Luiz P. Ramos*, Claudiney S. Cordeiro, Maria Aparecida F. Cesar-Oliveira,

Fernando Wypych, Shirley Nakagaki

Research Center in Applied Chemistry, Department of Chemistry, Federal University of Parana, Curitiba, Parana, Brazil

Corresponding author email: luiz. ramos@ufpr. br

OUTLINE

Introduction 255

Heteropolyacids 257

Zeolites 258

Clay Minerals 260

Clay Minerals Improving Acidity 262

Acid-Activated Clay Minerals in Biodiesel Production 263

Case 1 263

Case 2 263

Case 3 264

Layered Materials 265

Layered Double Hydroxides 265

Layered Hydroxide Salts 265

Layered Carboxylates 266

Layered Materials as Heterogeneous Catalysts in (Trans)Esterification Reactions 266

Polymeric Catalysts 269

Concluding Remarks 272

References 272

INTRODUCTION

It is well known that most of the products derived from the chemical industry involve a catalyst in at least one step of synthesis (Figueiredo and Ribeiro, 1987). However, traditional processes for chemical conversion have numerous inconveniences such as the generation of undesirable by-products and environmental pollu­tion. For this reason, civil groups as well as govern­mental agencies are pressing the industrial sector to overcome these problems by developing alternative processes in which waste generation is minimized or even eliminated. This concept is also part of the atom
economy theory proposed by Trost (1991), in which the majority of the atoms present in chemical reagents must be incorporated into useful products.

Many traditional processes based on homogeneous catalysis have been reviewed in order to minimize waste generation. In addition, many researchers have shown that heterogeneous catalysts are excellent alternatives to generate lower amounts of waste streams and also to improve the quality of coproducts, which may contribute with additional revenue for the overall pro­duction process.

The biodiesel industry is another important sector that is following the same strategic pathway (Cordeiro

Bioenergy Research: Advances and Applications

http://dx. doi. org/10.1016/B978-0-444-59561-4.00016-4

et al., 2012; Di Serio et al., 2008). Biodiesel is a biodegrad­able fuel derived from renewable sources that can be obtained by different routes such transesterification and esterification. The traditional transesterification process of oils and fats is based on the use of a homoge­neous system, with methanol as the transesterification agent and a base catalyst, usually an alkoxide or a hydroxide (NaOH or KOH) that generates the corre­sponding alkoxide in situ (Van Gerpen and Knothe,

2009) . Then, the synthetic mono alkyl esters can be used as biodiesel after suitable purification. The main problem of these processes is related to the required pu­rification steps of the mono alkyl esters as well as glyc­erin, which must be recovered in good condition due to their high commercial value.

Ideally, the biodiesel fuel must be free of residues formed in the chemical process like free and bonded glycerin, soaps and water, which are normally used in washing stages for purification. The presence of glycerin in the resulting biofuel is problematic because this pol­yol may undergo dehydration during combustion, pro­ducing a toxic unsaturated aldehyde named acrolein that is not only a dangerous atmospheric pollutant but also a reactive chemical that can be easily involved in condensation reactions, producing an accumulation of carbon deposits that may block filters and compromise the engine performance (Mittelbach and Tritthart,

1988) . Soaps and free fatty acids (FFAs) cause degrada­tion of engine components and free water can interfere with the biodiesel acid number and induce hydrolysis and biological contamination under nonadequate stor­age conditions (Ramos et al., 2003).

The traditional fatty acid esterification processes in homogeneous media uses Bransted acids such as sulfu­ric or hydrochloric as reaction catalysts (Ilgen et al., 2007). However, the extensive use of these catalysts may induce corrosion in reactor components and pipe­lines. Also, the purification of the monoesters produced in this way is also expensive and may require additional washing steps and distillation (Altiparmak et al., 2007).

A traditional sequence for biodiesel production in­volves (1) the recovery of vegetable oil by pressing and/or solvent extraction, (2) the oil pretreatment to adjust its properties for transesterification, (3) the trans­esterification process, (4) the purification by several stages of water washing and (5) the recovery of reaction coproducts, especially glycerin. Each of these steps adds costs to the overall process. Thus, the introduction of operation units that are able to reduce the contamination degree of mono alkyl esters and glycerin may be an important measure to make biodiesel more competitive from an economic and environmental point of view.

For these reasons, many researches had focused their efforts to substitute homogeneous catalysts for heteroge­neous ones. The biodiesel produced in a heterogeneous system is easily purified and glycerin is of high purity, diminishing the investment to achieve a suitable market specification (Ramos et al., 2003). Many classes of chem­ical compounds have been tested as solids for hetero­geneous catalysis to produce biodiesel by either esterification or transesterification processes. Among these, zeolites, ionic exchange resins, inorganic oxides, layered compounds, guanidines and metal complexes have been already used (Cordeiro et al., 2011).

In order to have a truly heterogeneous catalytic pro­cess, the solid catalyst must not leach into the reaction medium and it also needs to be stable under the reaction conditions and reusable. While using solid catalysts for biodiesel preparation, whether by esterification or trans­esterification, the most common catalyst classifications are solid Bronsted acids, Bransted bases or Lewis acids. The same solid catalyst, however, may present more than one of these sites and depending on the acidity or basicity of the solid, the catalytic performance can vary considerably (Sharma et al., 2011).

Recently, in addition to this primary classification, a number of other factors have been considered while developing solid catalysts for esterification or transester­ification reactions. The solids hydrophobicity, for once, is used to unveil the water tolerance. The knowledge of the pore and channel system is used to improve the mass transfer of the catalytic substrate, which for this kind of reaction presents a relatively high viscosity (Wilson and Lee, 2012).

Metal oxides, mainly calcium (CaO), magnesium (MgO) and strontium (SrO) oxides, are among the most extensively studied solid bases for heterogeneous cata­lytic processes (Sharma et al., 2011). Among all alkaline and alkaline earth metal oxides, CaO is the most widely studied. Many are the reasons to explain this fact, including its low cost, low toxicity and low solubility in methanol, which is the most commonly used primary alcohol for the catalytic transesterification of oils and fats (Sharma et al., 2011; Kusdiana and Saka, 2001).

CaO also has a long catalytic life, with high activity in many recycling processes under moderate reaction condi­tions (Lopez et al., 2007). Besides these advantages, CaO can be obtained from various and sometimes unusual natural sources. Naturally occurring minerals such as cal — cite (CaCOs) and several calcium salts (Lopez-Granados et al., 2010) as well as mollusk shells and egg shells (Cho and Seo, 2010) can be used as a source of CaO by calcination.

The impregnation of different alkaline salts in zeolites followed by appropriate thermal treatment can produce basic zeolites and the resulting solids have shown good activity as heterogeneous catalysts for transesterifica­tion. Studies have shown that the basicity of the resulted zeolite can be related to the electropositive nature of the exchanged alkaline cation (Philippou et al., 2000).

The infrequent use of acid catalysis in transesterifica­tion reactions, in comparison to the base catalysis, is in part justified by the lower catalytic activities of the acid compounds. On the other hand, acid catalysts are less sensitive to several contaminants such as water and FFAs, which in many cases can deactivate the base catalyst or drive the catalytic reaction to other products (Van Gerpen and Knothe, 2009).

Notwithstanding this apparent disadvantage of the acid catalyst, solid catalysts with Bransted or Lewis acid properties have been recently investigated in het­erogeneous processes. These solids are promising solid catalysts for the replacement of strong inorganic acids that, although effective in both esterification and trans­esterification homogeneous catalytic systems, have serious adverse factors such as corrosion of the reaction vessels. Furthermore, the use of strong inorganic acids leads to medium — and long-term environmental prob­lems (Helwani et al., 2009a). Thus, the possibility of using solids with acid properties, rather than highly corrosive liquids, therefore replacing homogeneous pro­cesses by heterogeneous ones, may be advantageous since higher catalytic efficiencies may be obtained in more sustainable conversion processes. These are likely to outweigh the higher costs that are often associated with the rational synthesis and use of suitable solid acids.

