Hydrogenases in cyanobacteria have been studied for over 35 years (Benemann and Weare, 1974; Hallenbeck and Benemann, 1978) and many variations of hydroge — nases have been described in different bacterial phyla (Vignais and Billoud, 2007). These enzymes are frequently classified into three different groups: nitroge — nase, the reversible hydrogenase (Hox), and the uptake hydrogenase (Hup) (Ghirardi et al., 2007).

HUP—HYDROGEN UPTAKE ENZYME

Hup is a [NiFe] hydrogenase that occurs associated with the thylakoid membrane (Seabra et al., 2009). This enzyme shows the least sensitivity to oxygen among the three classes. Its function is in the oxidation of H2, returning the captured electrons to cellular electron transfer reactions. To date it has been found only in N2-fixing strains and appears to have an intimate rela­tionship with nitrogenase (Marreiros et al., 2013). Under natural conditions, nitrogenase functions to reduce atmo­spheric N2 to NH3, producing H2 in an unavoidable side reaction. It is thought that Hup functions to recycle the recently formed H2, which is oxidized back into protons or reacted with O2 in a respiratory oxyhydrogen reaction, protecting the nitrogenase from O2 inactivation, avoiding an excessive build up of H2 in the cell and recovering part of the ATP used in its formation (Bothe et al., 2010; Tamagnini et al., 2007). In the nitrogen-fixing cyanobacte­ria, transcription of the Hup-encoding genes hupSL is associated with the nitrogen depletion response and is under the regulation of the NtcA, the global nitrogen regulator (Weyman et al., 2008). Hup inactivation increases the production of H2 two — to threefold in most cyanobacteria (Ludwig et al., 2006; Tamagnini et al., 2007).

NITROGENASE—A GRATUITOUS HYDROGENASE

In nature this complex enzyme carries out a critical function, breaking the three covalent bonds of molecu­lar nitrogen (N2) providing ammonia to the cell and closing the nitrogen cycle. This process consumes a large amount of energy in the form of ATP and high — energy electrons (Eqn (22.1)), producing NH3 with the coproduction of hydrogen in an unavoidable side reaction.

N2 + 10H+ + 8e~ + 16ATP/2NH3 + H2 + 16ADP

(22.1)

The most common nitrogenase is the Mo-Fe nitrogenase, which is characterized by a complex iron-sulfur cluster containing molybdenum. While performing nitrogen fixation, up to one-fourth of the electron flux goes to­ward the reduction of hydrogen. Variations of this enzyme includes the substitution of the molybdenum by vanadium or iron (V-Fe and Fe—Fe nitrogenases, respectively), which, although a greater proportion of electrons are allocated to hydrogen production, in fact show a lower net flux of electrons to hydrogen since their overall reaction rates are much lower than that of the Mo-Fe enzyme, limiting the application of these variants in bioproduction systems. One option that is an interesting strategy for H2 production, to increase the electron flux into H2, is cultivation in the absence of N2, since nitrogenase turnover continues, but now the elec­tron flux goes totally toward hydrogen evolution. In addition, the growth arrest caused by the nutrient lim­itation is of interest as this decouples hydrogen evolu­tion from biomass production, therefore potentially leaving more energy available for H2 production (Benemann and Weare, 1974). Even so, the expression of an oxygen-sensitive enzyme in an O2 rich milieu is counter productive. To overcome this problem, temporal separation between N2 fixation and photosynthesis can be used, where during the day the photosynthetic machinery works toward the carbon fixation, which then can be consumed to power nitrogenase and consequently proton reduction. Interestingly, the peak of hydrogen production in indirect biophotolysis occurs when the cell is reilluminated, possibly due to a burst in ATP synthesis before the oxygen formed by PSII (Figure 22.1) reaches a toxic level for the nitrogenase.

Heterocyst forming species on the other hand can perform direct biophotolysis by carrying out nitrogen fixation in the differentiated cell during the day. The heterocyst can maintain an internal anoxic environment since the expression of PSII is repressed. Hydrogen production therefore is supported through the use of carbon compounds delivered by the neighboring vegetative cells.

REVERSIBLE HYDROGENASE (HOX)

In addition to nitrogenase, N2-fixing cyanobacteria can have a second hydrogen-evolving enzyme, the so-called reversible hydrogenase (Hox). This enzyme is a heteropentameric complex that is formed by a hydrog — enase module (HoxHY) and a diaphorase module (Hox — EFU), which transfers electrons from NAD(P)H to the hydrogenase module (Bothe et al., 2010). Like Hup, Hox is a [NiFe] hydrogenase, but in this case it shows a high sensitivity to O2. Its expression is totally indepen­dent from that of nitrogenase and varies among species. In some cases it is under the control of the circadian clock, where it is shown to promote hydrogen produc­tion in the dark (Hallenbeck and Benemann, 1978; Schmitz et al., 2001). The bidirectional hydrogenase is not taxon specific, being found in many different groups of cyanobacteria, and its location and organization in the chromosome are also heterogeneous. Recent studies regarding Hox transcription factors have elucidated many aspects of its regulatory mechanisms, which are reviewed elsewhere (Oliveira and Lindblad, 2009).

Research in ethanol has been targeted for the devel­opment of second-generation technology, including the strategy of SSF process, which combines in a single unit the cellulose enzymatic hydrolysis and the ethanol fermentation (Santos et al., 2010). In the SSF process, glucose released by cellulase action is directly converted to ethanol by the fermenting microorganisms, which al­leviates problems caused by the end product.

The consumption of glucose and the presence of ethanol in the culture medium would reduce the risk of undesired contamination by glucose-dependent or­ganisms. Recently, consolidated bioprocessing, which combines enzyme production, saccharification and fermentation in a single step, has gained recognition as a potential bioethanol production system, because the costs of capital investment, substance and other raw ma­terials, and utilities associated with enzyme production can be avoided using microorganisms with the capa­bility for efficient cellulose hydrolysis and ethanol pro­duction (Hasunuma and Kondo, 2012).

Recently, there are many reports that SSF is superior to the traditional saccharification and subsequent fermentation in the ethanol production because the SSF process can improve ethanol yields by removing end-product inhibition of saccharification process and decrease the enzyme loading. Moreover, SSF requires a single fermenter for the entire process and eliminates the need for separating reactors for saccharification and fermentation leading to reduce the investment cost (Boonsawang et al., 2012).

Difference between SHF and SSF is in an incipient step of their development. It is possible to note that a significant number of studies reported in Tables 3.2 and 3.3 are mak­ing a comparison between the two techniques, which shows that the SSF researches are trying to develop an effi­cient process to substitute the SHF method. On the other hand the starchy raw materials have a great use in SSF fermentation; this can be explained by the simplicity of this substrate compared to cellulosic (the efficiency of enzymatic hydrolysis is better) and the conditions of operation can be milder, facilitating the adaptation of a fermentation microorganism.

