Open access

Biofuels and Co-Products Out of Hemicelluloses

Written By

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

Submitted: August 18th, 2012 Published: March 20th, 2013

DOI: 10.5772/52645

Chapter metrics overview

3,590 Chapter Downloads

View Full Metrics

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 inserted in the diesel pool. Ethanol is usually produced out of fermentation of C6 sugars (although 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 Lavoieet 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 accompanied by lignin, hemicelluloses, extractives and, in case of agricultural biomass, proteins. 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 related 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 agricultural biomass as well as energy crops. In all cases, although the feedstock is rather inexpensive (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-generation biofuel requires specific pre-treatment that should be as versatile as efficient in order 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 average 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. With an actual market price of 0.48 USD per liter the value of this ethanol would range between 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 remaining 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 industry. 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.

Conversion of the carbohydrates is of course an important part of the process although; isolation 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 cellulosic fibres in order to produce the largest amount of pulp possible per ton of biomass. Nevertheless, 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 expensive 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, Schildet 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 bicarbonate [5]. Hot water extractions were also performed at temperature around 170 °C at different 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 performed 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 Liuet al. [9]. Utilisation of sulphuric acid, although very efficient to remove hemicellulose may also have an impact on cellulose thus reducing 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, instead of using only an aqueous mixture of ions, the process relies on the utilisation of a combination 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 butanol and certain organic acids have also been investigated to the same purposes. Recent work by Wanget 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 (sulphuric) 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 condition, 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 production 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 increase 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 formic acid. Glucose and xylose are often the most abundant sugars in hemicelluloses hydrolysis 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 detoxification 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 conversion 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).

Components(wt%) Energy crops Agricol residues Forestresidues Coniferous
Straw [15]
Stover [17]
Pine [19]
Glucan 38.5 55.5 39.2 36.2 52.4 36
Xylan 26.3 12.4 24.6 20.1 14.9 7.5
Galactan 1.16 - - 1.45 2.2 2.5
Mannan 0.13 - - - 2.3 8.2
Arabinan 3.41 - 1.9 3.0 0.9 1.6

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 production 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 approaches 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 molecule, one of the major side-reaction of furfural is polymerisation which influences the conversion 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 dehydration reactions under acid catalyst..

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]
NaCl, HCl 78% [27]
Aluminium chloride Hexahydrate 76% [28]
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]

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 aqueous 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.

Production of furfural itself is of course of significant interest because, amongst many factors, 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 selectivity 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-methyltetrahydrofuran (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-renewables.

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 reduction 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 decarboxylation 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 reported conversion with copper chromite in supercritical CO2.

Results reported in literature show that xylose, under an acid catalyst, tend invariably to dehydrate to furfural thus limiting the possibilities for side-products in such specific conditions. 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 catalyst, 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-catalyzed 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 reaction 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 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) pyrocatechol; (4) 3-methylbenzene-1,2-diol; (5) 4-methylbenzene-1,2-diol; (6) 3,4-dimethylbenzene-1,2-diol; (7) 2-methylbenzene-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 temperatures (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 sugars that were formed during the reaction, conversion of xylose was shown to be more efficient 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 xylose 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 temperature.

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 conversion of xylose to lactic acid could reach 64 % (molar) accompanied by glyceric acid. Although 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 conversion 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 glyceraldehyde as depicted in Figure 8 below. The possible reaction leading to methylglyoxal may involve an E2 reaction on C2 leading to removal of the hydroxyl group on C3 then a keto-enol rearrangement to methylglyoxal.

Figure 8.

Conversion of glyceraldehyde to methylglyoxal.

Onda et al. [56] achieved a conversion rate of more than 20 % when using xylose as a feedstock 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 convert 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 between the amount of catalyst (varying from 1-3 % wt.) of NaOH, KOH and Ca(OH)2 respectively. 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 performed 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 xylose (200 °C) in a Ca(OH)2 solution produced about 57 % (mol.) of lactic acid with 2,4-dihydroxybutanoic 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 chapter, 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 biologically based. Traditionally, the concept evolves around fermenting carbohydrate-based syrup 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 homofermentative 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 dehydrogenase (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 D-lactate-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 lactate 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 temperature 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 auxotroph 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) mutagenesis 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 (phosphoketolase), thus for a given strain, even if hexoses are fermented through an homofermentative 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 lactate for 2 moles of pentoses, but through the PK it decreases to 1:1 [61], thus, the conversion 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 dehydrogenase (L-LDH) gene was disrupted, leaving only the homologous D-lactate dehydrogenase (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. lactis, thereby shifting heterolactic fermentation to a homolactic one.

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, consequently 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 transforming C. utilis, disrupting the native pdc1 gene, and expressing heterologous LDH, XR (xylose reductase), XDH (xylitol dehydrogenase) and XK (xylulokinase) enzymes. Furthermore, to prevent the redox imbalance, they increased the XR’s NADH (reduced nicotinamide 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.

Strain Gen Eng Str Medium Process LA(g/L) Tf(h) Yield(g/g) Prd(g/L/h) Ref
E. mundtii QU 25 - Cellobiose Batch 119 106 0.83 1.12 [86]
- Xylose Batch 86.7 0.84 0.9 [85]
- Glucose/
Batch 35.1 15 0.91 2.99 [86]
Lactobacillus sp. RKY2 - Wood
27 - 0.9 6.7 [91]
Lb. bifermentas DSM 20003 - Wheat bran
Batch 62.8 60 0.83 1.17 [80]
Lb. casei NCIMB 3254 - Cassava
83.8 60 0.96 1.4 [92]
Lb. delbrueckii Uc-3 UV
Cellobiose Batch 90 40 0.9 2.25 [93]
Molasse Batch 166 40 0.95 4.15 [77]
Lb. lactis RM 2-24 UV
Cellobiose Batch 80 48 0.8 1.66 [94]
Cellulose Batch SSF 73 48 0.73 1.52
Lb. plantarum ΔldhL1-xpk1::tkt Disruption of endogenous LDH gene.
Replacment of endogenous PK (xpk1) gene with heterologous tkt to redirect the PK pathway to the PPP.
Arabinose Batch 38.6 28 0.82 1.37 [89]
Lb. plantarum ΔldhL1-xpk1::tkt-Δxpk2 Idem as above. Disruption of 2nd PK gene (xpk2) to terminate acetate production. Xylose Batch 41.2 60 0.89 0.67 [87]
Lb. rhamnosus ATCC 7469 - Paper
73 168 0.97 0.45 [95]
Lb. rhamnosus ATCC 9595 (CECT288) - Apple
Batch 32.5 6 0.88 5.41 [96]
L. lactis IO-1 - Xylose Batch 33.3 - 0.68 - [84]
S. cerevisiae recombinant Replacement of native pdc1 and pdc5 by heterologous bovine L-LDH gene. Glucose Batch 82.3 192 0.83 0.43 [97]
S. cerevisiae recombinant Disruption of pcd1 and adh1 genes. Expression of bovine L-LDH. Glucose Batch 71.8 65 0.74 1.1 [98]
K. lactis Disruption of PDC and PDH genes.
Expression of bovine L-LDH gene.
Glucose Semi-Batch 60 500 0.85 0.12 [99]
C. utilis Disruption of endogenous PDC gene.Expression of heterologous LDH, XR, XDH and XK. XR gene site-specific mutation for preferential NADH cofactor utilization Xylose Batch 66.7 78 0.79 1.62 [88]
Batch 93.9 78 0.91 2.18
P. stipitis Expression of LDH from L. helveticus. Xylose Batch 58 147 0.58 0.39 [90]

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

* No xylose consumption occurred

**SSF = simultaneous saccharification and fermentation

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. Racemic mixtures of lactic acid (most probably derived from chemical synthesis) can be evaluated 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 purity 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 efficiency 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 ultrasonic 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 catalytic 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, propylene glycol and glycerol, as depicted in Figure 10 below.

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 reduction 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 converted 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 given 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-copper 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), followed 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. Nevertheless, 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 isomerisation, 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 actually 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 species on the other hand present high yields, fast metabolism and many GRAS (generally recognized 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 co-substrate 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 metabolize both substrates. This functionality was achieved in E. coli [140]by replacing the putative 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 putative XK (xylB gene) silenced.

Other well suited candidates for such a bioconversion would be LAB, offering the advantage 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 transporter 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-pathogenic and anaerobic nature of L. lactis is a notable advantage.

