7.2.1 Lignocellulosic Biomass—Raw Material for Second Generation Biofuel

Lignocellulosic feedstocks consist of mainly cellulose, hemicellulose, and lignin and can be found in the cell walls of almost all plant-derived materials, such as wood and grass, agricultural residues, and municipal solid wastes. The relative composition of the lignocellulosic material, however, varies greatly, depending on source (Chandel and Singh 2011; Garrote et al. 1999; Mosier et al. 2005) and for an overview, the weight percentage of dry biomass of representative lignocellu — losic materials are listed (Table 7.1).

Cellulose (b-1-4-glucan), a linear polymer of glucose units, is the major component of the lignocellulose (accounting up to 50 % of the total plant dry weight), the most abundant form of biologically fixed carbon in the biosphere, and a primary target for biofuels that are metabolites from microbial conversions (as in bioethanol production). It is hence a material of high interest to utilize well, but also a very recalcitrant material, making its utilization difficult. A major challenge is still to manage to convert lignocellulose in high yields to fermentable sugars

(see also Sect. 7.3. Lignocellulosics requires pretreatment for degradation) and to follow this with an efficient process that reduces the oxygenated carbohydrates to fuel molecules (Chundawat et al. 2011). In the process to obtain fermentable sugars, microbial GHs are used as catalysts to obtain saccharification (hydrolysis) of different polysaccharides in the biomass (explained more in the sections below). The microbial GHs are catalysts designed to degrade complex carbohydrate polymers into mono or oligosaccharides, that allow uptake and metabolism by the microorganism selected as cell-factory for the conversion into the desired biofuel, even if the microorganism on its own is not capable to degrade the polymeric carbohydrate forms.

It has been predicted that based on available land, the energy potential of lignocellulosics worldwide allows an energy outtake of approximately 100 EJ/ annum (1 EJ = 1 x 1018 J, covering woody biomass, straw, and energy crops) (Parikka 2004), which is to be compared to the global energy demand (425 EJ in

2001) (Lewis and Nocera 2006) showing that approximately one-quarter of the current demand can be obtained, and thus additional resources are needed to cover a shift from fossil to renewable resources. A means to increase the possible overall energy outtake is to also turn to biomasses from marine environments.

Changing the structure of biomass, typically increasing the enzyme accessible surface area, and reducing the degrees of polymerization of biomass, are possible by physical pretreatments such as size reduction (Zhu and Pan 2010; Harun et al. 2011). Different types of milling (e. g., ball milling, hammer milling, colloid milling, two-roll milling, and vibro energy milling), irradiations (e. g., by micro­waves, gamma rays, electron beams, and ultrasonications), and extrusion (sub­jecting the biomass to heating, mixing, and shearing) are used for this propose (Taherzadeh and Karimi 2008; Zheng et al. 2009).

Modification of biomass structure with a single physical treatment is typically not enough for efficient enzymatic hydrolysis, although it can be enough for improvements in biogas production. Thus, the physical treatments are used prior to (or together with) chemical and biological treatments (Taherzadeh and Karimi 2008; Yu et al. 2009).

Size reduction is used prior to most chemical pretreatments. Although it can affect the efficiency of the process, it heavily impacts process economy. Most chemical pretreatments are not successful without size reduction. However, in explosive, organosolv, and solvent processes large particles may be used (Zhu and Pan 2010; Shafiei et al. 2012). Explosive pretreatments, such as steam explosions, need less energy than mechanical size reduction. However, the explosive pre­treatments are not easily possible in laboratory investigations and there are some limitations in their scalability. Furthermore, they are not much effective for soft­woods (Zhu and Pan 2010). In some organosolv processes, such as ethanol organosolv pretreatment, which is one of the most effective methods, size reduction is not necessary. The process is also effective for softwoods (Pan et al. 2005, 2006a, c, 2007a, b). However, the process is not yet considered as an alternative process for large-scale pretreatment of lignocelluloses.

Size reduction can also be performed after chemical pretreatment, as refereed post-chemical pretreatment size reduction. Post-chemical pretreatment size reduction has different advantages compared to that before chemical pretreat­ments. Besides more effectiveness, the main advantage is lower mechanical energy consumption. On the other hand, without pre-size reduction, it is possible to work with denser solids and consequently higher solid per liquid ratio in the process, resulting in more concentrated hemicellulose sugar liquid. Furthermore, separation of fibers from the pretreated mixture is easier after pretreatment (Zhu and Pan

2010) . However, post size reduction is not applicable in all pretreatments.

Different irradiation processes have also been shown to improve the digest­ibility of lignocelluloses (Fernandez-Cegri et al. 2012). Treatment of biomass with high energy irradiation can modify the structure (Bak et al. 2009). However, its application is limited to less recalcitrant biomass such as rice straw (Bak et al.

2009) . Ultrasonication, on the other hand, has been used at large scale for the improvement of digestibility of different organic material and sludge resulting in higher yield of biogas and lower amounts of residual sludge (Pham et al. 2009; Elbeshbishy et al. 2011).

Combined irradiation (mainly ultrasonic and microwave) and chemical pre­treatments also have been shown to improve digestibility than a single chemical pretreatment. The irradiations can work in conjunction with NaOH pretreatment (Rodrigues et al. 2011; Singh et al. 2011), ionic liquid pretreatment (Ha et al. 2011; Ninomiya et al. 2012), and ammonia pretreatment (Chen et al. 2012).

Fungi, white-rot basidiomycetes in particular, require H2O2 to allow the extra­cellular peroxidase enzymes to function in lignin degradation. The H2O2 is pro­vided by oxidases that are produced by the fungus and act by reducing molecular O2 to H2O2 alongside the oxidation of a co-substrate (Dashtban et al. 2009; Isroi et al. 2011). Two such oxidases are glyoxal oxidase (GLOX; EC 1.2.3.5) and aryl alcohol oxidase (AAO; EC 1.1.3.7). GLOX is a copper-containing enzyme found in many white-rot fungi, for example P. chrysosporium, and can oxidise a variety of co-substrates, typically simple aldehydes (Isroi et al. 2011; Martrnez et al. 2005). Some of these substrates are natural substances produced by the metabolism of the fungus, for example, glyoxal and methylglyoxal (Isroi et al. 2011). AAO, a flavoenzyme first discovered in P. eryngii, acts upon specific metabolites of the white-rot fungi to give rise to H2O2. Chlorinated anisyl alcohols are among the substrates oxidised by this enzyme as well as aromatic aldehydes released during lignin degradation in the presence of aryl alcohol dehydrogenase (AAD; EC 1.1.1.91) (Isroi et al. 2011; Martrnez et al. 2005).

Starch is a one of the best and most high yielding feedstock for ethanol production, but yeast S. cereviciae cannot utilize it directly. Hydrolysis is required to produce ethanol from starch by fermentation. Starch was traditionally hydrolyzed by acids, but the specificity of the enzymes, their inherent mild reaction conditions, and the absence of secondary reactions have made the amylases to be the catalysts generally used for this process. There are two steps present in hydrolysis of starch using amylases. First, these starch suspensions should be brought to high temper­atures (90-110 °C) for the breakdown of starch kernels. The product of this first step, called liquefaction, is a starch solution containing dextrines and small amounts of glucose. In second step, the liquefied starch is subject to saccharification at lower temperatures (60-70 °C) through glucoamylase obtained generally from Aspergillus niger or Rhizopus species (Pandey et al. 2000; Shigechi et al. 2004). Apar and O zbek (2004) provide information about the effects of operating conditions on the enzymatic hydrolysis of corn starch using commercial a-amylase. In previous years, the possibility of hydrolyzing starch at low temperatures for achieving energy savings is being investigated (Robertson et al. 2006).

Potential Starchy Substrates for Ethanol Production Corn:

Ethanol is produced almost exclusively from corn in the USA. Corn is milled for extracting starch, which is enzymatically treated for obtaining glucose syrup. Then, this syrup is fermented into ethanol. There are two types of corn milling in the industry: wet and dry. During the wet-milling process, corn grain is separated into its components. Starch is converted into ethanol and the remaining compo­nents are sold as co-products. During dry-milling, grains are not fractionated and all their nutrients enter the process and are concentrated into a distillation co­product utilized for animal feed called Dried Distiller’s Grains with solubles (DDGS) (Gaulati et al. 1996).

Wheat:

Generally in Europe, ethanol is mostly produced from beet molasses; in some countries like France it is also produced from wheat by a process similar to that of corn. Some efforts have been made for optimizing fermentation conditions. For example, Wang et al. (1999) have determined the optimal fermentation tempera­ture and specific gravity of the wheat mash. Soni et al. (2003) have optimized the conditions for starch hydrolysis using a-amylase and glucoamylase obtained by solid-state fermentation of wheat bran.

Cassava:

Cassava is an important alternative source of starch for ethanol production and for production of glucose syrups. Cassava is the tuber that has gained most interest due to its availability in tropical countries being one of the top ten more important tropical crops. Ethanol production from cassava can be accomplished using either the whole cassava tuber or the starch extracted from it. Starch extraction can be carried out through a high-yield large-volume industrialized process as the Alfa Laval extraction method (FAO 2004), or by a traditional process for small — and mid-scale plants. This process can be considered as the equivalent of the wet­milling process for ethanol production from corn. The production of cassava with high starch content (85-90 % dry matter) and less protein and minerals content is relatively simple.

