Download Plant cell walls to ethanol

Biochem. J. (2012) 442, 241–252 (Printed in Great Britain)
241
doi:10.1042/BJ20111922
REVIEW ARTICLE
Plant cell walls to ethanol
Douglas B. JORDAN*1 , Michael J. BOWMAN*, Jay D. BRAKER*, Bruce S. DIEN*, Ronald E. HECTOR*, Charles C. LEE†,
Jeffrey A. MERTENS* and Kurt WAGSCHAL†
*USDA Agricultural Research Service, National Center for Agricultural Utilization Research, Peoria, IL 61604, U.S.A., and †USDA Agricultural Research Service, Western Regional
Research Center, Albany, CA 94710, U.S.A.
Conversion of plant cell walls to ethanol constitutes second
generation bioethanol production. The process consists of several
steps: biomass selection/genetic modification, physiochemical
pretreatment, enzymatic saccharification, fermentation and
separation. Ultimately, it is desirable to combine as many of the
biochemical steps as possible in a single organism to achieve
CBP (consolidated bioprocessing). A commercially ready CBP
organism is currently unreported. Production of second generation
bioethanol is hindered by economics, particularly in the cost of
pretreatment (including waste management and solvent recovery),
the cost of saccharification enzymes (particularly exocellulases
and endocellulases displaying kcat ∼ 1 s − 1 on crystalline
cellulose), and the inefficiency of co-fermentation of 5- and
6-carbon monosaccharides (owing in part to redox cofactor
imbalances in Saccharomyces cerevisiae).
INTRODUCTION
glycoside hydrolases or esterases that mediate their hydrolysis.
Barring pretreatment, the yields of monosaccharides from native
biomass hydrolysed with cellulases, xylanases and associated
enzymes are only on the order of 20 %. An efficient pretreatment
will increase the yield to over 80 %. (ii) Next there is enzymatic
saccharification. This step requires several enzymes to release
the major fermentable monosaccharides. The requisite enzymes
should display superior hydrolysis kinetics on natural substrates
and good stability properties. (iii) Finally there is fermentation.
This step needs to be robust, and capable of rapid and efficient
fermentation of all the sugars available (i.e. both 5- and 6-carbon
sugars). The second and third steps may be combined or run
separately to achieve the following processes: SHF (separate
hydrolysis and fermentation), where cellulosic and hemicellulosic
components of cell walls are separately hydrolysed prior to
presenting them to the fermenting organism; SSF (simultaneous
saccharification and fermentation), where enzymatic cellulose
hydrolysis and hexose fermentation occur concurrently in the
same vessel; SSCF (simultaneous saccharification and cofermentation), where cellulose and hemicellulose hydrolysis, and
hexose and pentose fermentation occur concomitantly in the
same vessel; and CBP (consolidated bioprocessing), where a
single organism carries out saccharification and fermentation.
Considerable effort has been applied to engineering an ideal CBP
organism (reviewed in [6]), but currently there are no reports of a
commercially viable CBP organism suitable for the operation.
Economical conversion of the polysaccharides that constitute
cell walls of plants to second generation bioethanol may seem
a straightforward exercise in chemical conversion. However, over
the last several years the processes involved in the conversion
of cellulose and hemicellulose, which compose the bulk of the
biomass feedstock, have been closely associated with the word
‘recalcitrance’ [1], to emphasize obstacles that can impede the
conversion. Recalcitrance of the conversion stems from the key
word ‘economical’, as each step in the conversion can be costly
and drive production costs to exceed those of its transportation fuel
competitors, such as those derived from fossil fuels (e.g. gasoline,
diesel and natural gas) or those derived from starch, sucrose and
vegetable oils (e.g. first generation bioethanol and biodiesel). The
combined strength of the glycosidic bonds of cellulose [2] and
its associated crystal structure dictate application of either harsh
physiochemical conditions or use of several specific enzymes.
In comparison with production of first generation bioethanol
(derived from corn starch and cane sugar), the harsh physiochemical conditions add considerable expense to construction of
saccharification reactors and downstream processing, and the use
of enzymes adds the expense of preparation of numerous enzymes,
particularly the cellulases. In recent years, protein production
efficiency of cellulases has been increased more than 10-fold, and
this alone makes enzymatic saccharification more economical
than physiochemical methods [3–5].
Conversion of cell walls into bioethanol can be viewed as
occurring in three operational steps. (i) First there is biomass
production and pretreatment. The feedstock of biomass will
depend on local availability to determine whether excess crop
stover (e.g. corn, wheat and sorghum) or a dedicated energy
crop (e.g. switchgrass and Miscanthus) is harvested. Pretreatment
consists of mechanical, heat and chemical conditions that aim at
maximizing exposure of the chemical bonds of biomass to the
Key words: cellulose, co-fermentation, economics, hemicellulose,
lignin, physiochemical pretreatment.
PLANT CELL WALL STRUCTURE
Plant tissue varies widely in structure and composition and,
as might be expected, in its response to pretreatment
and enzymatic saccharification [7]. Plants comprise primary and
secondary cell walls, both of which are fortified by cellulose
microfibrils. Primary cell walls typically contain cellulose,
Abbreviations used: CBP, consolidated bioprocessing; CE, carbohydrate esterase; DP, degree of polymerization; GH, glycoside hydrolase; 4NPA,
4-nitrophenyl-α-L-arabinofuranose; 4NPX, 4-nitrophenyl-β-D-xylopyranoside; PK, phosphoketolase; PPP, pentose phosphate pathway; SHF, separate
hydrolysis and fermentation; SSCF, simultaneous saccharification and co-fermentation; SSF, simultaneous saccharification and fermentation.
1
To whom correspondence should be addressed (email [email protected]).
c The Authors Journal compilation c 2012 Biochemical Society
242
Figure 1
D. B. Jordan and others
Polymeric structures of cellulose and hemicellulose chains
Cellulose consists of repeating β-(1,4)-linked D-glucose residues. Adjacent D-glucose residues are flipped, making cellobiose the fundamental repeating unit. The major hemicelluloses are shown.
