Value added products from hemicellulose: Biotechnological...
Transcript of Value added products from hemicellulose: Biotechnological...
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Value added products from hemicellulose: Biotechnological perspective Vishnu Menon, Gyan Prakash and Mala Rao*
Division of Biochemical Sciences, National Chemical Laboratory, Pune 411008, India
*Corresponding Author
Dr. Mala Rao, [email protected] , Phone : +91-20-25902228
Fax No: +91-20-25902648
Division of Biochemical Sciences
National Chemical Laboratory
Pune - 411 008,
India.
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Abstract Lignocellulosic biomass represents a renewable, widespread and low cost source of
sugars which ca be used as substrate for the production of specialilty chemicals. Various
agricultural residues contain about 20-30% hemicellulose, the second most abundant
biopolymer found in nature. They are heterogeneous polymer of pentoses (xylose and
arabinose) hexoses (mannose, glucose and galactose) and sugar acids. Xylans is the major
component of hemicellulose and are heteropolysaccharides with homopolymeric
backbone chains of 1, 4 linked β-D-xylopyranose units. The conversion of hemicellulose
to bioethanol and other value added products is challenging problem.and is foreseen to be
essential for the economic success of lignocellulosic ethanol. The xylose from the
hemicellulose hydrolysate can be fermented to ethanol/ xylitol/lactic acid and other
valuable products by pentose fermenting microbes. Xylitol a non caloric sugar has
important applications in pharmaceuticals and food industries due to high sweetening
power, non- and anticariogenicity property. . The acid or enzymatic hydrolysis of
hemicellulose to fermentable sugars will be outlined. The enzymatic and fermentation
routes for the conversion of hemicellulose into useful products will be described.
Keywords
Hemicellulose, Hemicellulases, Hydrolysis, Fermentation route, Biochemicals,
Recombinant approaches
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1 Introduction
In collation to first generation bioethanol from starch and molasses, the
development of second generation bioethanol from lignocellulosic biomass serves many
advantages from both energy and environmental concerns. Efficient conversion of
lignocellulosic materials is a global challenge for eco-friendly value addition. Cellulose is
the primary constituent of lignocellulosic biomass and is the most abundant organic
compound available. The estimated global annual production of biomass is 1×1011 tons
[1] and is considered to be one of the significant renewable resources for the production
of biofuel and value added products. Unlike fossil fuels, cellulosic ethanol produced
through fermentation of sugars is a renewable energy source. Biomass in general consists
of 40-50% cellulose, 25 to 30% hemicellulose and 15 to 20% lignin. The effective
utilization of all the three components would play a significant role in economic viability
of the cellulose to ethanol process [2].
In Indian scenario, bagasse and starchy biomass are expected to be primary
feedstocks in preference to other materials such as lipids. The setting up of biorefinary
requires that technologies be developed for biomass pretreatment, hydrolysis of cellulose
and hemicellulose and creation of microbes by recombinant DNA techniques for the
production of desired biochemicals and further their strain improvement. The biorefinary
concept is being built on two different platforms by National Renewable Energy Lab
(NREL) to promote different product areas. The sugar platform based on biochemical
conversion process focuses on the fermentation of sugar extracted from biomass
feedstocks and the other platform is based on thermo chemical conversion processes and
focuses on the gasification of biomass feedstocks and byproducts from the conversion
process (www.nrel.gov/biomass/biorefinery.html). Biorefineries using fibrous
lignocellulosic biomass as feed stock will produce C6 and C5 sugars (predominantly
xylose and arabinose) and lignin.
Various agricultural residues contain about 20-30% hemicellulose, the second
most abundant biopolymer found in nature. They are heterogeneous polymer of pentoses,
hexoses and sugar acids. In recent years, bioconversion of hemicellulose has received
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much attention because of its practical applications in various agro-industrial processes,
for the production of fuels and chemicals, delignification of paper pulp, digestibility
enhancement of animal feedstock, clarification of juices, and improvement in the
consistency of beer [3]. The present review is a comprehensive state-of-the-art describing
the acid and enzymatic hydrolysis of hemicellulose to fermentable sugars. The article
also covers the recent developments in the biotechnological and recombinant aspects for
the fermentation of hemicellulosic hydrolysates to value added products.
2 Structures of Hemicelluloses Hemicelluloses are heterogeneous polysaccharides, which are located between the
lignin and cellulose fibres and, depending on wood species, constitute about 20-30% of
the naturally occurring lignocellulosic plant biomass [4]. They are composed of both
linear and branched heteropolymers of D-xylose, L-arabinose, D-mannose, D-glucose, D-
galactose, L-rhamnose, L-fucose and D-glucuronic acid, which may be acetylated or
methylated. Most hemicelluloses contain two to six of these sugars [5]. They are usually
classified according to the main sugar residues in the backbone. Homopolymers of
xylose, so-called homoxylans only occur in seaweeds (red and green algae). The two
main hemicelluloses in wood are xylans and glucomannans both of which are present in
softwood, whereas in hardwood, xylan is the major component. Xylan is the second most
abundant polysaccharide, comprising up to 30% of the cell wall material of annual plants,
15-30% of hardwoods and 7-10% of softwoods [6]. Xylan occurs as a
heteropolysaccharide with a homopolymeric backbone chain of 1,4-linked β-D-
xylopyranose units, which consists of O-acetyl, α-L-arabinofuranosyl, α-1, 2-linked
glucuronic or 4-O-methylglucuronic acid substituents. In hardwoods, xylan exists as O-
acetyl- 4-O-methylglucuronoxylan with a higher degree of polymerization (150-200) than
softwoods (70-130), which occurs, as arabino-4-O-methylglucuronoxylan.
Approximately one in ten of the β-D-xylopyranose backbone units of hardwood xylan are
substituted at C-2 position with 1,2-linked 4-O-methyl-α-D-glucuronic acid residue,
whereas 70% are acetylated at C-2 or C-3 positions or both [4]. Softwood xylans are not
acetylated but the 4-O-methylglucuronic acid and the β-arabinofuranose residues are
attached to the C-2 and C-3 positions, respectively of the relevant xylopyranose backbone
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units [7]. Birch wood (Roth) xylan contains 89.3% xylose, 1% arabinose, 1.4% glucose,
and 8.3% anhydrouronic acid [8]. Rice bran neutral xylan contains 46% xylose, 44.9%
arabinose, 6.1% galactose, 1.9% glucose, and 1.1% anhydrouronic acid [9]. Wheat
arabinoxylan contains 65.8% xylose, 33.5% arabinose, 0.1% mannose, 0.1% galactose,
and 0.3% glucose [10]. Corn fiber xylan is one of the complex heteroxylans containing β-
(1,4)-linked xylose residues [11]. It contains 48–54% xylose, 33–35% arabinose, 5–11%
galactose and 3-6% glucuronic acid [12]. The xylan of grasses occurs as arabino-4-O-
methylglucuronoxylan with a degree of polymerisation of 70 and contains β-
arabinofuranosyl side chains linked to C-2 or C-3 positions, or both, of the β-D-
xylopyranose main chain residues. The 1,2-linked 4-O-methyl-α-D-glucuronic acid is
found less frequently in grasses than in hardwood xylan. Hardwood xylan contains 2-5%
by weight of O-acetyl groups linked to C-2 or C-3 positions of the xylopyranose units,
and 6% of the arabinosyl side chains are themselves substituted at position 5 with
feruloyl groups, whereas 3% are substituted with p-coumaroyl residues [13]. The relative
proportions of the various components of grass arabinoxylan vary from species to species
and from tissue to tissue within a single species. The main mannan group in the cell walls
of higher plants is galacto-glucomannan with the backbone composed of 1, 4-β-linked D-
glucose and D-mannose units that are distributed randomly within the molecule.
Softwood glucomannan has a backbone composed of β-1, 4- linked D-glucopyranose and
D-mannopyranose units, which are partially substituted by α-galactose units and acetyl
groups. Hardwoods contain a small amount of glucomannan (2-5%) that has no galactose
or acetic acid side groups present [14]. Table 1 describes the composition of
representative hemicellulosic feedstocks. Xylans are usually available in large amounts as
by-products of forest, agriculture, agro-industries, wood and pulp and paper industries.
2.1 Arabinoxylans
Arabinoxylans represent the major hemicellulose structures of the cereal grain cell
walls and are similar to hardwoods xylan but the amount of L-arabinose is higher. The
linear β-(1,4)-D-xylopyranose backbone is substituted by α-L-arabinofuranosyl units in
the positions 2-O and/or 3-O and by α-D-glucopyranosyl uronic unit or its 4-O-methyl
derivative in the position 2-O [40, 41]. O-acetyl substituents may also occur [42,43].
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Arabinofuranosyl residues of arabinoxylan may also be esterified with hydroxycinnamic
acids residues, e.g., ferulic and p-coumaric acids [43]. Dimerization of esterified phenolic
compounds may also lead to inter- and intra-molecular cross-links of xylan. The physical
and/or covalent interactions with other cellwall constituents, restricts the extractability of
xylan. In lignified tissues for example, xylan is ester linked through its uronic acid side
chains to lignin [44].
2.2 Glucuronoxylans
Glucuronoxylans (O-acetyl-4-O-methylglucuronoxylan) are the main
hemicellulose of hardwoods, which may also contain small amounts of glucomannans
(GM). In hardwoods, it represent 15– 30% of their dry mass and consist of a linear
backbone of β-D-xylopyranosyl units (xylp) linked by β-(1,4) glycosidic bonds. Some
xylose units are acetylated at C2 and C3 and one in ten molecules has an uronic acid
group (4-O-methylglucuronic acid) attached by α-(1,2) linkages. The percentage of acetyl
groups ranges between 8% and 17% of total xylan, corresponding, in average, to 3.5–7
acetyl groups per 10 xylose units [45]. The 4- O-methylglucuronic side groups are more
resistant to acids than the Xylp and acetyl groups. The average degree of polymerization
(DP) of glucuronoxylan is in the range of 100–200 [46]. Besides these main structural
units, it may also contain small amounts of L-rhamnose and galacturonic acid. The later
increases the polymer resistance to alkaline agents.
2.3 Arabinoglucuronoxylans
Arabinoglucuronoxylans (arabino-4-O-metylglucuronoxylans) are the major
components of non-woody materials (e.g., agricultural crops) and a minor component for
softwoods (5–10% of dry mass). They also consist of a linear β-(1,4)-D-xylopyranose
backbone containing 4-O-metil-α-D-glucopiranosyl uronic acid and α- L-
arabinofuranosyl linked by α-(1,2) and α-(1,3) glycosidic bonds [47,48]. The typical ratio
arabinose: glucuronic acid:xylose is 1:2:8 [45]. Conversely to hardwood xylan,
arabinoglucuronoxylan might be less acetylated, but may contain low amounts of
galacturonic acid and rhamnose. The average DP of AGX ranges between 50 and 185
[46].
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2.4 Xyloglucans
Xyloglucan is a predominant hemicellulose found in the primary cell wall of
dicots and several monocots ([49], consists of a β-(1→ 4) - linked β-D-Glcp (β-D-
glucopyranose) backbone having up to 75% of the β-D-Glcp residues covalently linked to
α-D-Xylp (α-D-xylopyranose) at the O-6 position. While the extent of xylose substitution
varies considerably, the glucose backbone structure is typically substituted by 50% or
75% xylose [50]. Xylose can be further substituted by galactose or arabinose and the
galactose may be fucosylated. The glucose, xylose and galactose residues are present in
the ratio of 2.8: 2.25:1.0. In addition, xyloglucans can contain O-linked acetyl groups
[51, 52]. Xyloglucans interact with cellulose microfibrils by the formation of hydrogen
bonds, thus contributing to the structural integrity of the cellulose network [49, 53]. XG
is also a storage polymer in the seed endosperm of many plants. Tamarind kernel powder
(TKP) a rich source of hemicellulose, galactoxyloglucan (GXG) is a crude extract
derived from the seeds of Tamarindus indica Linn., an abundantly grown tree of
Southeast Asia, India, Mexico, and Costa Rica [54].
2.5 Galactoglucomannans
Galactoglucomannans (O-acetyl-galactoglucomannans) are the main
hemicelluloses of softwoods, accounting up to 20–25% of their dry mass [46]. It consist
of a linear backbone of β-D-glucopyranosyl and β-D-mannopyranosyl units, linked by β-
(1,4) glycosidic bonds, partially acetylated at C2 or C3 and substituted by a-D-
galactopyranosyl units attached to glucose and mannose by α-(1,6) bonds. Some are
water soluble, presenting in that case higher galactose content than the insolubles [55].
Acetyl group’s content of galactoglucomannan is around 6%, corresponding, to 1 acetyl
group per 3–4 hexoses units. Glucomannans occur in minor amounts in the secondary
wall of hardwoods (<5% of the dry wood mass) [45]. They have linear backbone of β-D-
glucopyranosyl (Glcp) and β-D-mannopyranosyl (Manp) units but the ratio Glcp:Manp is
lower. The extent of galactosylation governs the association of galactoglucomannan and
glucomannans to the cellulose microfibrils and hence, their extractability from the cell
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wall matrix [44]. The average DP of galactoglucomannans ranges between 40 and 100
[46].
2.6 Complex heteroxylans
Complex heteroxylans are present in cereals, seeds, gum exudates and lucilages
and they are structurally more complex [56]. The β-(1,4)-D-xylopyranose backbone is
substituted with single uronic acid and arabinosyl residues and also with various mono-
and oligoglycosyl side chains.
3 Separation and saccharification of hemicellulose Without a profitable use of the hemicellulose fraction, bioethanol is too expensive
to compete in commercial markets [57]. Therefore to foster the commercial production of
lignocellulosic ethanol, bioconversion of the hemicelluloses into fermentable sugars is
essential. The initial step for the production of bioethanol is pre-treatment. The
hemicelluloses are commonly removed during the initial stage of biomass processing
aiming to reduce the structural complexity for further enzymatic cellulose hydrolysis.
Various pretreatment processes are available to fractionate, solubilize, hydrolyse and
separate cellulose, hemicellulose and lignin. These include concentrated acid [58], dilute
acid [11], alkaline [59], SO2 [60], hydrogen peroxide [61], steam explosion
(autohydrolysis) [62], ammonia fiber explosion (AFEX) [63], wet-oxidation [64], lime
[65], liquid hot water [66], CO2 explosion [67], organic solvent treatments [68], ionic
liquids [69, 70] and supercritical fluids [71]. The main processes for the selective
separation of hemicelluloses from biomass include the use of acids, water (liquid or
steam), organic solvents and alkaline agents. The later two are not selective towards
hemicellulose as they also remove lignin, which in turn can hinder the valorisation
process, e.g., fermentation or bioconversion, as the lignin-derived compounds are usually
microbial growth inhibitors. Therefore, acid/water/steam pretreatments are the most
commonly applied technologies yielding a selectively solubilisation of hemicelluloses
and producing hemicellulose-rich liquids totally or partially hydrolysed to oligomeric and
monomeric sugars and cellulose-enriched solids for further bioprocessing [72].
Depending on the operational conditions degradation products are also formed, both from
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sugars (furan and its derivates and weak acids) and, to a less extent, from lignin
(phenolics) [73]. These compounds may also inhibit the fermentation processes, leading
to lower ethanol yields and productivities; however a prior detoxification step is essential.
The following section describes few selected pretreatments wherein most of the
hemicellulose is obtained in the form of oligosaccharides and monomeric sugars.
3.1 Acid hydrolysis
3.1.1 Dilute acid hydrolysis
The dilute acid hydrolysis of the biomass is, by far, the oldest technology for
converting the biomass to ethanol. The first attempt at commercializing a process for the
ethanol from the wood was carried out in Germany in 1898. It involved the use of dilute
acid to hydrolyze the cellulose to glucose, and was able to produce 7.6 liters of ethanol
per 100 kg of wood waste (18 gal per ton). The hydrolysis occurs in two stages to
accommodate the differences between the hemicellulose and the cellulose [74] and to
maximize the sugar yields from the hemicellulose and cellulose fractions of the biomass.
The first stage is operated under milder conditions to hydrolyze the hemicellulose, while
the second stage is optimized to hydrolyze the more resistant cellulose fraction. The
liquid hydrolyzates are recovered from each stage, neutralized, and fermented to ethanol.
National Renewable Energy Laboratory (NREL) a facility of US Department of
energy (DOE) operated by Midwest Reseach institute, Bettelle, NREL outlined a process
where the hydrolysis is carried out in two stages to accommodate the differences between
hemicellulose and cellulose. NREL has reported the results for a dilute acid hydrolysis of
softwoods in which the conditions of the reactors were as follows: Stage 1: 0.7% sulfuric
acid, 190°C, and a 3-minute residence time; Stage 2: 0.4% sulfuric acid, 215°C, and a 3-
minute residence time. The liquid hydrolyzates are recovered from each stage and
fermented to alcohol. Residual cellulose and lignin left over in the solids from the
hydrolysis reactors serve as boiler fuel for electricity or steam production. These bench-
scale tests confirmed the potential to achieve yields of 89% for mannose, 82% for
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galactose and 50% for glucose. The fermentation with Saccharomyces cerevisiae
achieved ethanol conversion of 90% of the theoretical yield [75].
Typical sulphuric acid concentrations for hemicellulose hydrolysis are in the
range 0.5–1.5% and temperatures above 121–160°C. From hemicelluloses, dilute-acid
processes yield sugar recoveries from 70% up to >95% [76-79]. The advantages of dilute-
acid hydrolysis are the relatively low acid consumption, limited problem associated with
equipment corrosion and less energy demanding for acid recovery. Under controlled
conditions, the levels of the degradation compounds generated can also be reduced [72].
