2.0. review of literature 2.1 brief history of ethanol...
TRANSCRIPT
2.0. REVIEW OF LITERATURE
2.1 Brief history of ethanol production
Ethanol is an alcohol made through the fermentation of plant sugars from agricultural crops
and biomass resources (NEVC, 1998). With rapid depletion of the world reserves of
petroleum, ethanol in recent years has emerged as one of the alternative liquid fuel and has
generated immense activities of research in the production of ethanol and its environmental
impact. Production of alcoholic beverages is in fact as old as human civilization. The
production of pure ethanol apparently begins in the 12-14th century along with improvement
of distillation. During the middle ages, alcohol was used mainly for production of medical
drugs but also for the manufacture of painting pigments. The knowledge of using starchy
materials for ethanol production was first employed in the 12th century in typical beer
countries like Ireland. Ethanol was one of the most popular lamp illuminants used in 1850s
and approximately 90 million gallons ethanol was produced in the United States. But due to
the tax imposition on ethanol to assist in financing the civil war and the cheaper price of
kerosene, it quickly replaced ethanol as the premier illuminant in 1861 (Morris, 1993). It was
only in the 19th century that this trade became an industry with enormous production figures
due to the economic improvements of the distilling process. It was at the beginning of the
20th century that it had become known that alcohol might be used as fuel for various
combustion engines, especially for automobiles. In the 1970‟s, the interest in fuel ethanol was
renewed due to the oil crisis. Nearly 25 federal agencies administered various ethanol
programs and the National Alcohol Fuels Commission was established to study the potential
for alcohol based fuels (Lansing, 1983). Ethanol gained further support in 1980 when
Chrysler, Ford and General Motors released statements that ethanol with blends of up to 10%
would be covered in their vehicle warranties (RFA, 1998). It‟s market grew from less than a
billion litres in 1975 to more than 39 billion litres in 2006 and is expected to reach 100 billion
litres in 2015 (Licht, 2006). Interest in the use of biofuels worldwide has grown strongly in
recent years due to the limited oil reserves, concerns about climate change from greenhouse
gas emissions and the desire to promote domestic rural economies.
2.2 Ethanol and its characteristics
Bioethanol or fuel alcohol refers to ethyl alcohol produced by microbial fermentation (as
opposed to petrochemically-derived alcohol) that is used as a transportation biofuel. It is
produced through distillation of the ethanolic wash emanating from fermentation of biomass-
derived sugars and can be utilized as a liquid fuel in internal combustion engines, either neat
or in petrol blends (Walker, 2011). Table 2.1 summarises some of the important
characteristics of ethanol as a fuel source.
Table 2.1 Physico-chemical characteristics of ethanol as a liquid fuel.
Parameter
Characteristic properties
Molecular formula C2H5OH
Molecular mass 46.07 g/mol
Appearance Colourless liquid
Water solubility
(between –117°C and 78°C)
∞ (miscible)
Density 0.789 kg/l
Boiling temperature 78.5°C (173°F)
Freezing point –117°C
Flash point
12.8°C
(lowest temperature of ignition)
Ignition temperature 425°C
Explosion limits Lower 3.5% (v/v) Upper 19%(v/v)
Vapour pressure @ 38°C 50 mm Hg
Higher heating value (at 20°C) 29,800 KJ/kg
Lower heating value (at 20°C) 21,090 KJ/kg
Specific heat Kcal/Kg 60°C
Acidity (pKa) 15.9
Viscosity 1.200 mPa.s (20°C)
Refractive index (nD) 1.36 (25°C)
Octane number 99 Source: (Walker, 2011)
The high octane number of ethanol makes its blend achieve the same octane boosting
or anti-knock effect as petroleum derived aromatics like benzene. Aside high octane number
ethanol has a high evaporation heat and high flammability temperature that influences the
engine performance positively and increases the compression ratio. The blend E85 consisting
of 15% unleaded gasoline and 85% ethanol has a prevalent usage as alternative fuel because
of its advantage over pure ethanol which has a high risk of cold starting problem.
2.3 Bioethanol feedstocks
There is a growing interest worldwide to find out new and cheap carbohydrate sources for
production of bioethanol (Mohanty et al., 2009). For a given production line, the comparison
of the feedstocks includes several issues (Gnansounou et al., 2005) (1) chemical composition
of the biomass (2) cultivation practices (3) availability of land and land use practices (4) use
of resources (5) energy balance (6) emission of greenhouse gases, acidifying gases and ozone
depletion gases (7) absorption of minerals to water and soil (8) injection of pesticides (9) soil
erosion (10) contribution to biodiversity and landscape value losses (11) farm-gate price of
the biomass (12) logistic cost (transport and storage of the biomass) (13) direct economic
value of the feedstocks taking into account the co-products (14) creation or maintenance of
employment and (15) water requirements and water availability. Bioethanol feedstocks can
be divided into three major groups: (1) First generation feedstocks (2) Second generation
feedstocks and (3) Third generation feedstocks.
2.3.1 First generation feedstocks
First generation bioethanol feedstocks come from agricultural cereal and sugar crops that are
also sources of human (and animal) food (Fig. 2.1). The bioethanol produced by fermentation
of sugars such as sugarcane (Macedo et al., 2008; Leite et al., 2009), sugar beet (Ogbonna et
al., 2001; Icoz et al., 2009), sorghum (Mamma et al., 1995; Prasad et al., 2007a; Yu et al.,
2008), whey (Domingues et al., 2001; Gnansounou et al., 2005; Silveira et al., 2005; Dragone
et al., 2009) and molasses (Roukas, 1996) and starchy feedstocks such as grains viz. maize
(Gaspar et al., 2007; Persson et al., 2009), wheat (Nigam, 2001), root crops such as cassava
(Amutha and Gunasekaran, 2001; Kosugi et al., 2009; Rattanachomsri et al., 2009) are
commonly known as first generation bioethanol.
Sugar crops need only a milling process for the extraction of sugars to fermentation
(not requiring any step of hydrolysis), becoming a relatively simple process of sugar
transformation into ethanol. In this process, ethanol can be fermented directly from cane juice
or beet juice or from molasses generally obtained as a by product after the extraction of sugar
(Icoz et al., 2009).
In processes that use starch from grains like corn, saccharification is necessary before
fermentation. In this step, starch is gelatinized by cooking and submitted to enzymatic
hydrolysis to form glucose monomers, which can be fermented by microorganisms (Mussatto
et al., 2010). First generation bioethanol have played an important role in establishing the
infrastructure and policy drivers, required to support renewable transport fuels in the
international market place (EIA, 2008). There are examples of various first generation crops
having various amount of ethanol production (Table 2.2). However its growth and
development is limited due to (i) competition with food and fibre production for the use of
arable land (ii) regionally constrained market structures (iii) lack of well managed
agricultural practices in emerging economies (iv) high water and fertilizer requirements and
(v) a need for conservation of bio-diversity.
Fig. 2.1 Bioethanol from first generation feedstocks.
Sugar Crops:
Beet and Cane
Starch Crops:
Cereals, Roots and
Tubers
Gelatinization and
Saccharification:
To glucose, maltose
and malto dextrins
Juice Extraction:
To sucrose
Fermentation
Medium
Supplements:
N2, P,
vitamins, etc.
Fermentation
Bioethanol
Table 2.2 Ethanol yields from first generation crops.
Crop Ethanol yield (per hectare of cultivable
land)
Sweet sorghum 4.0-6.5
Wheat
4.8
Sugar beet 3.3-3.8
Potato
2.0-2.9
Chicory
2.0-3.9
Jerusalem artichoke 4.0-4.7
Source: (Gatel and Cormack, 1986; Abbas, 2010)
2.3.2 Second generation feedstocks
Exploitation of first generation feedstocks for future bio-fuel production is ultimately
unsustainable due to food security and land-use issues. Second-generation bioethanol refers
to fuel alcohol produced from non-food biomass sources, such as lignocellulose, the most
abundant form of carbon on the earth. The various forms of lignocellulosic feedstocks can be
grouped into six main categories (Table 2.3) and bioethanol potential of different
lignocellulosic substrates is represented in Table 2.4.
Table 2.3 Lignocellulosic biomass categories.
Biomass Category Common Examples
Industrial cellulosic waste
Municipal solid waste
Agricultural residues
Dedicated herbaceous
Hardwoods
Softwoods
Saw mail and Paper mill waste, Furniture
industry discards
Newsprint and office waste paper
Wheat straw, Corn Stover, Rice hulls,
Sugarcane bagasse
Biomass Alfalfa hays, Switch grass,
Bermuda grass, Reed canary grass,
Timothy grass
Aspen, Poplar
Pine, Spruce Source: (Sun and Cheng, 2002; Lin and Tanaka, 2006; Sanchez and Cardona, 2008)
Table 2.4 Ethanol production by different microorganisms on various lignocellulosic
substrates.
Substrate Sugars in
enzymic
hydrolysate
(g/l)
Fermenting
microorganisms
Ethanol
production
(g/l) or
yield (g/g)
Reference
Corn Stover
42 P. stipitis CBS 6054 15 Agbogbo and
Wenger, 2007
L. camara 73 S. cerevisiae VS3 42.0 Pasha et al., 2007
Prosopis
juliflora
(mesquite)
37.41 S. cerevisiae 18.52 Gupta et al., 2009
Sugarcane
bagasse
30.29 C. shehatae
NCIM3501
8.67 Chandel et al.,
2007b
Wheat straw 54.96 S. cerevisiae 25.14 Han et al., 2009
S. Spontaneum 53.91±0.44 S. cerevisiae VS3 22.85±0.44 Chandel et al.,
2009b
Rice straw 60 S. cerevisiae 12.34 Sukumaran et al.,
2009
News paper 38.21 S. cerevisiae 14.77 Kuhad et al.,
2010a
Sugarcane
bagasse
42.4 S. cerevisiae MA-R4 17.8 ± 0.69 Silva et al., 2010
Food waste 164.8 S. cerevisiae H058 81.50 Yan et al., 2010
Cashew apple
bagasse
15 S. cerevisiae 5.6 Rodrigues et al.,
2011
Rice straw (RS) 62.7mg/g RS Z.mobilis 0.86 Kumar and
Puspha, 2012
Rice straw (RS) 28mg/g RS S. cerevisiae 11 Belal, 2013
They account for nearly 50% of world biomass with an estimated annual production
of 10 to 50 billion tons, making lignocellulose arguably the most abundant and renewable
organic component of the biosphere (Claassen et al., 1999). Lignocellulosic biomass in the
form of wood and agricultural residues is virtually inexhaustible, since their production is
based on the photosynthetic process which is about 60% of the total biomass produced
(Kuhad et al., 1997). It was estimated that terrestrial plants produce about 1.3×1010
metric
tons per annum which is energetically equivalent to about two-thirds of the world‟s energy
requirement (Kim and Yun, 2006). Moreover, agricultural residuals or by-products are
annually renewable, abundantly available and account for more than 180 million tons per
year (Kapdan and Kargi, 2006). These lignocellulosic biomass includes woody material
(Ballesteros et al., 2004), straws (Huang et al., 2009; Silva et al., 2010), agricultural waste
(Lin and Tanaka, 2006) and crop residues (Hahn-Hagerdal et al., 2006). Second-generation
biofuels are expected to reduce net carbon emission, increase energy efficiency and reduce
energy dependency, potentially overcoming the limitations of first-generation biofuels
(Antizar-Ladislao and Turrion-Gomez, 2008). For countries where cultivation of energy
crops for bioethanol is difficult, lignocellulosic biomass offers an attractive option (Cardona
and Sanchez, 2007). The other major benefits of switching to cellulosic ethanol are its
renewable nature, long term sustainability, low net carbon emission, high energy efficiency,
low energy dependency, increase in national security and diversifying rural economies (IEA,
2008b). Polysaccharides present in lignocellulosic materials including cellulose and
hemicellulose are of great interest as feedstocks for second generation ethanol production. A
schematic for the conversion of biomass to fuel is shown in Fig. 2.2.
Fig. 2.2 General outline of the lignocellulose to bioethanol production process
2.3.3 Third generation feedstocks
Third-generation biofuels are produce from algal biomass, which has a very distinctive
growth yield as compared with classical lignocellulosic biomass (Brennan and Owende,
2010). Microalgae have broad bioenergy potential as they can be used to produce liquid
transportation and heating fuels, such as biodiesel and bioethanol. Microalgae provide
carbohydrates (in the form of glucose, starch and other polysaccharides), proteins and lipids
for the production of biofuels. They are recognised as one of the oldest living organisms, are
Lignocellulosic biomass
Pretreatment
Hydrolysis Fermentation
Distillation/ Separation
Separation
Bioethanol
thallophytes i.e. lacking roots, stems and leaves have chlorophyll a as their primary
photosynthetic pigment and lack a sterile covering of cells around the reproductive cells
(Brennan and Owende, 2010). Algae structures are primarily for energy conversion without
any development beyond cells and their simple development allows them to adapt to
prevailing environmental conditions and prosper in the long term. Third generation biofuels
derived from microalgae are considered to be a viable alternative energy resource that is
devoid of the major drawbacks associated with first and second generation biofuels such as
(1) microalgae are capable of all year round production (2) they can be cultivated in brackish
water on non-arable land and therefore may not incur land-use change, minimising associated
environmental impacts (Searchinger et al., 2008) (3) microalgae are able to produce 15-300
times more oil for biodiesel production than traditional crops on an area basis. Furthermore
compared with conventional crop plants which are usually harvested once or twice a year,
microalgae have a very short harvesting cycle (≈1-10 days depending on the process),
allowing multiple or continuous harvests with significantly increased yields (Schenk et al.,
2008) (4) they grow in aqueous media but need less water than terrestrial crops therefore
reducing the load on freshwater sources (Dismukes et al., 2008) (5) with respect to air quality
maintenance and improvement, microalgae biomass production can effect bio fixation of
waste CO2 (1 kg of dry algal biomass utilise about 1.83 kg of CO2) (Chisti, 2007) (6)
nutrients for microalgae cultivation (especially nitrogen and phosphorus) can be obtained
from wastewater, therefore, apart from providing growth medium, there is dual potential for
treatment of organic effluent from the agri-food industry (Cantrell et al., 2008) (7) algae
cultivation does not require herbicides or pesticides application (Rodolfi et al., 2008) (8) they
can also produce valuable co-products such as proteins and residual biomass after oil
extraction which may be used as feed or fertilizer (Spolaore et al., 2006).