CONCLUDING REMARKS

As can be seen from the tables above, there is a growing interest in ethanol production from agroindus­trial residues of a variety of sources including grains, straws, stalks and husks such as cotton, barley, triticale, wheat, coffee, rice, canola, sugarcane and other fruits and vegetables. In terms of volume, lignocellulosic material is the predominant raw material for second — generation ethanol. However, the production costs associated with the use of lignocellulosic ethanol is high, making it necessary to develop an efficient process for hydrolysis and fermentation, where the use of simul­taneous saccharification and hydrolysis is seen a prom­ising technology, but there is also the necessity to genetically modify a microorganism to grow at high temperatures or obtain an enzyme to carry out the hy­drolysis at normal fermentation temperature. Low-cost biomass residues offer excellent perspective for large — scale application of ethanol.

Acknowledgments

The authors thank CAPES for the scholarships and SCIT-RS and CNPq for the financial support of this work.

As a perspective for the future development of bioenergy-related databases, we ask: what do we need from newly developed databases? Nucleic Acids Research publishes a prestigious annual Database Spe­cial Issue since 20 years ago. Most databases published there for a particular class of proteins such as plant TFs (Guo et al., 2008), peroxidases (Fawal et al., 2012), transporters (Saier et al., 2006) and peptidases (Rawlings et al., 2012), all provide the following data or functional­ities: (1) a general classification of targeted protein families, manually collected references, a list of charac­terized proteins curated from the literature and/or pre­dicted member proteins; (2) secondary data derived from further in-depth bioinformatics analysis, such as computer-based functional annotation (e. g. Gene Ontology or protein domain annotation), sequence alignment, phylogenetic trees, predicted protein struc­tures etc.; (3) simple Web-based BLAST search against the sequence database and text search using keywords;

(4) long-term maintenance to update regularly with new data; and (5) plenty of documentation such as help, FAQ or tutorial pages. These could be considered as criteria for a good protein family database.

Although the plant biomass formation-related data­bases listed in Table 6.2 are all very useful, none of them have sufficiently integrated various functional omics data. Biologists working on one model plant often want to take advantage of these data to study their inter­ested genes, e. g. investigate fully sequenced plant ge­nomes to look for orthologs, or transcriptome data (microarray and RNA-seq) for expression profiles or look for coexpressed genes and go to the upstream re­gions for candidate cis-regulatory motifs; all these ana­lyses have to be done using individual bioinformatics tools or servers, which often requires expert knowledge to run or to interpret the results. In addition, many of the databases are outdated and none of them have included all CWR genes. For example, Purdue’s database is an excellent resource, but it does not include many of the newly characterized CWR genes such as TF family NAC, WRKY, MYB members shown to control lignin synthesis; many of the newly characterized CAZyme families such as GT43, 61, 75, transporters for NDP — sugars and monolignols; miRNAs; DUF (domain of un­known function) families etc. It includes neither much annotation data nor any search functionalities.

Therefore, the future plant CWR gene databases should aim to include all experimentally characterized CWR genes from any organisms, associated sequences and functional descriptions collected from the published literature, e. g. those listed in Table 6.1. Such character­ized gene list could be highly useful for annotating sequenced bioenergy plants such as switchgrass, poplar, maize, sorghum and Eucalyptus grandis. The CWR gene repertories for these organisms will be highly valuable for the bioenergy research community as people are trying to select candidate CWR genes to knock down or overexpress for developing transgenic plants in these model organisms. Gene families for CWR genes and other extensive secondary bioinformatics data should also be included in the databases, particularly phylog — eny (used to identify orthologs from homologs), pre­dicted cis-regulatory element, conserved coexpression network modules of known CWR genes, expression profiling, coexpressed gene list including noncoding RNAs, genomic location, gene neighborhood, epige — nomics, protein—protein interactions, structures, subcel­lular locations, single-nucleotide polymorphism, indels, etc. Similar databases should also be developed for plant CAZymes and include the above bioinformatics-derived data types. The reason is that CAZyDB now only covered 2 (A. thaliana and Oryza sativa) of over 40 sequenced plant and green algal genomes, not to mention there are more incomplete genomes and tran — scriptomes (ESTs and RNA-seq data).

References

Biodiesel is derived from plant and animal lipids. Lipids are subdivided in two main classes based on their chemical characteristics: polar and nonpolar (neutral) lipids. Neutral lipids include the tri — and diglycerides, waxes, and isoprenoid-type lipids. Monoglycerides divide neutral lipids from polar lipids. Polar lipids include phospholipids (e. g. phosphatidylinositol and phosphatidylethanolamine), free fatty acids, and glyc­erol. Desirable feedstocks for biodiesel production are composed of a higher proportion of saturated fatty
acyl neutral, rather than polar lipids. Compared to ani­mal fats and other seed-based oils, many microalgal spe­cies have been reported to contain a relatively greater proportion of polar lipids to neutral lipids (triglycerides) and the predominance of long-chain polyunsaturated fatty acids (greater than C18). However, several species of microalgae have been shown to produce various lipids, hydrocarbons, and other complex oils suitable for biodiesel production (Banerjee et al., 2002; Guschina and Harwood, 2006). To accurately predict yields from microalgae, it is critical to understand the lipid composi­tion of the feedstock. The fluorescence probe Nile Red is often used to monitor neutral lipid composition within microalgae. However, Nile Red cannot provide informa­tion regarding carbon chain length or saturation of fatty acids. Gas chromatography is often utilized for the iden­tification of specific fatty acids and the separation, identi­fication and quantification of specific lipid classes by High-performance liquid chromatography—evaporative light scattering detection (HPLC-ELSD) has recently been described (Jones et al., 2012). An informed real­time understanding of the lipid composition of the cul­ture may lead to better cultivation practices, which can drive the accumulation of desirable lipids and ultimately higher biodiesel yields.

The oil to biodiesel conversion process is termed transesterification (Figure 10.5). During transesterifica­tion, an alcohol (e. g. methanol and ethanol) is reacted with vegetable oil (fatty acid) in the presence of catalyst. Catalysts include alkalis (e. g. KOH and NaOH) or acids (e. g. H2SO4) to produce fatty acid methyl esters (FAME) or fatty acid ethyl esters and glycerol. Generally, meth­anol is preferred for transesterification because it is less expensive than ethanol. Transesterification requires 3 mol of alcohol for every 1 mol of triglyceride to pro­duce 1 mol of glycerol and 3 mol of methyl esters. This

reaction is reversible in nature and eventually arrives at equilibrium (Fukuda et al., 2001). The produced bio­diesel is immiscible and thus easily separated from glyc­erol by phase partitioning the biodiesel in a nonpolar solvent such as hexane or heptane. The solvent is later recovered by distillation. Transesterification is an inexpensive way of transforming the large, branched molecular structure of the vegetable oils into smaller, straight-chain molecules of the type required in regular diesel combustion engines.

Using microalgae as a feedstock, biodiesel can be pro­duced from extracted algal oils or by direct conversion of the biomass. The production of biodiesel from extracted microalgal oil proceeds as described above. For direct conversion of the biomass to biodiesel, the microalgae are first concentrated to a paste-like consis­tency. The cells are then incubated in methanol or ethanol in the presence of a strong acid or base at an elevated temperature. In this process, fatty acids derived from not only triglycerides but also diglycerides and free fatty acids are transesterified to biodiesel. The remaining residue contains starch and proteins, which can further be processed into ethanol, animal feed, or used as a feedstock in an anaerobic fermenter.

