The potential of microbial processes for lignocellulosic biomass conversion to
ethanol: a review
Davide Dionisi,* Materials and Chemical Engineering group, School of Engineering, University of Aberdeen,
Aberdeen, AB24 3UE, UK
James A. Anderson, Surface Chemistry and Catalysis group, School of Natural and Computing Sciences/
Materials and Chemical Engineering group, School of Engineering, University of Aberdeen, Aberdeen, AB24
3UE, UK
Federico Aulenta, Water Research Institute, National Research Council (CNR-IRSA), Via Salaria km 29.300
C.P. 10, 00015 Monterotondo (RM), Italy
Alan McCue, Surface Chemistry and Catalysis group, School of Natural and Computing Sciences, University of
Aberdeen, Aberdeen, AB24 3UE, UK
Graeme Paton, Department of Plant and Soil Science, University of Aberdeen, Cruickshank Building,
St. Machar Drive, Aberdeen, AB24 3UU, UK
Abstract
BACKGROUND: This paper assesses the feasibility of a single- or multi-stage process
entirely based on microbial cultures, with no or minimal non-biological pretreatment and with
no external enzyme addition, for the conversion of lignocellulosic materials into ethanol. The
considered process involves three distinct microbial processes, which can be possibly
combined in one single reaction stage: a) lignin hydrolysis; b) cellulose and hemicelluloses
hydrolysis; c) glucose fermentation to ethanol. This paper critically reviews the literature on
the three microbial processes and compares the rates of microbial processes with the ones
of the alternative physico-chemical pretreatment processes.
RESULTS: There is a large number of microbial species that can perform each of the three
processes required for the conversion of lignocellulosic biomass to ethanol, although only
one species has been unquestionably reported, so far, to be able to hydrolyse lignin under
anaerobic conditions; another challenge is controlling the anaerobic fermentation of glucose
to ethanol with mixed cultures; the rates of the microbial processes reported so far in the
literature are generally lower than the rates obtained with physico-chemical pretreatments.
CONCLUSIONS: While in principle the whole process from lignocellulosic biomass to
bioethanol can be carried out with existing, non engineered microorganisms, there is need
for further research to obtain rates and yields which are commercially attractive.
Keywords: bioethanol; cellulose; lignin; microbial hydrolysis; mixed cultures
* email: [email protected], phone: +44 (0)1224 272814
1
1. Introduction
“Second generation” bioethanol produced from lignocellulosic biomass is gaining increasing
interest, since it can avoid the fuel vs. food competition which is intrinsic in “first generation”
bioethanol produced from starch- or sugar-based materials such as corn or sugarcane.1
Lignocellulosic materials such as the organic fraction of municipal solid wastes, agricultural
wastes, forestry residues, etc, represent an almost unlimited resource, which could be
potentially used for bioethanol production.2 However, production of bioethanol from
lignocellulosic materials poses significant technical and economical challenges. According to
a UN panel of experts,3 in 2012 biofuels production from lignocellulosic feedstock accounted
for some 140 million litres per year, only 0.15% of the total production. The world’s first
commercial scale cellulosic ethanol plant is considered to be the one in Crescentino, Italy, by
Beta Renawables, with a full capacity of 75 million litres a year, which was started up in
2013.4
The bioethanol production process from lignocellulosic materials usually consists of several
steps: pretreatment to break the lignin structure and make cellulose (and/or hemicellulose)
available for hydrolysis, hydrolysis to hydrolyse cellulose to glucose, fermentation to convert
glucose into ethanol and separation processes for ethanol purification.5,6 As far as lignin
hydrolysis is concerned, the most widely adopted pretreatments are steam explosion, acid
hydrolysis, or ammonia fibre expansion (AFEX).7 As an example, the Iogen demonstration
plant (Canada), which has an average productivity of about 300 m3 ethanol/yr receives
wheat straw, corn stover and bagasse as feedstocks and uses a modified steam explosion
process
to
make
cellulose
more
accessible
for
(http://www.iogen.ca/technology/cellulosic-ethanol.html)).
the
These
following
chemical
steps
(Iogen,
or
physical
pretreatment processes usually give good lignin hydrolysis and they make cellulose and
hemicelluloses free for the next hydrolysis stages. However, they require conditions of high
temperature and/or high pressure, or the addition of significant amounts of chemicals. These
conditions make these processes expensive and limit their economic attractiveness at
commercial scale. Also, the severe operating conditions used in these processes may
generate toxic substances which inhibit the following stages and therefore a detoxification
stage is often necessary.8 The subsequent step of the process is cellulose and
hemicellulose hydrolysis to generate glucose and other sugars. This step is usually carried
out enzymatically, using commercially available or on-site produced cellulase enzymes. For
example, the Iogen demonstration plant mentioned above and the Abengoa demonstration
plant, (Abengoa, http://www.abengoa.com/web/en/innovacion/casos_exito/) which uses
wheat and barley straw as feedstock, both use enzymatic hydrolysis. Finally, ethanol
production from glucose is usually carried out using pure cultures of selected species,
2
usually the yeast Saccharomyces cerevisiae. While this yeast allows obtaining a very high
ethanol yield and high ethanol productivity, the use of pure cultures has some
disadvantages: pure cultures usually have a very narrow substrate spectrum, e.g. nonengineered S. cerevisiae cannot metabolise xylose, the main component in hemicellulose,
and this limits the range of feedstock that can be used; pure cultures of fermentative
microorganisms usually cannot hydrolyse cellulose, and this requires the addition of the
cellulose hydrolysis stage; the use of pure cultures requires sterilisation of the fermentation
vessel and of all the process lines leading to them, and this causes additional costs.
All of the described factors contribute to the high cost of ethanol production from
lignocellulosic biomass. This paper investigates, by critically analysing the relevant literature,
the feasibility of an alternative process for ethanol production from lignocellulosic biomass. In
contrast to the physico-chemical pretreatment and hydrolysis processes described above,
the process investigated in this paper is entirely, or almost entirely, based on microbial
processes, i.e. based on the physical contact of microorganisms with the substrate. In the
investigated process, the three distinct processes required to convert lignocellulose to
ethanol, i.e. lignin hydrolysis, cellulose and hemicellulose hydrolysis and glucose and other
sugars fermentation, are all carried out by different microorganisms, which could co-exist in
the same reactor, or could be present in different reactors in sequence. Obviously, the option
of a single reactor where all the different microbial species co-exist would be preferable from
an economic point of view. This would constitute an open (undefined) mixed culture, where
the microorganisms responsible for the various processes are selected from a mixed-culture
inoculum due to the applied process conditions (e.g. nature of the feedstock, residence time,
pH, temperature). The process investigated in this study has many aspects in common with
simultaneous saccharification and cofermentation (SSCF) and consolidated bioprocessing
(CBP) processes, described by Lynd et al.9 In SSCF an aerobic reactor is used for cellulase
production, and the produced cellulases are then used in a subsequent anaerobic reactor
where cellulose hydrolysis and sugars (both hexoses and pentoses) fermentation to ethanol
takes place. In CBP the production of cellulases and the fermentation of sugars take all
place in the same anaerobic reactor. However, there are two main differences between
SSCF and CBP process and the process investigated in this study (Figure 1): SSCF and
CBP are usually thought to come after a chemical-physical pretreatment step for lignin
hydrolysis, while the process considered here includes microbial lignin pretreatment; SSCF
and CBP are usually thought as pure cultures processes, using native microorganisms or
genetically engineered ones, while the process studied in this paper is aimed at using open
mixed microbial cultures.
3
Microbial processes based on open mixed cultures are gaining increasing interest due to
their lower cost and higher flexibility compared to the traditional pure culture processes.10 A
well known industrial process involving the use of open mixed cultures is anaerobic digestion
of organic materials to methane,11 whereas more recently the use of mixed cultures to
produce hydrogen,12 ethanol13 and biodegradable plastics14-16 has been investigated. An
interesting mixed culture process is the MixAlco process,17 which converts biomass to
carboxylic acids salts, which are then chemically converted to hydrocarbon fuels.
An entirely microbial process, if proven feasible, would have obvious cost advantages
compared to existing processes for lignocellulosic ethanol production, due to the use of
atmospheric pressure, temperature close to ambient, and no addition of expensive
chemicals or enzymes. The aim of this paper is to review the literature on the individual
processes that are necessary to obtain microbial conversion of lignocellulosic materials into
bioethanol, i.e. microbial lignin and cellulose hydrolysis and glucose and xylose fermentation
by mixed cultures. This review for the first time reports rate values from the literature studies
and critically discusses the effect of process parameters, in order to help identifying the
conditions that maximise the rates and yields of the microbial processes.
