Depolymerization of Alcell lignin using Lewis acid catalysts NiCl2 or FeCl3 yielded gas, solid and liquid products including the formation of ether-soluble mono­mers under different reaction conditions. Both catalysts favor condensation reactions leading to insoluble resi­dues. The low yields of organic monomers were domi­nated by phenolics over ketones and aldehydes (Hepditch and Thring, 2000).

Pyrolysis

Pyrolysis of isolated lignins gives a different product distribution than pyrolysis of wood of other lignocellu — losic materials. Lignin pyrolysis occurs in a wider temperature range (e. g. 160—900 °C) compared to poly­saccharides (e. g. 220—400 °C) (Yang et al., 2007). Further­more, the amount of char from isolated lignins is significantly higher compared to whole biomass pyrolysis. Solid acid catalysts such as H-Zeolite Socony Mobil-5 can effectively shift the products toward more deoxygenated compounds. Different isolated lignins py — rolyzed at temperature ranges of 500—800 °C yielded bio-oil, gas and char of 16—70%, 3—39%, and 17—81%, respectively (Azadi et al., 2013). Several researchers showed that inorganic alkaline catalysts such as NaOH can facilitate depolymerization of lignin by pyrolysis and influence the product composition (Amen-Chen et al., 2001).

Recently, an international study of fast pyrolysis of lignin was undertaken with contribution from 14 labora­tories. Based on the results it was concluded that an impure lignin containing up to 50% carbohydrates behaves like whole biomass, while a purified lignin was difficult to process in the fast pyrolysis reactors and produced a much lower amount of a more enriched aromatic bio-oil. It was concluded that for highly pure lignin feedstocks new reactor designs will be required other than the typical fluidized bed fast pyrolysis sys­tems (Nowakowski et al., 2010).

Upgrading of lignin pyrolysis oil by catalytic hydro­deoxygenation (HDO) is often used as described by de Wild et al. (2009). More stable oil due to partial removal of oxygen is an important upgrading property. Bu et al.

(2012) made a review on the catalytic HDO upgrading of lignin-derived phenols from biomass pyrolysis. This study shows that further investigation of HDO is needed to improve catalysts and optimize operation conditions, further understanding of kinetics of complex bio-oils, and availability of sustainable and cost-effective hydrogen sources. Further HDO treatments are dis­cussed in the next session.

Anellotech (2010) has developed a technology plat­form using catalytic pyrolysis for the claimed inexpen­sive production of chemicals and transportation fuels from nonfood biomass. Vispute et al. (2010) claim that all chemical conversions can be performed in one reactor, using an inexpensive catalyst. Target green chemicals are BTX.

Hanshu Ding*, Feng Xu

Department of Protein Chemistry, Novozymes Inc., Davis, California, USA Corresponding author email: hdin@novozymes. com; fxu@novozymes. com

OUTLINE

Industrial Phytochemicals 353

Overview 353

Colorants (Pigment, Dye, and Ink) 355

Dietary, Nutraceutical, Food or Feed Additives 356

Bioactive or Pharmaceutical Phytochemicals 356

Phytochemicals for Personal Care or Other Uses 358

Production of Industrial Phytochemicals 358

Extraction and Isolation from Specific Plants 358

Coproduction from Processing (Biorefinery) of Staple Crops 358

Production from Cultured Plant Cells 360

Production from Microbial Fermentation 360

Production from Algae via Aquaculture 361

Coproduction of Phytochemicals in Bioenergy

Processes 361

Coproduction from Starch — or Sugar-Based Bioenergy

Processes 361

Coproduction from Plant Oil-Based Bioenergy Processes 361

Coproduction from Lignocellulose (Biomass)-Based Bioenergy Processes 362

Coproduction from Bio-Oil, Syn-Gas, or Algal Bioenergy Processes 362

Colocation of Fermentative Phytochemicals Production with Bioenergy Processes 363

Utilization of Phytochemical Production By-Products for Bioenergy 363

References 363

INDUSTRIAL PHYTOCHEMICALS

Overview

Phytochemicals may be defined as chemicals derived or derivable from plants. Phytochemical sources may include not only unprocessed trees, crops, grains, fruits, nuts, vegetables and legumes, but also processed plant-derived materials such as starch, sugars and oil. Phytochemicals of commercial interest have demon­strated or suspected utilities for dietary, bioactive, therapeutic and industrial technical uses. Based on chemical structure, major groups of phytochemicals include (Figure 20.1) the following:

1. Carotenoids (carotenes or xanthophylls, e. g. a- or b-carotene, b-cryptoxanthin, lycopene, lutein, and zeaxanthin) and homologs (e. g. crocin)

2. Flavonoids (anthocyanins, flavanols, flavanones, flavonols, flavones, and isoflavones), and condensed tannin and xanthones

3. Other phenolic/quinonics (e. g. tocopherols, curcumin, resveratrol, carminic acids, alizarin, purpurin, lignans (dimeric phenyl propanoid), tannic acid, thymol, and capsaicin)

4. Alkaloids (e. g. caffeine, nicotine, quinine, vinblastine, and opiates) and other N-contained compounds (e. g. chlorophylls, flavins, betalain, indole-3-carbinol, galanthamine, and indigo)

Bioenergy Research: Advances and Applications http://dx. doi. org/10.1016/B978-0-444-59561-4.00020-6

FIGURE 20.1 Major groups of phytochemicals. Source: Drawings from ChemSpider and Sigmaaldrich. com are used in this and Figures 20.2—20.4. (For color version of this figure, the reader is referred to the online version of this book.)