While the focus of this chapter has been on biolipids it is important to note that any biomass can be converted to "bio-oil" via a high-temperature process known as pyrol­ysis. This "bio-oil" also known as Synfuel or Sunfuel is currently only produced on a small scale and it very much belongs to the second-generation biofuels, as it is a way of generating fuel from a range of biomass including straw, wood or other materials high in lignin, which are difficult to convert to bioethanol. The potential for mass production remains enormous. The production of this biomass to liquid or BtL fuel can vary in complexity and can vary depending on the individual needs, but it essentially comprises the following steps.

Gasification

Gasification is a form of incomplete combustion in which a fuel is burnt in an oxygen-deficient atmosphere. An energy-rich gas, consisting principally of methane, CO and hydrogen, is formed but heat release is mini­mized. Thus an energy-rich fuel (biomass) is converted into an energy-rich gas. There are differing processes for gasification. For example, a description of the

Carbo-V process first developed by Chloren Industries but now owned by Linde Engineering GmbH was out­lined in the Biofuel Technology Handbook (Rutz and Janssen, 2007). This involves low-temperature gasifica­tion, where low-temperature pyrolysis with air or oxy­gen at 400—500 °C allows the continuous production of a gas containing both tars (volatile component) and char (carbon solids). This is followed by a high — temperature gasification, where the gas is further oxidized (again hypostoichiometrically) in a combustion chamber. The third part involves blowing the pulverized char into the hot gasification medium. Pulverized char and gasification medium react endothermically in the gasification reactor and are converted into a raw synthe­sis gas. Other gasification processes can be found, such as the recently developed Bioliq®, which was formed by Lurgi AG (Frankfurt Germany) with Karlsruhe Insti­tute of Technology (Karlsruhe Germeny).

Cleaning Process

After gasification, it is usual to have many impurities and thus cleaning remains one of the most important and most technical challenges. Remaining tars tend to be refractory and difficult to remove by thermal or phys­ical processes. Generally, the impurities in biosyngas pro­duced from the gasifier can be grouped into three types: (1) organic impurities, such as tars, benzene, toluene, and xylenes; (2) inorganic impurities, such as O2, NH3, HCN, H2S, Carbonyl sulfide (COS), and HCl; and (3) other im­purities, such as soot and dust. Both thermal cracking, which involves the addition of steam and oxygen at 200—1000 ° C, and catalytic cracking at lower tempera­tures is possible, as is low-temperature scrubbing with an oil-based medium may all encompass the process. A multicontaminant syngas treatment process created by Southern Research Institute, Birmingham, Alabama, USA, uses a candle filter, which can be catalytic, closely coupled with the gasifier. A variety of sorbents is injected into the gasifier or between the gasifier and filter to remove various contaminants (e. g. alkali metals, sulfur species, and halides) both by reaction in the gas phase and on the filter cake. Catalysts may be incorporated into the candle filter or the filter may be coated with a catalyst to crack tar and ammonia depending on the oper­ating temperature of the candle filter. An outline of the process can be seen in Figure 12.1.

Synthesis

Two methods are available for this production step, but the Fischer-Tropsch (FT) synthesis is the most widely known. It was developed at the Kaiser-Wilhelm Institute for Research on Coal (Muhlheim/Ruhr) in 1925. In Germany, coal to liquid fuels have been

produced with the help of FT synthesis since 1938. During the process, CO and H2, with the aid of a catalyst, will form hydrocarbons. A variety of catalysts exist, but the most common are usually transition metals such as cobalt. In the case of biomass, however, an iron catalyst is often favored (Hu et al., 2012).

The other process is the methanol-to-gasoline® method, in which the syngas is first transformed into methanol as an intermediate state. In a following step, fuels can be obtained from this compound. Finally, after separating the produced liquid hydrocarbons into heavy, medium and light fractions, these hydrocarbons are refined and blended to achieve the desired fuel properties.

CONCLUSION

The search for a sustainable supply of fuel that does not contribute to global warming has consumed envi­ronmental scientists for decades. While it is unlikely there is a "silver bullet" solution to the pending energy crisis the use of biolipids has enormous potential to meet a large proportion of the global transport fuel requirements. Similarly, no Single lipid source is pro­duced in sufficient quantities to impact on the world’s fuel supplies; therefore, a combination of all biolipids outlined above will be required if biolipids are to be a realistic alternative to petroleum-based fuels. While plant-derived biolipids currently dominate the liquid bioenergy markets, microalgae remain the most prom­ising source of biolipids in the future. The limited land
usage requirement and efficient carbon fixing capabil­ities of microalgae make them the ideal choice as a source of biolipids; however, there are a number of stumbling blocks to be overcome before algal biofuels are a commercial reality. These include the challenge of growing algae at industrial scale to meet the increasing demand for liquid transport fuel, the energy input involved in harvesting and dewatering algae and finally the cost and environmental impact of efficiently extract­ing biolipids from algae. These challenges are far from insurmountable, however, and each challenge is being tackled by numerous academic institutions and increas­ingly, by large, multinational energy, food and industrial chemical companies. This concerted effort with regard to algae biofuels, coupled with the more established plant — and animal-based biofuel industries can supply a significant portion of the world’s energy needs in the future.

The use of LDH as catalysts for transesterification reactions is less common if compared to the use of LDOs derived from LDH by calcination. However, the t-butoxide intercalated Mg/Al LDH (LDH/t-BU) was shown to be catalytically active for production of b-ketoesters by transesterification with primary, second­ary and tertiary alcohols (Choudary et al., 2000).

Serio et al. (2006) synthesized Mg/Al LDHs by copre­cipitation at pH 10. After washing and drying, the LDHs were calcined at 500 °C for 14 h to produce the corre­sponding oxides. Besides, two samples of oxides identi­fied as MgO-1 and MgO~2 were obtained by calcination of Mg(OH)2 and (MgCO3)4 Mg(OH)2 at 400 °C. All these oxides were tested as catalysts for soybean oil methanol — ysis. Reactions carried out with 10 wt% catalyst at 100 °C yielded about 80% of products using the LDO solids and less than 20% with both MgO-1 and MgO~2. The higher activity of LDO, with respect to other catalysts, was justi­fied by the presence of a higher concentration of very strong base sites and large pores that favored the reaction by rendering the active sites more accessible to the bulky triglyceride molecules. In another study (Serio et al.,

2007) , an LDO obtained in the same way was used in the methanolysis of soybean oil with and without the addition of 10 wt% of its weight in oleic acid. The reaction was carried out at 180 °C for 1 h with 5 wt% catalyst using commercial MgO as a reference material. The methanol — ysis of neutral soybean oil was catalyzed with LDO and MgO and the yields were 92% and 75%, whereas the cor­responding values for the acidified soybean oil were 80.3% and 76.6%, respectively.