In principle, an alternative microorganism-based approach to convert lignocellulosic
materials into ethanol is to use genetically modified microorganisms, as opposed to open
mixed cultures of naturally occurring species. The rationale behind this is that the ability to
hydrolyse lignin and cellulose/hemicelluloses and to ferment sugars to ethanol with high
yields could be introduced into microorganisms using metabolic engineering techniques.
Without attempting to review the vast area of metabolic engineering for ethanol production,
which has been reviewed elsewhere,18,19 this paper compares the use of open mixed
cultures and of genetically modified microorganisms for lignin and cellulose hydrolysis and
for glucose and xylose fermentation to ethanol.
4
2. Lignocellulosic biomass
The main components of lignocellulosic biomass are lignin, cellulose and hemicelluloses.
Lignin is an aromatic polymer where the substituents are connected by ether and carboncarbon linkages. The main building blocks in lignin are p-coumaryl alcohol (p-hydroxyphenyl
propanol), coniferyl alcohol (guaiacyl propanol) and sinapyl alcohol (syringyl propanol).20 The
molecular weight of lignin is variable but is typically very high, 600-1000 kDa.21 Cellulose is a
polysaccharide composed of (1-4) linked D-glucose units, with molecular weight up to >500
kDa.22 Hemicellulose is a polysaccharide composed of various carbohydrate monomers,
mainly xylan, arabinose, mannose and glucose, present in different ratios in the various
materials. The molecular weight of hemicelluloses is usually lower than the one of
cellulose.23 The composition of various lignocellulosic materials, potential feedstock for
ethanol production, is reported in Table 1. Values in Table 1 are to be considered as
orientative values only, because the measured lignin content in a given biomass species is
highly dependent on the biomass history and on the measurement method used.24
The hemicellulose content in the feedstock is important due to the fact that hemicellulose
hydrolysis generates monosaccharides other than glucose, which need to be converted to
ethanol in order to avoid yield loss in the process. The conversion of non-glucose sugars to
ethanol poses significant challenges as discussed later in this paper. However,
hemicellulose hydrolysis is not considered to be a significant challenge, since hemicellulose
is hydrolysed more easily than cellulose,25 and it is expected that a mixed microbial culture
that hydrolyses cellulose would also be able to hydrolyse hemicelluloses.9 Therefore in this
paper the hydrolysis of only lignin and cellulose is discussed.
5
3. Microbial degradation of lignocellulosic biomass
The aim of this section is to identify, on the basis of the existing literature, whether there are
microorganisms which are able to catalyse the various steps required for lignocellulosic
biomass conversion to ethanol, i.e. lignin hydrolysis, cellulose hydrolysis and sugars
fermentation to ethanol. Both pure culture and mixed cultures of naturally existing
microorganisms and genetically modified microorganisms are considered. The rates of
microbial processes are reported and compared to the rates of the alternative chemicalphysical processes.
3.1 Lignin
3.1.1 Anaerobic conditions
High molecular weight lignin is generally considered as non biodegradable, or only
biodegradable at insignificant rates, in the absence of oxygen,21,26,27 even though there is
some evidence of the contrary.28,29 On the other hand, there is enough evidence to suggest
that the various lignin building blocks, constituted by the various aromatic compounds as
monomers or oligomers, are readily metabolised under anaerobic conditions.30,31 The very
limited evidence of anaerobic lignin biodegradation reported in the literature can be
attributed21 to non-lignin components or to low-molecular weight materials (<600 Da).
However, Benner et al.29 using mixed cultures, reported anaerobic lignin biodegradation
rates of up to 37% of the aerobic rates, for several lignocellulosic marine or wetland plants.
Silanikove and Brosh28 reported 45-58% anaerobic metabolisation of lignin in wheat-straw,
by the rumen bacteria in the goats’ gastrointestinal tract.
Only very recently, a convincing evidence of lignin degradation under anaerobic conditions
has been presented.32 The authors observed that the majority (85%) of insoluble switchgrass
biomass, that had not been previously chemically treated, was converted at 78°C by the
anaerobic bacterium Caldicellulosiruptor bescii. Interestingly, the glucose/xylose/lignin ratio
did not substantially change over the incubation period (3 successive cycles of 5 days each)
providing an indication that the three major biomass components, including lignin, were
solubilised and/or metabolized at comparable rates. A mass balance revealed that lignin was
not assimilated, only carbohydrates served as carbon and energy sources. Lignin
degradation was confirmed by gas-chromatography-mass spectrometry analyses which
revealed the presence of lignin-derived aromatic compounds, such as syringylglycerol,
guaiacylglycerol, and phenolic acids, in the spent culture broth.
6
Indirect evidence of lignin degradation/hydrolysis under anaerobic conditions can be
obtained observing anaerobic digestion studies, which are always carried out using open
mixed microbial cultures, with lignocellulosic materials as feedstock (Table 2). In general,
methane production has been reported with many lignocellulosic materials, even though the
possible lignin degradation and its extent are not usually measured. Methane production has
been reported even with feedstocks with high lignin content, such as Mirabilis and Ipomoea
fistulosa leaves33 and woody biomass such as poplar.34 Several studies investigated the
effect of lignin content on the anaerobic degradability of lignocellulosic biomass. Triolo et
al.35 observed that the higher the lignin content in the raw material the lower the methane
production. However, Tong et al.36 found only a poor correlation between the lignin content
of various lignocellulosic substrates and their methane production rate. This indicates that
biodegradation of lignocellulosic substrates is not only affected by the lignin content, but also
by other factors such degree of association between lignin and carbohydrates, cellulose
crystallinity and others.37 According to Turick et al.,34 one of the reasons why many
investigators observed only poor methane production from woody or other highly lignified
biomass is that the time length of the tests is not long enough. These authors carried out
tests for 100 days and observed substantial methane production from woody biomass.
Interestingly, they observed that most of the methane production occurred after more than
50 days from the start of the test, and they explained this behaviour with the need of the
microorganisms to become able to degrade lignin, making the cellulosic materials initially
shielded by lignin available.
Overall, even though so far only limited evidence for anaerobic lignin metabolization or
solubilisation is available, it is apparent that a significant fraction of the cellulose or
hemicellulose carbohydrates in lignocellulosic materials becomes available during anaerobic
digestion, and this indicates that likely lignin hydrolysis occurs under anaerobic conditions.
3.1.2 Aerobic conditions
In contrast to anaerobic conditions, there is wide literature evidence that lignin is
biodegraded under aerobic conditions. Various species of bacteria and fungi have been
reported to biodegrade lignin (Table 3). Fungi, in particular white-rot fungi, have been
studied more extensively than bacteria for their lignin biodegradation ability, and they are
generally considered more interesting than bacteria as a pretreatment of lignocellulosic
materials at industrial scale.38 Aerobic biodegradation of lignin has also been reported in
mixed culture studies, usually carried out in composting environments (Table 4). While
substantial lignin degradation has been reported for a wide range of lignocellulosic
substrates, usually the treatment times are quite long and lignin degradation is not complete.
7
The literature evidence so far indicates that lignin cannot be used as a sole carbon and
energy source, but requires an additional substrate to support microbial growth.21 The
growth substrate can be the glucose or carbohydrates units contained in the cellulose or
hemicellulose inside the lignin matrix,39 or can be externally added. This is important from
the process point of view, since it is expected that microbial lignin depolymerisation may
require the use of part of the cellulose and hemicellulose as growth substrate for the
microorganisms, making it not available for conversion to ethanol. However, in a mixed
culture environment with real lignocellulosic biomass as feedstock, other carbon and energy
sources than polysaccharides may be available (e.g. proteins) and so the loss of
carbohydrates for ethanol production may be avoided.
3.1.3 Effect of pH
The optimum pH for aerobic lignin degradation by fungi is approx. 4.0-4.5 and significantly
lower biodegradation rates are observed when the pH is lower than 4 or above 5.21,40 pH in
the range 4-5 is considered the optimum for the growth of most white-rot fungi. For bacteria,
no difference in lignin degradation rate was observed in the pH range 5.3-7.8, for a mixed
culture.41 These studies seem to indicate that the optimum pH for lignin biodegradation
coincides with the usual optimum pH for microorganism growth on carbon substrates.
3.1.4 Effect of feedstock particle size
The biodegradation of lignin occurs extracellularly and by decreasing the particle size it is
expected that the surface to volume ratio of lignocellulosic biomass increases, so an
increase in lignin biodegradation rate is expected. Limited experimental investigation has
been carried out however on the effect of feedstock particle size on lignin biodegradation
and usually studies are carried out with a single feedstock size, which is in the mm 42-44 or
cm45,46 range or not reported.47 Zimmerman and Broda48 observed higher lignin degradation
for straw pre-treated with a vibratory ball mill (particle size 2-5 m) than for straw ground
with a blender (particle size 0.5-1 mm). Even though still limited, investigation on the effect of
particle size has been carried out in the area of anaerobic digestion of lignocellulosic
biomass. Mshandete et al.49 observed that total fibre degradation during anaerobic digestion
to methane increased from 31% to 70% when the feedstock was grinded to 2 mm fibres
compared to untreated fibres (larger than 100 mm). Sharma et al.33 observed that the
quantity of biogas produced increased when the feedstock particle size was reduced. The
range of sizes in their study was 0.1-30 mm. However, they did not investigate specifically
lignin degradation.