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-phosphogluconate dehydrogenase, (15) ribulose-5P 3-epimerase, (16) xylulose-5P phosphoketolase, (17) phosphotransacetylase, (18) acetaldehyde dehydrogenase, (19) alcohol dehydrogenase; (20) pentose kinase, (21) pentose phosphate epimerase or isomerase, (22) acetate kinase. CoA coenzyme A.

Early work done on Corynebacterium glutamicum showed another alternative for the production of xylitol but the necessity of inserting gluconate as co-substrate for NADPH (nicotinamide adenine dinucleotide phosphate) regeneration rendered the application non economical [122,124]. Sasaki et al. [141] developed a C. glutamicum recombinant achieving simultaneous 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 extensive 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):

  • introduction homologous pentose transporter (araE);

  • disruption of the native lactate deshydrogenase (ldhA);

  • expression of single-site mutant XR from C. tenuis;

  • disruption of XK native gene (xylB);

  • disruption of phosphoenolpyruvate-dependent fructose phosphotransferase (ptsF gene; PTSfru).

Strain Genetic Engineering Strategy Yieldg/g Xyloseg/L Xylitol(g/L) Tf(h) Prd(g/l/h) ProcessStrategy Reference
Candida athensensis SB18 - 83% 250 207.8 175 1.15 Batch limited O2 [144]
87% 300 256.5 250 0.97 Fed Batch limited O2
79% 200 151.71 156 0.97 Batch limited O2
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. tropicalis KCTC 10457 - 87% 200 172 48 3.66 Batch limited O2 [147]
C. tropicalis KFCC 10960 - 93% 270 251 55 4.56 Fed Batch [148]
C. tropicalis KCTC 10457 - 90% 260 234 48 4.88 Fed Batch [147]
C. guilliermondii - 73% 250 - - - Fed Batch limited O2 [149]
C. tropicalis - 82% 750 189 58 4.94 Fed Batch/ Cell recylcing/ Glucose cosubstrate/ limited O2 [150]
C. tropicalis - 69% 100 - - 5.7 Cell recycling/ limited O2 [151]
C. tropicalis - 85% 214 182 15 12 cell recycling/ limited O2 [147]
S. cerevisiae Expression heterologous XR gene from P. stipitis. 95% 190 - - 0.4 Fed batch/ Glucose cosubstrate [152]
Corynebacterium glutamicum CtXR7 Expression of araE pentose transporter gene. Disruption of ldhA. Single site mutation of heterologous XR gene. Disruption of xylB& PTSfru genes. - 120 166 21 7.9 Fed batch/ Glucose cosubstrate/ 40g/L dry cell [135]
D. hansenii NRRL Y-7426 - 38% 45 19.7 72 0.274 Batch/ Detoxified grape marc hydrolysates [153]
S. cerevisiae Overexpression ALD6 & ACS1 genes.Expression of P. stipitis XR gene. ~100% 20 91.3 60 1.76 Fed batch/ Glucose cosubstrate [154]
Lactobacillus brevis NZ9800 Expression of P. stipitis XR gene.Expression of Lb. brevisxylT symporteur. ~100% 160 75 41 2.72 Fed batch/ Glucose cosubstrate [129]
C. tropicalis - 83% 80 96.5 120 1.01 Fed batch/ Corn Cob hydrolysates/limited O2 [155]
C. tropicalisSS2 Xylitol-assimilation deficient strain by chemical mutagenesis. 93% 100 220 70 3.3 Fed batch/ aerobic [156]
C. trolpicalis JH030 - 71% 45 31.1 80 0.44 Batch/ Rice straw hydrolysates [157]

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. tannophilus whose XDH shows a higher activity with NADP+ than NAD+ [158] proposed a theoretical 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 anaerobic conditions, the theoretical yield drops to 0.875. These yields follow the equations (1) and (2) below respectively:

126 xylose + 3 O 2 + 6 ADP + 6 P i + 48 H 2 O 114 xylitol + 6 ATP + 60 CO 2 E1
48 xylose + 15 H 2 O 42 Xylitol + 2 ethanol + 24 CO 2 E2

Owing the better yield both in xylitol and ATP (adenosine triphosphate) under oxygen-limited xylitol production, aeration is a crucial parameter. As a general trend, xylitol production 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 xylose to xylitol (95 % of theoretical). It has been suggested that the incapacity of S. cerevisiae 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 reported 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 original strain by 21 %. This was done by disrupting XDH putative gene, and over-expressing homologous glucose-6-phosphate dehydrogenase (G6PDH) and 6-phosphogluconate dehydrogenase (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 propylene 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 hydrogen 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 reported 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).

Figure 12.

Simplified conversion of xylose to xylonic acid

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 % selectivity with a conversion rate of 21 mmol/min/g (Au) in a continuous reactor. Nevertheless, contrarily 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).

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 reduction 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 by-product 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 simultaneously to xylonic acid and xylitol using electrotechnologies. Such process could be to a certain extent compared to the Cannizarro reaction where the original aldehyde is acting as redox reagent.

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

Figure 13.

Simplified scheme for the conversion of xylose xylaric acid

Conversion of C5 sugars and to a smaller extent of xylose into aldaric acids has been described in literature in a few reports. Kiely et al. [176] reported that a conversion up to 83 % xylose 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 tetrahydroxysuccinate. 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.

Severer oxidizing conditions leads to a breakage of the carbon-carbon bonds in the carbohydrate 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 hydroxide 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 molybdenum 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.

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 corylophilum, Pichiaquer cuum [185], Pseudomonas sp. [189, 190], Pullularia pullulans [191], Gluconobacter and Caulobacter [192, 193].

In metabolic pathways, xylose is converted to xylonate via 2 key enzymes. First, a xylose dehydrogenase (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 xylonate either spontaneously or by an enzyme with lactonase activity [194, 195]. It is hypothesized 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 biofuels 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 discussed above, the metabolic pathways implied by xylonate have been investigated in the recent 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 Trichoderma 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 pathway 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 research is at its genesis and significant efforts will be required for the creation of a highly productive and effective xylonate production biocatalyst.

At this point it is rather hard to verify the potential or the economic value of oxidation products 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 simple 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 plasticizer 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 difficult to reach economic at an industrial level.

Figure 15.

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

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). Production of ethanol thus requires isolation of cellulose from lignocellulosic matrix, then hydrolysis 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 lignocellulosic 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 technological challenge since it either requires enzymes, ionic liquids or strong acids that are rather expensive 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 technological 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 challenge 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 highly 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 problem 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, biologically as well as chemically into a wide variety of added value products and “green” compounds. 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 furfural 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 polymerisation 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 racemic mixture of D- and L-lactic acid and thus reducing the market value of the product, particularly 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 production 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.

Reduction of xylose can lead to many different products including xylitol for lower severity 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 results. The main problem with the xylitol market is that although it is increasing, it is fairly 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 opportunity for the production of ethylene glycol and propylene glycol, two very important products 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 potential 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 plasticizer. 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.



We would like to acknowledge Enerkem, Greenfield Ethanol, CRB Innovations and the Ministry of Natural Resources of Quebec for financial support of the Industrial Chair in Cellulosic Ethanol.