2.3.4 Rice straw as substrate
Rice (Oryza sativa) is a major crop grown worldwide with an annual productivity around 800
million metric tons that corresponds with large production of rice straw (Wati et al., 2007). In
terms of total production, rice is the third most important grain crop in the world behind
wheat and corn. For every ton of harvested grain, about 1.35 tons of rice straw remains in the
field which generate huge amount of straw annually (Kadam et al., 2000). It gives an
estimation of about 650-975 million tons of rice straw produced per year globally and a large
part of this is going as cattle feed and rest as waste. The disposal of rice straw is a problem
due to the huge bulk quantity, slow degradation rate and harboring of diseases. Moreover, it
cannot be used as animal feed due to its low digestibility; low protein, high lignin and silica
content (Kausar et al., 2010). The straw is removed from the field by burning which is a
common practice all over the world. The impact of open field burning of paddy straw on air
quality has led to legislation, which will help in future to check this practice and will save
plant nutrients. In the search for viable alternatives of biofuels, paddy straw has been pursued
as suitable lignocellulosic waste for ethanol production in a process involving chemical
pretreatment followed by enzymatic hydrolysis (Wati et al., 2007). Rice straw has the
potential to produce bioethanol as it is a source that does not directly influence the price of
the rice itself as a food source. It has several characteristics that make it a potential feedstock
for fuel ethanol production. It has high cellulose and hemicellulose content that can be readily
hydrolyzed into fermentable sugars. In terms of chemical composition, the straw
predominantly contains cellulose (32-47%), hemicellulose (19-27%) and lignin (5-24%)
(Maiorella, 1983; Zamora and Crispin, 1995; Garrote et al., 2002; Saha, 2003). The pentoses
are dominant in hemicellulose, in which xylose is the most important sugar (14.8-20.2%)
(Maiorella, 1983; Roberto et al., 2003). As per Karimi et al., (2006) 1 kg rice straw will
contain 390 g of cellulose. This amount is theoretically enough to produce 220 g or 283 ml of
ethanol, however considering the practically achievable best yield as 74%, it could produce
208 ml of ethanol from a cellulose content of 1 kg rice straw. Its annual production is about
731 million tons which is distributed in Africa, Asia, Europe and America. This amount of
rice straw can potentially produce 205 billion litres bioethanol per year (Balat et al., 2008). In
Asia, it is a major field-based residue that is produced in large amounts (667.59 million
metric tons). In fact, this total amount equalling 668 million metric tons could produce
theoretically 282 billion litres of ethanol if the technology is available.
2.4 Status of bioethanol production
2.4.1 Worldwide status of bioethanol production
Bioethanol production worldwide has increased considerably since the oil crisis in 1970
(Campbell and Laherrere, 1998). Its market grew from less than a billion litres in 1975 to
more than 65 billion litres in 2008 (Biofuels Platform, 2010) and is expected to reach 100
billion litres in 2015 (Licht, 2006). According to IEA (2008b) the total worldwide demand for
oil is projected to rise by 1% per year mostly due to increasing demand in energy market of
developing countries, especially India (3.9%/year) and China (3.5%/year). Global production
of bioethanol increased from 17.25 billion litres in 2000 (Balat, 2007) to over 46 billion litres
in 2007 (REN21, 2008). Bioethanol production in 2007 represented about 4% of the 1300
billion litres of gasoline consumed globally (REN21, 2008). The United States, Brazil and
several EU member states have the largest programs promoting biofuels in the world.
National biofuels policies tend to vary according to available feedstock for fuel production
and national agriculture policies. With all of the new government programs in America, Asia
and Europe in place, total global fuel bioethanol demand could grow to exceed 125 billion
litres by 2020 (Demirbas, 2007).
Bio-energy ranks second (to hydropower) in renewable U.S. primary energy
production and accounts for 3% of the U.S. primary energy production (James and Barry,
2007). The United States is the world‟s largest producer of bioethanol fuel, accounting for
nearly 47% of global bioethanol production in 2005 and 2006 (Balat and Balat, 2009). The
"Biofuels Initiative" in the US Department of Energy (US DOE, 2004), strives to make
cellulosic ethanol cost-competitive by 2012 and supposedly to correspond and account for
one third of the U.S. fuel consumption by 2030. In 2007, the U.S. president signed the Energy
Independence and Security Act of 2007 (EISA, 2007), which requires 34 billion litres of
biofuels (mainly bioethanol) in 2008 increasing steadily to 57.5 billion litres in 2012 and to
136 billion litres in 2022. Similar to Brazil, the US is also a big investor in bioethanol
research (Solomon et al., 2007) and has increased the ethanol production from 6.16 billion
litres or 1.63 billion gallons in 2000 to 39.3 billion litres or 10.4 billion gallons in 2009,
representing a 7-fold increase (Petrova and Ivanova, 2010). Currently over 95% of ethanol
production in the United States comes from corn, while the rest is made from wheat, barley,
cheese whey and beverage residues (Solomon et al., 2007). However, it is expected that about
1.53 billion litres or 405 million gallons of cellulosic ethanol will be produced by the end of
2012 (Solomon et al., 2007).
The EU has also adopted a Biomass Action Plan that sets out measures to increase the
development of biomass energy from wood, wastes and agricultural crops by creating market
based incentives and removing barriers to the development of markets. Implementation of the
plan will help the EU to cut its dependence on fossil fuels, reduce greenhouse gas emissions
and stimulate economic activity in rural areas. In 2003, the European Union adopted two
biofuels directives. These directives set targets for the share of renewable fuels in the
transport fuel market (2% by the end of 2005 and 5.75% by the end of 2010) (EC Directive,
2003). The 2005 target was not achieved but the industry is growing rapidly and it is
expected that the 2010 target will be achieved. On 23 January 2008, the European
Commission proposed a binding minimum target of 10% for the share of biofuels in transport
that envisages a 20% share of all renewable energy sources in total energy consumption by
2020 (EC, 2008). The bioethanol sectors in many EU member states have responded to policy
initiatives and have started growing rapidly. Bioethanol production increased by 71% and
consumption reached 2.44 billion litres in 2007 (Tokgoz, 2008). The potential demand for
bioethanol as a transportation fuel in the EU is estimated at about 12.6 billion litres in 2010
(Zarzyycki and Polska, 2007).
Brazil is the world's largest exporter of bioethanol and second largest producer after
the United States. With regard to bioethanol, the share of the US in the global production is
50% and Brazil provides 39% of the total global supply, while the share of OECD-Europe is
5% (Gnansounou, 2010). Since Brazil is one of the most developed nations in ethanol
production, almost all the Brazilian vehicles use either pure ethanol or the blend of gasoline
and ethanol (75:25) (Mussatto et al., 2010; RFA, 2010). The high percentage in which
ethanol is added to gasoline in Brazil is also an effort on part of the government to reduce the
imports of oil (Prasad et al., 2007). As a result of these efforts, ethanol production in Brazil
has substantially risen from 555 million litres (1975/76) to 16 billion litres (2005/06)
(Orellana and Bonalume Neto, 2006; Souza, 2006). Production has been expected to rise
from 15.4 billion litres in 2004 to 26.0 billion litres by 2010. Ethanol from sugarcane
provides 40% of automobile fuel in Brazil and approximately 20% is exported to the US, EU
and other markets (Greenergy, 2007).
There are more than 10 ethanol biofuel facilities either in operation or under
construction in Canada and 130 plants in the United States as of 2006 (Allan et al., 2006;
Parcell and Westhoff, 2006). In eastern Canada and the US, corn is used as the feedstock
while in western Canada wheat is used. Brazil produces a large amount of ethanol from
sugarcane and many vehicles in that country have been built to run directly on ethanol fuel.
In Europe, ethanol is produced in Sweden, Denmark, Germany, the United Kingdom, France,
Italy and Spain. Many Asian countries such as China, India, Japan and Indonesia are also
developing ethanol production capacity (Allan et al., 2006; Worldwatch Institute, 2006).
2.4.2 India’s status of bioethanol production
India is a country with a positive outlook towards renewable energy technologies and
committed to the use of renewable sources to supplement its energy requirements. The
country is one among the few nations to have a separate ministry for renewable energy which
address the development of biofuels along with other renewable energy sources. In the year
2003, the Planning Commission of the Government of India brought out an extensive report
on the development of biofuels (Planning Commission, 2003) and, bioethanol and biodiesel
were identified as the principal biofuels to be developed for the nation. The Ethanol Blended
Petrol Programme (EBPP) launched by the Government of India in January 2003 made it
mandatory to blend petrol with 5% of ethanol in the states of Andhra Pradesh, Goa, Gujarat,
Haryana, Karnataka, Maharashtra, Punjab, Tamil Nadu, Uttar Pradesh and Uttaranchal and in
the union territories of Daman and Diu, Dadra and Nagar Haveli and Chandigarh (except
Jammu and Kashmir, north-eastern states and Island territories). The National Biofuel Policy
released in December 2009 by the Ministry of New and Renewable Energy (MoNRE, 2009)
envisages a target of complete blending of 5% ethanol by 2011-12 and then gradually raise it
to 10% by 2016-17 and to 20% after 2017.
In India, ethanol is mainly produced from sugarcane molasses but the substrate has to
compete with the food demand and therefore cannot supply the required amount of ethanol.
Therefore, the nation needs to develop bioethanol technologies, which use biomass feedstock
that does not have food or feed value. The most appropriate bioethanol technology for the
nation would be to produce it from lignocellulosic biomass such as rice straw, rice husk,
wheat straw, sugarcane tops and bagasse, municipal waste and forest waste (Sukumaran et
al., 2009). According to Kim and Dale (2004), the total bioethanol production from plant
biomass is estimated to be 491 GL/year globally. India alone has the capacity to produce 25%
i.e.123 GL/year of the total world ethanol production, if the entire lignocellulosic residues
available are used for ethanol production. Hence, to contemplate a bioethanol production
plant, the lignocellulosic biomass assessment with geographical distribution and accurate
information on availability of biomass in different parts of the country is a pre-requisite. With
this in view during the Ninth Plan, the Ministry had sponsored 500 taluka level biomass
assessment studies in 23 states to compile data on availability of lignocellulosic biomass. As
an extension to this effort, a project for preparation of “Biomass Resource Atlas for India”
has been jointly sponsored to Indian Institute of Science (IISc), Bangalore and Regional
Remote Sensing Service Centre (RRSSC), Bangalore which aims at integration of the data on
biomass availability obtained from taluka-level studies and from other reliable sources with
information on crop distribution pattern derived from GIS-based maps provided by RRSSC.
2.5 Lignocelluloses
Lignocellulosic plant biomass is an important renewable carbon resource for the biorefinery
industry and is thus considered a sustainable and environment friendly alternative to the
current petroleum platform (Wongwilaiwalina et al., 2010). Lignocellulosic biomass such as
agricultural residues and herbaceous energy crops, consists mainly of three different types of
biopolymers i.e. cellulose, hemicellulose, lignin and pectins (Ragauskas et al., 2006; van
Maris et al., 2006) (Table 2.5).
Table 2.5 Polymer composition of lignocellulosic biomass.
Polymers
Content in
lignocellulose (%)
Major monomers
Cellulose
Hemicellulose
Lignin
Pectins (when present)
33-51
19-34
20-30
2-20
Glucose
Xylose, Glucose, Mannose,
Arabinose, Rhamnose, Galactose
Aromatic alcohols
Galacturonic acid and Rhamnose
Source: (van Maris et al., 2006)
No matter what plant it comes from, lignocellulosic biomass is composed of a complex
mixture of cellulose, hemicellulose and lignin (Fig. 2.3).
Fig. 2.3 Molecular component of plant cell wall structure (Source: Rubin, 2008).
The composition of these constituents may vary from one plant species to another
(Table 2.6). For example, hardwood has more cellulose constituent while wheat straw and
leaves have more hemicellulose constituent.
Table 2.6 Composition data of several lignocellulosic materials for bioethanol
production.