Xylan consists of repeating β-(1,4)-linked D-xylose residues with the potential for L-arabinose or acetyl substitutions at either the 2-O or 3-O positions or both; 2-O substitution with 4-hydroxy- or
4-methyl-glucuronic acid; and added complexity from substitutions (hexose, pentose and/or phenolics) on the L-arabinose side-chain residues [18,189–192]. Mannan consists of a β-(1,4)-linked
D-mannose backbone, whereas the glucomannan backbone has both D-mannose and D-glucose residues with β-(1,4) linkages. Xyloglucan has a β-(1,4)-linked D-glucose backbone with D-xylose
side chains, and the mixed-linkage β-glucan backbone has both β-(1,4)- and β-(1,3)-linked glucose residues.
hemicellulose (xyloglucans), pectin and proteins. In grasses,
glucuronoarabinoxylan, which is cross-linked by diferulate,
substitutes for the pectin [8]. Cellulose microfibres are crisscrossed within the cell wall with closer alignment and spacing
in primary cell walls than in secondary cell walls. Secondary cell
walls are composed of cellulose, hemicellulose and lignin and
constitute the majority of cell wall mass; for example, 70–80 %
weight of corn stover is present in secondary cell walls [9].
Cellulose
Cellulose, the most abundant biopolymer, is formed by β-(1,4)linked D-glucoses, where adjacent D-glucoses are flipped making
cellobiose the fundamental repeating unit (Figure 1) [10]. The
linear flat polymer allows for extensive hydrogen bonding within
and between chains, as well as van der Waals stacking interactions
between chains [11]. Glucan chains occur in hexagonal arrays of
36 (3 nm×5 nm width) with exceptionally high DPs (degrees
of polymerization) [12]. For corn stover, the mean DP is 6000
in primary walls and 14 000 in secondary walls [13]. Formation
of the arrays is controlled by transmembrane assembly complexes,
termed rosettes, which ensure that individual glucans are aligned
into crystallized fibrils to maximize tensile strength and to fit
tightly enough to exclude water. The lattice structure is polymorphic, where native cellulose is defined as crystal form I, but
following chemical treatment can be transformed to crystal forms
II, III, V or X [14]. Crystallinity is thought to influence biomass
recalcitrance and varies widely among plant cell walls from
approximately 40–50 % in plant cellulose to 65–80 % in bacterial
and algal cellulose (reviewed in [15]). Crystallinity measurements
can vary depending upon the analytical technique used for
measurement and, in the case of pretreated plant cell walls, upon
the removal of low-crystalline components (e.g. xylan) [14].
c The Authors Journal compilation c 2012 Biochemical Society
Hemicellulose
Hemicelluloses constitute 20–30 % of the biomass of
dicotyledonous plants, such as trees, up to 50 % for some tissues
of monocotyledonous plants, and approximately 25 % of the
available biomass of the bioenergy-specific crops Miscanthus,
switchgrass, fescue and fibre sorghum [16–19]. Hemicellulose
bonds to the surface of the cellulose microfibrils and forms a
matrix between fibres, where it plays the dual role of keeping
fibres from aggregating and adding flexion to the cell wall
[20]. Hemicellulose polysaccharides are shorter than those of
cellulose (DP of 200 or less) and they are often branched, with
short chains containing other sugars, acetyl groups and phenolic
groups (Figure 1). Structural heterogeneity is a hallmark of
hemicelluloses, arisen due to the physical benefits of resistance to
environmental degradation.
A procedural definition of hemicellulose is the polysaccharides
that are extractable from plant cell walls by alkaline solution
(e.g. 4–24 % potassium hydroxide). Hemicellulose composition
varies with plant species and tissue type and it can be classified
into four major groups [21] (Figure 1) based on polysaccharide
composition [18,22,23]. (i) Mannans comprise galactomannan,
glucomannan and galactoglucomannan. Galactomannans have
β-1,4-mannose backbones with α-1,6-galactose branches;
glucomannans contain both mannose and D-glucose β-1,4-linked
backbones; and galactoglucomannans have β-1,4-mannose and
β-1,4-glucose backbones with α-1,6-galactose branches attached
to the mannose backbone. For example, the secondary cell walls
of conifers (i.e. softwoods) consist of galactoglucomannan [10–
30 % (w/w)]. (ii) Mixed-linkage glucans comprise a backbone
of D-glucose residues having both β-1,3 and β-1,4 linkages. For
example, the primary cell walls of grasses contain 2–15 % (w/w)
of mixed-linkage glucans. (iii) Xylans have β-1,4-linked D-xylose
Cell walls to ethanol
backbones that may include arabinan and glucuronic acid side
chains. For example, in grasses, the primary cell walls contain
20–40 % (w/w) glucuronoarabinoxylan, whereas the secondary
cell walls contain 40–50 % (w/w) glucuronoarabinoxylan. In
dicots (e.g. hardwoods), the secondary cell walls contain 20–
30 % glucuronoxylan. (iv) Xyloglucan has a β-1,4-glucan
backbone with xylose-containing branches that can contain other
monosaccharide substitutions, such as galactose, arabinose and
fucose. For example, the primary cell walls of conifers (i.e.
softwoods) contain 10 % (w/w) xyloglucans and the primary cell
walls of dicots (e.g. hardwoods) contain 20–25 % (w/w).
Xylans are the most abundant class of hemicelluloses, with
glucuronoarabinoxylan being the main target for enzymatic
saccharification for renewable bio-feedstock production.
Glucuronoarabinoxylan (e.g. from corn stover) is composed
of a β-(1,4)-linked D-xylose polymer backbone (xylan) with
L-arabinose and glucuronic acid side chains [24]. Extensive
acetylation may occur and the L-arabinose side chains
can be esterified with ferulic acid that in turn cross-links
glucuronoarabinoxylan, hemicelluloses and lignin (Figure 1).
