerequisite for binding of N2 to the active site (Burgess and Lowe, 1996). Besides reducing N2, nitrogenase can reduce a number of other substrates with triple bonds. Importantly, in the absence of N2 as a substrate, nitrogenase exclusively catalyzes ATP-dependent reduction of H+ to H2 (Benemann and Weare, 1974; Pickett, 1996).

8H+ + 8e~ + 16ATP / 16(ADP + P;) + 4H2 (21.3)

Indeed, in terms of H2 production by N2 fixation in het­erocystous cyanobacteria the N2 is a much more potent inhibitor than O2 (Yeager et al., 2011). This is logical,
due to the fact that heterocysts can protect enzymes from external O2. Since the replacement of N2 with argon (Ar) gas is an expensive approach for optimi­zation of H2 production, an alternative method to genet­ically modify the catalytic site of the nitrogenase enzyme has been chosen as more appropriate. Recently, site — directed mutations have been introduced to several amino acid residues coordinating the Mo—Fe active site of the nzfl-enzyme in attempts to direct the electron flow selectively to H2 production in atmospheric N2 condition (Masukawa et al., 2010). Importantly, several mutant strains demonstrated nearly similar rate of H2 produc­tion under N2 and Ar atmosphere. Moreover, these strains accumulated significantly high levels of H2 under atmospheric N2 as compared to the reference strains.

Alternative Nitrogenases

In addition to the conventional Mo-nitrogenase, nifl, N2-fixing microorganisms possess also alternative nitro- genases: second type of Mo-nitrogenase, nif2, vanadium (V)-nitrogenase, vnf, and Fe-nitrogenase, anf (Bothe et al., 2010). Presence of the Fe-nitrogenase in cyanobac­teria has not yet been documented.

Mo-nitrogenase 2, encoded by nifHDK2, is expressed in both vegetative cells and heterocysts of Anabaena variabilis under N2-fixing and anaerobic conditions (Schrautemeier et al., 1995; Thiel et al., 1995). Unicellular Chroococcidiopsis, inhabiting in a gypsum rock, where the shards provide a microaerobic, low light environment, also possesses the alternative nif 2 system. Based on the phylogenetic analysis of nifH sequences, it has been suggested that nif 2 is characteristic of unicellular or fila­mentous nonheterocystous cyanobacteria fixing N2 only under microaerobic conditions (Boison et al., 2004). Weyman and coworkers reported the amino acid substi­tution in nifD2 as a first step toward the development of nitrogenase mutants in A. variabilis, which produces large amounts of H2 in N2 containing atmosphere (Weyman et al., 2010).

V-nitrogenase has a V-Fe cofactor in the active site. It is encoded by vnfHDGK genes and is expressed in het­erocysts only under Mo-deficient conditions, in the pres­ence of V (Kentemich et al., 1988; Thiel, 1993; Thiel et al., 1995). Biochemical and spectroscopic investigations of purified proteins isolated from Azotobacter vinelandii have revealed a mechanistic difference between the Mo—Fe and V—Fe catalytic site and in H2 evolution mechanisms (Lee et al., 2009).

N2 + 12H+ + 12e~ + 24ATP / 2NH3 + 24ADP + 24P; + 3H2

As can be seen from the Eqns (21.2) and (21.4), the distri­bution of electrons and protons are different for Mo — and V-nitrogenases. V-nitrogenase can produce three
times more H2 per mole of N2 reduced compared to Mo-nitrogenase (Eady, 1996). For this reason, the pro­duction of H2 by vnf system is likely to be more efficient and it is, therefore, worth searching for organisms pos­sessing alternative nitrogenases. For a long time the presence of alternative nitrogenases was confirmed only for A. variabilis and Anabaena azollae (Ni et al., 1990; Thiel,

1993) . Recent screening of 14 different cyanobacterial strains has revealed 8 strains with nif2, and 4 strains with vnf nitrogenases (Masukawa et al., 2009), suggest­ing that alternative hydrogenases are not unique.

