1.5. Optimization of carrier flow rate and sample volume

Good peak reproducibility was achieved when samples were injected into the air flow as carrier. When samples were injected into 0.14 mol L-1 HNO3, used as the carrier, there was a rise in the baseline (as expected, due to increase of the blank), followed by a fall due to cool­ing of the metal or ceramic tubes. This cooling was significant, since no transient signals were obtained following injection of standards, indicating that the temperature within the tubes was insufficient to atomize the analyte, which remained dispersed in the carrier solu­tion. This confirmed the findings of earlier work that the use of air (or other gas) as the carri­er avoids dilution and dispersion of the sample. Here, all analyses were performed using air as the carrier, not only because it was less expensive than use of a solution, and minimized waste generation, but also because it enabled the TS-FF-AAS system to be used to determine copper, which would not have been possible using a solution as the carrier.

Figure 2 shows the influence of the carrier (air) flow rate, in the range 9.0-18.0 mL min-1, on the absorbance values obtained using 50 |jL of a standard of 200 |jg Cu L-1in 0.14 mol L-1 HNO3, using both tubes. In the case of the metal tube, lower absorbance values were ob­tained at low flow rates, because the sample arrived slowly at the atomizer, increasing the measurement duration and resulting in an unpredictable and erratic vaporization. Hence, as the flow rate was increased, the absorbance also increased due to a more homogeneous va­porization of the sample [23,27,58].

This increase proceeded up to a carrier flow rate of 12.0 mL min-1, above which there was no significant variation in absorbance. The highest absorbance value was obtained at a flow rate of 18.0 mL min-1, which was therefore selected as the best flow rate to use with the metal tube.

When the ceramic tube was used, maximum absorbance was achieved at a carrier flow rate of 9.0 mL min-1. At higher flow rates, the residence time of the liquid in the heated section of the ceramic capillary was considerably diminished, reducing the time available for evapora­tion of the liquid, so that the sample was not delivered in the form of vapor/aerosol, but rather as a flow of liquid. The temperature within the tube decreased, and the color of the tube changed from ruby-red to opaque grey. It was also possible to see droplets emerging from the atomizer tube. Hence, the absorbance values did not increase, while greater varia­bility in the signal resulted in elevated standard deviation values. A flow rate of 3.0 mL min-1was selected, at which the absorbance signal was maximized, and the standard devia­tion was minimized.

The sample volume was varied between 50 and 200 |jL, using carrier flow rates of 18.0 and 9.0 mL min-1 for the metal and ceramic tubes, respectively. The results (Figure 3) re­vealed that for both tubes a sample volume of 50 |jL generated the highest absorbance value, with a low standard deviation, reflecting good repeatability in the experimental measurements. When 100 |jL of sample was used, there was a slight cooling of the ceram­ic capillary, and consequently of the atomization tubes, while there was no increase in the absorbance values. At a sample volume of 200 |jL, the ceramic capillary and the tube were substantially cooled, and there was no homogeneous thermospray formation, with erratic generation of droplets that acted to disperse the light radiation (probably to a large degree, since the deuterium lamp was unable to fully correct the resulting background signal). The unpredictable atomization resulted in very high standard deviation values. Using air as the carrier, a sample volume of 50 |jL was selected for the subsequent meas­urements, due to greater atomization homogeneity, satisfactory absorbance for a 30 mg Cu L-1 standard, and a low SD value.

Plant biomass can be used as a sustainable source of organic carbon to create bioenergy, ei­ther directly in the form of heat and electricity, or as liquid biofuels produced by thermo­chemical or biochemical methods or their combination [12]. In contrast to fossil energy sources, which are the result of long-term transformation of organic matter, plant biomass is created via photosynthesis using carbon dioxide as a source of carbon and sunlight as a source of energy and therefore is rapidly produced. The world annual production of bio­mass is estimated to be 146 billion metric tons [13], which could contribute 9-13% of the global energy supply yielding 45±10 EJ per year [14, 15].

Lignocellulose, which is stored in plant cell walls makes up a significant part of biomass representing 60-80% of woody tissue of stems, 15-30% of leaves or 30-60% of herbal stems [16]. Since it is not digestible for human beings, its use as a feedstock for bioprocesses does not compete with food production as in the case of sugar or starch raw materials.

All lignocellulose consist of three main polymeric components — cellulose, non-cellulosic carbohydrates (predominantly represented by hemicellulose) and lignin; its proportion and structure differs for different types of biomass (Tablel) and it is also influenced by variety, climatic conditions, cultivation methods and location. Minor components of the cell wall are represented by proteoglycans, pectin, starch, minerals, terpenes, resins tannins and waxes.