This structural heterogeneity indicates six requisite enzyme
activities for complete glucuronoarabinoxylan saccharification: endoxylanase, β-xylosidase, α-arabinofuranosidase, αglucuronidase, acetylxylan esterase and ferulic acid esterase.
Lignin
Lignin principally consists of p-hydroxyphenyl-, guaiacyl- and
syringyl-phenylpropanoid units that are polymerized by radical
chemistry coupling reactions to form covalent ether and alkyl
linkages [9]. Grasses and hardwoods contain largely guaiacyland syringyl-phenylpropanoid units, with only trace amounts
of p-hydroxyphenyl-phenylpropanoid units; softwoods largely
contain syringyl-phenylpropanoid units. Most pretreatments do
not extensively degrade lignin because of the nature of its linkages.
Studies on lignin-reduced mutants of alfalfa and sorghum plants
have demonstrated that lower lignin content is positively correlated with enzymatic digestibility following pretreatment [25,26].
Pectin
Pectin is a complex heteropolysaccharide that hydrates and
further cements the primary cell wall matrix. It accounts
for 30–40 % of non-cellulosic polysaccharides in the primary
cell walls of herbaceous dicotyledons and non-graminaceous
monocots with significantly lesser amounts found in grasses,
woody tissue and secondary cell walls [27,28]. Pectin
consists of long homogalacturonan chains of α-(1-4)-linked
D-galacturonic acid and is often esterified with methyl or
acetyl groups. Homogalacturonan is interspersed with the
branched polysaccharides rhamnogalacturonan I (primarily),
rhamnogalacturonan II and xylogalacturonan [29]. It is desirable
to hydrolyse pectin because it blocks cellulases, xylanases and
xylan-debranching enzymes from reaching their substrates. Also,
pectin is an important aspect in the conversion of citrus waste
and sugar beet pulp into ethanol, where the polysaccharides are
abundant [30,31].
PRETREATMENT PROCESSES
Pretreatment can include physical, chemical and thermal
processes, where most pretreatment conditions combine all three
effects. For example, biomass can be treated in a steam explosion
reactor with dilute sulfuric acid as a catalyst. The biomass is
mixed with dilute sulfuric acid, heated with direct steam and
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quenched by rapid depressurization; the final step increases
surface area and reduces particle size. Meanwhile the dilute
acid hydrolyses the hemicellulose and reduces the DP of the
lignin. Microscopic studies of the affect of pretreatment on cell
wall properties emphasized that only subtle changes are needed
to expose microfibrils sufficiently for cellulase action and that
collapsing cell wall structure either leads to the same or worse Dglucose yields [32]. Since pretreatment can have negative consequences, owing to the chemicals released from the native
plant material or as side products of the pretreatment, that
impede enzyme hydrolysis [33–39] and microbial fermentation
[40–42], less harsh treatments are called for. For example,
switchgrass pretreated with dilute ammonium [8 % (w/v), 180 ◦ C
for 20 min] was hydrolysed to D-glucose at 80 % efficiency using
commercial cellulases (B.S. Dien, unpublished work). Many
additional pretreatments have been applied, including ammonia
fibre explosion [43,44], alkaline peroxide [45–47], liquid hot
water [44,48], sulfuric acid/sulfite solutions [49,50] and cellulose
solvents (i.e. concentrated phosphoric acid and room temperature
liquid ionic solutions) [51–54]. Recent pretreatment studies
[54a,54b] have reinforced previous work [55,56] which indicates
that the major barrier to cellulase activity on pretreated biomass is
mass transfer limitations of cellulases binding to the microfibrils,
a feature termed porosity. The significance of modifications
to the fibril structure, while observed to benefit conversion of
pure samples of cellulose, remains uncertain for more complex
lignocellulosic samples, with the exception of pretreatments that
dissolve cellulose.
THE ENZYMES
Enzymes that catalyse deconstruction of cellulose
Deconstruction of crystalline cellulose can be achieved by
the activity of three enzymes: cellobiohydrolase (exocellulase,
E.C. 3.2.1.91 and E.C. 3.2.1. − ), endoglucanase (endocellulase, E.C. 3.2.1.4) and β-glucosidase (E.C. 3.2.1.21). The
first two enzymes act on cellulose. In comparison with many
other glycoside hydrolases acting on their natural substrates, the
cellulases are known for their low catalytic activity, particularly
on crystalline cellulose [3–5]. Several of the endoglucanases and
cellobiohydrolases from Trichoderma reesei display kcat values of
2–20 s − 1 on amorphous cellulose at 25 ◦ C [57], corresponding to
6-fold lower rates on crystalline cellulose [58]. A similar rate, 3.5
s − 1 , has been calculated for T. reesei CEL7A cellobiohydrolase
acting on crystalline cellulose [59–61]. Endoglucanases display
rates of similar magnitude on crystalline cellulose: 0.22 s − 1 at
45 ◦ C [62], 0.5 s − 1 at 80 ◦ C [63] and 1.4 s − 1 at ∼ 90 ◦ C [64]. In
recent years, the protein production efficiency has been improved
by at least 10-fold, which has been variously reported to decrease
the cost of the cellulases to approximately $0.10 [3], $0.20 [4] or
$0.50 [5] per gallon of ethanol produced.