3.1.2 Concentrated Acid Hydrolysis
This process is based on concentrated acid decrystallization of the cellulose
followed by the dilute acid hydrolysis to sugars at near theoretical yields. The separation
of acid from the sugars, acid recovery, and acid re-concentration are critical unit
operations. The fermentation converts sugars to ethanol. A process was developed in
Japan in which the concentrated sulfuric acid was used for the hydrolysis and the process
was commercialized in 1948. The remarkable feature of their process was the use of
membranes to separate the sugar and acid in the product stream. The membrane
separation, a technology that was way ahead of its time, achieved 80% recovery of acid
[80]. The concentrated sulfuric acid process has also been commercialized in the former
Soviet Union. However, these processes were only successful during the times of the
national crisis, when economic competitiveness of the ethanol production could be
ignored. Concentrated hydrochloric acid has also been utilized and in this case, the
prehydrolysis and hydrolysis are carried out in one step. Generally, acid hydrolysis
procedures give rise to a broad range of compounds in the resulting hydrolysate, some of
which might negatively influence the subsequent steps in the process. A weak acid
hydrolysis process is often combined with a weak acid prehydrolysis.
In 1937, the Germans built and operated commercial concentrated acid hydrolysis
plants based on the use and recovery of hydrochloric acid. Several such facilities were
successfully operated. During World War II, researchers at USDA's Northern Regional
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Research Laboratory in Peoria, Illinois further refined the concentrated sulfuric acid
process for corncobs. They conducted process development studies on a continuous
process that produced a 15%-20% xylose sugar stream and a 10%-12% glucose sugar
stream, with the lignin residue remaining as a byproduct. The glucose was readily
fermented to ethanol at 85%-90% of theoretical yield. The Japanese developed a
concentrated sulfuric acid process that was commercialized in 1948. The remarkable
feature of their process was the use of membranes to separate the sugar and acid in the
product stream. The membrane separation, a technology that was way ahead of its time,
achieved 80% recovery of acid [80]. R&D based on the concentrated sulfuric acid
process studied by USDA (and which came to be known as the "Peoria Process") picked
up again in the United States in the 1980s, particularly at Purdue University and at
Tennessee Valley Authority (TVA) [81]. Among the improvements added by these
researchers were 1) recycling of dilute acid from the hydrolysis step for pretreatment, and
2) improved recycling of sulfuric acid. Minimizing the use of sulfuric acid and recycling
the acid cost-effectively are critical factors in the economic feasibility.
(http://www1.eere.energy.gov/biomass/printable_versions/concentrated_acid.html). The
conventional wisdom in the literature suggests that the Peoria and TVA processes cannot
be economical because of the high volumes of acid required [82]. The improvements in
the acid sugar separation and recovery have opened the door for the commercial
application. Two companies namely Arkenol and Masada in the United States are
currently working with DOE and NREL to commercialize this technology by taking
advantage of niche opportunities involving the use of biomass as a means of mitigating
waste disposal or other environmental problems
(http://www1.eere.energy.gov/biomass/concentrated_acid.html). Minimizing the use of
the sulfuric acid and recycling the acid cost-effectively are the critical factors in the
economic feasibility of the process. U.S.Pat. No. 5,366,558 [83] uses two "stages" to
hydrolyze the hemicellulose sugars and the cellulosic sugars in a countercurrent process
using a batch reactor, and results in poor yields of glucose and xylose using a mineral
acid. Further, the process scheme is complicated and the economic potential on a large-
scale to produce inexpensive sugars for fermentation is low. U.S. Pat. No. 5,188,673 [84]
employs concentrated acid hydrolysis which has benefits of high conversions of biomass,
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but suffers from low product yields due to degradation and the requirement of acid
recovery and recycles. Sulfuric acid concentrations used are 30-70 weight percent at
temperatures less than 100°C. Although 90% hydrolysis of the cellulose and
hemicellulose is achieved by this process, the concentrated acids are toxic, corrosive and
hazardous and require reactors that are resistant to corrosion. In addition, the
concentrated acid must be recovered after the hydrolysis to make the process
economically feasible [85]. A multi-function process for hydrolysis and fractionation of
lignocellulosic biomass to separate hemicellulosic sugars using mineral acid like sulfuric
acid, phosphoric acid or nitric acid has been described [86]. A process for treatment of
hemicellulose and cellulose in two different configurations is described [87].
Hemicellulose is treated with dilute acid in a conventional process. The cellulose is
separated out from the "prehydrolyzate" and then subjected to pyrolysis at high
temperatures. Further, the process step between the hemicellulose and cellulose reactions
require a drying step with a subsequent pyrolysis high temperature step at 400-600°C for
conversion of the cellulose to fermentable products.
3.2 Steam explosion
Autohydrolysis and steam explosion processes selectively hydrolyze
hemicellulose from lignocellulosic biomass. The process is economic and ecofriendly as
it doesn’t involves chemical catalyst. A relatively high hemicellulose recovery in the
range of 55–84%, together with low levels of inhibitory by-products, has been obtained
through autohydrolysis [88-91]. Cellulose and lignin are not significantly affected,
yielding a cellulose- and lignin-rich solid phase together with a liquid fraction with a
relative low concentration of potential fermentation inhibitors. The main drawback of this
process is that the solubilised hemicellulose appears mainly in oligomeric form [16, 22,
90, 92, 93,]. In principle steam explosion is one of the attractive pretreatment methods
that can cause disintegration of the material, thereby creating a large surface area on
which cellulase enzyme complex can act upon. Simultaneously hemicellulose is
separated during the steam explosion process thereby improving the accessibility to the
enzymes and enhancement of the over all lignocellulose degradation [94]. Most steam
treatments yield high hemicellulose solubility (producing mainly oligosaccharides) along
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with slight lignin solubilisation. Studies without added catalyst report sugars recoveries
between 45% and 69% [95-98]. Cara et al., [99] have reported a 30% increase in
maximum pentose yield for olive tree prunings when steam explosion was followed by
acid hydrolysis.
3.3 Enzymatic hydrolysis The most promising method for hydrolysis of polysaccharides to monomer sugars
is by use of enzymes, i.e., cellulases and hemicellulases. Moreover, hemicellulases
facilitate cellulose hydrolysis by exposing the cellulose fibres, thus making them more
accessible [100]. The enzymatic hydrolysis of the lignocellulosic biomass is preceeded by
a pretreatment process in which the lignin component is separated from the cellulose and
hemicellulose to make it amenable to the enzymatic hydrolysis. The lignin interferes with
the hydrolysis by blocking the access of the cellulases to the cellulose and by irreversibly
the binding hydrolytic enzymes. Therefore, the removal of the lignin can dramatically
increase the hydrolysis rate [101]. Recently the enzymatic hydrolysis of lignocellulosic
biomass has been optimized using enzymes from different sources and mixing in an
appropriate proportion using statistical approach of factorial design. A twofold reduction
in the total protein required to reach glucan to glucose and xylan to xylose hydrolysis
targets (99% and 88% conversion, respectively), thereby validating this approach towards
enzyme improvement and process cost reduction for lignocellulose hydrolysis [102].
3.3.1 Hemicellulose degrading enzymes
3.3.1.1 Xylanases and Mannanases Hemicellulases are multi-domain proteins and generally contain structurally
discrete catalytic and non-catalytic modules. The most important non-catalytic modules
consist of carbohydrate binding domains (CBD) which facilitate the targeting of the
enzyme to the polysaccharide, interdomain linkers, and dockerin modules that mediate
the binding of the catalytic domain via cohesion-dockerin interactions, either to the
microbial cell surface or to enzymatic complexes such as the cellulosome [100]. Based on
the amino acid or nucleic acid sequence of their catalytic modules hemicellulases are
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either glycoside hydrolases (GHs) which hydrolyse glycosidic bonds, or carbohydrate
esterases (CEs), which hydrolyse ester linkages of acetate or ferulic acid side groups and
according to their primary sequence homology they have been grouped into various
families. Most studies on hemicellulases have focused until now on enzymes that
hydrolyse xylan [7]. Enzymes that hydrolyse mannan have been largely neglected, even
though it is an abundant hemicellulose, therefore the application of mannanases for
catalysing the hydrolysis of β-1,4 mannans is as important as the application of
xylanases.
Due to the complex structure of hemicelluloses, several different enzymes are
needed for their enzymatic degradation or modification. The two main glycosyl
hydrolases depolymerising the hemicellulose backbone are endo-1, 4- β-D-xylanase and
endo-1, 4-β-D-mannanase [5]. Endo-1,4-β-xylanase cleaves the glycosidic bonds in the
xylan backbone, bringing about a reduction in the degree of polymerization of the
substrate. Xylan is not attacked randomly, but the bonds selected for hydrolysis depend
on the nature of the substrate molecule, i.e., on the chain length, the degree of branching,
and the presence of substituents [103]. Initially, the main hydrolysis products are β-D-
xylopyranosyl oligomers, but at a later stage, small molecules such as mono-, di- and
trisaccharides of β-D-xylopyranosyl may be produced [104]. These enzymes are
produced by fungi, bacteria, yeast, marine algae, protozoans, snails, crustaceans, insect,
seeds, etc. [105]. Filamentous fungi are particularly interesting producers of xylanases
since they excrete the enzymes into the medium and their enzyme levels are much higher
than those of yeasts and bacteria [7]. Aspergillus niger, Humicola insolens,
Termomonospora fusca, Trichoderma reesei, T. longibrachiatum, T. koningii have been
used as industrial sources of commercial xylanases. Nevertheless, commercial xylanases
can also be obtained from bacteria, e.g., from Bacillus sp. Xylanases have many
commercial uses, such as in paper manufacturing, animal feed, bread-making, juice and
wine industries, or xylitol production [104, 106]. Several investigations so far have
indicated that xylanases are usually inducible enzymes [107], and different carbon
sources have been analysed to find their role in effecting the enzymatic levels. Xylanase
biosynthesis is induced by xylan, xylose, xylobiose or several β-D-xylopyranosyl
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residues added to the culture medium during growth [7, 104, 108]. However, constitutive
production of xylanase has also been reported [109]. Catabolite repression by glucose is a
common phenomenon observed in xylanase biosynthesis [7].
The main sugar moiety of galactoglucomannans (GGM) is D-mannose, but for its
complete breakdown into simple sugars, the synergistic action of endo-1,4-β-mannanases
(EC 3.2.1.78) and exoacting β-mannosidases (EC 3.2.1.25) is required to cleave the
polymer backbone. Additional enzymes, such as β-glucosidases (EC 3.2.1.21), α-
galactosidases (EC 3.2.1.22) and acetyl mannan esterases are required to remove side
chain sugars that are attached at various points on mannans [110, 112]. The property of
mannanolysis is widespread in the microbial world. A vast variety of bacteria,
actinomycetes, yeasts and fungi are known to be mannan degraders [113, 114].
Mannanases of microbial origin have been reported to be both induced as well as
constitutive enzymes and are usually being secreted extracellularly [110]. Although a
number of mannanase-producing bacterial sources are available, only a few are
commercially exploited as wild or recombinant strains, of these, the important ones are:
Bacillus sp., Streptomyces sp., Caldibacillus cellulovorans, Caldicellulosiruptor Rt8B,
Caldocellum saccharolyticum [115-117].
3.3.1.2 Accessory enzymes for hemicellulose hydrolysis
Since xylan is a complex component of the hemicelluloses in wood, its complete
hydrolysis requires the action of a complete enzyme system, which is usually composed
of β-xylanase, β-xylosidase, and enzymes such as α-L-arabinofuranosidase, α-
glucuronidase, acetylxylan esterase, and hydroxycinnamic acid esterases that cleave side
chain residues from the xylan backbone. Figure 1 describes the mechanism of xylanolytic
enzymes involved in the degradation of hemicellulose. All these enzymes act
cooperatively to convert xylan to its constituents [5]. Xylanases attack randomly the
backbone of xylan to produce both substituted and non-substituted shorter chain
oligomers, xylobiose and xylose. Xylosidases are essential for the complete breakdown
of xylan as they hydrolyse xylooligosaccharides to xylose [118]. The enzymes
arabinosidase, α-glucuronidase and acetylxylan esterase act in synergy with the xylanases
16
and xylosidases by releasing the substituents on the xylan backbone to achieve a total
hydrolysis of xylan to monosaccharides [4]. Table 2 lists the microorganisms producing
different hemicellulases [119].
3.3.1.3 Xyloglucanases
Many endoglucanases have been reported to hydrolyze xyloglucan as a substrate
analog [120], however few endo-β-1,4-glucanases have high activity toward xyloglucan,
with little or no activity towards cellulose or cellulose derivatives [121, 122]. They have
been assigned a new EC number (EC 3.2.1.151) and designated as xyloglucanase,
xyloglucan hydrolase (XGH), or xyloglucan-specific endo-β-1,4-glucanases (XEGs)
belonging to families 5, 12, 44, and 74, according to a recent classification of glycoside
hydrolases (GHs) available at http://afmb.cnrs-mrs.fr/CAZY/ [123–126]. They represent
a new class of polysaccharide degrading enzymes which can attack the backbone at
substituted glucose residues. Among these enzyme families, xyloglucanases placed in
GH family 74 are known to have high specific activity towards xyloglucan, with
inversion of the anomeric configuration, and both endo-type and exo-type hydrolases
have been found in several microorganisms [127– 134]. The exo-type enzymes recognize
the reducing end of xyloglucan oligosaccharide (oligoxyloglucan reducing- end-specific
cellobiohydrolase, EC 3.2.1.150, from Geotrichum sp. M128 [129] and oligoxyloglucan
reducing- end-specific xyloglucanobiohydrolase from Aspergillus nidulans [134]),
whereas the endo-type enzymes hydrolyze xyloglucan polymer randomly. In addition,
XEG74 from Paenibacillus sp. KM21 and Cel74A from Trichoderma reesei have been
reported to have endo-processive or dual-mode endo-like and exo-like activities [127,
132]. The complete hydrolysis of galactoxyloglucan requires an accessory enzyme, β-
galactosidase which cleaves the galactose residue attached to the branched xylose moiety
in the β-D-glucopyranose backbone [135]. The xyloglucan and the enzymes responsible
for its modification and degradation are finding increasing prominence, reflecting the
drive for diverse biotechnological applications in fruit juice clarification, textile
processing, cellulose surface modification, pharmaceutical delivery and production of
food thickening agents [136].
17
3.3.2 Role of thermostable hemicellulases in hydrolysis
Thermostable enzymes are gaining wide industrial and biotechnical interest due to
the fact that they are more stable and thus generally better suited for harsh process
conditions. Thermostability is expressed as a measure of the half-life of enzyme activity
at elevated temperatures [137]. Thermostable enzymes are produced both by thermophilic
and mesophilic organisms. Although thermophilic microorganisms are a potential source
for thermostable enzymes, the majority of industrial thermostable enzymes originate from
mesophilic organisms. Thermophilic bacteria have, however, received considerable
attention as sources of highly active and thermostable enzymes. Thermostable enzymes in
the hydrolysis of lignocellulosic materials have several potential advantages: higher
specific activity (decreasing the amount of enzyme needed), higher stability (allowing
elongated hydrolysis times) and increased flexibility for the process configurations [138].
The two first characteristics would expectedly improve the overall performance of the
enzymatic hydrolysis even at the range of conventional enzymes active at around 50°C.
Thus, carrying out the hydrolysis at higher temperature would ultimately lead to
improved performance, i.e. decreased enzyme dosage and reduced hydrolysis time and,
thus, potentially decreased hydrolysis costs. Thermostable enzymes would expectedly
also allow hydrolysis at higher consistency due to lower viscosity at elevated
temperatures and allow more flexibility in the process configurations. In addition to
improved performance in the hydrolysis of lignocellulosic substrates, thermophilic
enzymes allow the design of more flexible process configurations [138]. Menon et al,
[139] have analyzed the comparative data on hydrolysis of hemicellulosic substrates, oat
spelt xylan and wheat bran hemicellulose at different temperatures with thermostable
xylanase from alkalothermophilic Thermomonospora sp. At higher temperature, rapid
hydrolysis of hemicellulosic substrates were achieved, favouring a reduction in process
time and enzyme dosage.
18
4 Production of biochemicals from hemicellulosic hydrolysates
4.1 Fermentation of hemicellulosic hydrolysates to fuel ethanol
D-xylose, the predominant sugar of hemicellulose can be produced from
xylan/hemicellulose by the action of acid/enzymes. D-xylose is not readily utilized as D-
glucose for the production of ethanol by microorganisms such as Saccharomyces
cerevisiase. Bacteria, yeast and mould differ in their mode of conversion of D-xylose to
xylulose, the intial step in xylose fermentation. Few yeasts uses the enzymes xylose
reductase to reduce xylose to xylitol which is subsequently oxidized to D-xylulose by
xylitol dehydrogenase. Yeasts such as Pichia, Klyveromyces, Pachysolen are reported to
ferment xylose to ethanol under micro-aerophilic condition. The general requirements of
an organism to be used in ethanol production is that it should give a high ethanol yield, a
high productivity and be able to withstand high ethanol concentrations in order to keep
distillation costs low. In addition to these general requirements, inhibitor tolerance,
temperature tolerance and the ability to utilize multiple sugars are also essential. Figure 2
describes the catabolism of hemicellulose derived monosaccharide.
Essentially, there are three different types of processes namely: Separate
Hydrolysis and Fermentation (SHF), Direct Microbial Conversion (DMC) and
Simultaneous Saccharification and Fermentation (SSF). SSF has been shown to be the
most promising approach to biochemically convert cellulose to ethanol in an effective
way [140].