  1. 1. Lavoie JM, Marie-Rose S, Lynch D. Non-homogeneous residual feedstocks to biofuels and chemicals via the methanol route.Biomass Conversion and Biorefinery 2012; 1-6, ISSN: 2190-6823.
  2. 2. Lavoie JM, Beauchet R, Berberi V, Chornet M. Biorefining Lignocellulosic Biomass via the Feedstock Impregnation Rapid and Sequential Steam Treatment. Biofuel’s engineering process technology.Intech publisher 2011; 685-714, ISBN: 978-953-307-480-1.
  3. 3. Lavoie JM, Chornet M, Chornet E. Biomass refineries: relationships between feedstock and conversion approach. Energy and sustainability II, WIT press 2009; 35-48, DOI:10.2495/ESUS090041.
  4. 4. Schild G, Sixta H. Sulfur-free dissolving pulps and their application for viscose and lyocell.Cellulose 2011;18(4) 1113-1128, DOI:10.1007/s10570-011-9532-0.
  5. 5. Song T, Pranovich A, Bjarne H. Characterisation of Norway spruce hemicelluloses extracted by pressurised hot-water extraction (ASE) in the presence of sodium bicarbonate. Holzforschung 2011;65(1) 35-42, ISSN: 0018-3830.
  6. 6. Song T, Pranovich A, Holmbom B. Effects of pH control with phthalate buffers on hot-water extraction of hemicelluloses from spruce wood. Bioresource technology 2011 ; 102 (22) 10518-10523;ISSN: 0960-8524.
  7. 7. Duarte GV, Ramarao BV, Amidon TE. Polymer induced flocculation and separation of particulates from extracts of lignocellulosic materials. Bioresource technology 2010;101(22) 8526-8534, PMID: 20605092.
  8. 8. Peng F, Ren JL, Xu F, Bian J, Peng P, Sun RC. Comparative study of hemicelluloses obtained by graded ethanol precipitation from sugarcane bagasse. Journal of agricultural and food chemistry 2009;57(14) 6305-6317, DOI:10.1021/ jf900986b.
  9. 9. Liu W, Hou Q, Mao C, Yuan Z, Li K. Hemicelluloses Prior to Aspen Chemithermomechanical Pulping: Pre-Extraction, Separation, and Characterization. Journal of Agricultural and Food Chemistry 2012;60(19) 4880-4885, DOI: 10.1021/jf300787b.
  10. 10. Wang K, Yang H, Guo S, Tang Y, Jiang J, Xu F, Sun RC. Organosolv fractionation process with various catalysts for improving bioconversion of triploid poplar. Process Biochemistry 2012;47(10) 1503-1509, DOI: 10.1016/j.procbio.2012.06.002.
  11. 11. Brosse N, El Hage R, Sannigrahi P, Ragauskas A. Dilute sulphuric acid and ethanol organosolv pretreatment of Miscanthus x Giganteus. Cellulose Chemistry and Technology 2010;44(1-3) 71-78,ISSN:05769787.
  12. 12. Lavoie JM, Capek-Menard E, Gauvin H, Chornet E. Production of pulp from Salix viminalis energy crops using the FIRSST process. Bioresource Technology 2010;101(13) 4940-4946, DOI:10.1016/j.biortech.2009.09.021.
  13. 13. Lavoie JM, Beauchet R, Berberi V, Chornet M. Biorefining quasi homogeneous biomass via the Feedstock Impregnation Rapid and Sequential Steam Treatment. Biofuel’s engineering process technology 2011; Intech publishing 685-715, ISBN:978-953-307-480-1.
  14. 14. Lavoie JM, Beauchet R. Biorefinery of Cannabis sativa using one-and two-step steam treatments for the production of high quality fibres. Industrial Crops and Products 2012;37(1) 275-283, DOI:10.1016/j.indcrop.2011.11.016.
  15. 15. Pasangulapati V, Ramachandriya KD, Kumar A, Wilkins MR, Jones CL, Huhnke RL. Effects of cellulose, hemicellulose and lignin on thermochemical conversion characteristics of the selected biomass. Bioresource Technology. 2012;114 663–669, ISSN: 0960-8524.
  16. 16. Sathitsuksanoh N, Zhu Z, Zhang YHP. Cellulose solvent- and organic solvent-based lignocellulose fractionation enabled efficient sugar release from a variety of lignocellulosic feedstocks. Bioresource Technology 2012;117 228–233, ISSN: 0960-8524
  17. 17. Xu J, Zhang X, Cheng JJ. Pretreatment of corn stover for sugar production with switchgrass-derived black liquor. Bioresource Technology 2012;111 255–260, ISSN: 0960-8524
  18. 18. Jun A, Tschirner UW, Tauer Z. Hemicellulose extraction from aspen chips prior to kraft pulping utilizing kraft white liquor. Biomass and bioenergy 2012;37 229-236, ISSN:09619534.
  19. 19. Rana D, Rana V, Ahring BK. Producing high sugar concentrations from loblolly pine using wet explosion pretreatment. Bioresource Technology 2012;121 61–67, ISSN: 0960-8524
  20. 20. Marcotullio G, de Jong W. Furfural formation from D-xylose: the use of different halides in dilute aqueous acidic solutions allows for exceptionally high yields. Carbohydrate Research 2011;346(11) 1291-3, ISSN:1873-426X.
  21. 21. Nimlos MR, Qian X, Davis M, Himmel ME, Johnson DK. Energetics of xylose decomposition as determined used quantum mechanics. J. Phys. Chem. A.2006;110(42) 11824-11838, ISSN:1089-5639.
  22. 22. Lessard J, Morin JF, Wehrung JF, Magnin D, Chornet E. High Yield Conversion of Residual Pentoses into Furfural via Zeolite Catalysis and Catalytic Hydrogenation of Furfural to 2-Methylfuran. Top Catal. 2010;53(15-18) 1231-1234, ISSN:10225528.
  23. 23. Bugrayev A, Al-Haq N, Okopie RA, Qazi A, Suggate M, Sullivan AC, Wilson JRH. Covalently linked ethylmercaptophenyl sulfonic acid and ethylmercaptobenzyl sulfonic acid silica materials - Synthesis and catalytic activity. Journal of Molecular Catalysis A: Chemical 2008;280(1-2) 96-101, ISSN: 1381-1169.
  24. 24. Tao F, Song H, Chou L. Efficient process for the conversion of xylose to furfural with acidic ionic liquid. Canadian Journal of Chemistry 2011;89(1) 83-87, DOI: 10.1139/V10-153.
  25. 25. Rong C, Ding X, Zhu Y, Li Y, Wang L, Qu Y, Ma X, Wang Z. Production of furfural from xylose at atmospheric pressure by dilute sulfuric acid and inorganic salts.Carbohydrate Research 2012;350(1) 77-80, ISSN:1873-426X.
  26. 26. Lima S, Neves P, Antunes MM, Pillinger M, Ignatyev N, Valente AA. Conversion of mono/di/polysaccharides into furan compounds using 1-alkyl-3-methylimidazolium ionic liquids. Appl. Catal. A: Gen 2009;363(1-2)93-99, DOI: 10.1016/j.apcata.2009.04.049.
  27. 27. Gürbüz EI, Wettstein SG, Dumesic JA. Conversion of Hemicellulose to Furfural and Levulinic Acid using Biphasic Reactors with Alkylphenol Solvents. ChemSusChem 2012;5(2) 383-387, DOI:10.1002/cssc.201100608.
  28. 28. Yang Y, Hu CW, Abu-Omar MM. Synthesis of Furfural from Xylose, Xylan, and Biomass Using AlCl3 6H2O in Biphasic Media via Xylose Isomerization to Xylulose. ChemSusChem 2012;5(2) 405-410, DOI:10.1002/cssc.201100688.
  29. 29. Agirrezabal-Telleria I, Larreategui A, Requies J, Güemez MB, Arias PL. Furfural production from xylose using sulfonic ion-exchange resins (Amberlyst) and simultaneous stripping with nitrogen. Bioresource Technology 2011;102(16) 7478-7485, DOI:10.1016/j.biortech.2011.05.015.
  30. 30. Lima S, Antunes MM, Fernandes A, Pillinger M, Ribeiro MF, Valente AA. Catalytic cyclodehydration of xylose to furfural in the presence of zeolite H-Beta and a micro/mesoporous Beta/TUD-1 composite material. Applied Catalysis, A: General 2010;388(1-2) 141-148, ISSN: 0926-860X.