Others:

Besides corn and wheat, ethanol can be produced from rye, barley, triticale (Wang et al. 1997), and sorghum (Prasad et al. 2007). For these cereals, some pretreatments have proved to be useful. Abd-Aziz (2002) suggested the utilization of sago palm for ethanol production in the case of Malaysia. The ethanol pro­duction from bananas and banana wastes using commercial a-amylase and glu — coamylase has been studied by Hammond et al. (1996). In their work, an ethanol yield of 0.5 L EtOH/kg dry matter of ripe bananas was obtained. The processing of starch-containing food wastes by adding malt to the pulverized feedstock has been patented (Chung and Nam 2002). One of the most promising crops for fuel ethanol production is sweet sorghum, which produces grains with high starch content, stalks with high sucrose content and leaves, and bagasse with high lignocellulosic content. In addition, this crop can be cultivated in both temperate and tropical countries requiring only one-third of the water needed for cane cropping and half of the water required by corn. Moreover, it is tolerant to the drought, flooding, and saline alkalinity (Winner Network 2002). Grassi (1999) reports that from some varieties of sweet sorghum, the following productivities can be obtained: 5 ton/Ha grains, 8 ton/Ha sugar, and 17 ton dry matter/Ha lignocellulosics. The estimated price for fuel ethanol production from this feedstock is US$200-300/m3, whereas the corresponding one for sugarcane ethanol is 260, for corn ethanol is 300-420, and for lignocellulosic ethanol is 450.

One of the most concerns from the rapidly increasing demand for biodiesel is the unregulated conversion of lands and/or cropping systems into oil palm (or called ‘‘land-use change’’). This in turn may cause the drawbacks to the ecosystems and society such as the substantial release of CO2 into the atmosphere from carbon stock change and food-fuel conflicts. So far, land-use change has not yet been much relevant for palm biodiesel production in Thailand as biodiesel demand is small scale and FFB used currently originate from the old palm plantations. However, to satisfy the future anticipated CPO demand for food and palm bio­diesel production, both oil palm productivity improvement and expansion of palm plantations are necessary and land-use change impacts would become important.

In the study, four possible direct land-use changes (dLUCs) originate into oil palm in Thailand including (1) tropical forest land in the northeast, (2) cropland such as cassava plantation in the northeast or the east, (3) grassland which is assumed to represent the available set-aside land in Thailand, and (4) old rubber fields in the south are considered in the analyses. Those land-use change scenarios are from the onsite surveys and/or interviews with farmers (Siangjaeo et al. 2011; JGSEE 2010) accompanied with the policy to encourage the new oil palm plan­tations of the RTG which specified in the palm oil industry and oil palm devel­opment plan (years 2008-2012) (NCGEB 2009). Based on the IPCC guidelines for calculating GHG emissions caused by dLUCs (IPCC 2006), the GHG perfor­mances of biodiesel after accounting for GHG consequences of dLUCs are esti­mated as shown in Table 2.2.

The results indicate that there is a wide range of GHG performance of biodiesel if dLUC is taken into account in the system boundary. The worst case found was for the conversion of tropical forest land to oil palm which would increase the released GHG of palm biodiesel production around 5-8 times as compared to the case where LUC is excluded, due to loss of biomass carbon stock. This in turn will cause biodiesel to have higher GHG emissions than conventional diesel by around 240-269 %. Nevertheless, it must be noted that forest land conversion is unlikely to occur in Thailand as it is illegal and restricted by government. The conversions of field crop (e. g. cassava), old rubber field, and set-aside land to oil palm on the other hand would bring about GHG benefits i. e., the gain of biomass carbon stock and/or the increase in soil organic carbon stock. The life-cycle GHG emissions of those three scenarios range from 3 to 29 g CO2 eq/MJ and the net avoided GHG emissions compared to diesel are ranged 56-91 %. Therefore, policy measures to regulate the new oil palm plantations by encouraging only the suitable land types are important to maintain the environmental sustainability of palm biodiesel production in the future.

Lignocellulosic biomass refers to plant biomass. All plant cells are surrounded by an extracellular matrix known as the cell wall, a polysaccharide-rich matrix which is a major component of terrestrial plants. The plant cell wall is a composite structure and is divided into three layers; the middle lamella, the primary wall and the secondary wall (Carpita and Gibeaut 1993; Somerville et al. 2004) (Figs. 4.1 and 4.2). The middle lamella is the most external of all three layers and acts as a separating panel between two cells (Heredia et al. 1995). It is composed mainly of pectic substances and is the first boundary component to be formed by the cell during cytoplasmic division (Heredia et al. 1995; Hernon et al. 2010). The primary and secondary cell walls differ in function and in composition.

Operating at high solid content in the enzymatic hydrolysis process is crucial for large-scale development of bioproduct and biofuel production processes. The aim of utilizing high solid content is to reach high sugar concentrations and subse­quently high concentrations of fermentation products, such as ethanol (Jprgensen et al. 2007a; Hodge et al. 2009). Furthermore, maintaining high substrate con­centrations throughout the conversion process is important for the energy balance and economic viability of biofuels production. Obtaining high concentration of fermentation product reduces global production cost since downstream processing and water consumption can be lowered. In case of ethanol production, distillation increases significantly the energy demand of the process, especially when ethanol concentrations are below 4 % (Ohgren et al. 2006).

In general, higher substrate loadings results in higher concentration of sugars. However, it has been shown that enzyme performance gradually decreases as substrate concentration increased. This can be attributed to enzyme inhibition by end products or toxics, presence of high concentrations of lignin and mass transfer limitations (Jprgensen et al. 2007a; Kristensen et al. 2009). In addition, some recent studies have reported a decrease in the adsorption capacity of cellulase enzymes to cellulose at high substrate loadings due to the effect of hydrolysis products (Kristensen et al. 2009; Wang et al. 2011). To overcome these barriers, different process configurations and strategies have been suggested to increase solids concentration in bioconversion processes.

Operating at high initial solids content (above 10-15 % (w/w)) involves tech­nical barriers. Viscosity of the pretreated materials is usually very high, which implies mass transfer limitations and mixing difficulties. Operating fed-batch processes by adding fresh substrate when viscosity decreases has been shown as an effective strategy to increase substrate concentrations in fermentations processes (Ballesteros et al. 2002; Varga et al. 2004). Another possibility is carrying out a prehydrolysis prior to initiate the simultaneous saccharification and fermentation (SSF) process. Using this strategy, the enzymes act at optimum temperature and reduce viscosity, which can result in higher substrates loadings (Rosgaard et al. 2007; Manzanares et al. 2011). A recent advance for operating at high consistency is the development of novel bioreactors with improved mixing capacity and low energy consumption (Jprgensen et al. 2007b; Zhang et al. 2009).

Another problem when operating at high substrate concentration is product inhibition. Cellobiose, glucose, and hemicellulose-derived sugars have been shown to inhibit the enzymes action (Xiao et al. 2004). In SSF processes, sugars released by the action of the enzymes are converted directly to ethanol by the fermenting microorganism, which reduces end-product inhibition (Ballesteros et al. 1994; Olsson et al. 2006). Constant removal of glucose during the process has been also proposed (Andric et al. 2010).

Degradation compounds originated from carbohydrates and lignin during the pretreatment affect the enzymatic hydrolysis (Tengborg et al. 2001; Ganna-Aparicio et al. 2006) and the fermenting microorganisms (Palmqvist and Hahn-Hagerdal 2000a; Oliva et al. 2003; Oliva et al. 2004). At high substrate loadings, the concentration of these compounds increases, therefore their influence in the bioconversion process can become more significant. Washing the pretreated material has been typically employed to eliminate toxic compounds and increase enzymatic hydrolysis and fermentation yields. To avoid washing and use the whole slurries, detoxification strategies such as laccase treatments have been studied to reduce the concentration of phenolic compounds and increase substrate concentrations in fermentation (Moreno et al. 2012).

Different articles reported the utilization of high substrate concentrations for ethanol production. Using wet oxidized and steam exploded corn stover, a sub­strate consistency of 15 % and 10-30 % dry matter (DM) in fermentation experiments, respectively, was studied (Varga et al. 2004; Lu et al. 2010). Using corn stover pretreated by combination of stream explosion and alkaline hydrogen peroxide, it was reached a solids loading of 30 % (Yang et al. 2010). With hydrothermal pretreated and steam pretreated wheat straw, it was possible to carry out hydrolysis and SSF at high substrate concentrations up to 20-30 % (Jprgensen 2009; Ballesteros et al. 2011), and with steam pretreated spruce it could be reached a consistency of 14 % (Hoyer et al. 2010).