Cellulose is a homopolymer of 500-1 000 000 D-glucose units (e. g. 10 000 units in wood, 15 000 in native cotton) linked by p-1,4-glycosidic bonds [19, 26, 29]; the cellulose chains (200-300) are grouped together to form cellulose fibres. The strong inter-chain hydrogen bonds between hydroxyl groups of glucose residues in radial orientation and the aliphatic hydrogen atoms in axial positions creates a semi-crystalline structure resistant to enzymatic hydrolysis; weaker hydrophobic interactions between cellulose sheets promote the forma­tion of a water layer near the cellulose surface, which protects cellulose from acid hydrolysis [30]. Cellulose originating from different plants has the same chemical structure, but it dif­fers in crystalline structure and inter-connections between other biomass components. Mi­crofibrils made of cellulose are surrounded by covalently or non-covalently bound hemicellulose, which is a highly branched heteropolymer made from 70-300 monomers units of pentoses (xylose, arabinose), hexoses (galactose, glucose, mannose) and acetylated sugars (e. g. glucuronic, galacturonic acids). Unlike cellulose, hemicellulose is not chemically homogenous and its composition depends on the type of material — hardwood contains pre­dominantly xylans while softwood consists mainly of glucomannans [17, 23, 29,31]. Lignin, an amorphous heteropolymer of three phenolic monomers of phenyl propionic alcohols, namely p-coumaryl, coniferyl and sinapylalcohol, creates a hydrophobic filler, which is syn­thesized as a matrix displacing water in the late phase of plant fibre synthesis, and forms a layer encasing the cellulose fibres. Its covalent crosslinking with hemicellulose and cellulose forms a strong matrix, which protects polysaccharides from microbial degradation, makes it resistant to oxidative stress, and prevents its extraction by neutral aqueous solvents [31]. Forest biomass has the highest content of lignin (30-60% and 30-55 % for softwoods and hardwoods, respectively), while grasses and agricultural residues contain less lignin (10-30% and 3-15% respectively) [17].

There are several groups of lignocellulosic plant biomasses that can be exploited as a feed­stock for bioprocessing. Woody biomass is represented mainly by hardwoods (angiosperm trees, e. g. poplar, willow, oak, cottonwood, aspen) and softwoods (conifers and gymno — sperm trees e. g. pine, cedar, spruce, cypress, fir, redwood) together with forest wastes such as sawdust, wood chips or pruning residues. Nowadays the trend in this area is to use fast growing trees (poplar, willow) with genetically changed wood structures e. g. lower lignin content [32]. The advantage of forest biomass is its flexible harvesting time, thus avoiding long storage periods, and its high density, contributing to cost-effective transportation. Agri­cultural residues are represented mainly by corn stover or stalks, rice and wheat straw or sugarcane bagasse. The world’s annual production of rice straw, wheat straw and corn straw that can be exploited for bioethanol production is 694.1, 354.3 and 203.6 million tons, respectively [20]. In the USA, 370 million and 350-450 million tons of forest biomass and ag­ricultural wastes respectively are produced per year [17]. Although agrowastes are partly reutilized, e. g. as animal fodder, bedding, domestic fuel, used for cogeneration of electricity or reused in agriculture, a large fraction is still disposed as waste and is left in the fields; this can be utilized as a raw material for biofuels production. Sugarcane is nowadays one of the most important feedstocks for production of 1st generation bioethanol and also one of the plants with the highest photosynthetic efficiency, yielding around 55 tons of dry matter per hectare annually (approx. 176 kg/ha/day). Sugar cane bagasse, the fibrous lignocellulosic material remaining as waste is mostly used as a solid fuel in sugar mills or distilleries but due to its high cellulose content (Table 1) it can be reutilized as a feedstock for production of 2nd generation bioethanol. In the sugarcane season of 2010/11, the total sugar cane crop reached almost 1.627 billion tons (on 23 million hectares), which corresponds to 600 million tons of wet sugar cane bagasse [33]. Minor, but also important residues are the leaves, called sugarcane trash, amounting to 6-8 tons per hectare of sugarcane crop [34]. Another group of lignocellulosic biomass, herbaceous energy crops and grasses, which are represented pre­dominantly by switch grass, alfalfa, sorrel or miscanthus [24], are interesting due to their low demands on soil quality, low-cost investments, fast growth, low moisture content, high yield per hectare (e. g. 20 t/ha for miscanthus) and high carbohydrate content (Table 1). Be­sides lignocellulosic plant materials, other low-cost large volume feedstocks such as munici­pal solid waste, municipal wastewater, food-processing waste or waste from the paper industry can be utilized for bioethanol production. Mixed municipal recovery solid waste (MSW) consists of approximately 55% mineral waste, 6% of metallic waste, 5% animal and vegetable waste (food residues, garden waste), 3% of paper and cardboard waste and 31% of others [35]. In the EU alone, the annual production of municipal wastes amounts 2.6 million tons, 65% of which is derived from renewable resources [35, 36]. The main challenge in its bioprocessing is its heterogeneous composition. To be used for ethanol production, degrada­ble fractions of MSW should be separated after sterilization; cellulosic material (paper, wood or yard waste) represents approximately 60% of the dry weight of typical MSW as shown in Table 1 [25, 37]. Beside the solid wastes, lignocellulose extracted from municipal wastewater treatment processes can also be used as low-cost feedstock for biofuel production [26]. In Canada, 6.22 Mt of sugar could be annually produced using municipal sludge/biosolids and livestock manures [26]. Municipal wastewaters, which include faecal materials, scraps of toi­let paper and food residues, should be pre-treated to separate solid and liquid fractions, the former of which is processed further to gain simple sugars. Primary sludge contains more cellulose compared to activated sludge (Table 1) because it is consumed in the activated sludge process and is further degraded by anaerobic digestion processes in the sewage dis­posal plant [26]. When talking about industrial wastes as 2nd generation raw materials for biofuels, wastes from cellulose/paper production cannot be neglected. Paper sludge is waste solid residue from wood pulping and papermaking processes and is represented by poor — quality paper fibres, which are too short to be used in paper machines. It is attractive as a raw material for bioprocessing mainly due to its low cost (it is currently disposed of in land­fills or burned), its high carbohydrate content (Table 1) and its structure, which doesn’t re­quire any pretreatment [8, 27, 28]. Another waste is represented by sulphite waste liquor (SWL), a solution of monomeric sugars formed during the sulfite pulping process by disso­lution of lignin and most hemicelluloses. About 1 ton of solid waste is dissolved in SWL (11-14% solids) per ton of pulp and its annual production is around 90 billion litres [38]. SWL is usually burned after its concentration and evaporation, but since it’s main compo­nents are sugars and lignosulfonates, its use as a raw material for bioethanol production has potential. Chemical composition of SWL (a spectrum of fermentable sugars, inhibitors, nu­trients and minerals) differs significantly with the type of wood and technological proce­dures, e. g. concentration of the main sugars in SWL (% of dry matter) ranges for xylose from 3 to 5 % in soft wood (spruce, western hemlock) up to 21 % in eucalyptus, the highest con­centration of galactose and glucose around 2.5 % is in soft wood SWL, content of mannose can reach values of almost 15 % in soft wood SWL [39—42]. SWL cannot be fermented with­out careful pretreatment — stripping off free sulfur dioxide and simultaneous concentration, steaming, removing inhibitors, adding nutrients, and adjusting the pH [43].