Furthermore, the research of acid catalysts has also been driven by the possible use of waste cooking oil and other cheap and widely available raw materials for biodiesel production. For such materials, the catalyst must be suitable for acting in the presence of high water and acid content, properties that are often found in low cost feedstocks. In general, solids with high acid proper­ties usually meet these prerequisites (Oliveira et al., 2010; Zhao et al., 2012).

The present work presents a discussion about the most important classes of inorganic solids and poly­meric materials that have been applied in the synthesis of (m)ethyl monoesters through esterification or transes­terification. However, biological systems such as immo­bilized lipases are not treated in this book chapter. Luckily, highly qualified reviews have been published recently on this specific subject (Di Serio et al., 2008; Fjer- baek et al., 2009; Tana et al., 2010).

The conversion of technical lignin into these monoaromatic chemicals is assumed to be a long-term application (Holladay et al., 2007). Increased worldwide research activities can be observed in this area where predominantly thermochemical approaches are under study to convert lignin model compounds and depoly — merize technical lignins into the desired aromatic com­pounds. In general, lignin depolymerization can not only be performed in aqueous and organic phases, but also in dry form. Complex mixtures are the result in which the individual mass yields barely exceeds few percent. Mostly, C—O—C bonds are cleaved, while the C—C linkages in the lignin structures are very resistant to cleavage. The use of catalysts seems to be a necessity and these activities have been recently reviewed (Zakzeski et al., 2010; Gallezot, 2012; Azadi et al., 2013) showing the following main routes for technical lignin depolymerization in (mono)aromatic chemicals.

Base-catalyzed Depolymerization

Most work related to base-catalyzed depolymeriza­tion (BCD) originates from the pulp and paper industry where these alkaline processes are used to depolymerize and liberate lignin from the lignocellulosic matrix as described in the previous sections. Besides extensive cleavage of the b-O-4 linkages under BCD conditions the methoxyl contents in lignin decrease with the severity of alkaline conditions. However, repolymeriza­tion of liberated lignin fragments to condensation products may occur. Alcell organosolv lignin depolymer­ization in alkali (0—4%) yielded 7—30% liquid products. The maximum concentration of identified phe­nols was 4.4%, mostly syringol (2.4%) and a limited amount of guaiacol when less severe conditions were applied. Catechol was found at higher pH and tempera­tures (Thring, 1994). More recently, Yuan et al. (2010) studied the base-catalyzed degradation of kraft lignin in water—ethanol at 220—300 °C, with phenol as the capping agent into oligomers with a negligible char and gas production. Under the conditions applied lignin could not be degraded completely into lignin monomers.

Base-catalyzed lignin depolymerization with the addition of boric acid greatly facilitates the depolymer­ization of lignin in water, increase product selectivity and boric acid acts as a capping agent to suppress addi­tion and condensation reactions (Roberts et al., 2011).

Putrescine apparently has a specific role in skin phys­iology and neuroprotection (Janne et al., 2005). Fermen­tative production of putrescine can be achieved by manipulating arginine decarboxylase (ADC) pathway or ornithine decarboxylase (ODC) pathway in E. coli (Figure 19.4(a)) and C. glutamicum (Figure 19.4(b)) and of which, the ODC pathway is preferable as it comprises only a single reaction compared to two or three reactions of the ADC pathway. To increase L-ornithine formation, its conversion to L-arginine may be blocked; however, this results in unfavorable auxotrophy for L-arginine. Thus, the maintenance of prototrophy with concomitant high L-ornithine supply is a focus in strain construction. The pathway for biosynthesis of L-arginine and L-orni- thine, the substrates of the initial decarboxylase reac­tions in the ADC and ODC pathway, respectively, are similar in E. coli and C. glutamicum. There is some differ­ence in ornithine synthesis between them; C. glutamicum has a cyclic pathway while E. coli has a linear pathway (Glansdorff and Xu, 2007). The cyclic pathway is economical in terms of metabolic cost for ornithine syn­thesis when compared to linear pathway, because in linear pathway there is a concomitant hydrolysis of acetyl-CoA to acetic acid. The L-ornithine was then con­verted to citrulline by ornithine carbamoyl phosphate transferase ArgF (EC 2.1.3.3). The synthesis of all en­zymes in the pathway is subject to repression by L-argi — nine, which is mediated by the repressor ArgR in E. coli and C. glutamicum (Glansdorff and Xu, 2007). In order to use a microorganism in industrial fermentations or bio­transformations the organism should possess high toler­ance to the desired product. Concentrations of up to 66 g/l putrescine reduced the growth rate of C. glutami — cum by 34% and that of E. coli by 78% (Schneider and Wendisch, 2010).

In order to overproduce putrescine in E. coli, several attempts have been done so far. The ADC pathway is completed by agmatinase SpeB, which hydrolyzes agmatine to putrescine and urea. While urea cannot be reused by E. coli, putrescine can be utilized by E. coli as a sole carbon source. The overexpression of ODC genes speC (b2965) and of speF (b0693) in the wild — type genetic background led to comparable results as 0.72 or 0.87 g/l of putrescine accumulated in batch cul­tures (Eppelmann, 2006). The simultaneous overexpres­sion of speF and speAB, the ADC encoding gene speA as well as speB coding for the agmatinase of E. coli (b2938, b2937) increased putrescine accumulation up to 1.03 g/l (Eppelmann et al., 2006). In order to increase the putres — cine production a base strain was constructed, by inacti­vating the putrescine degradation and utilization pathways, and the engineered E. coli strain was able to produce 1.68 g/l of putrescine with a yield of 0.168 g/

FIGURE 19.4 (a) Engineered putrescine and cadaverine production pathways used in E. coli. GdH, glutamic acid dehydrogenase (EC1.4.1.4);

ArgA, amino acid N-acetyltransferase (EC 2.3.1.1); ArgB, acetylglutamic acid kinase (EC 2.7.2.8); ArgC, N-acetylglutamylphosphate reductase (EC 1.2.1.38); ArgD, acetylornithine aminotransferase (EC 2.6.1.11); ArgE, acetylornithinase (EC 3.5.1.16); ArgF, ornithine carbamoyltransferase (EC 2.1.3.3); ArgG, argininosuccinic acid synthetase (EC 6.3.4.5);ArgH, argininosuccinic acid lyase (EC 4.3.2.1); Pepck, phosphoenolpyruvic acid carboxykinase (EC 4.1.1.32); Ppc, phosphoenolpyruvic acid carboxylase (EC 4.1.1.31); Pyc, pyruvic acid carboxylase (EC 6.4.1.1); AspC, aspartic acid aminotransferase (EC 2.6.1.1); LysC, aspartokinase (EC 2.7.2.4); Asd, aspartic acid semialdehyde dehydrogenase (EC 1.2.1.11); MetL, ThrA bifunctional aspartokinase/ homoserine dehydrogenase (EC 2.7.2.4/1.1.1.3); DapA, dihydrodipicolinic acid synthase (EC 4.2.1.52); DapB, dihydrodipicolinic acid reductase (EC 1.3.1.26); DdH, meso-diaminopimelic acid dehydrogenase (EC 1.4.1.16); LysA, diaminopimelic acid decarboxylase (EC 4.1.1.20); ODC, ornithine decarboxylase (EC 4.1.1.17); ADC, arginine decarboxylase (EC 3.5.3.1).