8
3.1.5 Lignin degradation rates
The main limitation of microorganisms-based pretreatment processes for lignocellulosic
biomass conversion to ethanol is considered to be the low rate. However, very limited direct
information on the rate of lignin hydrolysis under aerobic conditions is present in the
literature. Table 5 reports lignin hydrolysis rates using fungi under aerobic conditions,
calculated by the authors of this paper on the basis of literature data. The maximum rate is
approx 0.1 g/L/h. Under anaerobic conditions, the only evidence of lignin degradation32 gives
a lignin degradation rate of 0.012 g/L/h. It is important to observe that the rates reported in
Table 5 have been obtained at lab scale with very small volumes and under non-optimised
conditions. Several variables could be in principle optimised to maximise the lignin
degradation rate: biomass concentration, oxygen concentration, pH and particle size of the
feedstock.
It is worth comparing the lignin degradation rates reported in Table 5 with the rates obtained
with non-biological pretreatment stages reported in the literature7 (Table 6). It is evident that
the lignin degradation rates obtained with fungi are in general at least one order of
magnitude lower than the ones obtained with chemical pretreatments. However, the process
conditions required by the chemical pretreatments are much more severe, with much higher
temperature and usually (with the exception of the hot water treatment) with the addition of
chemicals, which obviously cause higher process costs.
3.1.6 Lignin hydrolysis by genetically modified microorganisms
While the focus of genetic engineering for lignin hydrolysis has been on genetically
manipulating lignin biosynthesis in plants in order to reduce lignin content and make its
hydrolsysis easier,50,51 there are so far no reported attempts to genetically modify
microorganisms in order to make them capable of lignin hydrolysis.52 The reason for this is
probably the large number of enzymes which are potentially dedicated to lignin hydrolysis in
naturally occurring lignin-hydrolysing microorganisms, such as the white-rot fungus
Phanerochaete chrysosporium.53,54 It is possible, however, that only a few out of the whole
spectrum of lignases might be needed in industrial processes,52 therefore making genetic
engineering of microorganisms for lignin hydrolysis a more feasible option.
3.2. Cellulose
Unlike the case of lignin, a wide range of bacteria and fungi have been reported to hydrolyse
cellulose under anaerobic or aerobic conditions.
9
3.2.1 Anaerobic conditions
Table 7 summarises bacteria and fungi that have been reported to hydrolyse cellulose under
anaerobic conditions. Unlike the case of lignin, complete cellulose hydrolysis can be
obtained under anaerobic conditions, provided that the contact or residence time is
adequate. Table 8 reports literature evidence for cellulose degradation by mixed cultures
under anaerobic conditions, confirming that virtually complete cellulose hydrolysis is
possible. Most of the data in Table 8 refer to batch tests with an unacclimated inoculum, and
this explains the long time required for cellulose degradation.
The enzyme groups responsible for cellulose hydrolysis are very similar under anaerobic
and aerobic conditions but the spatial arrangement of the enzymes can be different.9 Under
anaerobic conditions cellulolytic enzymes are often bound to the external membrane of the
cell, even though in some cases they are present as free enzymes in the liquid medium.
Under aerobic conditions the enzymes are usually excreted in the liquid medium and are not
attached to the cell membrane.
3.2.2 Aerobic conditions
Table 9 summarises microorganisms which have been reported to hydrolyse cellulose under
aerobic conditions. Consistent with findings for anaerobic conditions, virtually complete
cellulose hydrolysis can be obtained under aerobic conditions. Similar evidence is obtained
for mixed cultures studies (Table 10), which usually refer to composting environments. An
interesting observation is that usually the rate of cellulose hydrolysis is comparable under
anaerobic and aerobic conditions.9 In terms of maximising the rate of cellulose hydrolysis,
this means that no preference should be given to aerobic compared to anaerobic conditions.
3.2.3 Effect of pH
For anaerobic and aerobic bacteria the optimum pH for cellulose hydrolysis is usually in the
range 6.5-8.0. For anaerobic bacteria, Shi and Weimer55 found an optimum pH of 6.5 for
cellulose hydrolysis with Ruminococcus flavefaciens, and Weimer56 with Fibrobacter
succinogenes found very little influence of pH on cellulose hydrolysis in the pH range 6.16.8. Using ruminal microbes under anaerobic conditions Hu et al.57,58 found no cellulose
hydrolysis at pH<5.5 and a very low rate at pH<6.0, the optimum pH being 7.0-7.5. Berquist
et al.59 reviewed various cellulolytic thermophilic bacteria, employing either aerobic or
anaerobic conditions, and reported an optimum pH in the range 7.0-8.1 in most cases. This
reported evidence is consistent with a study60 on the anaerobic hydrolysis of organic waste,
10
partially composed of lignocellulosic biomass, where an approximately 3-fold increase in the
hydrolysis rate was observed when the pH was increased from 5 to 7.
3.2.4 Effect of particle size
In general, as cellulose hydrolysis is dependent on the contact between the solid cellulose
and either the microorganisms or the excreted cellulolytic enzymes, it is expected that the
rate of cellulose hydrolysis should increase as cellulose particle size decreases, since the
area per unit volume increases. However, the beneficial effect of feedstock particle size
reduction is expected to depend on the substrate-to-microorganisms ratio, as well as on the
particle size.61 A few experimental studies have been conducted on the effect of particle size
on cellulose hydrolysis rate. Weimer et al.62 observed a decrease in the hydrolysis rate and
an increase in the induction time when the cellulose particle size increased. Similarly, Hu et
al.57 reported faster cellulose hydrolysis rate for 50 than for 100 m sized particles. Similar
evidence was obtained in continuous studies. Chyi and Dague63 observed a faster hydrolysis
rate with 20 than with 50 m particles.
3.2.5 Cellulose hydrolysis rates
Table 11 summarises microbial cellulose hydrolysis rates, calculated by the authors of this
paper on the basis of literature data. The reported rates for microbial cellulose hydrolysis
are, in general, higher than the corresponding rates for lignin hydrolysis (Table 5), therefore
indicating that the critical stage in the process is the lignin hydrolysis. Cellulose hydrolysis
rates up to about 0.3 g cellulose/L/h have been reported, both under aerobic and anaerobic
conditions. As observed for lignin degradation rates, the rates reported in Table 11 have
usually been obtained with very low volume systems and have not been optimised.
Important factors that can increase the rates of microbial cellulose hydrolysis are: biomass
concentration, cellulose concentration, reactor dilution rate, temperature, pH and cellulose
particle size, as discussed in previous sections. One of the most successful technologies for
cellulose hydrolysis is enzymatic hydrolysis, where the cellulolytic enzymes are externally
generated and added to the liquid mixture. Table 12 reports typical rates for enzymatic
cellulose hydrolysis. It is evident that enzymatic hydrolysis is generally faster than microbial
hydrolysis. However it has to be taken into account that enzymatic hydrolysis has a higher
capital and/or operational cost than microbial hydrolysis due to the need for external
generation or purchase of the enzymes.
11
3.2.6 Cellulose hydrolysis by genetically modified microorganisms
In recent years there has been a considerable interest in engineering microorganisms in
order to make them capable of hydrolysing cellulose.9 Particular attention has been given to
adding the cellulose-hydrolysis capability in microorganisms which are naturally able to
ferment glucose to ethanol with high yields. Several cellulase-encoding genes have been
expressed in various bacteria and yeasts, such as Zymomonas mobilis,64 Klebsiella
oxytoca,65,
66
Saccharomyces cerevisiae67 and others. However, while the results are in
general promising and encourage further research in this area, there is still no evidence that
a cellulose hydrolysis capability at rates which are high enough for a commercial process
has been engineered in genetically modified microorganisms. Most of the engineered
microorganisms reported so far have gained the capability of hydrolysing cellulose
derivatives but not native or crystalline cellulose. Genetically modified K. oxytoca exhibited
good capability to hydrolyse soluble carboxy-methyl cellulose65 or phosphoric acid-swollen
Avicel,66 but very limited ability to hydrolyse crystalline Sigmacell 50.65 The yeast S.
cerevisiae, expressing cellulases from Bacillus species, has been reported67 to have activity
on filter paper but was not able to grow in the absence of externally-added cellulases. The
same yeast has been engineered to hydrolyse phosphoric acid-swollen cellulose. 68, 69
3.3. Fermentation of carbohydrates to ethanol by mixed microbial cultures
Once cellulose and hemicellose have been hydrolysed, monomeric sugars need to be
fermented to ethanol. The main sugars that are present in the hydrolysis products of
lignocellulosic biomass are glucose, present in cellulose and in minor fractions in
hemicellulose, and xylose, which is often the main component of hemicellulose (Table 1).