Although lignocellulose biomass is cheap and predominantly comprises waste material, the logistics, handling, storage and transportation dramatically increases its cost and therefore its use directly on site is preferred over to its processing in a central plant [8]. Further price increases occur due to the character of material — most lignocelluloses mentioned above are not fermentable by common ethanol producers and must be decomposed and hydrolysed into simple sugars before fermentation is carried out.

Figure 4 illustrates the results obtained for the analytical curve in the concentration range 0.1-0.4 |jg Cu L-1 in 0.14 mol L-1HNO3, using the optimized conditions of the TS-FF-AAS sys­tem. The transient signals were repeatable, and (for both tubes) the curve was linear in the concentration range studied. A two-fold greater sensitivity was achieved using the ceramic tube.

Figure 5 illustrates the results obtained for the analytical curves constructed using concen­trations of Cu in the range 100-400 |jg L-1, with additions of analytein 0.14 mol L-1 HNO3 to equal volumes of sample, under the optimized TS-FF-AAS system conditions. The presence of 75.8 |jg Cu L-1 in the sample was calculated from curve (a), obtained using the ceramic tube. This value was slightly above the detection limit (Table 1), although below the concen­tration of the first point of the analytical curve. In the case of the metal tube (curve (b)), a Cu concentration of 80.0 |jg L-1 was below the detection limit for this tube, but was nevertheless in agreement with the result obtained for the ceramic tube.

A

0.30-| 0.25 0.20 0.15

0.10 0.05 0.00

0 100 200 300 400 500

[Cu] / mg L1

Figure 5. Regression lines fitted to the analytical curves of Cu in 1:1 mixtures of fuel samples and standards prepared in 0.14 mol L-1 HNO3, obtained using the ceramic tube (a) and the metal tube (b) Equations of the lines: A = 1.00×10-2 + 1.32×10-4 (Cu) (ceramic tube); A = 1.16×10-2 + 1.45×10-4 (Cu) (metal tube).

The analytical parameters obtained for the determination of Cu under the optimized condi­tions of the TS-FF-AAS system are provided in Table 2. The analytical curves were linear for a concentration range of 100-400 jag Cu L-1 in 0.14 mol L-1 HNO3. The system could be con­sidered to be sensitive, with characteristic concentrations of 8 and 15 jag Cu L-1 for the ce­ramic and metal tubes, respectively, and analysis frequencies (using HNO3 medium) of 26
and 100 determinations per hour, respectively. Better analytical performance of the system was achieved using the ceramic tube, compared to the metal tube. The data showed that the TS-FF-AAS technique was more sensitive than FAAS, with nine-fold (ceramic tube) and five-fold (metal tube) increases in sensitivity, relative to FAAS with pneumatic nebulization, for which the characteristic concentration was 77 |jg L-1. The increase in power of detectio- nobtained using the ceramic tube was around twice that for the metal tube. The sensitivity for determination of copper using the ceramic tube was therefore two-fold that obtained us­ing the metal tube.