(b) Engineered putrescine and cadaverine production pathways used in C. glutamicum. GdH, glutamic acid dehydrogenase (EC1.4.1.4); ArgJ, bifunctional ornithine acetyltransferase/N-acetylglutamic acid synthase (EC 2.3.1.35/2.3.1.1); ArgB, acetylglutamic acid kinase (EC 2.7.2.8); ArgC, N- acetylglutamylphosphate reductase (EC 1.2.1.38); ArgD, acetylornithine aminotransferase (EC 2.6.1.11); ArgE, acetylornithinase (EC 3.5.1.16); ArgF, ornithine carbamoyltransferase (EC 2.1.3.3); ArgG, argininosuccinic acid synthetase (EC 6.3.4.5); ArgH, argininosuccinic acid lyase (EC 4.3.2.1); Pepck, phosphoenolpyruvic acid carboxykinase (EC 4.1.1.32); Ppc, phosphoenolpyruvic acid carboxylase (EC 4.1.1.31); Pyc, pyruvic acid carboxylase (EC 6.4.1.1); AspC, aspartic acid aminotransferase (EC 2.6.1.1); LysC, aspartokinase (EC 2.7.2.4); Asd, aspartic acid semialdehyde dehydrogenase (EC 1.2.1.11); MetL, ThrA, bifunctional aspartokinase/homoserine dehydrogenase (EC 2.7.2.4/1.1.1.3); DapA, dihydrodipicolinic acid synthase (EC 4.2.1.52); DapB, dihydrodipicolinic acid reductase (EC 1.3.1.26); DdH, meso-diaminopimelic acid dehydrogenase (EC 1.4.1.16); LysA, dia­minopimelic acid decarboxylase (EC 4.1.1.20); ODC, ornithine decarboxylase (EC 4.1.1.17); ADC, arginine decarboxylase (EC 3.5.3.1).

g glucose. A further optimization by 25% was achieved by promoter exchange of genes encoding the enzymes converting L-glutamic acid into L-ornithine, as well as the exchange of speF—potE promoter (potE encodes the ornithine—putrescine antiporter) (Qian et al., 2009).

In contrast to E. coli, C. glutamicum is unable to degrade and utilize putrescine as a carbon source. The expression of genes of the ADC and ODC pathway from E. coli in the wild-type background of C. glutami — cum only led to minor amounts of putrescine. The dele­tion of argR and argF led to accumulation of L-ornithine but rendered the resulting strain arginine auxotrophic. When speC and speF from E. coli were expressed in the argR—argF deletion strain of C. glutamicum, produc­tion of 5 g/l putrescine resulted, which was about 50 times higher than strains endowed with the ADC pathway. To avoid costly supplementation with L-arginine and the strong feedback inhibition of the key enzyme N-acetylglutamate kinase (ArgB) by L-argi — nine, a plasmid addiction system for low-level argF expression was developed. This strain resulted in pu — trescine yields on glucose from less than 0.001 up to 0.26 g/g, the highest yield in bacteria reported to date and was named as PUT21. In fed-batch cultivation with C. glutamicum PUT21, a putrescine titer of 19 g/l at a volumetric productivity of 0.55 g/l h and a yield
of 0.16 g/g glucose was achieved (Schneider et al.,

2012) . Moreover, while plasmid segregation of the initial strain required antibiotic selection, plasmid segregation in C. glutamicum PUT21 was fully stable for more than 60 generations without antibiotic selection even in the pres­ence of L-arginine.

Cadaverine

Cadaverine can be overproduced by introduction of an overproduced lysine decarboxylase. The correspond­ing substrate, L-lysine, is synthesized in E. coli and C. glutamicum by similar pathways covering 10 enzy­matic steps initiating from the tricarboxylic acid cycle in­termediate oxaloacetate. The three initial steps in this pathway lead to aspartic acid semialdehyde, which is the branch point for biosynthesis of the amino acids, L-methionine, L-threonine, L-isoleucine and L-lysine (Figure 19.4). However, there were substantial differ­ences in the enzyme systems possessed by E. coli and C. glutamicum. When it is LysC from C. glutamicum that is additionally feedback inhibited by L-threonine, it was ThrA from E. coli that is subject to feedback inhibi­tion by L-threonine (Park and Lee, 2010). The tolerance of E. coli for cadaverine seems to be lower compared to pu — trescine. The biomass formed in the presence of 51 g/l

cadaverine was reduced by 30% in comparison to the same molar concentration of putrescine (Qian et al., 2011, 2009). Corynebacterium glutamicum was tested for growth on solid medium and grew even at concentra­tions of up to 31 g/l cadaverine (Mimitsuka et al., 2007).

Escherichia coli strains overexpressing the lysine decarboxylase gene cadA (b4131) in the wild-type ge­netic background led to accumulation of 0.8 g/l cadav — erine by growing cells. To avoid side reactions of enzymes active with putrescine toward cadaverine, a number of genes were deleted: the spermidine synthase gene speE, the spermidine acetyltransferase gene speG, the putrescine importer gene puuP, the putrescine aminotransferase gene puuA and ygjG, which encodes the initial enzyme of the second putrescine degradation pathway and is known to be active in vitro with cadav — erine. The resulting strain was able to accumulate 1.2 g/l cadaverine. Production of cadaverine was increased by 10% as a consequence of enhancing the flux of L-aspartic acid toward L-lysine by overexpression of dapA via pro­moter exchange. In fed-batch cultivation, this strain pro­duced 9.6 g/l cadaverine (Qian et al., 2011).

Cadaverine production in C. glutamicum was also achieved by insertional inactivation of homoserine de­hydrogenase gene, hom (cg1337, Figure 19.4, B-1) with cadA from E. coli. The resultant strain secretes 2.6 g/l cadaverine in the supernatant. The expression of cadA was driven by the strong kanamycin resistance gene promoter. But the strain was auxotrophic for L-methio — nine, L-threonine, and L-isoleucine (Mimitsuka et al., 2007). A different approach with biosynthetic lysine decarboxylase (LdcC) from E. coli led to 30% more cadaverine production than overexpression of cadA (Kind et al., 2010b). Later the C. glutamicum DAP-3c cadaverine-producing strain’s substrate spectrum was broadened for hemicellulose utilization by introducing xylA and xylB genes from E. coli (Buschke et al., 2011). Through various studies reasonable titers and produc­tivities were achieved for putrescine and cadaverine (Table 19.1).

CONCLUSION AND PERSPECTIVES

This chapter outlined the microbial production of amino acids, poly(amino acid)s and polyamines known so far. Biotechnological production of amino acids today serves a market with strong prospects of growth. In the foreground are the fermentation processes, which are now widely established in the production of proteino — genic amino acids; this can be extended to the produc­tion of other amino products like poly(amino acid)s and polyamines. The potential that will be leveraged in the future by modern methods and new findings in system biology will further stimulate and strengthen microbial production of amino products. Modern methods such as directed evolution will allow develop­ment of customized, highly selective, and stable en­zymes and whole cell biocatalysts, as well as efficient and ecologically sustainable production of the required products. It became a need to assess the feasibility of implementing, in addition to the established chemical processes, a biorefinery concept based on renewable raw materials. The poly(amino acid)s production does not have the luxury of background knowledge regarding the metabolic process leading to their synthe­sis when compared to the amino acids and polyamines. Even then, poly(amino acid)s was produced in recently good titers by using newly isolated strains and their genetically manipulated versions. However, the genetic engineering strategies were yet to attain maximal poten­tial in polyamine and poly(amino acid)s producing strains.

As discussed above, nonbiological production of hydrogen is energy intensive and often associated with the production of greenhouse gas. Biologically, hydrogen can be produced by a variety of microorgan­isms possessing one of several different hydrogenases. In the cyanobacteria, enzymes involved in hydrogen metabolism belong to one of the three families discussed above: Hox, Hup or nitrogenase. The uptake hydroge — nase (Hup) is not useful for hydrogen evolution since it is poised to work unidirectionally, toward the recy­cling of H2 into H+. When hydrogen is produced by a heterotrophic organism, an organic carbon source (ulti­mately derived from photosynthesis) is used to provide protons and chemical energy to fuel hydrogen evolu­tion. Ironically, this is also true for cyanobacteria carrying out direct or indirect biophotolysis, at least on the molecular level. As discussed above, a complete photosynthetic apparatus uses water as proton donor, releasing molecular oxygen (Figure 22.1). Thus, the high sensitivity of hydrogenase to this gas dictates that both reactions cannot occur in the same place at the same time. The solution found by Nature was the most obvious one: changing the timing (indirect biophotolysis) or the space (direct biophotolysis).