5. S-contained compounds (e. g. g-glutamylcysteines, (allyl)cysteine sulfoxides, and isothiocyanates)

6. Phytosterols (e. g. sitosterol, stigmasterol, campesterol or 4-desmethyl sterols), saponin, digoxin, and other terpenoids (e. g. artemisinin, paclitaxel, and camphor)

7. Polymeric carbohydrates (e. g. cellulose, hemicellulose, b-glucan, pectin, gum, inulin, and resistance starch), oligosaccharides (e. g. oligofructose), and lignin

8. Lipids and volatiles such as lecithin, essential oils, and menthol

9. Proteases such as bromelain and papain, as well as protease inhibitors

It is reported that about 8000 phenolics (including ~4000 flavonoids), ~ 20,000 terpenoids, ~ 10,000 alka­loids, ~700 carotenoids, and ~250 phytosterols are known, and many have been shown with various

functions (Watkins and Chaudhry, 2013). Comprehen­sive studies have been carried out on phytochemicals from whole grains, fruits and vegetables (Liu, 2007; Piironen et al., 2000).

Phytochemicals may have different physical, chemical and biological properties, thus suitable for different industrial usages as colorants (pigment, dye, and ink), dietary food/feed additives or nutraceuticals, bioactive/ pharmaceutical ingredients, personal care (cosmetic, perfume) agents, or other useful materials. For instance, carotenoids, polyphenols, flavonoids and tocopherols may be used as antioxidant or antiinflammatory agents; alkaloids may be used as analgesic, antispasmodic or mental disorder-relieving agents; and carotenoids may be used as coloring agents.

Phytochemicals are of great interest for industrial, technical, household, health care or other uses, due to their renewability, performance, safety, environment — friendliness, and diversity in structure and activity. The use of phytochemicals started at the dawn of humanity, has contributed to the civilization, and is reemerging along with the advancement of bioenergy, biobased chemicals, and biorefinery.

Colorants (Pigment, Dye, and Ink)

Many phytochemicals are chromophoric, reflecting lights that cover the visible wavelength range. Ubiqui­tous phytocolorants include chlorophyll (green) and carotenoids (yellow-red) from leaves and stems of plants, while more specific colorants may exist in flowers, fruits or other parts of plants (Figure 20.2). Colorants could also be produced from algae, bacteria, or fungi including the saprophytes (Gupta et al., 2011; Rymbai et al., 2011; Matthews and Wurtzel, 2007; Mortensen, 2006; Dufosse, 2006; Mapari et al., 2005; Adrio and Demain, 2003; Sengupta, 2003).

Colorants are used mostly as dyestuff, food/feed ad­ditives or cosmetic agents. Traditional plant-extracted/ derived dyestuffs include saffron from saffron crocus plant, madder (red) from madder plants (Rubia), and indigo from Indigofera plants (at present chemical synthesis from fossil feedstocks provides most indigo dyes, although microbial route has been explored).

Commonly used food or feed colorants derived from plants include extracts or isolates from specifically grown plants, such as bixin and norbixin (annatto), beta — lains (including betanin), curcumin (turmeric), crocin

(saffron), and carotenoids (including b-carotene, lutein, canthaxanthin and astaxanthin). The colorants also include extracts or isolates from agricultural residues, such as anthocyanins and carotenoids. The colorants may be produced microbially, as exemplified by the carotenoids such as astaxanthin, b-carotene, lutein, and riboflavin (Chattopadhyay et al., 2008).

Some plant-derived colorants, such as lutein and b-carotene, are used as cosmetic agents. In addition to plants, algae also produce colorants of industrial inter­est. For instance, phycobiliproteins have uses as natural dyes, cosmetic agents or food colorants (in addition to health applications) (Spolaore et al., 2006). Phytocolor­ants may also be used for thermoplastic (van den Oever et al., 2004).

Although the heterocyst/nitrogenase-based system has been the most studied, some other known cyanobac — terial hydrogen-producing reactions could poten­tially be used for biological hydrogen production. These include the unicellular and nonheterocystous filamentous cyanobacteria, which possess nitrogenase and are able to fix nitrogen in nature. Two strategies are employed to avoid oxygen inhibition. In some uni­cellular species, oxygen evolution and nitrogen fixation (or hydrogen production) are separated in time since photosynthesis and nitrogen fixation are under circa­dian control with photosynthesis taking place during the day and nitrogen fixation being maximal during the night period. The filamentous cyanobacterium Tri — chodesmium uses a strategy of spatial segregation where nitrogen fixation occurs in cells located in the middle of the bundle carrying out the oxygen-sensitive nitroge- nase reactions and the others carrying out the normal photosynthetic reactions (Berman-Frank, 2001).

