**Liquids**

**Chapter 1**

**Biofuels and Co-Products Out of Hemicelluloses**

Second generation biofuels are based on the utilisation of non-edible feedstock for the production either of ethanol to be inserted in the gasoline pool or of biodiesel to be insert‐ ed in the diesel pool. Ethanol is usually produced out of fermentation of C6 sugars (al‐ though other approaches does exist, see [1]) and the latter came, in first generation ethanol, from starch. In second-generation ethanol, the source of carbohydrate considered is usually cellulose, which, in turns, is obtained from lignocellulosic biomass. Recent work by Lavoie*et al*. [2] have depicted an overview of many types of lignocellulosic biomass and in most cases, cellulose, although a major component, is not the only one and is ac‐ companied by lignin, hemicelluloses, extractives and, in case of agricultural biomass, pro‐ teins. High grade biomass (as wood chips, sugar cane or even corn) are usually very expensive (more than 100 USD/tonne) because, in most part, of the important demand re‐ lated to those feedstock in industries and this is why cellulosic ethanol is more than often related to residual biomass. The latter includes but is not limited to residual forest and ag‐ ricultural biomass as well as energy crops. In all cases, although the feedstock is rather in‐ expensive (60-80 USD/tonne), it is composed of many different tissues (leaves, bark, wood, stems, etc.) making its transformation rather complex [3]. Industrialisation of second-gen‐ eration biofuel requires specific pre-treatment that should be as versatile as efficient in or‐ der to cope with the economy of scale that has to be implemented in order to make such

The whole economics of cellulosic ethanol relies first on ethanol, which has a commodity beneficiates from a quasi-infinite market as long as prices are competitive. Assuming aver‐ age cellulose content of 45-55 % (wt) in the lignocellulosic biomass, the ethanol potential of lignocellulosic biomass would range between 313-390 L per tonne of biomass converted.

> © 2013 Fuente-Hernández et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

© 2013 Fuente-Hernández et al.; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

distribution, and reproduction in any medium, provided the original work is properly cited.

Ariadna Fuente-Hernández, Pierre-Olivier Corcos,

Romain Beauchet and Jean-Michel Lavoie

http://dx.doi.org/10.5772/52645

**1. Introduction**

conversion economical.

Additional information is available at the end of the chapter

### **Biofuels and Co-Products Out of Hemicelluloses**

Ariadna Fuente-Hernández, Pierre-Olivier Corcos, Romain Beauchet and Jean-Michel Lavoie

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/52645

#### **1. Introduction**

Second generation biofuels are based on the utilisation of non-edible feedstock for the production either of ethanol to be inserted in the gasoline pool or of biodiesel to be insert‐ ed in the diesel pool. Ethanol is usually produced out of fermentation of C6 sugars (al‐ though other approaches does exist, see [1]) and the latter came, in first generation ethanol, from starch. In second-generation ethanol, the source of carbohydrate considered is usually cellulose, which, in turns, is obtained from lignocellulosic biomass. Recent work by Lavoie*et al*. [2] have depicted an overview of many types of lignocellulosic biomass and in most cases, cellulose, although a major component, is not the only one and is ac‐ companied by lignin, hemicelluloses, extractives and, in case of agricultural biomass, pro‐ teins. High grade biomass (as wood chips, sugar cane or even corn) are usually very expensive (more than 100 USD/tonne) because, in most part, of the important demand re‐ lated to those feedstock in industries and this is why cellulosic ethanol is more than often related to residual biomass. The latter includes but is not limited to residual forest and ag‐ ricultural biomass as well as energy crops. In all cases, although the feedstock is rather in‐ expensive (60-80 USD/tonne), it is composed of many different tissues (leaves, bark, wood, stems, etc.) making its transformation rather complex [3]. Industrialisation of second-gen‐ eration biofuel requires specific pre-treatment that should be as versatile as efficient in or‐ der to cope with the economy of scale that has to be implemented in order to make such conversion economical.

The whole economics of cellulosic ethanol relies first on ethanol, which has a commodity beneficiates from a quasi-infinite market as long as prices are competitive. Assuming aver‐ age cellulose content of 45-55 % (wt) in the lignocellulosic biomass, the ethanol potential of lignocellulosic biomass would range between 313-390 L per tonne of biomass converted.

© 2013 Fuente-Hernández et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 Fuente-Hernández et al.; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

With an actual market price of 0.48 USD per liter the value of this ethanol would range be‐ tween 150-187 USD per tonne of biomass processed. Since the latter is more expensive to process (first isolation of cellulose then hydrolysis of cellulose) and considering the fact that the feedstock is itself expensive, there is a necessity to get an added value out of the remain‐ ing 55-45 % (wt) content. This residual carbon source is composed mostly of hemicelluloses and of lignin. The latter is a very energetic aromatic-based macromolecule, that has a high calorific value explaining why many processes converting such biomass (as some pulp and paper processes) relies on the combustion of lignin to provide part of the energy for the in‐ dustry. It could also serve as a feedstock for the production of added-value compounds and although the subject is very pertinent to the field, it is out of the scope of this review, which focuses mostly on C5 sugars derived from hemicelluloses.

technique allows simultaneous removal both for lignin and hemicelluloses. However, in‐ stead of using only an aqueous mixture of ions, the process relies on the utilisation of a com‐ bination of ions (usually alkaline) in a 50/50 mixture of aqueous organic solvent. In most cases, the solvent is methanol for obvious economic reasons although other solvents as buta‐ nol and certain organic acids have also been investigated to the same purposes. Recent work by Wang*et al*. [10]have shown that in an organosolv process using different solvent as well as different catalyst with poplar, sodium hydroxide was shown to be the best catalyst for hemicellulose removal from the pulp. Recent work by Brosse *et al*. [11] also showed that for *Miscanthus Gigantheus*, an ethanol organosolv process combined with an acid catalyst (sul‐ phuric) lead to removal of most of the hemicelluloses and lignin from the original biomass.

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5

Finally, another approach that could lead to isolation of hemicellulose from a lignocellulosic matrix is steam processes. This technique relies on impregnation of the feedstock with water (either catalyzed or not) then treatment under pressure at temperature ranging from 180-230

C for a certain period of time after which pressure is relieved suddenly thus creating an "explosion" of the feedstock. Such process could lead, depending on the operating condi‐ tion, to the isolation of either hemicellulose or lignin in two steps or in a single step. Our team has demonstrated the feasibility of both processes for different substrates [12-14].

Independently of the substrate or the technique used for the isolation of the hemicelluloses, conversion of lignocellulosic biomass, either for the production of paper or for the produc‐ tion of biofuels requires a complete utilization of the carbon compound found in biomass. Once the hemicelluloses are isolated from the original feedstock, they can undergo different types of transformation leading to different added value compounds that could lead to in‐

Hemicelluloses account for 15-35 % of lignocellulosic biomass dry weight [2] and they are usually composed of different carbohydrates as well as small organic acids as acetic and for‐ mic acid. Glucose and xylose are often the most abundant sugars in hemicelluloses hydroly‐ sis although mannose, arabinose and galactose might also be present in lower concentrations. The carbohydrate compositions of some lignocellullosic biomass are shown in Table 1. Whilst the C6 sugars could easily be fermented to ethanol following detoxifica‐ tion of the mixture, C5 sugars remains hard to convert to ethanol, mostly because classical yeasts don't metabolise them and the genetically modified organism that ferment C5 sugars are usually slower than classical organisms used in the production of etanol from C6 sugars. Nevertheless, even if ethanol production may remain a challenge, other alternatives could be considered, both on the chemical and on the microbiological point of view, to allow con‐

Carbohydrates tend to react in acidic, basic, oxidative or reductive mediums and therefore, numerous do arise for the conversion of C5 sugars. Although many options are available, this review will focus solely on 4 different pathways: acid, base, oxidative, and reductive. Each of these pathways could be inserted in an integrated biorefinery process where each of

the fractions could be isolated and upgraded to high value compounds (see Figure 1).

crease the margin of profit for the industries in the field.

version of C5 sugar into added value products.

°

Conversion of the carbohydrates is of course an important part of the process although; iso‐ lation of hemicellulose for the lignocellulosic matrix is also crucial for such an approach and in consequence should also be briefly assessed. For years now, the pulp and paper industry have worked with lignocellulosic substrates and they have over the year developed many techniques allowing isolation of hemicelluloses. Chemical processes as soda pulping and kraft pulping allows isolation of both lignin and hemicellulose whilst protecting the cellulo‐ sic fibres in order to produce the largest amount of pulp possible per ton of biomass. Never‐ theless, in both chemical processes previously mentioned, the hemicellulose are rather difficult to reach since they are mixed with a variety of organic and inorganic compounds including lignin as well as the chemicals that were used for the pulping process. During the last decades, the pulp and paper industry have started to look toward other processes that could allow a preliminary removal of hemicelluloses in order to avoid a complicated and ex‐ pensive isolation after a chemical pulping process.

Amongst the techniques used for prehydrolysis, treatments with hot water catalyzed or not have been investigated in details in literature. As an example, Schild*et al*. [4] performed a preliminary extraction with water (via auto-hydrolysis) or with alkaline water prior to soda pulping in order to recuperate the hemicellulose prior to pulping. Similar testing was also performed on northern spruce with pressurised hot water in the presence of sodium bicar‐ bonate [5]. Hot water extractions were also performed at temperature around 170 ° C at dif‐ ferent pH (the latter were adjusted with a phthalate buffer) and these experiments showed that control of pH was crucial in order to extract more of the hemicelluloses (up to 8 % wt on original biomass) [6]. Hot water extractions at similar temperature range have also been per‐ formed on maple [7] as well as on sugarcane bagasse [8]. Overall the hot water pretreatment may be a very promising approach for isolation of hemicelluloses although reported rates did not go far over 10 % because of the necessity to preserve the cellulosic fibres in order to avoid losses for papermaking. Acid catalyst has also been used as pretreatment to remove hemicellulose prior to pulping as reported by Liu*et al*. [9]. Utilisation of sulphuric acid, al‐ though very efficient to remove hemicellulose may also have an impact on cellulose thus re‐ ducing the pulp production rates.

Another process that could lead to isolation of hemicellulose is the organosolv process, which is to a certain extent comparable to classical chemical pulping in that sense that the technique allows simultaneous removal both for lignin and hemicelluloses. However, in‐ stead of using only an aqueous mixture of ions, the process relies on the utilisation of a com‐ bination of ions (usually alkaline) in a 50/50 mixture of aqueous organic solvent. In most cases, the solvent is methanol for obvious economic reasons although other solvents as buta‐ nol and certain organic acids have also been investigated to the same purposes. Recent work by Wang*et al*. [10]have shown that in an organosolv process using different solvent as well as different catalyst with poplar, sodium hydroxide was shown to be the best catalyst for hemicellulose removal from the pulp. Recent work by Brosse *et al*. [11] also showed that for *Miscanthus Gigantheus*, an ethanol organosolv process combined with an acid catalyst (sul‐ phuric) lead to removal of most of the hemicelluloses and lignin from the original biomass.

Finally, another approach that could lead to isolation of hemicellulose from a lignocellulosic matrix is steam processes. This technique relies on impregnation of the feedstock with water (either catalyzed or not) then treatment under pressure at temperature ranging from 180-230 ° C for a certain period of time after which pressure is relieved suddenly thus creating an "explosion" of the feedstock. Such process could lead, depending on the operating condi‐ tion, to the isolation of either hemicellulose or lignin in two steps or in a single step. Our team has demonstrated the feasibility of both processes for different substrates [12-14].

Independently of the substrate or the technique used for the isolation of the hemicelluloses, conversion of lignocellulosic biomass, either for the production of paper or for the produc‐ tion of biofuels requires a complete utilization of the carbon compound found in biomass. Once the hemicelluloses are isolated from the original feedstock, they can undergo different types of transformation leading to different added value compounds that could lead to in‐ crease the margin of profit for the industries in the field.

Hemicelluloses account for 15-35 % of lignocellulosic biomass dry weight [2] and they are usually composed of different carbohydrates as well as small organic acids as acetic and for‐ mic acid. Glucose and xylose are often the most abundant sugars in hemicelluloses hydroly‐ sis although mannose, arabinose and galactose might also be present in lower concentrations. The carbohydrate compositions of some lignocellullosic biomass are shown in Table 1. Whilst the C6 sugars could easily be fermented to ethanol following detoxifica‐ tion of the mixture, C5 sugars remains hard to convert to ethanol, mostly because classical yeasts don't metabolise them and the genetically modified organism that ferment C5 sugars are usually slower than classical organisms used in the production of etanol from C6 sugars. Nevertheless, even if ethanol production may remain a challenge, other alternatives could be considered, both on the chemical and on the microbiological point of view, to allow con‐ version of C5 sugar into added value products.

Carbohydrates tend to react in acidic, basic, oxidative or reductive mediums and therefore, numerous do arise for the conversion of C5 sugars. Although many options are available, this review will focus solely on 4 different pathways: acid, base, oxidative, and reductive. Each of these pathways could be inserted in an integrated biorefinery process where each of the fractions could be isolated and upgraded to high value compounds (see Figure 1).


proaches depending on the reactant as reported by Marcotullio *et al*. [20] using halogen ions and proceeding only via the aliphatic form or as reported by Nimlos *et al*. [21] either via an aliphatic or a cyclic pathway (D-xylopyranose). Many different types of acid catalyst, either Brønsted or Lewis have been tested for the production of furfural. Although most of the acids reported in literature have been efficient so far for the production of the targeted mole‐ cule, one of the major side-reaction of furfural is polymerisation which influences the con‐ version rates and the selectivity of most of the processes reported in literature. An example of the abundance of research on this specific conversion is shown in Table 2 for different de‐

> **Catalyst Conversion Reference** H-Mordenite 98% [22] Sulphonic acid/Silica surface 99% [23] 1-methylimidazole 91% [24] KI, KCl (dilute acid) 88% [20] NaCl, H2SO4 83% [25] 1-alkyl-3-methylimidazolium 84% [26]

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7

NaCl, HCl 78% [27]

Amberlyst 70 75% [29] Zeolite H-Beta 74% [30] MCM-22, ITQ-2 70% [31] FeCl3 71% [32] Nafion 60% [33] Keggin type acids 62% [34] Vanadyl pyrophosphate 53% [35]

C under proportional

Aluminium chloride Hexahydrate 76% [28]

**Table 2.** Molar conversion to furfural in relationship with the catalyst used for the dehydration of xylose to furfural

pressure allowing the mixture to remain liquid. Many researches also use a co-solvent, often toluene in order to isolate furfural from the aqueous mixture. The reason why toluene is so popular to this purpose is mostly related to the fact that toluene has affinity for fufural thus

Heterogeneous catalyst has been proven to be very efficient for the process [22,23] although polymerisation tend to reduce the surface activity thus leading to a short-term deactivation of the catalyst. On the other hand, homogeneous catalyst was also shown to be efficient but at this point the whole technique relies on how the organic solvent is dispersed in the aque‐ ous mixture. Reducing the size of the organic solvent particles in water (or vice-versa) to the maximum should allow the best transfer between the aqueous phase to the organic phase,

For these reactions, the temperature is generally between 140-240 °

hydration reactions under acid catalyst..

under acid catalyst.

inhibiting its polymerization.

**Table 1.** Carbohydrate composition of some lignocellulosic biomass.

**Figure 1.** Potential utilization of hemicelluloses in an optimized conversion process for residual lignocellulosic biomass where C6 sugars are converted to ethanol, lignin and extractives to other added value products.

In this review, emphasis will be made on the recent work made for each of these conversion pathways both on the chemical and on the biochemical pathways. The review will focus on these 4 approaches also for their generally simple nature that would make them adaptable to an industrial context. These results will be compared to classical fermentation processes to produce ethanol with different types of organisms that can metabolise C5 sugars.

#### **2. Conversion of xylose under an acid catalyst**

#### **2.1. The chemical pathway**

Either in cyclic or aliphatic form, xylose then tends to dehydrate thus leading to the produc‐ tion of furfural whilst losing three molecules of water. Although this approach could explain the formation of furfural, it is not the sole options and many detailed reports have shown, by correlating the intermediaries with the actual structure, could be formed by many ap‐ proaches depending on the reactant as reported by Marcotullio *et al*. [20] using halogen ions and proceeding only via the aliphatic form or as reported by Nimlos *et al*. [21] either via an aliphatic or a cyclic pathway (D-xylopyranose). Many different types of acid catalyst, either Brønsted or Lewis have been tested for the production of furfural. Although most of the acids reported in literature have been efficient so far for the production of the targeted mole‐ cule, one of the major side-reaction of furfural is polymerisation which influences the con‐ version rates and the selectivity of most of the processes reported in literature. An example of the abundance of research on this specific conversion is shown in Table 2 for different de‐ hydration reactions under acid catalyst..


**Table 2.** Molar conversion to furfural in relationship with the catalyst used for the dehydration of xylose to furfural under acid catalyst.

For these reactions, the temperature is generally between 140-240 ° C under proportional pressure allowing the mixture to remain liquid. Many researches also use a co-solvent, often toluene in order to isolate furfural from the aqueous mixture. The reason why toluene is so popular to this purpose is mostly related to the fact that toluene has affinity for fufural thus inhibiting its polymerization.

Heterogeneous catalyst has been proven to be very efficient for the process [22,23] although polymerisation tend to reduce the surface activity thus leading to a short-term deactivation of the catalyst. On the other hand, homogeneous catalyst was also shown to be efficient but at this point the whole technique relies on how the organic solvent is dispersed in the aque‐ ous mixture. Reducing the size of the organic solvent particles in water (or vice-versa) to the maximum should allow the best transfer between the aqueous phase to the organic phase, assuming of course that furfural has suitable affinity for the solvent and that the partition coefficient favours the solvent.

Lange [45] patented a process using palladium and titanium oxide whilst Zheng et al. [46] worked with a copper alloy. Value for Me-THF could be estimated from the price of THF which is around 3000 USD/tonne [47] and the gap between the value of furfural and Me-THF could justify the process although hydrogen value can be estimated to be around 4.5

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9

Another potentially interesting approach for a transformation of furfural would be decar‐

Many researches have focused on decarboxylation including work by Zhang *et al*. [48] who mentioned decarboxylation with potassium-doped palladium, and Stevens *et al*. [49] who re‐

Results reported in literature show that xylose, under an acid catalyst, tend invariably to de‐ hydrate to furfural thus limiting the possibilities for side-products in such specific condi‐ tions. The acids could be Brønsted or Lewis type, all lead to the production of furfural

Although furfural is a very common route for the conversion of xylose under an acid cata‐ lyst, furfural itself is rarely related to microorganisms in that sense that it is often considered as an inhibitor instead of a metabolite. Nevertheless, to the best of our knowledge, no report

The interaction between xylose and bases, either Brønsted or Lewis, is rather less reported in the literature when compared to the acid conversion of xylose to furfural indicated in the previous section. Many very different reactions have been reported as in the case of Popoff and Theander [50] that have quantified the cyclic compounds produced after a base-cata‐

peculiars in comparison to other work made on the subject (see Figure 5) since most of the reported compounds are aromatics. The presence of aromatics may be a result that the reac‐

C for 4 hours. The produced compounds are rather

C.

USD/Kg (estimated with the actual price of natural assuming reforming of the latter).

boxylation to furan. The general process is depicted in Figure 4 below.

ported conversion with copper chromite in supercritical CO2.

furthermore when temperature are raised above 150 °

mentioned a biological conversion of xylose to furfural.

**3. Conversion of xylose under a base catalyst**

**Figure 4.** Decarboxylation of furfural to furan.

**2.2. The biological pathway**

**3.1. The chemical pathway**

lyzed reaction of pure D-xylose at 96 °

Production of furfural itself is of course of significant interest because, amongst many fac‐ tors, this chemical is commonly used in the industry as a solvent (mostly in oil chemistry). The average world production for furfural is 250 000 t/y and the actual market price evolves around 1000 USD/t [36] with recent market value reported to be closer to 1600 USD/tonne [37]. Furfural can also be a gateway to other products that could be used either as biofuels or as biomolecules. Example of such would be furfuryl alcohol via partial reduction of furfural (see Figure 2 below).

**Figure 2.** Reduction of furfural to furfuryl alcohol.

Furfuryl alcohol is also of interest since it is used as resins, adhesives and wetting agent, it has been mentioned that most of the 250 Kt/y of the furfural production is oriented toward production of furfuryl alcohol. The market value of this compound has been reported to be around 1800-2000 USD/tonne [38] and many reports in open literature mentions high selec‐ tivity for the conversion of furfural with iridium and ruthenium catalyst [39], rhodium [40], iron [41] and with zirconium oxide [42].

Another possible target for the transformation of furfural is for the production of 2-meth‐ yltetrahydrofuran (Me-THF) (see Figure 3). The latter is actually accredited as an additive for fuel and therefore, the possible market is virtually very important. It is also used in the petroleum industry to replace tetrahydrofuran (THF) that usually comes from non-re‐ newables.

**Figure 3.** Reduction of furfural to 2-methyltetrahydrofuran.

Reduction of furfural to Me-THF seems to represent an important challenge since there is fewer reports mentioned in literature on the subject, as compared, as an example, to the re‐ duction of furfural to furfuryl alcohol. Wabnitz *et al*. [43, 44] patented a one and two step process allowing conversion of furfural to Me-THF under a palladium-based catalyst and a mixture of palladium and copper oxide and chromium oxide as for the two step process. Lange [45] patented a process using palladium and titanium oxide whilst Zheng et al. [46] worked with a copper alloy. Value for Me-THF could be estimated from the price of THF which is around 3000 USD/tonne [47] and the gap between the value of furfural and Me-THF could justify the process although hydrogen value can be estimated to be around 4.5 USD/Kg (estimated with the actual price of natural assuming reforming of the latter).

Another potentially interesting approach for a transformation of furfural would be decar‐ boxylation to furan. The general process is depicted in Figure 4 below.

**Figure 4.** Decarboxylation of furfural to furan.

Many researches have focused on decarboxylation including work by Zhang *et al*. [48] who mentioned decarboxylation with potassium-doped palladium, and Stevens *et al*. [49] who re‐ ported conversion with copper chromite in supercritical CO2.

Results reported in literature show that xylose, under an acid catalyst, tend invariably to de‐ hydrate to furfural thus limiting the possibilities for side-products in such specific condi‐ tions. The acids could be Brønsted or Lewis type, all lead to the production of furfural furthermore when temperature are raised above 150 ° C.

#### **2.2. The biological pathway**

Although furfural is a very common route for the conversion of xylose under an acid cata‐ lyst, furfural itself is rarely related to microorganisms in that sense that it is often considered as an inhibitor instead of a metabolite. Nevertheless, to the best of our knowledge, no report mentioned a biological conversion of xylose to furfural.

#### **3. Conversion of xylose under a base catalyst**

#### **3.1. The chemical pathway**

The interaction between xylose and bases, either Brønsted or Lewis, is rather less reported in the literature when compared to the acid conversion of xylose to furfural indicated in the previous section. Many very different reactions have been reported as in the case of Popoff and Theander [50] that have quantified the cyclic compounds produced after a base-cata‐ lyzed reaction of pure D-xylose at 96 ° C for 4 hours. The produced compounds are rather peculiars in comparison to other work made on the subject (see Figure 5) since most of the reported compounds are aromatics. The presence of aromatics may be a result that the reac‐ tion time was long and the isomerisation that was required in order to induce such reaction was efficient. Johansson and Samuelson [51] tested the effect of alkali treatments (NaOH) on birch xylan and contrarily to the previous research; they found that the treatment led to the production of a variety of organic acids. Testing on untreated xylene showed that most of the organic acids were already obtained from xylans and the most distinctive impact was observed after a 2 day test at 40 ° C where the concentrations of L-galactonic and altronic acids increased significantly which could be related to a less severe treatment of xylans that also include C6 sugars.