The fermentation of glucose and xylose by mixed microbial cultures is reviewed in sections
3.3.1 and 3.3.2, respectively, while section 3.3.3 covers the use of genetically modified
microorganisms. Other sugars are also present in the hydrolysis products of lignocellulosic
biomass, e.g. arabinose, mannose, galactose, but usually in lower amounts than glucose
and xylose and their fermentation is not discussed here. It is worth observing that various
studies have been carried out on the anaerobic fermentation of arabinose by mixed cultures,
however they were mainly aimed at hydrogen and not ethanol production (e.g.
70,71
).
Fermentation of arabinose to ethanol has been mainly investigated by means of genetically
modified microorganisms(e.g. 72,73).
3.3.1 Fermentation of glucose
In an anaerobic mixed microbial culture glucose can be fermented to several different end
products, as summarised in Figure 2. Ethanol can be produced directly from glucose, and
12
then be converted to acetate or organic acids, which can then be converted to methane.
Lactate, butyrate and acetate can also be produced directly from glucose through microbial
action under anaerobic conditions (Figure 2). Conversely, propionate typically derives from
the conversion of lactate; under methanogenic conditions propionate, once produced, can
be further converted into acetate and H2 (provided that methanogens keep the H2 partial
pressure below 10-5 atm).
Table 13 lists some bacterial or fungal species which are able to convert glucose to ethanol
and some species which are able to convert ethanol to organic acids. The stoichiometry of
the key reactions hereafter discussed, i.e. glucose fermentation to ethanol and ethanol
conversion to acetic acid, are reported below:
ethanol production from glucose C6 H12O6  2C2 H 5OH  2CO2
ethanol conversion to acetic acid C2 H 5OH  H 2O  CH 3COOH  2H 2
If ethanol is the desired product, the operating conditions of the fermentation should be
chosen in order to maximise the rate of the ethanol-producing reactions and to minimise the
rate of ethanol consuming reactions. The anaerobic fermentation by mixed cultures to
methane is a well known process and is widely used in industry.74 Also, relatively wide
attention has been given to the anaerobic fermentation to organic acids such as acetate,
propionate and butyrate, since they are often found as intermediates in the fermentation to
methane and can, undesirably, accumulate in the liquid medium. However, much less focus
has been given to the mixed culture fermentation to ethanol. In the next sections, the
available information on the effect of process operating conditions on the anaerobic
fermentation of glucose to ethanol is reviewed.
3.3.1.1 Effect of pH
A few studies have investigated the effect of pH on the anaerobic fermentation of glucose to
ethanol and, while it seems that pH has an important effect on ethanol production, there is
still no clear evidence on the optimum pH range to drive the fermentation process towards
ethanol, rather than acetate and methane. Based on thermodynamic considerations,
Rodriguez et al.75 predicted that acidic pH values, below 5.5, should favour ethanol
production, while at pH values higher than 6.5, acetate should be the only product in the
liquid medium. In agreement with the theory that conversion to ethanol is favoured by acid
pH values, Ren et al.76 found that in a continuous reactor in the pH range 4.3-4.9 ethanol
concentration increased at lower pH, and in this pH range ethanol and acetate were always
the main fermentation products. In the same paper, in a batch study in the pH range 3.0-5.5,
13
the authors reported the highest ethanol concentration at pH 5.0, observing a much lower
ethanol production at pH 5.5. In a continuous study in the pH range 4.0-7.0,77 the highest
ethanol concentration was found at pH 6.0, but in this case, ethanol was not the main
fermentation product, the main products being butyrate and acetate. Hwang et al.78 found
that acetate and ethanol were the main fermentation products at pH 4.5-5.0, while at pH 5.06.0 propionate and acetate were the main products.
However, other studies found higher ethanol yield at neutral or basic pH values. Temudo et
al.13 investigated anaerobic fermentation of glucose with mixed cultures in a chemostat in a
range of pH 4-8.5. They found that in the pH range 6.25-8.5 acetate and ethanol were the
main fermentation products, in approximately equal molar ratio, while in the pH range 4-5.5
very little ethanol was produced and acetate and butyrate were the main fermentation
products. However, in this study the dilution rate at pH 4-5.5 was also different than the one
at pH 6.25-8.5 and this could also have affected the results. In general agreement with their
findings, Zoetemeyer et al.79 found that acetate and ethanol were the main products of
anaerobic fermentation at pH 8.0, while at pH values below 7, the main product became
butyrate. In a chemostat study in the pH range 5-8,80 the highest ethanol concentration was
found at pH 8, but in this study ethanol was a minor fermentation product, the main ones
being acetate and propionate.
Overall, analysis of the literature indicates that further study is needed to address the effect
of pH on ethanol yield from anaerobic fermentation of glucose.
3.3.1.2 Effect of temperature
The effect of temperature on glucose fermentation to ethanol by mixed cultures is potentially
particularly interesting. In general, the rates of all microorganism-mediated processes
increase with temperature, up to the maximum temperature which is tolerable by the
microorganisms. Microorganisms used in anaerobic fermentations can be classified as either
mesophilic (optimum temperature <45 OC) or thermophilic (optimum temperature >45 OC).
An interesting advantage of thermophilic over mesophilic bacteria when using mixed cultures
for ethanol production is that among thermophilic bacteria there are many microorganisms
which are able to convert glucose to ethanol but only very few which are able to oxidise
ethanol to acetate or other organic acids.81 Therefore, it is expected that higher ethanol
yields and rates might be obtained under thermophilic than mesophilic conditions.
Other advantages of thermophilic conditions (adapted from Wiegel)81 are the following:
a) lower use of the substrate for biomass production, therefore increasing ethanol yield;
14
b) pathogens do not grow at temperature higher than 60 OC;
c) since microbial processes generate heat, higher temperatures may be easier to maintain
than lower ones;
d) ethanol can be continuously distilled from the fermentation vessel by using a moderate
vacuum
On the other hand, thermodynamic calculations82 show that at higher temperatures the
reactions that generate hydrogen become more favourable. Since glucose oxidation to
acetate or butyrate and ethanol oxidation to acetate generate hydrogen, these reactions
become more favourable at higher temperatures, potentially leading to higher ethanol loss.
The most comprehensive study on the effect of temperature on the acidogenic fermentation
of glucose has been carried out by Zoetemeyer et al.83 The authors operated a chemostat at
pH 5.8 in the temperature range 20-60 OC. At temperatures up to 50 OC, butyrate and
acetate were the main products, and the ethanol yield was quite low (0.10-0.20 mol
ethanol/mol glucose). At 55
O
C, on the other hand, ethanol was the main fermentation
product, with a yield of 0.8 mol/mol glucose.
In general, analysis of the literature shows that the effect of temperature on anaerobic
fermentation to ethanol is potentially very important and deserves further investigation.
3.3.1.3 Effect of hydrogen partial pressure
Hydrogen concentration in the liquid phase or hydrogen partial pressure in the gas phase
(the two are proportional via Henry’s law), which can be manipulated by sparging with an
inert gas or by changing the process pressure, is an important variable that can affect the
spectrum of product distribution in anaerobic fermentation. The effect of hydrogen partial
concentration is twofold: a) hydrogen levels affect the NADH/NAD ratio and therefore the
feasibility of the biochemical pathways that determine product formation;84 b) certain
fermentation reactions which generate hydrogen (Figure 2) are close to the thermodynamic
equilibrium and hydrogen concentration (as well as pH) can determine whether they are
feasible or not.
The effect of hydrogen concentration on methane formation is well known: hydrogen
concentration has to be maintained at very low values in order for the conversion of organic
acids to acetate to occur, which is thermodynamically unfeasible at high hydrogen
concentrations, and this require a close syntrophy between hydrogen producing and
hydrogen consuming microorganisms.
15
However, when methane is not the desired product, very little is known on the effect of
hydrogen concentration on the spectrum of product distribution. The biochemical model by
Rodriguez et al.84 predicts that hydrogen partial pressures above approx 0.4 atm should lead
to butyrate as main fermentation product, while lower hydrogen pressures would give
acetate. In those simulations, carried out at pH 7, no ethanol formation was predicted, since
the model only predicted ethanol formation at acidic pH values. Considering the conversion
of ethanol to acetate, this reaction is thermodynamically feasible, at pH 7, only for a
hydrogen partial pressure lower than approx 0.15 atm.85 Therefore, hydrogen pressures
higher than this value should prevent ethanol oxidation to acetate and therefore decrease
ethanol losses. The experimental study by Mizuno et al.,86 showed that a reduction in
hydrogen partial pressure from 0.5 to 0.05 atm increased the rate of hydrogen production by
more than 50%, however very little effect was observed on the composition of the liquid
effluent, the main products being acetic and butyric acids, with much lower amounts of
ethanol.