Unlike the direct use of LDOs, Xi and Davis (2008) rehydrated the LDO and tested the resulting material as catalyst for transesterification. The experiments started with the coprecipitation of an Mg/Al LDH with an Mg:Al molar ratio of four. The material was calcined at 500 °C under nitrogen atmosphere to form the LDO and then rehydrated with vapor under nitro­gen atmosphere. The crystallinity of the resulting rehy­drated LDH was lower than that of the initial LDH. The absence of CO2 in the rehydration process avoided formation of carbonate ions. Hence, the counterion in the LDH structure was the hydroxyl ion. For this reason, the hydrated LDH had more Bransted sites than a typical LDH. This material was subsequently used in the methanolysis of tributyrine and the yield of monoes­ters was around 80% when the reaction conditions involved 136.5 g of methanol, 43.0 g of tributyrine and 0.25 g of catalyst at 60 °C for 400 min.

Zeng et al. (2008) synthesized various LDHs with different Mg:Al molar ratios by coprecipitation and ripened them at 65 °C. The solid LDHs were washed and dried at 90 °C to be subsequently calcined in a muffle at 673—1073 °C for 7 h, with the resulting oxides being tested in the transesterification of refined colza oil. The catalytic activity was correlated with the tempera­ture and time of calcination as well as with the Mg:Al molar ratio. The best yield (90.5%) was obtained from the oxide with Mg:Al molar ratio of three that was calcined at 500 ° C for 12 h. In this case, the transesterifi­cation was carried out with 1.5% of catalyst in relation to the oil mass, a methanol:oil molar ratio of 6:1 and stir­ring at 300 rpm for 4 h at 65 °C. In addition, the reuse assays showed that the catalytic activity was kept for six cycles with a slight decrease in ester yield after each cycle.

Mg/Al LDOs were also tested by Xie et al. (2006) in the transesterification of soybean oil with methanol. The precursor was synthesized by coprecipitation at pH 7. The material was calcined for 8 h at different tem­peratures and the obtained LDO was tested in the trans­esterification of soybean oil with a methanol:oil molar ratio of 15:1, 7.5% of catalyst and heating under reflux. The Mg:Al molar ratio of three yielded 67% of ester, which was the best result if compared to other molar ratios of 2.0, 2.5,3.5 and 4.0. The calcination temperature also influenced the catalytic activity. Actually, the calci­nation temperature affected the basic strength of the oxides as determined by the Hammett method. When the calcination temperature was increased from 300 °C to 500 °C, the methyl ester yield reached a maximum of 66%. The highest yield was attributed to the achieve­ment of the highest basicity after calcination. According to XRD, this oxide corresponded to the MgO periclase phase. Temperatures above 500 °C transformed the crystalline phase into spinel with less basicity and also less catalytic activity. Calcination below 500 °C led Al3+ to replace Mg2+ sites and the basicity Al bonded to O2~ is lower than that of Mg bonded to O2~. For the LDH with an Mg:Al molar ratio of three, calcination at 500 °C led to the optimal basicity for catalytic applica­tions in the methanolysis of soybean oil.

Cantrell et al. (2005) reported the use of layered materials for the catalytic transesterification of glyc­erin tributyrate. For this purpose, a series of [Mg^l xj

Alx(OH)2]x+ (COs)^ compounds with the x value rang­ing from 0.25 to 0.55 were calcined at 500 °C for 3 h under wet N2 flux (95% humidity). Also, pure Al2O3 and samples of magnesium-impregnated calcined hydrotalcite were used as reference materials and no catalytic activity was detected in any of these com­pounds. On the other hand, the LDOs improved their catalytic efficiency with an increase in their magnesium content, achieving a maximum ester yield of 74.8% with 25% of magnesium in the LDO structure. The reactions were always performed at the same experimental condi­tions (60 °C for 3 h), in which pure MgO yielded only 11% of esters.

In another study, heterogeneous catalytic processes were developed for the alcoholysis of triglycerides using LDOs that were impregnated with alkaline metals (Trakarnpruk and Porntangjitlikit, 2008). Mg/Al-NO3 LDHs were synthesized by coprecipitation and calcined at 450 °C for 35 h. The resulting oxide was added to a potassium acetate solution in order to impregnate the oxide with potassium ions. The material was recovered from the solution, dried at 100 °C for 12 h and calcined again at 500 °C for 2 h. The potassium content of the resulting powder was 1.5%. FAMEs with a 96.9% ester content and methyl ester yields of 86.6% were obtained with these solids in reactions carried out at 100 °C for 6 h, using 7% of catalyst and a methanol:oil molar ratio of 30:1.

Liu et al. (2007) carried out the catalytic conversion of chicken fat to methyl esters using oxides that were derived from the Mg6Al2(CO3)(OH)16’4H2O hydrotalcite by calcination at different temperatures (400—800 °C) for 8 h. As a result, the effect of the calcination temperature on the catalytic performance of the oxide was confirmed, as already described by Xie et al. (2006). High yields of 94 wt% were obtained when the LDH was calcined at 550 ° C and the reaction was carried out at 120 °C for 6 h with a catalyst loading of 0.04 mg/l. The catalyst activity decreased slightly in the first recycling stage but dropped to only 60% of the original value after the fourth consec­utive reaction cycle. However, the original activity could be totally recovered by calcination of the spent catalyst in air.

Antunes et al. (2008) catalyzed the methanolysis of soybean oil with Mg/Al and Zn/Al oxides that were obtained by calcination of the corresponding LDH at 450 °C for 12 h. Transesterification was performed for

7 h at 70,100 and 130 °C with a methanol:oil molar ratio of 55:1. The highest activity was detected at 130 °C and the yield at this temperature was 80% with MgO, 70% with Mg/Al LDO, 63% with Zn/Al LDH, 30% with ZnO, and 11% with A^O3.

Ilgen et al. (2007) used LDOs derived from Mgg — Al2(OH)16CO3-4H2O for the catalytic conversion of canola oil into methyl esters. The LDH was prepared by coprecipitation of magnesium and aluminum carbon­ate salts at pH 10 and ripened for 18 h. After separation, the solid compound was dried at 80 °C and then calcined at 500 °C for 16 h. The LDO with particle diam­eter of 150—177 mm gave a 63% ester yield when the re­action was carried out at 60 °C with methanol:canola oil molar ratio of 6:1. Higher molar ratios of 9:1, 12:1 and 16:1 decreased the ester yields to values lower than 60%, and when LDOs with other particle sizes (125, 125—150 and 150—177 mm) were used, the best perfor­mance was obtained in the range of 125—150 mm. In the same report, the use of n-hexane as a cosolvent was shown to be detrimental to methanolysis. Also, methanol resulted in better ester yields than ethanol.

Barakos et al. (2008) calcined Mg/Al-CO3 at 350 °C for 6 h and tested it for methanolysis of cotton oil. Sam­ples with 95% esters were obtained for reactions carried out at 180 °C, using methanol:oil molar ratios of 6:1 wt% and 1 wt% of catalyst at 2200 kPa.

Albuquerque et al. (2008) prepared calcium and mag­nesium mixed layered hydroxides by coprecipitation from which LDO catalysts were generated. The LDH was calcined at 800 °C and the resulting oxides were tested in the catalytic methanolysis of sunflower oil at 60 °C. Higher yields of 92.4% were obtained in methanol­ysis after 3 h using a methanol:oil molar ratio of 12:1 and a 2.5 wt.% of the solid catalyst with a 3.8 Mg:Ca ratio.