4.1.1 Separate hydrolysis and fementation (SHF)
SHF is a conventional two step process where the hemicellulose is hydrolysed
using the enzymes to form the reducing sugars in the first step and the sugars, thus
formed, are fermented to the ethanol in the second step using Pichia or Zymomonas. The
advantage of this process is that each step can be carried out at its optimum conditions.
Table 3 summarizes the ethanol production from hemicellulosic hydrolysates. A recent
study by Wilkins et al, [149] reveals an ethanol production by Kluyveromyces sp from D-
xylose at 40°C and 45°C with a yield of 0.15g/g and 0.08g/g respectively under anaerobic
19
condition. H. polymorpha could ferment both glucose and xylose up to 45°C [150].
Debaryomyces sp used both pentoses and hexoses to similar extents in sugar mixtures
and a preference for one carbohydrate did not inhibit the consumption of other. This
could be important, since the usual substrates of hemicelluloses, which often consists of
sugar mixtures [151].
4.1.2 Direct microbial conversion (DMC)
This process involved three major steps, namely enzyme production, hydrolysis of
the lignocellulosic biomass and the fermentation of the sugars all occurring in one step
[152]. The relative lower tolerance of the ethanol is the main disadvantage of this
process. A lower tolerance limit of about 3.5% has been reported as compared to 10% of
ethanologenic yeasts. Acetic acid and lactic acid are also formed as the by-products in
this process in which a significant amount of carbon get utilized [153]. Neurospora
crassa is known to produce ethanol directly from the cellulose/hemicellulose, since it
produces both the cellulase and the xylanase and also has the capacity to ferment the
sugars to ethanol anaerobically [154].
4.1.3 Simultaneous saccharification and fermentation (SSF)
The saccharification of the lignocellulosic biomass by the enzymes and the
subsequent fermentation of the sugars to ethanol by the yeast such as Saccharomyces or
Zymomonas takes place in the same vessel in this process [155]. The compatibility of
both saccharification and fermentation process with respect to various conditions such as
pH, temperature, substrate concentration etc. is one of the most important factors
governing the success of the SSF process. The main advantage of using SSF for the
ethanol bioconversion is enhanced rate of hydrolysis of lignocellulosic biomass (cellulose
and hemicellulose) due to removal of end product inhibition. Several inhibitory
compounds are formed during hydrolysis of the raw material, the hydrolytic process has
to be optimized so that inhibitor formation can be minimized. When low concentrations
of inhibitory compounds are present in the hydrolyzate, detoxification is easier and
fermentation is cheaper. The choice of a detoxification method has to be based on the
degree of microbial inhibition caused by the compounds. As each detoxification method
20
is specific to certain types of compounds, better results can be obtained by combining
two or more different methods. Another factor of great importance in the fermentative
processes is the cultivation conditions, which, if inadequate, can stimulate the inhibitory
action of the toxic compounds. Thermotolerant yeast is an added advantage for SSF and
thermotolerant yeast strains, e.g. Fabospora fragilis, Saccharomyces uvarum, Candida
brassicae, C. lusitaniae, and Kluyveromyces marxianus, have been evaluated for future
use in SSF, to allow fermentation at temperatures closer to the optimal temperature for
the enzymes. However, in all these cases saccharification of pure cellulose (e.g.
Sigmacell-50) or washed fibers, in defined fermentation medium, were applied. SSF of
cellulose with mixed cultures of different thermotolerant yeast strains have also been
carried out [156]. However, there is scarcity of literature from SSF experiments in which
hemicelluloses have been used together with thermotolerant strains. Menon et al., [139]
have reported for the first time the SSF experiments using hemicellulosic substrates and
thermotolerant Debaryomyces hansenii. Maximum ethanol concentrations of 9.1 g/L and
9.5 g/L were obtained in SSF with oat spelt xylan and wheat bran hemicellulose
respectively. These concentrations were attained in 36h for OSX and 48h for WBH from
the onset of SSF. The increased ethanol yield in SSF systems is evidently due to removal
of xylose formed during hydrolysis which causes end product inhibition.
4.2 Production of xylitol from hemicellulosic hydrolysates
The conversion of xylose to value-added product xylitol will have significant role
in the economic viability of cellulose bioconversion process. The pretreatment (acid or
enzyme) of hemicellulose predominantly liberates xylose and arabinose and
hydrogenation products are xylitol and arabinitol. Xylitol is a naturally occurring five-
carbon sugar alcohol [157], has important applications in pharmaceuticals and food
industries due to high sweetening properties, non- and anticariogenicity property and
microbial growth inhibition capacity [158, 159]. Xylitol is used as sugar substitute for
diabetics, as it does not require insulin for its metabolism [160]. Recently it has been
demonstrated that xylitol can prevent acute otitis media (AOM) [161] in children. The
value of xylitol market is currently $340 million with applications in mouthwashes,
21
toothpastes and chewing gums as well as in foods for special dietary uses. Global xylitol
consumption was 43,000 t in 2005, the U.S. and Western Europe accounting for 30 and
37% of the total xylitol consumption [162]. Currently the major product of xylan which is
of considerable importance is xylitol derived mainly from agricultural and wood residues
respectively.
Xylitol is produced from xylan-rich biomass by both chemical and biological
methods [163]. The chemical process adapted so far is not ecofriendly and hydrogenation
of xylose demands huge production costs in terms of temperature and pressure input as
well as the formation of by-products that require expensive separation and purification
steps. In addition, xylose should be purified before it is hydrogenised, which further
increases the capital investment and costs for xylitol production [164]. Certain microbial
species have potential to utilize xylose as carbon source by converting it to xylitol. Yeasts
naturally produce xylitol as an intermediate during D-xylose metabolism. Xylose
reductase (XR) is typically an NADPH or NADH-dependent enzyme which oxidizes
xylose to xylitol [165], while enzyme xylitol dehydrogenase (XDH) requires NAD+
[166] for reduction of xylitol to xylulose. The cofactors imbalance results in the secretion
of xylitol as a xylose fermentation by-product [167]. Various bioconversion methods
have been exploited for the production of xylitol from hemicellulose using
microorganisms or their enzymes [168]. Xylitol production from hemicellulosic
hydrolysates using various microorganisms is illustrated in Table 4. Xylitol production by
fermentation is usually operated under mild conditions and could achieve higher yield
and will lead to an ecofriendly process. Hence the biological process for xylitol
production is receiving wider attention.
4.3 Production of other value added products from hemicellulosic hydrolysates
4.3.1 Butanediol
2,3-Butanediol, also known as 2,3-butylene glycol (2,3-BD) is a valuable
chemical feedstock because of its application as a solvent, liquid fuel, and as a precursor
22
of many synthetic polymers and resins [3]. Butanediol is produced during oxygen-limited
growth, by a fermentative pathway known as the mixed acid-butanediol pathway [175].
The 2,3-BD pathway and the relative proportions of acetoin and butanediol serve to
maintain the intracellular NAD/NADH balance under changing culture conditions. All of
the sugars commonly found in hemicellulose and cellulose hydrolysates can be converted
to butanediol, including glucose, xylose, arabinose, mannose, galactose, and cellobiose.
The theoretical maximum yield of butanediol from sugar is 0.50 kg per kg. With a
heating value of 27,200 J/g, 2,3-BD compares favorably with ethanol (29,100 J/g) and
methanol (22,100 J/g) for use as a liquid fuel and fuel additive [176].
Hexose and pentose can be converted to 2,3-BD by several microorganisms
including Aeromonas [177], Bacillus [178], Paenibacillus [179], Serratia, Aerobacter
[180], Enterobacter [11] and also Klebsiella [181–186]. Among these, especially
Klebsiella is often used for 2,3-BDO production because of its broad substrate spectrum
and cultural adaptability. At first, 2,3-BD was produced worldwide from glucose or
xylose by fermentation. Starch [177,187], molasses [188], water hyacinth [189] and
Jerusalem artichoke tubers [190] were also tested for their suitability as substrates for 2,3-
BD fermentation. Some studies have been conducted utilizing the hemicellulose portion
of forest residues like wood for 2,3-BD production [191]. Cheng et al, [186] have
reported production of 2, 3-butanediol from corn cob hydrolysate by fed batch
fermentation using Klebsiella oxytoca. A maximal 2,3-butanediol concentration of 35.7
g/l was obtained after 60 h of fed-batch fermentation, giving a yield of 0.5 g/g reducing
sugar and a productivity of 0.59 g/h l.
4.3.2 Ferulic acid and Vanillin
Ferulic acid is the major phenolic acid of cereal grains such as wheat, triticale,
and rye [192, 193, 194]. It can exist as an extractable form, as free, esterified, and
glycosylated phenolic constituents [192] as well as an insoluble-bound occurring in the
outer layers of wheat grains [195]. Alkaline hydrolysis is reported to release ferulic acid
from the insoluble form [195, 196]. Also, enzyme preparations are used for the same
23
purpose [197]. There are numerous reports on antiradical, oxidase inhibitory,
antiinflammatory, antimicrobial,anticancer activities of ferulic acid and its derivatives in
the literature data [198-200]. Feruloyl esterases (FAEs) act synergistically with xylanases
to hydrolyze ester-linked ferulic (FA) and diferulic (diFA) acid from cell wallmaterial
and therefore play a major role in the degradation of plant biomass [201]. FAEs are used
as a tool for the release of Ferulic acid from agroindustrial byproducts such as wheat bran
[202-216], maize bran [217-219], maize fiber [210], brewer’s (or barley) spent grain
[202,220-227], sugar beet pulp [216,217,228-231], coastal bermudagrass [232], oat hulls
[233-235], jojobameal [236], wheat straw[217], coffee pulp [217] and apple marc [217].
Vanillin can be obtained by fermentation using suitable microorganisms in the stationary
growth phase [237-240]. The microbial transformation of ferulic acid is recognized as
one of the most attracting alternatives to produce natural vanillin. Bacteria belonging to
different genera are able to metabolize ferulic acid as the sole carbon source, producing
vanillin, vanillic acid and protocatechuic acid as catabolic intermediates [241,242].
Vanillin is industrially used as fragrance in food preparations, intermediate in the
productions of herbicides, antifoaming agents or drugs [243], ingredient of household
products such as air fresheners and floor polishes, and, because of its antimicrobial and
antioxidant properties [244,245], also as food preservative [246]. The much higher price
of the natural vanillin compared to the synthetic vanillin has been leading to growing
interest of the flavor industry in producing it from natural sources by bioconversion
[243,247–250]. Pseudomonas fluorescens was shown to produce vanillic acid from
ferulic acid [251,252], with formation of vanillin as an intermediate [253]. Promising
vanillin concentrations were obtained from ferulic acid by Amycolatopsis sp. [250,254]
and Streptomyces setonii [241,255,256]. However, the process development is difficult
because of the well known slow growth of actinomycetes and high viscosity of broths
fermented by them; therefore, the construction of new recombinants strains of quickly
growing bacteria able to overproduce vanillin is attractive. The biotechnological process
to produce vanillin from various agro by-products had been investigated using different
microorganisms as biocatalysts. Aspergillus niger I-1472 and Pycnoporus cinnabarinus
MUCL39533 were used in a two-step bioconversion using sugar-beet pulp [257,258],
24
maize bran [259], rice bran oil [260], and wheat bran [261]. Wheat bran [262] and corn
cob [263] is recently reported as a good substrate for biovanillin production by
Escherichia coli JM 109/pBB1.
4.3.3 Lactic acid
Lactic acid is widely used in food, pharmaceutical and textile industries. It is also
used as a source of lactic acid polymers which are being used as biodegradable plastics
[264,265]. The physical properties and stability of polylactides can be controlled by
adjusting the proportions of the L(+)- and D(−)-lactides [266]. Optically pure lactic acid
is currently produced by the fermentation of glucose derived from corn starch using
various lactic acid bacteria [267,268]. However, the fastidious lactic acid bacteria have
complex nutritional requirements [269] and the use of corn is not favoured as the
feedstock competes directly with the food and feed uses. The use of lignocellulose
biomass will significantly increase the competitiveness of lactic acid-based polymers
compared to conventional petroleum based plastics.
Lactobacillus spp. are used extensively in industry for starch-based lactic acid
production, the majority of which lack the ability to ferment pentose sugars such as
xylose and arabinose [266]. Although, Lactobacillus pentosus, L. brevis and Lactococcus
lactis ferment pentoses to lactic acid, pentoses are metabolized using the phosphoketolase
pathway which is inefficient for lactic acid production [270,271]. In the phosphoketolase
pathway, xylulose 5-phosphate is cleaved to glyceraldehyde 3-phosphate and acetyl-
phosphate. With this pathway, the maximum theoretical yield of lactic acid is limited to
one per pentose (0.6 g lactic acid per g xylose) due to the loss of two carbons to acetic
acid. Sugarcane bagasse [272], steam exploded wood [273], soybean stalk [274], corncob
molasses [275], trimming vine shoots [276] and wheat straw [270] hydrolysate are used
for lactic acid production. John et al, [277] have reported the production of lactic acid
from agro residues using Lactobacillus delbrueckii in solid state and simultaneous
saccharification and fermentation [278].
25
4.3.4 Furfural
Furfural is commercially synthesized from lignocellulosic biomass in a process
which involves acid hydrolysis of the pentosans or xylans, present in the hemicelluloses
of some agriculture residues and hardwoods into pentoses or xylose and successive
cyclodehydration of the latter to form furfural. The second step of dehydration is
comparatively slower than that of mineral acid hydrolysis. There is no synthetic route
available for the production of furfural. The potential furfural yield for typical feedstock
is expressed in terms of kg of furfural per metric ton of dry biomass. It is reported to be
220 for corncobs, 170 for bagasse, 160 for cornstalks, 160 for sunflower hulls, and
approximately 150–170 for hardwoods. Furfural, derived from renewable biomass, is
used for the production of a wide spectrum of important non- petroleum derived
chemicals. World market for furfural currently is estimated to be about 200,000 to
210,000 tpa of which about 60–62% is used for the production of furfuryl alcohol.
Furfural is well known for its thermosetting properties, physical strength, and corrosion
resistance. It is mainly used for the production of resin which accounts for 70% of the
market. Currently, commercial production of furfural suffers from technological
challenges and maintenance problems [279]. The use of conventional acid catalysts
(mineral acids such as H2SO4 or HCl) for the production of furfural poses serious
environmental problems and subsequently results in few undesirable side reactions
causing the formation of huge amounts of toxic waste [280]. Hence, there is a need to
develop ecofriendly catalytic processes to eliminate environmental corrosion as well as
waste-disposal problems. Coniochaeta ligniaria NRRL30616 metabolizes furfural and 5-
hydroxymethylfurfural (HMF) as well as aromatic acids, aliphatic acids and aldehydes.
NRRL30616 grew in corn stover dilute-acid hydrolysate and converted furfural to both
furfuryl alcohol and furoic acid [281].
4.3.5 Butanol
Butanol is a 4-carbon alcohol (butyl alcohol), an advanced biofuel, offers a
number of advantages and can help accelerate biofuel adoption in countries around the
world. It can be produced through processing of domestically grown crops, such as corn
26
and sugar beets, and other biomass, such as fast-growing grasses and agricultural waste
products. Biobutanol's primary use is as an industrial solvent in products such as lacquers
and enamels and is also compatible with ethanol blending and can improve the blending
of ethanol with gasoline. Using fermentation to replace chemical processes in the
production of butanol depends largely on the availability of inexpensive and abundant
raw materials and efficient conversion of these materials to solvents. Solventogenic
Acetone Butanol Ethanol (ABE)-producing Clostridia have an added advantage over
many other cultures as they can utilize both hexose and pentose sugars [282], which are
released from wood and agricultural residues upon hydrolysis, to produce ABE. Parekh et
al. [283] produced ABE from hydrolysates of pine, aspen, and corn stover using
Clostridium acetobutylicum P262. Similarly Marchal et al. [284] used wheat straw
hydrolysate and C. acetobutylicum, while Soni et al. [285] used bagasse and rice straw
hydrolysates and C. saccharoperbutylacetonicum to convert these agricultural wastes into
ABE. Qureshi et al, [286] have studied the production of butanol from corn fibre
hydrolysate using Clostridium beijerinckii BA101. Sun and Liu, [287] have reported the
production of butanol from sugar maple hemicellulose hydrolysate using Clostridia
acetobutylicum ATCC824.
4.3.6 Biohydrogen
Hydrogen is considered to be an ideal energy alternative for the future. Compared
with the conventional hydrogen generation process (thermochemical and
electrochemical), bio-hydrogen production processes are more environmentfriendly and
less-energy intensive. Hydrogen could be produced from renewable materials, such as
waste water, organic wastes, corn straws and waste water sludge, etc. Biohydrogen has
the potential to considerably reduce costs and environmental impact as it can be produced
with sunlight and minimal nutrients or organic waste effluents. Hydrogen (H2) producing
microorganisms can be rapidly grown in bioreactors with relatively small energy and
environmental footprints, making biohydrogen a renewable and low impact technology.
Biological hydrogen, biohydrogen, production may provide a renewable, more
sustainable alternative but has yet to reach a scale large enough for consideration in
replacing a significant portion of the hydrogen supply [288].Various hemicellulosic
27
hydrolysates from wheat straw [289,290], Corn stover [291], sugarcane bagasse [292],
Sweet sorghum [293], Corn straw [294] have been evaluated for biohydrogen production.