  31. 31. Antunes MM, Lima S, Fernandes A, Pillinger M, Ribeiro MF,Valente AA. Aqueous-phase dehydration of xylose to furfural in the presence of MCM-22 and ITQ-2 solid acid catalysts. Applied Catalysis, A: General 2012; 417-418 (29) 243-252, DOI:10.1016/j.apcata.2011.12.046.
  32. 32. vom Stein T, Grande PM, Leitner W, Dominguez de Maria P. Iron-Catalyzed Furfural Production in Biobased Biphasic Systems: From Pure Sugars to Direct Use of Crude Xylose Effluents as Feedstock. ChemSusChem 2011;4(11) 1592-1594, DOI:10.1002/cssc.201100259.
  33. 33. Lam E, Majid E, Leung AC, Chong JH, Mahmoud KA, Luong JH. Synthesis of Furfural from Xylose by Heterogeneous and Reusable Nafion Catalysts. ChemSusChem2011;4(4) 535-541, DOI:10.1002/cssc.201100023.
  34. 34. Dias AS, Pillinger M, Valente AA. Liquid phase dehydration of D-xylose in the presence of Keggin-type heteropolyacids. Appl. Catal. A. 2005;285(1-2) 126-131, DOI: 10.1016/j.apcata.2005. 02.016.
  35. 35. Sádaba I, Lima S, Valente AA, López Granados M. Catalytic dehydration of xylose to furfural: vanadyl pyrophosphate as source of active soluble species. Carbohydrate Research 2011;346(17) 2785-2791, DOI:10.1016/j.carres. 2011.10.001.
  36. 36. Win DT. Furfural-Gold from Garbage. AU J.T. 2005;8(4) 185-190, techno/2005/apr05/vol8no4_abstract04.pdf.
  37. 37. Jinan ZZ International Trade Co., Ltd.
  38. 38. Zibo Pulisi Trading Co., Ltd.
  39. 39. Zhou Z, Ma Q, Zhang A, Wu M. Synthesis of water-soluble monotosylated ethylenediamines and their application in ruthenium and iridium-catalyzed transfer hydrogenation of aldehydes. Applied Organometallic Chemistry 2011;25(12)856-861, DOI:10.1002/aoc.1824.
  40. 40. Ajjou AN, Pinet JL. The biphasic transfer hydrogenation of aldehydes and ketones with isopropanol catalyzed by water-soluble rhodium complexes. Journal of Molecular Catalysis A: Chemical 2004;214(2)203-206, ISSN:1381-1169.
  41. 41. Li H, Luo H, Zhuang L, Dai W, Qiao M. Liquid phase hydrogenation of furfural to furfuryl alcohol over the Fe-promoted Ni-B amorphous alloy catalysts. Journal of Molecular Catalysis A: Chemical 2003;203(1-2)267-275, DOI:10.1016/S1381-1169(03)00368-6.
  42. 42. Inada K, Shibagaki M, Nakanishi Y, Matsushita H. The catalytic reduction of aldehydes and ketones with 2-propanol over silica-supported zirconium catalyst. ChemInform 1994;25(15) no pages, DOI: 10.1002/chin.199415090.
  43. 43. Wabnitz T, Breuninger D, Heimann J, Backes R, Pinkos R. Method for one-step synthesis of 2-methyltetrahydrofuran from furfural and catalyst 2009; PCT Int. Appl., WO/ 2009/003882 A1 20090108.
  44. 44. Wabnitz T, Breuninger D, Heimann J, Backes R, Pinkos R. Production of 2-methyltetrahydrofuran from furfural over two catalysts in a structured bed 2009; PCT Int. Appl., WO/2009/003881 A1 20090108.
  45. 45. Lange JP. Process for the hydrogenolysis of furfuryl derivatives PCT Int. Appl.,2009; PCT/EP2009/057899, WO/2009/156439 A1 20091230, (June 2009).
  46. 46. Zheng HY, Zhu YL, Teng BT, Bai ZQ, Zhang CH, Xiang HW, Li YW. Towards understanding the reaction pathway in vapor phase hydrogenation of furfural to 2-methylfuran. Journal of Molecular Catalysis A: Chemical 2006;246(1-2) 18-23, DOI:10.1016/j.molcata.2005.10.003.
  47. 47. TPFTZ Qingshan Int'l Business Technology Co., Ltd.
  48. 48. Zhang W, Zhu Y, Niu S, Li Y. A study of furfural decarbonylation on K-doped Pd/Al2O3 catalysts. Journal of Molecular Catalysis A: Chemical 2011;335(1-2) 71-81, ISSN:1873-314X.
  49. 49. Stevens JG, Bourne RA, Twigg MV, Poliakoff M. Real-Time Product Switching Using a Twin Catalyst System for the Hydrogenation of Furfural in Supercritical CO2. Angewandte Chemie International Edition 2010; 49(47), ISSN:1433-7851.
  50. 50. Popoff T, Theander O. Reactions of D-xylose and D-glucose in alkaline aqueous solutions.Carbohydrate Research1976;48(1) 13-21, DOI:10.1016/S0008-6215(00)83510-7.
  51. 51. Johansson MH, Samuelson O. Reducing end groups in brich xylan and their alkaline degradation. Wood Science and Technology 1977; 11(4) 251-163, ISSN:0043-7719.
  52. 52. El Khadem HS, Ennifar S, Isbell HS. Contribution of the reaction pathways involved in the isomerization of monosaccharides by alkali. Carbohydrate Research 1987;169 13-21, ISSN: 0008-6215.
  53. 53. Jackson JE, Miller DJ, Marincean S. Process for the preparation of lactic acid and glyceric acid from aqueous solutions of hexoses and pentoses in the presence of an excess of a strongly basic anionic exchange resin 2007; U.S. Pat. Appl. Publ., 20070066844.
  54. 54. Epane G, Laguerre JC, Wadouachi A, Marek D. Microwave-assisted conversion of D-glucose into lactic acid under solvent-free conditions. Green Chemistry 2010;12(3) 502-506, DOI:10.1039/B922286C.
  55. 55. Rahubadda A, Montoya A, Haynes BS. C5 sugar decomposition products under hot compressed water conditions [online]. In:CHEMECA 2011: Engineering a Better World: Sydney Hilton Hotel, NSW, Australia, 2011;18-21 Barton, A.C.T.: Engineers Australia, 2011: 698-706. Conference paper. Availability:;dn=168748429919269;res=IELENGISBN: 9780858259676.
  56. 56. Onda A, Ochi T, Kajiyoshi K, Yanagisawa K. A new chemical process for catalytic conversion of D-glucose into lactic acid and gluconic acid. Applied Catalysis, A: General 2008;343(1-2) 49-54, DOI:10.1016/j.apcata.2008.03.017.
  57. 57. Ma C, Jin F,Cao J,Wu B. Hydrothermal Conversion of Carbohydrates into Lactic Acid with Alkaline Catalysts: 4th International Conference onBioinformatics and Biomedical Engineering (iCBBE), June 18-20, 2010, Chengdu, China, 1-4, DOI 10.1109/ICBBE.2010.5516468.
  58. 58. Aspinall GO,Greenwood CT,Sturgeon RJ. The degradation of xylans by alkali. J. Chem. Soc. 1961;3667-3674, DOI 10.1039/JR9610003667.
  59. 59. Yang BY. Montgomery R. Alkaline degradation of fructofuranosides.Carbohydrate Research 1996;280(1) 47-57, DOI: 10.1016/0008-6215(95)00233-2.
  60. 60. Raharja S, Rigal L, Barre L, VidalPF. Design of a continuous process for the alkaline treatment of xylose into lactic acid. The Canadian Journal of Chemical Engineering 1997;75(5)913-920, DOI: 10.1002/cjce.5450750511.
  61. 61. Okano K, Tanaka T, Ogino C, Fukuda H, Kondo A. Biotechnological production of enantiomeric pure lactic acid from renewable resources: Recent achievements, perspectives, and limits. Applied Microbiology and Biotechnology 2010;85(3)413-423, DOI:10.1007/s00253-009-2280-5.
  62. 62. Yadav AK, Chaudhari AB, Kothari RM. Bioconversion of renewable resources into lactic acid: An industrial view. Critical reviews in biotechnology 2011;31(1) 1-19, PMID: 20476870.
  63. 63. Abdel-Rahman MA, Tashiro Y, Sonomoto K. Lactic acid production from lignocellulose-derived sugars using lactic acid bacteria: Overview and limits. Journal of Biotechnology 2011;156(4) 286-301, DOI:10.1016/j.jbiotec. 2011.06.017.
  64. 64. Datta R, Henry M. Lactic acid: Recent advances in products, processes and technologies-A review. Journal of Chemical Technology and Biotechnology 2006;81(7) 1119-1129, DOI: 10.1002/jctb.1486.
  65. 65. Garvie EI. Bacterial lactate dehydrogenases. Microbiological reviews 1980;44(1) 106-139. PMCID: PMC373236
