The hydrolysis methods most commonly used are acid and enzymatic. Both dilute and concentrated acid is used. Dilute acid treatment is employed for the degra­dation of hemicellulose leaving lignin and cellulose network in the substrate. Other treatments are alkaline hydrolysis or microbial pretreatment with white-rot fungi (Phaenerochate chrysosporium, Cyathus stercoreus, Cythus bulleri, and Pycnoporous cinnabarinus, etc.) preferably act upon lignin leaving cellulose and hemicellulose network in the residual portion. However, during both treatment processes, a considerable amount of carbohydrates are also degraded; hence, the carbohydrate recovery is not satisfactory for ethanol production. The dilute acid process is conducted under high temperature and pressure and has reaction time in the range of seconds or minutes. The concentrated acid process uses relatively mild temperatures, but at high concentration of sulfuric acid and a minimum pres­sure involved, which only creates by pumping the materials from vessel to ves­sel. Reaction times are typically much longer than for dilute acid. In dilute acid hydrolysis, the hemicellulose fraction is depolymerized at lower temperature than the cellulosic fraction. Dilute sulfuric acid is mixed with biomass to hydrolyze hemicellulose to xylose and other sugars. Dilute acid is interacted with the bio­mass, and the slurry is held at temperature ranging from 120-220 °C for a short period of time. Thus, hemicellulosic fraction of plant cell wall is depolymerized and will lead to the enhancement of cellulose digestibility in the residual solids (Nigam 2002; Sun and Cheng 2002; Dien et al. 2006; Saha et al. 2005). Dilute acid hydrolysis has some limitations. If higher temperatures (or longer residence time) are applied, the hemicellulosic derived monosaccharides will degrade and give rise to fermentation inhibitors like furan compounds, weak carboxylic acids, and phenolic compounds (Olsson and Hahn-Hagerdal 1996; Klinke et al. 2004;

Larsson et al. 1999). These fermentation inhibitors are known to affect the etha­nol production performance of fermenting microorganisms (Chandel et al. 2007). In order to remove the inhibitors and increase the hydrolyzate fermentability, sev­eral chemicals and biological methods have been used. These methods include overliming (Martinez et al. 2000), charcoal adsorption (Chandel et al. 2007), ion exchange (Nilvebrant 2001), detoxification with laccase (Martin et al. 2002; Chandel et al. 2007), and biological detoxification (Lopez et al. 2004). The detoxi­fication of acid hydrolyzates has been shown to improve their fermentability; however, the cost is often higher than the benefits achieved (Palmqvist and Hahn — Hagerdal 2000; von Sivers et al. 1994). Dilute acid hydrolysis is carried out in two stages—First-stage and two-stage.

Enzymatic hydrolysis using cellulases does not generate inhibitors, and the enzymes are very specific for cellulose. Cellulases, mainly derived from fungi like Trichoderma reesei and bacteria like Cellulomonas fimi, are a mixture of at least three different enzymes: (1) endoglucanase which attacks regions of low crystallinity in the cellulose fiber, creating free chain-ends; (2) exoglucanase or cellobiohydrolase which degrade the molecule further by removing cellobiose units from the free chain-ends; and (3) в-glucosidase which hydrolyzes cellobi — ose to produce glucose, in much smaller amounts. Cellulase enzyme, however, has been projected as a major cost contributor to the lignocellulose-to-ethanol technology. The main challenges are the low glucose yield and high cost of the hydrolysis process. Cellulase costs around 15-20 cents per gallon ethanol as compared to only 2-4 cents per gallon ethanol for amylases used in the starch-to — ethanol process, and ethanol production from sugarcane and molasses bypasses this cost entirely. The cellulase requirement is also much higher than for amyl­ases. The complex three-dimensional (crystalline) structure of cellulose slows down the rate of hydrolysis dramatically; starch hydrolysis by amylases is 100 times faster than cellulose hydrolysis by cellulases under industrial processing conditions.

The US Department of Energy (DOE), in 1999, signed contracts (worth $17 million and $12.3 million, respectively) with Genencor International and Novozymes Inc., to increase cellulase activity and bring down the cost of cellulose enzymes. Novozymes has launched CellicTM, a new class of cellulases, which is claimed to be more cost-effective and four times more active compared to the previously produced cellulases (www. bioenergy. novozymes. com). On similar lines, Genencor International has announced the launch of Accellerase 1,500 on February 23, 2009. This is claimed to have significantly improved formulation and higher activity resulting in higher ethanol yields and robust operation in a wider variety of processes. The product is based on Genencor’s patented gene expression Technology.

Enzymatic hydrolysis is highly specific and can produce high yields of rela­tively pure glucose syrups, without generation of glucose degradation products. Utility costs are low, as the hydrolysis occurs under mild reaction conditions. The process is compatible with many pretreatment options, although purely physical methods are typically not adequate (Graf and Koehler 2000; Sun and Cheng 2002).

Many experts see enzymatic hydrolysis as key to cost-effective ethanol produc­tion in the long run (US DOE 2003). Although acid processes are technically more mature, enzymatic processes have comparable projected costs and the potential of cost reductions as technology improves (Lynd et al. 1999).