Macedo et al. (2006) prepared (Al2O3)4(SnO) and (Al2O3)4(ZnO) LDOs from the corresponding Sn/Al and Zn/Al LDHs and both present catalytic activity in the alcoholysis of soybean oil, even when branched alco­hols were used. Yields higher than 80% were obtained with methanol after 4 h at 60 °C and the recycling tests indicated that these materials did not lose their catalytic activity.

Shumaker et al. (2008) used LDO catalysts to convert soybean oil in methyl esters. The LDH precursors (Mg/Al, Fe/Al and Li/Al) were obtained by coprecipi­tation and subsequently calcined at 450 ° C for 2 h. The best catalytic performance was obtained with the oxide derived from [LiAl2(OH)6](CO3)0,5.nH2O, reaching a conversion of 83.1% in 2 h with a methanol:oil molar ratio of 15:1. Under the same conditions, the LDO derived from the Mg/Al precursor yielded only 13.6% of products. The same catalysts were also tested in the methanolysis of glyceryl tributyrate. The reactions were carried out at 65 °C under reflux for 1 h with 20 mmol of glyceryl tributyrate, 600 mmol of methanol and 0.1 g of catalyst. The Li/Al LDO gave yields higher than 98% while the LDOs with Mg/Al and Mg/Fe yielded only 32% and 23.9%, respectively. These results were close to the 37.1% yield that was achieved with MgO under the same conditions. These authors also observed the influence of the calcination temperature on the catalytic performance and concluded that the optimal temperature to obtain the best synthetic LDOs is between 450 °C and 500 °C.

Ngamcharussrivichai et al. (2007) used CaO. ZnO mixed oxides as heterogeneous catalysts for the metha — nolysis of palm kernel oil. A layered hydroxide formed by a mixture of the divalent cations (Ca2+ and Zn2+) was coprecipitated in alkaline media. The mixed hydro­xide was then subjected to calcination between 600 °C and 900 °C for 2—6 h. Ester yields higher than 94% were obtained with this catalyst after 1 h at 60 °C using a methanol:oil molar ratio of 30:1 and a catalyst loading of 10 wt%. Also, the mixed oxide was shown to have a Ca:Zn molar ratio of 0.25.

LDHs containing Zn/Al and Mg/Al with different counterions (nitrate, chlorite and carbonate) and M2+/M3+ ratios were synthesized by Cordeiro et al. (2012) and used as catalysts in the esterification of fatty acids with methanol. The best conversion of 97 wt% was obtained with Zn5AlCl for a reaction that was carried at 140 °C with a methanol:lauric acid molar ratio of 6:1 and 2 wt% of the solid catalyst. However, all the LDHs tested were converted in situ to layered carboxyl — ates, which preserved their catalytic activity even after several consecutive cycles of reuse.

LDH compounds containing Mg2+, Ni2+ and Al3+ were synthesized by Wang and Jehng (2011) and calcined at 500 °C for 10 h to produce heterogeneous LDO catalysts for biodiesel production. The best condi­tion for synthesis involved the use of a methanol:soy — bean oil molar ratio of 21, 0.3 wt% of catalyst, 105 °C and 1200 rpm for 4 h, when an 87% conversion of soy­bean oil to methyl esters was obtained. The observed catalytic efficiency was related to the basicity and Mg content of the Mg/Al/Ni catalysts.

Corma et al. (2005) also applied LDOs in the transes­terification of monoesters with glycerol. LDHs were initially calcined at 450 ° C under nitrogen flux for 8 h to produce LDOs that were immediately rehydrated in N2 atmosphere to avoid the presence of CO2. MgO was also synthesized from magnesium oxalate by calci­nation at 500 °C for 6 h and used as a control. The LDOs containing Li/Al had a performance better than those containing Mg /Al or MgO due to formation of stronger Lewis basic sites, since Li+ ions, which are more electro­positive than magnesium, increase the charge density of the oxygen. Based on this, alumina was impregnated with KF and the resulting material revealed basicity even higher than that of the Li/Al LDO. The catalytic conversion of glycerol to glycerin oleate with KF/ Al2O3 was 98% with monoester selectivity of 69%.

Cordeiro et al. (2008) showed that the LHS zinc hy­droxide nitrate [ZHN, Zn5(OH)8(NO3)2-2H2O] can be used as a heterogeneous catalyst for the esterification of fatty acids and for the transesterification of vegetable oils. For the transesterification reaction carried out at 150 °C for 2 h with 5 wt% of ZHN and a methanol:palm oil molar ratio of 48:1, the resulting ester layer contained

95.7 wt% of methyl esters and the purity of the glycerin layer was as high as 93 wt%. Also, when esterification was carried at 140 °C for 2 h with a methanol:lauric acid molar ratio of 4:1, the final ester layer contained

97.4 wt% of methyl laurate. In addition, these authors were able to demonstrate that ZHN turned into zinc laurate—(C12H23O2)2Zn—during the reaction course and this new layered material was held responsible for the observed catalytic activity.

LDHs containing Zn/Al and Mg/Al with different counterions and M2+/M3+ ratios were used as cat­alysts in the esterification of fatty acids with methanol. All LDHs were synthesized by coprecipitation and high conversion rates were obtained depending on the reaction condition. For instance, a 97 wt% conversion of lauric acid to methyl laurate was obtained using a methanol:fatty acid molar ratio of 6:1 and 2 wt% of cata­lyst at 140 °C for 2 h. However, all LDHs were also converted in situ into layered carboxylates and this new material was responsible for the observed catalytic activity, which was preserved even after several consec­utive cycles of reuse (Cordeiro et al., 2012).

Reinoso et al. (2012) used zinc carboxylates (acetate, laurate, palmitate, stearate and oleate) as catalysts for the methanolysis of soybean oil. Methyl ester conver­sions as high as 98 wt% were obtained for yields in the range of 84 wt% when the reaction was carried out for 2 h with a 30:1 methanol:oil molar ratio and a catalyst loading of 3 wt% in relation to the oil mass.

Jacobson et al. (2008) developed a solid catalyst by immobilizing zinc stearate in silica using the sol—gel method. The resulting solids contained 6 wt% of zinc and a total available surface area of 35 m2/g. These solids were shown to be catalytically active in the meth — anolysis of used frying oil with high acid number (~15%). High yields of 98wt% were obtained at 200 °C for 10 h using a methanol:oil molar ratio of 18:1 and 3 wt% of catalyst in relation to the mass of the start­ing material.

Lisboa et al. (2012) described the synthesis and char­acterization of layered copper(II), manganese(II), lantha — num(III) and nickel(II) laurates as well as their catalytic activity in the methyl and ethyl esterification of lauric acid. Conversions between 80 wt% and 90 wt% were observed for all catalysts when methanol was used for esterification, whereas only manganese laurate gave a reasonable catalytic activity of about 75 wt% with the use of ethanol. In general, the best results were obtained at temperatures around 140 °C. Also, the structure of copper(II) and lanthanum(III) laurates was shown to be preserved after three consecutive reaction cycles.