Feedstock Content (dry wt %)
Cellulose Hemicellulose Lignin Reference
Hardwoods
Eucalyptus
Globules
46.30 25.83 22.90 Garrote et al., 2007
Acacia dealbata 50.50 19.30 21.90 Mumoz et al., 2007
Poplar 44.05 15.71 20.95 Pan et al., 2006
Black locust 41.61 17.66 26.70 Hamelinck et al., 2005
Softwoods
Salix 42.50 25.00 26.00 Sassner et al., 2008
Spruce 44.00 24.60 27.50 Sassner et al., 2008
Pine 44.55 21.90 27.67 Hamelinck et al., 2005
Agro-industrial residues
Corn stover 40.00 29.60 23.00 Sassner et al., 2008
Corn cobs 34.40 40.75 18.80 Parajo et al., 2004
Rice husks 36.70 20.05 21.30 Parajo et al., 2004
Barley husks 21.40 36.62 19.20 Parajo et al., 2004
Rye straw 41.10 30.20 22.90 Gullon et al., 2010
Oat straw 39.40 27.10 17.50 Nigam et al., 2009
Rice straw 36.20 19.00 9.90 Nigam et al., 2009
Wheat straw 32.90 24.00 8.90 Nigam et al., 2009
Corn stalks 35.00 16.80 7.00 Nigam et al., 2009
Cotton stalks 58.50 14.40 21.50 Nigam et al., 2009
Soya stalks 34.50 24.80 19.80 Nigam et al., 2009
Sunflower stalks 42.10 29.70 13.40 Nigam et al., 2009
Sugarcane
bagasse
40.00 27.00 10.00 Nigam et al., 2009
Ethiopian
Mustard
32.70 21.90 18.70 Gonzalez-Garcia et al., 2010
Flax shives 47.70 17.00 26.60 Gonzalez-Garcia et al., 2010
Hemp hurds 37.40 27.60 18.00 Gonzalez-Garcia et al., 2010
Dedicated energy crops
Alfalfa stems 27.50 23.00 15.80 Gonzalez-Garcia et al., 2010
Switch grass 31.98 25.19 18.13 Hamelinck et al., 2005
Waste papers
Newspaper 61.30 9.80 12.00 Kim and Moon, 2003
2.5.1 Cellulose
Cellulose is a polysaccharide composed of linear glucan chains that are linked together by β-
1,4-glycosidic bonds with cellobiose residues as the repeating unit at different degrees of
polymerization depending on resources and packed into micro fibrils which are held together
by intramolecular hydrogen bonds as well as intermolecular van der Waals forces (Zhao et
al., 2011). The chemical formula of cellulose is (C6H10O5) n and the structure of one chain of
the polymer is presented in the Fig. 2.4.
Fig. 2.4 Structure of Cellulose molecule
It has high degree of polymerization (DP) from 100-20,000 which is water insoluble
and recalcitrant to hydrolysis into its individual glucose subunit because of tightly packed,
highly crystalline structure with straight, stable supra-molecular fibres of great tensile
strength and low accessibility in its polymer form (Demain et al., 2005). About 33% of all
plant matter is composed of cellulose. Cellulose does not melt with temperature, but its
decomposition starts at 1800C. There are several types of cellulose in wood, crystalline and
non-crystalline and accessible and non-accessible. Most wood-derived cellulose is highly
crystalline and may contain as much as 65% crystalline regions. The remaining portion has a
lower packing density and is referred to as amorphous cellulose. Accessible and non-
accessible refer to the availability of the cellulose to water, microorganisms, etc. The surfaces
of crystalline cellulose are accessible but the rest of the crystalline cellulose is non-accessible,
whereas most of the non-crystalline cellulose is accessible but part of the non-crystalline
cellulose is so covered with both hemicelluloses and lignin that it becomes non-accessible
(Rowell et al., 2005; Kuhad et al., 2011a). Concepts of accessible and non-accessible
cellulose are very important in moisture sorption, pulping, chemical modification, extractions
and interactions with microorganisms. Amorphous cellulose is degraded at a much faster rate
where as crystalline cellulose is highly resistant to microbial attack and enzymatic hydrolysis
(Zhang et al., 2006).
2.5.2 Hemicellulose
Hemicelluloses are heterogeneous group of polysaccharides with the β-(1-4) linked backbone
structure of pentose (C5) sugars, such as xylose and arabinose and hexose (C6) sugars,
including mannose, galactose and glucose as the repeating units which have the same
equatorial configuration at C1 and C4 (Fig. 2.5) (Scheller and Ulvskov, 2010). Hemicellulose
is more easily hydrolyzed than cellulose (Zaldivar et al., 2001). It was estimated that
hemicellulose account on average for about 22% of softwood, 26% of hardwood and 30% of
various agricultural residues (Zhang et al., 2007). The chemical composition and
hemicellulose content usually depends on the plant materials, growth stage and growth
conditions (Niehaus et al., 1999). Unlike cellulose, hemicelluloses are not chemically
homogenous (Saha, 2003). Hemicelluloses usually consist of more than one type of sugar
unit and called accordingly e.g., Galactoglucomanan, Arabinoglucuronoxylan,
Arabinogalactan, Glucuronoxylan, Glucomannan, etc. The hemicelluloses also contain
acetyl- and methyl-substituted groups (Rowell et al., 2005). The hemicellulose from
hardwood and agricultural residues are typically rich in xylan while on the other hand,
softwood contains more mannan and less xylan (Kuhad et al., 1997; Perez et al., 2002;
Kapoor et al., 2007; Olofsson et al., 2008; Moxley et al., 2009; Kuhad et al., 2011a). The
principle hemicelluloses in softwoods (about 20%) are Galactoglucomannans. The backbone
is linear or slightly branched chain of β-(1-4) linked D-mannopyranose and D-glucopyranose
units. In addition to Galactoglucomannans , softwoods hemicellulosic structures also contain
Arabinoglucuronoxylans (5-10%) and Arabinogalactan thus introducing the 5-carbon
monosaccharide Arabinose and Xylose (Sjostrom, 1981). The major hemicellulose in
hardwood is xylan. The backbone of xylan consists of β-(1-4) linked Xylopyranose units.
Hardwoods also contain Glucomannan with the backbone of β-(1-4) linked D-
mannopyranose and D-glucopyranose units. Unlike softwood xylan, hardwood xylan does not
contain arabinose units (Christane laine, 2005).
Hemicellulose is insoluble in water at low temperature. However, its hydrolysis starts
at a temperature lower than that of cellulose, which renders it soluble at elevated
temperatures. The presence of acid highly improves the solubility of hemicellulose in water.
Fig. 2.5 Repeating units of hemicelluloses (Source: Scheller and Ulvskov, 2010)
2.5.3 Lignin
Lignin is a highly branched polyphenolic, amorphous polymer with wide range of functional
groups consisting of phenyl propanoid monomers of coniferyl, sinapyl and p-coumaryl
alcohols (Fig. 2.6) (Vivekanand et al., 2008). Alkyl-aryl, alkyl-alkyl and aryl-aryl ether bonds
link these phenolic monomers together (Kumar et al., 2009). These three aromatic alcohols
give rise to guaiacyl units, synringyl units and p-hydrophenyl units whose proportion also
differ among hardwoods, softwoods and herbaceous biomass (Tomas pejo et al., 2008).
Dividing higher plants into two categories, hardwood (angiosperm) and softwood
(gymnosperm), it has been identified that lignin from softwood is made up of more than 90%
of coniferyl alcohol with the remaining being mainly p-coumaryl alcohol units. Contrary to
softwoods, lignin contained in hardwood is made up of varying ratios of coniferyl and sinapyl
alcohol type of units.
Phenolic hydroxyl, Benzylic hydroxyl and Carbonyl groups are attached as functional
groups to the phenyl propanoid skeleton of lignin (Dence and Lin 1992; Chang and Chang
1995). It provides structural rigidity to plant cell wall by forming firm linkages with cellulose
and hemicellulose (Bajpai and Bajpai 1992; Record et al., 2003). It is present in the cellular
wall to give structural support, impermeability and resistance against microbial attack and
oxidative stress (Sanchez, 2009). The structure of lignin makes it the most recalcitrant
substance of the three and currently there are no processing methods that make it available to
fermentation. However, as its energy content is high, it can be separated from cellulose and
hemicellulose and then be burned to produce electricity, but it can also be used to produce
other chemicals from its constituents (Chang, 2007).
As lignin is amorphous heteropolymer is also non-water soluble and optically
inactive; all this makes the degradation of lignin very tough. Lignin, just like hemicellulose,
normally starts to dissolve into water around 1800C under neutral conditions (Bobleter,
1994). The solubility of the lignin in acid, neutral or alkaline environments depends however
on the precursor (p-coumaryl alcohol, coniferyl alcohol, sinapyl alcohol or combinations of
them) of the lignin (Grabber, 2005).
Fig. 2.6 Schematic representation of building blocks of Lignin polymer: 1. p-Coumaryl
alcohol 2. Coniferyl alcohol 3. Sinapyl alcohol.
2.6 Ethanol production from lignocellulosic biomass
For a lingocellulosic ethanol process to be economically competitive with starch or sugar
based processes, all of the sugars present in the cellulose and hemicellulose have to be
available to the fermenting organism (Hahn-Hagerdal et al., 2007a). However, due to the
structure and composition of the plant cell wall as described above, this process entails a
much higher degree of complexity, leading to high ethanol production costs (Cardona and
Sanchez, 2007). The bioconversion of lignocellulosics to ethanol consists of two main
processes: hydrolysis of lignocellulosic carbohydrate to fermentable reducing sugars and
fermentation of the sugars to ethanol (Fig. 2.7). The hydrolysis is usually catalyzed by
cellulase enzymes and the fermentation is carried out by yeasts or bacteria. The presence of
lignin and hemicellulose in lignocellulosic materials make the access of cellulase enzymes
difficult, thus reducing the efficiency of the hydrolysis (Himmel et al., 2007). Pretreatment of
lignocellulosic biomass prior to hydrolysis can significantly improve the hydrolysis
efficiency by removal of lignin and hemicellulose, reduction of cellulose crystallanity and
increase of porosity (McMillan, 1994; Palmqvist and Hahn-Hagerdal, 2000a, b; Sun and
Cheng, 2002; Mosier et al., 2005; Kumar et al., 2009; Kuhad et al., 2011a).
Fig. 2.7 Schematic representation of process for bioethanol production from
lignocellulosic biomass (Source: Kuhad et al., 2011a).
2.7 Pretreatment of lignocellulosic biomass
Pretreatment of lignocellulose is required to alter its complex structure, thereby increasing its
surface area which facilitates rapid and efficient hydrolysis of the polymer to fermentable
sugars (Chen et al., 2007). The pretreatment needed to render the native lignocellulosic
materials susceptible to enzymatic hydrolysis is one of the most important stages of
bioethanol production owing to its economic cost (Eggeman and Elander, 2005). An „„ideal‟‟
pretreatment should fulfill the following conditions (Romani et al., 2010) (i) simple and
economical operation (ii) limited requirements of energy, process water and chemicals (iii)
limited corrosion (iv) ability to alter the structure of lignocellulosic materials (v) selectivity
towards polysaccharide losses (vi) high recovery of valuable hemicelluloses derived products
(vii) limited production of undesired degradation products (for example, phenolic acids,
furfural, or 5-hydroxymethylfurfural) (viii) production of substrates with high cellulose
content and susceptibility towards enzymatic hydrolysis (ix) generation of high quality lignin
or lignin-derived products (x) limited generation of wastes. Since lignocellulosic materials
have complex structures, their pretreatment is not simple (Pauly and Keegstra, 2008). There
is not a general agreement on which pretreatment can be considered as the best one (Romani
et al., 2012). Pretreatment methods can be classified as physical, chemical, physico-chemical
and biological (Galbe and Zacchi, 2007), which are summarized in Fig. 2.8.
Fig. 2.8 Different pretreatment methods for lignocellulosic biomass.
2.7.1 Physical pretreatment
Several mechanical and non-mechanical methods can be used for the physical pretreatment of
biomass. Mechanical methods involve biomass comminution by a combination of chipping,
grinding and milling to reduce biomass size and cellulose crystallanity (Kumar et al., 2009).
The energy required for mechanical pretreatment depends on the final particle size and
biomass characteristics. However, in most cases this energy consumed is higher than the
theoretical energy present in the biomass (Sun and Cheng, 2002; Kumar et al., 2009). Non-
mechanical methods such as irradiation have also been tested. Irradiation by e.g., gamma
rays, electron beam and microwaves can improve enzymatic hydrolysis of lignocelluloses
(Taherzadeh and Karimi, 2008). The cellulose component of the lignocellulose materials can
be degraded by irradiation to fragile fibres and low molecular weight oligosaccharides and
even cellobiose (Kumakura and Kaetsu, 1983). This method is, however, far too expensive to
be used in a full-scale process and doubts remain about its feasibility (Galbe and Zacchi,
•Ozonolysis
•Acid hydrolysis
•Alkaline Hydrolysis
•Oxidative delignification
•Organosolv Process
•Fungi
•Bacteria
•Steam explosion
•Ammonia fibre explosion
•CO2 explosion
•Liquid hot water
•Milling
•Pyrolysis
Physical Pretreatment
Physicochemical
Pretreatment
Chemical
Pretreatment
Biological
Pretreatment
2007). Pyrolysis has also been evaluated as a physical pretreatment method. When biomass is
treated at temperatures above 3000C, cellulose rapidly decomposes to gaseous products and
residual char. At lower temperatures, the decomposition is much slower and less volatile
products are formed. The high temperatures used and the cooling costs of the system,
however, render pyrolysis is an extremely expensive method (Bridgwater et al., 1999).
2.7.2 Chemical pretreatment
Chemical pretreatment involves the use of different chemical agents such as ozone, acids,
alkalis, hydrogen peroxide and organic solvents to release lignin and degrade the
hemicellulose (Sanchez and Cardona, 2008).
2.7.2.1 Ozonolysis
The most significant effect of treating lignocellulosic biomass with ozone is on the
degradation of lignin. Ozone pretreatment effectively decreases the amount of lignin and thus
increases the in vitro digestibility of the biomass (Kumar et al., 2009). Hemicellulose is
partially degraded while the cellulose is hardly affected (Silverstein et al., 2007). Ozonolysis
pretreatment has the following advantages: (1) it effectively removes lignin (2) it does not
produce toxic residues for the downstream processes and (3) the reactions are carried out at
room temperature and pressure (Garcia-Cubero et al., 2009). The efficiency of ozone
treatment can also be affected by insufficient reaction time, low ozone concentration and
uneven ozone distribution throughout the lignocellulosic material (Silverstein et al., 2007).