4.3.7 Chitosan
Chitosan, as an important kind of biomacromolecules, has protective and
supportive functions in cell walls of zygomycetes [295]. Chitosan synthesis is a complex
process, which involves changes of many intermediate metabolites and energy charges in
cells. Chitosan itself has found numerous applications in food, cosmetics and
pharmaceutical industries because of their unique properties such as nontoxicity,
biodegradability, biocompatibility, film-forming and chelating properties along with their
antimicrobial activity [296-298]. Tai et al, [299] have studied the potential of
hemicellulose hydrolysate of corn straw for the production of chitosan by Rhizopus
oryzae.
4.3.8 Xylo-oligosaccharides
Xylo-oligosaccharides are xylose-based oligomers that may have variable
proportions of substitute groups like acetyl, uronic, and phenolic acids, depending on the
lignocellulosic from which they are extracted and the process of production. Substituted
xylo-oligosaccharides have some specific characteristics that are driving research efforts
to develop applications in fields related to the food and pharmaceutical industries. They
may be used as soluble dietary fiber because of its low calorific value and acceptable
organoleptic properties. Furthermore, they are non-carcinogenic and act as prebiotics
promoting the growth of beneficial Bifidobacteria in the colon [300,301], and are
considered a possible ingredient in functional foods [302,303]. Some studies point to the
beneficial effect of xylo- ligosaccharides may have on reducing the risk of colon cancer
[304-306]. Recently, xylo-oligosaccharides extracted by autohydrolysis of bamboo have
been found to posses a cytotoxic effect on human leukemia cells [307]. Also xylo-
oligosaccharides can be used as a source of xylose for the production of xylitol, a well-
known low-calorie sweetener [308].
28
5 Recombinant approaches for ethanol and xylitol production
Recombinant DNA methods are being currently used for overexpression of
cellulases and hemicellulases for lignocellulose hydrolysis and production of bioproducts.
Saccharomyces cerevisiae and Z.mobilis have been explored for utilizing lignocellulosic
biomass by genetic manipulations. Several metabolic and evolutionary engineering
techniques have been applied to: increase substrate range, like in S. cerevisiae and Z.
mobilis; maximize ethanol production like in E. coli; supply other important traits to
improve lignocellulose conversion to ethanol and xylitol. In general, approaches involve
genetic engineering towards those specific traits, often followed by strain optimization
through adaptation. Since the molecular basis for ethanol and inhibitor tolerance is not
fully understood, random mutagenesis and evolutionary engineering have also been
applied to improve those traits. Moreover, as a result of technological developments,
systems biology approaches have been recently applied to characterise the functional
genomics of microorganisms and to evaluate the impact of metabolic and evolutionary
engineering strategies. This advanced characterisation (genomics, transcriptomics,
proteomics, metabolomics) is already contributing to better understand physiological
responses and to identify crucial targets for metabolic engineering [309-313]. Table 5
illustrates the production of xylitol and ethanol using recombinant strains.
6 Future prospects
The implementation of lignocellulosic material for the development of alternative
energy is an important aspect for the lucrative gratification of biofuel vision. For an
economically viable bioconversion process it is necessary to utilize both the cellulosic
and hemicellulosic fractions of biomass. The research focus on cellulosic ethanol is
tremendously enhanced all over the world. On the other hand, corn and other grains can
be readily hydrolyzed to glucose with high yield by low-cost enzymes. However this
causes the debate on food versus fuel. Therefore a hemicellulose based process, wherein
hemicellulose can be readily hydrolyzed to value added products is foreseen to be
significant from this aspect. The fermentation of C6 sugars by Saccharomyces is a well
29
established technology whereas the utilization of C5 sugars is a challenging area.
Extensive research on xylose fermenting yeasts such as Pachysolen tannophilus, Candida
shehatae and Pichia stiptis, has been conducted. However, low ethanol tolerance,
conversion rates and catabolite repression in xylose conversion due to glucose need to be
addressed. Another potential ethanologen is recombinant Zymomonas mobilis in view of
its ability to ferment both xylose and glucose. E. coli strain developed by Ingram’s group
deserves special recognition, as it not only ferments all five sugars (glucose, xylose,
arabinose, mannose and galactose) present in the synthetic sugar mixtures to ethanol but
also performs competently in real hydrolyzates like that of Pinus. Saccharomyces LNH-
ST is another promising recombinant capable of fermenting dilute acid treated corn fiber
hydrolyzates.
Pervasive research needs to be performed to develop an efficient cost-effective
and eco-friendly pretreatment methods, enzymes or cocktails of various enzymes for
complete conversion of hemicellulose to monomers, efficient and robust microorganisms
for fermentation of hemicellulosic hydrolysates to value added products in a cost-
competitive way. Although bioethanol production has been greatly improved by new
technologies there are still challenges that need further investigations. These challenges
include maintaining a stable performance of the genetically engineered yeasts in
commercial scale fermentation operations and integrating the optimal components into
economic ethanol production systems.
Metabolic engineering and other classical techniques such as random mutagenesis
address the further enhancement of microorganism capabilities by adding or modifying
traits such as tolerance to ethanol and inhibitors, efficient hydrolysis of
cellulose/hemicellulose, thermotolerance, reduced need for nutrient supplementation and
improvement of sugars transport. The improvement achieved in the fermentation step
with the help of metabolic engineering is just one of the aspects of an integrated process.
Keeping a realistic perspective one can conclude that several pieces still remain to be
properly assembled and optimized before an efficient industrial configuration is acquired.
30
Acknowledgements
MR acknowledges the financial support from CSIR Emeritus Scheme.
References 1. Smeets, E.M.W., Faaij, A.P.C., Lewandowski, I.M., Turkenburg, W.C., A bottom-
up assessment and review of global bio-energy potential to 2050, Prog Energy Combustion Sci, 33(2007), 56-106
2. Pandey A, Handbook of plant based biofuels, CRC press, Taylor & Francis Group, FL, 2009
3. Saha, B.C., Hemicellulose bioconversion, J Ind Microbiol Biotechnol, 30(2003), 279-291
4. Gilbert, H.J., Hazlewood, G.P., Bacterial cellulases and xylanases, J. Gen. Microbiol, 139(1993), 187-194
5. Biely, P., Biochemical aspects of the production of microbial hemicellulases. In: Hemicellulose and Hemicellulases (Coughlan, M.P. and Hazlewood, G.P., Eds.), pp. 29-51. Portland Press, Cambridge, 1993
6. Ball, A.S., McCarthy, A.J., Production and properties of xylanases from actinomycetes, J. Appl. Bacteriol, 66(1989), 439-444
7. Kulkarni, N., Shendye, A., Rao, M., Molecular and Bioechnological aspects of xylanases, FEMS Microbiology reviews, 23(1999), 411-456
8. Kormelink, F.J.M., Voragen, A.G., Degradation of different (glucurono)arabino]xylans by a combination of purified xylan-degrading enzymes, Appl Microbiol Biotechnol, 38(1993), 688– 695
9. Shibuya, N., Iwasaki, T., Structural features of rice bran hemicellulose, Phytochemistry, 24(1985), 285–289
10. Gruppen, H., Hamer, R.J., Voragen, A.G.J., Water-unextractable cell wall material from wheat flour. 2. Fractionation of alkali-extracted polymers and comparison with waterextractable arabinoxylans, J Cereal Sci, 16(1992), 53–67
11. Saha, B.C. and Bothast, R.J., Pretreatment and enzymatic saccharification of corn fiber, Appl Biochem Biotechnol, 76(1999), 65–77
12. Doner, L.W., Hicks, K.B., Isolation of hemicellulose from corn fiber by alkaline hydrogen peroxide extraction, Cereal Chem, 74(1997), 176–181
13. Atkins, E.D.T., Three dimensional structure, interactions and properties of xylans. In: Xylan and Xylanases (Visser, J., Beldman, G., Someren, M.A.K. and Voragen, A.G.J., Eds.), Elsevier, Amsterdam, 1992, 21-39.
14. Biswas, S., Basak, P.K. and Kaushik, N., Bioprocess and Bioproducts- Emerging Trends, TIFAC, New Delhi, 2009
15. Torget, R., Walter, P., Himmel, M., Grohmann, K., Dilute-acid pretreatment of corn residues and short-rotation woody crops, Appl. Biochem. Biotechnol, 28– 29(1991), 75–86
16. Kabel, M.A., Carvalheiro, F., Garrote, G., Avgerinos, E., Koukios, E., Parajo, J.C., Girio, F.M., Schols, H.A., Voragen, A.G.J., Hydrothermally treated xylan rich
31
byproducts yield different classes of xylo-oligosaccharides, Carbohydr. Polym, 50(2002), 47–56
17. Ropars, M., Marchal, R., Pourquie, J., Vandecasteele, J.P., Large-scale enzymatic hydrolysis of agricultural lignocellulosic biomass. 1. Pretreatment procedures, Bioresour. Technol, 42 (1992), 197–204
18. Garrote, G., Dominguez, H., Parajo, J.C., Kinetic modelling of corncob autohydrolysis, Process Biochem, 36 (2001), 571–578
19. Torget, R.W., Kim, J.S., Lee, Y.Y., Fundamental aspects of dilute acid hydrolysis/ fractionation kinetics of hardwood carbohydrates. 1. Cellulose hydrolysis, Ind. Eng. Chem. Res, 39 (2000), 2817–2825
20. Lee, J., Biological conversion of lignocellulosic biomass to ethanol, J. Biotechnol, 56 (1997), 1–24
21. Rubio, M., Tortosa, J.F., Quesada, J., Gomez, D., Fractionation of lignocellulosics. Solubilization of corn stalk hemicelluloses by autohydrolysis in aqueous medium, Biomass Bioenerg, 15 (1998), 483–491
22. Allen, S.G., Schulman, D., Lichwa, J., Antal, M.J., Laser, M., Lynd, L.R., A comparison between hot liquid water and steam fractionation of corn fiber, Ind. Eng. Chem. Res, 40 (2001), 2934–2941
23. Vila, C., Garrote, G., Dominguez, H., Parajo, J.C., Hydrolytic processing of rice husks in aqueous media: A kinetic assessment, Collect. Czechoslovak Chem. Commun, 67 (2002), 509–530
24. Dekker, R.F.H., Wallis, A.F.A., Enzymic saccharification of sugarcane bagasse pretreated by autohydrolysis steam explosion, Biotechnol. Bioeng, 25 (1983), 3027–3048
25. Aguilar, R., Ramirez, J.A., Garrote, G., Vazquez, M., Kinetic study of the acid hydrolysis of sugar cane bagasse, J. Food Eng, 55 (2002), 309–318
26. Neureiter, M., Danner, H., Thomasser, C., Saidi, B., Braun, R., Dilute-acid hydrolysis of sugarcane bagasse at varying conditions, Appl. Biochem.Biotechnol, 98–100(2002), 49–58
27. Grohmann, K., Torget, R., Himmel, M., Optimization of dilute acid pretreatment of biomass, Biotechnol. Bioeng. Symp, 15(1985), 59–80
28. Magee, R.J., Kosaric, N., Bioconversion of hemicelluloses, Adv. Biochem. Eng./ Biotechnol, 32(1985), 61–93
29. Taherzadeh, M.J., Eklund, R., Gustafsson, L., Niklasson, C., Liden, G., Characterization and fermentation of dilute-acid hydrolyzates from wood, Ind. Eng. Chem. Res, 36 (1997), 4659–4665
30. Fengel, D., Wegener, G., Wood: Chemistry, Ultrastructure, Reactions. Walter de Gruyter and Co., Berlin, 1983
31. Pereira, H., Variability in the chemical composition of plantation Eucalypts (Eucalyptus globulus Labill), Wood Fiber Sci, 20 (1988), 82–90
32. Miranda, I., Pereira, H., Kinetics of ASAM and kraft pulping of eucalypt wood (Eucalyptus globulus), Holzforschung, 56 (2002), 85–90
33. Garrote, G., Dominguez, H., Parajo, J.C., Mild autohydrolysis: an environmentally friendly technology for xylooligosaccharide production from wood, J. Chem. Technol. Biotechnol, 74 (1999), 1101–1109
32
34. Conner, A.H., Kinetic modeling of hardwood prehydrolysis. Part I. Xylan removal by water prehydrolysis, Wood Fiber Sci, 16 (1984), 268–277
35. Schell, D.J., Ruth, M.F., Tucker, M.P., Modeling the enzymatic hydrolysis of dilute-acid pretreated Douglas fir, Appl. Biochem. Biotechnol, 77–79(1999), 67–81
36. Torget, R., Hsu, T.A., Two temperature dilute-acid prehydrolysis of hardwood xylan using a percolation process, Appl. Biochem. Biotechnol, 45–46(1994), 5–22
37. Tengborg, C., Stenberg, K., Galbe, M., Zacchi, G., Larsson, S., Palmqvist, E., Hahn-Hagerdal, B., Comparison of SO2 and H2SO4 impregnation of softwood prior to steam pretreatment on ethanol production, Appl. Biochem. Biotechnol, 70–72(1998), 3–15
38. Soderstrom, J., Galbe, M., Zacchi, G., Effect of washing on yield in one- and two-step steam pretreatment of softwood for production of ethanol, Biotechnol. Prog, 20 (2004), 744–749
39. Soderstrom, J., Pilcher, L., Galbe, M., Zacchi, G., Two-step steam pretreatment of softwood with SO2 impregnation for ethanol production, Appl. Biochem.Biotechnol, 98(2002), 5–21
40. Brillouet, J.M., Joseleau, J.P., Utille, J.P., Lelievre, D., Isolation, purification, and characterization of a complex heteroxylan from industrial wheat bran, J. Agri. Food Chem, 30(1982), 488–495
41. Shibuya, N., Iwasaki, T., Structural features of rice bran hemicellulose, Phytochemistry, 24(1985), 285–289
42. Ishii, T., Acetylation at O-2 of arabinofuranose residues in feruloylated arabinoxylan from bamboo shoot cell-walls, Phytochemistry, 30 (1991), 2317–2320
43. Wende, G., Fry, S.C., O-feruloylated, O-acetylated oligosaccharides as sidechains of grass xylans, Phytochemistry, 44 (1997), 1011–1018
44. Ebringerova, A., Hromadkova, Z., Heinze, T., Hemicellulose, Adv. Polym. Sci, 186 (2005), 1–67
45. Alen, R., Structure and chemical composition of wood. In: Stenius, P. (Ed.), Forest Products Chemistry. Fapet Oy, Helsinki, 2000
46. Pereira, H., Graca, J., Rodrigues, J.C., Wood chemistry in relation to quality. In: Barnett, J.R., Jeronimidis, G. (Eds.), Wood Quality and Its Biological Basis. Blackwell Publishing, Oxford, 2003