In indirect biophotolysis the cell uses the chemical energy stored through the capture of sunlight, as

NADPH and ATP (Figure 22.1), to fix CO2 into organic compounds. These energy reserve molecules are then consumed in the dark to drive cellular metabolism, including nitrogen fixation by nitrogenase. The separa­tion of these reactions occurs naturally in several cyano — bacterial species by circadian control and in these strains dark hydrogen production by either nitrogenase or the bidirectional hydrogenase is frequently reported (Praba — haran et al., 2010; Troshina et al., 2002). An interesting characteristic found in many of these strains is a burst of hydrogen production when cells are reilluminated. This phenotype was characterized as a function of the bidirectional hydrogenase and hydrogen production ceases quickly as the O2 produced by photosystem II (Figure 22.1) accumulates in the cell, inactivating hydrogenase The production of H2 is thought to serve as an electron sink, helping the cell return to the proper redox state for carrying out the light reactions. In prac­tice, indirect biophotolysis could possibly be done as a large-scale production using a two-stage cultivation system. In a first stage, the cells are cultivated in the light and biomass is formed through photosynthesis. When the desired cell concentration is achieved and the cells have stored enough fixed carbon, a dark anaerobic culti­vation could follow, favoring proton reduction to hydrogen by hydrogenase. Thus, the water-splitting reaction is separated from H2 production in time and space. This system has being already demonstrated, where nitrogen limitation was also used to induce glycogen accumulation and increase hydrogen produc­tion yield in the second stage through the nitrogenase enzyme (Huesemann et al., 2009). In a similar approach with Synechococcus sp., the carbon accumulated in the first stage was converted into hydrogen in a second stage by a [NiFe] hydrogenase (McNeely et al., 2010).

H2 PRODUCTION BY HETEROCYSTOUS CYANOBACTERIA

Solar energy capture and hydrogen evolution by some filamentous cyanobacterial strains proceeds natu­rally in the presence of oxygen by confining the oxygen-sensitive processes to the heterocyst, a cell type that emerged shortly after the oxygenation of the earth’s atmosphere in what has been called the Oxygen Catas­trophe or Great Oxidation Event 2.6 billion years ago (Kumar et al., 2010; Mariscal and Flores, 2010). In this case the evolved hydrogen is produced by nitrogenase whose expression is restricted to the heterocyst under normal aerobic conditions (Murry et al., 1984). A num­ber of mechanisms are employed to protect nitrogenase from oxygen damage; heterocysts lack photosystem II so do not produce oxygen, gas diffusion into the heterocyst is restricted by a unique cell wall structure, and hetero­cysts possess a very active membrane-bound respiratory system that consumes trace amounts of entering oxygen.

Even so, some continual synthesis of nitrogenase is necessary to replace oxygen-damaged nitrogenase (Murry et al., 1983).

As discussed above, since heterocysts lack a complete photosynthetic apparatus, the necessary reductant is derived from fixed carbon imported from the neigh­boring vegetative cells through specialized interconnect­ing pore structures (Mariscal and Flores, 2010). The imported sugar is sucrose (Lopez-Igual et al., 2010) and it is metabolized though the oxidative pentose pathway (Summers et al., 1995) Thus, hydrogen produc­tion by heterocysts is essentially indirect biophotolysis on a microscopic scale, and since the energy captured by photosynthesis is first stored as fixed carbon, the maximal possible theoretical conversion efficiencies are reduced.

However, this system has been attractive due to its inherent robustness and has been studied for almost four decades (Benemann and Weare, 1974). Very reason­able conversion efficiencies, sustained for days to weeks, were achieved in early studies using nitrogen-limited cultures. Under laboratory conditions where higher effi­ciencies can be expected, conversion efficiencies (total incident light energy to free energy of hydrogen pro­duced) were shown to be 0.4% (Weissman and Bene- mann, 1977). Cultures incubated under natural sunlight (Figure 22.3) were able to attain an average con­version efficiency of 0.1% (Miyamoto et al., 1979a). Remarkably, even though there have been a large num­ber of studies since, very little improvement in yields

has been obtained. Thus, recent reports of conversion efficiencies found =0.7% under laboratory conditions (Berberoglu, 2008; Sakurai and Masukawa, 2007; Yoon et al., 2006) and 0.03—0.1% with natural sunlight (Sakurai and Masukawa, 2007; Tsygankov et al., 2002). Similar low efficiencies have been found with thermo­philic strains, which at least have the possible advantage of requiring less cooling (Miyamoto et al., 1979b, c). There should be room for improvement as theoretical efficiencies with this nitrogenase-based system have been calculated to be around 4.6% (Hallenbeck, 2011).

Since observed conversion efficiencies are lower than predicted, different strategies might be employed in order to improve overall performance, which is critically important since light conversion efficiencies directly impact on the photobioreactor footprint (doubling effi­ciency should halve the required surface area for the same amount of fuel production). For one thing, genetic engineering could be applied to optimizing the size of the photosynthetic antenna, since part of the reduction in efficiency is thought to be due to inefficient use of light energy at high intensities where more photons are captured than can be used and the excess energy is wasted. Another point that could be addressed is the hydrogen producing catalyst. Since half of the photon requirement is needed to provide ATP to nitrogenase ac­tion, replacing it with a hydrogenase, which does not require ATP for proton reduction, should in principle have an energy sparing effect. In a recent attempt to verify this, the [FeFe] hydrogenase from Shewanella onei — densis was expressed in Anabaena sp. under the control of a heterocyst-specific promoter with the required matu­ration genes (Gartner et al., 2012). Although it could be shown that active hydrogenase was made under the proper conditions, the increase in hydrogen production above the levels due to the coexisting nitrogenase was disappointingly small. Of course, under these condi­tions the two enzymes compete for the reductant; the true test would be to do this in a strain lacking nitroge — nase activity. Finally, it might in principle be a possible way to increase hydrogen production by increasing het­erocyst frequency. However, heterocyst frequency might already be close to optimal since even in long-term studies the H2/O2 ratio is close to the desired stoichiom­etry of two, what one would expect for optimal coupling between oxygen-generating photosynthesis in the vege­tative cells and hydrogen production by heterocysts.

Irmene Ortiz*, Rodolfo Quintero

Departamento de Procesos y Tecnologia, Universidad Autonoma Metropolitana — Cuajimalpa, Mexico D. F., Mexico

*Corresponding author email: irmene@correo. cua. uam. mx

OUTLINE

Trends in Pretreatments 62

Other Pretreatments 62

Pretreatment Modeling 65

Environmental and Economical Aspects 65

Concluding Remarks 66

References 66

LIGNOCELULLOSIC BIOMASS

Lignocellulosic biomass is composed primarily of cel­lulose, hemicelluloses (mainly xylan), lignin and smaller amounts of other compounds. Typically, the composi­tion of lignocellulosic biomass by weight is 40—50% cel­lulose, 20—40% hemicellulose, 10—30% lignin and other components such as minerals, oils, soluble sugars, pec­tins, proteins, and ashes (Jorgensen et al., 2007; Lewis et al., 2005; Wyman et al., 2005).

Cellulose, hemicelluloses and lignin are present in varying amounts in the different parts of the plant and they are intimately associated to form the structural framework of the plant cell wall; also, the content of the different sugars of the hemicelluloses varies signifi­cantly between different plants depending on plant spe­cies, age and growth conditions (Jorgensen et al., 2007).

Cellulose is the most abundant constituent of the plant cell wall; its linear structure enables the formation of both intra — and intermolecular hydrogen bonds
resulting in the aggregation of chains into elementary crystalline fibrils of 36 cellulose chains, while hemicellu — loses are complex branched heterogeneous polysaccha­rides composed of monomeric residues: D-glucose, D-galactose, D-mannose, D-xylose, L-arabinose, D-glucur — onic acid and 4-O-methyl-D-glucuronic acid; and lignin is a complex amorphous network formed by polymeri­zation of phenyl propane units and constitutes the most abundant nonpolysaccharide fraction in lignocel — lulose (Jorgensen et al., 2007; Lewis et al., 2005).