The unicellular cyanobacterium Cyanothece has been the subject of a number of recent studies demonstrating prolonged hydrogen production in the light mediated by nitrogenase. In one study, considerable hydrogen production (up to 465 mmol per milligram of chlorophyll per hour) was shown, the growth conditions were very stringent and hydrogen production was only observed when the culture was submitted to nitrogen starvation, sparged with argon to remove any oxygen formed through photosynthesis, supplemented with glycerol and cultivated under low light (Bandyopadhyay et al., 2010; Min and Sherman, 2010). Glycerol, in addition to serving as a possible additional energy source to sup­port nitrogenase activity, appears to release nitrogenase from diurnal control (Aryal et al., 2013). Another recent study found appreciable hydrogen and oxygen produc­tion with nitrogen-depleted cultures that were incu­bated under continuous illumination (Melnicki et al.,

2012) . Light saturation curves and photosynthesis inhi­bition studies indicate that the hydrogen is evolved indi­rectly from the fixed carbon produced through photosynthesis. Here again, the requirements for contin­uous illumination (it can hardly be energetically positive to produce hydrogen using artificial illumination) and for argon sparging raise serious hurdles to practicality. Thus, although a nice proof of principle, such a system would hardly be economically viable.

Many cyanobacteria also possess Hox, a soluble reduced nicotinamide adenine dinucleotide (NADH)- linked [NiFe] hydrogenase. This reversible hydrogenase is capable of hydrogen evolution, in particular when dark-adapted cells are reilluminated (Schwarz et al., 2010). As discussed above, this forms an electron valve, readjusting the poise of the photosynthetic apparatus, but activity is quickly inhibited with renewed oxygen evolution. A recent survey showed that a diversity of cya­nobacteria contains this enzyme and that there is great variability in both the amounts of hydrogen made by this enzyme and the pattern of hydrogen evolution (Kothari et al., 2012). This enzyme is also responsible for evolution during dark fermentation of endogenous reserves, principally glycogen, and hydrogen production by this pathway can be enhanced through lowering of the hydrogen partial pressure (Ananyev et al., 2012). At least in Synechocystis, hydrogen production by Hox can be increased by eliminating the master regulator AbrB2, which normally represses synthesis of Hox (Dutheil et al., 2012; Leplat et al., 2013). In a recent attempt to increase hydrogen production, heterologous expression of the [FeFe] hydrogenase from Clostridium acetobutylicum was carried out in the non-nitrogen-fixing cyanobacte­rium Synechococcus (Ducat et al., 2011). Active hydroge — nase was formed under proper conditions, but in vivo light-driven hydrogen production from this system was significant only when the cultures were incubated under an inert atmosphere and oxygenic photosynthesis was completely inhibited.

Ethanol

While hydrogen production, or at least direct bio­photolysis, can be driven directly by photosynthesis, all other biofuels must use the capacity of cyanobacteria to drive carbon dioxide fixation with photosynthetically derived energy, ATP and reductant. However, once fixed by the Calvin-Benson-Bassham cycle, the newly recycled carbon can be converted to useful biofuels through the introduction of novel (to cyanobacteria) metabolic pathways (Angermayr et al., 2009). The first such cyanobacterial-derived biofuel that was demonstrated was ethanol (Deng and Coleman, 1999; Dexter and Fu, 2009), and its production is the only cyanobacterial — produced biofuel under active investigation and com­mercial development (Algenol Biofuels: http://www. algenolbiofuels. com/). Algenol Biofuels is presently claiming production at "around $1.00 per gallon using sunlight, carbon dioxide and saltwater at production levels above 9000 gallons of ethanol per acre per year". At an average solar insolation for Florida of 19.8 MJ/day and since ethanol has a higher heating value of 29.7 MJ/ kg, this translates to a claim of a very impressive 2.8% conversion efficiency. Now another company, Joule Unlimited (http://www. jouleunlimited. com/), has step­ped into the picture, offering to sell SunFlow-E through its fuel company, Joule Fuels. Their process uses geneti­cally modified thermophilic cyanobacterium containing Moorella alcohol dehydrogenase, and their Web site claims are even more spectacular with targets of up to

25,0 gallons per acre (7.8% conversion efficiency) and $0.60 per gallon at full-scale commercial production.

Cyanobacteria can naturally produce relatively min­ute amounts of ethanol so at the simplest level, creating a cyanobacterium that produces higher levels of ethanol involves boosting flux through the ethanol pathway through the introduction of the key enzymes for conver­sion of pyruvate, generated by glycolysis of the fixed carbon, to ethanol, pyruvate decarboxylase (pdc) and alcohol dehydrogenase (adh). This alone is sufficient to produce low millimolar levels of ethanol in the medium upon prolonged (5—10 days) incubation and growth. Further increases, obviously necessary for practical pro­duction, have been achieved through a variety of means, including better transcriptional control and further metabolic engineering. Most of this development work is being done at private enterprise laboratories, but a recent published report (Gao et al., 2012) shows that impressive increases in yields can be achieved by inte­grating a foreign pdc and a native adh into the genome of Synechocystis and abolishing carbon flux into polyhy — droxybutyrate synthesis.

One of the most promising emerging biorefinery plat­forms is the biochemical path that focuses on fermenta­tion of sugars extracted from lignocellulosic feedstocks (Carvalheiro et al., 2008). This technology involves three basic steps: (1) conversion of biomass to sugar or other fermentation-feedstock, (2) bioconversion of these biomass intermediates using biocatalysts, and (3) pro­cess products to yield added value chemicals, fuel — grade ethanol and other fuels, heat and/or electricity (Carvalheiro et al., 2008).