47. Timell, T.E., Wood hemicelluloses, Adv. Carbohydr. Chem. Biochem, 20 (1965), 409–483
48. Woodward, J., Xylanases: Functions, Properties and Applications. Ellis Horwood Ltd., Chichester, 1984
49. de Vries, R.P., Visser, J., Aspergillus enzymes involved in degradation of plant cell wall polysaccharides, Microbiol. Mol. Biol Rev, 65 (2001), 497
50. Powlowski, J., Mahajan, Sonam., Schapira, M., and Master, E.R. Substrate recognition and hydrolysis by a fungal xyloglucan-specific family 12 hydrolase. Carbohydr Res., 344(2009), 1175-1179
51. Maruyama, K., Goto, C., Numata, M., Suzuki, T., Nakagawa, Y., Hoshino, T., Uchiyama,T., O-acetylated xyloglucan in extracellular polysaccharides from cell suspension cultures of Mentha, Phytochemistry, 41 (1996), 1309–1314
33
52. Sims, I.M., Munro, S.L.A., Currie, G., Craik, D., Bacic, A., Structural characterisation of xyloglucan secreted by suspension-cultured cells of Nicotiana plumbaginifolia, Carbohydr. Res, 293 (1996), 147–172
53. Carpita, N.C., Gibeaut, D.M., Structural models of primary-cell walls in flowering plants – consistency of molecular-structure with the physical properties of the walls during growth, Plant J, 3 (1993), 1–30
54. Goyal, P., Kumar, V., and Sharma, P., Carboxymethylation of tamarind kernel powder, Carbohydr Polym, 69(2007), 251-255
55. Timell, T.E., Recent progress in the chemistry of wood hemicelluloses, Wood Sci. Technol, 1(1967), 45–70
56. Stephen, A.M., Other plant polysaccharides. In: Aspinall, G.O. (Ed.), The Polysaccharides. Academic Press, New York, 1983
57. Wyman, C.E., Biomass ethanol: technical progress, opportunities and commercial challenges, Annu.Rev.Energy.Environ, 24(1999), 189-226
58. Goldstein, I.S., Easter, J.M., An improved process for converting cellulose to ethanol, Tappi, 75(1992), 135–140
59. Koullas, D.P., Christakopoulos, P.E., Kekos, D., Koukios, E.G., Macris, B.J., Effect of alkali delignification on wheat straw saccharification by Fusarium oxysporum cellulases, Biomass Bioenergy, 4(1993), 9–13
60. Clark, D.P., Mackie, K.L., Steam explosion of the softwood Pinus radiata with sulphur dioxide addition. 1. Process optimization, J Wood Chem Technol, 7(1987), 373–403
61. Gould, J.M., Alkaline peroxide delignification of agricultural residues to enhance enzymatic saccharification, Biotechnol Bioeng, 26(1984), 46–52
62. Fernandez-Bolanos, J., Felizon, B., Heredia, A., Rodriguez, R., Guillen, R., Jimenez, A., Steam-explosion of olive stones: hemicellulose solubilization and enhancement of enzymatic hydrolysis of cellulose, Bioresour Technol, 79(2001), 53–61
63. Dale, B.E., Leong, C.K., Pham, T.K., Esquivel, V.M., Rios, L., Latimer, V.M., Hydrolysis at low enzyme levels: application of the AFEX process, Bioresour Technol, 56(1996), 111–116
64. Schmidt, A.S., Thomsen, A.B., Optimization of wet oxidation pretreatment of wheat straw, Bioresour Technol, 64(1998), 139–151
65. Kaar, W.E., Holtzaple, M.T., Using lime pretreatment to facilitate the enzymatic hydrolysis of corn stover, Biomass Bioenergy, 18(2000), 189–199
66. Laser, M., Schulman, D., Allen, S.G., Lichwa, J., Antal, M.J. Jr., Lynd, L.R.A., Comparison of liquid hot water and steam pretreatments of sugar cane bagasse for bioconversion to ethanol, Bioresour Technol, 81(2002), 33–44
67. Dale, B.E., Moreira, M.J., A freeze-explosion technique for increasing cellulose hydrolysis, Biotechnol Bioeng Symp, 12(1982), 31–43
68. Chum, H.L., Johnsoon, D.K., Black, S., Organosolv pretreatment for enzymatic hydrolysis of poplars: 1, enzyme hydrolysis of cellulosic residues, Biotechnol Bioeng, 31(1988), 643–649
69. Ohno, H., Fukaya, Y., Task specific ionic liquids for cellulose technology, Chem.Lett, 38 (2009), 2–7
34
70. Zavrel, M., Bross, D., Funke, M., Buchs, J., Spiess, A.C., High-throughput screening for ionic liquids dissolving (ligno-)cellulose, Bioresour. Technol, 100(2009), 2580–2587
71. Miyafuji, H., Nakata, T., Ehara, K., Saka, S., Fermentability of water-soluble portion to ethanol obtained by supercritical water treatment of lignocellulosics, Appl. Biochem. Biotechnol, 121(2005), 963–971
72. Girio, F.M., Fonseca, C., Carvalheiro, F., Duarte, L.C., Marques, S., Bogel-Lukasik, R., Hemicelluloses for fuel ethanol: A review, Biores. Technol, 101(2010), 775-800
73. Olsson, L., Hahn-Hagerdal, B., Fermentation of lignocellulosic hydrolysates for ethanol production, Enzyme Microb. Technol, 18 (1996), 312–331
74. Harris, J.F., Baker, A.J., Conner, A.H., Jeffries, T.W., Minor, J.L., Patterson, R.C., Scott, R.W., Springer, E.L., Zorba, J., Two-Stage Dilute Sulfuric Acid Hydrolysis of Wood: An Investigation of Fundamentals. General Technical Report FPL-45, U.S. Forest Products Laboratory, Madison, Wisconsin, 1985
75. Nguyen, Q., Milestone Completion Report: Evaluation of a Two-Stage Dilute Sulfuric Acid Hydrolysis Process. Internal Report, National Renewable Energy Laboratory, Golden, Colorado, 1998
76. Allen, S.G., Schulman, D., Lichwa, J., Antal, M.J., Jennings, E., Elander, R., A comparison of aqueous and dilute-acid single-temperature pretreatment of yellow poplar sawdust, Ind. Eng. Chem. Res, 40 (2001), 2352–2361
77. Carvalheiro, F., Duarte, L.C., Medeiros, R., Girio, F.M., Optimization of brewery’s spent grain dilute-acid hydrolysis for the production of pentose-rich culture media, Appl. Biochem. Biotechnol, 113–116 (2004), 1059–1072
78. Marzialetti, T., Olarte, M.B.V., Sievers, C., Hoskins, T.J.C., Agrawal, P.K., Jones, C.W., Dilute acid hydrolysis of Loblolly pine: a comprehensive approach, Ind. Eng. Chem. Res, 47 (2008), 7131–7140
79. Monavari, S., Galbe, M., Zacchi, G., The influence of solid/liquid separation techniques on the sugar yield in two-step dilute acid hydrolysis of softwood followed by enzymatic hydrolysis, Biotechnol. Biofuels, 2 (2009), 6
80. Wenzl, H.F.J., The Acid Hydrolysis of Wood. In The Chemical Technology of Wood, Academic Press, New York, 1970
81. Broder, J.D., Barrier, J.W., Lightsey, G.R., Conversion of cotton trash and other residues to liquid fuel. In liquid fuels from renewable resources: Proceedings of an alternative energy conference (Cundiff, J.S., ed), American Society of Agricultural Engineers, St. Joseph, MI, 1992
82. Wright, J.D., d'Agincourt, C.G., "Evaluation of Sulfuric Acid Hydrolysis Processes for Alcohol Fuel Production." Biotechnology and Bioengineering Symposium, No 14, John Wiley and Sons, New York, 1984, 105-123
83. Brink, D.L., Enzymatic hydrolysis of biomass material, US Patent: 5366558, 1997
84. Clausen, E.C., Gaddy, J.L., Concentrated sulfuric acid process for converting lignocellulosic materials to sugars, US patent: 5188673, 1993
35
85. Sivers, M.V., Zacchi, G., A techno-economical comparison of three processes for the production of ethanol from pine, Bioresour. Technol, 51(1995), 43–52
86. Torget, R.W., Padukone, N., Hatiz, C., Wyman, C.E., Hydrolysis and fractionation of lignocellulosic biomass, US patent: 6022419, 2000
87. Scott, D.S., Piskorz, J., Process for the production of fermentable sugars from biomass, US patent: 4880473, 1989
88. Aoyama, M., Seki, K., Saito, N., Solubilization of bamboo grass xylan by steaming treatment, Holzforschung, 49 (1995), 193–196
89. Carvalheiro, F., Silva-Fernandes, T., Duarte, L.C., Girio, F.M., Wheat straw autohydrolysis: process optimization and products characterization, Appl. Biochem. Biotechnol, 153 (2009), 84–93
90. Conner, A.H., Lorenz, L.F., Kinetic modeling of hardwood prehydrolysis. Part III. Water and dilute acetic-acid prehydrolysis of southern red oak, Wood Fiber Sci, 18 (1986), 248–263
91. Garrote, G., Parajo, J.C., Non-isothermal autohydrolysis of Eucalyptus wood, Wood Sci. Technol, 36 (2002), 111–123
92. Carvalheiro, F., Esteves, M.P., Parajo, J.C., Pereira, H., Girio, F.M., Production of oligosaccharides by autohydrolysis of brewery’s spent grain, Bioresour. Technol, 91 (2004), 93–100
93. Garrote, G., Dominguez, H., Parajo, J.C., Mild autohydrolysis: an environmentally friendly technology for xylooligosaccharide production from wood, J. Chem. Technol. Biotechnol, 74 (1999), 1101–1109
94. Wei, Sun Zhan, Chen, Zhang Hang, Hue, Yan Wang, Yu, Run Ma, Study on enzymatic hydrolysis of steam treated straw using a ball mill shaker, J. Beijing Univ. Chem. Technol, 33 (2006), 26–30
95. Ballesteros, I., Oliva, J.M., Negro, M.J., Manzanares, P., Ballesteros, M., Enzymic hydrolysis of steam exploded herbaceous agricultural waste (Brassica carinata) at different particule sizes, Process Biochem, 38 (2002), 187–192
96. Heitz, M., Capek-Menard, E., Koeberle, P.G., Gagne, J., Chornet, E., Overend, R.P., Taylor, J.D., Yu, E., Fractionation of Populus tremuloides at the pilot-plant scale – optimization of steam pretreatment conditions using the stake-II technology, Bioresour. Technol, 35 (1991), 23–32
97. Martin, C., Marcet, M., Thomsen, A.B., Comparison between wet oxidation and steam explosion as pretreatment methods for enzymatic hydrolysis of sugarcane bagasse, Bioresources, 3 (2008), 670–683
98. Ruiz, E., Cara, C., Manzanares, P., Ballesteros, M., Castro, E., Evaluation of steam explosion pre-treatment for enzymatic hydrolysis of sunflower stalks, Enzyme Microb. Technol, 42 (2008), 160–166
99. Cara, C., Ruiz, E., Ballesteros, M., Manzanares, P., Negro, M.J., Castro, E., Production of fuel ethanol from steam-explosion pretreated olive tree pruning, Fuel, 87 (2008), 692–700
100. Shallom, D., Shoham, Y., Microbial hemicellulases, Curr. Opin. Microbiol, 6 (2003), 219–228
36
101. McMillan, J.D., Pretreatment of lignocellulosic biomass. In: Himmel ME,. Baker JO, Overend RP, Editors, Enzymatic Conversion of Biomass for Fuels Production, American Chemical Society, Washington, DC, 1994, 292–324
102. Berlin, A., Gilkes, N., Kilburn, D., Bura, R., Markov, A., Skomarovsky, A., Okunev, O., Gusakov, A., Maximenko, V., Gregg, D., Sinitsyn, A., Saddler, J., Evaluation of novel fungal cellulase preparations for ability to hydrolyze softwood substrates – evidence for the role of accessory enzymes, Enzyme Microb Technol, 37(2005), 175–184
103. Li, K.C., Azadi, P., Collins, R., Tolan, J., Kim, J.S., Eriksson, K.E.L., Relationships between activities of xylanases and xylan structures, Enzyme Microb. Technol, 27 (2000), 89–94
104. Polizeli, M.L.T.M., Rizzatti, A.C.S., Monti, R., Terenzi, H.F., Jorge, J.A., Amorim, D.S., Xylanases from fungi: properties and industrial applications, Appl. Microbiol. Biotechnol, 67 (2005), 577–591
105. Sunna, A., Antranikian, G., Xylanolytic enzymes from fungi and bacteria, Crit. Rev. Biotechnol, 17 (1997), 39–67
106. Collins, T., Gerday, C., Feller, G., Xylanases, xylanase families and extremophilic xylanases, FEMS Microbiol. Rev, 29 (2005), 3–23
107. Beg, Q.K., Kapoor, M., Mahajan, L., Hoondal, G.S., Microbial xylanases and their industrial applications: a review, Appl. Microbiol. Biotechnol, 56 (2001), 326–338
108. Kumar, R., Singh, S., Singh, O.V., Bioconversion of lignocellulosic biomass: biochemical and molecular perspectives, J. Ind. Microbiol. Biotechnol, 35 (2008), 377–391
109. Khasin, A., Alchanati, I., Shoham, Y., Purification and characterization of a thermostable xylanase from Bacillus stearothermophilus T-6, Appl. Environ. Microbiol, 59 (1993), 1725–1730
110. Dhawan, S., Kaur, J., Microbial mannanases: an overview of production and applications, Crit. Rev. Biotechnol, 27 (2007), 197–216
111. Wyman, C.E., Potential synergies and challenges in refining cellulosic biomass to fuels, chemicals, and power, Biotechnol. Prog, 19 (2003), 254–262
112. Paulechka, Y.U., Kabo, G.J., Blokhin, A.V., Vydrov, O.A., Magee, J.W., Frenkel, M., Thermodynamic properties of 1-butyl-3-methylimidazolium hexafluorophosphate in the ideal gas state, J. Chem. Eng. Data, 48 (2003), 457–462
113. Talbot, G., Sygusch, J., Purification and characterization of thermostable β-mannanase and α-galactosidase from Bacillus stearothermophilus, Appl. Environ. Microbiol, 56 (1990), 3505–3510
114. Hatada, Y., Takeda, N., Hirasawa, K., Ohta, Y., Usami, R., Yoshida, Y., Grant, W.D., Ito,S., Horikoshi, K., Sequence of the gene for a high-alkaline mannanase from an alkaliphilic Bacillus sp. strain JAMB-750, its expression in Bacillus subtilis and characterization of the recombinant enzyme, Extremophiles, 9 (2005), 497–500
115. Morris, D.D., Reeves, R.A., Gibbs, M.D., Saul, D.J., Bergquist, P.L., Correction of the β-mannanase domain of the cell C pseudogene from Caldocellulosiruptor saccharolyticus and activity of the gene product on kraft pulp, Appl. Environ. Microbiol, 61 (1995), 2262–2269
116. Sunna, A., Gibbs, M.D., Chin, C.W.J., Nelson, P.J., Bergquist, P.L., A gene encoding a novel multidomain β-1,4-mannanase from Caldibacillus cellulovorans
37
and action of the recombinant enzyme on kraft pulp, Appl. Environ. Microbiol, 66 (2000), 664–670
117. Zhang, Q., Yan, X., Zhang, L., Tang, W., Cloning, sequence analysis, and heterologous expression of a β-mannanase gene from Bacillus subtilis Z-2, Mol. Biol, 40 (2006), 368–374
118. Deshpande, V., Lachke, A., Mishra, C., Keskar, S. and Rao, M., Mode of action and properties of xylanase and L- xylosidase from Neurospora crassa, Biotechnol. Bioeng, 26(1986), 1832-1837
119. Howard, R. L., Abotsi, E., Jansen van Rensburg, E. L., Howard, S., Lignocellulose biotechnology: issues of bioconversion and enzyme production, African Journal of Biotechnology, 2(2003), 602-619
120. Vincken, J.P., Beldman, G., Voragen, A.G., Substrate specificity of endoglucanases: what determines xyloglucanase activity?, Carbohydr Res, 298(1997), 299–310
121. Edwards, M., Dea, I.C., Bulpin, P.V., Reid, J.S., Purification and properties of a novel xyloglucan-specific endo-(1-4)-β-D-glucanase from germinated nasturtium seeds (Tropaeolum majus L.), J Biol Chem, 261(1986), 9489–9494
122. Pauly, M., Andersen, L.N., Kauppinen, S., Kofod, L.V., York, W.S., Albersheim, P., Darvill, A., A xyloglucanspecific endo-β-1,4-glucanase from Aspergillus aculeatus: expression cloning in yeast, purification and characterization of the recombinant enzyme, Glycobiology, 9(1999), 93–100.