6.4 Conclusion

Efficient utilization of lignocellulosic materials in a biorefinery depends on the advances in pretreatment technologies, enzyme saccharification, and fermentation of sugars to fuels and chemicals. Optimization of pretreatment and enzymatic hydrolysis processes is crucial to make bioconversion processes from lignocellu — losic biomass viable and cost-effective. The aim of the pretreatment is increasing the digestibility of carbohydrates while minimizing degradation of sugars and generation of toxic compounds. The pretreatment has to be adapted to the different raw materials and should be validated at large scale. The cost and efficiency of enzyme products still represents a major bottleneck to improve the economy of industrial biorefineries. To reduce costs of enzymatic hydrolysis processes, it is required the optimization of enzymatic mixtures in order to increase sugars pro­duction yields, reduce pretreatment severity, and decrease enzyme dosages. Complexity of lignocellulosic substrates involves that enzyme cocktails should be adapted for each raw material and type of pretreatment. In addition, operating at high solid content should be considered as a key issue for biofuels production. Finally, the integration of all the process steps has a remarkable importance to increase overall process efficiency and promote large-scale development. The type of biomass and pretreatment determines the process configuration requirements for hydrolysis and fermentation as each step has a large impact on all subsequent stages.

References

Agrawal P, Verma D, Daniell H (2011) Expression of Trichoderma reesei b-mannanase in tobacco chloroplasts and its utilization in lignocellulosic woody biomass hydrolysis. PLOS ONE 6(12):e29302. doi:10.1371/journal. pone.0029302

Alvira P, Tomas-Pejo E, Ballesteros M, Negro MJ (2010) Pretreatment technologies for an efficient bioethanol production process based on enzymatic hydrolysis: a review. Bioresour Technol 101:4851-4861

Alvira P, Negro MJ, Ballesteros M (2011a) Effect of endoxylanase and alpha-arabinofuranosidase supplementation on the enzymatic hydrolysis of steam exploded wheat straw. Bioresour Technol 102:4552-4558

Alvira P, Tomas-Pejo E, Negro MJ, Ballesteros M (2011b) Strategies of xylanase supplemen­tation for an efficient saccharification and cofermentation process from pretreated wheat straw. Biotechnol Prog 27:944-950

Andric P, Meyer AS, Jensen PA, Dam-Johansen K (2010) Effect and modelling of glucose inhibition and in situ glucose removal during enzymatic hydrolysis of pretreated wheat straw. Appl Biochem Biotechnol 160:280-297

Bajpai P (2004) Biological bleaching of chemical pulps. Crit Rev Biotechnol 24:1-58

Baker JO, King MR, Adney WS, Decker SR, Vinzant TB, Lantz SE, Nieves RE, Thomas SR, Li LC, Cosgrove DJ, Himmel ME (2000) Investigation of the cell-wall loosening protein expansin as a possible additive in the enzymatic saccharification of lignocellulosic biomass. Appl Biochem Biotechnol 84-86:217-223

Ballesteros I, Negro MJ, Oliva JM, Cabanas A, Manzanares P, Ballesteros M (2006) Ethanol production from steam-explosion pretreated wheat straw. Appl Biochem Biotechnol 130:496­508

Ballesteros I, Negro MJ, Oliva JM, Saez F, Manzanares P, Ballesteros M (2011) Increased ethanol concentration in saccharification and fermentation media of steam-exploded cereal straw. XIX. In: International symposium on alcohol fuels (ISAF), Development and utilization of alcohols fuels, to promote sustainability

Ballesteros I, Oliva JM, Carrasco JE, Ballesteros M (1994) Effect of media supplementation of ethanol production by simultaneous saccharification and fermentation process. Appl Biochem Biotechnol 45-46:283-294

Ballesteros I, Oliva JM, Negro MJ, Manzanares P, Ballesteros M (2002) Ethanol production from paper material using a simultaneous saccharification and fermentation system in a fed-batch basis. World J Microb Biot 18(6):559-561

Ballesteros M (2010) Enzymatic hydrolysis of lignocellulosic biomass. In: Waldron K (ed) Bioalcohol production. Biochemical conversion of lignocellulosic biomass. Woodhead Publishing, UK, pp 159-177

Banerjee G, Scott-Craig JS, Walton JD (2010) Improving enzymes for biomass conversion: a basic research perspective. Bioenerg Res 3:82-92 Bayer EA, Henrissat B, Lamed R (2008) The cellulosome: a natural bacterial strategy to combat biomass recalcitrance. In: Himmel ME (ed) Biomass recalcitrance. Deconstructing the plant cell wall for bioenergy. Blackwell Publishing, USA, pp 407-435 Bayer EA, Belaich JP, Shoham Y, Lamed R (2004) The cellulosomes: multienzyme machines for degradation of plant cell wall polysaccharides. Annu Rev Microbiol 58:521-554 Beg QK, Kapoor M, Mahajan L, Hoondal GS (2001) Microbial xylanases and their industrial applications: a review. Appl Microbiol Biotechnol 56:326-338 Benko Z, Siika-aho M, Viikari L, Reczey K (2008) Evaluation of the role of xyloglucanase in the enzymatic hydrolysis of lignocellulosic substrates. Enzyme Microb Technol 43:109-114 Berlin A, Gilkes N, Kilburnn D, Bura R, Markov A, Okunev O, Gusarov A, Maximenko V, Gregg D, Saddler J (2005) Evaluation of novel fungal cellulase prepration for ability to hydrolyze softwood substrate-evidence of the role of accessory enzymes. Enzyme Microb Technol 37:175-184

Berlin AB, Maximenko V, Gilkes N, Saddler J (2006) Optimization of enzymes complexes for lignocellulose hydrolysis. Biotechnol Bioeng 97(2):287-296 Bourbonnais R, Paice MG (1990) Oxidation of non-phenolic substrates. An expanded role for laccase in lignin biodegradation. FEBS Lett 267:99-102 Carvalheiro F, Duarte LC, Gfrio FM (2008) Hemicellulose biorefineries: a review on biomass pretreatments. J Sci Ind Res 67:849-864

Chandra RP, Bura R, Mabee WE, Berlin A, Pan X, Saddler JN (2007) Substrate pretreatment: the key to effective enzymatic hydrolysis of lignocellulosics? Adv Biochem Eng/Biotechnol 108:67-93

Cosgrove DJ (2000) Loosening of plant cell walls by expansins. Nature 407:321-326 Crepin VF, Faulds CB, Connerton IF (2004) Functional classification of microbial feruloyl esterases. Appl Microbiol Biotechnol 63:647-652 Dahlman O, Jacobs A, Berg J (2003) Molecular properties of hemicelluloses located in the surface and inner layers of hardwood and softwood pulps. Cellulose 10:325-334 Decker SR, Siika-Aho M, Viikari L (2008) Enzymatic depolymerization of plant cell wall hemicellulases. In: Himmel ME (ed) Biomass Recalcitrance. Deconstructing the Plant Cell Wall for Bioenergy. Blackwell Publishing, USA, pp 407-435 Dias AA, Freitas GS, Marques GSM, Sampaio A, Fraga IS, Rodrigues MAM, Evtuguin DV, Bezerra RMF (2010) Enzymatic saccharification of biologically pretreated wheat straw with white-rot fungi. Bioresour Technol 101:6045-6050 DOE. US Department of Energy (2003) Energy information agency. Available from: http:// www. exxonmobil. com/corportae/energy_outlook. aspx Eriksson T, Karlsson J, Tjerneld F (2002) A model explaining declining rate in hydrolysis of lignocellulose substrates with cellobiohydrolase I (Cel 7A) and endoglucanase I (Cel7B) of Trichoderma reesei. Appl Biochem Biotechnol 101:41-60 Esteghlalian AR, Svivastava V, Gilkes N, Gregg DJ, Saddler JN (2001) An overview of factors influencing the enzymatic hydrolysis of lignocellulosic feedstocks. In: Himmel ME, Baker W, Saddler JN (eds) Glycosyl hydrolases for biomass conversion. ACS, USA, pp 100-111 Faulds CB, Mandalari G, Curco RB, Bisignano G, Christakopoulos P, Waldron KW (2006) Synergy between xylanases from glycoside hydrolase family 10 and family 11 and a feruloyl esterase in the release of phenolic acids from cereal arabinoxylan. Appl Microbiol Biotechnol 71:622-629

Foreman PK, Brown D, Dankmeyer L, Dean R, Diener S, Dunn-Coleman NS, Goedegebuur F, Houfek TD, England GJ, Kelley AS, Meerman HJ, Mitchell T, Mitchinson C, Olivares HA, Teunissen PJ, Yao J, Ward M (2003) Transcriptional regulation of biomass-degrading enzymes in the filamentous fungus Trichoderma reesei. J Biol Chem 278:31988-31997 Garaa-Aparicio MP, Ballesteros I, Gonzalez A, Oliva JM, Ballesteros M, Negro MJ (2006) Effect of inhibitors release during steam-explosion pretreatement of barley straw on enzymatic hydrolysis. Appl Biochem Biotechnol 129:278-288 Garaa-Aparicio MP, Ballesteros M, Manzanares P, Ballesteros I, Gonzalez A, Negro MJ (2007) Xylanase contribution to the efficiency of cellulose enzymatic hydrolysis of barley straw. Appl Biochem Biotechnol 136-140:353-366