The first step involves a pretreatment process the goal of which is to alter or remove structural and composi­tional impediments to hydrolysis in order to improve the rate of enzyme hydrolysis and increase yields of fermentable sugars from cellulose or hemicellulose (Mosier et al., 2005). The effectiveness of enzymatic hydrolysis of pretreated lignocellulosic biomass can be significantly enhanced if lignin and its derivatives are removed or effectively modified before adding enzymes because lignin and its derivatives interfere with the path for cellulases action and they are also toxic to microor­ganisms, slowing down enzymatic hydrolysis (Qing et al., 2010; Yang and Wyman, 2008).

The ideal pretreatment process produces a disrupted, hydrated substrate that is easily hydrolyzed but avoids the formation of sugar degradation products and fermentation inhibitors (Agbor et al., 2011). Further­more, there is an overall consensus on the several tech­nical, operational and economical characteristics that a successful pretreatment should accomplish, including

(1) maximize the production of highly digestible solids that enhances sugar yields during enzyme hydrolysis;

(2) avoid the degradation of sugars including those derived from hemicellulose; (3) not require the addition of toxic compounds or minimize their use; (4) fermenta­tion compatibility, minimizing the formation of inhibi­tors for the enzymes or microorganisms in the subsequent steps; (5) effectiveness at low moisture con­tent; (6) broad applicability for multiple crops, sites ages and harvesting times; (7) not required size reduction of biomass; (8) maximize the production of other valuable by-products, e. g. lignin; (9) to be cost-effective by oper­ating in reactors of moderate size, minimizing the heat and power requirements, chemicals and capital equip­ment; and (10) be scalable to industrial size (Alvira et al., 2010; Brodeur et al., 2011; Jorgensen et al., 2007; Yang and Wyman, 2008).

This chapter reviews the advances in the most studied pretreatments and those recently proposed in scheme of the biochemical biorefinery, including kinetics, mecha­nistic and economical models proposed for describing some of these pretreatment processes.

Keto acids are organic acids with ketone functional group on the second carbon, typically known as a-carbon, and are present in microorganisms as intermediate pro­ducts of amino acids production pathways, and degra­dation of biosynthesized amino acids to alcohols is what is commonly known as Ehrlich pathway (Figure 7.1). Different strains of S. cerevisiae and yeasts belonging to genera such as Endomycopsis, Candida, and Hansenula are known to produce higher alcohols via keto acid pathways (Singh and Kunkee, 1976; Cronk et al., 1979).

In addition to ethanol and CO2 production, S. cerevisiae produces a variety of relatively low — molecular weight flavor compounds such as alcohols, diacetyl, esters, organic acids, organic sulfides, and carbonyl compounds during fermentation (Ter Schure

Amino acid

‘ <

tt-Keto acid (e. g. L-glutamic acid)

<‘

Aldehyde (e. g. isobutyraldehyde)

У

Alcohol (e. g. isobutanol)

FIGURE 7.1 Simplified biochemistry of branched chain higher alcohol production from amino acids.

et al., 1998; Hazelwood et al., 2008). These compounds are formed via the Ehrlich pathway involving branched — chain amino acids such as isoleucine, leucine, methio­nine, phenylalanine, tryptophan, tyrosine, and valine, and biocatalysts such as transaminase, decarboxylase, and alcohol dehydrogenase (ADH) (Figure 7.1, Table 7.1 Ryan and Kohlhaw, 1974). This pathway is prevalent in yeast and is especially active when yeast is cultivated in growth medium whose carbon source is solely amino acids. Notably, catabolism of isoleucine, leucine, methi­onine, phenylalanine, tryptophan, tyrosine, and valine by S. cerevisiae via Ehrlich pathway generates

2- methylbutanol (active amyl alcohol), 3-methylbutanol (isoamyl alcohol), methionol, 2-phenylethanol, trypto — phol, p-hydroxyphenyl ethanol, and isobutanol, respec­tively (Figure 7.1; Table 7.1). These relatively long-chain alcohols are often referred to as fusel oils or fusel alcohols. During ethanolic fermentation by S. cerevisiae, small quan­tities of these alcohols (fusel oil) are produced (Singh and Kunkee, 1976). Whereas this mixture of alcohols may contribute flavor and body to wines, it can produce an off-flavor in wines when the acceptable concentration threshold is exceeded. Indeed, the catabolism of amino acids in S. cerevisiae and its regulation has been studied extensively (Ter Schure et al., 1998; Hazelwood et al., 2008; Dickinson et al., 1998,2000).