2.7.2.2 Acid pretreatment
Pretreatment with acid hydrolysis can result in improvement of enzymatic hydrolysis of
lignocellulosic biomasses to release fermentable sugars (Kumar et al., 2009). There are two
types of acid hydrolysis process commonly used dilute and concentrated acid hydrolysis. The
dilute acid process is conducted under high temperature and pressure and has reaction time in
the range of seconds or minutes. The concentrated acid process uses relatively mild
temperatures, but at high concentration of acid and a minimum pressure involved (Chandel et
al., 2007). Mineral acids such as H2SO4 and HCl have been used to pretreat the
lignocellulosic materials. Although concentrated mineral acids (Hydrochloric acid (HCl),
Sulphuric acid (H2SO4) and Nitric acid (HNO3)) are powerful agents for cellulose hydrolysis
but they are toxic, corrosive and hazardous and require reactors that are resistant to corrosion.
Moreover, the recovery of concentrated acid is problematic enough to make the process
economically feasible (Sivers and Zacchi, 1995; Torget et al., 2000). Whereas, dilute acid
hydrolysis has been successfully developed for pretreatment of lignocellulosic materials. The
dilute sulphuric acid pretreatment can achieve high reaction rates and significantly improves
cellulose hydrolysis (Esteghlalian et al., 1997; Sun and Cheng, 2002; Cara et al., 2008; Rocha
et al., 2009; Gupta et al., 2011a). The use of dilute acid has been successfully developed for
the pretreatment of lignocellulose (Sun and Cheng, 2002). Recently the focus of dilute acid
hydrolysis processes remained on using less severe conditions and achieves high yields of
xylan to xylose conversion. This is necessary to achieve favorable overall process economics
because of xylan which accounts up to one third of the total carbohydrate in many
lignocellulosic materials (Gupta et al., 2009; Kuhad et al., 2010a). Removal of hemicellulose
enhances cellulose digestibility in the residual solids and glucose yields of up to 100% can be
obtained when the hemicellulose is completely hydrolyzed (Kumar et al., 2009). There are
primarily two types of dilute acid pretreatment processes: (i) a high temperature (>1600C)
continuous-flow process used for low solids loadings (i.e. weight of solids/weight of reaction
mixture equals 5-10%) and (ii) a low temperature (<1600C) batch process used for high solids
loadings of about 10-40% (Esteghlalian et al., 1997; Taherzadeh and Karimi, 2008).
Although dilute acid pretreatment can significantly enhance cellulose hydrolysis, it has been
shown that the hydrolysate may be difficult to ferment because of the presence of toxic
substances (Galbe and Zacchi, 2007). Furthermore, the combined costs of building non-
corrosive reactors, using high pressures, neutralizing and conditioning the hydrolysate prior
to hydrolysis and fermentation all contribute to make dilute acid pretreatment a more
expensive process than, for example, steam explosion or the AFEX method (Kumar et al.,
2009).
2.7.2.3 Alkaline pretreatment
This form of pretreatment utilises alkaline solutions such as NaOH, KOH, NH4OH or
Ca(OH)2 (Taherzadeh and Karimi, 2008). Sodium hydroxide is the most commonly studied
pretreatment alkali and is seen as an alternative to sulphuric acid (Silverstein et al., 2007;
Kumar et al., 2009). Compared to acid processes, alkaline pretreatment causes less sugar
degradation and much of the caustic salts can be recovered or regenerated. Alkaline
pretreatment also requires lower temperatures and pressures than other pretreatment
technologies. However, it is much slower and the pretreatment times are in the order of hours
and sometimes days rather than minutes and seconds (Kumar et al., 2009). The mechanism of
alkali pretreatment is thought to be saponification of intermolecular ester bonds cross linking
xylan, lignin and other hemicelluloses (Silverstein et al., 2007). Vaccarino et al., (1987)
studied the effects of SO2, Na2CO3, and NaOH pretreatments on the enzymatic digestibility
of grape marc and the greatest degrading effects were obtained by pretreatment with 1%
NaOH solution at 1200C. Silverstein et al., (2007) studied the effectiveness of sulphuric acid,
sodium hydroxide, hydrogen peroxide and ozone pretreatments for enzymatic conversion of
cotton stalks. Dilute NaOH treatment causes the biomass to swell, leading to an increase in
internal surface area, a decrease in cellulose crystallinity and degree of polymerization, as
well as a separation of structural linkages between lignin and carbohydrates (Sun and Cheng,
2002). Compared with acid or oxidative reagents, alkali treatment appears to be the most
effective method in breaking the ester bonds between lignin, hemicellulose and cellulose and
avoiding fragmentation of the hemicellulose polymers (Gaspar et al., 2007). Alkaline
pretreatment is however, less effective for softwoods when the lignin content is above 26%
(Yamashita et al., 2010). Recently, Hu and coworkers, (2008) used microwave and radio
frequency based dielectric heating in the alkali pretreatment of switch grass to enhance its
enzymatic digestibility. In this strategy, switch grass could be treated on a large scale at high
solid loading with uniform temperature distribution (Hu et al., 2008; Hu and Wen, 2008).
2.7.2.4 Oxidative delignification
The oxidative delignification process involves the addition of an oxidizing compound such as
H2O2 (hydrogen peroxide) or peracetic acid to the biomass in a water suspension (Hendriks
and Zeeman, 2009). Lignin degradation is catalyzed by the peroxidase in the presence of
H2O2. Azzam, (1989) reported a significant increase in the susceptibility of sugarcane
bagasse to enzyme hydrolysis after pretreatment with hydrogen peroxide. About 50% of the
lignin and most of the hemicellulose was solubilized when treated with 2% H2O2 at 300C for
8 h (Azzam, 1989). This helped to achieve a 95% glucose recovery from cellulose in the
subsequent hydrolysis with cellulase. A total sugars yield of 604 mg/g, corresponding to 94%
of theoretical, was obtained after alkaline peroxide pretreatment and enzymatic
saccharification of barley straw (Saha and Cotta, 2010). Besides, alkaline peroxide (Bjerre et
al., 1996; Lissens et al., 2004; Martin et al., 2008), the chlorite oxidation and wet oxidation
are also used as promising oxidative delignifying pretreatments. Bjerre et al., (1996) used
wet oxidation and alkaline hydrolysis of wheat straw (20g straw/l, 1700C, 5-10 min) and
achieved 85% conversion yield of cellulose to glucose. Whereas, the sodium chlorite
treatment yielded approximately 90% delignification in woody material (Prosopis juliflora;
Lantana camara) (Gupta et al., 2009; Kuhad et al., 2010b). Inhibitors such as furfural and
hydroxy-methylfurfural were not observed following oxidative delignification treatment
(Kumar et al., 2009). However, hydrogen peroxide decomposes in the presence of water at
high temperatures and this may lead to a decreased solubilization of lignin and xylan
(Silverstein et al., 2007).
2.7.2.5 Organosolv process
The organosolv process (organosolvation) is a promising pretreatment strategy that employs
an organic or aqueous organic solvent mixture with inorganic solvent catalysts such as HCl or
H2SO4 to break the internal lignin and hemicellulose bonds (Zhao et al., 2009). Methanol,
ethanol, acetone, ethylene glycol and tetrahydrofurfuryl alcohol (THFA) are common organic
solvents that can be used in the process (Chum et al., 1988; Thring et al., 1990). Organic
acids such as oxalic, acetylsalicylic and salicylic acids can also be used as catalysts in the
organosolvation process (Sarkanen et al., 1980). Cellulose is partially hydrolyzed into smaller
fragments that remain insoluble in the liquor, hemicellulose is hydrolyzed mostly into soluble
components such as oligosaccharides, monosaccharides and acetic acid, while lignin is
hydrolyzed primarily into lower molecular weight fragments that dissolve in the aqueous
ethanol liquor (Kumar et al., 2009). After pretreatment, the solvents used need to be drained
from the reactor, evaporated, condensed and recycled to reduce operating costs. Moreover,
removal of solvents from the system is necessary to prevent them inhibiting enzyme
hydrolysis, growth of microorganisms as well as fermentation (Itoh et al., 2003; Xu et al.,
2003; Zhao et al., 2009; Kuhad et al., 2011a) .
2.7.3 Physico-chemical pretreatment
Pretreatments that combine both chemical and physical processes are referred to as
physicochemical processes (Chandra et al., 2007).
2.7.3.1 Steam explosion (Autohydrolysis)
Steam pretreatment is one of the most widely used methods to pretreat lignocellulose. This
method was formerly known as „steam explosion‟ because it was believed that an explosive
action on the fibres was necessary to render the material amenable to hydrolysis. However, it
is more likely that the hemicellulose is hydrolyzed by the acetic acid and other acids released
during the steam pretreatment (Mosier et al., 2005). The process causes hemicellulose
degradation and lignin transformation due to high temperature, thus increasing the potential
of cellulose hydrolysis (Lee et al., 2009, Boluda-Aguilar et al., 2010). High pressure and
consequently high temperature, typically between 160 and 2600C for a few seconds (e.g., 30
s) to several minutes (e.g., 20 min) were used in steam explosion (Varga et al., 2004; Kurabi
et al., 2005; Ruiz et al., 2006). The factors that affect steam-explosion pretreatment are
residence time, temperature, chip size and moisture content (Duff and Murray, 1996; Wright,
1998). The advantages of steam explosion pretreatment include the low energy requirement
compared to mechanical comminution and no recycling or environmental costs are associated
(Sun and Cheng, 2002; Kumar et al., 2009). Limitations of steam explosion include
destruction of a portion of the xylan fraction, incomplete disruption of the lignin-
carbohydrate matrix and generation of compounds that may be inhibitory to microorganisms
used in fermentation processes (Mackie et al., 1985; Palmqvist and Hahn-Hagerdal, 2000a, b;
Chandel et al., 2007a, Jurado et al., 2009).
2.7.3.2 Ammonia Fibre Explosion (AFEX)
AFEX is another type of physico-chemical pretreatment in which lignocellulosic materials
are exposed to liquid ammonia at high temperature and pressure for certain time and then the
pressure is suddenly decreased (Teymouri et al., 2005; Lee et al., 2010). This pretreatment
method is similar to the steam pretreatment process, operates at high pressures (Balan et al.,
2009). The effective parameters in the AFEX process are ammonia loading, temperature,
water loading, blow down pressure, time and number of treatments (Holtzapple et al., 1991).
The AFEX process produces only a pretreated solid material, while some other pretreatments
such as steam explosion produce slurry that can be separated in solid and liquid fractions
(Mosier et al., 2005). In ammonia fibre/freeze expansion (AFEX) process, a 5-15% ammonia
solution flows through a column reactor that is packed with biomass at 1ml/cm2 for 14 min at
temperatures between 160 and 180°C (Mosier et al., 2005). However, the AFEX process was
not very effective for the plant material with high lignin content such as Lantana camara (28-
35% lignin) and aspen chips (25% lignin). Hydrolysis yield of AFEX-pretreated newspaper
and aspen chips was reported as only 40% and below 50% respectively (McMillan, 1994).
One of the major advantages of AFEX pretreatment is no formation of some types of
inhibitory by-products which are produced during the other pretreatment methods such as
furans in dilute-acid and steam explosion pretreatment. However, part of phenolic fragments
of lignin and other cell wall extractives may remain on the cellulosic surface. Therefore,
washing with water might be necessary to remove part of these inhibitory components,
although increasing the amount of wastewater from the process (Chundawat et al., 2007).
However, there are some disadvantages in using the AFEX process compared to some other
processes. AFEX is more effective on the biomass that contains less lignin and the AFEX
pretreatment does not significantly solubilize hemicellulose compared to other pretreatment
processes such as dilute-acid pretreatment. Furthermore, ammonia must be recycled after the
pretreatment to reduce the cost and protect the environment (Wyman 1996; Eggeman and
Elander, 2005).
2.7.3.3 CO2 explosion
Similar to steam and ammonia explosion pretreatment, the CO2 explosion is also used for
pretreatment of lignocellulosic materials (Schacht et al., 2008). It was hypothesized that,
because CO2 forms carbonic acid when dissolved in water, the acid increases the hydrolysis
rate. Carbon dioxide molecules are comparable in size to water and ammonia and should be
able to penetrate small pores accessible to water and ammonia molecules. Carbon dioxide
was suggested to be helpful in hydrolyzing hemicellulose as well as cellulose (Kumar et al.,
2009). Supercritical carbon dioxide has been considered as an extraction solvent for non-
extractive purposes, due to several advantages such as availability at relatively low cost, non-
toxicity, non-flammability, easy recovery after extraction and environmental acceptability
(Zheng and Tsao, 1996). Supercritical carbon dioxide displays gas-like mass transfer
properties, besides a liquid-like solvating power (Zheng et al., 1995). It was shown that in the
presence of water, supercritical CO2 can efficiently improve the enzymatic digestibility of
aspen (hardwood) and southern yellow pine (softwood) (Kim and Hong, 2001). The
delignification with carbon dioxide at high pressures can be improved by co-solvents such as
ethanol-water or acetic acid-water and can efficiently increase the lignin removal (Pasquini et
al., 2005). The yields from CO2 explosion of lignocellulosics were relatively low compared to
steam or ammonia explosion pretreatment (Zheng et al., 1998; Kim and Hong, 2001; Mosier
et al., 2005; Kumar et al., 2009). Zheng and coworkers, (1998) compared CO2 explosion
with steam and ammonia explosion and found that CO2 explosion was more cost effective
than ammonia explosion and did not cause the formation of inhibitory compounds.