Gfrio FM, Fonseca C, Carvalheiro F, Duarte LC, Marques S, Bogel-Lukasik R (2010) Hemicelluloses for fuel ethanol: a review. Bioresour Technol 101:4775-4800 Guillen F, Martmez AT, Martmez MJ (1992) Substrate specificity and properties of the aryl — alcohol oxidase from the ligninolytic fungus Pleurotus eryngii. Eur J Biochem 209:603-611 Guillen F, Martmez MJ, Munoz C, Martmez AT (1997) Quinone redox cycling in the ligninolytic fungus Pleurotus eryngii leading to extracellular production of superoxide anion radical. Arch Biochem Biophys 339:190-199

Hahn-Hagerdal B, Karhumaa K, Fonseca C, Spencer-Martins I, Gorwa-Grauslund MF (2007) Towards industrial pentose-fermenting yeast strains. Appl Microbiol Biotechnol 74:937-953 Harris PV, Welner D, McFarland KC, Re E, Navarro Poulsen JC, Brown K, Salbo R, Ding H, Vlasenko E, Merino S, Xu F, Cherry J, Larsen S, Lo Leggio L (2010) Stimulation of lignocellulosic biomass hydrolysis by proteins of glycoside hydrolase family 61: structure and function of a large, enigmatic family. Biochemistry 49:3305-3316 Hendriks ATWM, Zeeman G (2009) Pretreatments to enhance the digestibility of lignocellulosic biomass. Bioresour Technol 100:10-18

Henrissat B, Davies G (1997) Structural and sequences-based classification of glycoside hydrolases. Curr Opin Struct Biol 7:637-644

Himmel ME, Andey WS, Baker JO, Nieves RA, Thomas SR (1996) Cellulases: structure, function, and applications. In: Wyman CE (ed) Handbook on bioethanol production and utilization. Taylor and Francis, UK, pp 143-161 Himmel ME, Ding SY, Johnson DK, Adney WS, Nimlos MR, Brady JW, Foust TD (2007) Biomass recalcitrance: engineering plants and enzymes for biofuels production. Science 315:804-807

Himmel ME, Picataggio SK (2008) Our challenge is to acquire deeper understanding of biomass recalcitrance and conversion. In: Himmel ME (ed) Biomass recalcitrance. Deconstructing the Plant Cell Wall for Bioenergy. Blackwell Publishing, USA, pp 1-6 Hodge DB, Karim MN, Schell DJ, McMillan JD (2009) Model-based fed-batch for high-solids enzymatic cellulose hydrolysis. Appl Biochem Biotechnol 152(1):88-107 Hoyer K, Galbe M, Zacchi G (2010) Effects of enzyme feeding strategy on ethanol yield in fed — batch simultaneous saccharification and fermentation of spruce at high dry matter. Biotechnol Biofuels 3:14

Hu J, Arantes V, Saddler JN (2011) The enhancement of enzymatic hydrolysis of lignocellulosic substrates by the addition of accessory enzymes such as xylanase: is it an additive or synergistic effect? Biotechnol Biofuels 4:36

Igarashi K, Uchihashi T, Koivula A, Wada M, Kimura S, Okamoto T, Penttila M, Ando T, Samejima M (2011) Traffic jams reduce hydrolytic efficiency of cellulase on cellulose surface. Science 333:1279-1282

Jin M, Lau MW, Balan V, Dale BE (2010) Two-step SSCF to convert AFEX-treated switchgrass to ethanol using commercial enzymes and Saccharomyces cerevisiae 424A(LNH-ST). Bioresour Technol 101:8171-8178

J0rgensen H (2009) Effect of nutrients on fermentation of pretreated wheat straw at very high dry matter content by saccharomyces cerevisiae. Appl Biochem Biotechnol 153:44-57 J0rgensen H, Kristensen JB, Felby C (2007a) Enzymatic conversion of lignocellulose into fermentable sugars: challenges and opportunities. Biofuel Bioprod Bior 1:119-134

J0rgensen H, Olsson L (2006) Production of cellulases by Penicillium brasilianum IBT 20888- Effect of substrate on hydrolytic performance. Enzyme Microb Technol 38:381-390 J0rgensen H, Vibe-Pedersen J, Larsen J, Felby C (2007b) Liquefaction of lignocellulose at high — solids concentrations. Biotechnol Bioeng 96:862-870 Juhasz T, Szengyel Z, Reczey K, Siika-Aho M, Viikari L (2005) Characterization of cellulases and hemicellulases produced by Trichoderma reesei on various carbon sources. Process Biochem 40:3519-3525

Jurado M, Prieto A, Martmez-Alcala A, Martmez AT, Martmez MJ (2009) Laccase detoxification of steam-exploded wheat straw for second generation bioethanol. Bioresour Technol 100:6378-6384

Jonsson JL, Palmqvist E, Nilvebrant N-O, Hahn-Hagerdal B (1998) Detoxification of wood hydrolysates with laccase and peroxidase from the white-rot fungus Trametes versicolor. Appl Microbiol Biotechnol 49:691-697

Kalyani D, Dhiman SS, Kim H, Jeya M, Kim I-W, Lee J-K (2012) Characterization of a novel laccase from the isolated Coltricia perennis and its application to detoxification of biomass. Process Biochem 47:671-678

Kersten PJ (1990) Glyoxal oxidase of Phanerochaete chrysosporium: Its characterization and activation by lignin peroxidase. Proc Natl Acad Sci USA 87:2936-2940 Kolb M, Sieber V, Amann M, Faulstich M, Schieder D (2012) Removal of monomer delignification products by laccase from Trametes versicolor. Bioresourc Technol 104:298­304

Kovacs K, Macrelli S, Szakacs G, Zacchi G (2009) Enzymatic hydrolysis of steam-pretreated lignocellulosic materials with Trichoderma viride enzymes produced in-house. Biotechnol Biofuels 2:14

Kristensen JB, Felby C, J0rgensen H (2009) Yield-determining factors in high-solids enzymatic hydrolysis of lignocellulose. Biotechnol Biofuels 2(1):11 Kubicek CP (1992) The cellulase proteins of Trichoderma reesei: structure, multiplicity, mode of action and regulation of formation. Adva Biochem Eng 45:1-27 Kuhar S, Nair LM, Kuhad RC (2008) Pretreatment of lignocellulosic material with fungi capable of higher lignin degradation and lower carbohydrate degradation improves substrate acid hydrolysis and the eventual conversion to ethanol. Can J Microbiol 54:305-313 Kumar P, Barrett DM, Delwiche MJ, Stroeve P (2009) Methods for pretreatment of lignocellulosic biomass for efficient hydrolysis and biofuel production. Ind Eng Chem Res 48:3713-3729

Kumar R, Wyman CE (2008) Effect of enzyme supplementation at moderate cellulase loadings on initial glucose and xylose release from corn stover solids pretreated by leading technologies. Biotechnol Bioeng 102(2):457-467 Kumar R, Wyman CE (2009a) Does change in accessibility with conversion depend on both the substrate and pretreatment technology? Bioresour Technol 100:4193-4202 Kumar R, Wyman CE (2009b) Effect of xylanase supplementation of cellulase on digestion of corn stover solids prepared by leading pretreatment technologies. Bioresour Technol 100:4203-4213

Kumar R, Wyman CE (2009c) Effects of cellulase and xylanase enzymes on the deconstruction of solids from pretreatment of poplar by leading technologies. Biotechnol Prog 25:302-314 Kurasin M, Valjamae P (2011) Processivity of cellobiohydrolases is limited by the substrate. J Biol Chem 286:169-177

Laureano-Perez L, Teymouri F, Alizadeh H, Dale BE (2005) Understanding factors that limit enzymatic hydrolysis of biomass. Appl Biochem Biotechnol 121:1081-1099 Larsson S, Cassland P, Jonsson LF (2001) Development of a Saccharomyces cerevisiae strain with enhanced resistance to phenolic compounds inhibitors in lignocellulose hydrolysates by heterologous expression of laccase. Appl Environ Microbiol 67:1163-1170 Lu Y, Wang Y, Xu G, Chu J, Zhuang Y, Zhang S (2010) Influence of high solid concentration on enzymatic hydrolysis and fermentation of steam-exploded corn stover biomass. Appl Biochem Biotechnol 160:360-369

Mansfield SD, Mooney C, Saddler JN (1999) Substrate and enzyme characteristics that limit cellulose hydrolysis. Biotechnol Prog 15:804-816 Manzanares P, Negro MJ, Oliva JM, Saez F, Ballesteros I, Ballesteros M, Cara C, Castro E, Ruiz E (2011) Different process configurations for bioethanol production from pretreated olive pruning biomass. J Chem Technol Biotechnol 86(6):881-887 Martmez AT (2002) Molecular biology and structure-function of lignin-degrading heme peroxidases. Enz Microb Technol 30:425-444