123. Henrissat, B., A classification of glycosyl hydrolases based on amino acid sequence similarities, Biochem J, 280(1991), 309–316
124. Henrissat, B., Bairoch, A., New families in the classification of glycosyl hydrolases based on amino acid sequence similarities, Biochem J, 293(1993), 781–788
125. Henrissat, B., Bairoch, A., Updating the sequencebased classification of glycosyl hydrolases, Biochem J, 316(1996), 695–696
126. Takuya, I., Katsuro, Y., Ayako, H., Kiyohiko, I., Masahiro, S., Substrate recognition by glycoside hydrolase family 74 xyloglucanase from the basidiomycete Phanerochaete chrysosporium, FEBS Journal, 274 (2007) 5727–5736
127. Yaoi, K., Nakai, T., Kameda, Y., Hiyoshi, A., Mitsuishi, Y., Cloning and characterization of two xyloglucanases from Paenibacillus sp. strain KM21, Appl Environ Microbiol, 71(2005), 7670–7678
128. Yaoi, K., Mitsuishi, Y., Purification, characterization, cDNA cloning, and expression of a xyloglucan endoglucanase from Geotrichum sp. M128, FEBS Lett, 560(2004), 45–50
129. Yaoi, K., Mitsuishi, Y., Purification, characterization, cloning, and expression of a novel xyloglucan-specific glycosidase, oligoxyloglucan reducing end-specific cellobiohydrolase, J Biol Chem, 277(2002), 48276–48281
130. Irwin, D.C., Cheng, M., Xiang, B., Rose, J.K., Wilson, D.B., Cloning, expression and characterization of a family-74 xyloglucanase from Thermobifida fusca, Eur J Biochem, 270(2003), 3083–3091
38
131. Hasper, A.A., Dekkers, E., van Mil, M., van de Vondervoort, P.J., de Graaff, L.H., EglC, a new endoglucanase from Aspergillus niger with major activity towards xyloglucan, Appl Environ Microbiol, 68(2002), 1556– 1560
132. Grishutin, S.G., Gusakov, A.V., Markov, A.V., Ustinov, B.B., Semenova, M.V., Sinitsyn, A.P., Specific xyloglucanases as a new class of polysaccharide-degrading enzymes, Biochim Biophys Acta, 1674(2004), 268–281
133. Chhabra, S.R., Kelly, R.M., Biochemical characterization of Thermotoga maritima endoglucanase Cel74 with and without a carbohydrate binding module (CBM), FEBS Lett, 531(2002), 375–380
134. Bauer, S., Vasu, P., Mort, A.J., Somerville, C.R., Cloning, expression, and characterization of an oligoxyloglucan reducing end-specific xyloglucanobiohydrolase from Aspergillus nidulans, Carbohydr Res, 340(2005), 2590– 2597
135. Benko, Z., Siika-aho, M., Viikari, L., Reczey, K., Evaluation of the role of xyloglucanase in the enzymatic hydrolysis of lignocellulosic substrates, Enzyme Microb Technol, 43(2008), 109-114
136. Gloster, T.M., Ibatullin, F.M., Macauley, K., Eklo, J.M., Roberts, S., Turkenburg, J.P., Bjørnvad, M.E., Jørgensen, P., Danielsen, S., Johansen, K.S., Borchert, T.V., Wilson, K.S., Brumer, H., Davies, G.J., Characterization and Three-dimensional Structures of Two Distinct Bacterial Xyloglucanases from FamiliesGH5 and GH12*, J Biol Chem, 282(2007), 19177-19189
137. Fullbrook, P.D., Practical limits and prospects (kinetics). In: Godfrey T, West S (eds) Industrial enzymology, 2nd edn. MacMillan, London, 1996, 508–509
138. Viikari, L., Alapuranen, M., Puranen, T., Vehmaanperä, J., Siika-aho, M., Thermostable Enzymes in Lignocellulose Hydrolysis, Adv.Biochem. Engin/Biotechnol, 108(2007), 121–145
139. Menon, V., Prakash, G., Prabhune, A., Rao, M., Biocatalytic approach for the utilization of hemicellulose for ethanol production from agricultural residue using thermostable xylanase and thermotolerant yeast, Bioresource technology, 101(2010), 5366-5373
140. Wright, J.D., Wyman, C.E., Grohmann, K., Simultaneous saccharification and fermentation of lignocellulose, Appl. Biochem. Biotechnol, 18(1988), 75-90
141. Rudolf, A., Baudel, H., Zacchi, G., Hahn-Hagerdal, B., Liden, G., Simultaneous saccharification and fermentation of steam-pretreated bagasse using Saccharomyces cerevisiae TMB3400 and Pichia stipitis CBS6054, Biotechnol. Bioeng, 99 (2008), 783-790
142. Telli-Okur, M., Eken-Saracoglu, N., Fermentation of sunflower seed hull hydrolysate to ethanol by Pichia Stipitis, Bioresour.Technol, 99(2008), 2162-2169
143. Cheng, K.K., Cai, B., Zhang, J., Ling, H., Zhou, Y., Ge, J., Xu, J., Sugarcane bagasse hemicellulose hydrolysate for ethanol production by acid recovery process, Biochemical Engineering Journal, 38(2008), 105-109
144. Gupta, R., Sharma, K.K., Kuhad, R.C., Separate hydrolysis and fermentation (SHF) of Prosopis juliflora, a woody substrate, for the production of cellulosic ethanol by Saccharomyces cerevisiae and Pichia stipitis-NCIM 3498, Bioresour.Technol, 100(2009), 1214-1220
39
145. Kumar, A., Singh, L.K., Ghosh, S., Bioconversion of lignocellulosic fraction of water-hyacinth (Eichhornia crassipes) hemicellulose acid hydrolysate to ethanol by Pichia stipitis, Bioresour.Technol, 100(2009), 3293-3297
146. Diaz, M.J., Ruiz, E., Romero, I., Cara, C., Moya, M., Castro, E., Inhibition of Pichia stipitis fermentation of hydrolysates from olive tree cuttings, World.J.Microbiol.Biotechnol, 25(2009), 891-899
147. Cho, D.H., Shin, S-J., Bae, Y., Park, C., Kim, Y.H., Enhanced ethanol production from deacetylated yellow poplar acid hydrolysate by Pichia stipitis, Bioresource Technology, 101 (2010) 4947–4951
148. Menon, V., Prakash, G., Rao, M., Enzymatic hydrolysis and ethanol production using xyloglucanase and Debaromyces hansenii from tamarind kernel powder: galactoxyloglucan predominant hemicellulose, Journal of Biotechnology 148 (2010) 233–239
149. Wilkins, M.R., Mueller, M., Eichling, S., Banat, I.M., Fermentation of xylose by the thermotolerant yeast strains Kluyveromyces marxianus IMB2, IMB4, and IMB5 under anaerobic conditions, Process.Biochem, 43(2008), 346-350
150. Ryabova, O. B., Chmil, O.M., Sibirny, A.A., Xylose and cellobiose fermentation to ethanol by the thermotolerant methylotrophic yeast Hansenula polymorpha, FEMS Yeast Res, 4(2003), 157–164
151. Breuer, U., Harms, H., Debaryomyces hansenii - an extremophilic yeast with biotechnological potential, Yeast, 23(2006), 415-437
152. Hogsett, D.A., Ahn, H.J., Bernardez, T.D., South, C.R., Lynd, L.R., Direct microbial conversion: prospects, progress and obstacles, Appl. Biochem. Biotechnol, 34/35(1992), 527-541
153. Klapatch, T.R., Hogsett, D.A.L., Baskaran, S., Pal, S., Lynd, L.R., Organism development and characterization for ethanol production using thermophilic bacteria, Appl. Biochem. Biotechnol, 45/46(1994), 209-213
154. Deshpande, V.V., Keskar, S., Mishra, C., Rao, M., Direct conversion of cellulose/hemicellulose to ethanol by Neurospora crassa, Enz. Microb. Technol, 8(1986), 149
155. Glazer, N.A., Nikaido, H., Ethanol. In: Microbial Biotechnology. Freeman WH. and company, San Fransisco, 1995, 359-391
156. Olofsson, K., Bertilsson, M., Lidén, G., A short review on SSF – an interesting process option for ethanol production from lignocellulosic feedstocks, Biotechnology for Biofuels, 1(2008), 7
157. Hyvonen, L., Koivistoinen, P. and Voirol, F., Food technological evaluation of xylitol, Adv. Food. Res, 28(1982), 373–403
158. Edgar, W.M., Sugar substitutes, chewing gum and dental caries-a review, Br Dent J, 184(1998), 29-32
40
159. Makinen, K.K., Dietary prevention of dental caries by xylitol—clinical effectiveness and safety, J. Appl. Nutrition, 44(1992), 16–28
160. Emodi, A., Xylitol, its properties and food application, Food Technology, 32(1978), 20–32
161. Uhari, M., Kontiokari, T., Niemela, M., A novel use of xylitol sugar in preventing acute otitis media, Pediatrics, 102(1998), 879–884
162. Kadam, K.L., Chin, C.Y., Brown, L.W., Flexible biorefinery for producing fermentation sugars, lignin and pulp from corn stover, J. Ind. Microbiol. Biotechnol, 35(2008), 331-34
163. Winkelhausen, E. and Kuzmanova, S., Microbial conversion of D-xylose to xylitol, J. Ferment.Bioeng, 86 (1998), 1-14.
164. Santos, J.C., Mussatto, S.I., Dragone, G., Converti, A., Silva, S.S., Evaluation of porous glass and zeolite as cells carriers for xylitol production from sugarcane bagasse hydrolysate, Biochem. Eng. J, 23(2005), 1–9
165. Verduyn, C., Van Kleef, R., Frank, J., Schreuder, H., van Dijken, J.P. and Scheffers, W.A., Properties of the NAD(P)H-dependent xlose reductase from the xylose fermenting yeast, Pichia stipitis, Biochem. J, 226(1985), 669-677
166. Rizzi, M., Harwart, K., Erlemann, P., Bui-Thanh, N.A. and Dellweg, H., Purification and properties of the NAD+-xylitol-dehydrogenase from the yeast Pichia stipitis, J. Ferment. Bioeng, 67(1989), 20-24
167. Kotter, P., and Ciriacy, M., Xylose fermentation by Saccharomyces cerevisiae, Appl. Microbiol. Biotechnol, 38(1993), 776-783
168. Nigam, P., Singh, K., Processes of fermentative production of Xylitol - a sugar substitute, Process biochemistry, 30(1995), 117-124
169. Villarreal, M.L.M., Prata, A.M.R., Felipe, M.G.A., Silva, J.B. Almeida E., Detoxification procedures of eucalyptus hemicellulose hydrolysate for xylitol production by Candida guilliermondii, Enzyme and Microbial Technology, 40 (2006) 17–24
170. Rao, R.S., Jyothi, C.P., Prakasham, R.S., Sarma, P.N., Rao, L.V., Xylitol production from corn fiber and sugarcane bagasse hydroysates by Candida tropicalis, Bioresource Technol, 97(2006), 1974–1978
171. Carvalheiro, F., Duarte,L.C., Medeiros, R., Girio, F.M., Xylitol production by Debaryomyces hansenii in brewery spent grain dilute-acid hydrolysate: effect of supplementation, Biotechnology Letters, 29(2007), 1887-1891
172. Liaw, W.C., Chen, C.S., Chang, W.S., Chen, K.P., Xylitol production from rice straw hemicellulose hydrolyzate by polyacrylic hydrogel thin films with immobilized Candida subtropicalis WF79, J Biosci Bioeng, 105(2008), 97–105
173. Canilha, L., Carvalho, W., Felipe, M.G.A., Silva, J.B.A., Xylitol production from wheat straw hemicellulosic hydrolysate:hydrolysate detoxification and carbon source used for inoculum preparation, Brazilian Journal of Microbiology, 39 (2008), 333-336
41
174. Cheng, K.K., Zhang, J.A., Chavez, E., Li, J-P., Integrated production of xylitol and ethanol using corncob, Applied Microbiology and Biotechnology, 87(2010), 411-417
175. Kosaric, N., Magee, R.J., Blaszczyk, R., Redox potential measurement for monitoring glucose and xylose conversion by K. pneumoniae, Chem Biochem Eng Q, 6(1992), 145–152
176. Tran, A.V., Chambers, R.P., The dehydration of fermentative 2,3-butanediol into methyl ethyl ketone, Biotechnol Bioeng, 29(1987), 343–351
177. Willetts, A., Butane 2,3-diol production by Aeromonas hydrophila grown on starch, Biotechnol Lett, 6(1984), 263–8
178. Groleau, D., Laube, V.M., Martin, S.M., The effect of various atmospheric conditions on the 2,3-butanediol fermentation from glucose by Bacillus polymyxa, Biotechnol Lett, 7(1985), 53–8
179. Nakashimada, Y., Marwoto, B., Kashiwamura, T., Kakizono, T., Nishio, N., Enhanced 2,3-butanediol production by addition of acetic acid in Paenibacillus polymyxa, J Biosci Bioeng, 90(2000), 661–4
180. Kosaric, N., Velikonja, J., Liquid and gaseous fuels from biotechnology: challenges and opportunities, FEMS Microbiol Rev 16(1995), 111–142
181. Yu, E.K.C., Saddler, J.N., Enhanced production of 2,3-butanediol by Klebsiella pneumoniae grown on high sugar concentrations in the presence of acetic acid, Appl Environ Microbiol, 44(1982), 777–84
182. Qureshi, N., Cheryan, M., Effect of lactic acid on growth and butanediol production by Klebsiella oxytoca, J Industrial Microbiol, 4(1989), 453–6
183. Cao, N.J., Xia, Y.K., Gong, C.S., Tsao, G.T., Production of 2,3-butanediol from pretreated corncob by Klebsiella oxytoca in the presence of fungal cellulose, Appl Biochem Biotechnol, 63(1997), 129–39
184. Qin, J.Y., Xiao, Z.J., Ma, C.Q., Xie, N.Z., Liu, P.H., Xu, P., Production of 2,3-Butanediol by Klebsiella Pneumoniae using glucose and ammonium phosphate, Chinese J Chem Eng, 14(2006), 132–6
185. Ji, X.J., Huang, H., Li, S., Du, J., Lian, M., Enhanced 2,3-butanediol production by altering the mixed acid fermentation pathway in Klebsiella oxytoca,. Biotechnol Lett, 30(2008), 731–4
186. Cheng, K.K., Liu, Q., Zhang, J-A., Li, J-P., Xu, J-M., Wang, G-H., Improved 2,3-butanediol production from corncob acid hydrolysate by fed-batch fermentation using Klebsiella oxytoca, Process Biochemistry, 45 (2010), 613–616
187. Perego, P., Converti, A., Borghi, M.D., Effects of temperature, inoculum size and starch hydrolyzate concentration on butanediol production by Bacillus licheniformis, Bioresour Technol, 89(2003), 125–31
188. Afschar, A.S., Bellgardt, K.H., Rossell, C.E., The production of 2,3-butanediol by fermentation of high test molasses, Appl Microbiol Biotechnol, 34(1991), 582–5
189. Motwani, M., Seth, R., Daginawala, H.F., Khanna, P., Microbial-production of 2,3- butanediol from water hyacinth, Bioresour Technol, 44(1993), 187–95
190. Sun, L.H.,Wang, X.D., Dai, J.Y., Xiu, Z.L., Microbial production of 2,3-butanediol from Jerusalem artichoke tubers by Klebsiella pneumoniae, Appl Microbiol Biotechnol, 82(2009), 847–52
42
191. Grover, B.P., Garg, S.K., Verma, J., Production of 2,3-butanediol from wood hydrolysate by Klebsiella pneumoniae, World J Microbiol Biotechnol, 6(1990), 328–32
192. Weidner, S., Amarowicz, R., Karamać, M., Dąbrowski, G., Phenolic acids in caryopses of two cultivars of wheat, rye and triticale that display different resistance to pre-harvest sprouting, European Food Research and Technology, 210(1999), 109–113
193. Weidner, S., Amarowicz, R., Karamać, M., Frączek, E., Changes in endogenous phenolic acids during development of Secale cereale caryopses and after dehydration treatment of unripe rye grains, Plant Physiology and Biochemistry, 38(2000), 595–602
194. Weidner, S., Krupa, U., Amarowicz, R., Karamać, M., Abe, S., Phenolic compounds in embryos of triticale caryopses at different stages of development and maturation in normal environment and after dehydration treatment, Euphytica, 126(2002), 115–122
195. Kim, K.H., Tsao, R., Yang, R., Cui, S.W., Phenolic acid profiles and antioxidant activities of wheat bran extracts and the effect of hydrolysis conditions, Food Chemistry, 95(2006), 466–473
196. Steinhart, H., Renger, A., Influence of technological processes on the chemical structure of cereal dietary fiber, Czech Journal of Food Sciences, 8(2000), 22–24
197. Moore, J., Cheng, Z.H., Su, L., Yu, L.L.L., Effects of solid-state enzymatic treatments on the antioxidant properties of wheat bran, Journal of Agriculture and Food Chemistry, 54(2006), 9032–9045
198. Ou, S.Y., Kwok, K.C., Ferulic acid: pharmaceutical functions, preparation and applications in foods, Journal of the Science of Food and Agriculture, 84(2004), 1261–1269
199. Hirata, A., Murakami, Y., Atsumi, T., Shoji, M., Ogiwara, T., Shibuya, K., Ito, S., Yokoe, I., Fujisawa, S., Ferulic acid dimer inhibits lipopolysaccharide- stimulated cyclooxygenase-2 expression in macrophages, In Vivo, 19(2005), 849–853
200. Wang, F., Yang, L.X., Huang, K.X., Li, X.K., Hao, X.J., Stockigt, J., Zhao, Y., Preparation of ferulic acid derivatives and evaluation of their xanthine oxidase inhibition activity, Natural Product Research, 21(2007), 196–202
201. Topakas, E., Vafiadi, C., Christakopoulos, P., Microbial production, characterization and applications of feruloyl esterases, Process Biochemistry, 42 (2007), 497–509
202. Bartolome, B., Faulds, C.B., Kroon, P.A., Waldron, K., Gilbert, H.J., Hazlewood, G., Williamson, G., An Aspergillus niger esterase (ferulic acid esterase III) and a recombinant Pseudomonas fluorescens subsp. cellulosa esterase (XylD) released a 5-50 dihydrodimer (diferulic acid) from barley and wheat cell walls, Appl Environ Microbiol, 63(1997), 208–12
203. MacKenzie, C.R., Bilous, D., Ferulic acid esterase activity from Schizophyllum commune, Appl Environ Microbiol, 54(1988), 1170–3
204. Faulds, C.B., Williamson, G., Release of ferulic acid from wheat bran by a ferulic acid esterase (FAE III) from Aspergillus niger, Appl Microbiol Biotechnol, 43(1995), 1082–7
43
205. Ralet, M.C., Faulds, C.B., Williamson, G., Thibault, J-F., Degradation of feruloylated oligosaccharides from sugar-beet pulp and wheat bran by ferulic acid esterase from Aspergillus niger, Carbohydr Res, 263(1994), 257–69
206. Topakas, E., Christakopoulos, P., Faulds, C.B., Comparison of mesophilic and thermophilic feruloyl esterases: characterization of their substrate specificity for methyl phenylalkanoates, J Biotechnol,115(2005), 355–66