2. Conclusions

The TS-FF-AAS system can be used to determine copper at low concentrations, using either metal (Inconel) or ceramic (Al2O3) tubes as atomizers. Following optimization considering the most important experimental variables affecting atomization, these systems provided significantly improved detection limits for Cu determination, with nine-fold (ceramic tube) and five-fold (metal tube) increases in sensitivity, compared to traditional FAAS with pneu­matic nebulization. The TS-FF-AAS technique is simple, fast, effective, and inexpensive. It requires low volumes of sample (as little as 50 |jL) and reagents, and reduces waste genera­tion. The method offers a useful new alternative for the determination of copper in alcohol.

Acknowledgments

The authors thank UFOP and CNPq for financial assistance.

Author details

Fabiana Aparecida Lobo1, Fernanda Pollo2, Ana Cristina Villafranca2 and Mercedes de Moraes2

1 UFOP — Universidade Federal de Ouro Preto, Brazil

2 UNESP — Universidade Estadual Paulista, Brazil

A prerequisite for ethanol production from lignocellulose is to break recalcitrant structure of material by removal of lignin, and to expose cellulose, making it more accessible to cellulo­lytic enzymes by modifying its structure; this happens in the pretreatment process. Basical­ly, lignocellulose processing into fermentable sugars occurs in two steps: a) pretreatment yielding a liquid fraction that is mostly derived from hemicellulose and lignin and a solid fraction rich in cellulose, b) further enzymatic or chemical hydrolysis of the solid (wet) cellu­lose fraction to yield fermentable sugars.

Delignification (extraction of lignin by chemicals) is an essential prerequisite for enzymatic digestion of biomass; it disrupts the lignin polymeric structure, leading to biomass swelling and increase in its surface area and enables contact of cellulolytic enzymes with cellulose fi­bres. Although some pretreatment methods do not lead to a significant decrease in lignin content, all of them alter its chemical structure making biomass more digestible even though it may contain the same amount of lignin as non-pretreated biomass [29]. Hemicellulose is often dissolved during pretreatment because it is thermosensitive and easily acid-hydro­lysed due to its amorphous branched structure; the liquid fraction obtained after pretreat­ment thus contains mainly pentose sugars (D-xylose, D-arabinose) originating from hemicelluloses, and strains fermenting pentose sugars must be used for its processing into ethanol as discussed later. The solid wet fraction obtained after pretreatment contains pre­dominantly cellulose and needs further processing to yield fermentable sugars.

The conversion of lignocellulose into fermentable sugars is more difficult to achieve than conversion of starch; starchy material is converted from a crystalline to an amorphous struc­ture at temperatures of 60-70°C, while lignocellulose is more resistant — a temperature of 320°C and a pressure of 25 MPa is needed to achieve its amorphous structure in water [17]. Therefore complete decomposition of cellulose is rarely attainable. Although lignocellulose pretreatment is an energy-intensive process, which contributes significantly to the price of the final product (18-20% of the total cost of lignocellulosic bioethanol is attributed to pre­treatment) [8], it is a necessary expense because enzymatic hydrolysis of non-pretreated ma­terial provides less than 20% of the theoretical maximum yield of fermentable sugars for the majority of lignocellulose feedstocks [44]. The resistance of biomass to enzymatic attack is characterized by a number of physical variables such as lignin content, crystallinity index (ratio of crystalline to amorphous composition of cellulose), degree of polymerization, chain length, specific surface area, pore volume or particle size [31], which are material specific; e. g. pretreatment of woody biomass differs considerably from agriculture biomass, while paper sludge doesn’t need any processing.

Efficient pretreatment of biomass is characterized by an optimum combination of variables which leads to effective disruption of the complex lignocellulosic structure, removes most of the lignin, reduces cellulose crystallinity and increases the surface area of cellulose that is accessible to enzymatic attack. At the same time, it should minimize the loss of sugars, limit the formation of toxic compounds, enable the recovery of valuable components (e. g. lignin or furfural), use high solids loading, be effective for many lignocellulosic materials, reduce energy expenses, minimize operating costs and maximize the sugar yield in the subsequent enzymatic processing [45—47]. Pretreatment efficiency is usually assessed as: a) total amount of recoverable carbohydrates analysed as concentration of sugars released in the liquid and solid fraction after pretreatment, b) conversion of cellulose, expressed as the amount of sug­ars released by enzymatic hydrolysis of the solid phase, c) fermentability of released sugars, expressed as the amount of ethanol produced in the subsequent fermentation or d) its toxici­ty (concentration of inhibitory compounds released by sugar and lignin decomposition) ana­lysed by HPLC or measured as the ability of test strains to grow.