Biofuels produced from native lignocellulose are known as second-generation biofuels. In this process the cellulose is converted into glucose, which is easily fermented to ethanol, while the hemicellulosic fraction is converted into monomeric sugars (mainly pentoses), a fermentation that is considerably harder to accomplish (Dias et al., 2011). The physicochemical and structural compositions of native lignocellulose are, however, recalcitrant to direct enzymatic hydrolysis of cellulose (Mosier et al., 2005). Therefore, a pretreatment step is

Bioenergy Research: Advances and Applications http://dx. doi. org/10.1016/B978-0-444-59561-4.00004’8

invariably required to render the cellulose amenable to enzymatic hydrolysis (Zheng et al., 2009).

The total estimated availability of usable biomass in the world is about 2 billion dry tons per year (Lewis et al.,

2005) . Therefore, the enormous potential of second- generation fuels and the increasing interest toward devel­oping effective, low-cost and environmentally friendly pretreatments for breaking down the close association of the structures of the biomass.

і

Thaddeus Chukwuemeka Ezeji1’*, Nasib Qureshi2, Victor Ujor1

1The Ohio State University, Department of Animal Sciences and Ohio State Agricultural Research and Development Center (OARDC), Wooster, OH, USA, 2United States Department of Agriculture, a National Center for Agricultural Utilization Research, ARS, Bioenergy Research, Peoria, IL, USA

Corresponding author email: ezeji.1@osu. edu

OUTLINE

BACKGROUND/INTRODUCTION

Isobutanol (Inte’mational Union of Pure and Applied Chemistry nomenclature: 2-methylpropan-1-ol) is a branched four-carbon alcohol [(CH3^CHCH2OH], with a boiling point of 108 °C, a melting point of —108 °C, and a relative density of 0.806 at 15 °C (Budavari,

1996) . It is also known as isobutyl alcohol or 2-methyl-

1- propanol. Isobutanol has a vapor pressure of 10.43 mm Hg or 13.9 hPa at 25 °C (Daubert and Danner, 1985) and a water solubility of 85.0 g/l at 25 °C (Valvani et al., 1981). These properties reveal that isobutanol is lighter than, and also soluble in water. While isobutanol is produced industrially via carbonylation (incorpora­tion of carbon monoxide into organic/inorganic com­pounds) of propylene, isobutanol can be produced biologically via fermentation of glucose with a potential to use lignocellulosic biomass. Isobutanol is naturally
produced in low amounts by Saccharomyces cerevisiae as a degradation product of valine. The first report of biolog­ical production of isobutanol was by Dickinson et al. (1998) who demonstrated that S. cerevisiae was able to produce isobutanol using 13C-labeled valine as substrate. It was hypothesized that the product of valine transami­nation, a-ketoisovalerate, had four potential routes to isobutanol, which include (1) catalysis of a-ketoisovalerate by branched-chain a-keto acid dehy­drogenase to produce isobutyryl-CoA and subsequently isobutanol; (2) catalysis of a-ketoisovalerate to isobutanol by pyruvate decarboxylase (PDC); (3) reduction of a-ketoisovalerate to a-hydroxyisovalerate by a-ketoiso — valerate reductase; and (4) use of the PDC-like enzyme encoded by YDL080c to produce isobutanol. Given the fact that riddance of branched-chain a-keto acid dehydro­genase activity in an lpd1 disruption mutant did not — prevent the formation of isobutanol, S. cerevisiae cell

a Mention of trade names or commercial products in this article is solely for the purpose of providing scientific information and does not imply recommendation or endorsement by the United States Department of Agriculture. USDA is an equal opportunity provider and employer.

Bioenergy Research: Advances and Applications http://dx. doi. org/10.1016/B978-0-444-59561-4.00007-3

homogenates could not convert a-hydroxyisovalerate to isobutanol, and a strain with a disrupted PDC-like gene, YDL080c, produced wild-type levels of isobutanol, hence, routes 1, 3, and 4 were eliminated in S. cerevisiae. Notably, elimination of PDC activity in a pdc1 pdc5 pdc6 triple mutant abolished isobutanol production thus, buttressing the notion that this route is the right channel to isobutanol biosynthesis.

As a feedstock chemical, isobutanol is used for the production of isobutyl acetate, which is subsequently used for the production of lacquers. It also finds use as a direct solvent, production of amino resins, isobutyl amines, and acrylate and methyl acrylate esters. The largest use for isobutanol is for the production of zinc dialkyldithiophosphates, an additive for lube oils, greases and hydraulic fluids, in which it functions as an antiwear and antioxidant additive. The second most significant applications of isobutanol are in the produc­tion of isobutyl acetate and as a solvent, primarily for surface coatings and adhesives (Bizziari et al., 2002).

Recent advances in liquid biofuel technology (Mariano et al., 2011,2012; Atsumi et al., 2008), depletion of petroleum reserves, global population growth, envi­ronmental and energy security concerns, have revived research efforts aimed at producing environmentally friendly liquid fuel chemicals. Indeed, global population is projected to reach 8.92 American billion by 2050 and world energy use may increase 53% by 2035. Conse­quently, there is an exigent need to source for or develop new fuels to fill potential shortfalls and, possibly replace our fast depleting petroleum reserves. Between 1980 and 2010, efforts have been focused on engineering microor­ganisms to make production of ethanol from biomass more efficient for use as a biofuel. Compared to ethanol, longer chain alcohols (e. g. n-propanol, n-butanol and isobutanol) have greater energy content, lower vapor pressure, and lower hygroscopicity, which make them superior alternatives to ethanol as a biofuel (Ladisch, 1991; Ezeji et al., 2005).

Isobutanol has the potential to substitute gasoline or serve as a gasoline supplement and can be produced from domestically abundant biomass sources including lignocellulosic biomass. Lignocellulosic biomass, which may contain xylan, arabinan, galactan, glucuronic, ace­tic, ferulic, and coumaric acids, is the most abundant renewable resource on the planet (Koukiekolo et al.,

2005) and has great potential as a substrate for isobuta­nol production (Higashide et al., 2011). Substrate cost has long been recognized to have significant influence on biofuel price and has been identified as a major factor affecting economic viability of n-butanol production by fermentation (Qureshi and Blaschek, 2000). Production of isobutanol from low-cost lignocellulosic biomass which does not compete with food crops may be critical for cost-effective fermentative production of isobutanol.

Whereas majority of producing microorganisms including S. cerevisiae use glucose as preferred substrate for growth and alcohol production, recent advances in genetic engineering have made it possible to metaboli — cally engineer these microorganisms to expand their substrate range to include pentose sugar components of lignocellulosic biomass hydrolysates such as xylose and arabinose, as in the case of ethanologenic microor­ganisms such as Escherichia coli (Dien et al., 1999, 2000), Zymomonas mobilis (Zhang et al., 1995; Deanda et al., 1996), and S. cerevisiae (Jin et al., 2005; Wisselink et al., 2007; Garcia Sanchez et al., 2010). This chapter describes the biochemistry of isobutanol production from biomass, latest developments in isobutanol production technology, and efforts directed toward development of more efficient and cost-effective processes for isobuta­nol production from biomass.

Biodiesel is a proven fuel. The conversion of vege­table oil to biodiesel was first described as early as 1853 by Patrick Duffy, many years before the first diesel engine became functional (Duffy, 1853). Rudolf Diesel’s engine was built several years later, running for the first time on August 10, 1893 using nothing but peanut oil feedstock. In a 1912 speech, Diesel said, "the use of vege­table oils for engine fuels may seem insignificant today but such oils may become, in the course of time, as important as petroleum and the coal-tar products of the present time."

Fossil fuel-derived petrodiesel is produced from the fractional distillation of fossil fuel crude oil. It contains ~75% saturated hydrocarbons and 25% aromatic hy­drocarbons (including naphthalenes and alkylben — zenes). Compared to petrodiesel, biodiesel molecules are comprised almost entirely FAME saturated, or monosaturated, hydrocarbons and ~ 5% aromatic com­pounds. Table 10.2 shows a comparison between the properties of biodiesel to petrodiesel. Biodiesel has a higher lubricity and thus better lubricating properties

than fossil diesel, which reduces wear on fuel systems and engine components. Biodiesel likewise has higher cetane ratings than today’s lower sulfur diesel fuels. The cetane number is a measure of a fuel’s ignition delay, or the time period between the start of injection and the first identifiable pressure increase during com­bustion of the fuel; the higher the cetane number the more easily the fuel will combust. Therefore higher ce­tane biodiesel should cause an engine to run more smoothly and quietly. Biodiesel’s higher flash point makes biofuel vehicles much safer in accidents than those powered by petrodiesel or gasoline. Biodiesel is biodegradable and nontoxic and also contains little to no sulfur, which makes it a much cleaner burning fuel compared to petrodiesel (Hai et al., 2000; Anderson et al., 2002; Hoekema et al., 2002; Choi et al., 2003; Grima et al., 2003; Zijffers et al., 2008; Brindley et al., 2011).