Martmez AT, Speranza M, Ruiz-Duenas FJ, Ferreira P, Camarero S, Guillen F, Martmez MJ, Gutierrez A, del Rfo JC (2005) Biodegradation of lignocellulosics: Microbiological, chemical and enzymatic aspects of fungal attack to lignin. Intern Microbiol 8:195-204 Martinez D, Berka RM, Henrissat B, Saloheimo M, Arvas M, Baker SE, Chapman J, Chertkov O, Coutinho PM, Cullen D, Danchin EGJ, Grigoriev IV, Harris P, Jackson M, Kubicek CP, Han CS, Ho I, Larrondo LF, De Leon AL, Magnuson JK, Merino S, Misra M, Nelson B, Putnam N, Robbertse B, Salamov AA, Schmoll M, Terry A, Thayer N, Westerholm-Parvinen A, Schoch CL, Yao J, Barabote R, Nelson MA, Detter C, Bruce D, Kuske CR, Xie G, Richardson P, Rokhsar DS, Lucas SM, Rubin EM, Dunn-Coleman N, Ward M, Brettin TS (2008) Genome sequencing and analysis of the biomass-degrading fungus Trichoderma reesei (syn. Hypocrea jecorina). Nature Biotechnol 26:553-560

Mayer AM, Staples RC (2002) Laccase: new functions for an old enzyme. Phytochem 60:551­565

Medve J, Karlsson J, Lee D, Tjerneld F (1998) Hydrolysis of microcrystalline cellulose by cellobiohydrolase I and endoglucanase II from Trichoderma reesei: adsorption, sugar production pattern, and synergism of the enzymes. Biotechnol Bioeng 59(5):621-634 Merino ST, Cherry J (2007) Progress and challenges in enzyme development for biomass utilization. Adv Biochem Eng Biotechnol 108:95-120 Moilanen U, Kellock M, Galkin S, Viikari L (2011) The laccase-catalyzed modification of lignin for enymatic hydrolysis. Enzyme Microb Technol 49:492-498 Moreno AD, Ibarra D, Fernandez JL, Ballesteros M (2012) Different laccase detoxification strategies for ethanol production from lignocellulosic biomass by the thermotolerant yeast Kluyveromyces marxianus CECT 10875. Bioresour Technol 106:101-109 Mukhopadhyay M, Kuila A, Tuli D, Banerjee R (2011) Enzymatic depolymerization of Ricinus communis, a potential lignocellulosic for improved saccharification. Biomass Bioenerg 35:3584-3591

Mosier N, Wyman CE, Dale BD, Elander RT, Lee YY, Holtzapple M, Ladisch CM (2005) Features of promising technologies for pretreatment of lignocellulosic biomass. Bioresour Technol 96:673-686

Numan MT, Bhosle NB (2006) Alpha-L-arabinofuranosidases: the potential applications in biotechnology. J Ind Microbiol Biotechnol 33:247-260 Ohgren K, Rudolf A, Galbe M, Zacchi G (2006) Fuel ethanol production from steam-pretreated corn stover using SSF at higher dry matter content. Biomass Bioenerg 30:863-869 Oliva JM, Ballesteros I, Negro MJ, Manzanares P, Cabanas A, Ballesteros M (2004) Effect of binary combinations of selected toxic compounds on growth and fermentation of Kluyver­omyces marxianus. Biotechnol Prog 20:715-720 Oliva JM, Saez F, Ballesteros I, Gonzalez A, Negro MJ, Manzanares P, Ballesteros M (2003) Effect of lignocellulosic degradation compounds in the steam explosion pretreatment on ethanol fermentation by thermotolerant yeast Kluyveromyces marxianus. Appl Biochem Biotechnol 105-108:141-153

Olsson L, Soerensen HR, Dam BP, Christensen H, Krogh KM, Meyer AS (2006) Separate and simultaneous enzymatic hydrolysis and fermentation of wheat hemicellulose with recombinant xylose utilizing Saccharomyces cerevisiae. Appl Microbiol Biotechnol 129­132:117-129

Olsson L, J0rgensen H, Krogh KBR, Roca C (2004) Bioethanol production from lignocellulosic material. In: Dumitriu S (ed) Polysaccharides: structural diversity and functional versatility. CRC Press, USA, pp 957-993

Palmqvist E, Hahn-Hagerdal B (2000a) Fermentation of lignocellulosic hydrolysates. II: inhibitors and mechanism of inhibition. Bioresour Technol 74:25-33 Palmqvist E, Hahn-Hagerdal B (2000b) Fermentation of lignocellulosic hydrolysates. I: inhibition and detoxification. Bioresour Technol 74:17-24

Palonen H, Viikari L (2004) Role of oxidative enzymatic treatments on enzymatic hydrolysis of softwood. Biotechnol Bioeng 86:550-557

Pan XJ, Gilkes N, Saddler JN (2006) Effect of acetyl groups on enzymatic hydrolysis of cellulosic substrates. Olzforschung 60:398-401

Pan X, Xie D, Gilkes N, Gregg DJ, Saddler JN (2005) Strategies to enhance the enzymatic hydrolysis of pretreated softwood with high residual lignin content. Appl Biochem Biotechnol 124:1069-1079

Panagiotou G, Olsson L (2007) Effect of compounds released during pre-treatment of wheat straw on microbial growth and enzymatic hydrolysis rates. Biotechnol Bioeng 96:250-258 Perez JA, Ballesteros I, Ballesteros M, Saez F, Negro MJ, Manzanares P (2008) Optimizing liquid hot water pretreatment conditions to enhance sugar recovery from wheat straw for fuel — ethanol production. Fuel 87:3640-3647

Persson I, Tjerneld F, Hahn-Hagerdal B (1991) Fungal cellulolytic enzyme production: a review. Process Biochem 26:65-74

Polizeli ML, Rizzatti AC, Monti R, Terenzi HF, Jorge JA, Amorim DS (2005) Xylanases from fungi: properties and indusrtial applications. Appl Microbiol Biotechnol 67(5):577-591 Poutanen K, Puls J (1989) The xylanolytic enzyme system of Trichoderma reesei. In: Lewis G, Paice M (eds) Biogenesis and biodegradation of plant cell wall polymers. ACS, USA, pp 630­640

Qi B, Chen X, Su Y, Wan Y (2011) Enzyme adsorption and recycling during hydrolysis of wheat straw lignocellulose. Bioresour Technol 102:2881-2889 Qi B, Luo J, Chen G, Chen X, Wan Y (2012) Application of ultrafiltration and nanofiltration for recycling cellulase and concentrating glucose from enzymatic hydrolyzate of steam exploded wheat straw. Bioresour Technol 104:466-472

Qing Q, Yang B, Wyman CE (2010) Xylooligomers are strong inhibitors of cellulose hydrolysis by enzymes. Bioresour Technol 101:9624-9630 Ragauskas AJ, Williams CK, Davison BH, Britovsek G, Cairney J, Eckert CA, Frederick WJ, Hallett JP, Leak DJ, Liotta CL, Mielenz JR, Murphy R, Templer R, Tschaplinski T (2006) The path forward for biofuels and biomaterials. Science 311:484-489 Raweesri P, Riangrungrojana P, Pinphanichakarn P (2008) Alpha-L-arabinofuranosidase from Streptomyces sp. PC22: purification, characterization and its synergistic action with xylanolytic enzymes in the degradation of xylan and agricultural residues. Bioresour Technol 99:8981-8986

Remond C, Aubry N, Cronier D, №ё! S, Martel F, Roge B, Rakotoarivonina H, Debeire P, Chabbert B (2010) Combination of ammonia and xylanase pretreatments: impact on enzymatic xylan and cellulose recovery from wheat straw. Bioresour Technol 101:6712-6717 Rosgaard L, Andric P, Dam-Johansen K, Pedersen S, Meyer AS (2007) Effects of substrate loading on enzymatic hydrolysis and viscosity of pretreated barley straw. Appl Biochem Biotechnol 143:27-40

Ruiz-Duenas FJ, Martmez AT (2009) Microbial degradation of lignin: How a bulky recalcitrant polymer is efficiently recycled in nature and how we can take advantage of this. Microbial Biotechnol 2:164-177

Ruiz-Duenas FJ, Martmez MJ, Martmez AT (1999) Molecular characterization of a novel peroxidise isolated from the ligninolytic fungus Pleurotus eryngii. Mol Microbiol 31:223-235 Saha BC (2000) Alpha-L-arabinofuranosidases: biochemistry, molecular biology and application in biotechnology. Biotechnol Adv 18:403-423 Saha BC (2003) Hemicellulose bioconversion. J Ind Microbiol Biotechnol 30:279-291 Saloheimo M, Paloheimo M, Hakola S, Pere J, Swanson B, Nyyssonen E, Bhatia A, Ward M, Penttila M (2002) Swollenin, a Trichoderma reesei protein with sequence with sequence

similary to the plant expansins, exhibits disruption activity on cellulosic materials. Eur J Biochem 269:4202-4211

Salvachua D, Prieto A, Lopez-Abelairas M, Lu-Chau T, Martinez AT, Martinez MJ (2011) Fungal pretreatment: An alternative in second-generation ethanol from wheat straw. Bioresour Technol 102:7500-7506

Sanchez OJ, Cardona CA (2008) Trends in biotechnological production of fuel ethanol from different feedstocks. Bioresour Technol 99:5270-5295 Selig MJ, Adney WS, Himmel ME, Decker SR (2009) The impact of cell wall acetylation on corn stover hydrolysis by cellulolytic and xylanolytic enzymes. Cellulose 16:711-722 Selig MJ, Knoshaug EP, Adney WS, Himmel ME, Decker SR (2008) Synergistic enhancement of cellobiohydrolase performance on pretreated corn stover by addition of xylanase and esterase activities. Bioresour Technol 99:4997-5005