By transamination of amino acids to a-keto acids followed by decarboxylation of a-keto acids to aldehydes, these aldehydes can undergo reduction reaction to produce alcohols (Figure 7.1; Table 7.1). Scientists are
exploiting this biosynthetic pathway to take advantage of the amino acid biosynthesis capability of producing microorganisms such as E. coli and S. cerevisiae to pro­duce fusel alcohol, of which isobutanol appears to be the most attractive. In particular, the specificity of decar­boxylases has been suggested to be an important factor influencing the composition of fusel alcohols (Harrison and Collins, 1968; Suomalainen and Keranen, 1967). Furthermore, the amount of fusel alcohols produced by different yeasts and specific ADH activities with the cor­responding alcohols as substrates was found to be related as well (Singh and Kunkee, 1976). In recent years, metabolic engineering strategy using heterologous hosts such as E. coli and Clostridium cellulolyticum to produce higher alcohols from glucose and cellulose, respectively, is under investigation (Atsumi et al., 2008; Higashide et al., 2011). Depending on the source, 2-ketoacid decar­boxylase (KDC, encoded by the kivd gene) and ADH (encoded by the adh2 gene), which play critical roles in fusel oil production, may have broad substrate specific­ities toward the catalysis of 2-ketoacids and generation of isobutanol (Figures 7.1 and 7.2). When these two genes, kivd from Lactococcus lactis and adh2 from S. cere — visiae, were cloned and overexpressed in E. coli, approx­imately six long-chain alcohols including 1-propanol,

1- butanol, isobutanol, 2-methyl-1-butanol, 3-methyl — 1-butanol, and 2-phenylethanol were produced (Atsumi et al., 2008). This strategy exploits the presence of a highly active amino acid biosynthetic pathway in the host microorganism, keto acid pathway, and the ability

FIGURE 7.2 Schematic diagram depicting pathways leading to valine and isobutanol biosynthesis in S. cerevisiae. Genes encoding enzymes that catalyze each step are indi­cated and are as follows: ADH2 (alcohol dehydrogenase), Batl and Bat2 (branched chain amino acid aminotransfer­ases), ILV2 (acetolactate synthase), ILV3 (dihydroxyacid dehydratase), ILV5 (acetohydroxyacid reductoisomerase), Kdc/kivd/Pdc, 2-ketoacid decarboxylase (pyruvate decar­boxylase); and PDA (pyruvate dehydrogenase complex). (For color version of this figure, the reader is referred to the online version of this book.)

Cytosol

of the host to reroute its 2-ketoacid intermediates for alcohol synthesis (Atsumi et al., 2008). The amount of individual alcohol produced is compared with the level of its corresponding ketoacid. For example, when alsS from Bacillus subtilis and ilvCD from E. coli were overex­pressed in E. coli, the resulting strain accumulated remarkable amounts of 2-ketoisovalerate (KIV) in the fermentation broth. Furthermore, when kivd and adh2 were co-expressed in this recombinant strain, approxi­mately 22 g/l isobutanol was produced over the course of 112 h of fermentation (Atsumi et al., 2008). Notably, AlsS of B. subtilis, kivd from L. lactis, adh2 from S. cerevi­siae, and YqhD (nicotinamide adenine dinucleotide phos­phate (NADPH)-dependent ADH) from E. coli have high affinity for pyruvate and 2-ketoacids, 2-ketoacids, isobu — tyraldehyde, and isobutyraldehyde, respectively.

Monomeric sugars can be converted to ethanol directly, while starches and cellulose first must be hy­drolyzed to fermentable sugars either enzymatically or chemically (Bashir and Lee, 1994). Like most biofuels processes, bioethanol production from microalgae be­gins with the concentration of algae. The algae are then further dried and ground to a powder. In the next step of the process, the algae mass is hydrolyzed and Saccharomyces cerevisiae yeast is added to the biomass to begin the fermentation process. The resulting fer­mented mash contains about 11—15% ethanol by volume as well as the nonfermentable solids from algae and yeast cells. Ethanol is then distilled off the mash at ~ 96% strength. Despite widespread knowledge of this fermentation process, the details of the conversion pro­cess of algal celluloses-to-bioethanol are only partially understood. Celluloses comprise a large fraction of algal cells walls. These molecules are tightly packed and enzymatic access is often limited without a pretreatment step (Figure 10.6).

Many authors have reported that it is essential to introduce a pretreatment stage to release and convert the complex carbohydrates entrapped in the cell wall into simple sugars necessary for yeast fermentation. Cel­lulose can be made more accessible by the addition of an acid (Figure 10.7). Arantes and Saddler (2010) have sug­gested a model where prior to hydrolysis of cellulose to

FIGURE 10.7

monomeric units, cellulases must adsorb onto the surface of the insoluble cellulose (Figure 10.8). The action of the cellulases serves to loosen tightly packed fibrous cellulosic networks and create additional access to cellulose chains buried within the fibrils. Then the synergistic action of exo- and endoglucanases cleave accessible molecules to form soluble cello-oligosaccharides, or oligomers of <6 sugar units. These oligosaccharides are quickly hydro­lyzed to primarily cellobiose, or two glucose molecules linked by a b (1/4) bond. Cellobiose hydrolyzation to glucose monomers is usually completed by the extraneous addition of b-glucosidase.

Once glucose monomers have been rendered, bio­ethanol from microalgal biomass can be produced through two distinct pathways: direct dark fermentation or yeast fermentation of saccharified biomass. Whereas direct dark fermentation yields are typically much lower, the yeast fermentation process is a very well — established, relatively high-yield, low-energy-intensive process. Because microalgae can be harvested multiple times a year, some species have been shown to
theoretically yield an order of magnitude more bio­ethanol compared to a land-based crop such as corn (Table 10.3). Further, using microalgae as a raw material is strongly advantageous as algae sugars may be derived from multiple sources—from intracellular starches and from the cellulosic cell wall. Nevertheless, to achieve higher yields, it is still necessary to screen for high starch-producing algal strains coupled with identifying mechanisms and culture conditions for inducing maximal accumulation of intracellular starches.