**Figure 6.** Major epimerisation products from 1-4 week reaction of D-xylose in a pH 11.5 KOH solution at room tem‐

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11

Xylose, as the other carbohydrates, is converted to smaller organic acids when reacted with a strong alkali medium. As an example, Jackson *et al*. [53] have demonstrated that the con‐ version of xylose to lactic acid could reach 64 % (molar) accompanied by glyceric acid. Al‐ though they did not used xylose but rather ribose and arabinose, they were able to reach conversions between 35-43 % into lactic acid using potassium hydroxide as catalyst under microwave irradiation [54]. Rahubadda *et al*. [55] have provided a mechanism for the con‐ version of xylose to lactic acid under a base catalyst. The simplified pathway is depicted in

**Figure 7.** Conversion of D-xylose to lactic acid via the methylglyoxal pathway.

perature.

Figure 7 below.

**Figure 5.** Cyclic and aromatics obtained from the based-catalysed treatment of D-xylose under a sodium hydroxide catalyst where (1) 2-hydroxy-3-methylcyclopent-2-enone; (2) 2-hydroxy-3,4-dimethylcyclopent-2-enone; (3) pyrocate‐ chol; (4) 3-methylbenzene-1,2-diol; (5) 4-methylbenzene-1,2-diol; (6) 3,4-dimethylbenzene-1,2-diol; (7) 2-methylben‐ zene-1,4-diol; (8) 1-(2,5-dihydroxyphenyl)ethanone; (9) 1-(3,5-dihydroxyphenyl)ethanone; (10) 1-(3,4 dihydroxyphenyl)ethanone; (11) 3,4-dihydroxybenzaldehyde; (12) 1-(2,3,4-trihydroxy -5-methylphenyl)ethanone; (13) 1-(2,3-dihydroxy-6-methylphenyl)ethanone.

El Khadem *et al*. [52] studied the effect of xylose conversion in an alkali medium at low tem‐ peratures (room) and for long periods (1-4 weeks) and one of the interesting features of his work was that the process did lead to the epimerization of sugars, but furthermore, it leads to the production of C6 sugars most probably from a reverse aldol reaction. Among the sug‐ ars that were formed during the reaction, conversion of xylose was shown to be more effi‐ cient to lyxose (18 %) and arabinose (15 %) with a decrease observed for most of the compounds between 1 and 4 weeks (see Figure 6). A vast majority (more than 50 %) of xy‐ lose remains on its original form and the reaction leads to the production of 1 % glucose and 2.5 % of sorbose, both are C6 sugars.

**Figure 6.** Major epimerisation products from 1-4 week reaction of D-xylose in a pH 11.5 KOH solution at room tem‐ perature.

Xylose, as the other carbohydrates, is converted to smaller organic acids when reacted with a strong alkali medium. As an example, Jackson *et al*. [53] have demonstrated that the con‐ version of xylose to lactic acid could reach 64 % (molar) accompanied by glyceric acid. Al‐ though they did not used xylose but rather ribose and arabinose, they were able to reach conversions between 35-43 % into lactic acid using potassium hydroxide as catalyst under microwave irradiation [54]. Rahubadda *et al*. [55] have provided a mechanism for the con‐ version of xylose to lactic acid under a base catalyst. The simplified pathway is depicted in Figure 7 below.

**Figure 7.** Conversion of D-xylose to lactic acid via the methylglyoxal pathway.

They mentioned in this report that methylglyoxal is most probably derived from glyceralde‐ hyde as depicted in Figure 8 below. The possible reaction leading to methylglyoxal may in‐ volve an E2 reaction on C2 leading to removal of the hydroxyl group on C3 then a keto-enol rearrangement to methylglyoxal.

*amylophilus*, *L. bulgaricus* and *L. leichmanii*. Mutant *Aspergillus niger* has also been reported to be effective at an industrial scale [64]. LAB have the particularity to possess an homo‐ fermentative metabolism producing only lactic acid as extracellular waste product, instead of the heterofermentative pathway yielding by-products such as aldehydes, organic acids and ketones. The catabolic pathway yielding lactic acid is essentially the same across all organisms; the pyruvate intermediate is converted to lactic acid by a lactate dehydrogen‐ ase (LDH). Thus for hexose sugars, the theoretical yield is 2 moles of lactate per mole of sugar (or 1g sugar for 1g lactate). This enzymatic catalysis has the advantage over its chemical counterpart to be stereospecific: both L-lactate-dehydrogenase (L-LDH) and Dlactate-dehydrogenase (D-LDH) exist, generating either L-lactate or D-lactate respectively [65]. Both are NAD-dependant (nicotinamide adenine dinucleotide) and may be found alone or together in wild lactate-producing microbial strains. Since optical purity of lac‐ tate is a major requirement for the lactate industry, research focuses on stereospecificity as

Biofuels and Co-Products Out of Hemicelluloses

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13

An efficient lactate producer has to display specific attributes, mainly the adaptability to low-cost substrates, high selectivity of desired enantiomer (L, D or both), high optimal tem‐ perature for decreased contamination risks, low pH tolerance and high performances (yield and productivity). LAB display appreciable performances, but lack a low pH tolerance, which implies uses of a pH control apparatus during the fermentation process. LAB optimal pH is near neutral, but the pKa of lactic acid being 3.8, an alkali agent, usually Ca(OH)2, must be used thus generating calcium lactate. After typical batch fermentation, the medium is acidified with H2SO4 therefore regenerating and purifying the lactic acid [64]. Another drawback of LAB is their requirement for a complex growth medium, since they are auxo‐ troph for certain amino acids and vitamins [71]. In order to overcome this problem, many fungi were also investigated for lactate production. Strains of *Rhizopus*, *Mucor* and *Monilla sp.* have shown potential whilst other fungi even displayed amylolytic activity, which could

Most researches still focuses on hexose conversion, and research group have optimized strains and process strategies in order to obtain high lactate titers, yields and productivities. Ding and Tan [75] developed a glucose fed-batch strategy using *L. casei* and generating up to 210 g/L of lactic acid with a 97 % yield. Chang *et al.* [76] proposed a continuous high cell density reactor strategy yielding a titer of 212.9 g/L and productivity of 10.6 g/L/h with *Lb. rhamnosus*. Dumbrepatil *et al.* [77] created a *Lb. delbrueckii* mutant by ultraviolet (UV) muta‐ genesis producing 166 g/L with productivity of 4.15 g/L/h in batch fermentation. Genetically engineered non-LAB biocatalysts yet have to match the performances of highly efficient wild LAB. In fact, *C. glutamicum*, *S. cerevisiae* and *E. coli* recombinant have been developed,

The search for lignocellulose-to-lactate biocatalysts have led to the discovery of many strains of pentose-utilizing LAB. *Lb. pentosus ATCC8041* [78, 79], *Lb. bifermentans DSM20003* [80], *Lb. brevis* [81], *Lb. Plantarum* [82], *Leuconostoc lactis* [83, 84], and *E. mundtii QU 25* [85, 86]. Lactic acid produced from xylose per say has been investigated by few [84,85, 87, 88], but with mitigated results, mainly due to the fact that the pentose-utilizing

much as yields and productivity [61,66-70].

lead to a direct starch-to-lactate conversion [72-74].

but with limited success [61].

**Figure 8.** Conversion of glyceraldehyde to methylglyoxal.

Onda *et al*. [56] achieved a conversion rate of more than 20 % when using xylose as a feed‐ stock with a carbon-supported platinum catalyst in alkali solution. In a recent report by Ma *et al*. [57], it was shown that using model compounds, different carbohydrates tend to con‐ vert into lactic acid at different levels. Fructose was shown to be more effectively converted to lactic acid than glucose and finally than xylose. The work also showed a correlation be‐ tween the amount of catalyst (varying from 1-3 % wt.) of NaOH, KOH and Ca(OH)2 respec‐ tively. Part of the work by Aspinall *et al*. [58] was aimed at the non-oxidative treatment of xylans from different substrates using sodium hydroxide as solvent. The reaction was per‐ formed at room temperature for 25 days and amongst the products that emerged from this reaction, a majority was acidic and lactic acid as well as formic acid were the two major products. Other work by Yang *et al*. [59] showed that higher temperature treatments of xy‐ lose (200 ° C) in a Ca(OH)2 solution produced about 57 % (mol.) of lactic acid with 2,4-dihy‐ droxybutanoic acid in second with 10 % (mol.). The same conversion patterns were observed by Raharja *et al*. [60] with production rates for lactic acid above 50 %.

#### **3.2. The biological pathway**

Amongst the different options for the conversion of xylose reported in the previous chap‐ ter, production of lactic acid via the microbial route is a vastly studied field [61-63] since currently, all of the production of lactic acid at an industrial scale in the world is biologi‐ cally based. Traditionally, the concept evolves around fermenting carbohydrate-based syr‐ up by homolactic organisms, mostly lactic acid bacteria (LAB). The most common carbohydrate-based substrates used to this purpose may be molasses, corn syrup, whey, sugarcane or even beet bagasse. Highly efficient LAB includes *Lactobacillus delbrueckii*, *L.* *amylophilus*, *L. bulgaricus* and *L. leichmanii*. Mutant *Aspergillus niger* has also been reported to be effective at an industrial scale [64]. LAB have the particularity to possess an homo‐ fermentative metabolism producing only lactic acid as extracellular waste product, instead of the heterofermentative pathway yielding by-products such as aldehydes, organic acids and ketones. The catabolic pathway yielding lactic acid is essentially the same across all organisms; the pyruvate intermediate is converted to lactic acid by a lactate dehydrogen‐ ase (LDH). Thus for hexose sugars, the theoretical yield is 2 moles of lactate per mole of sugar (or 1g sugar for 1g lactate). This enzymatic catalysis has the advantage over its chemical counterpart to be stereospecific: both L-lactate-dehydrogenase (L-LDH) and Dlactate-dehydrogenase (D-LDH) exist, generating either L-lactate or D-lactate respectively [65]. Both are NAD-dependant (nicotinamide adenine dinucleotide) and may be found alone or together in wild lactate-producing microbial strains. Since optical purity of lac‐ tate is a major requirement for the lactate industry, research focuses on stereospecificity as much as yields and productivity [61,66-70].

An efficient lactate producer has to display specific attributes, mainly the adaptability to low-cost substrates, high selectivity of desired enantiomer (L, D or both), high optimal tem‐ perature for decreased contamination risks, low pH tolerance and high performances (yield and productivity). LAB display appreciable performances, but lack a low pH tolerance, which implies uses of a pH control apparatus during the fermentation process. LAB optimal pH is near neutral, but the pKa of lactic acid being 3.8, an alkali agent, usually Ca(OH)2, must be used thus generating calcium lactate. After typical batch fermentation, the medium is acidified with H2SO4 therefore regenerating and purifying the lactic acid [64]. Another drawback of LAB is their requirement for a complex growth medium, since they are auxo‐ troph for certain amino acids and vitamins [71]. In order to overcome this problem, many fungi were also investigated for lactate production. Strains of *Rhizopus*, *Mucor* and *Monilla sp.* have shown potential whilst other fungi even displayed amylolytic activity, which could lead to a direct starch-to-lactate conversion [72-74].

Most researches still focuses on hexose conversion, and research group have optimized strains and process strategies in order to obtain high lactate titers, yields and productivities. Ding and Tan [75] developed a glucose fed-batch strategy using *L. casei* and generating up to 210 g/L of lactic acid with a 97 % yield. Chang *et al.* [76] proposed a continuous high cell density reactor strategy yielding a titer of 212.9 g/L and productivity of 10.6 g/L/h with *Lb. rhamnosus*. Dumbrepatil *et al.* [77] created a *Lb. delbrueckii* mutant by ultraviolet (UV) muta‐ genesis producing 166 g/L with productivity of 4.15 g/L/h in batch fermentation. Genetically engineered non-LAB biocatalysts yet have to match the performances of highly efficient wild LAB. In fact, *C. glutamicum*, *S. cerevisiae* and *E. coli* recombinant have been developed, but with limited success [61].

The search for lignocellulose-to-lactate biocatalysts have led to the discovery of many strains of pentose-utilizing LAB. *Lb. pentosus ATCC8041* [78, 79], *Lb. bifermentans DSM20003* [80], *Lb. brevis* [81], *Lb. Plantarum* [82], *Leuconostoc lactis* [83, 84], and *E. mundtii QU 25* [85, 86]. Lactic acid produced from xylose per say has been investigated by few [84,85, 87, 88], but with mitigated results, mainly due to the fact that the pentose-utilizing LAB do not perform as well in pentoses as in hexoses-rich metabolism. This phenomenon is most likely due to the fact that pentoses are metabolized in the PK pathway (phospho‐ ketolase), thus for a given strain, even if hexoses are fermented through an homofermen‐ tative route, pentose will yield heterofermentative products (i.e. acetic and lactic acid) [78, 89]. Nevertheless, Tanaka *et al.*[84] have shown that in addition to the PK, *L. lactis* could metabolize xylulose-5-phosphate (X5P), an intermediate pentose catabolite, through the pentose phosphate pathway (PPP). The theoretical yield through the PPP is 5 moles of lac‐ tate for 2 moles of pentoses, but through the PK it decreases to 1:1 [61], thus, the conver‐ sion advantage of the PPP is obvious. Okano *et al.* [87,89] demonstrated this approach by creating a pentoses-utilizing *Lb. plantarium* recombinant in which the native L-lactate de‐ hydrogenase (L-LDH) gene was disrupted, leaving only the homologous D-lactate dehy‐ drogenase (D-LDH) active. However, this strain produced both acetic and D-lactic acid; hence the PK gene (*xpk1*) was substituted by a heterologous transketolase (*tkt*) from *L. lac‐ tis*, thereby shifting heterolactic fermentation to a homolactic one.

**Strain Gen Eng Str Medium Process**

hydrolysates

bagasse

Sludge

pomace


Batch SSF

L. lactis IO-1 - Xylose Batch 33.3 - 0.68 - [84]

*DSM 20003* - Wheat bran

*<sup>3254</sup>* - Cassava

UV mutagenesis

UV mutagenesis

Disruption of endogenous LDH gene. Replacment of endogenous PK (xpk1) gene with heterologous *tkt* to redirect the PK pathway to the PPP.

Idem as above. Disruption of 2nd PK gene (xpk2) to terminate acetate production.

*ATCC 7469* - Paper

Replacement of native pdc1 and *pdc5* by heterologous bovine L-LDH gene.

*Lb. bifermentas*

*Lb. casei NCIMB*

*Lb. delbrueckii Uc-3*

*Lb. lactis RM 2-24*

*Lb. plantarum* ΔldhL1-xpk1::tkt

*Lb. plantarum* ΔldhL1-xpk1::tkt-Δxpk2

Lb. rhamnosus

Lb. rhamnosus *ATCC 9595 (CECT288)*

*S. cerevisiae* recombinant

**LA (g/L)**

Cellobiose Batch 90 40 0.9 2.25 [93] Molasse Batch 166 40 0.95 4.15 [77]

Arabinose Batch 38.6 28 0.82 1.37 [89]

Xylose Batch 41.2 60 0.89 0.67 [87]

Glucose Batch 82.3 192 0.83 0.43 [97]

73 168 0.97 0.45 [95]

Batch 32.5 6 0.88 5.41 [96]

Cellobiose Batch 80 48 0.8 1.66

Cellulose Batch SSF 73 48 0.73 1.52

Batch SSF\*\*

**Tf (h)**

Batch 62.8 60 0.83 1.17 [80]

**Yield (g/g)**

Biofuels and Co-Products Out of Hemicelluloses

83.8 60 0.96 1.4 [92]

**Prd (g/L/h)**

http://dx.doi.org/10.5772/52645

**Ref**

15

[94]

Modification of yeast strains in order to achieve xylose-to-lactate conversion has also been investigated, as an example Ilmen *et al.* [90] expressed the L-LDH gene from *L. helveticus* in *P. stipitis* and was able to reach a titer of 58 g/L of lactate with a yield of 58 %. These results were obtained despite the fact that no effort had been made to silence the native PDC/ADH (pyruvate decarboxylase/alcohol dehydrogenase) ethylic pathway, consequent‐ ly 4.5 g/L of ethanol was simultaneously produced as the endogenous PDC rivalled against the recombinant L-LDH for pyruvate. Tamakawa *et al.* [88] went further by trans‐ forming *C. utilis*, disrupting the native *pdc1* gene, and expressing heterologous LDH, XR (xylose reductase), XDH (xylitol dehydrogenase) and XK (xylulokinase) enzymes. Further‐ more, to prevent the redox imbalance, they increased the XR's NADH (reduced nicotina‐ mide adenine dinucleotide) affinity by site-directed mutagenesis. In batch culture this recombinant was able to yield titers up to 93.9 g/L of lactate at a yield of 91 %. Table 3 shows the most recent and most efficient strains developed for lactic acid production, both from hexoses and pentoses.




**4. Conversion of xylose under reducing conditions**

Xylose, as all the other carbohydrates that can be isolated from lignocellulosic biomass, has a carbonyl function that is susceptible to transformations, including reduction. One of the most common compounds that can be derived from xylose is xylitol, a pentahydroxy chiral

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Amongst the most reported catalysts in the literature are nickel and Raney nickel. According to Wisniak *et al*. [103] they are good catalysts for the production of xylitol from xylose with total conversion at 125 °C and 515 psi. In the same year, the authors published the use of ruthenium, rhodium and palladium for the reduction of xylose [104] concluding that the ef‐ ficiency of those metals was declining in the order Ru>Rh>Pd at temperatures around 100-125 °C under pressure. Mikkola *et al*. [105, 106] also used nickel as a catalyst by ultrason‐ ic process that generated close to 50 % conversion of xylose to xylitol. From this process was reported that an important problem was the deactivation of the catalyst. Utilisation of nickel also led to the publication of two patents, one in 2003 [107] and another in 2007 [108]. In the case of the first, the concept relied on the isomerization of D-xylose to L-xylose prior to cata‐

Ruthenium as well as ruthenium-based compounds has also been reported as catalysts for the reduction of xylose to xylitol. Ruthenium has been operated at temperatures between 90

[110]. Conversion rates for the latter have been reported to reach 35 % to xylitol for the latter with coproduction of glycerol and ethylene glycol. Ruthenium chloride (RuCl3) has also

been reported as a catalyst for the reduction of xylose to xylitol [111, 112].

C under pressure using ruthenium supported either on silica [109] or on carbon

**4.1. The chemical pathway**

compound as depicted in Figure 9.

**Figure 9.** Simplified conversion of D-xylose to D-xylitol.

lytic reduction under a nickel catalyst.

°

C and 110 °

*\*\*SSF = simultaneous saccharification and fermentation*

**Table 3.** Lactic acid concentration (LA), time of fermentation (Tf), yield and production rate for the most common microorganisms used for the biological conversion of xylose to lactic acid

Lactic acid seems to be, on the biological as well as on the chemical point of view the best possible compound that could be derived from a based-catalysed reaction of xylose. Race‐ mic mixtures of lactic acid (most probably derived from chemical synthesis) can be evaluat‐ ed to 1150 USD/tonne [100] whilst the pure isomer was reported to have a price market around 1750 USD/tonne [101]. As in many cases, the price will vary proportionally with pu‐ rity of the compound. Utilisation of lactic acid on the market is mostly related to polymers, food, pharmaceutical and detergents. The annual world demand for the compound should reach a little more than 367 Ktonnes/year by 2017 [102].

#### **4. Conversion of xylose under reducing conditions**

#### **4.1. The chemical pathway**

Xylose, as all the other carbohydrates that can be isolated from lignocellulosic biomass, has a carbonyl function that is susceptible to transformations, including reduction. One of the most common compounds that can be derived from xylose is xylitol, a pentahydroxy chiral compound as depicted in Figure 9.

**Figure 9.** Simplified conversion of D-xylose to D-xylitol.

Amongst the most reported catalysts in the literature are nickel and Raney nickel. According to Wisniak *et al*. [103] they are good catalysts for the production of xylitol from xylose with total conversion at 125 °C and 515 psi. In the same year, the authors published the use of ruthenium, rhodium and palladium for the reduction of xylose [104] concluding that the ef‐ ficiency of those metals was declining in the order Ru>Rh>Pd at temperatures around 100-125 °C under pressure. Mikkola *et al*. [105, 106] also used nickel as a catalyst by ultrason‐ ic process that generated close to 50 % conversion of xylose to xylitol. From this process was reported that an important problem was the deactivation of the catalyst. Utilisation of nickel also led to the publication of two patents, one in 2003 [107] and another in 2007 [108]. In the case of the first, the concept relied on the isomerization of D-xylose to L-xylose prior to cata‐ lytic reduction under a nickel catalyst.

Ruthenium as well as ruthenium-based compounds has also been reported as catalysts for the reduction of xylose to xylitol. Ruthenium has been operated at temperatures between 90 ° C and 110 ° C under pressure using ruthenium supported either on silica [109] or on carbon [110]. Conversion rates for the latter have been reported to reach 35 % to xylitol for the latter with coproduction of glycerol and ethylene glycol. Ruthenium chloride (RuCl3) has also been reported as a catalyst for the reduction of xylose to xylitol [111, 112].

Treatment of carbohydrates at a higher severity leads to the hydrogenolysis, implying not only the carbonyl compounds being reduce to alcohol but a breakage of the carbon-carbon bonds in the original carbohydrate. Recent work [113] shows that temperature above 250 ° C and pressure between 600-1000 psi, can lead to conversion of xylose to ethylene glycol, pro‐ pylene glycol and glycerol, as depicted in Figure 10 below.

 and *Corynebacterium* species that were able both to grow and produce xylitol with xylose as sole carbon source, although the reported yields were very low. In early work [123, 124], it was found that both *Corynebacterium* and *Enterobacter liquefaciens* strains were able to grow and produce xylitol from xylose although gluconate had to be present as cosubstrate. Never‐ theless, studies using wild bacterial strains for xylitol production are scares [122, 125-127]. In most metabolic pathways, bacteria go through direct xylose to xylulose conversion via iso‐ merisation, bypassing the xylitol intermediate. Subsequently, xylulose is phosphorylated in X5P and can be metabolized by most prokaryotes and eukaryotes via the PPP, or the PK

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19

Although the fact that yeast and fungi are generally more efficient xylitol producers than bacteria is widely recognized [129], certain highly productive species such as *Candida* are ac‐ tually known for their pathogenic nature [130]. Moreover, construction of recombinant yeasts by introduction of xylose reduction pathway in GRAS species such as *S. cerevisiae* have been accomplish, although these recombinant still have to match the productivities found using non-GMO organisms (genetically modified *organisms*) [131-134]. Bacterial spe‐ cies on the other hand present high yields, fast metabolism and many GRAS (generally rec‐ ognized as safe) species with recombinant strains often display higher efficiencies than their

It was found that the catabolic rate of xylose is usually enhanced by the presence of a cosubstrate such as glucose [136, 137]. However, most organisms preferentially use glucose to any other sugars due to allosteric competition in sugar transport and/or repression of other carbon catabolites [138, 139]. Thus, a suitable biocatalyst would have to simultaneously me‐ tabolize both substrates. This functionality was achieved in *E. coli* [140]by replacing the pu‐ tative cAMP-dependent receptor protein (CRP) with a cAMP-independent mutant, which also expressed a plasmid-based xylose transporter. Similarly, some authors [125] used this approach as well as inserting the heterologous XR gene and silencing the endogenous xylose isomerase (XI). Alternatively, heterologous XR and XDH may be introduced and the puta‐

Other well suited candidates for such a bioconversion would be LAB, offering the advant‐ age of an energy metabolism completely independent of their limited biosynthetic activity, thus their glycolysis pathways may be engineered without disturbing other key structural pathways [129]. By introduction of yeast XR gene, as well as a heterologous xylose trans‐ porter in *L. lactis,* they showed that bacterial productivity and yield might reach those of the best yeasts. Even if all xylose is not consumed when in high initial concentration, the non-

Early work done on *Corynebacterium glutamicum* showed another alternative for the produc‐ tion of xylitol but the necessity of inserting gluconate as co-substrate for NADPH (nicotina‐ mide adenine dinucleotide phosphate) regeneration rendered the application non economical [122,124]. Sasaki *et al.* [141] developed a *C. glutamicum* recombinant achieving si‐ multaneous co-utilization of glucose/xylose. This was done by introducing the pentose transporter area in *C. glutamicum* chromosomal DNA (deoxyribonucleic acid). *C. glutamicum* is a noticeable candidate for its non-pathogenic and gram-positive nature, as well as its ex‐

pathogenic and anaerobic nature of *L. lactis* is a notable advantage.

pathway in the case of heterolactic bacteria (Figure 11) [128].

non-altered counter-part [135].

tive XK (*xylB* gene) silenced.