Biodiesel has higher oxygen content than petrodie — sel, which can also reduce pollution emissions. How­ever, this benefit is offset by the fact that biodiesel is more likely to oxidize (react with oxygen), producing contaminants (gumming/sludge) that will plug fuel filters, leave deposits on injectors and cause injector pump problems. Further, continuous oxidization leads to the fuel becoming more acidic, which in turn causes corrosion on the components in the injection system. It will also dissolve fossil-diesel sludge built up over time and send it through fuel lines, plugging fuel fil­ters. Biodiesel cloud or gel point is higher than pump diesel, meaning that it tends to gel at low tem­peratures more readily which can lead to poor cold starting. Clearly, there are both benefits and draw­backs for using biodiesel in today’s automobile engines.

BIOETHANOL

First-generation bioethanol is usually produced by alcoholic fermentation of starch (e. g. corn and wheat) or sugar (e. g. sugarcane, sugar beet and sweet sorghum). Second-generation bioethanol feedstocks include ligno- cellulosic grasses, woody biomass, and algae. Bioethanol is an already well-established fuel in Brazil and the USA (Goldemberg, 2007). Owing to mandates enacted by the Brazilian government in 1976, all light-duty fleet vehi­cles are required to operate using a blend of gasoline and bioethanol fluctuating between 10% and 25%, or E10—E25. In 2003, the Brazilian car manufacturing in­dustry introduced flexible-fuel vehicles that can run on any proportion of gasoline (E20—E25 blend) and hy­drous ethanol (E100) (Horta Nogueira, 2004). Sales reached an impressive 92.3% share of all new cars and light-vehicle sales for 2009, and overall bioethanol pro­duction reached 5.5 billion U. S. liquid gallons.

Although the vast majority of bioethanol is produced by fermentation of corn glucose in the United States or sugarcane sucrose in Brazil (Rosillo-Calle and Cortez, 1998), bioethanol can be derived from any material that contains sugars, including microalgae. Unlike land-based food crops, the production of bioethanol from microalgae does not divert agricultural foods away from grocer’s shelves. This is especially true for corn and corn products, which serve as base ingredients of many processed foods. Further, microalgae can be cultivated in areas nonsuitable for traditional agricul­tural crops and can be harvested many times a year. Therefore, in the U. S., microalgae are generally thought to be the only practical alternative to current bioethanol crops such as corn and soybean (Chisti, 2007; Hu et al., 2008; Singh and Gu, 2010).

Matsumoto et al. (2003) screened several strains of marine microalgae with high-carbohydrate content and identified a total of 76 strains with a carbohydrate con­tent ranging from 33% to 53% . It has been estimated that approximately 46—140 kl of ethanol/ha year can be produced from microalgae (Mussatto, 2010). This yield is several orders of magnitude higher than yields obtained from other bioethanol feedstocks (Table 10.3).

W. J. Oosterkamp

Oosterkamp Oosterbeek Octooien, The Netherlands
email: willemjan@oosterkamp. org

OUTLINE

Introduction 204

Biodegradability 204

Addition of Macro — and

Micronutrients 204

Addition of Microbes 205

Addition of Enzymes 206

Pretreatments 207

Biological Pretreatment with

Enzymes 207

Chemical Pretreatment 207

Hot Water Treatment 207

Mechanical Pretreatment 207

Longer Retention Times 207

Energy Crops 207

Food Processing Residues 207

Rice Husks 207

Bagasse 207

Coffee Husks and Mucilage 208

Crop Residues 209

Spent Bedding 209

Kitchen and Garden Waste 209

Aquatic Weeds 209

Digestion Systems 211

Family-Size Biogas Plant 211

Wet Digesters 211

Scum Layer Digester 211

Solid Biomass Digester 212

Increase in Solids Content in Wet Digesters 212

Loading and Unloading of Digesters 212

Treatment of Digestate in Wet Digesters 212

Use of Methane 213

Chemical Conversion of Volatile Solids 213

Combustion 213

Gasification 213

Thermal Conversion of Volatile Solids 214

Slow Pyrolysis 214

Flash Pyrolysis 214

Discussion 214

Maximum Methane Yield 214

Nutrient Recycling 214

Soil Fertility 214

Digesters 214

Conclusions 214

References 215

Bioenergy Research: Advances and Applications http://dx. doi. org/10.1016/B978-0-444-59561-4.00013-9

INTRODUCTION

All-renewable energy resources are required to reduce dependency on fossil fuels from politically un­stable regions. Biomass is one such renewable energy resource. Farm and food processing residues are preferred but, where economic, energy plants can be used.

Biomass as such cannot replace fossil fuels. Such materials have to be converted into gas, liquid or elec­tricity. Biological volatilizing (anaerobic digestion) con­verts organic by-products and residues into methane and carbon dioxide, an energy source that can be used for cooking, the production of electricity and as trans­portation fuel.

In Asia there are over 10 million family-size anaerobic digestion plants utilizing manure and some straw. The biogas is used for cooking. There are significant health advantages in using biogas, compared to the local alter­native of the burning of cattle manure, leaves and wood inside the houses.

There are a few thousand centralized biogas plants in Europe that use manure with a whole range of easily digestible residues. Other biogas plants in Europe use sludge from wastewater cleanup plants. They convert the biogas into electricity and heat. Carbon dioxide is removed from the biogas in a number of recent plants; the gas is compressed and injected into the natural gas grid.

The digestate, after the production of biogas, should be used as an organic fertilizer. This will recycle the macro elements nitrogen, potassium, phosphorus and carbon to the soil. Recycling of carbon is essential for high soil productivity and will reverse the trend of lowering of crop yields (Hossain, 2001).

The energy content of the animal residues (mostly manure) produced worldwide is equivalent to an average power of 50—150 W per person (9—25EJ/a). The energy content of crop residues (mostly straw) is also 50—150 W per person (Hoogwijk et al., 2003). Worldwide energy consumption is 2.5 kW per person (500 EJ/a). Oil production worldwide is 1 kW per per­son (80 million barrels a day). Biogas from straw and manure can replace about 10—30% of the world oil pro­duction. This substitution can be doubled by the use of forest residues.

BIODEGRADABILITY

only part of it can be depolymerized into soluble com­ponents. Anaerobic digestion is a complex process that is slow compared to chemical processes. Chynoweth et al. (1987) have published on the processes involved in the anaerobic digestion of biomass. Hydrolytic bacte­ria break down the cellulose and hemicellulose into organic acids and neutral compounds. Hydrogen pro­ducing bacteria convert the acids into hydrogen. Homoacetogenic bacteria convert hydrogen into acetic acid. Methanogenic bacteria convert acetic acid into methane. A by-product in these conversions is carbon dioxide.

Anaerobic biodegradation potential assay is per­formed by mixing the material with digestate from an operating digester or by mixing the material with a defined nutrient medium according to Owen et al. (1979). The methane produced is measured at different times.

Chandler et al. (1980) made a correlation based on 15 different lingocellulosic materials.

yCH4 = a * (b — c * Zj) (13.1)

where

yCH4 is the methane yield in l/kg volatile solids (VS) a = 440 l/kg is the conversion between methane yield and VS reduction (Jerger et al., 1982). b = 0.83 fitted constant. c = 2.8 fitted constant. li is lignin fraction of the VS

This correlation gives a standard deviation of 80 l/kg VS for straws and woody biomass (Table 13.1).