  66. 66. John RP, Nampoothiri KM, Pandey A. Fermentative production of lactic acid from biomass: An overview on process developments and future perspectives. Applied Microbiology and Biotechnology 2007;74(3)524-534, DOI:10.1007/s00253-006-0779-6.
  67. 67. Wee Y, Kim J, Ryu H. Biotechnological production of lactic acid and its recent applications. Food Technology and Biotechnology 2006;44(2) 163-172, ISSN: 1330-9862.
  68. 68. Zhang ZY, Jin B, Kelly JM. Production of lactic acid from renewable materials by Rhizopus fungi. Biochemical engineering journal 2007;35(3) 251-263, ISSN:1369-703X.
  69. 69. Zhao B, Wang L, Li F, Hua D, Ma C, Ma Y, Xu P. Kinetics of D-lactic acid production by Sporolactobacillus sp. strain CASD using repeated batch fermentation. Bioresource technology 2010;101(16) 6499-6505, DOI: 10.1016/j.biortech.2010.03.069.
  70. 70. Zhao B, Wang L, Ma C, Yang C, Xu P, Ma Y. Repeated open fermentative production of optically pure L-lactic acid using a thermophilic Bacillus sp. strain. Bioresource technology 2010;101(16) 6494-6498, DOI: 10.1016/j.biortech.2010.03.051.
  71. 71. Salminen S, Ramos P, Fonden R. Substrates and lactic acid bacteria. Food Science and Technology (New York, NY, United States) 1993;58295-306. ISBN:0-8247-8907-5.
  72. 72. Hang YD. Direct fermentation of corn to L(+)-lactic acid by Rhizopus oryzae. Biotechnology Letters 1989;11(4) 299-300, DOI:10.1007/BF01031581.
  73. 73. Krištofíková L, Rosenberg M, Vlnová A, Šajbidor J, Čertík M. Selection of Rhizopus strains for L(+)-lactic acid and gamma-linolenic acid production. Folia Microbiologica (Prague, Czech Republic) 1991;36(5) 451-455, DOI:10.1007/BF02884065.
  74. 74. Miura S, Arimura T, Itoda N, Dwiarti L, Feng JB, Bin CH, Okabe M. Production of L-lactic acid from corncob. Journal of Bioscience and Bioengineering 2004;97(3) 153-157, ISSN: 1389-1723.
  75. 75. Ding S, Tan T. L-lactic acid production by Lactobacillus casei fermentation using different fed-batch feeding strategies. Process Biochemistry (Amsterdam, Netherlands) 2006;41(6) 1451-1454, ISSN: 1359-5113.
  76. 76. Chang HN, Kim N, Kang J, Jeong CM, Choi J, Fei Q, Kim BJ, Kwon S, Lee SY, Kim J. Multi-stage high cell continuous fermentation for high productivity and titer. Bioprocess and Biosystems Engineering 2011;34(4) 419-431, ISSN 16157591.
  77. 77. Dumbrepatil A, Adsul M, Chaudhari S, Khire J, Gokhale D. Utilization of molasses sugar for lactic acid production by Lactobacillus delbrueckii subsp. delbrueckii mutant Uc-3 in batch fermentation. Applied and Environmental Microbiology 2008;74(1) 333-335, DOI:10.1128/AEM.01595-07.
  78. 78. Bustos G, de la Torre N, Moldes AB, Cruz JM, Dominguez JM. Revalorization of hemicellulosic trimming vine shoots hydrolyzates trough continuous production of lactic acid and biosurfactants by L. pentosus. Journal of Food Engineering 2006;78(2) 405-412, ISBN:0260-8774.
  79. 79. Zhu Y, Lee YY, Elander RT. Conversion of aqueous ammonia-treated corn stover to lactic acid by simultaneous saccharification and cofermentation. Applied Biochemistry and Biotechnology 2007;137-140(1-2) 721-738, ISSN:0273-2289.
  80. 80. Givry S, Prevot V, Duchiron F. Lactic acid production from hemicellulosic hydrolyzate by cells of Lactobacillus bifermentans immobilized in Ca-alginate using response surface methodology. World Journal of Microbiology & Biotechnology 2008;24(6) 745-752, DOI:0.1007/s11274-007-9534-0.
  81. 81. Chaillou S, Bor Y, Batt CA, Postma PW, Pouwels PH. Molecular cloning and functional expression in Lactobacillus plantarum 80 of xylT, encoding the D-xylose-H+ symporter of Lactobacillus brevis. Applied and Environmental Microbiology 1998;64(12) 4720-4728, PMCID:PMC90914.
  82. 82. Helanto M, Kiviharju K, Leisola M, Nyyssölä A. Metabolic engineering of Lactobacillus plantarum for production of L-ribulose. Applied and Environmental Microbiology 2007;73(21) 7083-7091, DOI:10.1128/AEM.01180-07.
  83. 83. Ohara H, Owaki M, Sonomoto K. Xylooligosaccharide fermentation with Leuconostoc lactis. Journal of Bioscience and Bioengineering 2006;101(5) 415-420, ISSN:1389-1723.
  84. 84. Tanaka K, Komiyama A, Sonomoto K, Ishizaki A, Hall SJ, Stanbury PF. Two different pathways for D-xylose metabolism and the effect of xylose concentration on the yield coefficient of L-lactate in mixed-acid fermentation by the lactic acid bacterium Lactococcus lactis IO-1. Applied Microbiology and Biotechnology 2002;60(1-2) 160-167, ISSN:0175-7598.
  85. 85. Abdel-Rahman M, Tashiro Y. Zendo T, Sonomoto K. Effective (+)-Lactic Acid Production by Co-fermentation of Mixed Sugars. Journal of Biotechnology 2010;150(Supplement) 347-348, DOI:10.1016/j.jbiotec.2010.09.384.
  86. 86. Abdel-Rahman MA, Tashiro Y, Zendo T, Hanada K, Shibata K, Sonomoto K. Efficient homofermentative L-(+)-lactic acid production from xylose by a novel lactic acid bacterium, Enterococcus mundtii QU 25. Applied and Environmental Microbiology 2011;77(5) 1892-1895, DOI: 10.1128/AEM.02076-10.
  87. 87. Okano K, Yoshida S, Yamada R, Tanaka T, Ogino C, Fukuda H, Kondo A. Improved production of homo-D-lactic acid via xylose fermentation by introduction of xylose assimilation genes and redirection of the phosphoketolase pathway to the pentose phosphate pathway in L-lactate dehydrogenase gene-deficient Lactobacillus plantarum. Applied and Environmental Microbiology 2009;75(24) 7858-7861, DOI:10.1128/AEM.01692-09.
  88. 88. Tamakawa H, Ikushima S, Yoshida S. Efficient production of L-lactic acid from xylose by a recombinant Candida utilis strain. Journal of Bioscience and Bioengineering 2012;113(1) 73-75, ISSN:1389-1723.
  89. 89. Okano K, Yoshida S, Tanaka T, Ogino C, Fukuda H, Kondo A. Homo-D-lactic acid fermentation from arabinose by redirection of the phosphoketolase pathway to the pentose phosphate pathway in L-lactate dehydrogenase gene-deficient Lactobacillus plantarum. Applied and Environmental Microbiology 2009;75(15)5175-5178, ISSN: 0099-2240.
  90. 90. Ilmen M, Koivuranta K, Ruohonen L, Suominen P, Penttila M. Efficient production of L-lactic acid from xylose by Pichia stipitis. Applied and Environmental Microbiology 2007;73(1)117-123, DOI:10.1128/AEM.01311-06.
  91. 91. Wee Y, Ryu H. Lactic acid production by Lactobacillus sp. RKY2 in a cell-recycle continuous fermentation using lignocellulosic hydrolyzates as inexpensive raw materials. Bioresource technology2009;100(18)4262-4270, ISSN:0960-8524.
  92. 92. John RP, Nampoothiri KM, Pandey A. Simultaneous saccharification and fermentation of cassava bagasse for L-(+)-lactic acid production using Lactobacilli. Applied Biochemistry and Biotechnology 2006;134(3)263-272, ISSN:0273-2289.
  93. 93. Adsul M, Khire J, Bastawde K, Gokhale D. Production of lactic acid from cellobiose and cellotriose by Lactobacillus delbrueckii mutant Uc-3. Applied and Environmental Microbiology 2007;73(15) 5055-5057, DOI:10.1128/AEM.00774-07.