і і 2

Vasiliki Kachrimanidou, Nikolaos Kopsahelis, Colin Webb ,

Apostolis A. Koutinas1’*

^Department of Food Science and Human Nutrition, Agricultural University of Athens, Athens, Greece, 2School of Chemical Engineering and Analytical Science, University of Manchester, Manchester, England,

United Kingdom

*Corresponding author email: akoutinas@aua. gr

OUTLINE

Introduction 419

PHA Structure and Properties 420

PHA Production Integrated in Biorefinery Concepts 421

Valorization of Biodiesel Industry By-Products 424

Valorization of Second-Generation Bioethanol Industry By-Products 427

Valorization of By-Product Streams from Food Industries 427

PHA Production from Winery By-Products 428

PHB Production from Confectionery and Bakery Industry Waste Streams 429

PHB Production from Whey 430

Conclusions and Future Perspectives 430

References 430

INTRODUCTION

The imminent depletion of fossilized raw materials and increasing environmental concerns has paved the way toward the development of a sustainable bio-based economy. Biorefinery concepts constitute a significant aspect of the future bioeconomy era where renewable raw materials, such as widely available ligno — cellulosic biomass in conjunction with industrial by-products and waste streams, will be utilized for the production of value-added commercial products, including biofuels, chemicals, biodegradable polymers and antioxidants among others. However, the establish­ment of a new industrial sector is a difficult task not only
because of the viability of new technological advances but also because of the transition from the nonrenewable to the sustainable era should occur smoothly in order to avoid job losses and economic turmoil. A smooth transi­tion can be achieved through the integration of sustain­able processing schemes in those conventional industrial plants that generate waste and by-product streams suitable for bioconversion or green chemical conversion into value-added products.

Petroleum-derived plastics constitute an everyday commodity used mainly for packaging and disposable materials. According to the U. S. Environmental Protec­tion Agency, 31 million t of waste was generated in 2010, while only 8% of the total plastic waste was

Bioenergy Research: Advances and Applications http://dx. doi. org/10.1016/B978-0-444-59561-4.00024-3

(Bhubalana et al., 2010; Cavalheiro et al., 2012). PHAs can be categorized according to the number of carbon atoms into short-chain-length PHAs (monomers containing three to five carbon atoms) and medium-chain-length PHAs (monomers containing more than six carbon atoms) (Asbhy et al., 2011; Du et al., 2012). A historical overview of PHA research and industrial applications is presented in previ­ous publications (Solaiman et al., 2006; Verlinden et al., 2007; Du et al., 2012).

As previously mentioned, one of the most important advantages of PHAs is their biodegradability and biocompatibility. Under aerobic conditions, PHAs are degraded to carbon dioxide and water, while under anaerobic conditions, methane and water are the final products. Hence, these compounds can be utilized from various microorganisms living in soil and water as carbon source for their growth, without toxic effects to the environment.

PHB was the first member of the PHA family that was identified after isolation from Bacillus megaterium (Lemoigne, 1926). It can be produced by many bacterial strains (especially various strains of C. necator) in high concentrations (more than 150 g/l) and intracellular content (more than 80% on a dry weight basis) from commercial carbon sources (mainly glucose as well as fructose and sucrose) and starch hydrolysates (Ryu et al., 1997; Yu et al., 2003). In addition, the physical properties of PHB are similar to polypropylene. How­ever, the brittle and thermally unstable nature of PHB limits its commercial applications and constitutes one of the major reasons that have prevented its production in large-scale operations. The high crystallinity of PHB (55—80%), associated with the formation of large spher — ulites, is the main reason that causes the brittle nature of PHB. It should be stressed though that the applica­tion of appropriate processing methodologies could reduce the undesirable mechanical properties of PHB, which could be used for the production of ductile films (Barham et al., 1992). Furthermore, the molecular weight of the PHB homopolymer produced by many bacterial strains, under varying fermentation condition and utilization of different feedstocks may also result in a biopolymer with improved characteristics (Kusaka et al., 1999).

P(3HB-co-3HV) was the first copolymer of the PHA family that was identified and subsequently produced on industrial scale by Imperial Chemical Industries using a R. eutropha strain. The incorporation of 3HV units in different proportions in the copolymer by this R. eutropha strain was only possible after the addition of propionic acid as a carbon source precur­sor that induced the metabolic synthesis of 3HV units. The production of P(3HB-co-3HV) also demonstrated that it is feasible to alter the properties of PHAs by controlling fermentation conditions. For instance, the addition of increasing propionic acid concentra­tions during PHA accumulation results in increasing proportions of 3HV units (expressed as mol%) in the P(3HB-co-3HV) copolymer. In this way, it was demonstrated that the incorporation of 3HV units in the P(3HB-co-3HV) copolymer results in improved mechanical properties (Byrom, 1987; Choi and Lee,

1997) .