2.7.3.4 Liquid hot water pretreatment
Liquid hot water pretreatment is very similar to steam explosion, the major difference being
the explosive decompression of steam explosion pretreatment is replaced by controlled
cooling to keep the water in the liquid phase throughout the process (Weil et al., 1994). This
process has been shown to remove up to 80% of the hemicellulose and to enhance the
enzymatic digestibility of pretreated material in plant residue feedstocks, such as sugarcane
bagasse (Laser et al., 2002), corn stover (Mosier et al., 2005) and wheat straw (Perez et al.,
2008). Pressured reactors are used to keep the water in the liquid state at high reaction
temperatures, termed as “uncatalyzed solvolysis” by Mok and Antal, (1992). Various biomass
samples have been pretreated with compressed liquid water. The liquid hot water
pretreatment is attractive which eliminates the use of expensive chemicals/catalysts to
facilitate the hemicellulose de-polymerization; subsequently, there is no need for
neutralization or chemical recovery after the pretreatment. The resulting pretreated materials
are reported to be highly amicable to the enzymatic saccharification step (Taherzadeh and
Karimi, 2008).These various physical and chemical methods are summarized in Table 2.7.
Table 2.7 Summary of various processes used for the pretreatment of lignocellulosic
biomass.
Pretreatment
process
Description Advantages Issues Examples of
pretreated
materials
Mechanical
Comminution
Chipping,
grinding,
milling
Reduces
cellulose
crystallinity
Power
consumption
usually higher
than inherent
biomass energy
Wood and
forestry
wastes
(hardwood,
straw)
Steam
explosion
Saturated
steam at 160-
290 0C,
p = 0.69-4.85
Causes
hemicellulose
degradation and
lignin
Low xylose
recovery;
generation of
inhibitors of
Bagasse,
corn stalk,
wheat straw,
rice straw,
MPa for
several sec or
min, then
decompression
until atm.
pressure
transformation;
cost effective,
broadly
applicable to
different
feedstocks
downstream
processes,
washing required;
lignin binding by
cellulases slows
enzymatic
hydrolysis
barley straw,
sweet
sorghum
bagasse,
poplar, aspen
Ammonia fibre
explosion
(AFEX)
Anhydrous
ammonia -
NH3/biomass
1:1 at 70-900C/
15-20 atm
pressure,
followed by
rapid
decompression
Increases
accessible
surface area,
removes
lignin and
hemicellulose to
an extent;
does not
produce
inhibitors for
downstream
processes
Safety hazards of
dealing with
ammonia; need
for hemicellulases
to complete
conversion to C5
sugars; mixed
C5/C6 sugar
hydrolysate; not
efficient for
biomass with high
lignin content
Aspen wood
chips,
bagasse,
wheat straw,
barley straw,
rice hulls,
corn stover,
bermuda
grass, alfalfa
Liquid hot
water
(LHW)
Pressurized hot
water, p>5
MPa,
T= 70-2300C,
1-46 min;
solids load
<20%
Hydrolyse
cellulose and
lignin; remove
all cellulose
Monomeric
sugars that may
further
decompose to
furfural
Bagasse,
corn stover,
olive pulp,
alfalfa fibre
Acid
hydrolysis
Dilute (0.5-
3%) H2SO4,
HCl or HNO3,
at 130-2000C
/3-15 atm
pressure.
Conc. (10-
30%) H2SO4,
170-1900C
Hydrolyzes
hemicellulose to
xylose and
other sugars;
alters lignin
structure,
broadly
applicable to
different
feedstocks,
extensively
researched
High cost;
equipment
corrosion;
formation of toxic
substances, loss
of sugars; lignin
binding by
cellulases slows
hydrolysis
Bagasse,
corn stover,
wheat straw,
rye straw,
rice hulls,
switch grass,
bermuda
grass
Alkaline
hydrolysis
Dilute NaOH,
24 h, 600C;
Ca(OH)2,
4 h, 1200C;
0.05-0.15g/g
biomass
Removes
hemicellulose
and lignin;
increases
accessible
surface area
Long residence
times required;
irrecoverable
salts formed and
incorporated into
biomass
Hardwood,
bagasse, corn
stover,
straws with
low lignin
content (10-
18%), cane
leaves
Organosolv
process
Organic
solvents
(methanol,
ethanol,
Hydrolyzes
lignin and
hemicellulose
Solvents need to
be drained from
the reactor,
evaporated,
Poplar wood,
mixed
softwood
(spruce, pine,
acetone,
ethylene
glycol,
triethylene
glycol) or their
mixture with
1% of H2SO4
or HCl
condensed and
recycled; high
cost
douglas fir)
Source: (Sanchez and Cardona, 2008; Kumar et al., 2009; Sainz, 2009)
2.7.4 Biological pretreatment
Microorganisms can also be used to treat the lignocelluloses and enhance enzymatic
hydrolysis. The advantages of biological pretreatment of plant material over chemical and
mechanical pretreatment methods include (i) mild reaction conditions (ii) avoids the use of
toxic and corrosive chemicals (iii) higher product yield (iv) fewer side reactions (v) less
energy demand and (vi) less reactor resistance to pressure and corrosion (Lee, 1997; Kuhar et
al., 2008; Sanchez, 2009). In biological pretreatment process, microorganisms such as brown-
, white- and soft-rot fungi are used to degrade lignin and hemicellulose in waste materials
(Table 2.8). Biological treatment using various types of rot fungi, a safe and environmentally
friendly method, is increasingly being advocated as a process that does not require high
energy for lignin removal from a lignocellulosic biomass despite extensive lignin degradation
(Okano et al., 2005). Brown rots mainly attack cellulose, whereas white and soft rots attack
both cellulose and lignin. Lignin degradation by white-rot fungi occurs through the action of
lignin-degrading enzymes such as peroxidases and laccase (Lee et al., 2007). These enzymes
are regulated by carbon and nitrogen sources. White-rot fungi (WRF) are the most effective
for biological pretreatment of lignocellulosic materials. Some WRF have been reported to
degrade lignin selectively and this capability of selected WRF can be exploited for
delignification of plant materials without affecting much of cellulose (Kuhar et al., 2008;
Gupta et al., 2011b). Few studies have been reported on the pretreatment of plant biomass
with WRF for its affect on cellulose hydrolysis. According to Hatakka, (1983) 35% of the
wheat straw is convertible to reducing sugars when pretreated with Pleurotus ostreatus for 5
weeks. Taniguchi and co-workers, (2005) also observed a similar conversion rate in rice
straw pretreated with P. ostreatus for 60 days. Keller and co-workers, (2003) observed a 3 to
5 fold improvement in the enzymatic cellulose digestibility in corn stover pretreated with
Coriolus versicolor in more than 30 days. Thus, most of these fungal pretreatments have
suffered because of long incubation periods. Therefore, to economize microbial pretreatment
of lignocellulosics to improve the hydrolysis of carbohydrates to reducing sugars and to
eventually improve ethanol yield, there is a need to test more and more basidiomycetous
fungi for their ability to delignify the plant material quickly and efficiently (Kuhad et al.,
2011a).
Biological pretreatment in combination with other pretreatment technologies has also
been studied (Itoh et al., 2003, Balan et al., 2008). Itoh and colleagues, (2003) reported
production of ethanol by simultaneous saccharification and fermentation (SiSF) from beech
wood chips after bio-organosolvation pretreatments by ethanolysis and white-rot fungi,
Ceriporiopsis subvermispora, Dichomitus squalens, P. ostreatus and C. Versicolor. The yield
of ethanol obtained after pretreatment with C. subvermispora for 8 weeks was 0.294 g/g of
ethanolysis pulp and 0.176 g/g of beech wood chips. The yield was 1.6 times higher than that
obtained without the fungal treatments. The combined process enabled the separation of
lignin, cellulose and hemicellulose using only water, ethanol and wood-rot fungi. The
biological pretreatments saved 15% of the electricity needed for ethanolysis. In another
interesting approach, Balan et al., (2008) studied the effect of fungal treatment of rice straw
followed by AFEX pretreatment and enzymatic hydrolysis. They reported that treating rice
straw with white-rot fungus, followed by AFEX gave significantly higher glucan and xylan
conversions.
Table 2.8 Biological pretreatment of lignocellulosic substrates for enhanced
delignification and enzymatic digestibility.
S.No. Organism Substrate Reference
1. Phanerochaete
chrysosporium
Polymeric dyes Glenn and Gold, 1983
2. Merulius tremellosus Aspen wood Reid, 1985
3. Phanerochaete
chrysosporium,
Bjerkandera adusta,
Pleurotus ostreatus,
Phlebia tremellosus,
Trametes versicolor
Barley straw, wood pulp Bradley et al., 1989
4. Fusarium proliferatum Industrial lignins
(Polymeric kraft lignin,
Polymeric organosolv
lignin), Natural lignin
(Milled wood lignin)
Regalado et al., 1997
5. Pleurotus spp.,
Lentinus edodes
Milled tree leaves, Banana
peel, Apple peel,
Mandarin peel
Songulashvili et al., 2005
6. Aspergillus terreus,
Cellulomonas uda,
Trichoderma reesei,
Zymomonas mobilis,
Aspergillus awamori,
Cellulomonas cartae,
Bacillus macerans,
Trichoderma viride
Sugarcane trash Singh et al., 2008
7. Fungal isolate RCK-1 Wheat straw Kuhar et al., 2008
8. Phanerochaete
chrysosporium
Cotton stalks Jian et al., 2008
9. Echinodontium taxodii
2538 and Trametes
versicolor G20
Bamboo culms Zhang et al., 2007
10. Coriolus versicolor B1 Bamboo residues Zhang et al., 2007
11. Phanerochaete
chrysosporium
Oil palm empty fruit
bunch
Hamisan et al., 2009
12. Irpex lacteus Cornstalks Yu et al., 2010
13. Ceriporiopsis
subvermispora
Corn stover Wan and Li, 2010
14. Phanerochaete
chrysosporium
Rice straw Zeng et al., 2011
15. Ceriporiopsis
subvermispora,
Trametes versicolor
Rubber wood Nazarpour et al., 2013
2.8 Inhibitory compounds in lignocellulosic hydrolysate and their detoxification
During the pretreatment of lignocellulose, especially with dilute acid, numerous degradation
products are generated, many of which inhibit microbial growth and metabolism. The
inhibitors formed during pretreatment can be assigned into three main groups based on
origin: furan derivatives, weak acids and phenolic compounds (Palmqvist and Hahn-
Hagerdal, 2000a; Liu, 2006) (Fig. 2.9).
Furfural 5-hydroxymethyl furfural Acetic acid
(HMF) (Furfuryl alcohol)
Phenols Levulinic acid Formic acid
Fig. 2.9 Major types of inhibitors present in lignocellulosic hydrolysate (Source:
Mussatto and Roberto, 2004).
Aromatic compounds that occur from sugar degradation are predominantly furan
derivatives, the most prominent of which are furfural from pentoses and Hydroxymethyl
furfural (HMF) from hexoses. Furans are formed in high concentrations during severe acid
pretreatment conditions (Taherzadeh et al., 1997; Klinke et al., 2004) and are considered to
be the most potent inhibitors of yeast growth and fermentation (Olsson and Hahn- Hagerdal,
1996; Taherzadeh et al., 2000). Acetic acid is ubiquitous in hemicellulose hydrolyzates
where the hemicellulose and to some extent the lignin is acetylated. Hydrocarboxylic acids
such as glycolic acid and lactic acid are common degradation products of alkaline
pretreatment. Formic acid is produced from sugar degradation, whereas levulinic acid is
formed by 5-HMF degradation (Palmqvist and Hahn-Hagerdal, 2000b; Klinke et al., 2004).
Phenolic compounds are formed from lignin during dilute-acid hydrolysis (Clark and
Mackie, 1984). Phenolic compounds can also be formed from sugars (Popoff and Theander,
1976). In addition, some of the wood extractives are phenolic compounds (Sjostrom, 1993;
Rowell et al., 2005). Some of the phenolics are strongly inhibitory even at relatively low
concentrations while much higher concentrations are required for other phenolics to obtain an
inhibitory effect (Ando et al., 1986; Larsson et al., 2000). The most common phenolic
compounds found in lignocellulosic hydrolysates include 4-hydroxybenzaldehyde, 4-
hydroxybenzoic acid, vanillin, dihydroconiferyl alcohol, coniferyl aldehyde, syringaldehyde
and syringic acid (Klinke et al., 2004).
These compounds depending on their concentration in the hydrolysate can inhibit
microbial cell and affect the specific growth rate and cell mass yield per ATP. Furfurals and
hydroxymethyl furfurals (furans) are known to inhibit the glycolytic enzymes and the direct
inhibition of alcohol de-hydrogenase (ADH) contributes to the acetaldehyde excretion which
resulted in the prolonged lag phase in the microbes (Palmqvist and Hahn-Hagerdal, 2000a, b).
The phenolics cause partition in the biological membrane and loss of integrity thereby affect
the ability to serve as selective barrier and enzyme matrix. In general, degradation products
reduce enzymatic and biological activities, break down DNA, inhibit protein and RNA
synthesis and reduce ethanol yield (Modig et al., 2002). Therefore, to facilitate fermentation
processes, detoxification procedures are often required to remove inhibitory compounds from
the hydrolysate. However, these additional steps increase the costs and complexity of the
process and generate extra waste products (Liu, 2006).
Various detoxification methods including biological, physical and chemical ones have
been proposed to transform inhibitors into inactive compounds or to reduce their
concentration. However, the effectiveness of detoxification method depends both on the type
of hemicellulosic hydrolysate and on the species of microorganisms employed.