**Figure 10.** Simplified conversion of D-xylose to ethylene glycol, propylene glycol and glycerol as reported by Crabtree *et al*. [113].

Production of ethylene glycol and glycerol has also been reported by Guha *et al*. [110] as a side product of their xylitol production. Hydrogenolysis of xylitol is a logical suite for re‐ duction of xylose and specific work has been reported using different catalytic systems and experimental setups. As an example, it was recently reported [114] that xylitol could be con‐ verted into a mixture of polyols and different other products as formic acid and lactic acid as well as xylitol, which, according to the previously mentioned work in this chapter, is giv‐ en when xylose is submitted to a noble metal catalyst under hydrogen. In this specific case, the catalyst was platinum supported on carbon under a base-catalyzed matrix. Chopade *et al*. [115] also presented a patent reporting the conversion of carbohydrates (including xylose) into polyols using a ruthenium catalyst as did Dubeck and Knapp in 1984 [116].

In 2010 it was reported the use of nickel as a catalyst for hydrogenolysis of xylose [117] whilst Kasehagen [118] reported hydrogenolysis of carbohydrates under a nickel-iron-cop‐ per catalyst using a matrix of alkali salts with glycerol as the main product. The effects of nickel was studied by Wright [119] but this time using tungsten as a co-catalyst. Finally, there is a report about hydrogenolysis of carbohydrates under a rhenium catalyst [120].

#### **4.2. The biological pathway**

Only a few bacteria have been shown to naturally produce xylose as a metabolite. It has been showed [121] that a bacteria belonging to the genus *Gluconobacter* was able to produce xylitol from arabitol by way of a membrane-bound D-arabitol deshydrogenase (AraDH), fol‐ lowed by a soluble XDH. Rangaswamy *et al*. [122] isolated strains of *Serratia*, *Cellulomonas*  and *Corynebacterium* species that were able both to grow and produce xylitol with xylose as sole carbon source, although the reported yields were very low. In early work [123, 124], it was found that both *Corynebacterium* and *Enterobacter liquefaciens* strains were able to grow and produce xylitol from xylose although gluconate had to be present as cosubstrate. Never‐ theless, studies using wild bacterial strains for xylitol production are scares [122, 125-127]. In most metabolic pathways, bacteria go through direct xylose to xylulose conversion via iso‐ merisation, bypassing the xylitol intermediate. Subsequently, xylulose is phosphorylated in X5P and can be metabolized by most prokaryotes and eukaryotes via the PPP, or the PK pathway in the case of heterolactic bacteria (Figure 11) [128].

Although the fact that yeast and fungi are generally more efficient xylitol producers than bacteria is widely recognized [129], certain highly productive species such as *Candida* are ac‐ tually known for their pathogenic nature [130]. Moreover, construction of recombinant yeasts by introduction of xylose reduction pathway in GRAS species such as *S. cerevisiae* have been accomplish, although these recombinant still have to match the productivities found using non-GMO organisms (genetically modified *organisms*) [131-134]. Bacterial spe‐ cies on the other hand present high yields, fast metabolism and many GRAS (generally rec‐ ognized as safe) species with recombinant strains often display higher efficiencies than their non-altered counter-part [135].

It was found that the catabolic rate of xylose is usually enhanced by the presence of a cosubstrate such as glucose [136, 137]. However, most organisms preferentially use glucose to any other sugars due to allosteric competition in sugar transport and/or repression of other carbon catabolites [138, 139]. Thus, a suitable biocatalyst would have to simultaneously me‐ tabolize both substrates. This functionality was achieved in *E. coli* [140]by replacing the pu‐ tative cAMP-dependent receptor protein (CRP) with a cAMP-independent mutant, which also expressed a plasmid-based xylose transporter. Similarly, some authors [125] used this approach as well as inserting the heterologous XR gene and silencing the endogenous xylose isomerase (XI). Alternatively, heterologous XR and XDH may be introduced and the puta‐ tive XK (*xylB* gene) silenced.

Other well suited candidates for such a bioconversion would be LAB, offering the advant‐ age of an energy metabolism completely independent of their limited biosynthetic activity, thus their glycolysis pathways may be engineered without disturbing other key structural pathways [129]. By introduction of yeast XR gene, as well as a heterologous xylose trans‐ porter in *L. lactis,* they showed that bacterial productivity and yield might reach those of the best yeasts. Even if all xylose is not consumed when in high initial concentration, the nonpathogenic and anaerobic nature of *L. lactis* is a notable advantage.

Early work done on *Corynebacterium glutamicum* showed another alternative for the produc‐ tion of xylitol but the necessity of inserting gluconate as co-substrate for NADPH (nicotina‐ mide adenine dinucleotide phosphate) regeneration rendered the application non economical [122,124]. Sasaki *et al.* [141] developed a *C. glutamicum* recombinant achieving si‐ multaneous co-utilization of glucose/xylose. This was done by introducing the pentose transporter area in *C. glutamicum* chromosomal DNA (deoxyribonucleic acid). *C. glutamicum* is a noticeable candidate for its non-pathogenic and gram-positive nature, as well as its ex‐

**•** disruption of XK native gene (*xylB*);

**Strain Genetic Engineering**

**Strategy**


Expression heterologous XR gene from *P. stipitis*.

Expression of *araE* pentose transporter gene. Disruption of *ldhA*. Single site mutation of heterologous XR gene. Disruption of *xylB*& PTSfru genes.

Overexpression ALD6 & ACS1 genes.Expression of *P. stipitis* XR gene.

Expression of *P. stipitis* XR gene.Expression of *Lb. brevisxylT* symporteur.

PTSfru).

*Candida athensensis* SB18

*C. tropicalis* KCTC 10457

*C. tropicalis* KFCC 10960

*C. tropicalis* KCTC 10457

*S. cerevisiae*

*Corynebacterium glutamicum* CtXR7

*D. hansenii* NRRL Y-7426

*S. cerevisiae*

*Lactobacillus brevis* NZ9800

**•** disruption of phosphoenolpyruvate-dependent fructose phosphotransferase (*ptsF* gene;

**Xylose g/L**

*C. tropicalis* ASM III - 93% 200 130 120 1.08 Batch limited O2 [145] *Candida* sp. 559-9 - 99% 200 173 121 1.44 Batch limited O2 [146]

C. guilliermondii - 73% 250 - - - Fed Batch limited O2 [149]

*C. tropicalis* - 69% 100 - - 5.7 Cell recycling/ limited O2 [151] *C. tropicalis* - 85% 214 182 15 12 cell recycling/ limited O2 [147]

95% 190 - - 0.4


~100% 20 91.3 60 1.76

~100% 160 75 41 2.72


**Xylitol (g/L)**

**Tf (h)**

83% 250 207.8 175 1.15 Batch limited O2

79% 200 151.71 156 0.97 Batch limited O2




**Prd (g/l/h)**

87% 300 256.5 250 0.97 Fed Batch limited O [144] <sup>2</sup>

**Process Strategy**

Biofuels and Co-Products Out of Hemicelluloses

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Fed Batch/ Cell recylcing/ Glucose cosubstrate/ limited O2

> Fed batch/ Glucose cosubstrate

Fed batch/ Glucose cosubstrate/ 40g/L dry cell

Batch/ Detoxified grape marc hydrolysates

Fed batch/ Glucose cosubstrate

Fed batch/ Glucose cosubstrate

**Reference**

21

[150]

[152]

[135]

[153]

[154]

[129]

**Yield g/g**

*C. tropicalis* - 82% 750 189 58 4.94

**Figure 11.** Glycolysis and phosphoketolase (pentose phosphate) pathways in lactic acid bacteria (1) glucokinase, (2) phosphoglucose isomerase, (3) phosphofructokinase, (4) fructose 1,6-bisP aldolase, (5) triose-phosphate isomerase, (6) glyceraldehyde-3P dehydrogenase, (7) phosphoglycerate kinase, (8) phosphoglycerate mutase, (9) enolase, (10) pyruvate kinase, (11) lactate dehydrogenase, (12) hexokinase, (13) glucose-6P dehydrogenase, (14) 6-phosphogluco‐ nate dehydrogenase, (15) ribulose-5P 3-epimerase, (16) xylulose-5P phosphoketolase, (17) phosphotransacetylase, (18) acetaldehyde dehydrogenase, (19) alcohol dehydrogenase; (20) pentose kinase, (21) pentose phosphate epimer‐ ase or isomerase, (22) acetate kinase. *CoA* coenzyme A.

tensive use for amino and nucleic acid industrial synthesis [142, 143]. It was established [135] that xylitol productivity may be improved by disabling the xylitol import system (ptsF gene) and suggested that more work done on xylitol export system and redox balance may yield further improvements. Nevertheless, their CtXR7 *C. glutamicum* recombinant attained a productivity of 7.9 g/L/h and final xylitol concentration of 166 g/L after 21 h (see Table 4). This was achieved by (to date this is considered the best xylitol bacterial producer):




non-preferred carbon substrate for *S. cerevisiae* and do not provide sufficient energy for

*C. tropicalis* is a candidate of choice for xylitol production among the few native strains re‐ ported as the best xylitol producers to date (see Table 4) and this research for native strains and genetically engineered recombinant is still under way today [155-157]. As in *S. cerevisiae*, the PPP is the major NADPH biosynthesis pathway and efforts have been made to increase its flux. Ahmad *et al.* [165] recently successfully increased the metabolic flux toward PPP for NADPH regeneration, thereby enhancing xylitol production of the origi‐ nal strain by 21 %. This was done by disrupting XDH putative gene, and over-expressing homologous glucose-6-phosphate dehydrogenase (G6PDH) and 6-phosphogluconate dehy‐ drogenase (6-PGDH). Table 4 summarize the best xylitol producing strains found in the

Reduction of xylose either at low or at high severity thus producing either xylitol or polyols (including glycerol) is a process driven by the price of hydrogen. On the other hand, the market for small polyols as ethylene or propylene glycol may generate more opportunity than the xylitol market. Xylitol market value is between 3650 and 4200 USD/tonne [166] whilst ethylene glycol is reported at a market price of 980-1500 USD/tonne [167] and propy‐ lene glycol at 1500-1700 USD/tonne [168]. The market for each of the previously mentionned compound is around 100 Ktonnes/y for xylitol [169], 19 Mtonnes/y for ethylene glycol [170] and 1.4 Mtonnes/y for propylene glycol. Although the market for smaller polyols may seem to be larger, as an example conversion of xylose to ethylene glycol and propylene glycol would require 3 times as much hydrogen if compared to xylitol. Since the price for hydro‐ gen can be estimated roughly at 4.5-5 USD/Kg, the very concept of polyols production relies on the efficiency of the hydrogenolysis process therefore explaining why many of the report‐

Oxidation of xylose has been numerously reported in the literature although focus interest, both on the biological as well as chemical point of view has been focused toward a simple

Oxidation of xylose has been reported for a variety of different metallic catalyst including gold for high conversion rates [171]. Using a process performed a little higher than room temperature in a basic pH for 1 hour, they were able to reach a 78 % conversion of xylose to xylonic acid. Using comparable catalyst, Pruesse *et al*. [172] were able to reach 99 % selectivi‐ ty with a conversion rate of 21 mmol/min/g (Au) in a continuous reactor. Nevertheless, con‐ trarily to Bonrath, Pruesse and co-worker used a mixture of gold and palladium to perform

C as compared to 40 °

C).

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growth and metabolism [164].

literature up to date.

ed litterature in this chapter are patents.

oxidation of xylose to xylonic acid (see Figure 12).

this oxidation and temperature slightly higher (60 °

**5.1. The chemical pathway**

**5. Conversion of xylose under oxidizing conditions**

**Table 4.** Overview of the different strains allowing conversion of xylose to xylitol including yields, fermentation time (Tf), production (Prd) and the process strategy.

As previously discussed for ethanol, the redox imbalance that often occurs from XR/XDH preferential use of NADPH/NAD+ cofactors is a key factor for xylitol accumulation in the cell. In most yeast studied, it has been shown that XR has a marked preference for NADPH, while XDH has a quasi-unique specificity for NAD+ [126]. The main exception being *P. stipitis* who shows a nearly by-specificity for NAD(P)(H) for its XR and *P. tanno‐ philus* whose XDH shows a higher activity with NADP+ than NAD+ [158] proposed a the‐ oretical maximum xylitol yield in yeasts of 0.905 mol of xylitol per mol of xylose when NADH was efficiently used as cofactor by the XR or under aerobic condition where the NADH can be oxidized back to NAD+ in the respiratory chain. Otherwise, under anaero‐ bic conditions, the theoretical yield drops to 0.875. These yields follow the equations (1) and (2) below respectively:

$$126\text{ xylose} + 3\text{ O}\_2 + 6\text{ ADP} + 6\text{ P}\_i + 48\text{ H}\_2\text{O} \rightarrow 114\text{ xyltool} + 6\text{ ATP} + 60\text{ CO}\_2\tag{1}$$

$$48\text{ xylose} + 15\text{ H}\_2\text{O} \rightarrow 42\text{ Xylited} + 2\text{ ethanol} + 24\text{ CO}\_2\tag{2}$$

Owing the better yield both in xylitol and ATP (adenosine triphosphate) under oxygen-lim‐ ited xylitol production, aeration is a crucial parameter. As a general trend, xylitol produc‐ tion increases when oxygen is allowed in the medium under a certain threshold concentration [159]. This preference is yeast specific since for *P. stipitis* it is reported that the absence of dissolved oxygen is needed for optimal xylitol production; while *P. tannophilus* reaches maximum yields under anoxic conditions [160, 161].

Many strains of *S. cerevisiae* have been transformed for xylose utilization in the early 90's. As for xylose-to-xylitol, Hallborn *et al.* [152] reported a highly efficient conversion of xy‐ lose to xylitol (95 % of theoretical). It has been suggested that the incapacity of *S. cerevi‐ siae* to rapidly replenish its NADPH pool from its PPP during xylose metabolism is what causes the metabolic bottleneck [162, 163]. This is mainly due to the fact that xylose is a non-preferred carbon substrate for *S. cerevisiae* and do not provide sufficient energy for growth and metabolism [164].

*C. tropicalis* is a candidate of choice for xylitol production among the few native strains re‐ ported as the best xylitol producers to date (see Table 4) and this research for native strains and genetically engineered recombinant is still under way today [155-157]. As in *S. cerevisiae*, the PPP is the major NADPH biosynthesis pathway and efforts have been made to increase its flux. Ahmad *et al.* [165] recently successfully increased the metabolic flux toward PPP for NADPH regeneration, thereby enhancing xylitol production of the origi‐ nal strain by 21 %. This was done by disrupting XDH putative gene, and over-expressing homologous glucose-6-phosphate dehydrogenase (G6PDH) and 6-phosphogluconate dehy‐ drogenase (6-PGDH). Table 4 summarize the best xylitol producing strains found in the literature up to date.

Reduction of xylose either at low or at high severity thus producing either xylitol or polyols (including glycerol) is a process driven by the price of hydrogen. On the other hand, the market for small polyols as ethylene or propylene glycol may generate more opportunity than the xylitol market. Xylitol market value is between 3650 and 4200 USD/tonne [166] whilst ethylene glycol is reported at a market price of 980-1500 USD/tonne [167] and propy‐ lene glycol at 1500-1700 USD/tonne [168]. The market for each of the previously mentionned compound is around 100 Ktonnes/y for xylitol [169], 19 Mtonnes/y for ethylene glycol [170] and 1.4 Mtonnes/y for propylene glycol. Although the market for smaller polyols may seem to be larger, as an example conversion of xylose to ethylene glycol and propylene glycol would require 3 times as much hydrogen if compared to xylitol. Since the price for hydro‐ gen can be estimated roughly at 4.5-5 USD/Kg, the very concept of polyols production relies on the efficiency of the hydrogenolysis process therefore explaining why many of the report‐ ed litterature in this chapter are patents.

#### **5. Conversion of xylose under oxidizing conditions**

#### **5.1. The chemical pathway**

Oxidation of xylose has been numerously reported in the literature although focus interest, both on the biological as well as chemical point of view has been focused toward a simple oxidation of xylose to xylonic acid (see Figure 12).

Oxidation of xylose has been reported for a variety of different metallic catalyst including gold for high conversion rates [171]. Using a process performed a little higher than room temperature in a basic pH for 1 hour, they were able to reach a 78 % conversion of xylose to xylonic acid. Using comparable catalyst, Pruesse *et al*. [172] were able to reach 99 % selectivi‐ ty with a conversion rate of 21 mmol/min/g (Au) in a continuous reactor. Nevertheless, con‐ trarily to Bonrath, Pruesse and co-worker used a mixture of gold and palladium to perform this oxidation and temperature slightly higher (60 ° C as compared to 40 ° C).

**Figure 12.** Simplified conversion of xylose to xylonic acid

Copper has also been indirectly investigated for the conversion of xylose to xylonic acid in that sense that Van der Weijden *et al*. [173] used C5 sugars (including xylose) for the reduc‐ tion of copper sulfate in wastewater with very promising results. Although emphasis was not put on the carbohydrate itself, results showed that the reduction of copper from (II) to elemental was possible yet economical at larger scale. Xylonic acid was also observed as byproduct of xylose oxidation using chlorine, as a side reaction of lignin oxidation. In this work [174], the concentration of xylonic acid increased by a factor of 40 after the chlorination process. Interesting enough, the xylitol concentration also increased, which might lead to the conclusion that oxidation, was probably not the sole factor here and that side reactions as the Cannizarro reaction between two xylose molecules could have been occurring. Jokic *et al*. [175] showed that it was possible up to an efficiency of 80 % to convert xylose simultane‐ ously to xylonic acid and xylitol using electrotechnologies. Such process could be to a cer‐ tain extent compared to the Cannizarro reaction where the original aldehyde is acting as redox reagent.

Severer oxidizing conditions leads to a breakage of the carbon-carbon bonds in the carbohy‐ drate molecule leading to the production, mostly, of small organic acids as formic and acetic acid on glucose [180]. A simplified scheme of such a reaction is presented in Figure 14 below:

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**Figure 13.** Simplified scheme for the conversion of xylose xylaric acid

**Figure 14.** Simplified scheme for the conversion of xylose to formic acid under more severe oxidizing conditions.

An example of sever oxidation of xylose in a mixture of hydrogen peroxide and ammonium hy‐ droxide have been recently reported [181] with a conversion of 96 % at room temperature for 1 h. Similar conversion of xylose was reported [182] for a process using oxygen and a molybde‐

Further oxidation of xylose leads to a trihydroxydiacid, more specifically xylaric acid as de‐ picted in Figure 13 below.

Conversion of C5 sugars and to a smaller extent of xylose into aldaric acids has been descri‐ bed in literature in a few reports. Kiely *et al*. [176] reported that a conversion up to 83 % xy‐ lose into 2,3,4-trihydroxyglutaric acid was achievable in a reaction mixture composed of nitric acid and NaNO2. The side product of this reaction was reported to be disodium tetra‐ hydroxysuccinate. Conversion of xylose to xylaric adic was also reported [177] using oxygen under a platinum catalyst all of this in an alkali promoted medium. Comparable conversion process [178] was obtained without any alkali, though still performed the reaction in water at 90 °C under 75 psi of oxygen. The conversion for this process was 29 %. Fleche *et al*. [179] reported a maximum conversion of 58% once again using platinum supported on alumina.

**Figure 13.** Simplified scheme for the conversion of xylose xylaric acid

Severer oxidizing conditions leads to a breakage of the carbon-carbon bonds in the carbohy‐ drate molecule leading to the production, mostly, of small organic acids as formic and acetic acid on glucose [180]. A simplified scheme of such a reaction is presented in Figure 14 below:

**Figure 14.** Simplified scheme for the conversion of xylose to formic acid under more severe oxidizing conditions.

An example of sever oxidation of xylose in a mixture of hydrogen peroxide and ammonium hy‐ droxide have been recently reported [181] with a conversion of 96 % at room temperature for 1 h. Similar conversion of xylose was reported [182] for a process using oxygen and a molybde‐ num and vanadium catalyst. The reaction was done for 26 h at 353 K and 30 bar for a conversion of up to 54 % into formic acid with carbon dioxide as by-product.

At this point it is rather hard to verify the potential or the economic value of oxidation prod‐ ucts from xylose. Complete oxidation to formic acid could be the most suitable approach at this point since the market for xylonic and xylaric acid is not as well defined as for the sim‐ ple methanoic acid with its actual market value between 750-950 USD/tonne [200] and an annual world demand suspected to reach 573 Ktonnes in 2012 [201]. Conversion of xylaric acid into glutaric acid (pentanedioic acid) would lead to a very interesting market as a plas‐ ticizer but dehydration or reduction of the three central hydroxyl groups may be a challenge that could be winning at lab scale although a multiple synthesis pathway would be very dif‐

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ficult to reach economic at an industrial level.

**Figure 15.** Proposed pathway ford-xylose metabolism in *C. crescentus* [193].

#### **5.2. The biological pathway**

Xylonic acid synthesis from xylose has been reported for *Acetobacter* sp. [183], *Enterobacter cloacea* [184], *Erwinia* sp. [185, 186], *Fusarium lini* [187], *Micrococcus* sp. [188], *Penicillium cory‐ lophilum*, *Pichiaquer cuum* [185], *Pseudomonas* sp. [189, 190], *Pullularia pullulans* [191], *Glucono‐ bacter* and *Caulobacter* [192, 193].

In metabolic pathways, xylose is converted to xylonate via 2 key enzymes. First, a xylose de‐ hydrogenase (XD) oxidizes xylose to D-xylono-1,4-lactone (xylonolactone) using either NAD + or NADP+ as cofactor. This reaction is followed by the hydrolysis of xylonolactone to xylo‐ nate either spontaneously or by an enzyme with lactonase activity [194, 195]. It is hypothe‐ sized that *Pseudomonas* and *Gluconobacter sp.* both carry a membrane-bound pyrroloquinoline quinine (PQQ)-dependent XD and a cytoplasmic one [195, 196]. Stephens *et al.* [193] recently proposed a full xylose catabolic pathway for *C. crescentus*. Note that a similar pathway was proposed for arabinose yielding L-arabonate [197]. As shown in Figure 15, the proposed metabolic pathway for *C. crescentus* shows that xylonate is an intermediate in catabolic reactions that is quite different from the XI or XR/XDH previously discussed which were more intensively studied.