Cellobiohydrolases catalyse the release of cellobiose from
either the non-reducing end or the reducing end of cellulose,
depending on the cellobiohydrolase (Figure 2A). They do
so through a double-displacement mechanism, where the
stereochemistry at the anomeric centre is retained, or through
a single displacement mechanism, where the stereochemistry
at the anomeric centre is inverted [60]. Cellobiohydrolases
belong to GH (glycoside hydrolase) families 5, 6, 7, 9 and
48, according to the CAZy database (http://www.cazy.org),
which groups carbohydrate-active enzymes according to their
amino acid sequence homology [65]. The database lists X-ray
structures (PDB codes) and GenBank® accession information for
the enzymes discussed in the present review. Endoglucanases
c The Authors Journal compilation c 2012 Biochemical Society
244
Figure 2
D. B. Jordan and others
Molecules discussed in the text
(A) Hypothetical active site of a cellobiohydrolase that cleaves cellobiose from the non-reducing end. The scissile site is the glycosidic bond held between subsite − 1 and subsite + 1. Numbering
proceeds to the left of the scissile bond towards the non-reducing end of the oligoglucoside for negatively numbered subsites, and to the right towards the reducing end of the oligoglucoside for
positively numbered subsites. (B) Substrates of Selenomonas ruminantium β-xylosidase/α-arabinofuranosidase. (C) Overlay of D-xylopyranose and L-arabinofuranose showing that the two-ring
systems can occupy similar space.
catalyse the endohydrolysis of (1,4)-β-D-glucosidic linkages in
cellulose. The enzymes belong to families GH 5–10, 12, 16,
18, 19, 26, 44, 45, 48, 51, 74 and 124. GH61 was initially
described as an endoglucanase, but more recently has been
shown to lack GH activity [66]. Instead, members of the
family are copper mono-oxygenases that catalyse cleavage of
cellulose oxidatively, releasing cellodextrins [67–69]. The monooxygenases can accept reducing equivalents from ascorbate
or reduced dyes [67,69]. In situ, cellobiose dehydrogenase is
probably the electron donor [68,69]. In the presence of cellulose,
cellobiohydrolase and endoglucanase, the mono-oxygenase
and cellobiose dehydrogenase act synergistically to enhance the
cellulase activity by approximately 2-fold [68]. In the presence
of cellulose and β-glucoside, the mono-oxygenase enhances
the β-glucosidase manyfold [68]. β-Glucosidases act on soluble
cello-oligosaccharides, including cellobiose. The enzymes
catalyse the hydrolysis of terminal non-reducing β-D-glucosyl
residues through a double displacement mechanism, with release
of β-D-glucose. The enzymes belong to families GH 1, 3, 5, 9, 30
and 116. A GH3 β-glucosidase from Aspergillus oryzae expresses
typical catalytic parameters: kcat of 1000 s − 1 and K m of 2.0 mM
(pH 5.0, 50 ◦ C) [70]. Crude industrial mixtures of enzymes that
act on cellulose and soluble cello-oligosaccharides have been
reviewed recently [5].
Enzymes that catalyse deconstruction of hemicellulose
Below the focus is on the enzymes that operate on
glucuronoarabinoxylan. Descriptions of the enzymes that act
on the other forms of hemicellulose, as well as other plant
polysaccharides, can be found elsewhere [71]. Heterogeneity
of glucuronoarabinoxylan requires six distinct enzyme activities
for complete saccharification: endoxylanase (EC 3.2.1.8), βxylosidase (EC 3.2.1.37), α-arabinofuranosidase (EC 3.2.1.55),
α-glucuronidase (EC 3.2.1.131), acetylxylan esterase (EC
3.1.1.72) and ferulic acid esterase (EC 3.1.1.73). Economic
c The Authors Journal compilation c 2012 Biochemical Society
success of a biorefinery requires efficient utilization of the
hemicellulose carbohydrates that can be released by enzymatic
hydrolysis. In addition, xylan removal leads to greater enzymatic hydrolysis of cellulose, resulting in increased D-glucose
yield. The mechanisms by which xylan removal leads to
increased cellulose breakdown include increased accessibility to
the cellulose fibrils [72,73] and removal of xylo-oligosaccharides
[39], which inhibit cellulase enzyme activity. Similarly synergism
is a common theme for the enzymes that act in the deconstruction
of xylans, including glucuronoarabinoxylan. Inclusion of the
activities of the following individual enzymes enhances the rate of
the endoxylanase acting on glucuronoarabinoxylan: β-xylosidase
[74], α-glucuronidase [75], acetylxylan esterase [76] and ferulic
acid esterase [77,78].
Many industrially relevant endoxylanase genes have been
cloned from a wide array of bacteria and fungi [79]. The
majority of these enzymes are classified into the GH10 and GH11
families. Both categories of enzymes use a double-displacement
mechanism that retains the anomeric configuration. The GH10
endoxylanases have lower substrate specificity and, therefore, a
higher capacity to hydrolyse substituted xylan polymers. The
recombinant endoxylanase with the highest reported specific
activity is Xyn10B from Cellvibrio mixtus [80]. Acting on oat
spelt xylan, Xyn10B has a K m of 6.17 mg·ml − 1 and a kcat of 330 s − 1
at 37 ◦ C. There are also many endoxylanases that function
optimally at extreme conditions: pH 2 [81–83], pH 9–10 [84–86],
85–100 ◦ C [87–89] and 4 ◦ C [90,91].
Xylan 1,4-β-D-xylosidase (EC 3.2.1.37) catalyses the
hydrolysis of single D-xylose units from the non-reducing end of
xylo-oligosaccharides. It is classified in the CAZy database under
GH families 1, 3, 30, 39, 43, 51, 52, 54, 116 and 120 [65]. As well
as being required for the complete saccharification of xylan [92],
β-xylosidase has been demonstrated to act synergistically with
other hemicellulases in the degradation of xylan [74]. At present,
the preponderance of characterized β-xylosidases belong to GH
family 43. GH43 is targeted in part because this family catalyses
Cell walls to ethanol
Figure 3 Comparison of k cat values obtained for GH43 β-xylosidases acting
on X2 and 4NP
The β-xylosidases with literature values are from Selenomonas ruminantium (1), Bacillus
pumilus 12 (2), Bacillus pumilus IPO (3) and Bacillus halodurans C-125 (4) [92,97]. The
remaining β-xylosidases without literature values are from Bacillus subtilis subsp. subtilis str.
168 (5), Alkaliphilus metalliredigens QYMF (6), Bacillus sp. (7) and Bacillus sp. (8) (D.B.