The rate and extent of enzymatic hydrolysis of lignocellulosic biomass depends on enzyme loadings, hydrolysis periods, and structural features result­ing from pretreatments. The extent of cellulose conversion is dictated by the degree of attachment of the cellulase enzymes to the three-dimensional cel­lulose surface. Crystallinity of cellulose affects the initial rate of enzymatic hydrolysis while presence of lignin and acetyl groups limits the extent of cel­lulose hydrolysis due to unproductive binding of cellulase with lignin and the acetyl groups (Ghose and Bisaria 1979). Thus, an efficient pretreatment strat­egy that decreases cellulose crystallinity and removes lignin to the maximum extent can significantly decrease hydrolysis time as well as cellulase loading. The problem of non-specific binding of cellulases to lignin can be circum­vented by adding non-ionic surfactants like Tween 20. It has been shown that surfactant addition can increase the ethanol yield by 8 % and reduce cellulase loading by as much as 50 % while maintaining a constant ethanol yield. The proposed mechanism for this enhanced enzyme performance is the adsorption of the surfactant onto lignin surface, thereby preventing unproductive binding of the enzymes to lignin. The adsorption also helps in retaining the enzymes for recycle. Cheaper alternatives to non-ionic surfactants like Tween 20 can reduce the process cost considerably.

Another promising approach to enhancing enzymatic hydrolysis of pretreated biomass is the use of ultrasonic energy. The cavitational effect of ultrasound leads to biomass swelling and fragmentation, increasing the accessibility of cel — lulases to cellulosic substrate (Ebringerova and Hromadkova 2002). Intermittent ultrasonication of biomass before and after enzymatic hydrolysis has been shown to significantly increase the rate of reaction. Approximately 20 % increase in eth­anol yield and 50 % decrease in enzyme loading were observed on intermittent exposure of simultaneous saccharification and fermentation processes of mixed waste office paper to ultrasonic energy under selected conditions (Banerjee et al. 2010). Saccharomyces cerevisiae is by far the most commonly used micro­bial species for industrial ethanol production from sugar- and starch-based raw materials and is well adapted to the industrial scenario. It produces ethanol with stoichiometric yields and tolerates a wide spectrum of inhibitors and elevated osmotic pressure. Zymomonas mobilis has been projected as the future etha — nologen due to its high ethanol tolerance (up to 14 % v/v), energy efficiency, high ethanol yield (up to 97 % of theoretical), and high ethanol productivity. Complete substrate utilization is a prerequisite for economically competitive lignocellulosic fermentation processes. The native ethanologen S. cerevisiae is capable of fermenting only hexoses and cannot utilize pentoses like xylose, which is the main component of the hemicellulosic fraction of lignocellu — lose, and can contribute to as much as 30 % of the total biomass. Z. mobilis can only utilize glucose, fructose, and sucrose. Expanding the substrate range of whole-cell biocatalysts will greatly contribute to the economic feasibility of bioethanol production from renewable feedstock. The essential traits of a good lignocellulose-to-ethanol bioconverter are—utilization of both hexoses and pen­toses; high ethanol yields and productivity; minimum by-product formation; high ethanol tolerance; and tolerance to other inhibitors formed during biomass pre­treatment and hydrolysis. Four industrial benchmarks for ethanologenic strain development, which have the greatest influence on the price of lignocellulosic ethanol are—Process water economy, Inhibitor tolerance, Ethanol yield, Specific ethanol productivity (Banerjee et al. 2010).

The inability of S. cerevisiae and Z. mobilis to utilize pentose sugars neces­sitates the search for pentose utilizing strains. Two groups of microorganisms, i. e., enteric bacteria and some yeasts, are able to ferment pentoses, but with low ethanol yields (Bajpai and Margaritis 1982). Furthermore, xylose ferment­ing yeasts (Pachysolen tannophilus, Candida shehatae, and Pichia stipitis) are sensitive to high concentrations of ethanol (>40 g/l), require microaerophilic conditions, are highly sensitive to inhibitors, and are not capable of fermenting xylose at low pH.

Due to the lack of a natural microorganism for efficient fermentation of ligno — cellulose-derived substrates, there has been emphasis on constructing an efficient organism through metabolic engineering of different organisms (Banerjee et al. 2010). Metabolic engineering, by virtue of the recent molecular biology tools, has generated recombinant organisms displaying attractive features for the bioconver­sion of lignocelluloses to ethanol. The three most promising microbial species that have been developed by metabolic engineering in the last two decades are S. cer — evisiae, Z. mobilis, and Escherichia coli. The recombinant organisms developed for lignocellulose-to-ethanol process have been extensively reviewed in earlier works. Significant work on the metabolic engineering of E. coli has been com­pleted. The incorporation and expression of pyruvate decarboxylase and alcohol dehydrogenase II genes from Z. mobilis into E. coli under the control of a common (lac) promoter results in high ethanol yield due to co-fermentation of hexose and pentose sugars. Xylose isomerase (XI), encoded by the xylA gene, catalyzes the isomerization of xylose to xylulose in bacteria and some fungi. Recent develop­ments to improve XI activity and heterologous expression in S. cerevisiae through adaptation and metabolic manipulation have proven to be successful with ethanol yields approaching the theoretical maximum. Genes xylA and xylB, part of the xyl operon of E. coli, were introduced into Pseudomonas putida S12 under the transcriptional control of the constitutive tac promoter. This recombinant strain, after further laboratory evolution, could efficiently utilize the three, most abundant sugars in lignocellulose, glucose, xylose, and arabinose, as sole carbon sources. Whereas recombinant laboratory strains are useful for evaluating metabolic engi­neering strategies, they do not possess the robustness required in the industrial context. The strategies of producing industrial pentose fermenting strains involve the introduction of the initial xylose and arabinose utilization pathways and adap­tation strategies, including random mutagenesis, evolutionary engineering, and breeding.

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