207. Topakas, E., Stamatis, H., Biely, P., Kekos, D., Macris, B.J., Christakopoulos, P., Purification and characterization of a feruloyl esterase from Fusarium oxysporum catalyzing esterification of phenolic acids in ternary waterorganic solvent mixtures, J Biotechnol, 102(2003), 33–44
208. Topakas, E., Stamatis, H., Mastihubova, M., Biely, P., Kekos, D., Macris, B.J., Christakopoulos, P., Purification and characterization of a Fusarium oxysporum feruloyl esterase (FoFAE-I) catalysing transesterification of phenolic acid esters, Enzyme Microb Technol, 33(2003),729–37
209. Topakas, E., Stamatis, H., Biely, P., Christakopoulos, P., Purification and characterization of a type B feruloyl esterase (StFAE-A) from the thermophilic fungus Sporotrichum thermophile, Appl Microbiol Biotechnol, 63(2004), 686–90
210. Topakas, E., Vafiadi, C., Stamatis, H., Christakopoulos, P., Sporotrichum thermophile type C feruloyl esterase (StFaeC): Purification characterization, and its use for phenolic acid (sugar) ester synthesis, Enzyme Microb Technol, 36(2005), 729–36
211. Ferreira, L.M.A., Wood, T.M., Williamson, G., Faulds, C.B., Hazlewood, G., Gilbert, H.J., A modular esterase from Pseudomonas fluorescens subsp. cellulosa contains a non-catalytic binding domain, Biochem J, 294(1993), 349–55
212. Faulds, C.B., Williamson, G., Ferulic acid esterase from Aspergillus niger: purification and partial characterization of two forms from a commercial source of pectinase, Biotechnol Appl Biochem, 17(1993), 349–59
213. Bartolome, B., Faulds, C.B., Tuohy, M., Hazlewood, G.P., Gilbert, H.J., Williamson, G., Influence of different xylanases on the activity of ferulic acid esterase on wheat bran, Biotechnol Appl Biochem, 22(1995), 65–73
214. Kroon, P.A., Garcia-Conesa, M.T., Fillingham, I.J., Hazlewood, G.P., Williamson, G., Release of ferulic acid dehydrodimers from plant cell walls by feruloyl esterases, J Sci Food Agric, 79(1999), 428–34
215. Topakas, E., Christakopoulos, P., Enzymic release of phenolic antioxidants from plant cell wall material, NutrCos, 1–2(2004), 54–7
216. Thibault, J-F., Asther, M., Ceccaldi, B.C., Couteau, D., Delattre, M., Duarte, J.C., Faulds, C.B., Heldt-Hansen, H.P., Kroon, P., Lesage-Meessen, L., Micard, V., Renard, C.M.G.C., Tuohy. M., Van Hulle, S., Williamson, G., Fungal bioconversion of agricultural by-products to vanillin, Lebensm Wiss Technol, 31(1998), 530–6
217. Benoit, I., Navarro, D., Marnet, N., Rakotomanomana, N., Lesage-Meessen, L., Sigoillot, J-C., Asther, M., Asther, M., Feruloyl esterases as a tool for the release of phenolic compounds from agro-industrial by-products, Carbohydr Res, 341(2006), 1820–7
44
218. Faulds, C.B., Kroon, P.A., Saulnier, L., Thibault, J.F., Williamson, G., Release of ferulic acid from maize bran and derived oligosaccharides by Aspergillus niger esterases, Carbohydr Polym, 27(1995), 187–90
219. Shin, H-D., McClendon, S., Le, T., Taylor, F., Chen, R.R., A complete enzymatic recovery of ferulic acid from corn residues with extracellular enzymes from Neosartorya spinosa NRRL 185, Biotechnol Bioeng, 95(2006), 1108–15
220. Bartolome, B., Faulds, C.B., Williamson, G., Enzymic release of ferulic acid from barley spent grain, J Cereal Sci, 25(1997), 285–8
221. Bartolome, B., Gomes-Cordoves, C., Sancho, A.I., Diez, N., Ferreira, P., Soliveri, J., Copa-Patino, J.L., Growth and release of hydroxycinnamic acids from Brewer’s spent grain by Streptomyces avermitilis CECT 3339, Enzyme Microb Technol, 32(2003), 140–4
222. Faulds, C.B., Sancho, A.I., Bartolome, B., Mono- and dimeric ferulic acid release from brewer’s spent grain by fungal feruloyl esterases, Appl Microbiol Biotechnol, 60(2002), 489–94
223. Faulds, C.B., Zanichelli, D., Crepin, V.F., Connerton, I.F., Juge, N., Bhat, M.K., Waldron, K.W., Specificity of feruloyl esterases for water-extractable and water-unextractable feruloylated polysaccharides: influence of xylanase, J Cereal Sci, 38(2003), 281–8
224. Faulds, C.B., Mandalari, G., LoCurto, R., Bisignano, G., Waldron, K.W., Arabinoxylan and mono- and dimeric ferulic acid release from brewer’s grain and wheat bran by feruloyl esterases and glycosyl hydrolases from Humicola insolens, Appl Microbiol Biotechnol, 64(2004), 644–50
225. Faulds, C.B., Mandalari, G., LoCurto, R., Bisignano, G., Christakopoulos, P., Waldron, K.W., Synergy between xylanases from glycoside hydrolase family 10 and family 11 and a feruloyl esterase in the release of phenolic acids from cereal arabinoxylan, Appl Microbiol Biotechnol, 71(2006), 622–9
226. Moore, J., Bamforth, C.W., Kroon, P.A., Bartolome, B., Williamson, G., Ferulic acid esterase catalyses the solubilization of b-glucans, and pentosans from the starchy endosperm cell walls of barley, Biotechnol Lett, 18(1996), 1423–6
227. Sancho, A.I., Faulds, C.B., Bartolome, B., Williamson, G., Characterization of feruloyl esterase activity in barley, J Sci Food Agric, 79(1999), 447–9
228. Kroon, P.A., Williamson, G., Release of ferulic acid from sugar-beet pulp by using arabinanase, arabinofuranosidase and an esterase from Aspergillus niger, Biotechnol Appl Biochem, 23(1996), 263–7
229. Ferreira, P., Diez, N., Gutierrez, C., Soliveri, J., Copa-Patino, J.L., Streptomyces avermitilis CECT 3339 produces a ferulic acid esterase able to release ferulic acid from sugar beet pulp soluble feruloylated oligosaccharides, J Sci Food Agric, 79(1999), 440–2
230. Kroon, P.A., Faulds, C.B., Williamson, G., Purification and characterisation of a novel esterase induced by growth of Aspergillus niger on sugar-beet pulp, Biotechnol Appl Biochem, 23(1996), 255–62
231. Brezillon, C., Kroon, P.A., Faulds, C.B., Brett, G.M., Williamson, G., Novel ferulic acid esterases are induced by growth of Aspergillus niger on sugar-beet pulp, Appl Microbiol Biotechnol, 45(1996), 371–6
45
232. Borneman, W.S., Hartley, R.D., Morrison, W.H., Akin, D.E., Ljungdahl, L.G., Feruloyl and p-coumaroyl esterase from anaerobic fungi in relation to plant cell wall degradation, Appl Microbiol Biotechnol, 33(1990), 345–51
233. Yu, P., Maenz, D.D., McKinnon, J.J., Racz, V.J., Christensen, D.A., Release of ferulic acid from oat hulls by Aspergillus ferulic acid esterase and Trichoderma xylanase. J Agric Food Chem, 50(2002), 1625–30
234. Yu, P., McKinnon, J.J., Maenz, D.D., Racz, V.J., Christensen, D.A., The interactive effects of enriched sources of Asperillus ferulic acid esterase and Trichoderma xylanase on the quantitative release of hydroxycinnamic acids from oat hulls, Can J Anim Sci, 82(2002), 251–7
235. Yu, P., Mcnmon, J.J., Maenz, D.D., Olkowski, A.A., Racz, V.J., Christensen, D.A., Enzymic release of reducing sugars from oat hulls by cellulase, as influenced by Aspergillus ferulic acid esterase and Trichoderma xylanase, J Agric Food Chem, 51(2003), 218–23
236. Laszlo, J.A., Compton, D.L., Li, X-L., Feruloyl esterases hydrolysis and recovery of ferulic acid from jojoba meal, Ind Crop Prod, 23(2006), 46–53
237. Converti, A., De Faveri, D., Perego, P., Barghini, P., Ruzzi, M., Sene, L., Vanillin production by recombinant strains of Escherichia coli, Braz J Microbiol, 34(2003), 108–10.
238. Torre, P., De Faveri, D., Perego, P., Converti, A., Barghini, P., Ruzzi, M., Selection of co-substrate and aeration conditions for vanillin production by Escherichia coli JM109/pBB1, Food Technol Biotechnol, 42(2004), 193–6
239. Torre, P., De Faveri, D., Perego, P., Ruzzi, M., Barghini, P., Gandolfi, R., Bioconversion of ferulate into vanillin by Escherichia coli strain JM109/pBB1 in an immobilized cell reactor, Ann Microbiol, 54(2004), 517–27
240. De Faveri, D., Torre, P., Aliakbarian, B., Domínguez, J.M., Perego, P., Converti, A., Response surface modeling of vanillin production by Escherichia coli JM109/pBB1, Biochem Eng J, 36(2007), 268–75
241. Walton, N.J., Mayer, M.J., Narbad, A., Vanillin, Phytochemistry, 63(2003), 505–15 242. Burri, J., Graf, M., Lambelet, P., Löiger, J., Vanillin: more than a flavouring
agent—a potent antioxidant, J Sci Food Agric, 48(1989), 49–56 243. Davidson, P.M., Naidu, A.S., Phyto-phenols. In: Naidu AS, editor. Natural food
antimicrobial systems, Boca Raton, London: CRC Press; 2000 244. Gould, G.W., Industry perspectives on the use of natural antimicrobials and
inhibitors for food applications, J Food Protect, 1996, 82–6 245. Priefert, H., Rabenhorst, J., Steinbüchel, A., Biotechnological production of
vanillin, Appl Microbiol Biotechnol, 56(2001), 296–314 246. Serra, S., Fuganti, C., Brenna, E., Biocatalytic preparation of natural flavours and
fragrances, Trends Biotechnol, 23(2005), 193–8 247. Yoon, S., Li, C., Kim, J., Lee, S., Yoon, J., Choi, M., Production of vanillin by
metabolically engineered Escherichia coli, Biotechnol Lett, 27(2005), 1829–32 248. Overhage, J., Steinbüchel, A., Priefert, H., Harnessing eugenol as a substrate for
production of aromaticcompounds with a recombinant strains of Amycolatopsis sp. HR167, J Biotechnol, 125(2006), 369–76
46
249. Sutherland, J.B., Crawford, D.L., Pometto III, A.L., Metabolism of cinnamic, p-coumaric and ferulic acids by Streptomyces setonii, Can J Microbiol, 29(1983), 1253–7
250. Gurujeyalakshmi, G., Mahadevan, A., Dissimilation of ferulic acid by Bacillus subtilis, Curr Microbiol, 16(1987), 69–73
251. Andreoni, V., Bernasconi, S., Bestetti, G., Biotransformation of ferulic acid and related compounds by mutant strains of Pseudomonas fluorescens, Appl Microbiol Biotechnol, 42(1995), 830–5
252. Barghini, P., Montebove, F., Ruzzi, M., Schiesser, A., Optimal conditions for bioconversion of ferulic acid into vanillic acid by Pseudomonas fluorescens BF13 cells, Appl Microbiol Biotechnol, 49(1998), 309–14
253. Narbad, A., Gasson, M.J., Metabolism of ferulic acid to vanillin using a novel CoA-dependent pathway in a newly-isolated strain of Pseudomonas fluorescens, Microbiology, 144(1998), 1397–405
254. Rabenhost, J., Hopp, R., Process for the preparation of vanillin and suitable microorganisms, European Patent 0761817; 1997.
255. Müller, B., Münch, T., Muheim, A.,Wetli, M., Process for the preparation of vanillin, European Patent 0885968; 1998
256. Gunnarsson, N., Palmqvist, E.A., Influence of pH and carbon source on the production of vanillin from ferulic acid by Streptomyces setonii ATCC 39116, Develop Food Sci, 43(2006), 73–6
257. Lesage-Meessen, L., Stentelaire, C., Lomascolo, A., Couteau, D., Asther, M,, Moukha, S., Fungal transformation of ferulic acid from sugar beet pulp to natural vanillin, J Sci Food Agric, 79(1999), 487–90
258. Bonnina, E., Brunel, M., Gouy, Y., Lesage-Meessen, L., Asther, M., Thibault, J-F., Aspergillus niger I-1472 and Pycnoporus cinnabarinus MUCL39533, selected for the biotransformation of ferulic acid to vanillin, are also able to produce cell wall polysaccharide-degrading enzymes and feruloyl esterases, Enzyme Microb Technol, 28(2001), 70–80
259. Lesage-Meessen, L., Lomascolo, A., Bonnin, E., Thibault, J.F., Buleon, A., Roller, M., A biotechnological process involving filamentous fungi to produce natural crystalline vanillin from maize bran, Appl Biochem Biotechnol, 102–103(2002), 141–53
260. Zheng, L., Zhenga, P., Sun, Z., Bai, Y., Wang, J., Guo, X., Production of vanillin from waste residue of rice bran oil by Aspergillus niger and Pycnoporus cinnabarinus, Bioresour Technol, 98(2007), 1115–9
261. Thibault, J., Micard, V., Renard, C., Asther, M., Delattre, M., Lesage-Meessen, L., Fungal bioconversion of agricultural by-products to vanillin, LWT-Food Sci Technol, 31(1998), 530–6
262. Di Gioia, D., Sciubba, L., Setti, L., Luziatelli, F., Ruzzi, M., Zanichelli, D., Production of biovanillin from wheat bran, Enzyme Microb Technol, 41(2007), 498–505
263. Torres, B.R., Aliakbariana, B., Torrea, P., Peregoa, P., Domínguezb, J.M., Zilli, M., Converti, A.,Vanillin bioproduction from alkaline hydrolyzate of corn cob by Escherichia coli JM109/pBB1, Enzyme and Microbial Technology, 44 (2009), 154–158
47
264. Brown, S.F., Bioplastic fantastic, Fortune, 148(2006), 92–94 265. Datta, R., Tsai, S., Bonsignore, P., Moon, S., Frank, J.R., Technological and
economic potential of poly(lactic acid) and lactic acid derivatives, FEMS Microbiol. Rev, 16(1995), 221–231
266. Tsuji, F., Autocatalytic hydrolysis of amorphous-made polylactides: effects of L-lactide content, tacticity, and enantiomeric polymer blending, Polymer, 43(2002), 1789–1796
267. Carr, F.J., Chill, D., Maida, N., The lactic acid bacteria: a literature survey, Crit. Rev. Microbiol, 28(2002), 281–370
268. Hofvendahl, K., Hahn-Hagerdal, B., Factors affecting the fermentative lactic acid production from renewable resources, Enz. Microb. Technol, 26(2000), 87–107
269. Chopin, A., Organization and regulation of genes for amino acid biosynthesis in lactic acid bacteria, FEMS Microbiol. Rev,12(1993), 21–38
270. Garde, A., Jonsson, G., Schmidt, A.S., Ahring, B.K., Lactic acid production from wheat straw hemicellulose hydrolysate by Lactobacillus pentosus and Lactobacillus brevis, Bioresour.Technol, 81(2002), 217–223
271. Tanaka, K., Komiyama, A., Sonomoto, K., Ishizaki, A., Hall, S.J., Stanbury, P.E., Two different pathways for D-xylose metabolism and the effect of xylose concentration on the yield coefficient of lactate in mixed-acid fermentation by the lactic acid bacterium Lactococcus lactis IO-1, Appl. Microbiol. Biotechnol, 60(2002), 160–167
272. Patel, M., Ou, M., Ingram, L.O., Shanmugam, K.T., Fermentation of sugar cane bagasse hemicellulose hydrolysate to l(+)-lactic acid by a thermotolerant acidophilic Bacillus sp, Biotechnology Letters, 26(2004), 865-868.
273. Woiciechowski, A.L., Soccol, C.R., Ramos, L.P., Pandey, A., Experimental design to enhance the production of l-(+)-lactic acid from steam-exploded wood hydrolysate using Rhizopus oryzae in a mixed-acid fermentation, Process Biochemistry, 34(1999), 949-955
274. Xu, Z., Wang, Q., Wang, P., Cheng, G., Ji, Y., Jiang, Z., Production of lactic acid from soybean stalk hydrolysate with Lactobacillus sake and Lactobacillus casei, Proc. Biochem, 42 (2007), 89–92
275. Wang, L., Zhao, B., Liu, B., Yu, B., Ma, C., Su, F., Hua, D., Li, Q., Ma, Y., Xu, P., Efficient production of l-lactic acid from corncob molasses, a waste by-product in xylitol production, by a newly isolated xylose utilizing Bacillus sp. strain, Bioresource Technology, 101(2010), 7908-7915
276. Bustos, G., Torre, N., Moldes, A.B., Cruz, J.M., Domínguez, J.M., Revalorization of hemicellulosic trimming vine shoots hydrolyzates trough continuous production of lactic acid and biosurfactants by L. pentosus , Journal of Food Engineering, 78(2007), 405-412
277. John, R.P., Nampoothiri, M., Pandey, A., Solid state fermentation for lactic acid production from agro waste using Lactobacillus delbrueckii, Process biochemistry, 41(2006), 759-763
278. John, R.P., Anisha, G.S., Nampoothiri, M., Pandey, A., Direct lactic acid fermentation: Focus on simultaneous saccharification and lactic acid production, Biotechnology Advances, 27(2009), 145-152
48
279. Ajit Singh Mamman, A.S., Lee, J-M., Kim, Y-C., Hwang, I.T., Park, N-J., Hwang, Y.K., Chang, J-S., Hwang, J-S., Furfural: Hemicellulose/xylosederived biochemical, Biofuels, Bioprod. Bioref, 2(2008), 438–454
280. Va´zquez, M., Oliva, M., Te´llez-Luis, S.J., Ramı´rez, J.S., Hydrolysis of sorghum straw using phosphoric acid: Evaluation of furfural production, Bioresource Technology 98 (2007) 3053–3060
281. Nicholsa, N.N., Sharmab, L.K., Moweryb, R.A., Chamblissb, C.K., Walsumc, G.P., Diena, B.S., Itena, L.B., Fungal metabolism of fermentation inhibitors present in corn stover dilute acid hydrolysate, Enzyme and Microbial Technology, 42 (2008) 624–630
282. Singh, A., Microbial production of acetone and butanol. Microbial Pentose Utilization Current Applications in Biotechnology. Elsevier Science, New York, 197–220, 1995
283. Parekh, S.R., Parekh, R.S., Wayman, M., Ethanol and butanol production by fermentation of enzymatically saccharified SO2-prehydrolysed lignocellulosics, Enzyme Microbial. Technol, 10(1988), 660–668
284. Marchal, R., Rebeller, M., Vandecasteele, J.P., Direct bioconversion of alkali-pretreated straw using simultaneous enzymatic hydrolysis and acetone butanol production, Biotechnol. Lett, 6(1984), 523–528
285. Soni, B.K., Das, K., Ghose, T.K., Bioconversion of agro-wastes into acetone butanol, Biotechnol. Lett, 4(1982), 19–22.