Jordan and J.D. Braker, unpublished work).
hydrolysis using an inverting single-displacement mechanism.
This precludes transxylosylation [93], which could otherwise
impinge on efficiency, as in the case of high concentrations of
a competing nucleophile, such as xylobiose.
GH43 β-xylosidase SXA, isolated from the organism
Selenomonas ruminantium, a resident of bovine rumen [94,95],
has the highest reported kcat value (185 s − 1 at pH 5.3, 25 ◦ C) for
hydrolysis of xylobiose (X2, Figure 2B), with a K m of 2.1 mM
[96]. GH43 β-xylosidase, XylBH43, from Bacillus halodurans
acting on xylobiose exhibits a kcat of 117 s − 1 and a K m of 3.02 mM
at pH 6.5 and 25 ◦ C [97]. Owing presumably to the spatial
similarity of β-D-xylose and α-L-arabinofuranose (Figure 2C), βxylosidases ordinarily exhibit secondary α-arabinofuranosidase
activity [98,99]. Many β-xylosidases have been studied
exclusively with colorimetric model substrates. However, to
evaluate the enzymes’ potential performance in a saccharification
reactor, it is necessary to determine their activity on relevant
natural substrates, as demonstrated by β-xylosidase GbtXyl43A.
This enzyme exhibits greater catalytic efficiency in hydrolysing
4NPA (4-nitrophenyl-α-L-arabinofuranose) (Figure 2B) than
for 4NPX (4-nitrophenyl-β-D-xylopyranoside) (Figure 2B), but
displays insignificant α-arabinofuranosidase activity and is
limited to β-xylosidase activity on natural substrates [100]. Also,
it has been demonstrated that there is only a modest correlation
between the activity exhibited on a natural substrate, X2, and the
model substrate, 4NPX (Figure 3) (D.B. Jordan and J.D.Braker,
unpublished work) [92].
β-Xylosidases can be inhibited by D-xylose and D-glucose
at low millimolar levels, both of which could be present at
high concentrations in an industrial saccharification process.
This precludes the use of β-xylosidase in an SHF process
because the concentrations of D-glucose and D-xylose would
reach high concentrations (>1 M) in excess of 100-fold that of
K i . Increased attention has been directed towards characterizing
and engineering decreased monosaccharide inhibition of βxylosidases [101]. Also, there are several reports of D-xylosetolerant β-xylosidases from both fungal sources [102–104] and
bacterial sources [105,106].
245
α-Arabinofuranosidases (EC 3.2.1.55), classified in the CAZy
database under GH families 3, 43, 51, 54 and 62 [65],
catalyse the hydrolysis of terminal non-reducing L-arabinose
side chains from the xylan backbone, where they can be found
both singly- and doubly-substituted at C-2 and/or C-3 of the
xylopyranose backbone via α-1,2- and α-1,3-linkages [107].
α-Arabinofuranosidases that cleave L-arabinose exclusively
from glucuronoarabinoxylan polymers are termed glucuronoarabinoxylan arabinofuranohydrolases [108]. Similar to the βxylosidases, many of the α-arabinofuranosidases display secondary β-xylosidase activity [109,110]. α-Arabinofuranosidases have
generally been characterized kinetically on artificial substrates,
which does not correlate with activity on natural substrates. This
is illustrated in the recent study of two α-arabinofuranosidases,
AF30 and AF47, isolated from a fungal pathogen of preharvest
corn, where the relative kcat (AF47/AF30) is ∼ 2 for 4NPA
hydrolysis, whereas the specific activity ratio for corn fibre
glucuronoarabinoxylan hydrolysis (AF47/AF30) is ∼ 0.33 [111].
α-Arabinofuranosidase kcat values for hydrolysis of natural
substrates are rarely reported, a recent notable exception being the
modular GH43 glucuronoarabinoxylan arabinofuranohydrolase
from the ruminal bacterium Fibrobacter succinogenes. This
enzyme has reported kinetic parameters of a kcat of 240 s − 1
and K m of 4.1 mg·ml − 1 acting on natural glucuronoarabinoxylan
substrate for the wild-type enzyme, and moreover, a truncated
site-directed mutant resulted in an increased kcat of 630 s − 1 and
K m of 24 mg·ml − 1 [112].
The α-glucuronidases (EC 3.2.1.131) catalyse hydrolysis of
the 1,2-linked glucuronosyl side chains from xylan [113].
The glucuronosyl substitutions inhibit both enzymatic and
acidic hydrolysis of the xylan polymer [114]. In addition, the
glucuronosyl group can form covalent cross-links to lignin [115–
117]. The majority of α-glucuronidases are categorized as GH67
and remove only the glucuronosyl group that is attached to the
terminal residue at the non-reducing end of xylo-oligosaccharides.
The α-glucuronidase with the highest reported catalytic activity
(a kcat of 202 s − 1 at 40 ◦ C and pH 4.8) on a native substrate
(aldotetraouronic acid) is from Aureobasidium pullulans [118].
Most α-glucuronidases have an optimal pH of 4.5–6.5 and an
optimal temperature of 40–65 ◦ C. There are several examples
of α-glucuronidases that have acidic (pH 3–3.5) optima, but
none with an alkaline optimum [119–121]. The α-glucuronidase
from Thermotoga maritima has the highest reported temperature
optimum (85 ◦ C) [122]. Previously, a new category of αglucuronidases was discovered that acts on a polymeric substrate,
and these enzymes are classified in the GH115 family [123].
CEs (carbohydrate esterases)
CEs are organized into 16 families in the CAZy database.
The CE enzyme activities targeted for glucuronoarabinoxylan
saccharification are acetylxylan esterases (EC 3.1.1.72; CE
families 1–7, 12 and 15) and ferulic acid esterases (EC 3.1.1.73;
CE family 1). CEs belong mainly to the α/β hydrolase fold
superfamily, which includes hydrolases, dehalogenases, lipases
and peroxidases [124]. The active sites generally contain a serinehistidine-carboxylate catalytic triad, and exhibit modest catalytic
rates and much smaller rate enhancements than the GH enzymes.