These include the addition of activated charcoal, extraction with organic solvents, ion
exchange, ion exclusion, molecular sieves, over liming, intracellular acidification, yeast strain
variation and recombinant strains (Olsson and Hahn-Hagerdal, 1996; Rao et al., 2006). Over-
liming with a combination of high pH and temperature has for a long time been considered as
a promising detoxification method for dilute sulphuric acid-pretreated hydrolysate of
lignocellulosic biomass (Chandel et al., 2007a). This process has been demonstrated to help
with the removal of volatile inhibitory compounds such as furfural and hydroxymethyl
furfural (HMF) from the hydrolysate additionally causing a sugar loss (~10%) by adsorption
(Martinez et al., 2000; Ranatunga et al., 2000; Chandel et al., 2011a, b). The detoxification of
hemicellulose hydrolysate, by activated charcoal is known to be a cost effective with high
capacity to absorb compounds without affecting levels of sugar in hydrolysate (Mussatto and
Roberto, 2001; Chandel et al., 2007a; Canilha et al., 2008). The effectiveness of activated
charcoal treatment depends on different process variables such as pH, contact time,
temperature and the ratio of activated charcoal taken versus the liquid hydrolysate volume
(Prakasham et al., 2009). The effectiveness of any detoxification method depends on the type
of lignocellulosic hydrolysate, as each has a different level of toxicity depending on the raw
material and pretreatment conditions (Carvalho et al., 2006). Other alternative approaches to
detoxification include adapting the fermenting organism to the hydrolysate or isolating strains
from natural and industrial habitats or harsh environments. A newer approach is the
development of inhibitor-tolerant strains through genetic modification and metabolic
engineering. However, due to the synergistic interactions among inhibitors and poor
knowledge of the mechanisms of these interactions, it is not clear against which inhibitor
resistance is desired (Olsson and Hahn Hagerdal, 1996; van Maris et al., 2006; Sanchez and
Cardona, 2008). To facilitate the development of specific, efficient and cheap detoxification
methods, intense research is required to identify the key inhibitory substances as well as their
inhibitory mechanisms. This information will also enable the modification of pretreatment
and hydrolysis processes to minimize the formation of the most potent inhibitors (Olsson and
Hahn-Hagerdal, 1996).
2.9 Enzymatic hydrolysis
Enzymatic hydrolysis of cellulose is carried out by the cellulose-hydrolyzing enzyme
cellulases, a mixture of several enzymes that hydrolyze crystalline/amorphous cellulose to
fermentable sugars (Duff and Murray, 1996). The hydrolysis of cellulose by cellulolytic
enzymes has been investigated intensively since the early 1970s, with the objective of
developing a process for the production of ethanol. Over the past decades, a great amount of
research interest and effort has been generated in this area (Coughlan, 1992; Bjerre et al.,
1996; Schwald et al., 1989; Duff and Murray, 1996; Wright, 1998; Himmel et al., 1999).
The products of the hydrolysis are usually reducing sugars majorly glucose. The
utility cost of enzymatic hydrolysis is low compared to acid or alkaline hydrolysis because
enzyme hydrolysis is usually conducted at mild conditions (pH 4-6 and temperature 45-500C)
and does not have a corrosion problem (Kuhad et al., 2010, 2011b). Both bacteria and fungi
can produce cellulases for the hydrolysis of lignocellulosic materials. These microorganisms
can be aerobic or anaerobic, mesophilic or thermophilic. Bacteria belonging to Clostridium,
Cellulomonas, Bacillus, Thermomonospora, Ruminococcus, Bacteriodes, Erwinia,
Acetovibrio, Microbispora and Streptomyces can produce cellulases and among them
Cellulomonas fimi and Thermomonospora fusca have been studied extensively (Bisaria,
1991; Duff and Murray, 1996; Sun and Cheng, 2002). These microorganisms act on various
lignocellulosic substrates for the production of cellulases under different cultivation
conditions (Table 2.9).
Table 2.9 Cellulase production by different microorganisms on various lignocellulosic
substrates under different cultivation conditions.
Microorganism
Raw
material
used as
carbon
source
Cultivation
type
Enzyme titres (U/ml)
or
(U/g substrate)
Reference
Aspergillus
oryzae
MTCC 1846
Saccharum
spontaneum
SmF FPase 0.85 ± 0.07;
CMCase 1.25 ± 0.04;
Xylanase 55.56 ± 0.52
Chandel et al.,
2009
Bacillus subtilis Banana
waste
SSF 9.6 IU/g Tsao et al.,
2000
A. niger NRRL3 Wheat bran SSF Cellobiase 215 IU/g Weber and
Agblevor,
2005
Neurospora
crassa
Wheat straw SmF 19.7 U/ml Romero et al.,
1999
Penicillium
decumbans
Wheat straw SSF FPase 23 IU/ml Yang et al.,
2004
P. janthinellum
NCIM 1171
Sugarcane
bagasse
SmF FPase 0.55; CMCase
21.58; Xylanase 28.1
IU/ml
Adsul et al.,
2004
A. fumigatus
Wheat bran,
Sugarcane
bagasse
SmF/SSF CMCase 365 U/l;
FPase 47 U/g
Grigorevski-
Lima et al.,
2009
T. reesei
NRRL11460
Sugarcane
bagasse
SSF 154.58 U/g Singhania et
al., 2006
Bacillus spp. Organic
compost
SmF 1.333 mg glucose
released
ml-1
min-1
Mayende et al.,
2006
Humicola sp.
(Th10).
Paddy straw
and Soybean
trash
SSF FPase 11.43; CMCase
15.38; Cellobiase 90.2
Kumar et al.,
2008a
Fusarium
chlamydosporum
Sugarcane
bassage
SSF FPase 95.2 IU/g;
CMCase 281.8
IU/g;
Cellobiohydrolase,
182.4 IU/g; β-
glucosidase135.2 IU/g
Qin et al., 2010
P. citrinum Rice bran SSF Endoglucanase 2.04 ± Dutta et al.,
0.13; FPase
0.64 ± 0.16 IU/ml
2008
A. terreus AV49 Groundnut
shell
SmF CMCase 1.147 IU/ml;
FPase
0.175 U/ml
Vyas et al.,
2005
T. reesei Wheat bran SmF FPase 0.33 U/ml;
CMCase 0.43 U/ml
Gomes et al.,
2006
Commercial cellulases are mainly obtained from aerobic cultivations of Trichoderma
reesei and to a lesser extent Aspergillus niger (Prasad et al., 2007; Sanchez and Cardona,
2008). Other fungi that have been reported to produce cellulases include species of
Sclerotium rolfsii, P. chrysosporium and species of Trichoderma, Aspergillus, Schizophyllum,
Fusarium and Penicillium (Sternberg, 1976; Duff and Murray, 1996; Kuhad et al., 1999; Sun
and Cheng, 2002). Of all these fungal genera, Trichoderma has been most extensively studied
for cellulase production. The cellulase manufacturing companies together with their brand
names of cellulases with the compositional details have been summarized in Table 2.10.
Table 2.10 Major cellulase producers at commercial scale for biorefinery based
applications.
Company name
and
address
Microorganism
Brand names
of
enzymes
Applications of
the
formulation
Website
link
Novozymes A/S
Krogshoejvej 36
2880
Bagsvaerd
Denmark
Trichoderma
reesei,
T. longibrachiatum
and Aspergillus
niger
Cellic CTec2,
Cellic HTec2,
Celluclast,
Novozymes
188,
Viscozyme L
Lignocellulosic
substrate
hydrolysis
http://www.
novozymes.
com
Genencor,
Danisco US
Inc., Genencor
Division
3490 Winton
Place Rochester,
NY 14623, USA
T. reesei and
T.longibrachiatum
Spezyme CP,
Accelerase®15
00,
Multifect CL
Commercially
available
biomass enzyme
developed
specifically
for second
generation
biofuels
www.genen
cor.com
Dyadic
International 140
Intracoastal
Pointe Drive,
Suite 404 Jupiter,
Florida
T. longibrachiatum AlternaFuel®
100P,
AlternaFuel®
200P
Conversion of
lignocellulosic
biomass to
glucose for
fermentation
into ethanol
www.dyadi
c.com
33477-5094 USA
Amano Enzymes
Inc
Nishiki Naka-ku,
Nagoya,
460–8630, Japan
A. niger Cellulase DS,
Cellulase AP
30K
Saccharification
of
lignocellulosics
into fermentable
sugars
www.innov
adex.com
AB Enzymes
GmbH
Feldbergstrasse
78 64293
Darmstadt
Germany
T. longibrachiatum
/T. reesei
ROHAMENT
® CL
A special
thermo stable
cellulase for
biomass
saccharification
www.abenz
ymes.com
Maps (India)
Limited,
Ahmedabad,
India
Bacillus sp. Palkolase HT,
Palkolase
LT, Palkodex
Starch
liquefaction and
saccharification
www.mapse
nzymes.co
m
Specialty
Enzymes and
Biotechnologies
Co.,Chino, CA,
USA
Bacillus sp. SEBfuel G,
SEB Amyl
GA200,
CelluSEB Fuel
Liquefaction
and
Saccharification
www.specia
ltyenzymes.
com
Iogen, (Ottawa,
Canada)
T. reesei Ultra-Low
Microbial
(ULM)
Biomass
saccharification
http://www.
iogen.ca/
Biocatalysts
Limited,
Cardiff, CF15
7QQ,Wales, UK
Trichoderma sp. Cellulase 13P,
13L
Degradation of
cellulose
completely
www.biocat
alysts.com
The cellulase system contains of three major enzyme components: β-endoglucanase
(EC 3.2.1.4), β-exoglucanase (EC 3.2.1.91) and β-D-glucosidase (EC 3.2.1.21) (Bhat and
Bhat, 1997; Lynd et al., 2002) (Fig. 2.10). The exoglucanase act on the ends of the cellulose
chain and release β-glucoside as the end product; endoglucanase randomly attack the internal
O-glycosidic bonds, resulting in glucan chains of different lengths and the β-glycosidases act
specifically on the β-cellobiose disaccharides and produce glucose (Beguin and Aubert, 1994;
Kuhad et al., 1999; Kuhad et al., 2011b). β-glucosidase catalyzes cleavage of cellobiose,
which plays a significant role in the hydrolysis process, since cellobiose is an end-product
inhibitor of many cellulases including both exo- and endo-glucanases (Lee, 1997; Galbe and
Zacchi, 2002; Rabinovich et al., 2002; Sun and Cheng, 2002). β-glucosidase, in turn is
inhibited by glucose and therefore, enzymatic hydrolysis is sensitive to the substrate
concentration (Philippidis et al., 1993). In addition to substrate concentration, pretreatment of
cellulosic materials and hydrolyzing conditions such as temperature and pH are among
factors influencing the effectiveness of enzymatic hydrolysis. (Duff and Murray, 1996; Galbe
and Zacchi, 2002).
Fig. 2.10 Procedural mechanistic action of all three cellulases on the cellulose polymer.
Hydrolysis of the individual cellulose fibres to break it into smaller sugars by exo-
cellulase, breakage of the non-covalent interactions present in the crystalline structure
of cellulose by endo-cellulase and β-glucosidase finally hydrolyze the disaccharides or
cellobiose into glucose (Source: Chandel et al., 2011).
Structurally, cellulases typically have two separate domains: a catalytic domain (CD)
and a cellulose binding module (CBM), which is linked by a flexible linker region. The CBM
is comprised of approximately 35 amino acids and the linker region is rich in serine and
threonine (Divne et al., 1998). The nature of the lignocellulosic substrate changes during the
time course of enzymatic hydrolysis (Wang et al., 2006).
Enzymatic hydrolysis methods have shown distinct advantages over acid based
hydrolysis methods; the very mild process conditions give potentially higher yields, the
utility cost is low (no corrosion problems), therefore this is the method of choice for future
wood-to-ethanol process (Duff and Murray, 1996). Many experts see enzymatic hydrolysis as
key to cost-effective ethanol production in the long run. Although acid processes are
technically more mature, enzymatic processes have comparable projected costs and the
potential of cost reductions as technology improves.
Several factors can influence the enzymatic hydrolysis of cellulose. A low substrate
concentration would result in a low overall glucose yield (Hamelinck et al., 2005). An
increase in the substrate concentration would lead to an increased glucose yield as well as an
increased rate of reaction. However, a high substrate concentration can cause substrate
inhibition which would substantially decrease the rate of the hydrolysis and the extent of
substrate inhibition depends on the ratio of total substrate to total enzyme. A high cellulase
dosage would also significantly raise process costs (Prasad et al., 2007). The susceptibility of
cellulosic substrates to enzymatic hydrolysis depends on the structural feature of the
substrate, including cellulose crystallanity, degree of polymerization, surface area and lignin
content (Sun and Cheng, 2002; Taherzadeh and Karimi, 2008). Lignin interferes with
hydrolysis by acting as a shield, preventing access of cellulases to cellulose and
hemicellulose thereby resulting in extended reaction times to achieve high conversions. On
top of that, lignin irreversibly adsorbs a large portion of the cellulase rendering it unavailable
for further hydrolysis of cellulose (Qing et al., 2010). Therefore, removal of lignin during
pretreatment is essential to dramatically increase the hydrolysis rate (McMillan, 1994; Prasad
et al., 2007). Also, removal of hemicellulose increases the mean pore size of the substrate,
thereby increasing cellulase accessibility to cellulose (Hendriks and Zeeman, 2009).
To reduce the enzyme cost in the production of fuel ethanol from lignocellulosic
biomass, two aspects are widely addressed: optimization of the cellulases production and
development of a more efficient cellulase-based catalysis system. Additionally, protein
engineering and directed evolution are powerful tools that can facilitate the development of
more efficient thermophilic cellulases (Baker et al., 2005). Recycling and reuse of the
enzymes is also an attractive methodology to reduce enzymatic hydrolysis costs (Singh et al.,
1991; Ramos et al., 1993; Lee et al., 1995; Gregg et al., 1998; Sun and Cheng, 2002; Mosier
et al., 2005). The recovery of enzymes is largely influenced by adsorption of the enzymes
onto the substrate, especially to lignin. Another constraint in the recycling of the enzymes is
enzymes inactivation. There are several strategies to recover and reuse the cellulases. The
filtrate obtained after complete hydrolysis of the cellulose fraction can be concentrated by
ultra-filtration to remove sugars and other small compounds that may inhibit the action of the
enzymes (Tu et al., 2007). Another method for recycling enzymes is by immobilization,
which enables separation of the enzymes from the process flow. The principle of
immobilization is to fixate the carbohydrolytic enzymes onto a solid matrix either by
adsorption or grafting (Dourado et al., 2002, Mosier et al., 2005). The recycling techniques
are mostly tested at laboratory scale. Therefore, the ability to scale up the techniques, the
robustness and feasibility still needs to be demonstrated.