Since the identification and commercial production of P(3HB-co-3HV) copolymers, research has focused on the identification or modification of microbial strains capable of producing PHA copolymers without addition of carbon source precursors or the production of different PHA copolymers, with addition of carbon source precursors, containing two, three or four mono­mers that demonstrate desired properties (Madden et al., 2000; Loo et al., 2005; Koller et al., 2007a). For instance, the archeon Haloferax mediterranei accumu­lates 72.8% (w/w) of P-(3HB-co-3HV) that contains 6 mol% 3HV units directly from whey sugars, while it produces the terpolymer P(3HB-co-3HV-co-4HB) when it is supplemented with 3HV and 4HB precursors (Koller et al., 2007a). Table 24.1 presents the specific properties of various PHAs compared with major petroleum-derived plastics. Nowadays, it is widely accepted that PHA physical properties can vary from brittle PHB homopolymers with high crystallinity to flexible PHA copolymers with lower crystallinity, such as P(3HB-co-3HV) and P(3HB-co-3HHx), to elastic PHA copolymers, such as P(3HB-co-4HB) and P(3- hydroxyoctanoate-co-3-hydroxydecanoate) (Wolf et al., 2005; Whitehouse et al., 2006). In the last 30 years, PHAs have been identified as potential biopolymers for a wide spectrum of end-uses including food pack­aging, flushable hygiene products, tissue engineering applications, adhesives, agriculture and biocomposites (Wolf et al., 2005).

Pretreatments vary from hot-water extraction and steam pretreatments (often with an oxidant or other chemical) to weak and strong acid and alkali pretreat­ments (Sun and Cheng, 2002). Physical pretreatments include mechanical communition, milling and ultra­sound methods (Agbor et al., 2011; Balat, 2011), as well as irradiation.

Chemical pretreatment methods include ammonia fi­ber explosion (AFEX), organosolv treatment and the addition of either acid or alkali (Isroi et al., 2011; Ong, 2004; Dashtban et al., 2009). The use of acid as a catalyst, normally H2SO4, targets the hemicellulose to dissolve with lignin and cellulose remaining as solids, whereas the addition of alkali, normally NaOH, mainly targets lignin, leaving mainly cellulose as a solid with hemicel — luloses (Dashtban et al., 2009; Ong, 2004). Physicochem­ical pretreatments combine a mild chemical treatment with high pressure and temperature and include methods ranging from uncatalyzed solvolysis (hydro­thermolysis) to steam explosion with chemical additives such as carbon dioxide or sulfur dioxide, AFEX and "popping" techniques (Mosier et al., 2005; Pan et al., 2005; Wi et al., 2011). Recent developments include pretreatments based on alkali soaking (NaOH) coupled with extrusion (Karunanithy and Muthukumarappan, 2011). Steam explosion consists of steaming the lignocel — lulose at high pressure followed by either a rapid or slow reduction in pressure to dissolve the hemicellu — loses into solution and allow the cellulose and lignin to remain as solids (Ong, 2004; Dashtban et al., 2009). SO2 or CO2 can be used as catalysts, although SO2 can be highly toxic to downstream fermentation microor­ganisms (Ong, 2004).

Although physical and chemical pretreatments can effectively reduce the recalcitrance of lignocellulosic compounds within a shorter time frame, they result in many environmental and cost concerns for industries. They require high-energy input alongside high — pressure reactors and can produce toxic compounds and wastewaters (Isroi et al., 2011).

Biological pretreatment methods include the use of microorganisms in order to delignify the lignocellulose material (Ravichandra et al. 2013; Dashtban et al.,

2009) . The enzymes produced by the microorganisms selectively disrupt the fibril and lignin structures of the plant cell wall and provide the advantages of lower energy demands, minimal waste production and reduced effects on the environment (Isroi et al., 2011; Dashtban et al., 2009). Microbial delignification is a gentle and effective approach to remove up to 31.59% lignin from biomass such as corn stover (Wan and Li,

2010) but results in a low rate of downstream hydrolysis. Pretreatment times required for direct microbiological methods are lengthy, being typically from 18 to 35 d. Nonetheless, enzymatic delignification is an alternative option and different "-omics" technologies are likely to yield new enzymatic delignification systems from

different white rot and brown rot fungi (Martinez et al.,

2009) .

The method chosen for pretreatment is dependent upon the lignocellulosic material and the hydrolysis to be carried out afterward. If the hydrolysis step is accom­panied by microbial enzymes, which are optimized at a lower pH (4—6), an acidic pretreatment is preferred as the first step in the bioconversion process (Dashtban et al., 2009).