  94. 94. Singhvi M, Joshi D, Adsul M, Varma A, Gokhale D. D-(-)-Lactic acid production from cellobiose and cellulose by Lactobacillus lactis mutant RM2-24. Green Chemistry 2010;12(6)1106-1109, DOI:10.1039/b925975a.
  95. 95. Marques S, Santos JA. L Gírio, FM, Roseiro JC. Lactic acid production from recycled paper sludge by simultaneous saccharification and fermentation. Biochemical engineering journal 2008;41(3)210-216, DOI:10.1016/j.bej.2008.04. 018.
  96. 96. Gullon B, Yáñez R, Alonso JL, Parajó JC. L-Lactic acid production from apple pomace by sequential hydrolysis and fermentation. Bioresource technology 2008;99(2) 308-319, ISSN:0960-8524.
  97. 97. Ishida N, Saitoh S, Onishi T, Tokuhiro K, Nagamori E, Kitamoto K, Takahashi H. The effect of pyruvate decarboxylase gene knockout in Saccharomyces cerevisiae on L-lactic acid production. Bioscience, biotechnology, and biochemistry 2006;70(5)1148-1153, PMID: 16717415.
  98. 98. Tokuhiro K, Ishida N, Nagamori E, Saitoh S, Onishi T, Kondo A, Takahashi H. Double mutation of the PDC1 and ADH1 genes improves lactate production in the yeast Saccharomyces cerevisiae expressing the bovine lactate dehydrogenase gene. Applied Microbiology and Biotechnology 2009;82(5)883-890, ISSN:0175-7598.
  99. 99. Bianchi MM, Brambilla L, Protani F, Liu C, Lievense J, Porro D. Efficient homolactic fermentation by Kluyveromyces lactis strains defective in pyruvate utilization and transformed with the heterologous LDH gene. Applied and Environmental Microbiology 2001;67(12)5621-5625, ISSN: 0099-2240.
  100. 100. Tianjin Tiger International Trade Co., Ltd.
  101. 101. Qingdao Abel Technology Co., Ltd.
  102. 102. PRWeb Online visibility on Vocus, Global Lactic Acid Market to Reach 367.3 Thousand Metric Tons by 2017, According to New Report by Global Industry Analysts, Inc.
  103. 103. Wisniak J, Hershkowitz M, Stein S. Hydrogenation of Xylose to Xylitol, Ind. Eng Chem., Prod. Res. Develop. 1974;13(1)75, DOI:10.1021/i360049a015.
  104. 104. Wisniak J, Hershkowitz M, Stein S. Hydrogenation of Xylose over Platinum Group Catalysts. Ind. Eng. Chem. Prod. Res. Dev. 1974;13(4)232-236,DOI:10.1021/i360052a004.
  105. 105. Mikkola JP, Salmi T. Three-phase catalytic hydrogenation of xylose to xylitol - prolonging the catalyst activity by means of on-line ultrasonic treatment. Catalysis Today 2001;64(3-4)271-277, DOI: 10.1016/S0920-5861(00)00530-7.
  106. 106. Mikkola JP, Kubicka D, Kuusisto J, Granholm N, Salmi T, Holmbom B. Non-traditional three-phase reactor setup for simultaneous acoustic irradiation and hydrogenation. Journal of Chemical Technology and Biotechnology 2003;78(2-3)203-207, DOI:10.1002/jctb.739.
  107. 107. Heikkila H, Ojamo H, Tylli M, Ravanko V, Nurmi J, Haimi P, Alen R, Koivikko H. Preparation of L-xylose and its use for the production of xylitol. U.S. Pat. Appl. Publ., 20030097029, (May 2003).
  108. 108. Kuusisto J, Heikkilae H, Tylli M, Golde M, Riihimaeki T. Catalytic hydrogenation of sugar to sugar alcohol, using increasing reaction temperature and/or addition of monocarboxylic acid to maintain catalyst activity.Brit. UK Pat. Appl. GB 2437517(October2007).
  109. 109. Vanoppen D, Maas-Brunner M, Kammel U, Arndt JD. Preparation and use of silicon dioxide-supported ruthenium catalysts for saccharide hydrogenation, Ger. Offen. DE10128205(December 2002).
  110. 110. Guha SK, Kobayashi H, Hara K, Kikuchi H, Aritsuka T, Fukuoka A. Hydrogenolysis of sugar beet fiber by supported metal catalyst. Catalysis Communications 2011;12(11) 980-983, DOI:10.1016/j.catcom.2011.02.017.
  111. 111. Kwak BS, Lee BI, Kim TY, Kim JW, Lee SI. Process for preparation of sugar alcohols by ruthenium-catalyzed hydrogenation of sugars. PCT Int. Appl., WO 2005021475 A1 20050310 (2005).
  112. 112. Kwak BS, Lee BI, Kim TY, Kim JW, Lee SI. Method for preparing sugar alcohols using ruthenium- and zirconia- catalyzed hydrogenation of monosaccharides. PCT Int. Appl., WO 2006093364 A1 20060908 (2006).
  113. 113. Crabtree SP, Tyers DV. Hydrogenolysis of sugar feedstock. PCT Int. Appl., PCT/GB2004.004391, WO/2005/051874, (June 2005).
  114. 114. Sun J, Liu H. Selective hydrogenolysis of biomass-derived xylitol to ethylene glycol and propylene glycol on supported Ru catalysts. Green Chemistry 2011;13(1)135-142, DOI:10.1039/c0gc00571a.
  115. 115. Chopade SP, Miller DJ, Jackson JE, Werpy TA, Frye Jr JG, Zacher AH. Catalysts and process for hydrogenolysis of sugar alcohols to polyols. US Patent 6291725, (January 2001).
  116. 116. Dubeck M, Knapp GG. Two stage hydrogenolysis of carbohydrate to glycols using sulfide modified ruthenium catalyst in second stage. US Patent 4,476,331, (September 1984).
  117. 117. Holcomb DE. Catalyst and method for production of polyols by hydrogenolysis of carbohydrates. US Patent 7,692,001, (April 2010).
  118. 118. Kasehagen L. Hydrogenolysis of reducible sugars to obtaini a high percentage of glycerol. US Patent 3,396,199, (August 1968).
  119. 119. Wright LW. Hydrogenation and hydrogenolysis of carbohydrates with tungsten oxide promoted supported nickel catalyst. US Patent 3,965,199, (June 1976).
  120. 120. Werpy TA, Frye Jr JG, Zacher AH, Miller DJ. Hydrogenolysis of 6-carbon sugars and other organic compounds. US Patent 6,841,085, (January 2005).
  121. 121. Suzuki S, Sugiyama M, Mihara Y, Hashiguchi K, Yokozeki K. Novel enzymatic method for the production of xylitol from D-arabitol by Gluconobacter oxydans. Bioscience, biotechnology, and biochemistry 2002;66(12)2614-2620, DOI: 10.1271/bbb.66.2614.
  122. 122. Rangaswamy S, Agblevor FA. Screening of facultative anaerobic bacteria utilizing D-xylose for xylitol production. Applied Microbiology and Biotechnology 2002;60(1-2)88-93, ISSN:0175-7598.
  123. 123. Yoshitake J, Ohiwa H, Shimamura M, Imai T. Production of polyalcohol by a Corynebacterium species. I. Production of pentitol from aldopentose. Agricultural and Biological Chemistry 1971;35(6) 905-911.
  124. 124. Yoshitake J, Ishizaki H, Shimamura M, Imai T. Xylitol production by an Enterobacter species. Agricultural and Biological Chemistry 1973;37(10) 2261-2266.
  125. 125. Cirino PC, Chin JW, Ingram LO. Engineering Escherichia coli for xylitol production from glucose-xylose mixtures. Biotechnology and bioengineering 2006;95(6)1167-1176, ISSN:0006-3592.
  126. 126. Parajó JC, Domínguez H, Domínguez JM. Biotechnological production of xylitol. Part 1: Interest of xylitol and fundamentals of its biosynthesis. Bioresource technology 1998;65(3)191-201, DOI:10.1016/S0960-8524(98)00038-8.
  127. 127. Izumori K, Tuzaki K. Production of xylitol from D-xylulose by Mycobacterium smegmatis. Journal of Fermentation Technology 1988;66(1) 33-36, DOI:10.1016/0385-6380(88)90126-4.
  128. 128. Monedero V, Pérez-Martínez G, Yebra MJ. Perspectives of engineering lactic acid bacteria for biotechnological polyol production. Applied Microbiology and Biotechnology 2010;86(4)1003-1015, ISSN:0175-7598.
  129. 129. Nyyssoelae A, Pihlajaniemi A, Palva A, von Weymarn N, Leisola M. Production of xylitol from D-xylose by recombinant Lactococcus lactis. Journal of Biotechnology 2005;118(1)55-66, ISSN:0168-1656.