A different correlation was developed for straws and woody biomass.

y CH4 = a * (1 — Zi) * (1 — e—dt) (13.2)

d = f * (1 — g * li) is exponential factor. f = 0.025 fitted constant. g = 3 fitted constant.

This correlation assumes that biodegradation can be described as a first-order process. Shielding of cellulose and hemicellulose by lignin is reflected in the exponen­tial factor. This shielding eventually breaks down. This correlation performs better with a standard deviation of 32 l/kg VS.

This section describes the application of functional­ized polymers as catalysts for esterification and transes­terification reactions to produce biodiesel. Polymeric catalysts consist of functionalized polymeric matrixes or polymeric matrixes that can be used as solid supports for a variety of catalysts, constituting catalytic systems (Coutinho et al., 2004a, b; Guerreiro et al., 2010; Lee and Saka, 2010; Zieba et al., 2010). These materials have long been studied as heterogeneous catalysts in systems that traditionally employ acid or basic homoge­neous catalysts.

Biodiesel production can be carried out in the pres­ence of different types of catalysts. In the specific case of polymeric catalysts belonging to the class of function­alized polymers, their use in biodiesel synthesis has focused on acid catalysts, such as in the case of ion exchange resins (Ma and Hanna, 1999; Guerreiro et al., 2006; Knothe et al., 2006; Soldi et al., 2009; Rezende et al., 2008; Helwani et al., 2009b; Lee and Saka, 2010).

Acid-catalyzed triacylglycerol transesterification is not commercially applied as often as catalysis in basic medium because acid catalysis in homogeneous me­dium is around 4000 times slower than the base- catalyzed reaction. However, acid catalysts can perform esterification and transesterification simultaneously, producing biodiesel directly from oils with high acid number. These oils are not suitable for biodiesel produc­tion via alkaline catalysis, since the FFAs promptly react with the base, generating soaps that make the separation between the ester and glycerin difficult during the washing step (Bondioli et al., 1995; Schuchardt et al., 1998; Ma and Hanna, 1999; Vicente et al., 2004; Lotero et al., 2005; Meher et al., 2006; Rezende et al., 2008; Lee and Saka, 2010).

Several acid catalysts can be used in alcoholysis, espe­cially sulfonic and sulfuric acids (Hayyan et al., 2011). Although these catalysts afford high yields of mono alkyl esters, they require high temperatures and long reaction times to achieve a satisfactory conversion rate. Another disadvantage is that residual acid cata­lysts can contaminate the fuel and attack the metal parts of the engine, corroding it. To avoid this situation, acid catalysts must be completely eliminated from the final product, which demands many purification steps (Canakci and Gerpen, 1999).

Some types of organic polymers and ion exchange resins can be used as polymeric catalysts, behaving as heterogeneous catalysts for esterification and transester­ification reactions. Heterogeneous catalysts such as these reduce the number of biodiesel purification steps, facilitates catalyst reuse, and decreases production costs (Schuchardt et al., 1998; Choudary et al., 2000; Fukuda et al., 2001; Harmer and Sun, 2001; Ramos et al., 2003; Abreu et al., 2004; Suppes et al., 2004; Chouhan and Sarma, 2011).

The pioneering studies of Merrifield on the solid — phase synthesis of polypeptides and subsequent works have shown that polystyrene (PS) is a suitable polymeric support for catalysts and reagents (Merrifield, 1963; Frechet, 1981). Styrene and divinylbenzene (DVB) are among the monomers that are most often employed to prepare solid polymeric matrixes. Polystyrene — co-divinylbenzene (PS-DVB), an insoluble copolymer, results from styrene polymerization in the presence of varied amounts of DVB. The characteristics of this copolymer depend on the quantity of DVB present in the material (Kapura and Gates, 1973; Xia et al., 2012).

In many cases, the success of a heterogeneous catalyst relies on the features of the polymeric material. Bergbreiter (2002) proposed that some physicochemical properties should be considered when choosing the catalyst support, including the catalytic activity, surface area, porosity, and thermal and mechanical stability of the material in the conditions of the catalyzed reaction (Bergbreiter, 2002; Chouhan and Sarma, 2011).

In the field of polymer chemistry, the term resin is indistinctly employed to describe polymers with and without cross-links (Akelah and Sherrington, 1981; Sharma, 1995). Alternatively, in heterogeneous catalysis, the term resin refers to species consisting of long polymeric chains interconnected via cross-links, the so-called polymeric matrix. Polymeric matrixes are
tridimensional, insoluble, and porous; their ability to ex­change ions arises from the introduction of adequate functional groups into the polymeric support (Kunin et al., 1962; Akelah and Sherrington, 1981; Frechet, 1981; Harmer and Sun, 2001; Hart et al., 2002).

Polymeric matrix functionalization can be achieved in two ways: (1) monomers containing the desired func­tional groups (or precursors of this functional group) can be directly polymerized; (2) the polymeric support can be prepared first, and the functional group is intro­duced by chemical modification of the polymeric sup­port (Molinari et al., 1979; Kucera and Jancar, 1998; Harmer and Sun, 2001).

Coutinho and Rezende (2001) and Coutinho et al. (2004a) showed that supported species can be prepared by chemically modifying the copolymer base (polymeric support). These authors reported the sulfonation of a sup­port consisting of PS reticulated with DVB (Figure 16.7). The aromatic rings on the insoluble PS-DVB copolymer react with concentrated sulfuric acid in the presence of

1,2- dichloroethane; the latter compound expands the polymeric matrix and allows sulfonation of the internal surface as well. Most of the functional groups introduced into polymeric matrices concentrate inside the resin beads (Coutinho and Rezende, 2001).

Cationic resins can be used as an option for catalytic reactions involving mineral or sulfonic acids. In the presence of water, the cationic groups on the polymer display different acidity constants, as in the case of com­pounds with low molecular mass.

The catalytic performance of an ion exchange resin is associated with the concentration of functional groups and the physicochemical properties of the support. Therefore, compared with homogeneous catalysts, different factors affect the activity of resins.

The use of ion exchange resins as catalysts has many advantages: (1) despite being equivalent to strong

——— CH2-CH-CH2-CH-CH2——————-

——— CH2—CH—CH2-CH—CH2————

FIGURE 16.7 Sulfonation of a polymeric matrix consisting of polystyrene and divinylbenzene.

FIGURE 16.8 PS sulfonation with acetyl sulfate.

mineral acids, resins are less oxidizing and corrosive, since most of the catalytic sites are located inside the beads—therefore, they do not pose any hazards to the operator and are easy to store; (2) resins behave as selec­tive catalysts and enable reaction control; (3) catalysts with high purity are recovered at the end of the reaction by simple filtration; (4) resins do not require neutraliza­tion before being separated from the reaction medium, a step that usually reduces product yield; (5) resins elimi­nate the steps and equipment necessary to separate the catalyst and purify the product, simplifying continuous or batch procedures based on ion exchange resins; and

(6) if the resins undergo deactivation due to contamina­tion or prolonged use, they can be reactivated via a sim­ple procedure that does not release hazardous gases into the atmosphere (Saha and Sharma, 1996; Coutinho and Rezende, 2001; Harmer and Sun, 2001; Marquardt and Eifler-Lima, 2001; Mitsutani, 2002; Coutinho et al., 2003, 2004a, b; Kiss et al., 2006).

The main drawback of ion exchange resins is that their maximum operation temperature is relatively low. Literature suggests that they should be used below 125 °C to ensure long catalyst lifetime (John and Israel — stam, 1960; Akelah and Sherrington, 1981; Gimenez et al., 1987; Coutinho and Rezende, 2001; Rezende et al., 2008).

Aromatic compounds are easy to functionalize, espe­cially if they contain acid groups like sulfonic acids. The sulfonation of organic compounds containing benzene rings, including polymers, has been extensively re­ported (Ma and Hanna, 1999; Coutinho and Rezende, 2001; Coutinho et al., 2003, 2004a, b, 2006; Rezende et al., 2008; Soldi et al., 2009). Figure 16.8 represents the sulfonation PS (Soldi et al., 2009).