3.3.1.4 Effect of solids retention time
The solids retention time is a critical parameter for glucose fermentation with mixed cultures.
It is well known that the end-product of glucose fermentation by mixed cultures is methane, if
the digestion time or residence time is long enough.74,87,88 Therefore, glucose fermentation to
ethanol has to be carried out at relatively short residence times. However, within the region
of relatively short residence times, little systematic study has been carried out to investigate
whether the residence time affects the distribution of fermentation products. Zoetemeyer et
al.79 investigated the effect of residence time in the range 1.5-10 h (at 30 OC) and they
reported ethanol profiles for pH values of 5.69 and 6.44. They found that ethanol yield
tended to increase with longer residence times at pH 5.69 (up to 0.3 mol/mol), while it
tended to increase with shorter residence times at pH 6.44 (up to approx 0.2 mol/mol).
3.3.1.5 Rates and yields
Table 14 summarises ethanol production rates and yields in glucose fermentation studies by
mixed cultures. Only studies where the main target was ethanol or acids production are
considered here. It is evident that with mixed cultures ethanol yields of up to 0.8 mol
ethanol/mol glucose have been obtained. The maximum theoretical yield of ethanol on
glucose is 2 mol ethanol/mol glucose, assuming that all glucose is fermented to ethanol.
However, this maximum yield achievable in practice is lower than this, due to the fact that
some glucose is inevitably used for biomass growth. For the yeast Saccharomyces
cerevisiae the ethanol yield on glucose is typically in the range 1.6-1.9 mol ethanol/mol
glucose.89 The lower ethanol yield obtained with mixed cultures is due to the fact that part of
16
the glucose is fermented to other products, mainly acetate and in some cases other acids
such as propionate and butyrate. In order to develop commercial processes for ethanol
production with mixed cultures, the challenge is to determine process conditions that direct
glucose fermentation to ethanol, minimising both glucose and ethanol conversion to organic
acids. To this regard, it is important to understand which are the causes for the observed
variability in ethanol yield under similar process conditions. As an example, at a residence
time of 8 h, 30 OC, pH 6.25, Temudo et al.,13 observed a ethanol yield on glucose higher
than 0.6 mol/mol, while under similar conditions (residence time about 7 h, 30 OC, pH 6.44)
Zoetemeyer et al.79 found negligible ethanol yield. The reasons for the different behaviour
could be due to the presence or absence of nitrogen sparging, the use of different inocula,
the start-up procedure, the glucose concentration in the feed, etc.
In terms of ethanol productivity, high ethanol production rates of up to 1.5 g/L/h have been
reported with mixed cultures. This value is lower than ethanol productivity on glucose for
Saccharomyces cerevisiae, 3-18 g/L/h.89 However, considering that the literature studies
reported in Table 14 were not specifically aimed at maximising ethanol productivity, and that
ethanol productivity could be easily increased simply by increasing glucose concentration in
the feed, it seems that, with more lab- or pilot-scale investigation, ethanol productivity from
glucose with mixed cultures could reach the same or higher productivities currently obtained
with industrial processes.
3.3.2 Fermentation of xylose
Fermentation of xylose is much less known than glucose fermentation, in particular as far as
mixed cultures are concerned. In principle, the spectrum of substrates which can be
obtained by anaerobic fermentation of xylose is similar to that can be obtained from glucose
(Figure 2), even though the quantitative distribution of the products and the microbial species
involved may be different. The stoichiometry of xylose conversion to ethanol is the
following:90 C5 H10O5  1.67CH 3CH 2OH  1.67CO2
The theoretical maximum yield of ethanol from xylose is 1.67 mol ethanol/mol xylose, i.e.
virtually the same yield as glucose if expressed in mass terms (0.51 g ethanol/g xylose).
Table 15 reports several species of microorganisms which have been reported to convert
xylose into ethanol. Certain microbial species are able to produce ethanol from xylose with
almost the maximum yield, while other always generate other co-products, mainly
acetate.91,92 In terms of the considered process with mixed cultures, the operating conditions
have to be found that maximise ethanol yield, minimising the formation of other fermentation
17
by-products. However, while several recent studies have investigated the effect of operating
conditions on xylose fermentation to hydrogen,93-96 the only study which has investigated
xylose conversion to ethanol by mixed cultures is the one by Temudo et al.97 They compared
chemostat cultures grown on xylose or glucose as only carbon sources comparing ethanol
and acids production with the two substrates. They observed that the culture grown on
xylose produced much less ethanol than the one grown on glucose (0.05 mol ethanol/mol
xylose vs. 0.24 mol ethanol/mol glucose), the other main products being in both cases
acetate and butyrate. However, interestingly ethanol yield on xylose increased much when
xylose concentration in the feed increased from 4 to 10 g/l, from 0.05 to 0.69 mol
ethanol/mol xylose (the yield of butyrate was correspondingly much lower), but the reason
for this is not known. The authors also observed that the mixed culture grown solely on
xylose was immediately able to metabolise glucose when this substrate was added,
indicating that in a mixed culture with complex substrates such as real wastes, the same
microorganisms may be able to metabolise both glucose and xylose.
3.3.3 Fermentation of glucose and xylose to ethanol by genetically modified microorganisms
In general, the reason behind metabolic engineering of microorganisms in order to produce
ethanol from glucose and xylose is essentially to increase the range of substrates that can
be potentially converted to ethanol at high yield by a single microorganism. Indeed, native
strains of the yeast Saccharomyces cerevisiae are not able to utilise pentoses such as
xylose, therefore limiting the range of feedstock that can be used for ethanol production.
Other microorganisms such as Escherichia coli are on the other hand able to metabolise a
wider range of substrates but the native strains don’t produce ethanol as main fermentation
product.
The enteric microorganism Klebsiella oxytoca M5A1 converts xylose to ethanol, but also
produces organic acids (acetic, lactic, succinic). By metabolic engineering Ohta et al.98
increased the molar fraction of ethanol in the products of xylose fermentation from 62% to
90%. Similarly, using metabolic engineering on E. Coli KO11, Yomano et al.99 obtained
almost stoichiometric conversion of xylose to ethanol, with very minor production of organic
acids. Other researchers used metabolic engineering to increase the ethanol yield from
glucose in Lactobacillus sp.100 Since the most common microorganism used in industrial
bioethanol production is the yeast Saccharomyces cerevisiae, considerable effort has been
dedicated to engineering this microorganism to metabolise xylose.101 In general, good
success has been obtained, however the volumetric productivity obtained with recombinant
S. cerevisiae on xylose is still significantly lower than the one of the native strain on
glucose19. So far, the maximum ethanol productivity obtained for recombinant S. cerevisiae
18
on xylose is 0.5 g/l/h (lab scale study). Table 16 reports ethanol production rates and yields
from glucose and xylose by genetically modified microorganisms in selected literature
studies. In general, volumetric productivities of up to 2 g ethanol/l/h and almost quantitative
conversions of glucose and xylose to ethanol have been obtained, so indicating the success
of genetic engineering in generating microorganisms able to convert multiple sugars to
ethanol at high rate and yield.
19
4. Towards an integrated microbial process to convert lignocellulosic biomass to
ethanol: challenges and research opportunities
The literature reviewed in this paper shows that there are microorganisms which are able to
catalyse each of the three steps required for the conversion of lignocellulosic biomass into
ethanol: lignin hydrolysis, cellulose hydrolysis and glucose and xylose fermentation to
ethanol. Therefore, an entirely microbial process converting lignocellulosic biomass to
ethanol can, at least in principle, be considered. Compared to the alternative processes
which use high-pressure/high temperature conditions for lignin hydrolysis and external
enzymes addition for cellulose hydrolysis, an entirely microbial process at ambient pressure
and relatively low temperature would clearly give an important reduction in process costs.
An integrated (or single stage or consolidated) process which uses untreated lignocellulosic
biomass as feedstock and converts it to ethanol is clearly the most desirable option. This
process could be obtained using two alternative approaches: use of an open mixed culture,
where many different naturally-occurring microorganisms co-exist and carry out the various
steps, or use of a pure culture of a genetically modified microorganism which is able to carry
out all the required process steps. However, either approach is still far from becoming reality.
In this section the main challenges and research opportunities for the two approaches will be
discussed.