Although it might seem that the problem of lignocellulose pretreatment has been solved by the chemical pulping process, which has been used commercially for a long time to produce various paper products, the opposite is true; despite most lignin is removed in these proc­esses, they have been optimized to maintain the strength and integrity of cellulose fibres that are used for papermaking or as chemical feedstock and thus they are not easily hydro­lysed by enzymes. The traditional sulfite pulping process was first reported in 1857 where treatment of wood with a mixture of sulfur dioxide in hot water considerably softened the wood; in 1900 the sulfurous acid process was patented [6]. Nowadays chemical pulp pro­duction based on the sulphite method [38] use sulfurous acid and its salts (Ca2+, Mg2+, Na+ and NH4+) in combination with SO2 as a cooking liquor at temperatures of 120 — 150 °C. Sul — furous acid is an impregnation agent, improving the penetration of hydrolytic chemicals in­side the wood structure [48], and importantly, promotes sulfonation of lignin leading to formation of lignosulfonic acid and its salts, that are soluble [49, 50]. Combinations of salts and cooking conditions produce different qualities of cellulose and different compositions of the sulfite waste liquors. Possibility to optimize old sulphite pulping process to obtain high­er degree of saccharification of hard and softwoods had led to various modifications of proc­ess condition [48, 51—54]. So called SPORL technique is based on application of solution of bisulphate salts and sulfur dioxide (sulfurous acid) on biomass; sulfuric acid can also be added depending on lignin content (the higher amount of sulfuric acid is necessary for bio­mass with higher content of lignin, e. g. softwood, eucalyptus).

Many other processes have been investigated over the last decades in order to intensify lignocellulose pretreatment process by exploiting various physical, chemical and biological methods or their combination as reviewed elsewhere [29, 47] and summarized in Table 2.

volatility of ammonia, concerns low cost, ammonia

Acid treatments lead mainly to hydrolysis of hemicelluloses (pentose and hexose frac­tions) while alkaline treatments bring about lignin removal. Concentrated acids such as sulphuric or hydrochloric have been used as powerful agents to treat lignocelluloses, but due to their toxicity, corrosivity and necessity of recovery after hydrolysis, attention has shifted to milder conditions e. g. 0.5 % (v/v) sulfuric acid [57]. To improve cellulose hy­drolysis in dilute acid processes, higher temperatures are favoured [58] since at a moder­ate temperature, direct saccharification resulted in low yields. As demonstrated by Candido et al. [59] for bagasse, dilute acid hydrolysis is greatly influenced by reaction time; at 100°C in 10% v/v sulfuric acid, the loss of mass and hemicellulose content de­creased with time while soluble lignin concentration increased. Several modifications of the dilute acid hydrolysis method have been reported, e. g. acid hydrolysis with 1 % H2SO4 to remove hemicellulose and lignin followed by an alkaline step to increase the yield of cellulose. Methods based on the use of organosolv, wet oxidation, steam explo­sion or steam enriched with various impregnating agents (SO2 CO2, NH3) are also often used for lignocellulose pretreatment as summarized in Table 2. The principle of the orga — nosolv is mild hydrolysis of lignocellulose catalysed by sulfuric acid or sodium hydrox­ide in the reactor followed by extraction into ethanol at temperatures around 175 °C. Taking sugar cane bagasse as an example, the solid to liquid ratio can vary from 1 to 5 kg/l or lower, and solubilized lignin and hemicellulose appear in the liquid phase [34].

Wet oxidation is widely used in research and development technologies. Martin et al.

[60] compared wet oxidation of bagasse, which was mixed with water (ca. 6 % w/v dry bagasse) in a special autoclave under slightly alkaline conditions, with steam explosion.

In the wet oxidation procedure, slightly lower solubilisation of lignin, higher solubilisa­tion of hemicellulose and higher cellulose content in the solid phase (approx. 60 % w/w) was achieved in comparison with steam explosion (45 % w/w). The effect of steam en­richment with CO2 or SO2 proved promising results as for enzymatic hydrolysis of cellu­lose and the low content of inhibitors, especially 2-furalaldehyde and 5-hydroxymethyl-2- furalaldehyde.

In summary, biomass pretreatment is a key bottleneck in the bioprocessing of lignocellulose biomass and even though all methods have distinct advantages, as summarized in Table 2, the main problems are high energy consumption and low substrate loading, leading to low sugar recovery. However, increasing the biomass concentration leads to high solid slurries which are very viscous, with a pasta-like behaviour, creating a challenge for mixing, pump­ing and handling; this increases energy demands reflected in a higher price for the ethanol as well as concentrates toxic compounds, thus counteracting any potential benefits [61].

Although the pretreatment process disrupts the complex structure of the material and caus­es partial hydrolysis of cellulose, the content of fermentable sugars is still very low; further enzymatic degradation of the cellulose polymeric chain must be carried out to increase the concentration of glucose, which is utilized (optimally together with hemicellulose-derived monomers) in fermentation as shown in Figure 1.