2.10 Cultural conditions for the production of Carboxymethyl cellulase enzyme
Two main fermentation types that are generally used for the production of commercial
enzymes are submerged fermentation (SmF) and solid state fermentation (SSF) (Frost and
Moss, 1987). Two major differences are found when submerged and solid state conditions are
compared: (i) CMCase yield or productivity is higher in SSF than in SmF (ii) CMCase
location under SSF conditions is mostly extracellular, whilst it is bounded to the mycelium
under SmF conditions. Maximum CMCase activity expressed intracellularly is also 18 times
more in SSF than in SmF, while the extracellular activity is 2-5 times higher in SSF than
SmF.
2.10.1 Submerged fermentation (SmF)
Submerged fermentation involves the growth of the microorganism as a suspension in liquid
medium in which various nutrients are either dissolved or suspended as particulate solids in
many commercial media (Frost and Moss, 1987). Submerged fermentation is the preferred
method for production of most of the commercial enzymes principally because sterilization
and the process control are easier to engineer in these systems (Aunstrup et al., 1979). But
this technique is not only expensive but also of energy intensive.
2.10.2 Solid State Fermentation (SSF)
SSF technique comprising the cultivation of microorganisms on moist solid supports, either
on inert carriers or on insoluble substrates that can, in addition, be used as carbon and energy
source. The fermentation takes place in the absence or near absence of free water, thus being
close to the natural environment to which microorganisms are adapted. The aim of SSF is to
bring the cultivated fungi or bacteria into tight contact with the insoluble substrate and thus to
achieve the highest substrate concentrations for fermentation. The hyphal mode of fungal
growth and their good tolerance to low water activity (Aw) and high osmotic pressure
conditions give fungi major advantages over unicellular microorganisms in the colonization
of solid substrates and utilization of available nutrients (Krishna, 2005).
The moisture levels in SSF processes vary between 30 and 85%. SSF includes non-
aseptic conditions, use of raw materials as substrates, use of a wide variety of matrices
(which vary in composition, size, mechanical resistance, porosity and water holding
capacity), low capital cost, low energy expenditure, less expensive downstream processing
(in case, if extraction of the product is necessary, it requires less solvent and lower recovery
cost than SmF), less water usage and lower wastewater output, potential higher volumetric
productivity, higher concentration of the products, high reproducibility, lesser fermentation
space due to much higher volumetric loading of the substrate in SSF than in SmF because the
moisture level of the SSF is lower, resulting in compact fermentation facility, easier control
of contamination and generally simpler fermentation media (Mudget et al., 1986).
Solid substrate fermentation (SSF) is cheaper, less technology oriented and the
enzyme extraction is easier with the release of negligible amount of liquid effluent and
thereby produces less pollution as compared to other methods (Pandey and Radhakrishnan,
1993). Recently, a closer evaluation of these two processes in several research centers
throughout the world have revealed the enormous economical and practical advantages of
SSF over SmF. Enzyme titres are higher in SSF than in SmF when comparing the same strain
and fermentation broth. Stability of excreted enzymes and a low level of catabolic repression
are also higher in SSF technique (Lekha and Lonsane, 1994; Aguilar et al., 2007). Economic
analysis has indicated that SSF technology can considerably reduce the capital investment
and total product cost and increase profitability, thereby making it an ideal technology in
several industrial sectors (Castilho et al., 2000). Various nutritional and fermentation
parameters affect enzyme production both under submerged as well as solid state conditions
and thus they need to be optimized for maximum enzyme production.The fermentation
medium must meet the nutritional requirements of the microorganism. It basically contains
sources of carbon, nitrogen, minerals, additives and some growth factors as given below.
The sources and optimal concentration of carbon is an important factor for the
production of carboxymethyl cellulase enzyme. Different types of carbon sources (paddy
straw, wheat straw, sugarcane bagasse, jute stick, carboxymethylcellulose, corncobs,
groundnut shells, cotton, ball milled barley straw, delignified ball milled oat spelt xylan,
sulfite pulp, printed papers and mixed waste paper) have been reported for the production of
cellulase enzyme. Das et al., (2010) have reported maximum cellulase production by utilizing
CMC as carbon source by Bacillus sp.. Shabeb et al., (2010); Ariffin et al., (2008); Krishna,
(1999) and Robson and Chambliss, (1984) showed that addition of cellulose, filter paper,
CMC, starch or cellobiose to the fermentation medium favored cellulase production by
Cellulomonas sp., Clostridium and Bacillus sp.. Kumar et al., (2012) have reported maximum
cellulase activity when xylan and sucrose was used as carbon source by Bacillus cereus
MRK1.
Some investigators showed that agro-industrial residues such as rice bran, rice straw,
sugar cane bagasse and wheat bran could be used as substrates for cellulase production i.e.
Bacillus subtilis CBTK 106, Bacillus subtilis BL62 and Bacillus pumillus exhibited their
maximum cellulase productivity when wheat bran, banaba stalk and soyabean were added to
the production media respectively (Heck et al., 2002; Poorna and Prema, 2007). Ojumu et al.,
(2003) reported about some lignocellulosics which serves as carbon source for the production
of cellulase. Ikram-ul-Haq and Khan, (2006) have reported the use of wheat bran and
sugarcane bagasse for cellulase production. Kang et al., (2004) have reported higher enzyme
yields using different ratios of rice straw and wheat bran by Aspergillus sp. The amount of
carbon produced by cellulase is variable since the production of the cellulases is influenced
by substrates (carbon source) on the growth of the cellulolytic organisms. The important
cellulolytic fungus like Trichoderma sp. (Mandels and Reese, 1985); Penicillium sp. (Brown
et al., 1987); Sporotrichium sp. (Eriksson and Johnsrud, 1983); Aspergillus sp. (Kazuhisa,
1997); etc. have been reported to have cellulolytic activity.
Nitrogen sources are the secondary energy sources for the organisms which plays an
important role in the growth of the organism and enzyme production. A wide range of
nitrogenous compounds either organic or inorganic can affect the productivity of cellulase.
Organic nitrogen sources responded in positive cellulase activity more than inorganic ones in
general. Acharya and Chaudhary, (2011) have reported maximum cellulase activity when
yeast extract was added to the production medium as nitrogenous compound by B.
licheniformis WBS1 and Bacillus sp. WBS3. Kumar et al., (2012) have also reported the
maximum cellulase activity when yeast extract was added to the production medium while
Rathnan et al., (2013) have reported maximum cellulase activity when malt extract was added
as nitrogenous compound to the production medium. Sun et al., (2010) reported maximum
cellulase activity using corn-steep solid by Trichoderma sp. This was in correlation with
findings of many other workers whom found that maximum cellulase productivity was
obtained by Bacillus pumilus BpCRI 6, Pseudomonas flourescens, Monascus pupureus and
Streptomyces sp. BRC2 when tryptone was added as an organic source to the production
medium (Bakare et al., 2005; Chellapandi and Himanshu, 2008; Daniel et al., 2008).
Spiridonov and Wilson, (1998) found NH4 compounds are the most favourable nitrogen
sources for cellulase synthesis. Some other workers, found that maximum cellulase
productivity was obtained when ammonium phosphate was added to the production media by
Bacillus pumillus, Ruminococcus albus, Bacillus sp., Bacillus spp. B21 and Streptomyces sp.
BRC2 respectively (Wood et al., 1982; Kotchoni et al., 2003; Chellapandi and Himanshu,
2008). Though the addition of organic nitrogen sources such as beef extract and peptone
resulted in increased growth and enzyme production as was reported before, they were not an
effective replacement for inorganic nitrogen sources because of their higher cost (Tao et al.,
1999).
Enzymes being proteins contain ionizable groups consequently the pH of the culture
medium affects their structure and function (Frost and Moss, 1987). Most microbial
extracellular enzymes are produced in higher yield at optimum growth pH. Cellulase
production by various bacteria and fungi has been shown to be markedly dependent on pH.
Hydrogen ion concentration of the production medium strongly affects many enzymatic
processes and transport of compounds across the cell membrane. Song et al., (1985) observed
optimal cellulase production at pH 9.0 by Clostridium acetobutylium. Yang et al., (1995)
reported maximum cellulase production in pH range of 7-9 for Bacillus spp. Souichiro et al.,
(2004) reported optimum initial pH for growth and cellulose degradation of C.
straminisolvens sp. nov. at pH 7.5. Optimum pH for fungal cellulase varies from species to
species and with different substrates. This might be due to the fact that fungal cultures require
slightly acidic pH for their growth and enzyme biosynthesis (Haltrich et al., 1996). Tolan and
Foody, (1999) reported that cellulases which are active in the acidic pH range (4.8-6) are
considered to be suitable for industrial application such as stone washing denim (pH 4-7),
paper industry (pH 5), animal feed supplement (acidic pH) and textile industry. Das et al.,
(2008) also observed cellulase activity was optimum at pH 4.8 by Trichoderma reesei RUT-
C30. Baig et al., (2004) have reported pH 6.0 as optimum for maximum cellulase production
from Trichoderma lignorum using banana waste. Akiba et al., (1995) reported that the
optimum pH for cellulase activity of Aspergillus niger was between pH 6 to 7 while Sohail et
al., (2009) reported that the cellulase activity was maximum at pH 4 by Aspergillus niger
MS82. Such different results may appear because of the difference within the same genus.
Generally, the pH of the culture increased during the first two days of cellulase fermentation
by fungi due to utilization of hemicellulose and amorphous cellulose from lignocellulosic
materials for growth. After an active growth was achieved, the culture pH decreased due to
the formation of carboxylic groups and carbonic acids from lignin (Portjanskaja et al., 2006).
At this stage, the fungus has started to utilise the crystalline portion of cellulose and starts
secreting cellulase.
The temperature of the fermentation medium is one of critical factor that has profound
influence on the production of end product. The temperature requirement of the organism is
based on the nature of organisms. According to Yang et al., (1995) many Bacillus spp.
needed 32-370C for better production of cellulase. Immanuel et al., (2006) recorded
maximum CMCase activity in Cellulomonas, Bacillus and Micrococcus sp. at 400C. In
addition there were reports that the cellulase production by Aspergillus niger was observed
over a wide range of temperatures between 30 to 500C (Jaradat et al., 2008). Most work
concerning the effect of incubation temperature on growth of filamentous fungi supports the
finding that is within limits, increased incubation temperature results in increased growth rate
(Mandels et al., 1974; Brown et al., 1987). Shafique and Bajwa, (2009) observed maximum
cellulase production for T. reesei at optimum temperature of 300C. Lu et al., (2003) reported
that the cellulase production by temperature depends on the strain of microorganism.
2.11 Effect of UV mutagenesis
The electromagnetic spectrum consists of a number of different kinds of waves such as radio
waves, X-rays, infrared light, visible light, ultraviolet light and gamma rays. Being forms of
electromagnetic radiation, they all share a common ray-wave nature and in that they are all
examples of energy flowing through space without necessarily having any kind of medium.
The reason that ultraviolet light has such an effect on biological tissue is due primarily of its
interaction with deoxyribonucleic acid, or DNA, which is the "blueprint" for living cells.
Most biological growth and virtually all growth in cultures of bacteria, is the result of cells
splitting apart to form new cells (Wang and Taylor, 1991). When this occurs, the DNA of the
"parent" cell remains in the two new cells. However, the presence of ultraviolet light disrupts
the copying of DNA. In larger more complex cells like the ones found in human skin, this can
cause the DNA to incorrectly copy itself, resulting in a mutation that produces a different
kind of living cell. However, in the simpler cells of bacteria these mutations generally result
in an offspring cell that cannot perform the basic functions of life and therefore dies.
Eventually the bacteria are completely eradicated. Exposure to ultraviolet light may prohibit
growth and reproduction of bacterial cells.
Short wavelength ultraviolet light can damage chemical bonds in the bacterial cells'
DNA. With enough of the right kind of exposure, this damage occurs quickly so that the
bacteria cannot replicate and repair fast enough (Beck, 2004). As various strains of bacteria
have different biological properties, the time it takes for ultraviolet light to kill a culture of a
particular strain of bacteria can vary. Some variables are the thickness of the cell wall, the
composition of the cell wall and the speed at which the bacteria reproduce. For instance,
bacteria with thinner cell walls and quicker reproduction times tend to die after a shorter dose
of ultraviolet radiation. However, most cultures of bacteria die in a minute or less of exposure
(Goodsell, 2012).
Effect of UV mutagenesis was also analyzed on enzyme synthesis. UV rays caused
some genetic changes in microorganisms which may promote or repress the enzyme
producing genes. For example, Prabakaran et al., (2009) isolated three fungal strains from
sugarcane field and then subjected to UV mutation for highest enzyme activates production.
Among the three isolated and mutated strains, highest production of cellulases was observed
by Penicillium chrysogenum. Suntornsuk and Hang, (2008) reported that Rhizopus oryzae
when subjected to UV mutagenesis, resulted in the production of more glucoamylase as
compared to parent strain.