4.1. Open mixed cultures
The main challenges to be overcome for the development of a mixed culture process are the
following:
- Low rates of lignin and cellulose hydrolysis. The rates of lignin and cellulose hydrolysis
reported so far for microbial processes are lower than for chemical-physical or enzymatic
processes;
- Control of the anaerobic fermentation of sugars (mainly glucose and xylose) to ethanol. In a
mixed culture environment, fermentation of sugars can lead to many different products, in
addition to ethanol, i.e. other alcohols, volatile fatty acids (acetic, propionic, butyric, etc),
hydrogen or methane;
- Co-existence of lignin- and cellulose-hydrolysing and of ethanol-producing microorganisms
in the same vessel. For an integrated process the microbial populations responsible for lignin
and cellulose hydrolysis and the ones responsible for sugars fermentation should co-exist in
the same vessel. This might be possible or not, depending on the operating conditions of the
process and on the growth rate of the various microorganisms.
20
Some strictly interlinked research opportunities which may address the challenges above are
discussed below.
- Enrichment studies: As discussed in section 3.1.1, microbial hydrolysis of lignin is usually
considered to be difficult and slow and a factor that may limit the rate of hydrolysis is
adaptation of the microorganisms to the substrate. It is possible to hypothesise that once
mixed cultures have become adapted to a lignocellulosic substrate and have synthesised the
enzymes required for its hydrolysis, then the rate of hydrolysis should proceed faster.
Therefore a process can be envisaged, where a mixed culture is previously acclimated to the
lignocellulosic substrate (slow process) and then transferred in a continuous reactor, or
semi-continuous reactor such a Sequencing Batch Reactor, with a continuous, or semicontinuous, feed of the substrate (fast process). Having been previously acclimated, the
microbial culture should be able to remove the substrate at high rate. Investigation of this
process at lab-scale is possible but very few studies have been carried out. An example is
the study by Haruta et al.,102 where a stable microbial community able to degrade various
cellulosic and lignocellulosic substrates was generated from composting microorganisms by
acclimation on filter paper;
- Particle size reduction: reducing the feedstock particle size is expected to give higher rates
of hydrolysis, but the quantitative evidence for this effect is rather limited, especially as far as
ethanol production is concerned. Lab-scale studies specifically targeted at exploring and
quantifying the possible rate increase obtainable by particle size reduction are required;
- Reactor configuration and process parameters: the reactor used for the integrated process
can be operated under various configurations, e.g. continuous-flow with or without biomass
recycle, Sequencing Batch Reactor, etc. For each configurations, various process
parameters have to be specified, e.g. temperature, pH, hydraulic retention time, solids
retention time, length of cycle, length of the feed (the latter two only apply to Sequencing
Batch Reactors). The choice of these parameters can affect both the hydrolysis rate and the
spectrum of product distribution of sugars fermentation. E.g. it can be expected, in principle,
that in a Sequencing Batch Reactor the hydrolysis rate should be higher than in a
continuous-flow reactor due to the higher substrate concentration at the start of the cycle,
which is expected to give a higher reaction rate. However, no experimental proof of this in
the context of lignocellulosic biomass hydrolysis has been reported. Similarly, only limited
experimental investigation has been carried out regarding the effect of process parameters
on products distribution of sugars fermentation, examples are the studies by Temudo et
al.13,97 All these aspects deserve systematic investigation at lab-scale.
21
An interesting alternative to open mixed cultures is the use of selected mixed cultures, where
only selected species, responsible for different stages of the lignocellulosic biomass
conversion to ethanol, are inoculated in the reactor. A successful study using this approach
has been published very recently.103 The authors obtained 67% ethanol yield from pretreated
(dilute acid) wheat straw using a microbial culture composed of three naturally occurring
strains: Trichoderma reesei, Saccharomyces cerevisiae and Scheffersomyces stipitis. The
fungus T. reesei was responsible for cellulose and hemicellulose hydrolysis, while the yeasts
were responsible for ethanol production from glucose (S. cerevisiae) or pentoses (S. stipitis).
The authors utilised a biofilm membrane reactor with the presence in the same reactor of
aerobic, microaerophilic and anaerobic conditions, therefore allowing the co-existence of the
three different species.
4.2 Genetically modified microorganisms
Similarly to the use of open mixed cultures, the approach of using a single, genetically
modified, microorganism to convert lignocellulosic biomass to ethanol is still far from
becoming reality.
So far no attempt has been reported to introduce in microorganisms the ability to
hydrolyse/break down lignin and this is probably due to the complexity of the genome of the
native lignin degrading species. Therefore, so far the concept of metabolic engineering for
bioethanol production has been focused on the use of chemically or physically pretreated
feedstocks, where lignin has been hydrolysed and cellulose is available for microbial attack.
Considering metabolic engineering for cellulose hydrolysis, the ability to hydrolyse pretreated
cellulose has been introduced in various microbial strains, but so far very little success has
been reported with untreated crystalline cellulose. More success has been reported in the
increase of ethanol yield in microorganisms that are naturally able to hydrolyse cellulose, but
even in this case the rates are in the majority of cases very low. 104 So far, the main success
of genetic engineering for bioethanol production has been the development of
microorganisms which are able to convert multiple sugars to ethanol with high yields. While
this is an important step forward, the main issues of lignin and cellulose hydrolysis are still
far from being solved by means of genetically modified microorganisms.
Interesting research opportunities lie ahead in the following areas:
- Introduction of the lignin hydrolysis capability into microorganisms which are naturally able
to hydrolyse cellulose, or, as opposite strategy, introduction of the cellulose hydrolysis
capability into microorganisms which are naturally able to hydrolyse lignin;
22
- Improvement in the ability to hydrolyse crystalline cellulose with microorganisms which are
native ethanol producers;
- Increase in the ethanol yield for microorganisms which are naturally able to hydrolyse
cellulose.
23
5. Conclusions
This paper has reviewed the existing literature on microbial processes for lignin hydrolysis,
cellulose hydrolysis and glucose fermentation to ethanol. The main evidence from this study
is the following:
- there is a wide range of microorganisms that can perform each of the three steps required
for lignocellulosic biomass conversion into ethanol, i.e. lignin hydrolysis, cellulose hydrolysis
and glucose, or xylose, fermentation to ethanol;
- while there are many reported fungi species that are able to hydrolyse lignin under aerobic
conditions, there is only one recent study in the literature giving clear evidence of lignin
hydrolysis under anaerobic conditions. However, many mixed culture studies give an indirect
evidence that lignin can be at least partially degraded under anaerobic conditions. In
principle, if anaerobic lignin hydrolysis can be achieved, a single-stage process with mixed
microbial cultures including lignin and cellulose hydrolysis and glucose fermentation to
ethanol can be envisaged;
- cellulose and hemicelluloses hydrolysis can be carried out by many different microbial
species, both under aerobic and anaerobic conditions. Interestingly, the literature evidence
collected so far indicates no significant differences in the cellulose hydrolysis rate under
aerobic or anaerobic conditions;
- regarding anaerobic fermentation of sugars to ethanol, literature studies with mixed
cultures specifically targeted at ethanol production have been very limited and they have
reported a maximum yield of 0.8 mol ethanol/mol glucose, compared to the 2 mol
ethanol/mol glucose which is the theoretical maximum yield;
- metabolic engineering has been successful in generating microorganisms which are able to
convert a wider range of sugars to ethanol with high yields, however much more limited
success has been obtained by engineering microorganisms in order to combine cellulose
hydrolysis and high ethanol yield.
An integrated (or consolidated) process converting untreated lignocellulosic biomass to
ethanol can, at least in principle, be conceived according to two different approaches: use of
open mixed cultures of existing microorganisms or use of a pure culture of a genetically
modified microorganism. Regarding the use of open mixed cultures, the main challenges to
be overcome are: low rates of lignin and cellulose hydrolysis, control of the anaerobic
fermentation of sugars to ethanol and co-existence of different microbial populations in the
same reactor. Possible research areas which can help addressing these challenges are:
24
enrichment studies with microbial adaptation to the lignocellulosic substrate, investigation of
the effect of particle size reduction on the hydrolysis rates and investigation of the effect of
reactor configuration and operating parameters. Regarding the use of genetically modified
microorganisms the main challenges are the development of microorganisms which are able
to hydrolyse lignin and crystalline cellulose and convert the produced sugars to ethanol.
25
Table 1. Approximate composition (% of dry weight) for various lignocellulosic materials.