Most commercial enzyme preparations (the largest producers are Genencor, Novozymes or Spezyme) are produced by cultivation of Trichoderma resei as mixtures of enzymes with en — do-1,4-fi-D-glucanase (EC 3.2.1.4, hydrolysis of (1^4) glucosidic linkages inside the chain), exo-1,4- fi-glucosidase (EC 3.2.1.74, hydrolysis of (1^4) linkage in (1^4)-p-D-glucans to re­move successive glucose units), fi-glucosidase (EC 3.2.1.21, hydrolysis of terminal non-re­ducing p-D-glucosyl residues with release of p-D-glucose) and p-1,4-glucan cellobiohydrolase (EC 3.2.1.91, hydrolysis of (1^4)-p-D-glucosidic linkages in cellulose and cellotetraose releasing cellobiose from non-reducing ends of the chains) activities working in synergy.

In recent years, the efficiency of commercial enzyme mixtures has rapidly increased and permits high conversions of cellulose to glucose; e. g. 85% and 91% yields of glucose were reported for ionic liquid pretreated poplar and switchgrass [62] and 85% and 83% yields were achieved for acid pretreated poplar and rice straws respectively [17, 63, 64]. Although the differential between the price of amylolytic and cellulolytic enzymes is currently re­duced, the major difference is in dosing; about 40 -100 times more enzyme (based on protein weight) is required to breakdown cellulose compared to starch [29]. According to economic analyses, the conversion of biomass into fermentable sugars, which includes enzyme pro­duction and enzymatic hydrolysis together with indispensable pretreatment of biomass, comprises 33 % of the total cost [8, 17] and the estimated cost of cellulases is 50 cents per gallon (3.785 l) of ethanol, which is often comparable to the purchase cost of the feedstock [65]. For this reason attention has turned to further improvement of the composition and ac­tivity of enzyme cocktails, e. g. by constructing tailor-made multienzyme systems. It was shown that addition of xylanase and pectinase to alkali-pretreated biomass can reduce the negative effect of hemicellulose and pectin, which can restrict access of cellulases to the cel­lulose surface, while p-xylosidase can decompose xylobiose and polymerized xylooligomers to avoid inhibition of cellulolytic enzymes [22, 45]. Unfortunately, improved enzyme cock­tails are not generally applicable, e. g. an enzyme complex enriched with p-mannanase and amyloglucosidase improved digestibility of dried distillers grains, but this was not required for corn stover [22]. Furthermore, the rate and efficiency of enzymatic hydrolysis can be af­fected by enzyme adsorption to non-cellulolytic substrates, e. g. lignin through phenolic groups and hydrophobic interactions, which limits the accessibility of cellulose to cellulases [45, 47]. To reduce this effect, "designer cellulosomes" have been recently constructed [45].

The cellulosome is a large complex of cellulolytic enzymes, originally produced by anaero­bic bacteria [66], and has been engineered to comprise a recombinant chimeric scaffolding protein and many bound protein hybrids that have low lignin binding affinity. A different approach is represented by the addition of non-catalytic additives, e. g. surfactants (e. g. Tween, polyethylene glycol), polymers or proteins (bovine serum albumin, gelatine), which compete with cellulolytic enzymes for adsorption sites of lignin and thus prevent non-pro­ductive enzyme binding and can also facilitate enzyme recycling. Addition of expansins (plant proteins), expansin-like proteins or swollenin (fungal protein) promotes enhanced en­zymatic hydrolysis by disrupting hydrogen bonding between cellulose and other cell-wall polysaccharides [45]. Recycling of enzymes, e. g. by ultrafiltration, re-adsorbtion onto fresh substrate, enzyme immobilization onto various materials e. g. chitosan-alginate composite, chitosan-clay composite, Eupergit C, mesoporous silicates, silicagel or kaolin are other ap­proaches to reduce pretreatment costs [45].

The activity of cellulolytic enzymes can be reduced not only by ineffective binding, but also by feedback inhibition by glucose and cellobiose released by hydrolysis of cellulose as reviewed by Andric et al. [67] and by inhibitory effects of toxic products that may be released during pretreatment (type and concentration depends on biomass and process conditions) and can affect not only the rate and yield of saccharification but also sub­strate fermentability.