Alkaline pretreatment conditions could effectively saponify the
ester bonds, rendering the esterases redundant.
Acetylxylan esterases remove acetyl groups from the xylan
backbone. The acetyl groups can attach to the 2-O, 3-O or both
positions of the xylosyl monomer. Most acetylxylan esterases
are in families CE 1, 4 or 5. The enzymes were previously
demonstrated to preferentially remove acetyl groups from the
c The Authors Journal compilation c 2012 Biochemical Society
246
D. B. Jordan and others
2-O position [125]. However, in an aqueous environment, the
acetyl groups can easily migrate between the 2-O and 3-O
positions, thus it is not critical to use two esterases to target the
acetyl groups at both sites. Acetylxylan esterases have long been
recognized as being important for the enzymic saccharification
of acetylated biomass [126]. A kcat of 24 s − 1 at 50 ◦ C has been
determined for the enzyme from T. reesei acting on D-xylose
tetraacetate [127].
Ferulic acid esterases (EC 3.1.1.73) catalyse hydrolysis of
L-arabinose-ferulate ester bonds [128–130]. L-Arabinose side
chains of glucuronoarabinoxylan can be substituted with ferulic
acid (4-hydroxy-3-methoxycinnamic acid) at the C-5 and C-2
hydroxy groups of L-arabinose. There it can form ferulate bridges
consisting of dehydrodimers and dehydrotrimers of various
linkage configurations between carbohydrate chains, and between
carbohydrate chains and lignin (Figure 1). Cross-linking increases
plant cell wall structural rigidity, which simultaneously hinders
access to the hemicellulosic substrate by GH enzymes [131], and
has been likened to “the molecular equivalent of spot-welding a
steel-mesh frame” [132]. Most activity studies with ferulic acid
esterases have been conducted on methyl ester model substrates,
which display a maximum kcat of 200 s − 1 [133,134]. The turnover
number for the release of ferulic acid from hemicellulose derived
from rye grass is approximately 4 s − 1 at 37 ◦ C [135].
FERMENTATION
Economically competitive ethanol production from lignocellulosic materials requires efficient use of both the hexose and
pentose monosaccharides. Although there are other hexose
sugars, in addition to D-glucose, galactose and mannose (e.g.
glucuronic acid, galacturonic acid and rhamnose), depending
on the feedstock, they are present in small amounts relative
to the hemicellulose fraction consisting mainly of D-xylose.
L-Arabinose is also present in the hemicellulose fraction, but
again, it is a small portion of the hemicellulose relative to Dxylose. The recognition that utilization of the D-xylose fraction
will be required to make lignocellulosic ethanol cost-competitive
has resulted in a great deal of work to engineer pathways and
organisms to convert D-xylose.
Fermentative production of biofuels from lignocellulosic
feedstocks provides unique challenges for micro-organisms. In
addition to efficient pentose metabolism and inhibitor tolerance,
the ideal fermenting micro-organism would also maintain the
productivity measures of the current starch- and cane sugarbased ethanol systems, being tolerant to ethanol, low pH, high
osmolarity and high temperature (to lower the cooling cost of removing heat generated by the fermentation). No micro-organism
has been discovered that is capable of fermenting D-xylose at high
titres and high productivity using a homo-ethanol pathway [136].
As a result, several groups have undertaken research to isolate
and/or engineer, through directed and random methods, organisms
with well-defined properties and reliable genetic transformation
systems, such as Zymomonas mobilis [137] and Escherichia
coli [138,139], along with additional yeast species such as
Scheffersomyces stipitis [40]. Despite some success in engineering
these organisms for potential use in lignocellulosic biofuel
production, most are not as tolerant as Saccharomyces cerevisiae
towards ethanol or some of the inhibitors found in lignocellulosic
hydrolysates [140]. Additionally, while numerous organisms
naturally metabolize D-xylose, and some will ferment D-xylose to
ethanol, D-glucose fermentation rates are typically orders of magnitudes lower by these organisms compared with Saccharomyces.
Thus, owing to its high ethanol yield, high productivity under
anaerobic conditions and wide use in industrial fermentation
c The Authors Journal compilation c 2012 Biochemical Society
processes, S. cerevisiae remains the preferred organism for
converting biomass-derived monosaccharides into bioethanol.
Unfortunately, S. cerevisiae does not naturally ferment Dxylose. Several pathways that exist in nature for metabolizing
D-xylose (Figure 4) have been engineered into S. cerevisiae.
Most efforts towards engineering Saccharomyces yeasts for
D-xylose fermentation have focused heavily on reconstitution
of the two xylulose 5-phosphate-producing pathways (i.e. Dxylose reductase/xylitol dehydrogenase or D-xylose isomerase)
(reviewed in [141–144]). Native D-xylose-metabolizing fungi
typically use a two-step reduction/oxidation, whereas bacteria
usually employ a single isomerization of D-xylose. Both pathways
convert D-xylose into xylulose, which is phosphorylated by
xylulokinase to produce the PPP (pentose phosphate pathway)
intermediate xylulose 5-phosphate. A series of carbon-transfer
reactions in the non-oxidative branch of the PPP results in
the production of fructose 6-phosphate and glyceraldehyde 3phosphate, which can be metabolized by S. cerevisiae.