Minor components such as ash, proteins, etc are not included in the table
Cellulose and hemicellulose
Material
other
Lignin
glucose
xylose
arabinose
Ref
carbohydr
ates
Corn stover
21
40
22
3
1
105
Wheat straw
15
32
35-40
4-8
4-8
106, 107
Rice straw
10
41
15
5
2
45
Leaves
0
Paper
0-15
Newspaper
18-30
Switchgrass
23
32
20
4
<1
105
Poplar
29
40
15
1
2
105
Eucaliptus
28
50
11
<1
2
108
Pine
28
45
6
2
14
108
Spruce
28
45
7
1
15
109
Angiosperms
18-24
42-52
12-26
0.5-0.6
2-4
110
Conifers
27-32
43-46
5-10
0.5-2
9-14
110
95-100
85-99
0
0
106
0
60-80
26
106
106
Table 2. Evidence of anaerobic biodegradation of lignocellulosic materials by mixed cultures
Lignin content in the
Feedstock
feedstock (% of dry
Measure of degradation
weight)
Time
(days)
Ref
0.18-0.22 m3 CH4/kg
Sisal fibre
waste
8.6
VS, 30- 70% neutral
65
detergent fibres
days
49
reduction
0.16-0.25 m3 CH4/kg
Wheat straw
10
VS, 26-38% cellulose
reduction
0.24-0.36 m3 CH4/kg
Rice straw
11
VS, 34-48% cellulose
reduction
Mirabilis
leaves
0.29-0.34 m3 CH4/kg
20
reduction
0.39-0.43 m3 CH4/kg
Ipomoea
fistulosa
25
leaves
VS, 42-47% cellulose
reduction
Lignocellulosic
(woody)
33
weeks
8
33
weeks
8
33
weeks
8
33
weeks
0.13 (average) m3
2-5
CH4/kg VS
weeks
111
biomass
0.25-0.33 m3 CH4/kg VS
Wheat straw
Wheat straw
17
Corn stover
10
Wood grass
VS, 34-39% cellulose
8
27
17-36
112
days
0.30-0.33 m3 CH4/kg VS
70
(70-78% of TBMP)
days
0.36 m3 CH4/kg VS
70
(84% of TBMP)
days
0.29 m3 CH4/kg VS
70
(66% of TBMP)
days
36
36
36
Salix
0.27-0.31 m3 CH4/kg VS
eriocephala
(70-80% of cellulose
(pussy willow)
control)
Salix lucida
0.27-0.29 m3 CH4/kg VS
100
(shining willow)
(70-74% of cellulose
days
100
34
days
34
27
control)
Populus sp.
(hybrid poplar)
0.27 m3 CH4/kg VS
(70% of cellulose
control)
Platanus
0.32 m3 CH4/kg VS
occidentalis
(82% of cellulose
(sycamore)
control)
Water hyacinth
0.19-0.21 m3 CH4/kg VS
28
100
34
days
100
34
days
113
Table 3. Microorganisms reported to degrade lignin under aerobic conditions
Microbial species
Substrate
Extent of
degradation (%)
Time (days)
Ref
Bacteria
Pseudomonas
Kraft lignin
39
52
114
Poplar wood
47-57
30
115
Poplar wood
40-52
30
115
Xanthomonas spp.
Poplar wood
39-48
30
115
Mixed culture
Wood flour
80
40-60
41
Wood flour
20
40-60
41
Indulin lignin
3-4
35
116
Indulin lignin
3-4
35
116
Barley straw
29-52
21
27
Barley straw
36-48
21
48
spp.
Acinetobacter spp.
Pseudomonas
spp.
Pseudomonas
spp.
Streptomyces
badius
Streptomyces
viridosporous
Streptomyces
cyaneus
Thermomonospora
mesophila
Fungi
Pleurotus
Cotton stalks
40
30
42
Cotton stalks
60
30
42
Cotton stalks
28
14
47
Bamboo culms
24
28
43
Bamboo culms
9-24
28
43
Bamboo culms
5-19
43
Ganoderma spp.
Bamboo culms
5-16
43
Phanerochaete
Synthetic lignin
Up to 38
ostreatus
Phanerochaete
chrysosporium
Phanerochaete
chrysosporium
Echinodontium
taxodii 2538
Trametes
versicolor spp.
Trametes
ochracea spp.
29
35
40
chrysosporium
Ceriporia lacerata
Red pine
13
56
45
Stereum hirsutum
Red pine
15
56
45
Red pine
12
56
45
Polyporus
brumalis
30
Table 4. Evidence of aerobic biodegradation of lignocellulosic materials by mixed cultures
Substrate
% lignin (initial)
Ryegrass straw
12
Horse manure,
wheat straw
20
Canola residue
(Brassica
11
campestris)
Wheat leaves
Spruce
groundwood
6
degradation
7-27% lignin
degradation
12-43% lignin
degradation
17% lignin
degradation
83% lignin
degradation
Time (days)
Ref
45
117
47
118
154
119
224
120
45-69
121
135
122
45
123
50
124
90
125
9
102
CO2 evolution
23-27
10-40% of the
maximum
Sewage sludge
and green plant
Measure of
25
waste
37% lignin
degradation
Agricultural
25% lignin
organic waste
degradation
Wheat straw, root
vegetables
26% lignin
residues, bran and
degradation
soild
Olive-mill
70% lignin
wastewaters and
degradation
wheat straw
Rice straw
10
80% straw
degradation
31
Table 5. Lignin degradation rates, calculated by the authors of this paper based on literature
data.
Microorganism
Irpex lacteus
Degradation rate
Substrate
(g/L/h)
Wood chips of Pinus
Ref
0.007
126
0.014
126
0.004
126
0.020
126
Barley straw
0.0045
48
Barley straw
0.004
48
Cotton stalks
0.05
42
Cotton stalks
0.1
42
Bamboo culms
0.04
43
Bamboo culms
0.04
43
Bamboo culms
0.03
43
Cotton stalks
0.04-0.06
47
Poplar wood
0.0001-0.0002
115
strobes
Wood chips of
Irpex lacteus
Liriodendron
tulipifera
Trametes versicolor
Wood chips of Pinus
MrP 1
strobes
Trametes versicolor
MrP 1
Streptomyces
cyaneus
Thermonospora
mesophila
Pleurotus ostreatus
Phanerochaete
chrysosporium
Echinodontium
taxodii 2538
Trametes versicolor
G20
Ganoderma sp En3
Phanerochaete
chrysosporium
Acinetobacter spp.
Wood chips of
Liriodendron
tulipifera
32
Table 6. Rates of lignin degradation with non-biological pretreatments reported in the
literature 7
Pretreatment technology
Pretreatment
conditions
SO2-enhanced
Dilute acid
steam
explosion
Temperature
(OC)
Chemical
loading
Reaction time
(min)
140
1% H2SO4
180
0.05 gSO2/g
biomass
Liquid hot
Aqueous
water
ammonia
200
160
none
15% NH4OH
Lime
120
1 gCa(OH)2
+ 100 psi O2
40
10
10
60
240
4.8
22
23
11.9
1.6
Lignin
degradation
rate (g/L/h)
33
Table 7. Microorganisms reported to hydrolyse cellulose under anaerobic conditions
Extent of
Microorganism
Substrate
degradation
(%)
Ruminococcus albus
Avicel PH105
Time
(days)
Ref
30-70
0.5-2.5
127
Clostridium thermocellum
MN300
100
4
128
Ruminococcus flavefaciens
Sigmacell 20
54-87
0.3-2
55
Clostridium cellulolyticum
MN301
20-75
0.5-3
129
Clostridium thermocellum
MN300
45
4
130
Caldicellulosiruptor bescii
Switchgrass
85
15
32
~100
NR
131
SW40
80
5
132
MN300
100
5
132
Sigmacell 20
54-79
0.5-3
56
Bacteroides
succinogenes+Selenomonas
ruminantium
Ball milled
Whatman
no.1 Filter
paper
Clostridium
thermocellum+Clostridium
thermohydrosulfuricum
Clostridium
thermocellum+Clostridium
thermohydrosulfuricum
Fibrobacter Succinogenes
34
Table 8. Anaerobic degradation of cellulose by mixed cultures
Substrate
Time (days)
Ref
133
36
69
36
62% reduction in cellulose
13
133
Cellulose
68% reduction in cellulose
9 (HRT)
134
Sigmacell 50
80% reduction in cellulose
20
135
Cellulose powder
38-58% reduction in cellulose
5
136
Filter paper
BW200
Paper (hardwood and
softwood pulp)
Measure of degradation
99% (as CH4 production
compared to glucose)
100% (as CH4 production
compared to glucose)
35
Table 9. Microorganisms reported to hydrolyse cellulose under aerobic conditions
Microorganism
Substrate
Extent of
degradation (%)
Time (days)
Ref
Bacteria
Cellulomonas
uda JC3
Cellulomonas
uda JC3
Avicel
15
5
137
Solka-Floc
20
5
137
30
5
137
35
5
137
45
5
137
75
5
137
70
5
137
MN300
60
28
138
Whatman CF11
100
4
139
50-75
0.5-1.2
140
100
7
141
Cellulomonas
CC31
uda JC3
(Whatman)
Cellulomonas
Filter paper
uda JC3
(Whatman no 1)
Cellulomonas
uda JC3
MN300
Cellulomonas
Amorphous
uda JC3
cellulose
Cellulomonas
Whatman for
uda JC3
Chromatography
Cellulomonas
fermentans
Cytophaga sp.