Researches on highly efficient microbial xylonic acid production are scarce compared to bio‐ fuels or xylitol. Even if the identification of xylonate producing species began as early as 1938 [187], the first attempt to isolate a possible industrial biocatalyst was done by Buchert *et al.* [185], who identified *P. fragi ATCC4973* as a potentially high efficiency xylonate producer (92 % of initial sugar converted to xylonic acid with initial xylose concentration of 100 g/L). In further work, *P. fragi* and *G. oxydans* showed yields of over 95 % but the low tolerance of those native strains to inhibitors tends to be problematic for industrial uses [192]. As dis‐ cussed above, the metabolic pathways implied by xylonate have been investigated in the re‐ cent years [193,196]. The first recombinant microorganism engineered for the industrial production of xylonate was done by Toivari *et al.* [198]. By introducing the heterologous *Tri‐ choderma reesei xyd1* gene (coding for the NADP+ dependant XD) in *S. cerevisiae*, they were able to obtain up to 3.8 g/L xylonate with 0.036 g/L/h productivity and 40 % yield. Nygard *et al.* [195] engineered *K. lactis* by introducing *T. reesei xyd1* and deleting the putative *xyl1* gene coding for the XR. Up to 19 g/L xylonate where produced when grown on a xylose (40 g/L) and galactose (10.5 g/L) medium. The native ability of fast xylose uptake was an advantage, but high intracellular xylonate concentration was observed, which may indicate difficulties with product export. Liu *et al.* [199] used similar approach engineering *E. coli* by disrupting the native xylose metabolic pathways of XI and XK (as shown in Figure 16). The native path‐ way of xylonate was also blocked by disrupting xylonic acid dehydratase genes. The XD from *C. crescentus* was introduced and 39.2 g/L of xylonate from 40 g/L of xylose in minimal medium was obtained at high productivity 1.09 g/L/h. From these results it is clear that re‐ search is at its genesis and significant efforts will be required for the creation of a highly pro‐ ductive and effective xylonate production biocatalyst.

At this point it is rather hard to verify the potential or the economic value of oxidation prod‐ ucts from xylose. Complete oxidation to formic acid could be the most suitable approach at this point since the market for xylonic and xylaric acid is not as well defined as for the sim‐ ple methanoic acid with its actual market value between 750-950 USD/tonne [200] and an annual world demand suspected to reach 573 Ktonnes in 2012 [201]. Conversion of xylaric acid into glutaric acid (pentanedioic acid) would lead to a very interesting market as a plas‐ ticizer but dehydration or reduction of the three central hydroxyl groups may be a challenge that could be winning at lab scale although a multiple synthesis pathway would be very dif‐ ficult to reach economic at an industrial level.

**Figure 15.** Proposed pathway ford-xylose metabolism in *C. crescentus* [193].

conversion price is going to be higher than classical or first generation ethanol production. Keeping that fact in mind, the conversion of cellulose to glucose itself is a major technologi‐ cal challenge since it either requires enzymes, ionic liquids or strong acids that are rather ex‐ pensive to buy or expensive to recycle and since it is of outmost importance for the

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The remaining carbon content of lignocellulosic biomass is also an important factor to be considered. Since the maximum production of ethanol from the total feedstock could vary around 300-400 L per tonne, there is at this point a necessity to generate co-products from the biomass in order to make this whole process economic at the end thus coping for techno‐ logical problem as conversion of cellulose to glucose. Lignin is one of the most abundant macromolecule on earth bested only by cellulose. The aromatic nature of lignin is a chal‐ lenge for ethanol production but not for added value compounds as aromatic monomers that could displace actual monomers used in the polymer industry that are usually obtained

Hemicelluloses are also an important part of the lignocellulosic biomass. Hemicelluloses, contrarily to cellulose that is characterized by an amorphous and a crystalline part, are high‐ ly ramified and easy to hydrolyse. Usually, a simple diluted alkali solution, acidic solution or even hot water can allow conversion of hemicellulose to simple sugars. The major prob‐ lem with hemicellulose is the heterogeneous composition including but not limited to small acids and a variety of C6 and C5 sugars. Whilst the C6 sugars could be easily fermented to ethanol, pending reduction of the organic acids and other inhibitors, the C5 sugars require

Other than the classical fermentative pathway, C5 sugars can as well be converted, biologi‐ cally as well as chemically into a wide variety of added value products and "green" com‐ pounds. In this paper, we have identified 4 pathways for the conversion of C5 sugars but

Reaction of xylose under an acid catalyst is probably one of the most investigated fields in this domain. The target for this conversion being furfural, a well-known chemical as well as precursor for other compound as furan, Me-THF, THF and furfuryl alcohol, a reactant used in the polymer industry. The best approach for the conversion of xylose furfural, to the best of our knowledge, is chemical as no microorganism allowing conversion of C5 sugars to fur‐ fural has been identified so far. The conversion of xylose to furfural was reported to reach more than 95 % for both heterogeneous and homogeneous catalyst. On the other hand, the selectivity toward furfural is not always as efficient since the latter undergoes polymerisa‐

A basic catalyst leads to a conversion of C5 sugars to lactic acid although this pathway as not been deeply investigated in the literature. Lactic acid is a compound well in demand on the market but the limitations for the chemical transformation is the lack of stereospecificity of the products. Conversion of xylose under a base catalyst leads to the production of a race‐ mic mixture of D- and L-lactic acid and thus reducing the market value of the product, par‐ ticularly if the polymer industry is targeted. On the other hand, the biological conversion of

more specifically xylose, a common carbohydrate in biomass hemicelluloses.

tion in acidic medium, which often also leads to deactivation of the catalyst.

production of the ethanol, technology is to a certain extent limited by this reality.

from non-renewable materials.

speciality yeasts for fermentation.

**Figure 16.** D-xylose and D-xylonic acid metabolic pathways in *E. coli*. The symbol X denotes that the gene is disrupted.

#### **6. Conclusion**

Second-generation ethanol or "cellulosic ethanol" relies on the utilisation of lignocellulosic biomass as a source of carbohydrates via the "bio" conversion route (keeping in mind that other pathway, as thermocatalytic pathways, may also lead to cellulosic ethanol). Produc‐ tion of ethanol thus requires isolation of cellulose from lignocellulosic matrix, then hydroly‐ sis of cellulose to glucose prior to fermentation. Both of the previously mentioned steps represent challenges for industry, but the whole economic of the process is perhaps the most challenging part of cellulosic ethanol production. Cellulose is usually available in lignocellu‐ losic biomass in the 45-60 % range which, assuming a perfect conversion implies production of 300-400 L/tonne of lignocellulosic biomass processed. At an actual price of 0.48 USD/L, each ton of biomass has a potential value of about 150-200 USD/tonne of biomass processed.

The conversion of lignocellulosic biomass is rather more complex and to a certain extent more expensive than starch-based feedstock as corn and therefore, one can assume that the conversion price is going to be higher than classical or first generation ethanol production. Keeping that fact in mind, the conversion of cellulose to glucose itself is a major technologi‐ cal challenge since it either requires enzymes, ionic liquids or strong acids that are rather ex‐ pensive to buy or expensive to recycle and since it is of outmost importance for the production of the ethanol, technology is to a certain extent limited by this reality.

The remaining carbon content of lignocellulosic biomass is also an important factor to be considered. Since the maximum production of ethanol from the total feedstock could vary around 300-400 L per tonne, there is at this point a necessity to generate co-products from the biomass in order to make this whole process economic at the end thus coping for techno‐ logical problem as conversion of cellulose to glucose. Lignin is one of the most abundant macromolecule on earth bested only by cellulose. The aromatic nature of lignin is a chal‐ lenge for ethanol production but not for added value compounds as aromatic monomers that could displace actual monomers used in the polymer industry that are usually obtained from non-renewable materials.

Hemicelluloses are also an important part of the lignocellulosic biomass. Hemicelluloses, contrarily to cellulose that is characterized by an amorphous and a crystalline part, are high‐ ly ramified and easy to hydrolyse. Usually, a simple diluted alkali solution, acidic solution or even hot water can allow conversion of hemicellulose to simple sugars. The major prob‐ lem with hemicellulose is the heterogeneous composition including but not limited to small acids and a variety of C6 and C5 sugars. Whilst the C6 sugars could be easily fermented to ethanol, pending reduction of the organic acids and other inhibitors, the C5 sugars require speciality yeasts for fermentation.

Other than the classical fermentative pathway, C5 sugars can as well be converted, biologi‐ cally as well as chemically into a wide variety of added value products and "green" com‐ pounds. In this paper, we have identified 4 pathways for the conversion of C5 sugars but more specifically xylose, a common carbohydrate in biomass hemicelluloses.

Reaction of xylose under an acid catalyst is probably one of the most investigated fields in this domain. The target for this conversion being furfural, a well-known chemical as well as precursor for other compound as furan, Me-THF, THF and furfuryl alcohol, a reactant used in the polymer industry. The best approach for the conversion of xylose furfural, to the best of our knowledge, is chemical as no microorganism allowing conversion of C5 sugars to fur‐ fural has been identified so far. The conversion of xylose to furfural was reported to reach more than 95 % for both heterogeneous and homogeneous catalyst. On the other hand, the selectivity toward furfural is not always as efficient since the latter undergoes polymerisa‐ tion in acidic medium, which often also leads to deactivation of the catalyst.

A basic catalyst leads to a conversion of C5 sugars to lactic acid although this pathway as not been deeply investigated in the literature. Lactic acid is a compound well in demand on the market but the limitations for the chemical transformation is the lack of stereospecificity of the products. Conversion of xylose under a base catalyst leads to the production of a race‐ mic mixture of D- and L-lactic acid and thus reducing the market value of the product, par‐ ticularly if the polymer industry is targeted. On the other hand, the biological conversion of xylose to lactic acid is a well-known and extensively reported process for which the produc‐ tion was reported to reach 6.7 g/L/d for genetically modified organisms as, in this specific case, *Lactobacillus sp. RKY2*. According to the reports, the production of lactic acid would be more efficient by the biological approach since it can lead to a stereospecific and a higher market value.

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Reduction of xylose can lead to many different products including xylitol for lower severi‐ ty up to diols as ethylene glycol and propylene glycol at higher severity. It is ambiguous to determine at this point if either the chemical or the biological pathway is more efficient for the production of xylitol since reports on both pathways have shown promising re‐ sults. The main problem with the xylitol market is that although it is increasing, it is fair‐ ly small and therefore it is harder to fit in a new production of xylitol. On the other hand, a more severe reduction of xylose, leading to diols, could be a very interesting opportuni‐ ty for the production of ethylene glycol and propylene glycol, two very important prod‐ ucts in the chemical industry. The downside of this approach would be the production of glycerol as a side-product.

Finally, oxidation of xylose is, at this point, the approach with the lower potential for a rapid commercialisation since the market for xylonic acid and xylaric acid is hard to size at present. The conversion process, both chemical and biological seems to have significant po‐ tential in terms of scalability but the end usage is not well defined at this point. The best option would be to produce glucaric acid from xylaric acid, which could be used as a plasti‐ cizer. On the other hand, such a process, overall rather complicated, would add a significant cost for a product that would land in the commodity range.

#### **Acknowledgement**

We would like to acknowledge Enerkem, Greenfield Ethanol, CRB Innovations and the Min‐ istry of Natural Resources of Quebec for financial support of the Industrial Chair in Cellulo‐ sic Ethanol.

#### **Author details**

Ariadna Fuente-Hernández, Pierre-Olivier Corcos, Romain Beauchet and Jean-Michel Lavoie\*

\*Address all correspondence to: jean-michel.lavoie2@usherbrooke.ca

Industrial Research Chair on Cellulosic Ethanol (CRIEC), Département de Génie Chimique et de Génie Biotechnologique, Université de Sherbrooke, Sherbrooke, Québec, Canada

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**Chapter 2**

**Production of 2nd Generation of Liquid Biofuels**

Fluctuations in the price of oil and projections on depletion of accessible oil deposits have led to national and international efforts to enhance the proportion of energy derived from renewable sources (bioenergy) with special emphasis on the transport sector (e.g. according to Directive 2009/28 EC, by 2020, 20% of energy in EU-27 should be met from renewable sources and 10% should be used in transportation). To fulfil the legal requirements, wider exploitation of biofuels made from renewable feedstocks, as a substitute for traditional liq‐ uid fuels, will be inevitable; e.g. the demand for bioethanol in the EU is expected to reach 28.5 billion litres by 2020 [1], while in America 36 billion gallons of ethanol must be pro‐ duced by 2022 [2]. Bioethanol, which has a higher octane level then petrol but only contains 66% of the energy yield of petrol, can be used as blend or burned in its pure form in modi‐ fied spark-ignition engines [2]. This will improve fuel combustion, and will contribute to a reduction in atmospheric carbon monoxide, unburned hydrocarbons, carcinogenic emis‐ sions and reduce emissions of oxides of nitrogen and sulphur, the main cause of acid rain [2]. Butanol-gasoline blends might outcompete ethanol-gasoline ones because they have bet‐ ter phase stability in the presence of water, better low-temperature properties, higher oxida‐ tion stability during long term storage, more favourable distillation characteristics and lower volatility with respect to possible air pollution. Recently performed ECE 83.03 emis‐ sion tests [3] have shown negligible or no adverse effects on air pollution by burning buta‐ nol-gasoline blends (containing up to 30% v/v of butanol) in spark ignition engines of Skoda

Although most of the world's bioethanol is currently produced from starch or sugar raw materials, attention is increasingly turning to 2nd generation biofuels made from lignocellu‐ lose, e.g. agriculture and forest wastes, fast growing trees, herbaceous plants, industrial

> © 2013 Paulová et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

© 2013 Paulová et al.; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Leona Paulová, Petra Patáková,

http://dx.doi.org/10.5772/53492

**1. Introduction**

passenger cars.

Mojmír Rychtera and Karel Melzoch

Additional information is available at the end of the chapter


### **Production of 2nd Generation of Liquid Biofuels**

Leona Paulová, Petra Patáková, Mojmír Rychtera and Karel Melzoch

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/53492

#### **1. Introduction**

Fluctuations in the price of oil and projections on depletion of accessible oil deposits have led to national and international efforts to enhance the proportion of energy derived from renewable sources (bioenergy) with special emphasis on the transport sector (e.g. according to Directive 2009/28 EC, by 2020, 20% of energy in EU-27 should be met from renewable sources and 10% should be used in transportation). To fulfil the legal requirements, wider exploitation of biofuels made from renewable feedstocks, as a substitute for traditional liq‐ uid fuels, will be inevitable; e.g. the demand for bioethanol in the EU is expected to reach 28.5 billion litres by 2020 [1], while in America 36 billion gallons of ethanol must be pro‐ duced by 2022 [2]. Bioethanol, which has a higher octane level then petrol but only contains 66% of the energy yield of petrol, can be used as blend or burned in its pure form in modi‐ fied spark-ignition engines [2]. This will improve fuel combustion, and will contribute to a reduction in atmospheric carbon monoxide, unburned hydrocarbons, carcinogenic emis‐ sions and reduce emissions of oxides of nitrogen and sulphur, the main cause of acid rain [2]. Butanol-gasoline blends might outcompete ethanol-gasoline ones because they have bet‐ ter phase stability in the presence of water, better low-temperature properties, higher oxida‐ tion stability during long term storage, more favourable distillation characteristics and lower volatility with respect to possible air pollution. Recently performed ECE 83.03 emis‐ sion tests [3] have shown negligible or no adverse effects on air pollution by burning buta‐ nol-gasoline blends (containing up to 30% v/v of butanol) in spark ignition engines of Skoda passenger cars.

Although most of the world's bioethanol is currently produced from starch or sugar raw materials, attention is increasingly turning to 2nd generation biofuels made from lignocellu‐ lose, e.g. agriculture and forest wastes, fast growing trees, herbaceous plants, industrial

© 2013 Paulová et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 Paulová et al.; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

wastes or wastes from wood and paper processing. The concept of ethanol production from lignocellulose sugars is not new. Probably the first technical attempt to degrade polysac‐ charides in wood was carried out by the French scientist Henri Braconnot in 1819 using 90% sulfuric acid [4]. His findings were exploited much later, in 1898, with the opening of the first cellulosic ethanol plant in Germany, followed by another one in 1910 in the US [5, 6]. During World War II, several industrial plants were built to produce fuel ethanol from cellu‐ lose (e.g. in Germany, Russia, China, Korea, Switzerland, US), but since the end of the war, most of these have been closed due to their non-competitiveness with synthetically pro‐ duced ethanol [7]. In spite of all the advantages of lignocellulosic as a raw material (e.g. low and stable price, renewability, versatility, local availability, high sugar content, noncompeti‐ tiveness with food chain, waste revaluation) and extensive efforts of many research groups to reduce bottlenecks in technology of lignocellulosic ethanol production (e.g. energy inten‐ sive pretreatment, costly enzymatic treatment, need for utilization of pentose/hexose mix‐ tures, low sugar concentration, low ethanol concentration), large scale commercial production of 2nd generation bioethanol has not been reopened yet [8], although many pilot and demonstration plants operate worldwide [9]. Identically, only first generation biobuta‐ nol is produced in China (approx. annual amount 100 000 t) and Brazil (approx. annual amount 8 000 t) [10]. At the 2012 London Olympic Games, British Petrol introduced its three most advanced biofuels i.e. cellulosic ethanol, renewable diesel and biobutanol. At a demon‐ stration plant at Hull UK, biobutanol, produced by Butamax (joint venture of BP and Du‐ Pont) was blended at 24 % v/v with standard gasoline and used in BMW-5 series hybrids without engine modifications [11]. As the final price of both ethanol and 1-butanol produced by fermentation is influenced mostly by the price of feedstock, the future success of industri‐ al ABE fermentation is tightly linked with the cost of pre-treatment of lignocellulosic materi‐ al into a fermentable substrate.

structure differs for different types of biomass (Table1) and it is also influenced by variety, climatic conditions, cultivation methods and location. Minor components of the cell wall are

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represented by proteoglycans, pectin, starch, minerals, terpenes, resins tannins and waxes.

**Biomass Cellulose Hemicellulose Lignin Reference** Hardwood 45-47 25-40 20-55 [17, 18] Softwood 40-45 25-29 30-60 [17, 18] Wheat straw 30-49 20-50 8-20 [19-22] Rye straw 30.9 21.5 25.3 [21] Corn fibre 15 35 8 [23] Corn cobs 35-45 35-42 5-15 [22, 23] Corn stover 39-42 19-25 15-18 [22, 23] Corn straw 42.6 21.3 8.2 [20]

Rice straw 32-47 15-27 5-24 [20, 22, 23]

Sugarcane bagasse 40 24-30 12-25 [20, 22, 23] Switchgrass 30-50 10-40 5-20 [17, 23, 24] Bermuda grass 25-48 13-35 6-19 [22, 23] Cotton seed hairs, flax 80-95 5-20 0 [18, 22]

49 16 10 [25]

13.8 not identified not identified [26]

50-70 12-20 6-10 [17]

Rice hulls 24-36 12-19 11-19 [22]

Primary municipal sludge 29.3 not identified not identified [26]

Sawdust 45.0 15.1 25.3 [22]

Newspaper 40-55 25-40 18-20 [1, 17] Used office paper 55.7 13.9 5.8 [1] Magazine 34.3 27.1 14.2 [1] Cardboard 49.6 15.9 14.9 [1] Paper sludge 33-61 14.2 8.4-15.4 [27, 28] Chemical pulps 60-80 20-30 2-10 [18]

**Table 1.** Overview and composition of lignocellulosic biomass and other lignocellulosic sources

Municipal solid waste – separated fibre

Thickened waste activated

Waste paper from chemical pulps

sludge

#### **2. Characterization of 2nd generation feedstock**

Plant biomass can be used as a sustainable source of organic carbon to create bioenergy, ei‐ ther 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 bio‐ mass 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 (Table1) 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.


**Table 1.** Overview and composition of lignocellulosic biomass and other lignocellulosic sources

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 β-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 forma‐ tion 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 dif‐ fers in crystalline structure and inter-connections between other biomass components. Mi‐ crofibrils 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 pre‐ dominantly 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 syn‐ thesized 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].

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 pre‐ dominantly 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). Be‐ sides lignocellulosic plant materials, other low-cost large volume feedstocks such as munici‐ pal 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, degrada‐ ble 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 toi‐ let 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 dis‐ posal 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 poorquality 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 land‐ fills or burned), its high carbohydrate content (Table 1) and its structure, which doesn´t re‐ quire 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 disso‐ lution 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 compo‐ nents 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, nu‐ trients and minerals) differs significantly with the type of wood and technological proce‐ dures, e.g. concentration of the main sugars in SWL (% of dry matter) ranges for xylose from

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There are several groups of lignocellulosic plant biomasses that can be exploited as a feed‐ stock 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. Agri‐ cultural 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 ag‐ ricultural 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 pre‐ dominantly 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). Be‐ sides lignocellulosic plant materials, other low-cost large volume feedstocks such as munici‐ pal 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, degrada‐ ble 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 toi‐ let 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 dis‐ posal 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 poorquality 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 land‐ fills or burned), its high carbohydrate content (Table 1) and its structure, which doesn´t re‐ quire 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 disso‐ lution 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 compo‐ nents 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, nu‐ trients and minerals) differs significantly with the type of wood and technological proce‐ dures, 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 con‐ centration 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 with‐ out careful pretreatment - stripping off free sulfur dioxide and simultaneous concentration, steaming, removing inhibitors, adding nutrients, and adjusting the pH [43].

terial 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

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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 sug‐ ars 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 toxici‐ ty (concentration of inhibitory compounds released by sugar and lignin decomposition) ana‐

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 proc‐ esses, 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 hydro‐ lysed 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 pro‐ duction based on the sulphite method [38] use sulfurous acid and its salts (Ca2+, Mg2+, Na+

) 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 in‐ side 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 high‐ er degree of saccharification of hard and softwoods had led to various modifications of proc‐ ess 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 bio‐

paper sludge doesn´t need any processing.

and NH4

+

lysed by HPLC or measured as the ability of test strains to grow.

mass with higher content of lignin, e.g. softwood, eucalyptus).

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.

#### **3. Biomass disruption in pretreatment process**

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 cellulo‐ lytic enzymes by modifying its structure; this happens in the pretreatment process. Basical‐ ly, 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) cellu‐ lose 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 fi‐ bres. 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-hydro‐ lysed due to its amorphous branched structure; the liquid fraction obtained after pretreat‐ ment 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 pre‐ dominantly 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 struc‐ ture 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 pre‐ treatment) [8], it is a necessary expense because enzymatic hydrolysis of non-pretreated ma‐ terial 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 sug‐ ars 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 toxici‐ ty (concentration of inhibitory compounds released by sugar and lignin decomposition) ana‐ lysed 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 proc‐ esses, 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 hydro‐ lysed 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 pro‐ duction 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 in‐ side 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 high‐ er degree of saccharification of hard and softwoods had led to various modifications of proc‐ ess 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 bio‐ mass 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.