Sipos B, Benko Z, Dienes D, Reczey K, Viikari L, Siika-Aho M (2010) Characterisation of specific activities and hydrolytic properties of cell-wall-degrading enzymes produced by Trichoderma reesei Rut C30 on different carbon sources. Appl Biochem Biotechnol 161:347­364

Sipos B, Szilagyi M, Sebestyen Z, Perazzini R, Dienes D, Jakab E, Crestini C, Reczey K (2011) Mechanism of the positive effect of poly(ethylene glycol) addition in enzymatic hydrolysis of steam pretreated lignocelluloses. C R Biol 334:812-823 Sun Y, Cheng J (2002) Hydrolysis of lignocellulosic materials for ethanol production: a review. Bioresour Technol 83:1-11

Tabka MG, Herpoёl-Gimbert I, Monod F, Asther M, Sigoillot JC (2006) Enzymatic saccharification of wheat straw for bioethanol production by a combined cellulase xylanase and feruloyl esterase treatment. Enzyme Microb Tech 39:897-902 Taherzadeh MJ, Karimi K (2008) Pretreatment of lignocellulosic wastes to improve ethanol and biogas production: a review. Int J Mol Sci 9:1621-1651 Tengborg C, Galbe M, Zacchi G (2001) Reduced inhibition of enzymatic hydrolysis of steam — pretreated softwood. Enzyme Microb Tech 28:835-844 Tenkanen M, Makkonen M, Perttula M, Viikari L, Teleman A (1997) Action of Trichoderma reesei mannanase on galactoglucomannan in pine kraft pulp. J Biotech 57:191-204 Teter S, Xu F, Nedwin GE, Cherry (2010) Enzymes for biorefineries. In: Kamm B, Gruber PR, Kamm M (eds) Biorefineries-industrial processes and products. Wiley, USA, pp 357-383 Tomas-Pejo ME, Ballesteros M, Oliva JM, Olsson L (2010) Adaptation of the xylose fermenting yeast Saccharomyces cerevisiae F12 for improving ethanol production in different fed-batch SSF processes. J Ind Microbiol Biotechnol 37:1211-1220 Tomas-Pejo ME, Alvira P, Ballesteros M, Negro MJ (2011) Pretreatment technologies for lignocellulose-to-bioethanol conversion. In: Larroche C, Ricke SC, Dussap CG, Gnansounou E, Pandey A (eds) Biofuels: Alternative feedstocks and conversion processes. Academic, USA, pp 149-176

Tu M, Chandra RP, Saddler JN (2007) Evaluating the distribution of cellulases and the recycling of free cellulases during the hydrolysis of lignocellulosic substrates. Biotechnol Prog 23:398­406

Tu M, Saddler JN (2010) Potential enzyme cost reduction with the addition of surfactant during the hydrolysis of pretreated softwood. Appl Biochem Biotechnol 161:274-287 Varga E, Klinke HB, Reczey K, Thomsen AB (2004) High solid simultaneous saccharification and fermentation of wet oxidized corn stover to ethanol. Biotechnol Bioeng 88:567-574 Vries RP, Harry CMK, Charlotte HP, Jacques AEB, Jaap V (2000) Synergy between enzymes from Aspergillus involved in the degradation of plant cell wall polysaccharides. Carbohyd Res 327:401-410

Wang W, Kang L, Wei H, Arora R, Lee YY (2011) Study on the decreased sugar yield in enzymatic hydrolysis of cellulosic substrate at high solid loading. Appl Biochem Biotechnol 164:1139-1149

Wilson DB (2008) Aerobic microbial cellulase systems. In: Himmel ME (ed) Biomass recalcitrance. Deconstructing the plant cell wall for bioenergy. Blackwell Publishing, USA, pp 374-392

Wyman CE, Balan V, Dale BE, Elander RT, Falls M, Hames B, Holtzapple MT, Ladisch MR, Lee YY, Mosier N, Pallapolu VR, Shi J, Thomas SR, Warner RE (2011) Comparative data on effects of leading pretreatments and enzyme loadings and formulations on sugar yields from different switchgrass sources. Bioresour Technol 102:11052-11062

Xiao Z, Zhang X, Greff DJ, Saddler JN (2004) Effects of sugar inhibition on cellulases and b- glucosidase during enzymatic hydrolysis of softwood substrates. Appl Biochem Biotechnol 113-116:1115-1126

Yang M, Li W, Liu B, Li Q, Xing J (2010) High-concentration sugars production from corn stover based on combined pretreatments and fed-batch process. Bioresour Technol 101:4884­4888

Yang B, Wyman CE (2008) Pretreatment: the key to unlocking low-cost cellulosic ethanol. Biofuel Bioprod Bior 2:26-40

Yang R, Xu S, Wang Z, Yang W (2005) Aqueous extraction of corncob xylan and production of xylooligosaccharides. LWT-Food Sci Technol 38:677-682

Yang M, Zhang A, Liu B, Li W, Xing J (2011) Improvement of cellulose conversion caused by the protection of Tween-80 on the adsorbed cellulase. Biochem Eng J 56:125-129

Zhang J, Siika-aho M, Tenkanen M, Viikari L (2011) The role of acetyl xylan esterase in solubilisation of xylan and enzymatic hydrolysis of wheat straw and giant reed. Biotechnol Biofuels 4:60

Zhang X, Qin W, Paice MG, Saddler JN (2009) High consistency enzymatic hydrolysis of hardwood substrates. Bioresour Technol 100:5890-5897

Lignin, a very complex polymer, playing a cementing role to connect cells, increases the mechanical strength properties, and makes plant resistant against diseases and biodegradation by microorganisms. Lignin is sometimes referred as glue between hemicellulose and cellulose components; while sometimes the hemicellulose is referred as glue between lignin and cellulose. Anyway, hemi — cellulose and lignin are known to cover the surface of cellulose which adds structural strength to the cellulose matrix (Perez and Samain 2010). Softwoods (25-40 %) contain higher lignin than hardwoods (18-25 %) and agriculture resi­dues (10-20 %) (Fengel and Wegener 1984; McMillan 1992); however, the lignin content is not the only difference between softwoods and other lignocelluloses. The main distinction is originated from the difference in monomeric units and linkage types in lignin. This dissimilarity in the lignin content may result in significant differences in susceptibility of various pretreatment techniques between hardwoods and softwoods. Pretreatment of hardwoods and agriculture residues is usually less harsh than softwoods. The reason is the presence of higher number of vessels in the hardwoods and agriculture residues which permit greater heat and mass transfer into the biomass matrix (Cochard and Tyree 1990; Hepworth et al. 2002; Kim et al. 2011). Generally, easier penetration of chemicals, enzymes, and heat makes the hardwoods and agriculture residues easier for pretreatment than softwoods.

Lignin is a cross-linked polymer of hydroxyphenylpropanoid units connected by C-C and C-O-C linkages, in which over 10 inter-phenylpropane linkage types have been detected. There are several monomeric units and linkage types in lignin. There are two major classes of lignin, guaiacyl lignins (G-lignin) and syringyl lignins (S-lignin). They contain guaiacyl (G), Syringyl (S), and hydroxybenzal — dehyde (H) units (Lewis and Yamamoto 1990). Different lignocellulose materials with different age and cultivation conditions have different ratios of G, S, and H. The lower accessibility of plant vessels is partially the result of the occurrence of guaiacyl lignin type in the vessel walls. Therefore, not only the amount of lignin, but also the guaiacyl to syringyl ratio in lignin can affect the swelling of the cellulosic residue (Ramos et al. 1992). The principal structural elements in lignins have been largely investigated; however, many aspects of the lignin chemistry are still unclear.

For lignin synthesis in woody materials, a series of secondary reactions are recognized leading to cross-linking between lignin and hemicelluloses (Lee 1997). Biodegradation of lignin is a secondary metabolic process, occurring only under low levels of nitrogen (Lee 1997).

During plant biosynthesis, it is believed that lignin is not simply deposited between cellulose and hemicellulose, but is linked with at least part of them. These linkages are termed as lignin-polysaccharide complex (LPC) or lignin-carbohy­drate complex (LCC) (Chesson 1988). Thus, as a result of these linkages, it is almost impossible to completely separate or purify cellulose or hemicellulose from lignin, and to have lignin free of polysaccharides. Furthermore, lignin has a ten­dency of recondensation during delignification processes (Kim et al. 2003). Not only are van der Waals and H-bond involved, but chemical bonds such as covalent bonds are also detected between lignin and polysaccharides (Besombes and Mazeau 2005).

A scanning electron microscope (SEM) is an electron microscope that images a sample by scanning it with a beam of electrons in a raster scan pattern. The electrons bombard the atoms of the sample and the signal produced reflects information about the sample’s surface topography, composition and other prop­erties such as electrical conductivity.