  130. 130. Fridkin SK, Jarvis WR. Epidemiology of nosocomial fungal infections. Clinical microbiology reviews 1996;9(4)499-511, ISSN: 0893-8512.
  131. 131. Meinander NQ, Hahn-Haegerdal B. Fed-batch xylitol production with two recombinant Saccharomyces cerevisiae strains expressing XYL1 at different levels, using glucose as a cosubstrate: a comparison of production parameters and strain stability. Biotechnology and bioengineering 1997;54(4)391-399, ISSN:0006-3592.
  132. 132. Parajó JC, Domínguez H, Domínguez JM. Biotechnological production of xylitol. Part 2: Operation in culture media made with commercial sugars. Bioresource technology 1998;65(3)203-212, DOI:10.1016/S0960-8524(98)00036-4.
  133. 133. Lee WJ, Ryu YW, Seo JH. Characterization of two-substrate fermentation processes for xylitol production using recombinant Saccharomyces cerevisiae containing xylose reductase gene. Process Biochemistry (Oxford) 2000;35(10)1199-1203, DOI:10.1016/S0032-9592(00)00165-5.
  134. 134. Govinden R, Pillay B, Van Zyl WH, Pillay D. Xylitol production by recombinant Saccharomyces cerevisiae expressing the Pichia stipitis and Candida shehatae XYL1 genes. Applied Microbiology and Biotechnology 2001;55(1)76-80, ISSN:0175-7598.
  135. 135. Sasaki M, Jojima T, Inui M, Yukawa H. Xylitol production by recombinant Corynebacterium glutamicum under oxygen deprivation. Applied Microbiology and Biotechnology 2010;86(4)1057-1066, ISSN:0175-7598.
  136. 136. Akinterinwa O, Khankal R, Cirino PC. Metabolic engineering for bioproduction of sugar alcohols. Current opinion in biotechnology 2008;19(5)461-467, ISSN:0958-1669.
  137. 137. Nigam P, Singh D. Processes for fermentative production of xylitol - a sugar substitute. Process Biochemistry (Oxford) 1995;30(2)117-124, DOI:10.1016/0032-9592(95)80001-8.
  138. 138. Dien BS, Cotta MA, Jeffries TW. Bacteria engineered for fuel ethanol production: current status. Applied Microbiology and Biotechnology 2003;63(3)258-266, ISSN:0175-7598.
  139. 139. Hahn-Hagerdal B, Karhumaa K, Fonseca C, Spencer-Martins I, Gorwa-Grauslund M. Towards industrial pentose-fermenting yeast strains. Applied Microbiology and Biotechnology 2007;74(5)937-953, ISSN:0175-7598.
  140. 140. Khankal R, Chin JW, Cirino PC. Role of xylose transporters in xylitol production from engineered Escherichia coli. Journal of Biotechnology 2008;134(3-4)246-252, ISSN:0168-1656.
  141. 141. Sasaki M, Jojima T, Kawaguchi H, Inui M, Yukawa H. Engineering of pentose transport in Corynebacterium glutamicum to improve simultaneous utilization of mixed sugars. Applied Microbiology and Biotechnology 2009;85(1)105-115, ISSN:0175-7598.
  142. 142. Kinoshita S. Glutamic acid bacteria. In: Demain AL, Solomon NA (eds) Biology of industrial microorganisms. Benjamin/Cummins, London 1985;6 115-142.
  143. 143. Terasawa M, Yukawa H. Industrial production of biochemicals by native immobilization. Bioprocess technology 1993;16, Ind. Appl. Immobilized Biocatal.37-52.
  144. 144. Zhang J, Geng A, Yao C, Lu Y, Li Q. Xylitol production from D-xylose and horticultural waste hemicellulosic hydrolysate by a new isolate of Candida athensensis SB18. Bioresource technology 2012;105134-141, ISSN: 0960-8524.
  145. 145. Lopez F, Delgado OD, Martinez MA, Spencer JFT, Figueroa LIC. Characterization of a new xylitol-producer Candida tropicalis strain. Antonie van Leeuwenhoek 2004;85(4)281-286, ISSN:0003-6072.
  146. 146. Ikeuchi T, Azuma M, Kato J, Ooshima H. Screening of microorganisms for xylitol production and fermentation behavior in high concentrations of xylose. Biomass and Bioenergy 1999;16(5)333-339, DOI:10.1016/S0961-9534(99)00005-7.
  147. 147. Kwon S, Park S, Oh D. Increase of xylitol productivity by cell-recycle fermentation of Candida tropicalis using submerged membrane bioreactor. Journal of Bioscience and Bioengineering 2006;101(1)13-18, ISSN:1389-1723.
  148. 148. Oh DK, Kim SY. Increase of xylitol yield by feeding xylose and glucose in Candida tropicalis. Applied Microbiology and Biotechnology 1998;50(4)419-425, ISSN:0175-7598.
  149. 149. Ojamo H. Yeast xylose metabolism and xylitol production. VTT Publications 1994;176, ISBN:9513844145.
  150. 150. Choi J, Moon K, Ryu Y, Seo J. Production of xylitol in cell recycle fermentations of Candida tropicalis. Biotechnology Letters 2000;22(20)1625-1628, DOI:10.1023/A:1005693427389.
  151. 151. Granström TB. Biotechnological production of xylitol with Candida yeasts. Aalto University, Library Publication Archive, 160, ISBN:951-22-5997-4.
  152. 152. Hallborn J, Walfridsson M, Airaksinen U, Ojamo H, Hahn-Hagerdal B, Penttila M, Kerasnen S. Xylitol production by recombinant Saccharomyces cerevisiae. Biotechnology (Nature Publishing Company) 1991;9(11)1090-1095, PMID 1367625
  153. 153. Salgado JM, Rodríguez N, Cortés S, Domínguez JM. Effect of nutrient supplementation of crude or detoxified concentrated distilled grape marc hemicellulosic hydrolysates on the xylitol production by Debaryomyces hansenii. Preparative biochemistry & biotechnology 2012;42(1) 1-14, ISSN: 1082-6068.
  154. 154. Oh E, Bae Y, Kim K, Park Y, Seo J. Effects of overexpression of acetaldehyde dehydrogenase 6 and acetyl-CoA synthetase 1 on xylitol production in recombinant Saccharomyces cerevisiae. Biocatalysis and Agricultural Biotechnology 2012;1(1)15-19, DOI:10.1016/j.bcab.2011.08.011.
  155. 155. Li M, Meng X, Diao E, Du F. Xylitol production by Candida tropicalis from corn cob hemicellulose hydrolysate in a two-stage fed-batch fermentation process. Journal of Chemical Technology and Biotechnology 2012;87(3)387-392, DOI:10.1002/jctb.2732.
  156. 156. Jeon YJ, Shin HS, Rogers PL. Xylitol production from a mutant strain of Candida tropicalis. Letters in applied microbiology 2011;53(1)106-113, ISSN:0266-8254.
  157. 157. Huang C, Jiang Y, Guo G, Hwang W. Development of a yeast strain for xylitol production without hydrolysate detoxification as part of the integration of co-product generation within the lignocellulosic ethanol process. Bioresource technology 2012;102(3)3322-3329, ISSN:0960-8524.
  158. 158. Barbosa MFS, de Medeiros MB, de Mancilha IM, Schneider H, Lee H. Screening of yeasts for production of xylitol from D-xylose and some factors which affect xylitol yield in Candida guilliermondii. Journal of industrial microbiology 1988;3(4)241-251, DOI:10.1007/BF01569582.
  159. 159. Roseiro JC, Peito MA, Gírio FM, Amaral-Collaco MT. The effects of the oxygen transfer coefficient and substrate concentration on the xylose fermentation by Debaryomyces hansenii. Archives of Microbiology 1991;156(6) 484-490, DOI:10.1007/BF00245396.
  160. 160. Ligthelm ME, Prior BA, du Preez JC. The oxygen requirements of yeasts for the fermentation of D-xylose and D-glucose to ethanol. Applied Microbiology and Biotechnology 1988;28(1)63-68, ISSN:0175-7598.
  161. 161. Debus D, Methner H, Schulze D, Dellweg H. Fermentation of xylose with the yeast Pachysolen tannophilus. European Journal of Applied Microbiology and Biotechnology 1983;17(5) 287-291, DOI:10.1007/BF00508022.