A detailed understanding of the structure of technical lignins is critically important in order to direct the efforts toward their valorization (Glasser, 2000). Not surpris­ingly, there is in the literature significant evidence suggesting that the performance of purified technical lignins can be linked to their chemical structure (Gosselink et al., 2010; Berlin, 2011, 2012a, b; Chung and Washburn, 2012). However, it is recognized that there is a fundamental lack of knowledge in the under­standing of technical lignins as a polymer and their con­version to materials, so targeted modifications via refining, chemical modifications, or fractionation can be pursued to maximize their performance in formu­lated products (Baker and Rials, 2013). Hence, the importance of the lignin analytical methods employed
to study the structure of these lignins will be discussed in detail below.

Native lignin is an irregular heterogeneous polymer. The same applies to technical lignins with the particular­ity that the lignin heterogeneity is typically increased by the biomass processing. It is widely believed that the lignin structure is tridimensional; however, there is no solid evidence supporting this hypothesis. Some scien­tists question the latter claim (Ralph et al., 2004). Lignin is optically inactive. The repeated (monomeric) unit in lignin is the phenylpropane unit (or so-called the "C9-unit") of the p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) types (Figure 18.2). Coniferous lignins are predominantly of G-type. Hardwood lignins contain both G — and S-units. The H-unit content in woody lignin is usually low; however, they can significantly contribute to the structure of nonwoody lignins, for instance, from annual fibers. In addition, annual fiber lignins contain significant amounts of cinnamic and ferrulic acid derivatives attached to lignin predomi­nantly via ester linkages with the gamma hydroxyl of C9-units (Adler, 1977; Sakakibara, 1991; Ralph et al., 2004). The lignin C9-units contain different functional groups. The most common ones are aromatic methoxyl and phenolic hydroxyl, primary and secondary aliphatic hydroxyl, small amounts of carbonyl groups (of the aldehyde and ketone types) and carboxyl groups. The monomeric C9 lignin units are linked to form a polymer by C—O—C and C—C linkages. The most abundant lignin interunit linkage is the b-O-4 type of linkage (structures 1—4, and 7; Figure 18.2) comprising about 50% of the interunit linkages in lignin (ca. 45% in soft­woods and up to 60—65% in hardwoods). Other com­mon lignin interunit linkages are the resinol (P~P) (structure 6; Figure 18.2), phenylcoumaran (b-5) (struc­ture 5; Figure 18.2), 5-5′ (structure 12; Figure 18.2), and

FIGURE 18.2

4- O-5 (structure 11; Figure 18.2) moieties. The number of these structures varies in different lignins, but rarely ex­ceeds 10% of the total lignin moieties. The number of other lignin moieties is usually below 5% (Adler, 1977; Sakakibara, 1991; Balakshin et al., 2008).

The degree of lignin condensation (DC) is an impor­tant lignin characteristic as it is often correlated (nega­tively) with lignin reactivity. The definition of condensed lignin moieties found in the literature is not always clear. Most commonly, condensed lignin struc­tures are lignin moieties linked to other lignin units
via 2, 5 or 6 positions of the aromatic ring (in H-units also C-3 position). The most common condensed struc­tures are 5-50, b-5, and 4-O-5′ structures. Since the C-5 position of the syringyl aromatic ring is occupied by a methoxyl group and therefore it cannot be involved in condensation, hardwood lignins are less condensed than softwood lignins.

According to recent findings, almost all lignin in soft­wood and softwood pulps is linked to polysaccharides, mainly via hemicelluloses (Lawoko et al., 2005). The main types of lignin—carbohydrate (LC) linkages in
wood are phenyl glycoside bonds (A), esters (B) and benzyl ethers (C) (Figure 18.2; Helm, 2000; Koshijima and Watanabe, 2003; Balakshin et al., 2007; Balakshin et al., 2011; Balakshin et al., 2014). The occurrence of sta­ble LC bonds in native lignins is one of the main reasons preventing selective separation of the wood components in biorefining processes.

Technical lignins are obtained as a result of lignocellu — losic biomass processing. Technical lignins differ dramatically from the corresponding native ones as a result of a combination of multiple reactions including catalyzed biomass hydrolysis, condensation of lignin fragments, elimination of native lignin functional groups, formation of new functional groups, and others. They are appreciably more heterogeneous (in terms of chemical structure and molecular mass) than the native lignins. Technical lignins have a high variety of structural moieties present in rather small amounts (Balakshin et al., 2003; Liitia et al., 2003).

Technical lignins can be classified from different points of view (Table 18.2). From a practical point of view, there are technical lignins originated from pulp and paper industrial processes which are considered mostly waste products without controllable chemical properties. These are kraft and soda lignins (kraft and soda pulping, correspondingly) and lignosulfonates (sulfite pulping). On the other hand, there is a big group of technical lignins from various emerging biomass bio­refining processes such as different variations of AH, steam explosion (SE), and OS pretreatment, in particular.

In terms of the process chemistry, and, correspond­ingly, the lignin chemical structure, lignins can be derived from acid — or alkali-based processes. The former includes most of the emerging biomass biorefinery pre­treatments, such as AH, SE (except AFEX) and most of OS processes as well as lignosulfonates. Alkaline pro­cesses are kraft and soda pulping, AFEX pretreatment, and some OS processes. In addition to the process na­ture, the feedstock source has naturally an important impact on the structure of technical lignins.

Another consideration which can be used to classify technical lignins, especially in view of their application, is the presence or absence of sulfur in their structure. Therefore, kraft lignin, and, especially lignosulfonates, are sulfur-containing lignins whereas soda, OS, AH and SE lignin are sulfur-free or low-sulfur-containing lignins.

In terms of the chemical structure, native lignins un­dergo significant degradation/modification during biomass processing. Lignin degradation occurs predom­inantly via cleavage of b-O-4 linkages (although the mechanisms are different for different processes), which results in an increase of phenolic hydroxyls and a decrease in lignin molecular mass. The lignin degrada­tion also leads to a decrease in aliphatic hydroxyls, oxygenated aliphatic moieties and the formation of carboxyl groups and saturated aliphatic structures. In contrast to lignin degradation, some reversed reactions, such as lignin repolymerization/condensation, take place to some degree resulting in increase of lignin mo­lecular mass and decrease of its reactivity. These changes are common for most of the technical lignins although the degree of transformation varies significantly de­pending on the process conditions (temperature, time, pH, and others) and feedstock origin.

Each process provides the lignin with specific chemi­cal characteristics. First, the reaction mechanism is quite different in acidic and basic media. The cleavage of b-O — 4 linkages under alkaline conditions occurs via a quinone methide intermediate which results in the for­mation of coniferyl alcohol-type moieties as a primary reaction product (Figure 18.3). They are not accumulated in the lignin; however, they undergo further secondary rearrangement reactions forming various (aryl-) aliphatic acids. b-5 and b-1 type of linkages of the native lignin cannot be cleaved during the process but are transformed into stilbene-type structures (structure 30; Figure 18.2). The latter are stable and are accumulated in alkaline lignins. In addition, a significant amount of vinyl ether structures (structure 29; Figure 18.2) forms

during soda pulping and accumulates in lignin, in contrast to kraft lignin. Another relevant structural dif­ference between soda and kraft lignins, resulting from differences in the reaction mechanism, is the presence of aryl-glycerol type structures (structure 20; Figure 18.2) in the former. On the other hand, lignin undergoes demethylation reactions which result in formation of

o-quinone structures during kraft pulping (but not in the case of soda pulping). In addition, kraft lignins contain small amounts of organically bound sulfur, likely in the form of thiol compounds (Marton, 1971; Gierer, 1980; Gellerstedt, 1996; Balakshin et al., 2003). Kraft and soda lignins show significantly higher degree of condensation than the corresponding native lignins. However, this is the result of accumulation of condensed moieties of original native lignin rather than the result of extensive condensation reactions during pulping (Balak­shin et al., 2003). Kraft and soda lignins contain small amounts of carbohydrate and ash impurities. The amounts of these contaminants are dependent on feed­stock origin and are significantly higher in annual fiber lignins than in woody lignins.