For SEM analysis of grass biomass samples which were subjected to various mild acid pre-treatments as described earlier (a study done by the authors A. O’Donovan and V. K. Gupta), the treated grass biomass residues were oven dried at 60 °C and adhered onto a stainless steel specimen holder called a specimen stub with the aid of an adhesive carbon tab. As grass biomass is non-conductive it is coated with an ultrathin coating of electrically conducting material, deposited on the sample either by low-vacuum sputter coating or by high-vacuum evaporation. Non-conductive specimens tend to charge when scanned by the electron beam, and especially in secondary electron imaging mode, this causes scanning faults and other image artefacts. In the below images, the grass biomass was gold coated using a gold EM Scope SC500 Au coater but materials such as gold/palladium alloy, platinum, osmium, iridium, tungsten, chromium and graphite can also be used. The biomass must be electrically conductive, at least at the surface, and electrically grounded to prevent the accumulation of electrostatic charge at the surface. For additional information on SEM analysis refer to a review available on the Internet at http://serc. carleton. edu/research_education/geochemsheets/techniques/SEM. html. This online review also refers to the literature that further explores SEM.

SEM analysis is a useful tool to examine the effects of pre-treatments and enzymatic hydrolysis on the structure of the plant cell wall and has been used by several researchers for this purpose (Gomez et al. 2008; Jieben et al. 2011).

The gold coated biomass samples were analysed using a Hitachi S-570 SEM and suitably magnified images were recorded. Dilute acid pretreatment may affect biomass structure by solubilising or altering hemicelluloses, altering lignin struc­ture and increasing the available surface area and pore volume of the substrate. The effects of various mild acid pretreatments are shown in images A to N, Fig. 4.10.

Images A and B show untreated grass. It is clear that the cells are well struc­tured the fibres that make up the structure of the grass are connected very tightly. After treatment with just water (hot liquid pretreatment), the cells are generally still well structured and the fibres are still tightly connected (images C & D). After acid hydrolysis, the SEM images show the grass biomass has been affected by all acid pretreatments. In images E, F, G, H, K and L, the cells seem less structured and organised and the fibres seem less tightly connected. However, treatment with nitric acid seems to have had a very destructive effect on the grass biomass. This would correlate with the results of Fig. 4.9 which shows greatest sugar release was achieved by treatment with nitric acid. Image I shows how the grass fibres have completely come apart and in image J areas of the structure have become weak­ened, with pores starting to appear. Treatment with sulphuric acid also seems to

have been a very effective pretreatment as it is clear some cell tissues have been destroyed and very definite pores have appeared in the grass structure (images M and N). The destruction of the grass structure shown in these images may be attributed to the preferential degradation of the labile components such as hemi — celluloses and acid soluble lignin.

Pre-treating grass biomass with dilute acid is a favorable process as it helps remove the hemicelluloses fraction and disrupt the grass structure which allows greater accessibility for the cellulase enzymes. It may also help lead to less hemicelluloses and lignin content in the cellulose preparation for the acid hydrolysed perennial rye grass.

4.3 Conclusion

The effect of pretreatments is very dependent on the biomass composition. This chapter focused on reviewing in detail the composition of the grass plant cell wall. Mild acid pre-treatments were reviewed and several mild acid pretreatment con­ditions were tested. The resulting acid hydrolysate sugar yields were noted and the pretreated biomass residues were subjected to SEM analysis to take a closer look at the effects of acid pretreatments, specifically on the plant cell wall. These pre­treatment processes should make the lignocellulosic biomass more susceptible to enzymatic attack, where crystallinity of cellulose, its accessible surface area and protection by lignin and hemicellulose are the main factors in order to obtain an efficient hydrolysis.

Acknowledgments I would like to acknowledge Pierce Lalor and the Centre for Microscopy and Imaging at the National University of Ireland Galway for the use of and assistance with a Scanning Electron Microscope

References

Anderson W, Akin DE (2008) Structural and chemical properties of grass lignocelluloses related to conversion for biofuels. J Ind Microbiol Biotechnol 35:355-366 Alberts B, Bray D, Lewis J, Raff M, Roberts K, Watson JD (1989) Molecular biology of the cell, biochemical education, vol 18. Garland Publishing, New York. doi:10.1002/bmb.1990.5690180128 Begum P, Aubert JP (2000) Cellulases. In: Lederberg J (ed) Encyclopedia of microbiology. Academic, San Diego, pp 744-758

Bertoldo C, Antranikian G (2002) Starch-hydrolyzing enzymes from thermophilic archaea and bacteria. Curr Opin Chem Biol 6:151-160

Brett C, Waldron K (1996) Physiology and biochemistry of plant cell walls, 2nd edn. Chapman and Hall, London

Cara C, Ruiz E, Oliva JM, Saez F, Castro E (2008) Conversion of olive tree biomass into fermentable sugars by dilute acid pre-treatment and enzymatic saccharification. Bioresour Technol 99:1869-1876. doi:10.1016/j. biortech.2007.03.037 Carpita NC (1996) Structure and biogenesis of the cell walls of grasses. Ann Rev Plant Physiol Plant Molec Biol 47:445-476

Carpita NC, Gibeaut DM (1993) Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth. Plant J 3:1-30

Carpita NC, McCann M (2000) In: Buchanan BB, Gruissem W, Jones RJ (eds) Biochemistry and molecular biology of plants. American Society of Plant Biologists, Rockville, MA, pp 25-109 Comparot-Moss S, Denyer K (2009) The evolution of the starch biosynthetic pathway in cereals and other grasses. J Exp Bot 60:2481-2492. doi:10.1093/jxb/erp141 Cosgrove DJ (2005) Growth of the plant cell wall. Nature 6:850-861

Cowling EB (1974) Physical and chemical constraints in the hydrolysis of cellulose and lignocellulosic materials. Biotechnol Bioeng Symp 5:163-181 Ding SY, Himmel ME (2006) The maize primary cell wall microfibril: a new model derived from direct visualization. J Agric Food Chem 54:597-606 Dinges JR (2001) Molecular structure of three mutations at the maize sugary1 locus and their allele-specific phenotypic effects. Plant Physiol 125:1406-1418. doi:10.1104/pp.125.3.1406

DuBois M, Gilles KA, Hamilton JK, Rebers PA, Smith F (1956) Colorimetric method for determination of sugars and related substances. Anal Chem 28:350-356 (American Chemical Society). doi:10.1021/ac60111a017

Ebringerova A et al (2005) Hemicellulose. In: Heinze T (ed) Advances in polymer science, vol 186. Springer-Verlag, Berlin/Heidelberg, pp 1-67. doi:10.1007/b136812 Endler A, Persson S (2011) Cellulose synthases and synthesis in arabidopsis. Molecular Plant 4:199-211. doi: 10.1093/mp/ssq079

Faik A (2010) Xylan biosynthesis: news from the grass. Plant Physiol 153:396-402. doi:10.1104/ pp.110.154237

Gharpuray MM, Lee YH, Fan LT (1983) Structural modification of lignocellulosics by pre­treatments to enhance enzymatic-hydrolysis. Biotechnol Bioeng 25:157-172 Gomez LD, Bristow JK, Statham ER, McQueen-Mason SJ (2008) Analysis of saccharification in brachypodium distachyon stems under mild conditions of hydrolysis. Biotechnol Biofuels 1:15

Hendriks ATWM, Zeeman G (2009) Pretreatments to enhance the digestibility of lignocellulosic biomass. Bioresour Technol 100:10-18. doi:10.1016/j. biortech.2008.05.027 Heredia A, Jimenez A, Guillen R (1995) Composition of plant cell walls. Zeitschrift Fur Lebensmittel-Untersuchung Und-Forschung 200:24-31 Hernon AT, O’Donovan A, Shier MC (2010) Applications of Fungal Enzymes In Bioconversion. In: Gupta VK, Tuohy MG, Gaur RK (eds) Fungal biochemistry and biotechnology. Lambert Academic Publishing, Germany. ISBN no: 978-3-8433-5800-2 Iiyama K, Lam T, Stone B (1990) Phenolic-acid bridges between polysaccharides and lignin in wheat internodes. Phytochemistry 29:733-737

Ishii T (1999) The plant cell wall polysaccharide rhamnogalacturonan II self-assembles into a covalently cross-linked dimer. J Biol Chem 274:13098-13104. doi:10.1074/jbc.274.19.13098 Jieben T, Kefu C, Jun X, Jun L, Chuanshan Z (2011) Effects of Dilute acid hydrolysis on composition and structure of cellulose in Euliopsis Binata. Bioresources 6:1069-1078 Kato Y, Ito S, Iki K, Matsuda K (1982) Xyloglucan and r{beta}-D-glucan in cell walls of rice seedlings. Plant Cell Physiol 23:351-364

Kobayashi M, Matoh T, Azuma J (1996) Two chains of rhamnogalacturonan II are cross-linked by borate-diol ester bonds in higher plant cell walls. Plant Physiol 110:1017-1020. doi:10.1104/pp.110.3.1017

Kumar D, Murthy GS (2011) Impact of pre-treatment and downstream processing technologies on economics and energy in cellulosic ethanol production. Biotechnol Biofuels 4:27 Morikawa K, la Cour TF, Nyborg J, Rasmussen KM, Miller DL, Clark BF (1978) High resolution x-ray crystallographic analysis of a modified form of the elongation factor Tu: guanosine diphosphate complex. J Mol Biol 125:325-338