LX-7
Fungi
Trichoderma
viride
Trichoderma
reesei
Ball milled wood
cellulose (BW
200)
Solka Floc 200
36
Table 10. Aerobic degradation of cellulose by mixed cultures
Substrate
Measure of degradation
Time (days)
Ref
Avicel
97% of theoretical CO2
47
142
Avicel
84% of theoretical CO2
45
143
Cellulose
87% of initial cellulose
3
144
Avicel
83% of initial cellulose
100
145
Avicel
95% of theoretical cellulose
70
146
Cellulose filter paper
100% of initial cellulose
72
147
Filter paper
79% weight loss
4
102
37
Table 11. Cellulose degradation rates, calculated by the authors of the present paper on the
basis of literature data
Aerobic/anaer
Substrate
Degradation
obic
rate (g/L/h)
MN300
Anaerobic
0.005
128
Rumen microorganisms
Avicel PH102
Anaerobic
0.12
58
Rumen microorganisms
Avicel PH101
Anaerobic
0.05
57
MN300
Anaerobic
0.005
138
Sigmacell 20
Anaerobic
0.05-0.20
56
Sigmacell 20
Anaerobic
0.06-0.27
55
Avicel
Anaerobic
0.03
148
MN300
Anaerobic
0.07
132
MN300
Anaerobic
0.04
130
Trichoderma reesei
Solka Floc 200
Aerobic
0.30
141
Clostridium
MN301
Anaerobic
0.03-0.05
129
BW200
Aerobic
0.12-0.33
140
Microorganism
Clostridium
thermocellum
Cellulomonas
fermentans
Fibrobacter
succinogenes
Ruminococcus
flavefaciens
Clostridium
Ref
straminisolvens
Clostridium
thermocellum+Clostridi
um
thermohydrosulfuricum
Clostridium
thermocellum
cellulolyticum
Trichoderma viride
Cellulomonas uda JC3
CC31
0.05
Avicel
0.04
Solka Floc
0.044
Filter paper
Aerobic
Whatman no. 1
38
0.055
137
MN300
0.105
Whatman for
0.25
chromatography
Ruminococcus albus
Mixed culture
Clostridium
thermocellum
Avicel PH105
Anaerobic
0.16-0.33
Avicel
Anaerobic
0.001
Filter paper
Anaerobic
0.0025
Avicel
Anaerobic
0.10
127
149
39
150
Table 12. Cellulose hydrolysis rates with enzymatic hydrolysis reported in the literature
Cellulose hydrolysis
Cellulose type
Enzyme loading
Avicel
15 FPU/g cellulose
0.23
151
Solka Floc SW40
17.6E-3 IU/mL
0.08
152
Avicel
60 FPU/g cellulose
1.3
153
rate (g/L/h)
40
Ref
Table 13. Microorganisms which are able to convert glucose to ethanol and ethanol to
acetate under anaerobic conditions (adapted from 81, 82, 89, 154-159)
Microorganism
Microorganism type
Glucose to ethanol
Clostridium thermocellum
Bacterium
Clostridium thermohydrosulfuricum
Bacterium
Thermoanaerobium brockii
Bacterium
Sarcina ventriculi
Bacterium
Thermoanaerobacter ethanolicus
Bacterium
Ruminococcus albus
Bacterium
Saccharomyces cerevisiae
Yeast
Zymomonas mobilis
Bacterium
Aspergillus spp.
Fungus
Fusarium spp.
Fungus
Penicillum spp.
Fungus
Schizosaccharomyces pombe
Yeast
Kluyveromyces marxianus
Yeast
Ethanol to organic acids
Desulfotomaculum nigrificans
Bacterium
Pelobacter acetylenicus
Bacterium
Desulfovibrio spp.
Bacterium
Desulfobulbus propionicus
Bacterium
Pelobacter propionicus
Bacterium
41
Table 14. Glucose fermentation to ethanol by mixed cultures in chemostat studies
Residence
time (h)
a
pH
Temp.
(OC)
Ethanol yield
(mol/mol
glucose)
Ethanol
rate (g/L/h)
Other
main
Ref
productsa
8
6.25-8.5
30
0.55-0.70
0.07-0.09
acetate
5-12
8
37
0.1-0.25
0.02-0.08
6
4-7
36
0.08-0.17
0.025-0.05
3-4
8
30
0.7
0.5
acetate
79
72
5
35
0.18
0.02
acetate
78
1.5-10
5.8
20-60
0.1-0.8
1.5
13
acetate,
80
propionate
acetate,
77
butyrate
acetate,
83
butyrate
In all the studies the main products in the gas phase were hydrogen and carbon dioxide
42
Table 15. Microorganisms which are able to convert xylose to ethanol under anaerobic
conditions
Microorganism
Microorganism type
Bacillus macerans
Bacterium
Clostridium thermohydrosulfuricum
Bacterium
Thermoanaerobacter ethanolicus
Bacterium
Aerobacter aerogenes
Bacterium
Fusarium oxysporum
Fungus
Aeromonas hydrophila
Bacterium
Bacillus polymixa
Bacterium
Aerobacter indologenes
Bacterium
Brettanomyces spp.
Yeast
Candida shehatae
Yeast
Pachysolen tannophilus
Yeast
Pichia stipitis
Yeast
Monilia spp.
Fungus
Mucor spp.
Fungus
Neurospora spp.
Fungus
Paecilomyces spp.
Fungus
Polyporus spp.
Fungus
Rhizopus spp.
Fungus
43
Table 16. Glucose and xylose fermentation to ethanol by genetically modified
microorganisms
Microorganism
pH
Temp.
Ethanol yield
Ethanol rate
(OC)
(mol/mol sugar)
(g/L/h)
Ref
Glucose
Erwinia sp. SR38
Klebsiella oxytoca
M5A1
6.0
30
1.9
0.7
160
6.0
30
2.0
2.1
98
37
0.8
0.18
161
37
0.9
0.02
100
Lactobacillus casei
686
Lactobacillus
plantarum
Xylose
Klebsiella oxytoca
M5A1
Klebsiella oxytoca
M5A1
Escherichia coli LY160
Saccharomyces
cerevisiae 424A
Saccharomyces
cerevisiae MA-R5
Saccharomyces
cerevisiae DA24-16
Saccharomyces
cerevisiae ADAP8
6.0
30
1.6
2.0
98
6.0
30
1.7
1.3
162
6.5
37
1.6
0.9
99
30
0.7
0.13
163
30
1.2
0.50
164
30
1.3
1.3
165
30
1.1-1.4
0.03-0.07
44
166
A
Combined reactor for lignin
hydrolysis, cellulose
hydrolysis and carbohydrates
fermentation (anaerobic)
Lignocellulosic
biomass
Ethanol to purification
B
Lignocellulosic
biomass
Lignin
hydrolysis
cellulose
Cellulose
hydrolysis
Ethanol to
carbohydrates Fermentation purification
C
Lignocellulosic
biomass
Chemical or
physical
pretreatments
cellulose
Cellulase
production
cellulase
Cellulose
hydrolysis
and
fermentation
Ethanol to
purification
D
Lignocellulosic
biomass
Chemical or
physical
pretreatments
cellulose
Combined reactor for
cellulose hydrolysis and
carbohydrates fermentation
(anaerobic)
Ethanol to
purification
Figure 1. Possible schemes for conversion of lignocellulosic biomass to ethanol. A) and B)
are the entirely microbial processes considered in this study. Scheme C) is SSCF, scheme
D) is CBP. See text for definitions of abbreviations. Not all the material flows are shown in
the schemes and “cellulose” also include hemicellulose.
45
Glucose
(24 e-eq/mol)
Butyrate
(20 e-eq/mol)
2 x Ethanol
(12 e-eq/mol)
2 x Lactate
(12 e-eq/mol)
2 x H2
(2 e-eq/mol)
2/3 x
2x
2 x Acetate
(8 e-eq/mol)
2x
4x
4 x H2
(2 e-eq/mol)
2x
Acetate
4/3 x Propionate
(14 e-eq/mol)
4/3 x
4x
4 x H2
(2 e-eq/mol)
(8 e-eq/mol)
CH4
(8 e-eq/mol)
Figure 2. Possible products of anaerobic fermentation of glucose in a mixed culture
environment with associated electron flow. Electron equivalents (e-eq) represent the moles
of electrons which would be released upon complete oxidation.
46
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