The lignin chemistry originated from the emerging acid-based biomass biorefinery processes is very diverse (Glasser et al., 1983). The acid-based biomass biorefining can be catalyzed by addition of mineral or organic acids (from catalytic amounts to the use of organic acids as the reaction media) or without acid addition (autohydroly­sis) when organic acids are generated due to cleavage of acetyl groups of lignocellulosics as well as due to the formation of acidic reaction products. Technical lig­nins derived from biomass biorefinery processes have
been much less investigated than kraft lignins. More­over, a high diversity of lignins is expected in the future given the large number of technical biomass pretreat­ment processes under either R&D or industrial deploy­ment and the high variety of potential raw materials (softwoods, hardwoods, annual fibers, agricultural resi­dues, etc.) as compared to the relative uniformity of pulping processes.

The main pathway of lignin degradation under acidic conditions is the acidic hydrolysis of b-O-4 linkages (Figure 18.4). The major products of this reaction are the so-called Hibbert ketones (Wallis, 1971). The accu­mulation of Hibbert ketones in lignin results in relatively high content, as it compares to alkaline lignins, of carbonyl groups and the corresponding saturated aliphatic structures (Berlin et al., 2006). Although degra­dation of lignin under acidic condition occurs via vinyl ether intermediates, they do not accumulate in the lignin since vinyl ether structures are very unstable in acid me­dia. Significant amounts of olefinic moieties were observed in lignin obtained under acidic conditions, but their nature is different from the olefinic structures of kraft and soda lignins, their exact structure is still not well understood (Berlin et al., 2006). Moreover, lignin condensation reactions under acidic conditions are more significant than those occurring in alkaline pro­cesses. Acidic lignin condensation occurs predomi­nantly via 2 and 6 positions of the aromatic ring, in contrast to alkaline condensation which occurs predom­inantly at the C-5 position of the aromatic ring (Glasser et al., 1983). The DC is dependent on the acidity of the reaction media (pH and solvent used) and the process

Hibbert ketones

severity (temperature and time). An extreme example of highly modified technical lignins is the industrial AH lignin produced in Russia or Belarus which is obtained at 170-190 °C, 2-3 h with 1% H2SO4. AH lignin is insol­uble in polar organic solvents and alkaline solutions due to the strong condensation/polymerization occurred during the AH process. Hydrolysis lignin has high con­tent of phenolic hydroxyl groups and olefinic structures. In addition, it contains 10-30% residual carbohydrates and up to 20% lipophilic extractives (Chudakov, 1983). In contrast, a significant fraction of AH lignin obtained at very high reaction temperature (>220 °C) but short re­action time (<1 min) was soluble in 1 N NaOH (70-90% of AH lignin) and dioxane (50%); the amounts of carbo­hydrates in these soluble lignins were significantly lower, 2-4% (Glasser et al., 1983; Lora and Glasser,

2002) . SE lignin is also quite degraded in terms of cleav­age of b-O-4 linkages, but apparently much less condensed than AH lignins (Glasser et al., 1983; Robert et al., 1988; Li et al., 2009).

OS methods of lignin production are very diverse, where different organic solvents and reaction pH can be varied during the process. Each of these processes produces lignins that are very different in their physico­chemical and biochemical properties. Furthermore, the physical conditions (e. g. temperature, time, and pres­sure) and chemical conditions (e. g. pH and concentra­tion of solvents) under which these processes are conducted can drastically affect the molecular weight (Mw), chemical structure, and functional groups distri­bution in the generated lignin derivatives (Abdelkafi et al., 2011; Balakshin et al., 2013a, b). The most investi­gated OS delignification technology is the Alcell process
deployed at industrial scale in Eastern Canada during the 1980s. The Alcell process can be carried out in aqueous ethanol liquor at moderate acidity (no exoge­nous acid is added; the acidic pH results from formation of organic acids during the process). Alcell lignin is practically sulfur-free and has significantly lower amounts of carbohydrate and ash impurities compared to kraft lignins.

Lignosulfonates are a special class of technical lignins and they constitute the bulk of the commercial lignins for materials and chemical applications. Lignosulfonates are primarily isolated from sulfite spent liquors. Howev­er, sulfonated or sulfomethylated kraft lignins are often used in similar to lignosulfonates applications but they have different chemical properties, in particular the sul­fonic acid groups in lignosulfonates are located in the side alkyl chains, whereas in sulfonated kraft lignins they are found in the aromatic ring. Sulfur-containing lignins, in form of lignosulfonates or sulfomethylated kraft lignins, are commercialized by Borregaard (Sarpsborg, Norway), MeadWestvaco (Richmond, VA, USA), Tembec (Montreal, Quebec, Canada) and other smaller players for a wide variety of applications including dispersants for wettable powders, binders for granules and seed coatings, additives, etc. Although sulfur-containing lignins are generated during acid sul­fite pulping or are produced by sulfonation of kraft lignin, the chemistry of the process and correspondingly the lignin structure is quite specific. The main reaction in sulfite pulping is sulfonation of lignin side chain, pre­dominantly at the a-position of the propane chain as well as at the conjugated a-hydroxyl group. In addition, carbonyl groups also undergo sulfonation although the

mechanism is different from that for hydroxyl groups. Introduction of highly polar sulfonate groups into the lignin structure strongly increases its solubility in aqueous solutions. Most of the lignosulfonates contain about 1 sulfonate group per 2 monomeric units. Although strong degradation of lignin is not needed to transfer sulfonated lignin from solid phase into solution, it still takes place and the degree of degradation is dependent on the reaction conditions. Therefore, the molecular mass of lignosulfonates is very high and varies strongly. The average Mw of lignosulfonates has been reported in the range of 10,000—40,000 Da and a fraction with Mw up to 100,000 Da has been isolated. An increased number of phenolic hydroxyl groups can be observed in lignosulfonates and it is strongly depen­dent on reaction conditions (Glennie, 1971).

Bio-oil and syn-gas are fuels chemically converted from lignocellulosic feedstocks. Aggressively pursued and developed as viable bioenergy, bio-oil and syn-gas productions rely on thermal-chemical conversion (or pyrolysis) of biomass or other lignocellulosic feedstocks to combustible oily substances (comprising numerous oxygenated hydrocarbons) or H2—CO gas mix, respec­tively. Depending on process conditions, up to hundreds of compounds may be present in bio-oils, with numerous chemicals of interest (other than combusti­bility) among them (Abou-Zaid and Scott, 2012; Vender — bosch and Prins, 2010; Briens et al., 2008; Demirbas, 2009). These may include polyphenols, proanthocyani — dins, tannins, flavonoids or organic acids. Some of the (phenolic) compounds may possess biocidal activity, making them useful as pesticide, bactericide, antitermite

agent, or wood preservatives (Di Blasi et al., 2010). For instance, bio-oil made from tobacco can have antimic­robes or insect activity (Hossain et al., 2013). Bio-oil made from lignin may provide substances that replace formaldehyde-phenol resins in particle board (Vender — bosch and Prins, 2010).