  162. 162. Kötter P, Ciriacy M. Xylose fermentation by Saccharomyces cerevisiae. Applied Microbiology and Biotechnology 1993;38(6)776-783, DOI:10.1007/BF00167144.
  163. 163. Hallborn J, Gorwa MF, Meinander N, Penttilä M, Keränen S, Hahn-Haegerdal B. The influence of cosubstrate and aeration on xylitol formation by recombinant Saccharomyces cerevisiae expressing the XYL1 gene. Applied Microbiology and Biotechnology 1994;42(2-3)326-333, ISSN: 0175-7598.
  164. 164. Sonderegger M, Jeppsson M, Hahn-Hagerdal B, Sauer U. Molecular Basis for Anaerobic Growth of Saccharomyces cerevisiae on Xylose, Investigated by Global Gene Expression and Metabolic Flux Analysis. Applied and Environmental Microbiology 2004;70(4)2307-2317, DOI:10.1128/AEM.70.4.2307-2317.
  165. 165. Ahmad I, Shim WY, Jeon WY, Yoon BH, Kim J. Enhancement of xylitol production in Candida tropicalis by co-expression of two genes involved in pentose phosphate pathway. Bioprocess and Biosystems Engineering 2012;35(1-2)199-204, ISSN:1615-7591.
  166. 166. Shanghai Yanda Biotechnology Co., Ltd.
  167. 167. Wuhan Xuyadi Chemicals Co., Ltd.
  168. 168. Tianjin Yuanlong Chemical Industry Co., Ltd.
  169. 169.
  170. 170.
  171. 171. Bonrath W, Fischesser J. Novel reaction with a gold catalyst. WO2008148549 (December 2008).
  172. 172. Pruesse U, Heidkamp K, Decker N, Herrmann M, Vorlop KD. Processing of raw materials from renewable resources by selective oxidation with gold catalysts. Chemie Ingenieur Technik 2010;82(8) 1231-1237.
  173. 173. Van der Weijden RD, Mahabir J, Abbadi A. Reuter, MA. Copper recovery from copper (II) sulfate solutions by reduction with carbohydrates. Hydrometallurgy 2002;64(2)131-146, DOI:10.1016/S0304-386X(02)00031-2.
  174. 174. Ericsson T, Samuelson O. Treatment of Birch Xylan with Chlorine in Aqueous Solution. Acta Chemica Seandinavica В,Summary citation from AGRICOLA, the online catalog of the National Agricultural Library (NAL)1975;29(3) 309-314, DOI: 10.3891/acta.chem.scand.29b-0309.
  175. 175. Jokic A, Ristic N, Jaksic MM, Spasojevic M, Krstajic N. Simultaneous electrolytic production of xylitol and xylonic acid from xylose. Journal of applied Electrochemistry 1991;21(4) 321-326, ISSN: 0021-891X.
  176. 176. Kiely DE, Hash KR. Process for the oxidization of monosaccharides to glycosyl carboxylic acids using oxygen and nitric acid as co-oxidants. U.S. Pat. Appl. Publ, US 20080033205(February 2008).
  177. 177. Venema FR, Peters JA, van Bekkum H. Platinum-catalyzed oxidation of aldopentoses to aldaric acids. Journal of Molecular Catalysis 1992;77(1)75-85; DOI:10.1016/0304-5102(92)80186-K.
  178. 178. Boussie TR, Dias EL, Fresco ZM, Murphy VJ. Production of glutaric acid and derivatives from carbohydrate-containing materials. PCT Int. Appl., AU-A-2010259935, WO 2010/144871, (December 2010).
  179. 179. Fleche G. Process for the manufacture of xylaric acid and uses thereof US Patent 5731467, (March 1998).
  180. 180. Isbell HS, Frush HL, Martin ET. Reactions of carbohydrates with hydroperoxides: Part I. Oxidation of aldoses with sodium peroxide. Carbohydrate Research 1973;26(2)287-295, DOI:10.1016/S0008-6215(00)84515-2.
  181. 181. Pullanikat P, Jung SJ, Yoo KS, Jung KW. Oxidative degradation of reducing carbohydrates to ammonium formate with H2O2 and NH4OH. Tetrahedron Letters 2010;51(47)6192-6194, ISSN:0040-4039.
  182. 182. Wölfel R,Taccardi N,Bösmann A,Wasserscheid P. Selective catalytic conversion of biobased carbohydrates to formic acid using molecular oxygen. Green Chemistry 2011;13(10)2759-2763, DOI:10.1039/C1GC15434F.
  183. 183. Bernhauer K, Riedl-Tumova E. Oxidations by acetic acid bacteria. X. Oxidation of enantiomorphic forms of xylose and arabinose. Biochemische Zeitschrift 1950;321 26-30.
  184. 184. Ishizaki H, Ihara T, Yoshitake J, Shimamura M, Imai T. D-Xylonic acid production by Enterobacter cloacae. Nippon Nogei Kagaku Kaishi 1973;47(12) 755-761, DOI: 10.1271/nogeikagaku1924.47.755.
  185. 185. Buchert J, Viikari L, Linko M, Markkanen P. Production of xylonic acid by Pseudomonas fragi. Biotechnology Letters 1986;8(8) 541-546, DOI:10.1007/BF01028079.
  186. 186. Uchida K, Suzuki Y. Physiology of plant pathogens. 3. Production of D-galactonic acid, 2-keto-D-galactonic acid, L-arabonic acid, and D-xylonic acid by Erwinia milletiae. Nippon Nogei Kagaku Kaishi 1975;49(5) 257-262.
  187. 187. Hayasida A. Demonstration of pentose oxidase and catalase in fusaria. Biochemische Zeitschrift 1938;298169-178.
  188. 188. Ohsugi M, Tochikura T, Ogata K. Production of D-xylonic acid by Micrococcus species. Agricultural and Biological Chemistry 1970;34(3)357-363.
  189. 189. Lockwood LB, Nelson GEN. The oxidation of pentoses by Pseudomonas. Journal of Bacteriology1946;52(5)581-586.
  190. 190. Weimberg R. Pentose oxidation by Pseudomonas fragi. Journal of Biological Chemistry 1961;236(3)629-635, PMID 13783864.
  191. 191. Kiessling H, Lindberg B, McKay J. Some products of the metabolism of D-xylose by Pullularia pullulans. Acta chemica Scandinavica 1962;16 1858-1862, DOI: 10.3891/acta.chem.scand.16-1858.
  192. 192. Buchert J, Puls J, Poutanen K. Comparison of Pseudomonas fragi and Gluconobacter oxydans for production of xylonic acid from hemicellulose hydrolyzates. Applied Microbiology and Biotechnology 1988;28(4-5)367-372, DOI:10.1007/BF00268197.
  193. 193. Stephens C, Christen B, Fuchs T, Sundaram V, Watanabe K, Jenal U. Genetic analysis of a novel pathway for D-xylose metabolism in Caulobacter crescentus. Journal of Bacteriology 2007;189(5)2181-2185, ISSN: 0021-9193.
  194. 194. Yamanaka K, Gino M, Kaneda R. A specific NAD-D-xylose dehydrogenase from Arthrobacter sp. Agricultural and Biological Chemistry 1977;41(8)1493-1499, ISSN:0002-1369.
  195. 195. Nygard Y, Toivari MH, Penttila M, Ruohonen L, Wiebe MG. Bioconversion of d-xylose to d-xylonate with Kluyveromyces lactis. Metabolic engineering 2011;13(4)383-391, ISSN:1096-7176.
  196. 196. Meijnen J, de Winde JH, Ruijssenaars HJ. Establishment of oxidative D-xylose metabolism in Pseudomonas putida S12. Applied and Environmental Microbiology 2009;75(9)2784-2791, ISSN: 0099-2240.
  197. 197. Novick NJ, Tyler ME. (1982). L-Arabinose metabolism in Azospirillum brasiliense. Journal of Bacteriology 1982;149(1)364-367, PMCID PMC216631.
  198. 198. Toivari MH, Ruohonen L, Richard P, Penttilae M, Wiebe MG. Saccharomyces cerevisiae engineered to produce D-xylonate. Applied Microbiology and Biotechnology 2010;88(3)751-760, ISSN:0175-7598.
  199. 199. Liu H, Valdehuesa KNG, Nisola GM, Ramos KRM, Chung W. High yield production of D-xylonic acid from D-xylose using engineered Escherichia coli. Bioresource technology 2012;115244-248, ISSN:0960-8524.
  200. 200. Henan Premtec Enterprise Corporation
  201. 201.

Written By

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

Submitted: August 18th, 2012 Published: March 20th, 2013