286. Nasib Qureshi, N., Ezeji, T.C., Ebener, J., Dien, B.S., Cotta, M.A., Blaschek, H.P.,Butanol production by Clostridium beijerinckii. Part I: Use of acid and enzyme hydrolyzed corn fiber, Bioresource Technology, 99 (2008) 5915–5922
287. Sun, Z., Liu, S., Production of n-butanol from concentrated sugar maple hemicellulosic hydrolysate by Clostridia acetobutylicum ATCC824, Biomass and Bioenergy, 2010, 1-9
288. Brentner, L., Peccia, J., Zimmerman, J., Challenges in Developing Biohydrogen as a Sustainable Energy Source: Implications for a Research Agenda, Environ. Sci. Technol, 44(2010), 2243–2254
289. Kaparaju, P., Serrano, M., Thomsen, A.B., Kongjan, P., Angelidaki, I., Bioethanol, biohydrogen, biogas production from wheat straw in a biorefinery concept, Bioresource Technology, 100(2009), 2562-2568
290. Kongjan, P., Angelidaki, I., Extreme thermophilic biohydrogen production from wheat straw hydrolysate using mixed culture fermentation: Effect of reactor configuration, Bioresource Technology, 101(2010), 7789-7796
291. Cao, G., Ren, N., Wang, A., Lee, D-L., Guo, W., Liu, B., Feng, L., Zhao, Q., Acid hydrolysis of corn stover for biohydrogen production using Thermoanaerobacterium thermosaccharolyticum W16, International Journal of Hydrogen Energy, 34(2009), 7182-7188
292. Pattra, S., Sangyoka, S., Boonmee, M., Reungsang, A., Bio-hydrogen production from the fermentation of sugarcane bagasse hydrolysate by Clostridium butyricum, International Journal of Hydrogen Energy, 33(2008), 5256-5265
293. Ivanova, G., Rákhely, G., Kovács, K.L., Thermophilic biohydrogen production from energy plants by Caldicellulosiruptor saccharolyticus and comparison with related studies, International Journal of Hydrogen Energy, 34(2009), 3659-3670
49
294. Xu, J-F., Ren, N-g., Su, D-X., Qiu, J., Bio-hydrogen production from acetic acid steam-exploded corn straws by simultaneous saccharification and fermentation with Ethanoligenens harbinense B49, International Journal of Hydrogen Energy, 34(2010), 381-386
295. Goksungur, Y., Optimization of the production of chitosan from beet molasses by response surface methodology, J Chem Technol Biotechnol, 79(2004), 974–981
296. Rabea, E.I., Badawy, M.E., Stevens, C.V., Smagghe, G. and Steurbaut, W., Chitosan as antimicrobial agent: applications and mode of action, Biomacromolecules, 4(2003), 1457–1465
297. Synowiecki, J. and Al-Khateeb, N.A., Production, properties, and some new applications of chitin and its derivatives, Crit Rev Food Sci Nutr, 43(2003), 145–171
298. Silva, H.S.R.C., dos Santos, K.S.C.R. and Ferreira, E.I., Chitosan: hydrossoluble derivatives, pharmaceutical applications and recent advances, Quim Nova, 29(2006), 776–785
299. Tai, C., Li, S., Xu, Q., Ying, H., Huang, H., Ouyang, P., Chitosan production from hemicellulose hydrolysate of corn straw: impact of degradation products on Rhizopus oryzae growth and chitosan fermentation, Letters in Applied Microbiology, 51(2010), 278–284
300. Crittenden, R., Playne, M., Production, properties, and applications of food-grade oligosaccharides, Trends in Food Science and Technology, 7(1996), 353–361
301. Yuan, X., Wang, J., Yao, H., Feruloyl oligosaccharides stimulate the growth of Bifidobacterium bifidum, Anaerobe, 11(2005), 225–229
302. Cummings, J., Edmond, L., Magee, E., Dietary carbohydrates on health: do we still need the fibre concept?, Clinical Nutrition Supplements, 1(2004), 5–17.
303. Gibson, G., Fibre and effects on probiotics (the prebiotic concept), Clinical Nutrition Supplements, 1(2004), 25–31
304. Wollowski, I., Rechkemmer, G., Pool-Zobel, B., Protective role of probiotics and prebiotics in colon cancer, American Journal of Clinical Nutrition, 73(2001), 451–455
305. Hsu, C. K., Liao, J. W., Chung, Y. C., Hsieh, C. P., Chan, Y. C., Xylooligosaccharides and fructooligosaccharides affect the intestinal microbiota and precancerous colonic lesion development in rats, Journal of Nutrition, 134(2004), 1523–1528
306. Nabarlatz, D., Ebringerová, A., Montané, D., Autohydrolysis of agricultural by-products for the production of xylo-oligosaccharides, Carbohydrate polymers, 69(2007), 20-28
307. Ando, H., Ohba, H., Sakaki, T., Takamine, K., Kamino, Y., Moriwaki, S., Hot-compressed-water decomposed products from bamboo manifest a selective cytotoxicity against acute lymphoblastic leukemia cells, Toxicology in Vitro, 18(2004), 765–771
308. Rivas, B., Domínguez, J., Domínguez, H., Parajó, J. Bioconversion of posthydrolysed autohydrolysis liquors: an alternative for xylitol production from corncobs, Enzyme and Microbial Technology, 31(2002), 431–438
309. Bengtsson, O., Jeppsson, M., Sonderegger, M., Parachin, N.S., Sauer, U., Hahn- Hagerdal, B., Gorwa-Grauslund, M.F., Identification of common traits in improved
50
xylose-growing Saccharomyces cerevisiae for inverse metabolic engineering, Yeast, 25 (2008), 835–847
310. Grotkjaer, T., Christakopoulos, P., Nielsen, J., Olsson, L., Comparative metabolic network analysis of two xylose fermenting recombinant Saccharomyces cerevisiae strains, Metab. Eng, 7 (2005), 437–444
311. Jeffries, T.W., Van Vleet, J.R., Pichia stipitis genomics, transcriptomics, and gene clusters, FEMS Yeast Res, 9 (2009), 793–807
312. Karhumaa, K., Pahlman, A.K., Hahn-Hagerdal, B., Levander, F., Gorwa-Grauslund, M.F., Proteome analysis of the xylose-fermenting mutant yeast strain TMB 3400, Yeast, 26 (2009), 371–382
313. Van Vleet, J.H., Jeffries, T.W., Yeast metabolic engineering for hemicellulosic ethanol production, Curr. Opin. Biotechnol, 20 (2009), 300–306
314. Nyyssola, A., Pihlajaniemi, A., Palva, A., Weymarn, N., Leisola, M., Production of xylitol from d-xylose by recombinant Lactococcus lactis, J Biotechnol 118 (2005), 55–66
315. Ko, B.S., Rhee, C.H. Kim, J.H., Enhancement of xylitol productivity and yield using a xylitol dehydrogenase gene-disrupted mutant of Candida tropicalis under fully aerobic conditions, Biotechnol Lett 28 (2006), 1159–1162
316. Hibi, M., Yukitomo, H., Ito, M., Mori, H., Improvement of NADPH-dependent bioconversion by transcriptome-based molecular breeding, Appl Environ Microbiol 73 (2007), 7657–7663
317. Oh, Y.J., Lee, T.H., Lee, S.H., Oh, E.J., Ryu, Y.W., Kim, M.D., Seo, J.H., Dual modulation of glucose 6-phosphate metabolism to increase NADPH-dependent xylitol production in recombinant Saccharomyces cerevisiae, J Mol Catal B: Enzym 47 (2007), 37–42
318. Shi, N.Q., Davis, B., Sherman, F., Cruz, J., Jeffries, T.W., Disruption of the cytochrome c gene in xylose-utilizing yeast Pichia stipitis leads to higher ethanol production, Yeast, 15 (1999), 1021–1030
319. Mohagheghi, A., Evans, K., Chou, Y.C., Zhang, M., Cofermentation of glucose, xylose, and arabinose by genomic DNA-integrated xylose/arabinose fermenting strain of Zymomonas mobilis AX101, Appl. Biochem. Biotechnol, 98–100(2002), 885–898
320. Bengtsson, O., Hahn-Hagerdal, B., Gorwa-Grauslund, M.F., Xylose reductase from Pichia stipitis with altered coenzyme preference improves ethanolic xylose fermentation by recombinant Saccharomyces cerevisiae, Biotechnol. Biofuels, 2(2009), 9
321. Chen, K., Iverson, A.G., Garza, E.A., Grayburn, W.S., Zhou, S., Metabolic evolution of non-transgenic Escherichia coli SZ420 for enhanced homoethanol fermentation from xylose, Biotechnol. Lett, 32 (2010), 87–96
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Table 1 Composition of representative hemicellulosic feedstocks
Substrate Xylose Arabinose Galactose Mannose Rhamanose Uronic acid Reference
Agricultural and agro-industrial biomass Corn cob 28-30 3-5 1-1.2 - 1 3 15,16,17,18 Corn stover 15-25 2-3.6 0.8-2.2 0.3-0.4 - - 15,19,20 Corn stalks 26 4.1 <2.5 <3.0 - - 21 Corn fibre 21.6 11.4 4.4 - - - 22 Rice husk 17.7 1.9 - - - - 23 Rice straw 15-23 3-5 0.4 1.8 - - 22,20 Sugarcane bagasse 21-26 2.5-7 1.6 0.5-0.6 - - 24-26
Wheat straw 19-21 2.4-4 2-2.5 0-0.8 - - 27,20 Wheat bran 16 9 1 0 - - 16 Barley straw 15 4 - - - - 28 Hardwoods Birch 19-25 0.3-0.5 0.7-1.3 1.8-3.2 0.6 4-7 29,30 Maple 18-19 0.8-1 1.0 1.3-3.3 - 4.9 15,30 Eucalypt 14-19 0.6-1 1-1.9 1-2 0.3-1 2 16, 31-33 Oak 22 1 1.9 2.3 - 3 34 Aspen 18-28 0.7-4 0.6-2 0.9-2.5 0.5 5-6 27,29,30,35 Poplar 18-21 0.9-1.4 1 3.5 - 2.3-3.7 30, 36 Softwoods Spruce 5-10 1.2 1.9-4.3 9.4-15 0.3 1.5-6 29,30,37-39 Douglas fir 6 3 3.7 - - - 35 Pine 5-10 2-4 2-4 6-13 - 3-6 30
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Table 2 List of microorganisms with high specific activity (μmol.min-1.mg-1) for hemicellulases (119) ENZYME Organism Substrate Specific
activity Feruloyl esterase Clostridium
stercorarium Ethyl ferulate 88
β-1,4-xylosidase
Thermoanaerobacter ethanolicus
o-nitrophenyl-β-D-xylopyranoside
1073
Exo-β-1,4-mannosidase
Pyrococcus furiosus
p-nitrophenyl-β-D-galactoside
31.1
Endo-β-1,4-mannanase
Bacillus subtilis
Galactoglucomannan/ glucomannans/mannan
514
α-Larabinofuranosidase
Clostridium stercoarium
alkyl-α-arabinofuranoside/ aryl-α-arabinofuranoside/ Larabinogalactan/ L-arabinoxylan/ methylumbelliferyl-α-Larabinofuranoside
883
α-Glucuronidase
Thermoanaerobacterium saccharolyticum
4-O-methyl-glucuronosyl-xylotriose
9.6
α-Galactosidase Escherichia coli raffinose 27350 Endo-galactanase Bacillus subtilis arabinogalactan 1790 β-Glucosidase Bacillus polymyxa 4-nitrophenyl-β-D-
glucopyranoside 2417
Acetyl xylan esterase Fibrobacter succinogenes
Acetylxylan/ α-naphthyl acetate
2933
Feruloyl esterase
Aspergillus niger
Methyl sinapinate
156
Endo-1,4-β-xylanase Trichoderma longibrachiatum
1,4-β-D-xylan
6630
β-1,4-xylosidase
Aspergillus nidulans
p-nitrophenyl-β-D-xylopyranoside
107.1
Exo-β-1,4-mannosidase
Aspergillus niger
β-D-Man-(1-4)-β-D-GlcNAc-(1-4)-β-DGlcNAc- Asn-Lys
188
Endo-β-1,4-mannanase
Sclerotium rolfsii
Galactoglucomannan/mannans galactomannans/glucomannans/
380
Endo-α-1,5-arabinanase Aspergillus niger 1,5-α-L-arabinan 90.2 α-L-arabinofuranosidase Aspergillus niger 1,5-α-L- 396.6
53
arabinofuranohexaose/ 1,5-α- L-arabinotriose/ 1,5-L-arabinan/ α-Larabinofuranotriose
α-Glucuronidase
Phanerochaete chrysosporium
4-O-methyl-glucuronosyl-xylobiose
4.5
α-Galactosidase Mortierella vinacea melibiose 2000 Endo-galactanase Aspergillus niger 6593
β-glucosidase
Humicola insolvens
(2-hydroxymethylphenyl)-β-Dglucopyranoside
266.9
Acetyl xylan esterase
Schizophyllum commune
4-methylumbelliferyl acetate/ 4- nitrophenyl acetate
227
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Table 3 Ethanol production from hemicellulosic hydrolysates
Substrate Pretreatment Detoxification Microorganisms Ethanol (g/L) Ethanol yield (g/g) Reference
Bagasse Steam pretreatment - Pichia Stipitis CBS
6054 19.5 0.22 141
Sunflower seed
hull Acid hydrolysis Over-liming combined
with Na2SO3 Pichia Stipitis NRRLY-
7124 11 0.32 142
Bagasse Acid Hydrolysis Over-liming or electrodialysis
Pachysolen tannophilus DW 06 21 0.35 143
Prosopsis Julifora Acid hydrolysis Over-liming Pichia stipitis NCIM
3498 7.13 0.39 144
Water hyacinth
Acid hydrolysis - Pichia stipitis - 0.425 145
Olive Tree Acid hydrolysis Over-liming and Activated charcoal Pichia stipitis - 0.42 146
Yellow poplar wood meal
Alkaline pretreatment
Acid hydrolysis Over-liming Pichia stipitis 28.7 0.48 147
Oat spelt xylan Enzymatic hydrolysis - Debaromyces hansenii 7.55 0.42
139 Wheat bran Enzymatic
hydrolysis - Debaromyces hansenii 6.42 0.41
Tamarind kernel powder Acid hydrolysis Over-liming and
Activated charcoal Debaromyces hansenii 16 0.35 148
Tamarind kernel powder
Enzymatic hydrolysis - Debaromyces hansenii 5.18 0.39 148
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Table 4 Xylitol production from detoxified hemicellulosic hydrolysates
Substrate Pretreatment Detoxification Microorganisms Xylitol (g/L) Xylitol yield (g/g) Reference
Sugarcane bagasse Acid hydrolysis Activated charcoal Candida guilliermondii 35.5 0.72 164
Eucalyptus Acid hydrolysis Activated charcoal Ion exchange resins Candida guilliermondii 32.7 0.57 169
Corn fibre Acid hydrolysis Over-liming
Activated charcoal Ion exchange resins
Candida tropicalis 129 0.43 170
Brewery spent-grain Acid hydrolysis Over-liming Debaromyces hansenii 18 0.38 171
Rice straw Acid hydrolysis Over-liming Activated charcoal
Candida subtropicalis WF79 - 0.73 172
Wheat straw Acid hydrolysis Activated charcoal Candida guilliermondii FTI 20037 24.2 0.48 173
Corn cob Acid hydrolysis Over-liming, solvent
extraction Candida tropicalis
W103 68.4 0.7 174
56
Table 5 Production of xylitol and ethanol using recombinant strains Sugars Genetic manipulations Microorganism Yield and/or
productivity Reference
Xylitol Xylose + glucose Expressed P.stipitis XR L.lactis 2.7 g/l/h 314
Xylose + glycerol Disrupted XYL2 C.tropicalis 3.2 g/l/h 315
Xylose + glucose Expressed XylE and K.lactis XR, deleted xylA,yhbC-deficient
E.coli 0.81 g/l/h 316
Xylose + glucose Overexpreesed G6PDH, expreesed Xyl1, attenuated PGI activity
S.cerevisiase 0.34g (g cdw h)-1 317
Ethanol
Xylose Disrupted CRC 1 P.stipitis 0.46 g/g 318
Xylose + glucose Genomic DNA integration
Z.mobilis 0.4 g/g 319
Xylose Overexpreesed native P.stipitis XDH, xylulokinase, deleted GRE3
S.cerevisiae 0.39 g/g 320
Xylose Evolutionary engineering, increased PDH activity
E.coli 0.46 g/g 321
XylE- xylose permease; XR- xylose reductase; G6PDH- glucose-6-phoshate dehydrogenase; PGI- phosphoglucose isomerase; PDH- pyruvate dehydrogenase; XDH- xylitol dehydrogenase; CRC 1- cytochrome c
57
Mechanism of action of the hemicellulases
Figure 1 : Xylanolytic enzymes involved in the degradation of hard wood and soft wood xylan. Ac, acetyl group; Ara, α- arabinofuranose; MeGlcA, α-4-Omehtylglucuronic acid; Xyl xylose.
58
Figure 2 Microbial conversion of xylose to ethanol and xylitol
Acetyl
NADH NAD
Alcohol dehydrogenase
Pyruvate decarboxylase
Pyruvate formate lyase
Pyruvate Acetaldehyde ETHANOL
CO2
Aldehyde dehydrogenase
ATP
ADP
NADH
ATP
ADP
NAD
(TKL)
Transaldolase (TAL)
Transketolase (TKL)
D-Ribose - 5-
Glyceraldihyde-3- Phosphate
D-Fructose-6- Phosphate
NAD
NADH
D-Xylulose - 5- Phosphate
Xylose isomerase (XI)
Xylulose kinase (XK)
ADP
ATP
NAD(P) NAD(P)
Xylitol dehydrogenas
Xylose reductase (XR)
D - XYLOSE
XYLITOL
D-Xylulose
Hemicellulose Hemicellulase