In comparison to terrestrial feedstocks that contain lignin, certain species of microalgae and cyanobacteria have high potentiality for bioethanol production due to their high productivity rates, high biomass ferment­able carbohydrate content, and lack of lignin. Lignin is a recalcitrant substance (i. e. not easily degraded) present in the cell walls of terrestrial biomass that cannot be con­verted to bioethanol—its processing is a major impedi­ment for bioethanol production (Ragauskas et al.,

2006) . Microalgae’s potential can be highlighted by the fact that 75% of algal complex carbohydrates can be
hydrolyzed into a fermentable hexose monomer, and the fermentation yield of bioethanol is ~ 80% of the theoret­ical optimal value (Huntley and Redalje, 2007). Harun et al. (2009) have shown that the blue-green Chlorococum sp. produces a maximum bioethanol concentration of 3.83 g/l obtained from 10 g/l samples that are preex­tracted for lipids versus those that remain as dried intact cells. This indicates that microalgae can be used for the production of both lipid-based biofuels and ethanol bio­fuels from the same biomass as a means to increase their overall economic value (Jones and Mayfield, 2012). The microalgae Chlorella vulgaris and Porphyridium sp., particularly, have been considered as promising feed­stocks for bioethanol production because they can accu­mulate up to 37% and 54% (dry weight) of starch, respectively. The potential for simple, low-cost methods of bioethanol production from microalgae and cyano­bacteria are real. The next phase of biofuel research should develop improved methodologies to increase intracellular ethanol production efficiencies.

Agricultural waste materials like straw and the solid fraction of manure are lignocellulosic materials. These materials are strong, flexible and protected against decay. They consist of cellulose, hemicellulose and lignin. Lignin cannot be converted into biogas, and

Solubilization of lignocellulosic materials is inhibited by free fatty acids produced during hydrolyzation and subsequent acidogenesis. Increasing the number of methanogenic bacteria reduces the concentration of free fatty acids. Macronutrients nitrogen and phosphate

and the micronutrients S, Ca, Mg, Fe, Ni, Co, Mo, Zn, Mn, Se and Cu (Demirel et al.., 2011) are required for the multiplication of methanogenic bacteria. Scherer et al. (1983) determined the chemical composition of a number of methanogenic bacteria (Table 13.2). Scherer

(2011) advises also on the minimum concentration of ions in a digester. The basal medium of Guengoer — Demirci et al. (2004) and the recommendations of Speece (1987) are similar. Lebuhn et al. (2010) formulated a spe­cial cocktail. They commented that for maize silage Co should be added at 0.1 mg/kg VS and sodium at 30 mg/kg VS. Speece (1987) has a recommendation for Se. Se was not limiting in the tests by Lebuhn et al.

(2010) .

Straws, husks, bagasse and woody biomass are gener­ally deficient in macro — and micronutrients. The same holds for cattle manure in Asia as cattle feed mostly on rice straw. Manures in Europe and North America have an excess of nitrogen.

Jerger et al., 1982 found a 60% increase in methane yield in half the time in batch-fed anaerobic potential assays with extra-micronutrients (Table 13.3). They also added NH4CL and KH2PO4 to reduce the C/N ratio to 15 and the C/P ratio to 75. Similar results have been obtained by Komatsu et al. (2007). They obtained a methane yield of 280 l/kg VS with sewage sludge and rice straw at a hydraulic retention time of 20 days in a continuously operating digester at 36 °C. Somayaji et al. (1994) had 240 l/kg VS in 40 days for rice straw.

The addition of micronutrients has an effect of 10—70% on the methane production.

Sewage water cleanup sludges are a source of macro and micronutrients. Average primary sewage sludge has the right concentration for Ni and Mo. Co is an order of magnitude too low. In some sludges the concentrations of Fe, Co and Ni are too low (Speece, 1988). Concentra­tions for Ca, Fe, Zn, Mn and Cu are an order of magni­tude too high for its use in agriculture (Wolf et al., 2005).

Optimum nutrient conditions are cost-effective. Industrial fertilizers should be used, lacking organic sources of nitrogen and phosphate. For each kilogram of dry lignocellulosic biomass a maximum of 40 g of urea and 20 g of phosphate are required. Human urine is a good source of nitrogen and phosphate. Human feces are also good but require storage for more than 100 days in order to prevent the spread of illnesses. Eco — san toilets (Terefe and Edstram, 1999) separate urine and feces, so that urine can be directly used and feces stored for the required period.

ADDITION OF MICROBES

Op den Camp et al. (1991) describe an acidogenic reactor with rumen-derived bacteria. A hydraulic retention time of 12 h and a solids retention time of 72 h resulted in a methane yield of 440 l/kg VS for cel­lulose and 120 l/kg VS for barley and rye straw. The

methane was produced in a second reactor separated from the first by a filter with 0.03 mm pore size. The liquid without the free fatty acids was recycled to the first reactor. Soluble lignin products (humic acids) inhibited further degradation of the straws. The German company Ares Technology is performing tests

at pilot plant scale. Typical conversion yields are around 50% (Strecker, 2012).