Although some success has been reported on the
aforementioned pathways in S. cerevisiae, a number of
shortcomings remain and warrant exploration of additional
pathways. Another potential pathway to D-xylose metabolism
could be initiated by the enzyme D-xylose dehydrogenase
(Figure 4). This pathway splits at the intermediate 2-oxo-3deoxyxylonate where it is (i) cleaved by an aldolase to generate
pyruvate and glycolaldehyde, or (ii) acted on by two additional
enzymes to form α-oxoglutarate. This pathway has the potential
to avoid limitations imposed by low PPP flux in S. cerevisiae
because xylulose 5-phosphate is not formed. The initial enzyme
of the pathway, D-xylose dehydrogenase, has a higher affinity
(K m <4 mM) for D-xylose [145–148] than D-xylose reductases or
isomerases which have K m values of approximately 20–60 mM
[149–151]. D-Xylose isomerases with higher affinity for xylose
are common in bacteria; however, many of these bacterial Dxylose isomerases do not work in S. cerevisiae, and D-xylose
isomerases that have been shown to function in this yeast show
poor affinity for D-xylose. Considering the inefficient transport of
D-xylose into S. cerevisiae (see below), a pathway with increased
affinity for D-xylose could improve D-xylose utilization. Archaea
and some bacteria are the only organisms to date that have been
shown to contain complete pathways to metabolize D-xylose
via an initial oxidation [146–148]. D-Xylose dehydrogenase
has recently been expressed in Kluyveromyces lactis for the
production of xylonic acid [152], but engineering of the entire
pathway has not been reported.
Another possible route to explore is a branch of the xylulose 5phosphate route which includes cleavage of xylulose 5-phosphate
to acetyl phosphate, and glyceraldehyde 3-phosphate by the
enzyme PK (phosphoketolase) (Figure 4). Acetyl phosphate can
be converted into acetyl-CoA, which can be metabolized via the
TCA (tricarboxylic acid) cycle. Alternatively, it can be further
converted into ethanol by acetaldehyde dehydrogenase (acylating)
and alcohol dehydrogenase (not shown). The role of PK in Dxylose (and D-glucose) metabolism in certain bacteria has long
been established [153,154]. Later, PK activity was shown to be
induced in yeasts up to 70-fold by D-xylose [155], suggesting that
this branch plays an important role in D-xylose metabolism, at
least in certain yeasts. A PK pathway has been engineered into
S. cerevisiae, resulting in an increased ethanol yield [156]. Aside
from this work, little has been reported pursuing this approach.
In addition to exploring the D-xylose dehydrogenase and PK
routes to D-xylose metabolism, a number of additional problems
associated with the two main D-xylose metabolism pathways are
also in need of resolution. Additional concerns related to D-xylose
transport, redox imbalance, flux limitations through the PPP and
Cell walls to ethanol
Figure 4
D-Xylose
247
degradation pathways in nature
Xylulose 5-phosphate route: XR, D-xylose reductase (E.C. 1.1.1.21); XOH DH, xylitol dehydrogenase (E.C. 1.1.1.9); XI, D-xylose isomerase (E.C. 5.3.1.5); XK, xylulokinase (E.C. 2.7.1.17); RPE,
ribulose 5-phosphate-3-epimerase (E.C 5.1.3.1); TKL, transketolase (E.C. 2.2.1.1); AP, acylphosphatase (E.C. 3.6.1.7); AK, acetate kinase (E.C. 2.7.2.1); PTA, phosphotrans-acetylase (E.C. 2.3.1.8);
and ACS, acetyl-CoA synthetase (E.C. 6.2.1.1). Non-phosphorylated intermediate route: XDH, D-xylose dehydrogenase (E.C. 1.1.1.175, E.C. 1.1.1.179); XL, xylonolactonase (E.C. 3.1.1.68); XAD,
xylonate dehydratase (E.C. 4.2.1.82); KDXA, 2-oxo-3-deoxy xylonate aldolase (E.C. 4.1.2.28); KDXD, 2-oxo-3-deoxy xylonate dehydratase (E.C. 4.2.1.-); and αKGSADH, α-oxoglutaric semialdehyde
dehydrogenase (E.C. 1.2.1.3).
the role of gluconeogenesis must be resolved to further increase
D-xylose fermentation rates.
D-Xylose
transport
Saccharomyces yeasts do not possess D-xylose-specific
transporters. D-Xylose gains entry to the cell through the
HXT family of D-glucose transporters [157]. To overcome
this bottleneck, a number of groups have expressed D-xylose
transporters from bacterial, yeast and plant species into yeast
[158–163]. This approach has improved D-xylose uptake for some
of the strains. Unfortunately, progress in this area has been slow
because the number of known D-xylose transporters available
for expression and study in S. cerevisiae is limited. While kinetic
data for D-xylose transport in native D-xylose-metabolizing yeasts
have been available for decades [164–170], most of the D-xylosespecific transporters from these organisms have not been isolated.
Work with additional transporters, preferably passive transporters
that do not require the expenditure of energy, will shed additional
light on the efficiency of D-xylose transport and its role in
improving D-xylose fermentation.
Redox imbalance
Under anaerobic conditions, cofactor differences between the first
two enzymes in the fungal pathway result in NADPH depletion
and NADH accumulation [171]. NADH accumulation favours
the production of xylitol and glycerol at the expense of ethanol.
NADPH depletion results in decreased D-xylose reduction and
limits reducing power for generating cell biomass and for inhibitor
tolerance [172]. Multiple strategies have been investigated to
alter the redox balance. Protein engineering of the S. stipitis
xylose reductase to increase the use of NADH over NADPH by
the enzyme has shown some benefit [173–176]. However, many
of the strategies have failed to show significant improvement,
and others have intensified the problem [177,178]. Theoretically,
expression of a D-xylose isomerase would alleviate this imbalance
of cofactors [179]. However, most D-xylose isomerases expressed
in S. cerevisiae function poorly and the equilibrium does favour
D-xylose. NADPH may still be limiting in S. cerevisiae expressing
a D-xylose isomerase pathway due to its inability to recycle
D-xylose back to D-glucose 6-phosphate to regenerate NADPH
through the oxidative branch of the PPP [180].