**Pretreatment Condition Advantages Disadvantages Refe-**

times

Ozonolysis Room temperature, normal

hours

Combined acid and alkali pretreatment (formic acidaqueous ammonia, dilute sulphuric acidsodium hydroxide)

Combined acid and organic solvent (concentrated H3PO4 + aceton),

Ionic liquid (IL)pretreatment

Acid-catalyzed steam explosion pressure, reaction time -

Temperature <100 °C, cellulose recovered by addition of water, ethanol or

**Physicochemical pretreatment**

Steam explosion catalysed by addition of H2SO4 or SO2

pressure 0.7-4.8 MPa, reaction time 1-10 min followed by biomass

acetone

Steam explosion Temperature 160-240 °C,

explosion

recycle, continuous process, short residence

Lignin degradation, no inhibitors, ambient temperature

Cellulose digestion, fractionation of lignocellulose, most of non-cellulosic components removed,

high loading

structure disrupted, high yield of amorphous cellulose, lignin removed, reduced enzyme loading

Lignin extraction, low temperature, high biomass loading, high lignin solubility, cellulose dissolution, solvents recovered and reused, environmentally friendly

Extensive redistribution of lignin, high cellulose digestibility, cellulose swelling, limited use of

Decreased time and temperature compared to steam explosion

chemicals

Moderate temperatures Cellulose crystalline

**rence**

55

[45]

[29, 44, 45]

[29, 31, 45, 55]

[45]

Hemicellulose dissolved [21]

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Hemicellulose dissolved [45]

Cellulose recovered by addition of acetone,

Little lignin removed, incomplete destruction of biomass matrix, sugar decomposition (inhibitors), hemicellulose dissolved, high energy consumption

Inhibitors formation, hemicellulose dissolved, high

temperature

deionized water or alcohol, IL denaturates enzymes, IL must be washed before reused




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 hy‐ drolysis in dilute acid processes, higher temperatures are favoured [58] since at a moder‐ ate 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 de‐ creased 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 explo‐ sion 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 hydrox‐ ide 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. Martín 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 solubilisa‐ tion 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 en‐ richment with CO2 or SO2 proved promising results as for enzymatic hydrolysis of cellu‐ lose and the low content of inhibitors, especially 2-furalaldehyde and 5-hydroxymethyl-2-

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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, pump‐ ing 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 caus‐ es 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

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-ß-D-glucanase (EC 3.2.1.4, hydrolysis of (1→4) glucosidic linkages inside the chain), exo-1,4- ß-glucosidase (EC 3.2.1.74, hydrolysis of (1→4) linkage in (1→4)-β-D-glucans to re‐ move successive glucose units), ß-glucosidase (EC 3.2.1.21, hydrolysis of terminal non-re‐

monomers) in fermentation as shown in Figure 1.

furalaldehyde.

**Table 2.** Overview and main characteristics of methods leading to biomass pretreatment

Acid treatments lead mainly to hydrolysis of hemicelluloses (pentose and hexose frac‐ tions) 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 hy‐ drolysis in dilute acid processes, higher temperatures are favoured [58] since at a moder‐ ate 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 de‐ creased 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 explo‐ sion 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 hydrox‐ ide 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. Martín 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 solubilisa‐ tion 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 en‐ richment with CO2 or SO2 proved promising results as for enzymatic hydrolysis of cellu‐ lose 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, pump‐ ing 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 caus‐ es 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-ß-D-glucanase (EC 3.2.1.4, hydrolysis of (1→4) glucosidic linkages inside the chain), exo-1,4- ß-glucosidase (EC 3.2.1.74, hydrolysis of (1→4) linkage in (1→4)-β-D-glucans to re‐ move successive glucose units), ß-glucosidase (EC 3.2.1.21, hydrolysis of terminal non-re‐ ducing β-D-glucosyl residues with release of β-D-glucose) and β-1,4-glucan cellobiohydrolase (EC 3.2.1.91, hydrolysis of (1→4)-β-D-glucosidic linkages in cellulose and cellotetraose releasing cellobiose from non-reducing ends of the chains) activities working in synergy.

to avoid inhibition of cellulolytic enzymes [22, 45]. Unfortunately, improved enzyme cock‐ tails are not generally applicable, e.g. an enzyme complex enriched with β-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 af‐ fected 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 anaero‐ bic 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-pro‐ ductive enzyme binding and can also facilitate enzyme recycling. Addition of expansins (plant proteins), expansin-like proteins or swollenin (fungal protein) promotes enhanced en‐ zymatic 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 ap‐

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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 sub‐

Toxic products can generally be divided into three main groups – aliphatic acids, furan de‐ rivatives 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

proaches to reduce pretreatment costs [45].

**4. Toxic compounds released in pretreatment process**

strate fermentability.

**Figure 1.** Simplified diagram of production of liquid biofuels from lignocellulose biomass

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 re‐ duced, 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 pro‐ duction 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 ac‐ tivity 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 cel‐ lulose surface, while β-xylosidase can decompose xylobiose and polymerized xylooligomers to avoid inhibition of cellulolytic enzymes [22, 45]. Unfortunately, improved enzyme cock‐ tails are not generally applicable, e.g. an enzyme complex enriched with β-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 af‐ fected 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 anaero‐ bic 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-pro‐ ductive enzyme binding and can also facilitate enzyme recycling. Addition of expansins (plant proteins), expansin-like proteins or swollenin (fungal protein) promotes enhanced en‐ zymatic 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 ap‐ proaches 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 sub‐ strate fermentability.

#### **4. Toxic compounds released in pretreatment process**

Toxic products can generally be divided into three main groups – aliphatic acids, furan de‐ rivatives 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 bio‐ mass [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 com‐ pounds (their effect is intensified when present in combination) and the strain used for etha‐ nol 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 in‐ hibitors with high molecular weights affect expression and activity of sugar and ion trans‐ porters. 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 in‐ fluenced [75] due to its ability to detoxify the broth by reduction of furfural to furfuryl alco‐ hol, which is less toxic.

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, ρ-coumaric, ferulic and vanillic acids, together with vanillin, were effectively removed from a model solution of phenolic compounds by treatment with 0.01µM peroxidase (E.C. 1.11.7), re‐ sulting 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 im‐ provement of butanol yield achieved with *C. beijerinckii* BA101 [85]. Another popular ap‐ proach 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 meth‐ od 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

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Fermentation of lignocellulose hydrolysates is more complicated compared to fermentation of 1st generation feedstock (sugar cane juice, molasses, grains) for several reasons: a) pentose sugars (predominantly xylose) are present along with hexoses (mainly glucose, mannose, galactose) in the hydrolysate, b) toxic compounds released during pretreatment can influ‐ ence metabolic activity of the fermentation strain, c) low concentrations of fermentable sug‐ ars hamper the attainment of a high ethanol concentration. Because lignocellulose hydrolysates are poor in some nutrients (phosphorus, trace elements, and vitamins) they are usually supplemented, e.g. by addition of corn steep or yeast extract before being used as a substrate for fermentation. For an efficient process it is necessary to identify a strain that uti‐ lizes both pentose and hexose sugars, produces ethanol with a high yield and productivity and is tolerant to both inhibitors and ethanol. One of the main challenges is to simultaneous‐ ly co-ferment pentose and hexose sugars, but neither yeast *S. cerevisiae* nor the bacterium *Z. mobilis*, which are usually used for ethanol production, contain genes for expression of xy‐ lose reductase and xylitol dehydrogenase [89]. In order to enhance process effectiveness, cofermentation or sequential fermentation of hexoses and pentoses has been examined by combining good ethanol producers with strains naturally utilizing pentoses e.g. *Pichia stipi‐ tis, Candida shehatae, Pachysolen tannophillus, Klebsiella oxytoca.* However, xylose utilization is the rate limiting step due to catabolite repression by hexoses and the low availability of oxy‐ gen, and inhibition of pentose-utilizing strains by ethanol [90, 91]. Moreover, the yield of ethanol by co-fermentation is usually lower than with separate processes, e.g. yields of 0.5 g ethanol per g glucose (98% of theoretical) and 0.15 g/g xylose (29% theoretical) were ach‐ ieved by separate cultivation of *Z. mobilis* and *P. tannophillus* respectively, but in optimized

to treat hydrolysates with activated carbon [88].

**5.1. Ethanol fermentation**

**5. Fermentation of lignocellulosic substrates**

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 ρ-coumaric and ferulic acids present at a concentration of 0.3 g/l [76-78]. Solvent produc‐ tivity 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 hy‐ droxymethyl 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 calci‐ um 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, ρ-coumaric, ferulic and vanillic acids, together with vanillin, were effectively removed from a model solution of phenolic compounds by treatment with 0.01µM peroxidase (E.C. 1.11.7), re‐ sulting 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 im‐ provement of butanol yield achieved with *C. beijerinckii* BA101 [85]. Another popular ap‐ proach 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 meth‐ od 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].

#### **5. Fermentation of lignocellulosic substrates**

#### **5.1. Ethanol fermentation**

Fermentation of lignocellulose hydrolysates is more complicated compared to fermentation of 1st generation feedstock (sugar cane juice, molasses, grains) for several reasons: a) pentose sugars (predominantly xylose) are present along with hexoses (mainly glucose, mannose, galactose) in the hydrolysate, b) toxic compounds released during pretreatment can influ‐ ence metabolic activity of the fermentation strain, c) low concentrations of fermentable sug‐ ars hamper the attainment of a high ethanol concentration. Because lignocellulose hydrolysates are poor in some nutrients (phosphorus, trace elements, and vitamins) they are usually supplemented, e.g. by addition of corn steep or yeast extract before being used as a substrate for fermentation. For an efficient process it is necessary to identify a strain that uti‐ lizes both pentose and hexose sugars, produces ethanol with a high yield and productivity and is tolerant to both inhibitors and ethanol. One of the main challenges is to simultaneous‐ ly co-ferment pentose and hexose sugars, but neither yeast *S. cerevisiae* nor the bacterium *Z. mobilis*, which are usually used for ethanol production, contain genes for expression of xy‐ lose reductase and xylitol dehydrogenase [89]. In order to enhance process effectiveness, cofermentation or sequential fermentation of hexoses and pentoses has been examined by combining good ethanol producers with strains naturally utilizing pentoses e.g. *Pichia stipi‐ tis, Candida shehatae, Pachysolen tannophillus, Klebsiella oxytoca.* However, xylose utilization is the rate limiting step due to catabolite repression by hexoses and the low availability of oxy‐ gen, and inhibition of pentose-utilizing strains by ethanol [90, 91]. Moreover, the yield of ethanol by co-fermentation is usually lower than with separate processes, e.g. yields of 0.5 g ethanol per g glucose (98% of theoretical) and 0.15 g/g xylose (29% theoretical) were ach‐ ieved by separate cultivation of *Z. mobilis* and *P. tannophillus* respectively, but in optimized

co-fermentation, the yield was just 0.33 g ethanol/g sugar. The same yield was obtained in a 5-reactor process combining *P. stipitis* and *S. cerevisiae* [92], but it was enhanced to 0.49 g/g sugars (96% theoretical) by cultivation of an adapted co-culture of *S. cerevisiae*, *P. tannophilis* and recombinant *E. coli* in dilute-acid softwood hydrolysate [93]. In a subsequent process employing *P. stipilis* and *S. cerevisiae*, which was inactivated before *Pichia* inoculation to avoid oxygen competition, 75% of theoretical ethanol yield was achieved [94]. A different approach is represented by the use of a recombinant strain prepared either by cloning genes encoding xylose utilization into good ethanol producers or to construct synthetic pathways for ethanol production in pentose-utilizing hosts. Wild type yeasts can be genetically modi‐ fied to utilize xylose by introducing fungal genes encoding xylose reductase and xylitol de‐ hydrogenase or bacterial/fungal genes for xylose isomerase [95]. Yeast *S. cerevisiae* was transformed with the *xylA* gene from *Thermus thermophiles* and *Piromyces sp.* to produce xy‐ lose isomerase, but unfortunately, this enzyme was inhibited by xylitol, favouring instead, its formation. Recently a recombinant strain of *S. cerevisiae* expressing a heterologous *xylA* gene produced 0.42 g/g of ethanol from xylose [96]. A strategy using *xyl1* and *xyl2* genes from *P. stipitis* introduced into *S. cerevisiae* produced transformants that exclusively con‐ sumed xylose, but produced significant amounts of xylitol [97]. On the other hand, with re‐ combinant *Z. mobilis*, which carried *E. coli* genes encoding for xylose isomerase, xylulokinase, transketolase and transaldolase, 86% ethanol yield from xylose was achieved. Another strain of *Z. mobilis*, expressing genes *araABD* from *E. coli,* encoding L-arabinose iso‐ merase, L-ribulokinase, L-ribulose-5-P-4 epimerase together with genes for transketolase and transaldolase, was able to grow on arabinose with 98% ethanol yield. *E. coli*, which nat‐ urally utilizes a wide range of substrates including pentoses, was transformed by genes en‐ coding pyruvate decarboxylase and alcohol dehydrodenase, resulting in enhanced ethanol production [96]. Adaptation of recombinant strains to inhibitors can further increase the yield of ethanol, e.g. the ethanol yield achieved with a genetically engineered strain of *S. cer‐ evisiae* grown on bagasse hydrolysate was increased from 0.18 g/g to 0.38 g/g after adapta‐ tion [98]. Recombinant strains that not only consume pentoses but also hydrolyse hemicelluloses by co-expressing endoxylanase, β-xylosidase and β-glucosidase activities has recently been constructed [95] and yields of 0.41 g/g of ethanol were obtained from total sugars in a rice straw hydrolysate.

egies, e.g. maximizing dry matter by removing most hemicellulose and lignin, utilizing alternative bioreactors with novel mixing modes (e.g. peg mixer, shaking, gravitational tum‐ bling, hand stirring) or gradual dosing of substrate into the bioreactor (fed-batch), which en‐ ables the use of more substrate and thus increases the yield of ethanol above values achievable in batch mode. Moreover, the actual concentration of toxic substrates is reduced and yield and/or productivity is enhanced by controlled dosing of substrate and prolonged cultivation time, thus shortening unprofitable periods between batches [45, 89]. Feed rates should reflect the type of hydrolysate and strain. Continuous cultures usually using immo‐ bilized cells (to prevent their wash out from the bioreactor at high dilution rates) is another

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Integration strategies, which replace classical separate hydrolysis and fermentation process‐ es (SHF) by combining several process steps in one vessel represents another approach for lignocellulosic ethanol production. Simultaneous saccharification and fermentation (SSF), which combines enzymatic hydrolysis and fermentation in one step, permits an increased rate of cellulose hydrolysis by elimination of product inhibition (the released glucose is con‐ sumed by the microbial strain), an increased rate of sugar consumption, reduced contamina‐ tion due to the presence of ethanol and a reduced number of reactors. However, SSF is constrained by different temperature optima for each process (the cellulase optimum is usu‐ ally 40-50 °C, whereas the fermentation temperature usually cannot exceed 35 °C for most ethanol producers) and carbon source limitation in the early stages of the process. Several modifications of SSF to ease the problems and increase productivity have been published. These include the use of thermotolerant ethanol producers [100, 101], application of a presaccharification step [102] or the use of recombinant strains consuming both hexose and pentose sugars (a simultaneous saccharification and co-fermentation process (SSCF)) [103] in batch or fed-batch mode [104]. Consolidated bioprocessing (CBP), which combines cellulase production, cellulose hydrolysis and fermentation into a single step have been investigated as a way of reducing the cost of cellulolytic enzymes, increasing volumetric productivity and reducing capital investment [105]. Some biofuel companies (e.g. Mascoma and Qteros) have been founded based on this concept [105]. CBP microorganisms should combine high cellulase production and secretory capability, the ability to utilize a broad range of sugars, tolerance to high concentrations of salts, solvents and inhibitors, high ethanol productivity and yield, have a known genomic DNA sequence and developed recombinant technologies and ideally be usable as feed protein after fermentation [105]. There is a lack of native organ‐ isms that combine the ability to produce cellulolytic enzymes and be homoethanolic with high titres and yields. Although some thermophilic anaerobic bacteria e.g. *Clostridium ther‐ mocellum,* are high cellulase producers and utilize both pentose and hexose sugars, they have a low tolerance to ethanol ~30 g/l [106] and an insufficient yield ~0.2 g/g [107]. There‐ fore recombinant strains have been prepared by engineering cellulolytic microorganisms (e.g. *C. thermocellum, C. phytofermentans, C. cellulolyticum, T. reesei or F.oxysporum*) to produce ethanol. Knockout mutants of *Thermoanaerobacterium saccharolyticum* that lack lactic and ace‐ tic acid production exhibited an ethanol yield from xylose of 0.46 g/g [108], while recombi‐ nant *Geobacillus thermoglucosidasius* produced 0.42-0.47 g/g of ethanol from hexoses [65]. Another attempt, to create a recombinant cellulose-utilizing microorganism using non-cellu‐

strategy to increase process productivity [89].

In addition to the wide range of sugars, their low concentration in hydrolysates is problem‐ atic. Since ethanol recovery by distillation is only economically viable on the industrial scale for yields greater than 4% (w/w), which for most hydrolysates requires a dry mass concen‐ tration greater than 20% [45], the use of high substrate loading is needed. Effect of substrate concentration (unbleached hardwood pulp and organosolve pretreated poplar) on glucose concentration resulting from the enzyme hydrolysis was studied in [52] and [99]. In labora‐ tory scale after 48 h of enzymatic hydrolysis 158 g/l glucose in the hydrolyzate was reached, ethanol concentration after fermentation ranged between 50.4 and 63.1 g/l. The general problem for this kind of conversions is that high load of the pulp or pretreated lignocelulo‐ sic material gives rise to high viscosity and thus also to mixing and transport problems. These extremely high yields of glucose can be attributed to a very efficient peg mixer. Prob‐ lems connected with use of such high viscosity slurries can be overcome by various strat‐ egies, e.g. maximizing dry matter by removing most hemicellulose and lignin, utilizing alternative bioreactors with novel mixing modes (e.g. peg mixer, shaking, gravitational tum‐ bling, hand stirring) or gradual dosing of substrate into the bioreactor (fed-batch), which en‐ ables the use of more substrate and thus increases the yield of ethanol above values achievable in batch mode. Moreover, the actual concentration of toxic substrates is reduced and yield and/or productivity is enhanced by controlled dosing of substrate and prolonged cultivation time, thus shortening unprofitable periods between batches [45, 89]. Feed rates should reflect the type of hydrolysate and strain. Continuous cultures usually using immo‐ bilized cells (to prevent their wash out from the bioreactor at high dilution rates) is another strategy to increase process productivity [89].

Integration strategies, which replace classical separate hydrolysis and fermentation process‐ es (SHF) by combining several process steps in one vessel represents another approach for lignocellulosic ethanol production. Simultaneous saccharification and fermentation (SSF), which combines enzymatic hydrolysis and fermentation in one step, permits an increased rate of cellulose hydrolysis by elimination of product inhibition (the released glucose is con‐ sumed by the microbial strain), an increased rate of sugar consumption, reduced contamina‐ tion due to the presence of ethanol and a reduced number of reactors. However, SSF is constrained by different temperature optima for each process (the cellulase optimum is usu‐ ally 40-50 °C, whereas the fermentation temperature usually cannot exceed 35 °C for most ethanol producers) and carbon source limitation in the early stages of the process. Several modifications of SSF to ease the problems and increase productivity have been published. These include the use of thermotolerant ethanol producers [100, 101], application of a presaccharification step [102] or the use of recombinant strains consuming both hexose and pentose sugars (a simultaneous saccharification and co-fermentation process (SSCF)) [103] in batch or fed-batch mode [104]. Consolidated bioprocessing (CBP), which combines cellulase production, cellulose hydrolysis and fermentation into a single step have been investigated as a way of reducing the cost of cellulolytic enzymes, increasing volumetric productivity and reducing capital investment [105]. Some biofuel companies (e.g. Mascoma and Qteros) have been founded based on this concept [105]. CBP microorganisms should combine high cellulase production and secretory capability, the ability to utilize a broad range of sugars, tolerance to high concentrations of salts, solvents and inhibitors, high ethanol productivity and yield, have a known genomic DNA sequence and developed recombinant technologies and ideally be usable as feed protein after fermentation [105]. There is a lack of native organ‐ isms that combine the ability to produce cellulolytic enzymes and be homoethanolic with high titres and yields. Although some thermophilic anaerobic bacteria e.g. *Clostridium ther‐ mocellum,* are high cellulase producers and utilize both pentose and hexose sugars, they have a low tolerance to ethanol ~30 g/l [106] and an insufficient yield ~0.2 g/g [107]. There‐ fore recombinant strains have been prepared by engineering cellulolytic microorganisms (e.g. *C. thermocellum, C. phytofermentans, C. cellulolyticum, T. reesei or F.oxysporum*) to produce ethanol. Knockout mutants of *Thermoanaerobacterium saccharolyticum* that lack lactic and ace‐ tic acid production exhibited an ethanol yield from xylose of 0.46 g/g [108], while recombi‐ nant *Geobacillus thermoglucosidasius* produced 0.42-0.47 g/g of ethanol from hexoses [65]. Another attempt, to create a recombinant cellulose-utilizing microorganism using non-cellu‐ lolytic strains with high ethanol production have not been very successful; although some recombinant ethanologenic strains secreting some active cellulases have been prepared [106, 109, 110], their requirement for a nutrient rich medium and often sensitivity to end-product inhibition hamper their use [105].

**Substrate Pretreatment Microbial strain ABE concentration**

Wheat bran Diluted sulphuric acid *C. beijerinckii* ATCC 55025 12/32/0.16 [140]

*C. saccharoperbutylacetonicum*

*C. saccharoperbutylacetonicum*

*C. saccharobutylicum* 260 *C. butylicum 592 C. butylicum 592*

All fermentations were run in SHF mode i.e. sugar release and fermentation were separate processes.

**Table 3.** Selection of batch ABE fermentations in laboratory scale using lignocellulosic hydrolysates as a substrate

Until now, lignocellulosic substrates must be prehydrolysed for the ABE process. In the case of fermentation of lignocellulosic hydrolysate, usually containing low concentrations of fer‐ mentable sugars, one of the main bottlenecks in the ABE process, the low final titre of buta‐

Heat + enzyme *C. acetobutylicum* JB200 34/39/0.63 [130]

Diluted acetic acid *C. acetobutylicum* ABE 0801 19/32/0.10 [141]

ATCC 27022

ATCC 27022

Wheat straw Diluted sulphuric acid+ enzyme

Corn fiber Diluted sulphuric acid+

enzyme

enzyme

enzyme

+enzyme

enzyme

enzyme

Alkali + (NH4)2SO4 precipitation + activated carbon treatment +

Steam explosion, lyophilization + enzyme +4 fold concentration of released sugars

Diluted acid + overliming

hot water + overliming+

AFEX +overliming+

AFEX stands for ammonium fiber expansion process.