Weiss et al. (2009) isolated and multiplied hemicellu — lytic bacteria. These were immobilized on trace metal activated zeolite. Digestion of second-stage sludge from a biogas plant gave a methane yield of 215 l/kg VS after 34 days (35 °C) and 150 l/kg VS for the control.

Ed de Jong Richard J. A. Gosselink2

1Avantium Chemicals, Amsterdam, The Netherlands, 2Food and Biobased Research,
Wageningen UR, Wageningen, The Netherlands
Corresponding author email: ed. dejong@avantium. com

OUTLINE

Introduction 278

Occurrence and Composition of Lignocellulosic

Biomass 278

Storage Carbohydrates 280

Structural Carbohydrates 280

Cellulose 280

Hemicelluloses 280

Glucuronoxylans 280

Glucomannan 282

Xyloglucans 282

Galactoglucomannans 282

Arabinoglucuronoxylans 282

Arabinogalactan 282

Arabinoxylan 283

b-(1/3, 1/4)-Glucans 283

Complex Heteroxylans 283

Conclusions on Carbohydrate Feedstocks 283

Lignin 283

Pretreatment Technologies 286

Steam Explosion 286

Liquid Hot Water 288

Wet Oxidation 288

Dilute and Concentrated Acid Pretreatment 289

Alkaline (Lime) Pretreatment Process 289

Pretreatment Technologies Still at a Laboratory/

Conceptual Stage 290

Ammonia Fiber Explosion/Ammonia Recycle Percolation) 290

Ionic Liquids 291

Sub/Supercritical Treatments 291

Summary of Lignocellulosic Biomass Pretreatments 291

Lignocellulosic Biorefineries—Classification 292

C6 and C6/C5 Sugar Platform 295

Fermentation Products 295

Chemical Transformation Products 296

Lignin Platform 296

Importance of Furans and Aromatics as Building

Blocks for Chemicals and Fuels 297

Carbohydrate Dehydration 298

Introduction 298

Furfural Production and Applications 298

5-Hydroxymethylfurfural Formation from Hexose Feedstock 301

Relevance of 5-Hydroxymethylfurfural as a Platform Chemical 304

Conversion of Technical Lignins into

Monoaromatic Chemicals 305

Base-catalyzed Depolymerization 305

Acid-catalyzed Depolymerization 305

Pyrolysis 305

Oxidative Depolymerization 306

Reductive Hydrodeoxygenation 306

Solvolysis 307

Sub — and Supercritical Water 307

Supercritical Solvents 308

Ionic Liquids 308

Future Perspectives of Lignin Aromatics 308

Conclusions and Further Perspectives 309

References 309

Bioenergy Research: Advances and Applications http://dx. doi. org/10.1016/B978-0-444-59561-4.00017-6

INTRODUCTION

Around the world significant steps are being taken to move from today’s fossil-based economy to a more sus­tainable economy based on biomass. A key factor in the realization of a successful biobased economy will be the development of biorefinery systems allowing highly efficient and cost-effective processing of biological feed­stocks to a range of biobased products, and successful integration into existing infrastructure. The recent climb in oil prices and consumer demand for environmentally friendly products have now opened new windows of opportunity for biobased chemicals and polymers. Industry is increasingly viewing chemi­cal and polymer production from renewable resources as an attractive area for investment. Within the biobased economy and the operation of a biorefinery there are sig­nificant opportunities for the development of biobased building blocks (chemicals and polymers) and materials (fiber products, starch derivatives, coatings, resins, etc.). In many cases this happens in conjunction with the pro­duction of bioenergy or biofuels. The production of bio­based products could generate US$ 10—15 billion of revenue for the global chemical industry. The economic production of biofuels is often a challenge. The copro­duction of chemicals, materials, food and feed can generate the necessary added value.

The world is more and more confronted with the reduction of fossil oil reserves, strong fluctuations of fossil fuel prices and the increase in CO2 emissions and the ensuing problem of the greenhouse gas effect. Recent development on the production of shale gas at various places in the world might change this picture on the short term, but the disadvantages associated with fossil resources stay in place. These environ­mental, social and economic challenges have created the need for sustainable alternatives to fossil fuels and chemicals (Brown, 2003; Kamm et al., 2006). The use of plant biomass as starting material is one of the alternatives to reduce the dependency on fossil oil for transportation fuels and is the main alternative to replace petrochemicals. The biomass can be trans­formed into energy, transportation fuels, various chem­ical compounds and materials such as natural fibers by biochemical, chemical, physical and thermal processes (Brown, 2003; Huber et al., 2006; Gallezot, 2012; Climent et al., 2011a, b; Lichtenthaler and Peters, 2004). The fermentation and the chemical conversion of carbohy­drates into value-added compounds has received increasing interest in the last decade, and in a bio­refinery different advantages may be taken from both processes (Kamm et al., 2006; Gallezot, 2012; Climent et al., 2011a; Lichtenthaler and Peters, 2004; Spiridon and Popa, 2008; Lin and Huber, 2009; Stocker, 2008; Dhepe and Fukuoka, 2008). However, the poten­tial competition with food and feed applications and the consequent rise in feedstock prices is an important aspect to take into consideration. Therefore the use of lignocellulosic feedstocks (often referred to as second — generation feedstocks) is strongly advocated. In addi­tion to carbohydrates also substantial amounts of lignin is produced when using lignocellulosic feedstocks. In this chapter the composition of lignocellulosic biomass is discussed followed by an overview of the most important pretreatment and fractionation technologies. Especially the effect of the different technologies on the subsequent fermentative/chemocatalytic conver­sions is addressed. The importance is illustrated by an overview of the most important commercial as well as anticipated chemical building blocks from car­bohydrates and lignin with a special emphasis on the production of furan-based building blocks from carbo­hydrates and aromatic building blocks originating from lignin.