Nishitani K (1997) The role of endo xyloglucan transferase in the organization of plant cell walls. Int Rev Cytol 173:157-206

Nishitani K, Nevins D (1991) Glucuronoxylan xylanohydrolase. A unique xylanase with the requirement for appendant glucuronosyl units. J Biol Chem 266:6539-6543 Nishitani K, Nevins DJ (1989) Enzymic analysis of feruloylated arabinoxylans (Feraxan) derived from zea mays cell walls: II. fractionation and partial characterization of feraxan fragments dissociated by a Bacillus subtilis Enzyme (Feraxanase). Plant Physiol 91:242-248. doi:10.1104/pp.91.1.242

O’Neill MA, Eberhard S, Albersheim P, Darvill AG (2001) Requirement of borate cross-linking of cell wall rhamnogalacturonan II for arabidopsis growth. Science 294:846-849, (New York, N. Y.). doi:10.1126/science.1062319

Reiter WD (2002) Biosynthesis and properties of the plant cell wall. Curr Opin Plant Biol 5:536­542

Richardson S, Gorton L (2003) Characterisation of the substituent distribution in starch and cellulose derivatives. Anal Chim Acta 497:27-65 Ridley BL, Mohnen DA, O’Neill MA, O’Neill Mohnen (2001) Pectins: structure, biosynthesis, and oligogalacturonide-related signalling. Phytochemistry 57:929-967

Somerville C, Bauer S, Brininstool G, Facette M, Hamann T, Milne J, Osborne E et al. (2004) Toward a systems approach to understanding plant cell walls. Science 306:2206-2211, (New York, N. Y.). doi:10.1126/science.1102765

Sun Y, Cheng J (2002) Hydrolysis of lignocellulosic materials for ethanol production: a review. Bioresour Technol 83:1-11

Taherzadeh MJ, Karimi K (2008) Pre-treatment of lignocellulosic wastes to improve ethanol and biogas production: a review. Int J Mol Sci 9:1621-1651. doi:10.3390/ijms9091621 Teeri T (1997) Crystalline cellulose degradation: new insight into the function of cellobiohy — drolases. Tibtech 15:160-167

Timell TE (1964) Wood hemicellulose. Adv Carbohydr Chem 19:247-302 Tsmousis G (1991) Science and technology of wood; structure, properties, utilization. Van Nostrand Reinhold, New York

Vogel J (2008) Unique aspects of the grass cell wall. Curr Opin Plant Biol 11:301-307. doi:10.1016/j. pbi.2008.03.002

Vries RP, Visser J (2001) Aspergillus enzymes involved in degradation of plant cell wall polysaccharides. Microbiol Mol Biol Rev 65:49 Willats WG, McCartney LM, Mackie W, Knox JP (2001) Pectin: cell biology and prospects for functional analysis. Plant Mol Biol 47:9-27

Yang B, Wyman CE (2008) Pre-treatment: the key to unlocking low-cost cellulosic ethanol. Environ Eng 2:26-40. doi:10.1002/bbb

Zhao H, Kwak JH, Conrad Zhang Z, Brown HM, Arey BW, Holladay JE (2007) Studying cellulose fiber structure by SEM, XRD, NMR and acid hydrolysis. Carbohydr Polym 68:235 http://serc. carleton. edu/research_education/geochemsheets/techniques/SEM. html http://www. ccrc. uga. edu http://cellwall. genomics. purdue. edu http://nutrition. jbpub. com

Part III

Several species of cellulolytic fungi, such as Trichoderma reesei, naturally pro­duce a large repertoire of saccharolytic enzymes to digest lignocellulose effi­ciently, assimilate all lignocellulosic sugars, and convert these sugars into ethanol, showing that they naturally possess all pathways for conversion of lignocellulose into bioethanol (Chambergo et al. 2002; Lynd et al. 2002). It has been shown that a biorefinery consuming thousands of tons of biomass per day will require many tons of cellulase preparation to operate assuming that enzymes with far greater specific activity are not identified (Xu et al. 2009). Currently, only fungi naturally produce the required amounts of cellulase and some strains of T. reesei produce more than 100 g/L cellulase (Cherry and Fidantsef 2003). Thus, advantages of T. reesei as a CBP organism are: (1) the production of sufficient quantities of

Table 8.3 Characteristics of an ideal CBP organism

1. Ability to ferment all hexoses and pentoses present in lignocellulose

2. High product yield, titer, and productivity

3. High product and inhibitor tolerance

4. General robustness for industrial processes, cellulase production in toxic environment

5. Tolerance to low pH and high temperature

6. Amenability to DNA manipulation

7. High levels of heterologous protein production and secretion (if cellulolytic ability must be

engineered)

8. Concurrent fermentation of sugars (hexose and pentose co-utilization)

9. GRAS status

10. Recyclability in successive processes

11. Minimum nutrient supplementation requirement

Van Zyl et al. (2007) cellulases at reasonable cost, (2) several strains are established commercially, and (3) it can utilize all lignocellulose sugars for production of ethanol. Challenges to overcome before T. reesei can be considered as a CBP organism include the observations that ethanol yield, rate of production and tolerance are low, and that mixing during fermentation may require more energy owing to the filamentous cell morphology. Preliminary studies showed that T. reesei could produce cellulases when grown aerobically on cellulose that continued to degrade cellulose to sugars and ferment these sugars to ethanol when cultures were rendered anaerobic (Xu et al. 2009). However, acetic acid was produced as a major byproduct. The major limitation for efficient ethanol production by T. reesei does not lie in the absence of the relevant genes and pathways but are more likely related to the low expression of these genes or the activity of the enzymes encoded. Approaches to solving these problems are to enhance the expression of the relevant genes at the transcriptional level and/or to introduce heterologous genes that encode enzymes with higher activities and to knockout genes responsible for the production of byproducts. Recently, the laccase gene lacA from Trametes sp. AH28-2 was heterologously expressed in T. reesei under control of a constitutive promoter (Zhang et al. 2012). Transformants were identified that were able to secrete the recombinant laccase. Reducing sugar yields obtained from saccharification of corn residue by crude enzyme extracts prepared from the transformants increased by 31.3-71.6 %, respectively, compared to the host strain.

Another filamentous fungus, Fusarium oxysporum, also produces the enzymes required to break down cellulose and hemicellulose while simultaneously fer­menting the released sugars to ethanol albeit at relatively low yields (Anasontzis et al. 2011; Panagiotou et al. 2005). In SSF of a cellulosic substrate a F. oxysporum wild-type strain was able to grow in aerobic conditions and produced ethanol with a yield of 0.35 g/g cellulose under anaerobic conditions. The strain was also shown to effectively produce a complete system of hydrolytic enzymes when grown on various agro-industrial lignocellulose by-products, such as dry citrus peels, corn cob, and brewer’s spent grain with concomitant ethanol production (Anasontzis et al. 2011; Xiros et al. 2008). It was hypothesized that homologous overexpres­sion of cellulases and hemicellulases under constitutive control, could provide a higher breakdown rate of the biomass and thus increase the supply of sugars to the ethanol production pathway. To this end, the endoxylanase two of F. oxysporum, was overproduced under control of the constitutive Aspergillus nidulans gpdA promoter (Anasontzis et al. 2011). The fermentative performance of the trans­formants were evaluated and compared to that of the wild type in simple CBP systems using corn cob or wheat bran as sole carbon sources. Transformants produced approximately 60 % more ethanol compared to the wild type on corn cob and wheat bran likely due to the * 2-2.5-fold higher extracellular xylanase activities in the fermentation broths of the transformants.

High-temperature conversion process conditions potentially provide a significant energy saving since reactors would not have to be cooled to mesophilic conditions before inoculation and then reheated for distillation (Xu et al. 2010). Furthermore, it has been shown that a 10 °C increase in temperature approximately doubles enzymatic reaction rates, decreasing the amount of enzyme required (Ibrahim and El-diwany 2007). In addition, the use of reaction and fermentation temperatures in excess of 60 °C minimizes the risk of contamination. Since cellulose hydrolysis and sugar release is in most cases the rate-limiting step in a typical CBP process, high — temperature hydrolysis will be therefore be advantageous. Thermophilic bacteria capable of cellulose hydrolysis and ethanol production show great potential as CBP organisms (Xu et al. 2010). The cellulosome producing thermophilic Gram-positive anaerobic bacterium C. thermocellum is regarded as a potential CBP organism as it is very efficient at hydrolyzing crystalline cellulose (Lynd et al. 2002). While growth of wild-type strains is inhibited in the presence of ethanol concentrations above 2 % (v/v) strains have been evolved that remained viable at ethanol concentrations of up to 8 % (v/v) (Xu et al. 2010). This group also investigated the effect of some of these inhibitors on cellulosome activity of C. thermocellum. It was shown that that some organic acids actually promoted cellulolytic activity and that the C. thermocellum cellulosome could tolerate certain concentrations of furfural (up to 5 mM), p-hydroxybenzoic acid (up to 50 mM) and catechol (up to 1 mM). The C. thermo — cellum cellulosomes were also able to tolerate higher ethanol concentrations and temperatures than the T. reesei enzymes used commercially.