Fung Min Liew, Michael Kopke and Sean Dennis Simpson

Additional information is available at the end of the chapter http://dx. doi. org/10.5772/52164

1. Introduction

With diminishing global reserves of crude oil and increasing demand, especially from developing countries, the pressure on oil supply will grow. Although the 2007-2010 fi­nancial crisis brought down the price of crude oil (per barrel) from a record peak of US $145 in July 2008, factors such as recovering global economies and political instability in the Middle East have restored the price of crude oil to the US$100 mark. At current rate of consumption, the global reserves of petroleum are predicted to be exhausted within 50 years [1, 2]. This, coupled with the deleterious environmental impacts that result from accumulating atmospheric CO2 from the burning of fossil fuels, the development of af­fordable, and environmentally sustainable fuels is urgently required. Many countries have responded to this challenge by legislating mandates and introducing policies to stimulate research and development (R&D) and commercialization of technologies that allow the production of low cost, low fossil carbon emitting fuels. For instance, the Euro­pean Union (EU) has mandated member countries to a target of deriving 10% of all transportation fuel from renewable sources by 2020 [3]. Between 2005 and 2010, renewa­ble energies such as solar, wind, and biofuels have been increasing at an average annual rate of 15-50% [4]. Renewable energy accounted for an estimated 16% of global final en­ergy consumption in 2009 [4].

Biofuels have been defined as solid (bio-char), liquid (bioethanol, biobutanol, and biodie­sel) and gaseous (biogas, biosyngas, and biohydrogen) fuels that are mainly derived from biomass [5]. Liquid biofuels provided a small but growing contribution towards worldwide fuel usage, accounting for 2.7% of global road transport fuels in 2010 [4]. The world’s largest producer of biofuels is the United States (US), followed by Brazil and the EU [4]. In 2009, US and Brazil accounted for approximately 85% of global bioethanol

production while Europe generated about 85% of the world’s biodiesel [6]. The global market for liquid biofuels (bioethanol and biodiesel) increased dramatically in recent years, reaching US$83 billion in 2011 and is projected to US$139 billion by 2021 [7].

The use and production of biofuels has a long history, starting with the inventors Niko­laus August Otto and Rudolph Diesel, who already envisioned the use of biofuels such as ethanol and natural oils when developing the first Otto cycle combustion and diesel engines [6]. While fermentative production of ethanol has been used for thousands of years, mainly for brewing beer starting in Mesopotamia 5000 B. C., fermentative produc­tion of another potential biofuel butanol, has only been discovered over the last century, but had significant impact. During the World War 1, Chaim Weizmann successfully ap­plied a process called ABE (acetone-butanol-ethanol) fermentation using Clostridium aceto — butylicum to generate industrial scale acetone (for cordites, the propellant of cartridges and shells) from starchy materials [6, 8]. His contribution was later recognised in the Balfour declaration in 1917 and he became the first President of the newly founded State of Israel [6, 8]. Intriguingly, the enormous potential of butanol produced at that time was not realized and the substance was simply stored in huge containers [6]. ABE fer­mentation became the second biggest ever biotechnological process (after the ethanol fer­mentation process) ever performed, but the low demand of acetone following the conclusion of the war led to closure of all the plants [8]. Although ABE fermentation briefly made a comeback during the Second World War, increasing substrate costs and increasing stable supply of low cost crude oil from the Middle East rendered the tech­nology economically unviable. Recently, a resurgence of the technology is underway as some old plants are reopened and new plants are being built or planned in China, the US, the United Kingdom (UK), Brazil, France and Austria [6, 8].

Traditionally sugar substrates derived from food crops such as sugar cane, corn (maize) and sugar beet have been the preferred feedstocks for the production of biofuels. How­ever, world raw sugar prices have witnessed significant volatility over the last decade or so, ranging from US$216/ton in year 2000 to a 30 year high of US$795/ton in February 2011 due to global sugar deficits and crop shortfall [9]. This has created uncertainty and raised sustainability issues about its use as a feedstock for large scale biofuel production. This review aims to shed light on the use of syngas and industrial waste gas as feed­stocks, and the emerging field of gas fermentation to generate not only biofuels, but also other high-value added products. The advantages of gas fermentation over conventional sugar-based fermentation and thermochemical conversions, and their flexibility in utiliz­ing a spectrum of feedstocks to generate syngas will be discussed. The biochemistry, ge­netic and energetic background of the microorganisms that perform this bioconversion process will be critically examined, together with recent advances in systems biology and synthetic biology that offer growing opportunities to improve biocatalysts in terms of both the potential products that can be produced and their process performance. The key processes such as gasification, bioreactor designs, media formulation, and product recovery will be analysed. Finally, the state of commercialization of gas fermentation will be highlighted and an outlook will be provided.