Sulfonation significantly modifies the physical prop­erties of linear PS, especially the polarity. Hence, sulfo — nated PS should remain insoluble during biodiesel production. Soldi et al. (2009) studied methods to sulfo­nate linear PS and applied the resulting sulfonated material as heterogeneous polymeric catalyst to produce soybean methyl esters. Raw materials with different moisture degrees and the effect of different variables on the conversion rate have been investigated; biodiesel production from soybean oil and beef tallow led to significantly improved yields.

Recently, much interest has been taken in utilizing low-cost plant oil and fat containing a large amount of

FFAs. However, oils with high FFA content are difficult to transesterify using the commercially available alka­line catalyst (Zhang et al., 2003; Tesser et al., 2005; Marchetti and Errazu, 2008; Sharma et al., 2008; Liu et al., 2009; Tesser et al., 2010; Chouhan and Sarma, 2011; Shahid and Jamal, 2011; Borges and Diaz, 2012). Canakci and Van Gerpen (1999, 2001) found the transes­terification would not occur if the FFA content in the oil was beyond 3%. According to the research paper by Kouzu et al. (2011), the promising approach is to esterify FFA into FAMEs with the help of the solid acid catalyst, and there were some research papers studying utili­zation of several types of heterogeneous catalysts including sulfonated cation exchange resin (Russnueldt and Hoelderich, 2009; Tesser et al., 2010; Kouzu et al.,

2011) . With respect to utilization of the sulfonated resin for the preesterification of FFA, some researchers focused their attention on the macroreticular type but the use of two types of resins (macroreticular and gelular types) were also studied by other authors (Ramadhas et al., 2005; Soldi et al., 2009; Lam et al., 2010; Melero et al., 2010; Kouzu et al., 2011; Semwal et al., 2011; Li et al., 2012; Xia et al., 2012; Zhang et al., 2012a, b).

Feng et al. (2011) investigated the continuous esterifi­cation of FFAs from acidified oil with methanol by cation exchange resin in a fixed bed reactor to prepare biodiesel and the operational stability of continuous esterification by resin in the fixed bed reactor was also conducted (McNeff et al., 2008; Shibasaki-Kitagawa et al., 2010; Feng et al., 2011; Cheng et al., 2012).

According to Feng et al. (2010), from the viewpoint of cost savings, the use of cation exchange resins in hetero­geneous catalytic processes may be advantageous over enzymes and supercritical methanol (Feng et al., 2010). These resins are composed of copolymers of DVB and styrene containing sulfonic acid groups attached to ben­zene rings and these are the active sites for esterification and transesterification (Marchetti and Errazu, 2008; Rezende et al., 2008; Russnueldt and Hoelderich, 2009; Feng et al., 2010; Kouzu et al., 2011). However, other sul­fonated polymeric backbones such as Amberlyst®, Dow — ex® and Nafion®, a perfluorinated ion exchange resin, all of them having a very strong Bransted acid character, have also been used in these type of reactions (Ozbay et al., 2008; Talukder et al., 2009; Feng et al., 2010; Park et al., 2010; Galia et al., 2011; Yin et al., 2012; Zhang et al., 2012a). In general, cation exchange resins are

preferable for esterification (Gimenez et al., 1987; Chen et al., 1999; Coutinho et al., 2004b; Coutinho et al., 2006; Grob and Hasse, 2006), while anionic resins may be applied for transesterification of oils and fats (Shiba — saki-Kitakawa et al., 2007; Ren et al., 2012).

CONCLUDING REMARKS

Truly heterogeneous catalytic processes are attractive for many practical applications due to their recyclability, structural stability, high selectivity and good catalytic performance. However, all these properties are hardly achievable in a single catalytic system. In most cases, leachable catalytic species migrate to the reaction envi­ronment, causing a partial contamination of the final product as well as a loss in catalytic activity when the solids are applied in several consecutive reaction cycles. Moreover, in many situations found in the literature, the heterogeneity of catalytic systems is not approached with proper analytical methods, resulting in wrong conclusions and/or classification of the proposed solid catalyst. These usually arise from poor data on catalyst recovery and reuse, poor characterization of the catalyst structure and high leaching levels of catalytic species. Also, in many cases, no attempt is made to fully characterize these properties and solids are classified as heterogeneous catalysts just because they are partially filterable after reaction completion. One of such flaws was nicely demonstrated by Silva et al. (2013) using bismuth-containing mixed oxides. Apart from these ob­servations, the lack of suitable reaction controls such as in the case of TC in esterification reactions reveal unrealistic catalytic performance in reactions that are known to be autocatalytic under appropriate experi­mental conditions. Nevertheless, a number of rather attractive heterogeneous catalytic systems have been discovered so far for biodiesel applications, probably due to the wide scope of catalytic properties that are influential in both esterification and transesterification. However, many of these will never be able to reach in­dustrial applications because their benchmarking was never strong enough to support further investments at large scale.

There is a rather good correlation between different analytical methods used for the analysis of native lignin preparations, both in intralab (Evtuguin et al., 2001) and in interlab studies (Sakakibara, 1991; Capanema et al., 2004; Zhang and Gellerstedt, 2007; Balakshin et al.,

2008) . An exception is the 31P NMR analytical methodol­ogy of native lignins which yielded ca. 30% lower numbers for aliphatic hydroxyls (Pu et al., 2011) compared to other methods (Sakakibara, 1991; Balakshin et al., 2008).

A rather good correlation between various methods for technical lignins analysis has been also reported (Faix et al., 1994; Cateto et al., 2008). However, a compre­hensive review of published analytical data leads to a much less optimistic conclusion. Table 18.3 shows that significant variability in the structure of the same tech­nical lignins can be observed when these lignins are analyzed by independent methods, in contrast to what is seen with the analysis of native lignin preparations. This deviation might be caused by the interference of specific lignin moieties generated during the technical process on the results of each specific analytical method which, as a rule, was developed and validated for the analysis of native lignin preparations.

Various wet chemistry techniques for the analysis of lignin functional groups have been comprehensively reviewed previously (Lin and Dence, 1992; Zakis, 1994). Therefore, we will focus our discussion on major NMR spectroscopic techniques for the analysis of technical lig­nins, 31P and 13C NMR, as well as advanced NMR methods, which have received less attention.

As 31P NMR spectroscopy of derivatized lignins be­comes one of the most common techniques for lignin anal­ysis, it is important to critically evaluate the method in a comprehensive manner. There are two main

modifications of the 31P NMR lignin analysis. Originally,

2- chloro-1,3,2-dioxaphospholane (31P-I method) was sug­gested (Archipov et al., 1991) as the derivatizing agent in this method. Later, 2-chloro-4,4,5,5-tetramethyl-

1,3,2- dioxaphospholane (31P-II) was reported

(Kostukevich et al., 1993; Granata and Argyropoulos, 1995) to provide better signal separation and it is currently used as the major 31P method for lignin analysis. Although good agreement has been reported between the results obtained with these two derivatization re­agents (Granata and Argyropoulos, 1995), data reported later did not confirm this observation (Tables 18.3 and

18.4) . The results obtained with the 31P-II method tend to underestimate results compared to the data generated by all other analytical methods. The 31P-I method tends to report significantly higher amount of aliphatic and total hydroxyl groups if compared to the data obtained by the 31P-II methodology (Table 18.3), in contrast to the conclu­sions drawn in the original validation work (Granata and Argyropoulos, 1995).

The significantly lower numbers reported by the 31P-II NMR analysis, as it is compared to other analytical methods, in the structural analysis of lignins might be explained by the incomplete lignin derivatization with the phosphorylating reagent PR-II possibly due to steric hindrance of the bulky reagent containing four t-butyl groups. The use of PR-I yields apparently more quanti­tative results. However, the signal resolution in 31P-I is not high enough as it can clearly be seen in the publica­tion by Akim et al. (2001). In this publication, the signals of primary hydroxyls and 5-substituted phenolic hy­droxyls are heavily overlapped. The main conclusion derived from this observation is that even when the res­olution of a resonance signal is formally acceptable (Argyropoulos, 1994), one cannot conclude that the resolved signals are reporting the correct values.