Corn cobs Steam explosion + enzyme

Rice straw Alkali + (NH4)2SO4

Sugar cane bagasse

Cassava bagasse

Domestic organic waste

Dried distiller`s grain and solubles

Sweet sorghum

stem

XAD-4 resin treatment +

precipitation + activated carbon treatment +

**(g/l) /yield (%)**

*C. beijerinckii* P260 13/25/0.14 [139]

*C. beijerinckii* BA101 8/32/0.11 [127]

*C. acetobutylicum* 21/31/0.45 [131]

*C. acetobutylicum* DSM 792 9/26/0.08 [132]

12/35/0.20 13/32/0.20 12/32/0.20

**/productivity (g/l/h)**

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13/28/0.15 [88]

14/30/0.17 [88]

[78]

**Reference**

65

#### **5.2. ABE (acetone-butanol-ethanol) fermentation**

Different, so-called solventogenic species of the genus *Clostridium*, like *Clostridium acetobu‐ tylicum*, *Clostridium beijerinckii* or *Clostridium saccharoperbutylacetonicum,* can be used for 1 butanol production by ABE fermentation. The fermentation usually proceeds in two steps; at first butyric and acetic acids, along with hydrogen and carbon dioxide, are formed and then metabolic switching leads to the formation of solvents (mainly 1-buta‐ nol and acetone) and the cessation/slowdown of acid and gas production (for recent re‐ views see [111-114]). Industrial fermentative ABE (butanol) production, which has quite a long and impressive history connected with both World Wars, is nowadays carried out only in China and Brazil (estimated annual production of 100 000 t and 8 000 t from corn starch and sugar cane juice, respectively) [10]. However, many corporations such as BP, DuPont, Gevo, Green Biologics, Cobalt Technologies and others have declared their inter‐ est in this field. A unique example of the use of lignocellulosic hydrolysate on an indus‐ trial scale is the former Dukshukino plant (operated in the Soviet Union up to 1980s) producing acetone and butanol by fermentation. The plant was based on current "very modern" biorefinery concepts which assumed the conversion of complex feedstocks (hy‐ drolysates of agricultural waste + molasses or corn) into many valuable products i.e. in addition to solvents (acetone, butanol and ethanol), it was possible to produce liquid CO2, dry ice, H2, fodder yeast, vitamin B12 and biogas [115].

The most interesting approach to fermentation of any lignocellulosic substrate is probably consolidated bioprocessing (CBP) i.e. a method in which a single microorganism is used for both substrate decomposition and fermentation to produce the required metabolites. Although some clostridial species such as *Clostridium thermocellum* can utilise cellulosic substrates and produce ethanol [116, 117], the ABE fermentation pattern unfortunately cannot be produced using clostridia. However, *C.acetobutylicum* ATCC 824 possesses genes for various cellullases and a complete cellulosome [118-120]. But even if production of some cellulases by *C. acetobutylicum* ATCC 824 was induced by xylose or lichenan [118], cellulose utilization was not achieved, possibly because of insufficient or deficient synthesis of an unknown specific chaperone that could be responsible for correct secre‐ tion of cellulases [119]. Nevertheless as solventogenic *Clostridium* species are soil bacteria that differ significantly in fermentative abilities and genome sizes, it is not excluded that in the future, some solventogenic species with cellulolytic activity will be isolated from an appropriate environment. Recently, a new strain of *Clostridium saccharobutylicum* with hemicellulolytic activity and ABE fermentation pattern was found amongst 50 soil-borne, anaerobic, sporulating isolates [121].


All fermentations were run in SHF mode i.e. sugar release and fermentation were separate processes.

AFEX stands for ammonium fiber expansion process.

**Table 3.** Selection of batch ABE fermentations in laboratory scale using lignocellulosic hydrolysates as a substrate

Until now, lignocellulosic substrates must be prehydrolysed for the ABE process. In the case of fermentation of lignocellulosic hydrolysate, usually containing low concentrations of fer‐ mentable sugars, one of the main bottlenecks in the ABE process, the low final titre of buta‐ nol (caused by severe butanol toxicity towards bacterial cells), is of minor importance. In fact hydrolysates are very good substrates for clostridia that express extensive fermentative abilities [122, 123]and can utilise not only cellulose-derived glucose but also hemicellulose monomers (xylose, arabinose, galactose, mannose). Co-fermentation of various sugar mix‐ tures was described for *Clostridium beijerinckii* SA-1 (ATCC 35702) [124], *Clostridium acetobu‐ tylicum* DSM 792 [125], *C. acetobutylicum* ATCC 824 [126] and *C.beijerinckii* P260 [127] however, at the same time, catabolic repression of xylose utilization in the presence of glu‐ cose was demonstrated in *C. acetobutylicum* ATCC 824 [128, 129].

the opening of pilot and demonstration plants around the world (production capacity in millions of gallons for the year 2012 given in brackets) e.g. the POET demonstration plant in Iowa (0.02 from corn stover and cobs), Abengoa in Kansas (0.01 from corn stover), Blue Sugarsin Wyoming (1.3 from stover and cobs), Chempolis in Finland (3.7 from paper waste), Fiberight in Iowa (6.0 MSW), Iogen in Canada (0.48 from stover), Praj MATRIX in India (0.01 from cellulose), UPM-Kymemene/Mesto in Finland (0.68 from mixed cellulose) and in spite of several proclamations, none of them is operating at the industrial scale [9]. To make this possible, further reductions in processing costs will be necessary to achieve a product that is competitive with 1st generation bioethanol. Further process integration is required, including decreased energy demand during pretreatment, increased sugar concentration, higher enzyme activity and strain recycling. By-products, e.g. lignin separated after pretreatment procedure can be used to generate energy for ethanol plant operations (lignin has higher caloric value (25.4 MJ/kg) then the biomass it‐ self [8]) or used as a dispersant and binder in concrete admixtures, as an alternative to phenolic and epoxy resins, or as the principal component in thermoplastic blends, poly‐ urethane foams or surfactants [143]. A combination of 1st and 2nd generation feedstocks (e.g. corn cobs together with stover ) can eliminate bottlenecks and lead to product com‐ petitiveness. Higher bioethanol production costs can also be compensated for by political and economic instruments such as tax incentives (e.g. tax exemption on biofuels and higher excise taxes for fossil fuels) and legislation (mandatory blends) to enable ready ac‐ cess of 2nd generation biofuels to the market [30]. Butanol, as a second generation biofuel, might be produced via fermentation and used as an excellent fuel extender in addition to ethanol if the technological bottleneck of a low final concentration, yield and productivity could be overcome, and the assumption that suitable cheap waste pretreatments were

Production of 2nd Generation of Liquid Biofuels

http://dx.doi.org/10.5772/53492

67

The. review was performed thanks to financial support of the projects Kontakt ME10146 of Ministry of Education, Youth and Sport of Czech Republic and BIORAF No.TE01020080 of

Department of Biotechnology, Institute of Chemical Technology Prague, Prague, Czech Re‐

possible.

**Acknowledgement**

**Author details**

public

the Technological Agency of the Czech Republic.

Leona Paulová, Petra Patáková, Mojmír Rychtera and Karel Melzoch

\*Address all correspondence to: Leona.paulova@vscht.cz

An overview of fermentation parameters achieved in batch ABE fermentations of different hydrolysates is presented in Table 3. The most promising results were obtained by Lu et al. [130] using cassava bagasse and a mutant strain, *C.acetobutylicum* JB200; the results of Marchal et al. [131] were unique at the scale used (48 m3 ) as shown in Table 3. A frequent problem of lignocellulosic hydrolysates is a low final concentration of fermentable sugars caused by low density of the original substrate. This can be overcome by evaporation of the hydrolysate [132](see Table 3) or by addition of glucose and/or other carbohydrates present in the hydrolysate (this is only possible in laboratory scale experiments) [85,133-135]. In the case of glucose supplemented corn stover and switchgrass hydrolysates, final ABE concen‐ trations of 26 and 15 g/l were achieved [135]. With *C.beijerinckii* P260, use of diluted and Ca(OH)2 treated barley straw hydrolysate supplemented with glucose resulted in a solvent concentration of 27 g/l, a yield of 43% and productivity of 0.39 g/l/h [133]. In addition to ma‐ terials presented in Table 3, other substrates like diluted sulfite spent liquor supplemented with glucose [134], palm empty fruit bunches [136, 137] or hardwood [138] were used in the ABE process but in these cases, additional optimizations were necessary.

In addition to a batch fermentation arrangement, semi-continuous fermentation of enzymat‐ ically hydrolyzed SO2 pretreated pine wood using *C.acetobutylicum* P262 resulted in 18 g/l of solvents, a yield of 36% and solvent productivity of 0.73 g/l/h [142]. Further, fed-batch fer‐ mentation of wheat straw hydrolysate supplemented with varying concentrations of hydro‐ lysate sugars (glucose, xylose, arabinose and mannose) using *C.beijerinckii* P260 yielded a solvent productivity of 0.36 g/l/h if gas stripping was used [127]. In the cases shown in Table 3, enzyme hydrolysis preceeded fermentation, however simultaneous saccharification and fermentation (SSF) was also tested. In SSF of acid pre-hydrolyzed wheat straw using *C.beijer‐ inckii* P262 and solvent removal by gas stripping, 21 g/l of ABE was produced with a pro‐ ductivity of 0.31 g/l/h [127] Nevertheless, the solvent yield from hardwood using SSF was rather low, at 15% [138].

#### **6. Conclusion**

Intensive research over the last decades on lignocellulose-derived ethanol have focused mainly on intensification of biomass pretreatment, production of cellulolytic enzymes, and strain and process improvements, and have eliminated some of the main technologi‐ cal bottlenecks. Although a number of projects on 2nd generation bioethanol ended with the opening of pilot and demonstration plants around the world (production capacity in millions of gallons for the year 2012 given in brackets) e.g. the POET demonstration plant in Iowa (0.02 from corn stover and cobs), Abengoa in Kansas (0.01 from corn stover), Blue Sugarsin Wyoming (1.3 from stover and cobs), Chempolis in Finland (3.7 from paper waste), Fiberight in Iowa (6.0 MSW), Iogen in Canada (0.48 from stover), Praj MATRIX in India (0.01 from cellulose), UPM-Kymemene/Mesto in Finland (0.68 from mixed cellulose) and in spite of several proclamations, none of them is operating at the industrial scale [9]. To make this possible, further reductions in processing costs will be necessary to achieve a product that is competitive with 1st generation bioethanol. Further process integration is required, including decreased energy demand during pretreatment, increased sugar concentration, higher enzyme activity and strain recycling. By-products, e.g. lignin separated after pretreatment procedure can be used to generate energy for ethanol plant operations (lignin has higher caloric value (25.4 MJ/kg) then the biomass it‐ self [8]) or used as a dispersant and binder in concrete admixtures, as an alternative to phenolic and epoxy resins, or as the principal component in thermoplastic blends, poly‐ urethane foams or surfactants [143]. A combination of 1st and 2nd generation feedstocks (e.g. corn cobs together with stover ) can eliminate bottlenecks and lead to product com‐ petitiveness. Higher bioethanol production costs can also be compensated for by political and economic instruments such as tax incentives (e.g. tax exemption on biofuels and higher excise taxes for fossil fuels) and legislation (mandatory blends) to enable ready ac‐ cess of 2nd generation biofuels to the market [30]. Butanol, as a second generation biofuel, might be produced via fermentation and used as an excellent fuel extender in addition to ethanol if the technological bottleneck of a low final concentration, yield and productivity could be overcome, and the assumption that suitable cheap waste pretreatments were possible.

#### **Acknowledgement**

The. review was performed thanks to financial support of the projects Kontakt ME10146 of Ministry of Education, Youth and Sport of Czech Republic and BIORAF No.TE01020080 of the Technological Agency of the Czech Republic.

#### **Author details**

Leona Paulová, Petra Patáková, Mojmír Rychtera and Karel Melzoch

\*Address all correspondence to: Leona.paulova@vscht.cz

Department of Biotechnology, Institute of Chemical Technology Prague, Prague, Czech Re‐ public

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[106] Lee R. Lynd W. H. V. Z., John E. Mcbride, Mark Laser. Consolidated bioprocessing of cellulosic biomass: an update. Current Opinion in Biotechnology 2005(16), 577-583.

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**Chapter 3**

**Biofuels Ethanol and Methanol in OTTO Engines**

Today, humanity faces many environmental problems, one of which is atmospheric pollution that leads to greenhouse effect, ozone formation and to many health problems to human beings. Also, many countries around the world face the problem of energy shortage. At the same time we must not forget the need for clean air, clean fuel and biodegradable, renewable materials. Hazardous pollutants that lead to atmospheric pollution have many sources and automobile's exhaust emission is one of these. Petroleum-based products that have been used as fuels produce dangerous gas emissions. In order to decrease environmental impacts, scientists and many governments turned their attention to renewable fuels as alternatives to conventional fossil fuels and as oxygenates [1]. The beginning of the 21st century finds humans more familiar with the concept of sustainable development. We must prevent the degradation of our environment focusing in more friendly technologies. This need lead scientists to the use of other energy sources that can be used with the same efficiency but won't have damaging effect to the environment. The increased vehicle number that usually uses petroleum-based fuels results to dangerous emissions production such as carbon monoxide (CO), carbon dioxide (CO2), hydrocarbons (HC), nitrogen oxides (NOx) and others. These emissions besides the fact that lead to environmental degradation they also constitute a threat for human health. People's concern about the risks associated with hazardous pollutants results to an increased demand for renewable fuels as alternatives to fossil fuels [1,2]. Ethanol and methanol are alcohols that can be used as fuels instead of gasoline in automobile engines. For better understanding of the use of these two alcohols we must examine them separately. Fuel ethanol is an alternative fuel that is produced from biologically renewable resources that it can also be used as an octane enhancer and as oxygenate. Ethanol (ethyl alcohol, grain alcohol, ETOH) is a clear, colorless liquid alcohol with characteristic odor and as alcohol is a group of chemical compounds whose molecules contain a hydroxyl group, -OH, bonded to a carbon atom. Is produced with the process of fermentation of grains such as wheat, barley, corn, wood, or

> © 2013 Arapatsakos; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 Arapatsakos; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

distribution, and reproduction in any medium, provided the original work is properly cited.

Charalampos Arapatsakos

http://dx.doi.org/10.5772/52772

**1. Introduction**

Additional information is available at the end of the chapter


### **Biofuels Ethanol and Methanol in OTTO Engines**

Charalampos Arapatsakos

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/52772

#### **1. Introduction**

Today, humanity faces many environmental problems, one of which is atmospheric pollution that leads to greenhouse effect, ozone formation and to many health problems to human beings. Also, many countries around the world face the problem of energy shortage. At the same time we must not forget the need for clean air, clean fuel and biodegradable, renewable materials. Hazardous pollutants that lead to atmospheric pollution have many sources and automobile's exhaust emission is one of these. Petroleum-based products that have been used as fuels produce dangerous gas emissions. In order to decrease environmental impacts, scientists and many governments turned their attention to renewable fuels as alternatives to conventional fossil fuels and as oxygenates [1]. The beginning of the 21st century finds humans more familiar with the concept of sustainable development. We must prevent the degradation of our environment focusing in more friendly technologies. This need lead scientists to the use of other energy sources that can be used with the same efficiency but won't have damaging effect to the environment. The increased vehicle number that usually uses petroleum-based fuels results to dangerous emissions production such as carbon monoxide (CO), carbon dioxide (CO2), hydrocarbons (HC), nitrogen oxides (NOx) and others. These emissions besides the fact that lead to environmental degradation they also constitute a threat for human health. People's concern about the risks associated with hazardous pollutants results to an increased demand for renewable fuels as alternatives to fossil fuels [1,2]. Ethanol and methanol are alcohols that can be used as fuels instead of gasoline in automobile engines. For better understanding of the use of these two alcohols we must examine them separately. Fuel ethanol is an alternative fuel that is produced from biologically renewable resources that it can also be used as an octane enhancer and as oxygenate. Ethanol (ethyl alcohol, grain alcohol, ETOH) is a clear, colorless liquid alcohol with characteristic odor and as alcohol is a group of chemical compounds whose molecules contain a hydroxyl group, -OH, bonded to a carbon atom. Is produced with the process of fermentation of grains such as wheat, barley, corn, wood, or

© 2013 Arapatsakos; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 Arapatsakos; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

sugar cane. In the United States ethanol is made by the fermentation of corn [1-3]. By the reaction of fermentation simple sugars change into ethanol and carbon dioxide with the presence of zymase, an enzyme from yeast. Ethanol can also be made from cellulose that is obtained from agricultural residue and waste paper [1]. It is a high-octane fuel with high oxygen content (35% oxygen by weight) and when blended properly in gasoline produces a cleaner and more complete combustion. Ethanol is used as an automotive fuel either by itself or in blends with gasoline, such as mixtures of 10% ethanol and 90%gasoline, or 85% ethanol and 15% gasoline [3-6]. Many countries around the world use ethanol as fuel. For example, in Brazil ethanol is produced using as raw material sugarcane and many vehicles use ethanol as fuel. Also in Canada and in Sweden ethanol is highly promoted as fuel because of the many environmental benefits that ethanol has. When gasoline is used as fuel hydrocarbons (HC) escape to the atmosphere. Many hydrocarbons are toxic and some, such as benzene, cane cause cancer to humans. If ethanol is used as fuel hydrocarbons are not being produced because ethanol is an alcohol that does not produce HC when is burned. The reaction of hydrocarbons and nitrogen oxides that are produced from the gasoline burning, in the presence of sunlight leads to the formation of photochemical smog. The use of ethanol as fuel can contribute to the decrease of photochemical smog since it does not produces hydrocarbons [5-8]. Vehicles that burn petroleum fuels produce carbon monoxide (CO) because these fuels do not contain oxygen in their molecular structure. Carbon monoxide is a toxic gas that is formed by incom‐ plete combustion. When ethanol, which contains oxygen, is mixed with gasoline the combus‐ tion of the engine is more complete and the result is CO reduction [9-11].

4 2 2 2 2CH + 3H O CO + CO +7H - Synthesis ® gas (1)

Biofuels Ethanol and Methanol in OTTO Engines

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81

CO + CO + 7H 2CH OH +2H + H O 2 2 3 22 ® (2)

The main advantage of methanol as fuel is that is being produced from resources that can be found globally, while a large percentage of petroleum is located in Middle East. Furthermore, the materials needed for methanol production such as natural gas or biomass are renewable. This means that methanol can also be cheaper and more economically attractive than gasoline. When fossil fuels are used in automobiles produce exhaust emissions of hydrocarbons, carbon dioxide and other gases that contribute to the greenhouse effect. Methanol can give lower HC and CO emissions and besides that the vehicles that use methanol emit minimum particulate matter compared to gasoline, which usually has damaging effect to humans. In addition, methanol has high-octane content that promotes better the process of combustion. Another advantage of methanol is that if it does ignite can cause less severe fires to the vehicle because is less flammable than gasoline [4]. Some disadvantages that methanol has are the lower energy content compared to gasoline, the fact that is not volatile enough for easy cold starting and can damage plastic and rubber fuel system components. The vehicle that uses methanol for fuel must have a large storage tank because pure methanol burns faster than gasoline, and corrosion resistant, materials must be used for the storage equipment [14-16]. Renewable fuels such as ethanol and methanol will probably replace petroleum-based fuels in the near future because petroleum reserves are not sufficient enough to last many years. Also, the severe environ‐ mental problems around the world will eventually lead to the use of more environmentally friendly technologies. The question that is examined in this chapter is how the mixtures of gasoline-ethanol and gasoline-methanol behave in a four-stroke engine from the aspect of

The experimental measurements were carried out on a four-stroke, air-cooled engine. This is

native generator (230V/50Hz) with maximum electrical load approximately 1ΚW(picture 1). The engine according to the manufacturer uses as fuel gasoline. The engine functioned with‐ out load and under full load conditions (1KW) using different fuel mixtures: gasoline, gaso‐ line-10%ethanol, gasoline-20%ethanol, gasoline-30%ethanol, gasoline-40%ethanol, gasoline-50%ethanol, gasoline-60%ethanol, gasoline-70%ethanol gasoline-80%ethanol gaso‐ line-90%ethanol and 100% ethanol, gasoline-10% methanol, gasoline-20%methanol, gaso‐ line-30%methanol, gasoline-40%methanol, gasoline-50%methanol, gasoline-60%methanol, gasoline-70%methanol. During the tests, exhaust gases measurements, were also monitored for every fuel mixture and for every load conditions. Also, during the function of the engine the consumption was recorded for every fuel. There was lack of engine regulation concern‐

displacement that is connected with a phase single alter‐

emissions and fuel consumption.

a one-cylinder engine with 123cm3

**2. Experimental part**

Using renewable fuels, such as ethanol, there is also a reduction of carbon dioxide (CO2) in the atmosphere. Carbon dioxide is non-toxic but contributes to the greenhouse effect. Because of the fact that plants absorb carbon dioxide and give off oxygen, that balances the amount of CO2 that is formed during combustion absorbed by plants used to produce ethanol. That is why the use of ethanol will partially offset the greenhouse effect that is formed by carbon dioxide emissions of burning gasoline [11-13]. Ethanol, as an octane enhancer, can substitute benzene and other benzene-like compounds, which are powerful liver carcinogens, and reduce their emissions to the atmosphere. Besides the environmental benefits, production and use of ethanol, which is a renewable fuel, increases economic activity, creates job openings, stabilizes prices and can increase farm income. That is why ethanol as an automotive fuel has many advantages.

Methanol (CH3OH) is an alcohol that is produced from natural gas, biomass, coal and also municipal solid wastes and sewage. It is quite corrosive and poisonous and has lower volatility compared to gasoline, which means that is not instantly flammable. Usually methanol is used as a gasoline-blending compound, but it can be used directly as an automobile fuel with some modifications of the automobile engine.

Although there are many feedstocks that are being used for the production of methanol, natural gas is more economic. Methanol is produced from natural gas with a technology of steam reforming. By this method natural gas is transformed to a synthesis gas that is fed to a reactor vessel to produce methanol and water at the presence of a catalyst. The reac‐ tions(equation 1,2) that represent methanol production are the following [4]:

$$\text{2CH}\_4 + \text{3H}\_2\text{O} \rightarrow \text{CO} + \text{CO}\_2 + \text{7H}\_2 \text{ - Syrtheisi gas} \tag{1}$$

$$2\text{ CO} + \text{CO}\_2 + 7\text{H}\_2 \quad \rightarrow \quad 2\text{CH}\_3\text{OH} + 2\text{H}\_2 + \text{H}\_2\text{O} \tag{2}$$

The main advantage of methanol as fuel is that is being produced from resources that can be found globally, while a large percentage of petroleum is located in Middle East. Furthermore, the materials needed for methanol production such as natural gas or biomass are renewable. This means that methanol can also be cheaper and more economically attractive than gasoline. When fossil fuels are used in automobiles produce exhaust emissions of hydrocarbons, carbon dioxide and other gases that contribute to the greenhouse effect. Methanol can give lower HC and CO emissions and besides that the vehicles that use methanol emit minimum particulate matter compared to gasoline, which usually has damaging effect to humans. In addition, methanol has high-octane content that promotes better the process of combustion. Another advantage of methanol is that if it does ignite can cause less severe fires to the vehicle because is less flammable than gasoline [4]. Some disadvantages that methanol has are the lower energy content compared to gasoline, the fact that is not volatile enough for easy cold starting and can damage plastic and rubber fuel system components. The vehicle that uses methanol for fuel must have a large storage tank because pure methanol burns faster than gasoline, and corrosion resistant, materials must be used for the storage equipment [14-16]. Renewable fuels such as ethanol and methanol will probably replace petroleum-based fuels in the near future because petroleum reserves are not sufficient enough to last many years. Also, the severe environ‐ mental problems around the world will eventually lead to the use of more environmentally friendly technologies. The question that is examined in this chapter is how the mixtures of gasoline-ethanol and gasoline-methanol behave in a four-stroke engine from the aspect of emissions and fuel consumption.