PPP flux limitations
Although low flux through the PPP in S. cerevisiae is beneficial
for D-glucose fermentation, low flux through the non-oxidative
part of the PPP limits D-xylose fermentation. Elevated expression
of the non-oxidative branch enzymes has been used to increase
D-xylose fermentation [181]. It was also previously discovered
that strains used for production of ethanol from cane sugar have
c The Authors Journal compilation c 2012 Biochemical Society
248
D. B. Jordan and others
duplicate genes involved in thiamine (vitamin B1 ) and vitamin B6
synthesis [182]. Thiamine pyrophosphate is a cofactor of the nonoxidative PPP pathway enzyme transketolase, and this adaptive
duplication may be helpful for increasing flux through the PPP.
NADPH produced by the oxidative branch of the PPP pathway
is also used in detoxifying furfural and hydroxymethylfurfural,
and low flux through this pathway potentially further limits the
ability of Saccharomyces to efficiently ferment D-xylose from
lignocellulosic hydrolysates [172].
Requirement for gluconeogenesis
NADPH regeneration in yeast occurs mainly through the oxidative
branch of the PPP. Native D-xylose-utilizing yeasts appear to
recycle D-xylose back to D-glucose 6-phosphate to be used
for glucan synthesis of cell wall components and in the
oxidative branch of the PPP for NADPH production required
for anabolic reductive reactions. S. cerevisiae expressing the
S. stipitis D-xylose reductase/xylitol dehydrogenase pathway
was recently shown to be unable to induce the genes required
to regenerate NADPH in this manner, thus limiting D-xylose
utilization [180]. Resolving the redox imbalance by use of a
D-xylose isomerase pathway or introducing alternative NADPH
regeneration mechanisms will alleviate the need for recycling
D-xylose to D-glucose, but may not eliminate it. Regardless of
the pathway used to metabolize D-xylose, D-glucose will still
be required for the glucan component of the cell wall and
NADPH (via D-glucose 6-phosphate and the oxidative PPP) for
cell growth and inhibitor tolerance. Whereas native D-xyloseutilizing yeasts are able to induce enzymatic activity to produce Dglucose 6-phosphate from D-xylose, the transcriptional response
of S. cerevisiae to D-xylose is not optimized for this pathway,
and may actually induce genes that negatively affect D-xylose
utilization. A recent paper describes expression of a cellodextrin
transporter and an intracellular β-glucosidase in S. cerevisiae
grown in a medium containing cellobiose and D-xylose [183].
This strategy partially overcomes the problem by providing low
levels of D-glucose during D-xylose fermentation. SSCF can also
provide low levels of D-glucose during D-xylose fermentation,
allowing more efficient D-xylose uptake and fermentation.
D-Xylose-regulated
promoters
A wide variety of promoters are available for constitutive and
regulated expression of foreign genes in S. cerevisiae. In many
cases, however, constitutive expression is a waste of cellular
resources during the D-glucose phase, when D-xylose is not
metabolized. For example, the expression of genes for D-xylose
transport, or for improving redox imbalance during the Dxylose consumption phase, could induce an imbalance during
D-glucose fermentation. The ability to fine-tune the expression
of the multiple genes required for D-xylose fermentation, to
be expressed only when needed, will allow better control of
the genetically engineered pathways. Unfortunately, D-xyloseregulated promoters are not yet available for control of gene
expression in S. cerevisiae.
PATH FORWARD
Improving the commercial feasibility of second generation
bioethanol production requires integrating goals for feedstock
development, pretreatment, enzymatic saccharification, fermentation, waste treatment and process water recycling. In the
area of feedstock development, better analytical techniques will
c The Authors Journal compilation c 2012 Biochemical Society
allow for better understanding of the interrelationship among
the major cell components, hemicellulose, lignin, pectin and
cellulose. Immediate progress in feedstock development is likely
to be dominated by lignin modification because decreased
lignin content is highly correlated with improved enzymatic
saccharification yields, and considerable progress has been
made in determining lignin synthetic pathways [25,26]. Several
pretreatments are able to affect alteration of cellulose lattice
structure, of which room temperature ionic liquids are the newest
candidates; however, doubts remain regarding cost and the ability
to recycle these solvents. Most pretreatment strategies being
pursued [44,184] do not fully hydrolyse xylan in order to avoid
generation of high levels of simple sugars in the pretreatment step;
ultimately, such procedures maximize yield of monosaccharides
and minimize formation of furans [41,42]. Greater understanding
of the structure of residual fermentation oligosaccharides and their
impact on complete hydrolysis and/or enzyme activity may lead
to knowledge of additional enzyme activities needed to complete
saccharification. Newly discovered enzymes (formerly GH61)
that oxidatively cleave cellulose offer an exciting possibility for
obtaining large improvements in cellulose deconstruction rates
[66–68]. Natural enzyme sources, metagenomic DNA libraries
and genetically engineered libraries should be searched for
enzymes with improved turnover numbers on natural substrates,
increased tolerance to soluble inhibitors (e.g. D-glucose, D-xylose
and furans), and lower non-specific binding to lignin.
Improving the efficiency of D-xylose fermentation will remain
a major research focus. Although yield has been improved
considerably (to >0.4 g of ethanol/g of D-xylose) [185], specific
ethanol productivity (g of ethanol/g of cells per h) from D-xylose
still lags D-glucose fermentation [143] by an order of magnitude.
In the future, genetic engineering of strains will continue to be
a powerful tool to improve yeast strains, as work with native
D-xylose-utilizing yeasts has uncovered genes that may assist
with D-xylose metabolism that are either not present or regulated
improperly in D-xylose-grown S. cerevisiae. Additionally, a
number of microarray and proteomic studies have also shown that
multiple genes are regulated in strains that have been engineered
to ferment D-xylose, suggesting that D-xylose utilization will
depend on genes and/or pathways beyond what have already been
engineered into the strains [186–188] and will probably need
to be evolved/adapted simultaneously. Increased understanding
of how native D-xylose-utilizing yeasts efficiently metabolize Dxylose and re-engineering these additional pathways, along with
strategies for improved inhibitor tolerance, into robust industrial
S. cerevisiae strains will lead to further improvements in ethanol
productivity, yield and cost competitiveness.
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Received 28 October 2011/30 November 2011; accepted 2 December 2011
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