Although, a very high reproducibility has been re­ported in a specific intralaboratory study (Granata and Argyropoulos, 1995), the reproducibility of quantitative 31P NMR spectroscopy reported in independent interla­boratory studies is much lower (Table 18.4). Moreover, even in studies conducted at the same laboratory, one can observe significant divergences between the results reported earlier (Granata and Argyropoulos, 1995) and more recently (Xia et al., 2001) for the same lignin sam­ples (see SE aspen and poplar lignins in Table 18.3). As expected, the worst reproducibility of 31P NMR analytical methods has been observed for different types of 5-substituted phenolic hydroxyls (S-units and

5- condensed G units) in a hardwood technical lignin (Table 18.4) due to a very poor signal resolution.

Content of Major Lignin Functional Groups Determined by Different Analytical Methods

Therefore, in these cases, we consider erroneous report­ing separately S-units and 5-condensed G-units, it is more reasonable to report them as "5-substituted pheno — lics". The reproducibility of 31P NMR is overall better for major signals such as aliphatic hydroxyls (AliphOH), phenolic hydroxyls (PhOH), and total OH, especially for Indulin AT lignin. Surprisingly, it is still not of very high quality the results reported for Alcell lignin (Table

18.4) . In part, but not completely, this observation might be explained by inconsistencies in the lignin itself. In addition, 31P NMR reports much lower carboxyl numbers than wet chemistry methods and 13C NMR methods (Ta­ble 18.3). In fact, 13C NMR methods usually report the sum of carboxyl and ester structures in general. For instance, the significantly higher numbers reported by 13C NMR for Alcell lignin might be explained by the sig­nificant contribution of ester structures (predominantly ethyl esters) in this lignin. However, it is quite unreason­able to expect significant amounts of esters in kraft lig­nins isolated from high alkaline solutions. Therefore, it might be concluded that 31P NMR also underestimates the amounts of carboxyl groups in lignins.

In summary, it appears that 31P NMR-I does not pro­vide sufficient resolution even between major signals, PhOH and AliphOH, while 31P NMR-II yields signifi­cantly lower values. Moreover, separation of S-units and G-condensed (at C-5) structures is very ambiguous in 31P NMR-II analysis of technical lignins, consequently so is the evaluation of S/G ratio and the degree of condensation. The amount of 5-substituted (S + G— condensed) and 5-unsubstitued phenolic hydroxyl should be reported instead. Comparative data for the analysis of various technical lignins by the 31P-II NMR method are summarized in Table 18.5.

Due to the high degree of variability in the structure of lignins discussed above, it is difficult to make any gen­eral conclusions on the existing structural differences among the lignins originated from various technical pro­cesses (Tables 18.3—18.5). In addition to the variability linked to the feedstock origin and process variables, the numbers for different structural moieties reported can vary due to the particularities of the used analytical methodologies (Table 18.3). There is overall a lack of comprehensive comparison of technical lignins. In addi­tion, the existing comparative studies are limited to a few lignin functionalities only, such as amounts of phenolic, aliphatic hydroxyl groups, or methoxyl groups.

Y. Allahverdiyeva*, E. M. Aro, S. N. Kosourov

Department of Biochemistry, University of Turku, Turku, Finland
Corresponding author email: allahve@utu. fi

It is well known that the fossil energy resources are limited. Despite the fact that millions of years of photo­synthesis were required to ensure the fossil fuel forma­tion and accumulation, the current consumption of fossil fuels occurs at a rapid rate. Such utilization of fos­sil fuels creates extreme damage to the environment, increasing the CO2 level in atmosphere and leading to global warming and pollution on the Earth. Future sce­narios predict an increase in CO2 partial pressure in the atmosphere from the current levels of approximately 380, to about 750 and up to 1000 patm until the end of this century (Raupach et al., 2007).

ronmentally friendly and renewable energy sources. An efficient strategy for production of bioenergy would employ photosynthetic microorganisms, which are collectively, a significant player in the global carbon cycle. Cyanobacteria and green algae have inherited mechanisms for production of hydrogen, which pos­sesses all properties of a clean and efficient energy carrier. Although the natural production of hydrogen by these microorganisms is negligible at the current state, there is a huge potential for engineering and synthetic biology advances of cyanobacteria and green algae toward commercially profitable production of hydrogen and other biofuels.

Bioenergy Research: Advances and Applications http://dx. doi. org/10.1016/B978-0-444-59561-4.00021-8