#### **2. Experimental part**

The experimental measurements were carried out on a four-stroke, air-cooled engine. This is a one-cylinder engine with 123cm3 displacement that is connected with a phase single alter‐ native generator (230V/50Hz) with maximum electrical load approximately 1ΚW(picture 1). The engine according to the manufacturer uses as fuel gasoline. The engine functioned with‐ out load and under full load conditions (1KW) using different fuel mixtures: gasoline, gaso‐ line-10%ethanol, gasoline-20%ethanol, gasoline-30%ethanol, gasoline-40%ethanol, gasoline-50%ethanol, gasoline-60%ethanol, gasoline-70%ethanol gasoline-80%ethanol gaso‐ line-90%ethanol and 100% ethanol, gasoline-10% methanol, gasoline-20%methanol, gaso‐ line-30%methanol, gasoline-40%methanol, gasoline-50%methanol, gasoline-60%methanol, gasoline-70%methanol. During the tests, exhaust gases measurements, were also monitored for every fuel mixture and for every load conditions. Also, during the function of the engine the consumption was recorded for every fuel. There was lack of engine regulation concern‐ ing the stable air/fuel ratio. For this purpose, data Acquisition cart was used with the termi‐ nal wiring board with on-board Cold Junction The data acquisition card was installed at a PC. This particular measuring system and software completed a scanning cycle per channel every 0.1 second approximately. This measuring speed was considered adequate for the purpose of the experiment and the sampling capabilities of the chemical sensors. For the ex‐ haust gas (CO and HC) measurements a analyzer was used.

**Figure 1.** The illustration of the experimental unit

The figures of CO and HC emissions, for every fuel and for every load conditions, are represented below [4-7]:

**10%methanol**

**10%methanol**

**gasoline**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

is used as fuel

**CO(%)**

is used as fuel

**CO(%)**

**CO(%)**

**CO(%)**

**CO(%)**

**Figure 3.**The CO variation when mixture of gasoline-10%methanol **Figure 5.**The CO variation when mixture of gasoline-30%methanol

**CO(%)**

**30%methanol**

**Figure 4.** The CO variation when mixture of gasoline-20%methanol is used as fuel

**30%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**20%methanol**

**Figure 3.** The CO variation when mixture of gasoline-10%methanol is used as fuel

**10%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**gasoline**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**20%methanol**

is used as fuel is used as fuel

Biofuels Ethanol and Methanol in OTTO Engines

http://dx.doi.org/10.5772/52772

0 50 100 150 200 250 300 350 400 450 500

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**time(s)**

**Figure 3.**The CO variation when mixture of gasoline-10%methanol **Figure 5.**The CO variation when mixture of gasoline-30%methanol

is used as fuel is used as fuel

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

is used as fuel is used as fuel

**Figure 2.** The CO variation when gasoline is used as fuel. **Figure 4.** The CO variation when mixture of gasoline-20%methanol

**Figure 3.**The CO variation when mixture of gasoline-10%methanol **Figure 5.**The CO variation when mixture of gasoline-30%methanol

is used as fuel

**CO(%)**

**CO(%)**

**30%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**20%methanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**Figure 2.** The CO variation when gasoline is used as fuel. **Figure 4.** The CO variation when mixture of gasoline-20%methanol **Figure 2.** The CO variation when gasoline is used as fuel **Figure 2.** The CO variation when gasoline is used as fuel. **Figure 4.** The CO variation when mixture of gasoline-20%methanol

**CO(%)**

is used as fuel

**CO(%)**

**CO(%)**

**30%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**20%methanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**Figure 3.**The CO variation when mixture of gasoline-10%methanol **Figure 5.**The CO variation when mixture of gasoline-30%methanol

#### **10%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**gasoline**

**Figure 3.** The CO variation when mixture of gasoline-10%methanol is used as fuel

**CO(%)**

#### is used as fuel is used as fuel **20%methanol**

**30%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 2.** The CO variation when gasoline is used as fuel. **Figure 4.** The CO variation when mixture of gasoline-20%methanol **Figure 4.** The CO variation when mixture of gasoline-20%methanol is used as fuel

is used as fuel

**CO(%)**

**Figure 3.**The CO variation when mixture of gasoline-10%methanol **Figure 5.**The CO variation when mixture of gasoline-30%methanol

is used as fuel is used as fuel

**10%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**gasoline**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**CO(%)**

is used as fuel

**CO(%)**

**Figure 2.** The CO variation when gasoline is used as fuel. **Figure 4.** The CO variation when mixture of gasoline-20%methanol

is used as fuel is used as fuel

**10%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**gasoline**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**CO(%)**

**CO(%)**

#### **30%methanol**

0 50 100 150 200 250 300 350 400 450 500

**20%methanol**

**Figure 3.**The CO variation when mixture of gasoline-10%methanol **Figure 5.**The CO variation when mixture of gasoline-30%methanol **Figure 5.** The CO variation when mixture of gasoline-30%methanol is used as fuel

**50%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

is used as fuel is used as fuel

**50%methanol**

is used as fuel is used as fuel

is used as fuel. is used as fuel.

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

is used as fuel. is used as fuel.

**Figure 7.** The CO variation when mixture of gasoline-50%methanol **Figure 9.** The CO variation when mixture of gasoline-70%methanol

**CO(%)**

**CO(%)**

**CO(%)**

**CO(%)**

**Figure 7.** The CO variation when mixture of gasoline-50%methanol **Figure 9.** The CO variation when mixture of gasoline-70%methanol

**CO(%)**

**CO(%)**

**Figure 6.**The CO variation when mixture of gasoline-40%methanol **Figure 8.** The CO variation when mixture of gasoline-60%methanol

**CO(%)**

**CO(%)**

**60%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**70%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 7.** The CO variation when mixture of gasoline-50%methanol **Figure 9.** The CO variation when mixture of gasoline-70%methanol

is used as fuel is used as fuel

Biofuels Ethanol and Methanol in OTTO Engines

http://dx.doi.org/10.5772/52772

**40%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**50%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**70%methanol**

**60%methanol**

is used as fuel. is used as fuel.

**60%methanol**

**Figure 7.** The CO variation when mixture of gasoline-50%methanol is used as fuel.

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**70%methanol**

**Figure 8.** The CO variation when mixture of gasoline-60%methanol is used as fuel

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 6.**The CO variation when mixture of gasoline-40%methanol **Figure 8.** The CO variation when mixture of gasoline-60%methanol **Figure 6.** The CO variation when mixture of gasoline-40%methanol is used as fuel **Figure 6.**The CO variation when mixture of gasoline-40%methanol **Figure 8.** The CO variation when mixture of gasoline-60%methanol

**CO(%)**

**CO(%)**

**CO(%)**

is used as fuel is used as fuel

is used as fuel. is used as fuel.

**60%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**70%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

#### **50%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**40%methanol**

**Figure 7.** The CO variation when mixture of gasoline-50%methanol **Figure 9.** The CO variation when mixture of gasoline-70%methanol **Figure 7.** The CO variation when mixture of gasoline-50%methanol is used as fuel.

**CO(%)**

#### **60%methanol**

**70%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 6.**The CO variation when mixture of gasoline-40%methanol **Figure 8.** The CO variation when mixture of gasoline-60%methanol **Figure 8.** The CO variation when mixture of gasoline-60%methanol is used as fuel

is used as fuel is used as fuel

**40%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**50%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**CO(%)**

**CO(%)**

is used as fuel. is used as fuel.

**Figure 7.** The CO variation when mixture of gasoline-50%methanol **Figure 9.** The CO variation when mixture of gasoline-70%methanol

**CO(%)**

**40%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**50%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**CO(%)**

**CO(%)**

#### **70%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**60%methanol**

**Figure 7.** The CO variation when mixture of gasoline-50%methanol **Figure 9.** The CO variation when mixture of gasoline-70%methanol **Figure 9.** The CO variation when mixture of gasoline-70%methanol is used as fuel.

 is used as fuel. is used as fuel. Figure 2 represents CO emissions when the fuel that is used is gasoline. The engine functions without load at first and then (after 250s) functions under full load conditions (1KW). The average value of CO emissions during the function of the engine without load is 6,41%, while at full load conditions the average value of CO emissions is 8,7%. Following, a mixture of gasoline with 10% methanol is used (fig. 3) and the same test is conducted with this mixture. From figure 3 it is being observed that the average value of CO emissions without load conditions of the engine is 4,87%, while at full load conditions the percentage of CO emissions is 6,9%. The same tests are conducted while increasing the percentage of the methanol in the fuel, using the mixtures: gasoline-20%methanol(fig. 4), gasoline-30%methanol(fig. 5), gaso‐ line-40%methanol(fig. 6), gasoline-50%methanol(fig. 7), gasoline-60%methanol(fig. 8), and gasoline-70%methanol(fig. 9).

> The HC emissions when the fuel that is used is gasoline are represented at figure 10. As it was mentioned above, the engine functioned without load at first and then (after 250s approxi‐ mately) functioned under full load conditions (1KW). During the function of the engine without load the average value of HC emissions is 1091ppm, while at full load conditions the average value of HC emissions is 730ppm. The mixture of gasoline with 10% methanol is illustrated at figure 11. At this figure is being observed that the average value of HC emissions without load conditions of the engine is 496ppm, while at full load conditions the HC emissions is 613ppm. When the percentage of the methanol in the fuel increases: gasoline-20%metha‐ nol(fig. 11), gasoline-30%methanol(fig. 13), gasoline-40%methanol(fig. 14), gasoline-50%meth‐ anol(fig. 15), gasoline-60%methanol(fig. 16), and gasoline-70%methanol(fig. 17).

**Figure 10.** The HC variation when gasoline is used as fuel. **Figure 12.**The HC variation when mixture of gasoline-20%methanol

**Figure 10.** The HC variation when gasoline is used as fuel. **Figure 12.**The HC variation when mixture of gasoline-20%methanol

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**20%methanol**

**20%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**30%methanol**

**30%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 11.** The HC variation when mixture of gasoline-10%methanol **Figure 13.**The HC variation when mixture of gasoline-30%methanol

**Figure 11.** The HC variation when mixture of gasoline-10%methanol **Figure 13.**The HC variation when mixture of gasoline-30%methanol

is used as fuel.

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is used as fuel.

**10%methanol**

**10%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**gasoline**

**gasoline**

**HC(ppm)**

**HC(ppm)**

**Figure 10.** The HC variation when gasoline is used as fuel.

**HC(ppm)**

**HC(ppm)**

is used as fuel is used as fuel

is used as fuel is used as fuel

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 11.** The HC variation when mixture of gasoline-10%methanol is used as fuel

**20%methanol**

**20%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**30%methanol**

**30%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

#### **gasoline**

**gasoline**

**Figure 10.** The HC variation when gasoline is used as fuel. **Figure 12.**The HC variation when mixture of gasoline-20%methanol **Figure 10.** The HC variation when gasoline is used as fuel. **Figure 10.** The HC variation when gasoline is used as fuel. **Figure 12.**The HC variation when mixture of gasoline-20%methanol is used as fuel.

#### **10%methanol**

**10%methanol**

is used as fuel.

is used as fuel is used as fuel

is used as fuel is used as fuel

**Figure 11.** The HC variation when mixture of gasoline-10%methanol **Figure 13.**The HC variation when mixture of gasoline-30%methanol

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**Figure 11.** The HC variation when mixture of gasoline-10%methanol **Figure 13.**The HC variation when mixture of gasoline-30%methanol **Figure 11.** The HC variation when mixture of gasoline-10%methanol is used as fuel

#### **20%methanol 20%methanol**

**Figure 10.** The HC variation when gasoline is used as fuel. **Figure 12.**The HC variation when mixture of gasoline-20%methanol **Figure 12.** The HC variation when mixture of gasoline-20%methanol is used as fuel. **Figure 10.** The HC variation when gasoline is used as fuel. **Figure 12.**The HC variation when mixture of gasoline-20%methanol is used as fuel.

is used as fuel.

**10%methanol**

**10%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**gasoline**

**gasoline**

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

is used as fuel is used as fuel

is used as fuel is used as fuel

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**time(s)**

**Figure 14.** The HC variation when mixture of gasoline-40%methanol **Figure 16.**The HC variation when mixture of gasoline-60%methanol

**Figure 14.** The HC variation when mixture of gasoline-40%methanol **Figure 16.**The HC variation when mixture of gasoline-60%methanol

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**60%methanol**

**60%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**70%methanol**

**70%methanol**

**Figure 15.** The HC variation when mixture of gasoline-50%methanol **Figure 17.**The CO variation when mixture of gasoline-70%methanol

**Figure 15.** The HC variation when mixture of gasoline-50%methanol **Figure 17.**The CO variation when mixture of gasoline-70%methanol

is used as fuel. is used as fuel.

is used as fuel. is used as fuel.

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**40%methanol**

**40%methanol**

0 50 100 150 200 250 300 350 400 450 500

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**time(s)**

**50%methanol**

**50%methanol**

**Figure 14.** The HC variation when mixture of gasoline-40%methanol is used as fuel.

**Figure 15.** The HC variation when mixture of gasoline-50%methanol is used as fuel.

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

is used as fuel. is used as fuel.

is used as fuel. is used as fuel.

#### **30%methanol**

**Figure 11.** The HC variation when mixture of gasoline-10%methanol **Figure 13.**The HC variation when mixture of gasoline-30%methanol **Figure 13.** The HC variation when mixture of gasoline-30%methanol is used as fuel

**Figure 11.** The HC variation when mixture of gasoline-10%methanol **Figure 13.**The HC variation when mixture of gasoline-30%methanol

**60%methanol**

**60%methanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**70%methanol**

**70%methanol**

#### **40%methanol 40%methanol**

**Figure 14.** The HC variation when mixture of gasoline-40%methanol **Figure 16.**The HC variation when mixture of gasoline-60%methanol **Figure 14.** The HC variation when mixture of gasoline-40%methanol is used as fuel. **Figure 14.** The HC variation when mixture of gasoline-40%methanol **Figure 16.**The HC variation when mixture of gasoline-60%methanol is used as fuel. is used as fuel.

is used as fuel. is used as fuel.

is used as fuel. is used as fuel.

is used as fuel. is used as fuel.

**Figure 15.** The HC variation when mixture of gasoline-50%methanol **Figure 17.**The CO variation when mixture of gasoline-70%methanol

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**time(s)**

**Figure 15.** The HC variation when mixture of gasoline-50%methanol **Figure 17.**The CO variation when mixture of gasoline-70%methanol **Figure 15.** The HC variation when mixture of gasoline-50%methanol is used as fuel.

#### **60%methanol 60%methanol**

**Figure 14.** The HC variation when mixture of gasoline-40%methanol **Figure 16.**The HC variation when mixture of gasoline-60%methanol **Figure 16.** The HC variation when mixture of gasoline-60%methanol is used as fuel. **Figure 14.** The HC variation when mixture of gasoline-40%methanol **Figure 16.**The HC variation when mixture of gasoline-60%methanol is used as fuel. is used as fuel.

is used as fuel. is used as fuel.

**40%methanol**

**40%methanol**

0 50 100 150 200 250 300 350 400 450 500

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**time(s)**

**50%methanol**

**50%methanol**

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

**HC(ppm)**

is used as fuel. is used as fuel.

is used as fuel. is used as fuel.

**time(s)**

9

9

In the case of HC emissions there is also a decrease of emissions when the percentage of methanol in the fuel increases at idle and under full load conditions. There is an exception at the mixture gasoline-70%methanol where the average value of HC without load is 534ppm and under full load is 367ppm. These values are higher than the values that correspond to the mixture of gasoline-60%methanol (295ppm, 298ppm). This is explained by mentioning the fact that during the use of the mixture gasoline-70%methanol there was a malfunction of the engine that was cause by the bad mixture of the air with the fuel(gasoline-70%methanol), since the engine was not regulated(ratio air/fuel) for every mixture maintaining the adjustments for gasoline. Also it must reported that the addition of methanol in the fuel led to HC decrease for the same mixture but for different load conditions. When gasoline was used HC emissions were higher at no load conditions than at full load conditions(1KW), while during the use of gasoline-methanol mixtures this was reversed. This is due to the better combustion under full

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91

load conditions because methanol has higher octane number than gasoline [4-7].

the variation of those emissions can be better understood.

could not function properly.

**CO(%)**

**C O (% )**

It is important to mention that when mixture gasoline-80%methanol was tested the engine

The CO and HC emissions are represented in the figures below, for the mixtures: gasoline, gasoline-ethanol, for every fuel and for every load conditions. For these mixtures the average values of the emissions (CO, HC) are presented at the figures below. From the average values,

**10%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**Figure 18.** The CO variation for the gasoline-10% ethanol mixture is used as fuel

**Figure 18.** The CO variation for the gasoline-10% ethanol mixture is used as fuel

**20%ethanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 19.** The CO variation for the gasoline-20% ethanol mixture is used as fuel

11

**Figure 15.** The HC variation when mixture of gasoline-50%methanol **Figure 17.**The CO variation when mixture of gasoline-70%methanol **Figure 17.** The CO variation when mixture of gasoline-70%methanol is used as fuel.

**Figure 15.** The HC variation when mixture of gasoline-50%methanol **Figure 17.**The CO variation when mixture of gasoline-70%methanol

In the case of HC emissions there is also a decrease of emissions when the percentage of methanol in the fuel increases at idle and under full load conditions. There is an exception at the mixture gasoline-70%methanol where the average value of HC without load is 534ppm and under full load is 367ppm. These values are higher than the values that correspond to the mixture of gasoline-60%methanol (295ppm, 298ppm). This is explained by mentioning the fact that during the use of the mixture gasoline-70%methanol there was a malfunction of the engine that was cause by the bad mixture of the air with the fuel(gasoline-70%methanol), since the engine was not regulated(ratio air/fuel) for every mixture maintaining the adjustments for gasoline. Also it must reported that the addition of methanol in the fuel led to HC decrease for the same mixture but for different load conditions. When gasoline was used HC emissions were higher at no load conditions than at full load conditions(1KW), while during the use of gasoline-methanol mixtures this was reversed. This is due to the better combustion under full load conditions because methanol has higher octane number than gasoline [4-7].

It is important to mention that when mixture gasoline-80%methanol was tested the engine could not function properly.

The CO and HC emissions are represented in the figures below, for the mixtures: gasoline, gasoline-ethanol, for every fuel and for every load conditions. For these mixtures the average values of the emissions (CO, HC) are presented at the figures below. From the average values, the variation of those emissions can be better understood.

**20%ethanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 19.** The CO variation for the gasoline-20% ethanol mixture is used as fuel

#### **10%ethanol**

**Figure 18.** The CO variation for the gasoline-10% ethanol mixture is used as fuel **Figure 18.** The CO variation for the gasoline-10% ethanol mixture is used as fuel

**CO(%)**

**C O (% )**

**CO(%)**

#### **20%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**10%ethanol**

**Figure 19.** The CO variation for the gasoline-20% ethanol mixture is used as fuel **Figure 19.** The CO variation for the gasoline-20% ethanol mixture is used as fuel

#### **30%ethanol**

11

12

93

13

**30%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

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0 50 100 150 200 250 300 350 400 450 500

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**60%ethanol**

**Figure 22.** The CO variation when gasoline-50% ethanol mixture used as fuel

**Figure 22.** The CO variation when gasoline-50% ethanol mixture used as fuel

**Figure 23.** The CO variation when gasoline-60% ethanol mixture used as fuel

**50%ethanol**

**Figure 21.** The CO variation for the gasoline-40% ethanol mixture is used as fuel **Figure 21.** The CO variation for the gasoline-40% ethanol mixture is used as fuel

**40%ethanol**

**Figure 20.** The CO variation for the gasoline-30% ethanol mixture is used as fuel

**CO(% )**

**C O (% )**

**CO(%)**

**C O (% )**

12

**40%ethanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 21.** The CO variation for the gasoline-40% ethanol mixture is used as fuel

**Figure 20.** The CO variation for the gasoline-30% ethanol mixture is used as fuel **Figure 20.** The CO variation for the gasoline-30% ethanol mixture is used as fuel

13

#### **40%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**30%ethanol**

**Figure 21.** The CO variation for the gasoline-40% ethanol mixture is used as fuel **Figure 21.** The CO variation for the gasoline-40% ethanol mixture is used as fuel

**C O (% )**

#### **50%ethanol**

**60%ethanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 23.** The CO variation when gasoline-60% ethanol mixture used as fuel

**Figure 22.** The CO variation when gasoline-50% ethanol mixture used as fuel **Figure 22.** The CO variation when gasoline-50% ethanol mixture used as fuel

**C O (% )**

### **Figure 23.** The CO variation when gasoline-60% ethanol mixture used as fuel **Figure 23.** The CO variation when gasoline-60% ethanol mixture used as fuel

#### **70%ethanol**

13

14

15

**70%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

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95

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**100%ethanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 26.** The CO variation when gasoline-90% ethanol mixture used as fuel

**Figure 26.** The CO variation when gasoline-90% ethanol mixture used as fuel

**Figure 27.** The CO variation when 100% ethanol used as fuel

**90%ethanol**

**Figure 25.** The CO variation when gasoline-80% ethanol mixture used as fuel **Figure 25.** The CO variation when gasoline-80% ethanol mixture used as fuel

**80%ethanol**

**Figure 24.** The CO variation when gasoline-70% ethanol mixture used as fuel

**CO(% )**

**CO(% )**

**CO(%)**

**C O (% )**

14

**80%ethanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 25.** The CO variation when gasoline-80% ethanol mixture used as fuel

**Figure 24.** The CO variation when gasoline-70% ethanol mixture used as fuel **Figure 24.** The CO variation when gasoline-70% ethanol mixture used as fuel

**CO(%)**

#### **60%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**50%ethanol**

15

#### **80%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**70%ethanol**

**Figure 25.** The CO variation when gasoline-80% ethanol mixture used as fuel **Figure 25.** The CO variation when gasoline-80% ethanol mixture used as fuel

**C O (% )**

#### **90%ethanol**

**100%ethanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 26.** The CO variation when gasoline-90% ethanol mixture used as fuel **Figure 26.** The CO variation when gasoline-90% ethanol mixture used as fuel

**Figure 27.** The CO variation when 100% ethanol used as fuel

**CO(%**

 **)**

#### **100%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**90%ethanol**

**Figure 27.** The CO variation when 100% ethanol used as fuel **Figure 27.** The CO variation when 100% ethanol used as fuel

#### **10%ethanol**

**10%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

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0 50 100 150 200 250 300 350 400 450 500

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**Figure 31.** The HC variation when gasoline-40% ethanol mixture used as fuel

**40%ethanol**

**Figure 30.** The HC variation when gasoline-30% ethanol mixture used as fuel

**Figure 30.** The HC variation when gasoline-30% ethanol mixture used as fuel

**30%ethanol**

**Figure 29.** The HC variation when gasoline-20% ethanol mixture used as fuel **Figure 29.** The HC variation when gasoline-20% ethanol mixture used as fuel

**20%ethanol**

**Figure 28.** The HC variation when gasoline-10% ethanol mixture used as fuel

**HC (ppm)**

**HC (ppm)**

**HC (ppm)**

**HC (ppm)**

**20%ethanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 29.** The HC variation when gasoline-20% ethanol mixture used as fuel

**Figure 28.** The HC variation when gasoline-10% ethanol mixture used as fuel **Figure 28.** The HC variation when gasoline-10% ethanol mixture used as fuel

**HC (ppm)**

17

**20%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**10%ethanol**

**Figure 29.** The HC variation when gasoline-20% ethanol mixture used as fuel **Figure 29.** The HC variation when gasoline-20% ethanol mixture used as fuel