Toxic products can generally be divided into three main groups — aliphatic acids, furan de­rivatives and phenolic compounds [68—70] released by degradation of carbohydrates, and compounds arising from lignin. In acidic solutions, cellulose and hemicellulose are broken down into hexose and pentose sugars, which are further decomposed at high temperatures into furan derivatives represented mainly by 2-furaldehyde (furfural, FF) and 5-hydroxy- methyl-2-furaldehyde (hydroxymethylfurfural, HMF). Free aliphatic acids, represented mainly by acetic, formic or levulinic acids, are created by substituents cleaved from lignin and hemicelluloses within the pretreatment, or are produced by cells during fermentation, while phenolic derivatives (4-hydroxybenzoic acid, 3,4-dihydroxybenzoic acid or vanilin) arise mainly from lignin decomposition in alkaline solution [71]. About 40 lignocellulose degradation products have been identified in various hydrolysates [71], the type and amount depending on type of biomass and pretreatment conditions [68]; e. g. furfural, hy — droxymethylfurfural and levulinic acid occur in higher concentrations at low pH combined with high temperature and pressure [68, 71], while vanilin, vanilic, benzoic and 4-hydroxy — coumaric acids are formed under alkaline conditions at elevated temperatures and acetic acid is produced in significant concentrations independent of the process and type of bio­mass [71]. Although many studies on the effect of inhibitors on cellulolytic enzymes have been published, a general conclusion is not easy to draw because it is influenced not only by the type and origin of the enzyme preparation, but also by its dosing and the concentration of inhibitors. However, in general, compounds exhibiting higher hydrophobicity tend to be more inhibitory to cellulolytic enzymes, the greatest inhibitory effect being caused by acetic and formic acids [72—74], while the activity of enzymes is not practically influenced by levu — linic acid [73]. On the other hand, the presence of inhibitory compounds also affects ethanol productivity in the subsequent fermentation by influencing metabolic functions of ethanol producing strains. Inhibitory effects are described by type and concentration of toxic com­pounds (their effect is intensified when present in combination) and the strain used for etha­nol production, but generally, fermentation is mainly influenced by the presence of furan derivatives together with phenolic compounds and weak acids (at low pH). As reviewed elsewhere [70, 75], low molecular weight compounds are able to penetrate the cell, while in­hibitors with high molecular weights affect expression and activity of sugar and ion trans­porters. Growth and rate of ethanol production by Saccharomyces cerevisiae, the main ethanol producing strain, is significantly inhibited by furfural, while ethanol yield is almost not in­fluenced [75] due to its ability to detoxify the broth by reduction of furfural to furfuryl alco­hol, which is less toxic.

Surprisingly, in butanol production process, C. beijerinckii BA101, C. acetobutylicum P260, C. acetobutylicum ATCC 824, Clostridium saccharobutylicum 262 and Clostridium butylicum 592 were not sensitive towards sugar degradation products like furfural or hydroxymethylfur — fural (up to concentrations of 2-3 g/l) but its growth and solvent production were inhibited by p-coumaric and ferulic acids present at a concentration of 0.3 g/l [76—78]. Solvent produc­tivity and final solvent concentration in C. beijerinckii P260 were stimulated by addition of furfural or hydroxy methylfurfural (or both compounds) to the fermentation medium, at concentrations of up to 1 g/l [79]. C. acetobutylicum ATCC 824 metabolized furfural and hy­droxymethyl furfural into furfuryl alcohol and 2,5-bis-hydroxymethylfuran, respectively and these compounds positively influenced solvent production up to a concentration of 2 g/l. It was hypothesised that this biotransformation step, independent of initial furfural and HMF concentrations, might increase solventogenesis via an increased rate of regeneration of NAD+ [80]. Another possible inhibitor of phenolic origin, syringaldehyde, caused inhibition of solvent production by C. beijerinckii NCIMB 8052 over the whole range tested (0.2-1 g/l). This inhibition was probably caused by decreased expression and activity of coenzyme A transferase, which participated in utilization of butyric and acetic acids, because these acids accumulated in the medium [81].

The inhibitory effects of toxic compounds released by sugars and lignin degradation can be reduced in several ways, e. g. optimization of pretreatment conditions to minimize the formation of inhibitors, use of specific detoxification methods, e. g. precipitation by calci­um hydroxide (overliming) alone or in combination with sulphite addition, adsorption on charcoal, evaporation of the volatile fraction, extraction with ethyl acetate or diethyl ether, ion extraction, treatment with peroxidase (E. C. 1.11.7) and laccase (EC 1.10.3.2), or use of microbial strains with increased resistance to inhibitors (achieved by adaptation or prepared by genetic modification) [75, 82, 83]. Lignin degradation products, p-coumaric, ferulic and vanillic acids, together with vanillin, were effectively removed from a model solution of phenolic compounds by treatment with 0.01|oM peroxidase (E. C. 1.11.7), re­sulting in improved growth and butanol production by C. beijerinckii NCIMB 8052 [84]. Sulphuric acid-hydrolysed corn fiber was treated with XAD-4 resin, resulting in an im­provement of butanol yield achieved with C. beijerinckii BA101 [85]. Another popular ap­proach for detoxification of acid hydrolysates for butanol production is "overliming" i. e. addition of Ca(OH)2 in excess to hydrolysate [78, 85]. Although this detoxification meth­od has been known for a long time, its mode of action, especially in the case of butanol production, is not completely clear. Addition of Ca(OH)2 to an acid hydrolysate decreases furfural and HMF concentrations [86, 87] but does not affect acid concentrations; thus it is only possible to assume a beneficial neutralization effect. Furthermore it may be useful to treat hydrolysates with activated carbon [88].