Algae have attracted intensive research and develop­ment efforts for bioenergy production, because algal processes might directly be driven by photosynthesis (thus fixing CO2) or yield hydrocarbons (for drop-in refining or use as fuel). The main by-product from algal bioenergy production is the post-hydrocarbon-harvest algal mass, which might be used as feed or fertilizer. Some algae species can produce phenolics, terpenoids, carotenoids, alkaloids or sterols at significant levels (Huang and Ramaswamy, 2012; Brennan et al., 2012). Phytochemical productions from algae (as mentioned in Section (Production from Algae via Aquaculture)) might be combined with hydrocarbon production, to further valorize algal bioenergy processes.

Agricultural production is only one aspect of food systems. Sustainability in agriculture also needs to address more encompassing concepts such as food se­curity and land availability on a regional level. If agri­culture would switch to organic, 10—20% more land would be needed due to lower yields, given diets do not change and the same amount of wastage is pro­duced as today. However, dietary change is a key topic for sustainable food systems, as a large part of agricul­ture’s environmental charge stems from animal hus­bandry. Reducing meat, egg and dairy product consumption levels would greatly help to reduce envi­ronmental pressure from agricultural production. Focusing on feeding animals on grasslands and not on food crops such as soy and maize would reduce the need for land, as calorie production from crops is much more efficient than from animals. Furthermore, about 30% of agricultural production are lost or wasted globally (Godfray et al., 2010). Reducing this would also contribute to reduced agricultural land use. Such reduced land use would on the other hand reduce pressure to further increase yields. Organic production in combination with reduced waste and lower consumption of animal products that are mainly based on grassland feed (and some by-products of food pro­duction) thus comprise an optimal option for a sustain­able food system (e. g. preliminary results from the FAO-SOL-model, Schader et al., 2012).

WHAT IS SUSTAINABLE BIOENERGY
PRODUCTION?

Ways of Comparisons

When assessing the sustainability of energy crop pro­duction, the first criterion is usually its GHG perfor­mance with regard to a fossil baseline. For this comparison, the baseline fuel mix plays a crucial role, as the increasing importance of unconventional fossil fuel sources such as oil sands will increase GHG emis­sions from the baseline and its general environmental impacts, thus relatively improving the performance of bioenergy production (Faist Emmenegger et al., 2012). This bears the danger that biofuel options with increasing environmental impacts and less favorable GHG balances become relatively more sustainable. Here, we adopt a different focus as we are primarily interested in the sustainability of bioenergy production with reference to sustainability in agricultural produc­tion systems and in food systems in general. This takes all sustainability criteria into account and does not focus on the GHG balance.

Trent Chunzhong Yang1, Jyothi Kumaran2,3, Samuel Amartey4,
Miranda Maki5, Xiangling Li1,6, Fan Lu7, Wensheng Qin5’*
1Aquatic and Crop Resource Development, National Research Council Canada, Ottawa, ON, Canada, 2Human Health
Therapeutics, National Research Council Canada, Ottawa, ON, Canada, 3School of Environmental Sciences, University
of Guelph, Guelph, ON, Canada, ^Division of Biology, Imperial College of Science, Technology and Medicine, South
Kensington, London, UK, ^Department of Biology, Lakehead University, ON, Canada,

6College of Chinese Medicine, Guangzhou University of Chinese Medicine, Guangzhou, China,

7College of Bioengineering, Hubei University of Technology, Wuhan, Hubei Province, China
*Corresponding author email: wqin@Lakeheadu. ca

OUTLINE

Lignocellulosic Biomass and its Pretreatment 72

Nonbiological Pretreatment 72

Physical Pretreatments 72

Chemical Pretreatments 72

Physicochemical Pretreatments 72

Biological Pretreatment with Microorganisms 73

Potential Advantages over Nonbiological

Pretreatment 73

Biological Degradation of Lignin 73

Commonly used Microorganisms for Biological

Pretreatment 73

Natural Microorganisms and Practical Applications in Bioconversion 74

Application of White-Rot Fungus in Treatment of Different Biomasses 74

White-Rot Fungus Pretreatment of Biomass

for Animal Feed 75

White-Rot Fungus Pretreatment in Biological

Pulping 75

White-Rot Fungus Pretreatment of Biomass

for Biofiber 75

Brown-Rot Fungi 75

Soft-Rot Fungi 76

Bacteria 77

Genetically Modified Microorganisms for Biomass Conversion 77

Rational Engineering 77

Metabolic Engineering of Microbial Pathways

for Enhanced Bioproduct Production 78

Strategies of Using Microbial Pretreatment to Enhance Sugar Release for Biofuel and Bioproduct Production 79

Application of Microbial Pretreatment for Biogas Production 80

Application of Microbial Pretreatment for Biomass Conversion 81

Strategies for Microorganism Application

in Biomass 81

Commonly Used Microorganisms in Biomass

Conversion and Some Application Examples 82 Other Bioproducts Produced by Microbial Conversion of Biomass: Introduction 84

References 87

Bioenergy Research: Advances and Applications http://dx. doi. org/10.1016/B978-0-444-59561-4.00005-X

LIGNOCELLULOSIC BIOMASS AND ITS
PRETREATMENT

Lignocellulose is the primary building block of plant cell walls and is composed mainly of cellulose, hemicel — luloses, lignin and small quantities of pectin, proteins, extractives and ash. The cellulose, hemicelluloses and lignin are present in varying amounts in the different parts of the plant and are intimately associated to form the complex structural framework of the plant cell wall where cellulose and hemicellulose are bound together with lignin and other components to form a tight matrix. The composition of lignocellulose depends on plant species as well as growth conditions and age.

Lignocellulose biomass is a renewable, sustainable, abundant and cheap resource for producing renewable biofuels and bioproducts. However, their conversion into fermentable sugar before fermentation is a major hurdle due to its complex structure and recalcitrant nature. While hydrolysis of cellulose and hemicellulose yields fermentable sugars, they are not easily accessible due to the crystalline structure of cellulose and interfer­ence by the phenyl-propanoid polymer, lignin.

Bioconversion of carbohydrates from lignocellulosic feedstocks into fermentable sugars is a key challenge in the biorefinery process. Efficient, cost-effective and environmentally benign pretreatment and hydrolysis methods are required. The primary purpose of pretreat­ment is to change the architecture of the cell wall by delignification and disrupting the cellulose structure and making the lignocellulosic biomass accessible and reactive to allow high rates and yields on enzymatic hydrolysis. Pretreatment has been considered as one of the most expensive processing steps in biomass to fermentable sugar conversion (Mosier et al., 2005).

This article focuses mainly on biological conversion of biomass with microorganisms. However, nonbiolog­ical pretreatments, as well as the most frequently stud­ied and applied procedures, will also be discussed.