**HC (ppm)**

#### **30%ethanol**

**40%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**Figure 31.** The HC variation when gasoline-40% ethanol mixture used as fuel

**Figure 30.** The HC variation when gasoline-30% ethanol mixture used as fuel **Figure 30.** The HC variation when gasoline-30% ethanol mixture used as fuel

**HC (ppm)**

**HC (ppm)**

#### **Figure 31.** The HC variation when gasoline-40% ethanol mixture used as fuel **Figure 31.** The HC variation when gasoline-40% ethanol mixture used as fuel

#### **50%ethanol**

**50%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

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0 50 100 150 200 250 300 350 400 450 500

**time(s)**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 34.** The HC variation when gasoline-70% ethanol mixture used as fuel

**Figure 34.** The HC variation when gasoline-70% ethanol mixture used as fuel

**Figure 35.** The HC variation when gasoline-80% ethanol mixture used as fuel

**80%ethanol**

**Figure 33.** The HC variation when gasoline-60% ethanol mixture used as fuel **Figure 33.** The HC variation when gasoline-60% ethanol mixture used as fuel

**70%ethanol**

**60%ethanol**

**Figure 32.** The HC variation when gasoline-50% ethanol mixture used as fuel

**HC(ppm)**

**HC (ppm )**

**HC (ppm)**

**HC (ppm)**

**60%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**Figure 33.** The HC variation when gasoline-60% ethanol mixture used as fuel

**Figure 32.** The HC variation when gasoline-50% ethanol mixture used as fuel **Figure 32.** The HC variation when gasoline-50% ethanol mixture used as fuel

**HC (ppm)**

#### **40%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**30%ethanol**

19

#### **60%ethanol**

**Figure 32.** The HC variation when gasoline-50% ethanol mixture used as fuel

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**50%ethanol**

**Figure 33.** The HC variation when gasoline-60% ethanol mixture used as fuel **Figure 33.** The HC variation when gasoline-60% ethanol mixture used as fuel

**HC (ppm)**

#### **70%ethanol**

**80%ethanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 35.** The HC variation when gasoline-80% ethanol mixture used as fuel

**Figure 34.** The HC variation when gasoline-70% ethanol mixture used as fuel **Figure 34.** The HC variation when gasoline-70% ethanol mixture used as fuel

**HC(ppm)**

**HC (ppm )**

#### **80%ethanol**

**Figure 34.** The HC variation when gasoline-70% ethanol mixture used as fuel

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**70%ethanol**

**Figure 35.** The HC variation when gasoline-80% ethanol mixture used as fuel **Figure 35.** The HC variation when gasoline-80% ethanol mixture used as fuel

#### **90%ethanol**

19

20

**90%ethanol**

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

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101

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

Figures 18 - 28 present the CO variation when as fuel is used gasoline –ethanol and gasoline –methanol mixtures when the engine functioned without load and under full load condi‐ tions(1ΚW). From these figures is observed lower CO emissions when gasoline-ethanol mixtures are used compared to the mixtures gasoline-methanol, until the mixture of 70% ethanol and methanol. Over the 70% percentage of methanol the engine could not function and that is why there is no further presentation of comparative curves of CO emissions. It must also be mentioned that for the mixtures of gasoline –70% methanol, gasoline –90%ethanol and 100%ethanol the engine malfunctioned. The average values of CO emissions for the above

In the figures 28 - 37 is observed higher decrease of HC in the case were methanol is used, with exception of the use of gasoline –70%methanol mixture where the HC are higher compared to the mixture gasoline-70%ethanol. This is due to the malfunction that occurred during the use of gasoline-70%methanol mixture. There was also malfunction of the engine when the mixtures of gasoline-90%ethanol and 100%ethanol were used, which had as result the HC increase during the use of those mixtures. These observations are presented more clearly in the figure

mixtures and for both load conditions are presented in the figure 38 below [4-7]:

**Figure 37.** The HC variation when 100% ethanol used as fuel **Figure 37.** The HC variation when 100% ethanol used as fuel

**100%ethanol**

**Figure 36.** The HC variation when gasoline-90% ethanol mixture used as fuel

38 below [4-7]:

**HC(ppm)**

**HC (ppm )**

20

**100%ethanol**

0 50 100 150 200 250 300 350 400 450 500 **time(s)**

**Figure 36.** The HC variation when gasoline-90% ethanol mixture used as fuel **Figure 36.** The HC variation when gasoline-90% ethanol mixture used as fuel

**Figure 37.** The HC variation when 100% ethanol used as fuel

**HC(ppm)**

#### **100%ethanol**

**Figure 36.** The HC variation when gasoline-90% ethanol mixture used as fuel

0 50 100 150 200 250 300 350 400 450 500

**time(s)**

**90%ethanol**

**Figure 37.** The HC variation when 100% ethanol used as fuel **Figure 37.** The HC variation when 100% ethanol used as fuel

**HC (ppm )**

Figures 18 - 28 present the CO variation when as fuel is used gasoline –ethanol and gasoline –methanol mixtures when the engine functioned without load and under full load condi‐ tions(1ΚW). From these figures is observed lower CO emissions when gasoline-ethanol mixtures are used compared to the mixtures gasoline-methanol, until the mixture of 70% ethanol and methanol. Over the 70% percentage of methanol the engine could not function and that is why there is no further presentation of comparative curves of CO emissions. It must also be mentioned that for the mixtures of gasoline –70% methanol, gasoline –90%ethanol and 100%ethanol the engine malfunctioned. The average values of CO emissions for the above mixtures and for both load conditions are presented in the figure 38 below [4-7]:

In the figures 28 - 37 is observed higher decrease of HC in the case were methanol is used, with exception of the use of gasoline –70%methanol mixture where the HC are higher compared to the mixture gasoline-70%ethanol. This is due to the malfunction that occurred during the use of gasoline-70%methanol mixture. There was also malfunction of the engine when the mixtures of gasoline-90%ethanol and 100%ethanol were used, which had as result the HC increase during the use of those mixtures. These observations are presented more clearly in the figure 38 below [4-7]:

figure 38 below[4-7]:

Figures 18 - 28 present the CO variation when as fuel is used gasoline –ethanol and gasoline –methanol mixtures when the engine functioned without load and under full load conditions(1ΚW). From these figures is observed lower CO emissions when gasolineethanol mixtures are used compared to the mixtures gasoline-methanol, until the mixture of 70% ethanol and methanol. Over the 70% percentage of methanol the engine could not function and that is why there is no further presentation of comparative curves of CO emissions. It must also be mentioned that for the mixtures of gasoline –70% methanol,

methanol mixture **Figure 38.** The CO emission average value for every gasoline-ethanol and gasoline-methanol mixture

compared to the mixture gasoline-70%ethanol. This is due to the malfunction that occurred during the use of gasoline-70%methanol mixture. There was also malfunction of the engine when the mixtures of gasoline-90%ethanol and 100%ethanol were used, which Figure 39 shows the average values of HC for every mixture, when the engine functions without load and under full load conditions. It is being observed grater decrease of HC during the use of methanol in the fuel contrary to the use of ethanol.

22

23

**1131**

103

**1450**

**441**

**463**

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had as result the HC increase during the use of those mixtures. These observations are

**Figure 39.** The HC emission average value for every gasoline-ethanol and gasoline-

**<sup>290</sup> <sup>327</sup>**

*417 362*

**433**

<sup>331</sup> <sup>400</sup> <sup>357</sup>

**Figure 39.** The HC emission average value for every gasoline-ethanol and gasoline-methanol mixtures

**338**

gasoline 10 20 30 40 50 60 70 80 90 100 **% ethanol and %methanol in fuel**

**214**

*298*

**full load**

**<sup>197</sup>** <sup>295</sup>

**gasoline**

**70%methanol**

**234 281**

**ethanol-engine without**

**ethanol-engine under full load methanol -engine without load methanol-engine under**

**load**

534

*367*

**191 188**

Figure 39 shows the average values of HC for every mixture, when the engine functions without load and under full load conditions. It is being observed grater decrease of HC

**Engine rpm**

Also is shown HC emissions decrease compared to gasoline, while the percentage of methanol and ethanol in the fuel increases without load and under full electrical load conditions (1KW). At higher percentage of ethanol in the fuel 90%ethanol and 100%ethanol it is observed HC emissions increase, which is due to incomplete combustion. Indeed, during the tests of the mixtures: gasoline-70%methanol, gasoline – 90%ethanol and 100%ethanol, there was an engine malfunction mostly at without electrical load, as it was mentioned above. This malfunction is showed from the rounds per minute recording

> **0 50 100 150 200 250 300 350 400 450 500 time(s)**

> > **Engine rpm**

**0 50 100 150 200 250 300 350 400 450 500 time(s)**

during the use of methanol in the fuel contrary to the use of ethanol.

**Figure 40**. The rpm variation when used fuel gasoline

**Figure 40.** The rpm variation when used fuel gasoline

**Figure 41**. The rpm variation when used fuel gasoline –70%methanol

**507**

*495*

**450**

**590**

*558*

**570**

presented more clearly in the figure 38 below[4-7]:

**947**

**709**

*613*

<sup>470</sup> <sup>496</sup>

**730**

**1091**

methanol mixtures

**H C** 

**[ppm] ave rage value**

in the figures below:

**2400**

**2400**

**2500**

**2600**

**2700**

**2800**

**rpm**

**2900**

**3000**

**3100**

**2500**

**2600**

**2700**

**2800**

**rpm**

**2900**

**3000**

**3100**

**Figure 38.** The CO emission average value for every gasoline-ethanol and gasoline-

In the figures 28 - 37 is observed higher decrease of HC in the case were methanol is used, with exception of the use of gasoline –70%methanol mixture where the HC are higher

21 Also is shown HC emissions decrease compared to gasoline, while the percentage of methanol and ethanol in the fuel increases without load and under full electrical load conditions (1KW). At higher percentage of ethanol in the fuel 90%ethanol and 100%ethanol it is observed HC emissions increase, which is due to incomplete combustion. Indeed, during the tests of the mixtures: gasoline-70%methanol, gasoline – 90%ethanol and 100%ethanol, there was an engine malfunction mostly at without electrical load, as it was mentioned above. This malfunction is showed from the rounds per minute recording in the figures below:

presented more clearly in the figure 38 below[4-7]:

**Figure 39.** The HC emission average value for every gasoline-ethanol and gasoline-methanol mixtures

during the use of methanol in the fuel contrary to the use of ethanol.

**Figure 41**. The rpm variation when used fuel gasoline –70%methanol

#### Figure 39 shows the average values of HC for every mixture, when the engine functions **Engine rpm**

without load and under full load conditions. It is being observed grater decrease of HC

**Figure 39.** The HC emission average value for every gasoline-ethanol and gasoline-

**Engine rpm**

**0 50 100 150 200 250 300 350 400 450 500 time(s)**

**70%methanol**

**Figure 40**. The rpm variation when used fuel gasoline **Figure 40.** The rpm variation when used fuel gasoline

**2400**

**2500**

**2600**

**2700**

**2800**

**rpm**

**2900**

**3000**

**3100**

methanol mixtures

**2500**

**2600**

**2700**

**2800**

**rpm**

**2900**

**3000**

**3100**

**Engine rpm**

**Engine rpm**

**0 50 100 150 200 250 300 350 400 450 500 time(s)**

**gasoline**

**Figure 41.** The rpm variation when used fuel gasoline –70%methanol

**Figure 41**. The rpm variation when used fuel gasoline –70%methanol

**Engine rpm**

**0 50 100 150 200 250 300 350 400 450 500 time(s)**

**100%ethanol**

#### **Engine rpm**

23

**Figure 42**. The rpm variation when used fuel gasoline –90%ethanol

**2400**

**1000**

**1300**

**1600**

**1900**

**2200**

**rpm**

**2500**

**2800**

**3100**

**2500**

**2600**

**2700**

**2800**

**rpm**

**2900**

**3000**

**3100**

**Engine rpm**

**90% ethanol**

**0 50 100 150 200 250 300 350 400 450 500**

**time(s)**

**Engine rpm**

**0 50 100 150 200 250 300 350 400 450 500 time(s)**

During the tests the rounds per minute of the engine were recorded as it was mentioned above. The normal variation of the engine rpm appears in figure 40. The same variation that is illustrated in this figure corresponds to the mixtures gasoline until the mixtures gasoline-90% ethanol and gasoline-60%methanol, without any change. As it is presented in figure 39, the average value of the engine rpm without load (0-200s and 420-500s) is approximately 2990rpm while at full load conditions (200-420s) the average value of the engine rpm is 2880rpm. It must be noted that the engine has a round stabilizer. In figures 41, 42 and 43 the mixtures gaso‐ line-70%methanol, gasoline-90%ethanol and 100%ethanol are illustrated and irregular variation of the engine rpm is presented, which is caused from the engine malfunction. Higher irregular variation is observed at without load condition, and lower at full load conditions in the case of use ethanol. This malfunction is due to the smaller calorific value of methanol and ethanol than the gasoline, and to the fact that there is no adjustment of the air/fuel ratio during the use of gasoline-methanol and gasoline-ethanol mixtures. The initial adjustment that

Furthermore, during the tests the consumption of the fuel was recorded for every mixture separately and for every load conditions. The results of the consumption recording are

**100%ethanol**

Biofuels Ethanol and Methanol in OTTO Engines

http://dx.doi.org/10.5772/52772

105

**Figure 43**. The rpm variation when used fuel 100%ethanol

**Figure 43.** The rpm variation when used fuel 100%ethanol

corresponds to gasoline as fuel is maintained [6,7].

illustrated in the figure below:

24

24

**Figure 42**. The rpm variation when used fuel gasoline –90%ethanol **Figure 42.** The rpm variation when used fuel gasoline –90%ethanol

**Figure 43**. The rpm variation when used fuel 100%ethanol

**1000**

**1300**

**1600**

**1900**

**2200**

**rpm**

**2500**

**2800**

**3100**

#### **Engine rpm**

**0 50 100 150 200 250 300 350 400 450 500**

**time(s)**

**Engine rpm**

**90% ethanol**

**Figure 42**. The rpm variation when used fuel gasoline –90%ethanol

**2400**

**2500**

**2600**

**2700**

**2800**

**rpm**

**2900**

**3000**

**3100**

**Figure 43**. The rpm variation when used fuel 100%ethanol **Figure 43.** The rpm variation when used fuel 100%ethanol

During the tests the rounds per minute of the engine were recorded as it was mentioned above. The normal variation of the engine rpm appears in figure 40. The same variation that is illustrated in this figure corresponds to the mixtures gasoline until the mixtures gasoline-90% ethanol and gasoline-60%methanol, without any change. As it is presented in figure 39, the average value of the engine rpm without load (0-200s and 420-500s) is approximately 2990rpm while at full load conditions (200-420s) the average value of the engine rpm is 2880rpm. It must be noted that the engine has a round stabilizer. In figures 41, 42 and 43 the mixtures gaso‐ line-70%methanol, gasoline-90%ethanol and 100%ethanol are illustrated and irregular variation of the engine rpm is presented, which is caused from the engine malfunction. Higher irregular variation is observed at without load condition, and lower at full load conditions in the case of use ethanol. This malfunction is due to the smaller calorific value of methanol and ethanol than the gasoline, and to the fact that there is no adjustment of the air/fuel ratio during the use of gasoline-methanol and gasoline-ethanol mixtures. The initial adjustment that corresponds to gasoline as fuel is maintained [6,7].

Furthermore, during the tests the consumption of the fuel was recorded for every mixture separately and for every load conditions. The results of the consumption recording are illustrated in the figure below:

recording are illustrated in the figure below:

During the tests the rounds per minute of the engine were recorded as it was mentioned above. The normal variation of the engine rpm appears in figure 40. The same variation that is illustrated in this figure corresponds to the mixtures gasoline until the mixtures gasoline-90% ethanol and gasoline-60%methanol, without any change. As it is presented in figure 39, the average value of the engine rpm without load (0-200s and 420- 500s) is approximately 2990rpm while at full load conditions (200-420s) the average value of the engine rpm is 2880rpm. It must be noted that the engine has a round stabilizer. In figures 41, 42 and 43 the mixtures gasoline-70%methanol, gasoline-90%ethanol and 100%ethanol are illustrated and irregular variation of the engine rpm is presented, which is caused from the engine malfunction. Higher irregular variation is observed at without load condition, and lower at full load conditions in the case of use ethanol. This malfunction is due to the smaller calorific value of methanol and ethanol than the gasoline, and to the fact that there is no adjustment of the air/fuel ratio during the use of gasolinemethanol and gasoline-ethanol mixtures. The initial adjustment that corresponds to

> tion compared to gasoline increase when the percentage of the methanol and ethanol in the fuel was increased in both load conditions. Between the use of methanol and ethanol mixtures is observed higher increase of consumption when the mixtures of methanol are used due to the fact that methanol has lower calorific value compared to ethanol. From the aspect of emissions, when the mixtures of gasoline with methanol and ethanol are compared, there is grater reduction of emissions in the case where methanol is used. It can be said that this is caused because of the smaller carbon chain of the methanol molecule, which results to the better combustion of methanol. It is also observed that the engine functions with the mixtures of methanol until the use of 70%methanol mixture with gasoline, while with ethanol mixtures until 100% ethanol as fuel (with the initial adjustment of the air/fuel ratio that is made for gasoline). This is due to the fact that ethanol has higher octane number compared to methanol. Finally, it is important the fact that methanol and ethanol are a renewable fuels, which present emissions decrease compared to gasoline, when they are used, in a time period where petroleum reservations are depleted and the environmental pollution is one of the most

Biofuels Ethanol and Methanol in OTTO Engines

http://dx.doi.org/10.5772/52772

107

Department of Production and Management Engineering, Democritus University of Thrace,

[1] Arapatsakos I. C, Sparis D. P., "Testing the Two Stroke Engine Using Mixtures of Gasoline - Ethanol" International Journal of Heat & Technology, Vol. 16, pp. 57-63, 1998.

[2] Arapatsakos I. C, "Air and Water Influence of Two Stroke Outboard Engine Using Gasoline -Ethanol Mixtures" Transactions of SAE 2000-01-2973, Book SP-1565, 2000.

[3] Arapatsakos I. Charalampos, "Testing The Tractor Engine Using Diesel –Ethanol Mixtures Under Full Load Conditions" International Journal of Heat & Technology,

[4] Arapatsakos I. Charalampos, Karkanis N. Anastasios, Sparis D. Panagiotis, "Gas Emissions And Engine Behavior When Gasoline-Alcohols Mixtures Are Used" Journal

[5] Arapatsakos I. Charalampos, Karkanis N. Anastasios, Sparis D. Panagiotis, "Tests On A Small Four Engine Using As Fuel Gasoline-Bioethanol Mixtures" Transactions of

important problems that humanity faces [4-7].

**Author details**

Xanthi, Greece

**References**

Charalampos Arapatsakos\*

Vol. 19, n.1, 2001.

WIT, ISSN 13569-5886, 2003.

of Environmental Technology, Vol. 24, 2003.

**Figure 44.** The fuel consumption

25 **Figure 44**. The fuel consumption Figure 44 shows an increase of fuel consumption when the percentage of methanol and ethanol in the fuel increases than gasoline. Also, between the use of the mixtures of methanol and ethanol is observed small increase during the use of methanol because of the smaller calorific value that methanol has compared to ethanol. The smaller calorific value of methanol and ethanol compared to gasoline and also the lack of regulation (ratio air/fuel) of the engine, results to the consumption increase contrary to the use gasoline. This increase of consumption happens automatically for the rounds regulator that the engine has, for the maintaining of the rounds constant.

#### **3. Conclusion**

From the observations above is appeared that methanol and ethanol as mixture with gasoline results in an emissions (CO and HC) decrease when the engine functions without load and under full load conditions. There is an exception in the use of the mixtures: gasoline-70%meth‐ anol, gasoline –90% ethanol and 100%ethanol where there is observed an HC emissions increase because of the incomplete combustion and consequently due to engine malfunction. Also, it must be mentioned that the adjustment of the engine (air/fuel ratio) was that which referred to the use of gasoline as fuel. From the aspect of consumption, there was a consump‐ tion compared to gasoline increase when the percentage of the methanol and ethanol in the fuel was increased in both load conditions. Between the use of methanol and ethanol mixtures is observed higher increase of consumption when the mixtures of methanol are used due to the fact that methanol has lower calorific value compared to ethanol. From the aspect of emissions, when the mixtures of gasoline with methanol and ethanol are compared, there is grater reduction of emissions in the case where methanol is used. It can be said that this is caused because of the smaller carbon chain of the methanol molecule, which results to the better combustion of methanol. It is also observed that the engine functions with the mixtures of methanol until the use of 70%methanol mixture with gasoline, while with ethanol mixtures until 100% ethanol as fuel (with the initial adjustment of the air/fuel ratio that is made for gasoline). This is due to the fact that ethanol has higher octane number compared to methanol. Finally, it is important the fact that methanol and ethanol are a renewable fuels, which present emissions decrease compared to gasoline, when they are used, in a time period where petroleum reservations are depleted and the environmental pollution is one of the most important problems that humanity faces [4-7].

#### **Author details**

Charalampos Arapatsakos\*

Department of Production and Management Engineering, Democritus University of Thrace, Xanthi, Greece

#### **References**


[6] Charalampos Arapatsakos, Anastasios Karkanis, Panagiotis Sparis, "Gasoline – Ethanol, Methanol Mixtures And A Small Four-Stroke Engine" International journal of heat and technology Vol. 22, n.2 2004.

**Chapter 4**

**Analytical Methodology for**

Fabiana Aparecida Lobo, Fernanda Pollo,

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/52514

ute to environmental contamination [2-5].

**1. Introduction**

Ana Cristina Villafranca and Mercedes de Moraes

**Determination of Trace Cu in Hydrated Alcohol Fuel**

Many private and governmental initiatives have been established worldwide to identify viable alternatives to petroleum derivatives [1,2].The goals are to reduce dependence on imported en‐ ergy from non-renewable sources, while mitigating environmental problems caused by petro‐

Ethyl alcohol (ethanol) is considered to be a highly viable alternative fuel. Its production from biomass means that it can provide a source of energy that is both clean and renewable. The in‐ clusion of ethanol as a component of gasoline can help to reduce problems of pollution in many regions, since it eliminates the needto use tetraethyl lead (historically notorious as a highly tox‐

The quantitative monitoring of metal elements in fuels (including gasoline, alcohol, and die‐ sel) is important from an economic perspective in the fuel industry as well as in the areas of transport and environment. The presence of metalspecies (ions or organometallic com‐ pounds) in automotive fuels can cause engine corrosion, reduce performance, and contrib‐

The low concentrations of metals in fuels typically require the use of sensitive spectrometric an‐ alytical techniques for the purposes of quality control. Atomic absorption spectrometry (AAS) can be applied for the quantitative determination of many elements (metals and semi-metals) in a wide variety of media including fuels, foodstuffs, and biological, environmental, and geo‐ logical materials, amongst others. The principle of the technique is based on measurement of the absorption of optical radiation, emitted from a source, by ground-state atoms in the gas phase. Atomization can be achieved using a flame, electrothermal heating, or specific chemical

> © 2013 Lobo et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 Lobo et al.; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

distribution, and reproduction in any medium, provided the original work is properly cited.

leum products, and to develop national technologies in the alternative energy field.

ic trace component of the atmosphere in major cities) as an anti-knock additive.