2.12 Fermentation
After enzymatic hydrolysis, the lignocellulosic substrates are converted to monosaccharides,
which are further fermented to ethanol by microorganisms. Approximately 80% of the
ethanol produced in the world is still obtained from the fermentation and the rest comes
largely by synthesis from the petroleum product, ethylene (Lin and Tanaka, 2006). Ethanol
fermentation is a biological process in which sugars are fermented by microorganisms to
produce ethanol and CO2. As compared to starch and molasses, the fermentation of plant
biomass (lignocellulosic) hydrolysate is a complex process. Regarding fermentation systems
for lignocelluloses to ethanol operations, the following approaches can be employed
depending on the nature of the feedstock (a) Separate Hydrolysis and Fermentation (SHF)
involves four discrete process steps (b) Simultaneous Saccharification and Fermentation
(SiSF) which consolidates hydrolysis and fermentation of cellulose hydrolysis products into
one process step (c) Simultaneous Saccharification and Co-fermentation (SSCF) involves two
process steps: cellulase production and a second step in which cellulose hydrolysis and
fermentation of both cellulose and hemicellulose hydrolysis products occurs (d) Consolidated
Bioprocessing (CBP) also known as Direct Microbial Conversion (DMC), cellulase
production, hydrolysis and fermentation of products of both cellulose and hemicellulose
hydrolysis are accomplished in a single process step. All these processes require the
hydrolysis of pre-treated biomass (with cellulase and hemicellulase enzymes or microbes);
and fermentation of resultant hexose (Glucose, Mannose, Galactose) and pentose (Xylose,
Arabinose) sugars.
2.12.1 Separate Hydrolysis and Fermentation (SHF)
Enzymatic hydrolysis performed separately from fermentation step is known as separate
hydrolysis and fermentation (SHF) (Wingren et al., 2003). In this the pretreated biomass first
undergoes enzymatic hydrolysis (saccharification) followed by ethanolic fermentation
(Sanchez and Cardona, 2008) (Fig. 2.11). A major advantage of SHF is that hydrolysis and
fermentation can be performed at their optimum operating conditions. The enzymes are
however, end-product inhibited when cellobiose and glucose accumulate (Sun and Cheng,
2002; Hahn-Hagerdal et al., 2006).
CO2
Cellulase
Glucose
Fig. 2.11 General outline of Separate hydrolysis and Fermentation
Enzyme production
C6 Fermentation Pretreatment Enzymatic Hydrolysis
Recovery of
Bioethanol
Lignocellulosic Biomass
2.12.2 Simultaneous Saccharification and Fermentation (SiSF)
The idea of performing the enzymatic hydrolysis and fermentation simultaneously was put
forward by Gauss and coworkers in a patent from 1976 (Gauss et al., 1976). In SiSF,
hydrolysis and fermentation are performed in a single process unit allowing reducing sugars
produced to be immediately consumed by the fermenting organism (Fig. 2.12). Thus, the
effect of end-product inhibition by sugars is neutralized (Hahn-Hagerdal et al., 2006; Sanchez
and Cardona, 2008). SiSF also seems to decrease the inhibition of enzymes by toxic by-
products present in pre-hydrolysate after pretreatment (Tengborg et al., 2001). This improves
the overall ethanol yield and productivity. Furthermore, SiSF compared to the two-stage SHF
process has several other advantages that include (i) a lower enzyme requirement (ii) a
reduced risk of contamination, since glucose is removed immediately and ethanol is produced
(iii) a shorter process time (iv) less reactor volume because a single reactor is used and (v)
lower capital costs (Sun and Cheng, 2002). However, there are some drawbacks of the SiSF
process, one of which is the difficulty encountered with yeast recirculation due to the
presence of lignin residues in the hydrolysate (Ohgren et al., 2007). A major disadvantage of
SiSF is that the optimum temperature condition for enzyme hydrolysis (45-500C) is much
higher than what is required for fermentation (e.g., 300C for S. cerevisiae). Therefore, a
compromise temperature of around 380C is employed meaning hydrolysis is usually the rate-
limiting process in SiSF (Philippidis and Smith, 1995; Sun and Cheng, 2002).
CO2
Cellulase
Glucose
Fig. 2.12 General outline of Simultaneous Saccharification and Fermentation
Enzyme production
Pretreatment Cellulose hydrolysis
C6 Fermentation
Recovery of
Bioethanol
Lignocellulosic Biomass
2.12.3 Simultaneous Saccharification and Co-fermentation (SSCF)
An improvement of the SSF technology called SSCF (Simultaneous Saccharification and Co-
fermentation) is targeted at ethanol production from both hexose and pentose sugars in one
step (Hahn-Hagerdal et al., 2006; Zhang and Lynd, 2010) (Fig. 2.13). The hydrolyzed
hemicelluloses during pretreatment and the solid cellulose are not separated after
pretreatment allowing the hemicelluloses sugars to be converted to ethanol together with SSF
of the cellulose (Teixeira et al., 2000). SSCF offers increased potential for a more streamlined
processing and lower capital costs. The success of SSCF and co-fermentation of hexose and
pentoses in general requires the construction of genetically engineered microorganisms able
to co-ferment glucose and xylose concurrently with enzymatic hydrolysis of cellulose and
hemicellulose.
CO2
Cellulase
Glucose
Fig. 2.13 General outline of Simultaneous Saccharification and Co-fermentation
2.12.4 Consolidated Bioprocessing (CBP)
In Consolidated Bioprocessing (CBP), ethanol together with all of the required enzymes is
produced in a single bioreactor by a single microorganism‟s community (Fig. 2.14). The
process is also known as direct microbial conversion (DMC). It is based on utilization of
mono-co-cultures of microorganisms which ferment cellulose to ethanol. CBP seems to be an
alternative approach with outstanding potential and the logical endpoint in the evolution of
ethanol production from lignocellulosic materials. Application of CBP entails no operating
costs or capital investment for purchasing enzymes or its production (Hamelinck et al., 2005;
Lynd et al., 2005).
Enzyme production
Pretreatment Cellulose hydrolysis
C5 and C6 Fermentation
Recovery of
Bioethanol
Lignocellulosic Biomass
CO2
Fig. 2.14 General outline of Consolidated Bioprocessing
For several decades, microbial utilization of sugars obtained from the hydrolysis of
lignocellulosics for the production of fuel ethanol has been an active area of research (Jeffries
et al., 1994; Dien et al., 1997; Sreenath and Jeffries, 2000; Lawford and Rousseau, 2002).
This has been largely due to the absence of suitable ethanolgens that can utilise the mixture of
the various pentose, hexose and higher sugars present in hydrolysates (Singh and Mishra,
1995). Unlike sucrose and starch-based bioethanol, which is produced from one or two sugar
monomers; lignocellulose-based ethanol is obtained through fermentation of a mixed sugar
hydrolysate, i.e. hexoses and pentoses (Zaldivar et al., 2001; Hahn-Hagerdal et al., 2006). As
a result, for lignocellulose to be economically competitive with sugar cane or grains, all types
of sugars in cellulose and hemicellulose must be efficiently converted to ethanol (Jeffries,
2006; Hahn-Hagerdal et al., 2007a). The potential fermenting organism must have these traits
so that be able to meet such demands: (i) Broad substrate utilization range (ii) High ethanol
yield (greater than 90% of theoretical) (iii) High ethanol tolerance (at more than 40 g/l) (iv)
High ethanol productivity (v) Minimal by-product formation (vi) Minimal nutrient
supplementation required (vii) Increased tolerance to inhibitors (viii) Tolerance to acidic pH
and high temperature (ix) Tolerance to process hardiness (x) Tolerance to high osmotic
pressure (xi) Recyclable (xii) Simultaneous sugar utilization (Zaldivar et al., 2001; Dien et
al., 2003; Senthilkumar and Gunasekaran, 2005). Various bacteria, fungi and yeast able to
utilise hexoses and pentoses are summarized in (Table 2.11.) and (Table 2.12.) respectively.
Pretreatment Cellulase production,
hydrolysis and Co-
fermentation
Recovery of Bioethanol
Lignocellulosic Biomass
Table 2.11 List of microorganisms that can ferment hexose sugars.
Hexose Fermenting
Organisms Reference
Bacteria
Clostridium sporogenes Miyamoto, 1997
Zymomonas mobilis Miyamoto,1997
Klebsiella aerogenes Ingram et al., 1998
Escherichia coli LY01 Dien et al., 2003
Klebsiella oxytoca Matthew et al., 2005
Fungi and Yeast
Pachysolen tannophillus Abbi et al., 1996a
Kluyeromyces marxianus Ballesteros et al., 2004
Saccharomyces cerevisiae Kuhad et al., 2010b
Saccharomyces kudriazevii Belloch et al., 2008
Pichia stipitis Gupta et al., 2009
Rhizomucor pusillis Millati et al., 2005
Saccharomyces paradoxus Belloch et al., 2008
Table 2.12 List of microorganisms that can ferment pentose sugars.
Pentose fermenting
Organisms Reference
Bacteria
Bacillus macerans Dien et al., 2003
Bacillus polymyxa Singh and Mishra, 1993
Clostridium acetobutylicum El Kanouni et al., 1998
Clostridium thermosellum Herrero and Gomez, 1980
Lactobacillus casei Roukas and Kotzekidou,1998
Lactobacills pentosus Chaillou et al., 1999
Escherichia coli Yomano et al., 1998
Fungi and Yeast
Mucor indicus Millati et al., 2005
Mucor corticolous Millati et al., 2005
Pachysolen tannophilus Schneider et al., 1981
Pichia stipitis Gupta et al., 2009
Rhizopus oryzae Millati et al., 2005
Neurospora crassa Deshpande et al., 1986
Fusarium oxysporum Jeffries and Jin, 2004
Research has confirmed considerable differences in the uptake and utilization of the
various sugars by bacteria, yeast and molds (Jeffries et al., 1994; Picataggio et al., 1994; Ho
et al., 2000; Green et al., 2001; Zhang, 2002). Some studies reported that alcohol
fermentation using lignocellulosic hydrolysates has some technological problems such as
enzymatic hydrolysis reaction of cellulose which is about two orders of magnitude slower
than the average ethanol fermentation rate with yeast (Antoni et al., 2007). Historically, the
best known microbes used for ethanolic fermentation of hexoses such as glucose and
galactose have been yeasts. Of these, Saccharomyces cerevisiae (baker‟s yeast) is the
preferred choice due to its ability to produce ethanol up to concentrations reaching 18% (w/v)
and its high tolerance of up to 150 g ethanol l-1
(Claassen et al., 1999). Various studies have
been carried out using S. cerevisiae for the fermentation of lignocellulosic hydrolysates. An
enzymatic hydrolysate of Alfa-alfa when fermented with S. cerevisiae consumed more than
98 % sugars and caused 85% fermentation efficiency with ethanol productivity of 1.3 g/l/h
(Belkacemi et al., 1997). While as per another report, the enzymatic hydrolysate (180 g/l) of
washed steam exploded oak chips when employed for continuous fermentation with S.
cerevisiae, an ethanol concentration of (77 g/l) with an ethanol productivity (16.9 g/l/h) and
ethanol yield (0.43 g/g) was obtained (Lee et al., 1999). Wang and coworkers, (2004)
reported an ethanol production 41-46 g/l from various monomeric and oligomeric sugars i.e.
glucose (85 g/l), fructose (91.1 g/l) and sucrose (96.6 g/l) using S. cerevisiae. Later on Chen
et al., (2007) used fed batch enzymatic saccharification strategy to achieve 110 g/l sugar
concentration and when this hydrolysate was fermented with S. cerevisiae, almost 95.3 g/l
sugar was consumed to produce 45.7 g/l ethanol with an ethanol yield of 94%. In another
study, enzymatic hydrolysate of acid and alkali treated cashew apple bagasse on fermentation
with S. cerevisiae produced 20.0 g/l and 8.2 g/l ethanol with an ethanol productivity of 3.33
g/l/h and 2.7 g/l/h respectively (Rocha et al., 2009). Gupta et al., (2009) and Kuhad et al.,
(2010a) have achieved an ethanol yield of 0.48 g/g from the enzymatic hydrolysate of
pretreated P. juliflora and L. camara containing 36.5 and 37.5 g/l sugars respectively.
Bacterial ethanol fermentation can use all sugars derived from cellulosic biomass; however, it
suffers from catabolite repression. The widely studied Zymomonas mobilis is considered the
work horse of bacterial ethanol fermentation (Alterthum and Ingram, 1989). Streptococcus
fragilis and Kluyveromyces fragilis are used widely for commercial ethanol production (Pesta
et al., 2006). The thermophilic bacterium Clostridium thermocellum could readily hydrolyze
cellulosic biomass; degrade hemicellulose and cellulose for ethanol production (Lynd et al.,
2002; Wu et al., 2008). Cellulolytic microorganisms give significant cellulose hydrolysis but
after hydrolysis diversion towards different metabolic shifts gives mixed gaseous acidogenic
fermentation products (Lynd et al., 2002; Demain et al., 2005). Some studies reported that
after hydrolysis of lignocellulosic biomass, the produced pentose sugars (mainly D-xylose
and L-arabinose) create problem in yeast alcohol fermentation because yeast strains lack the
xylose utilization enzymes (mainly Xylose reductase and Xylitol dehydrogenase) (Hahn-
Hagerdahl et al., 2007). Thus, the efficient utilization of the xylose component of
hemicellulose in addition to hexoses offers opportunity to significantly reduce the cost of
bioethanol production (Goldemberg, 2007).
Xylose is the most prominent pentose sugar in the hemicellulose of hardwoods and
crop residues (25% of dry weight) and is second only to glucose in natural abundance,
whereas arabinose constitutes only 2-4% of dry weight, although it can reach up to 20% in
many herbaceous crops (McMillan and Boynton, 1994). Because of their high content in
lignocellulose, the efficient utilization of pentoses is important to significantly reduce
production costs (Prasad et al., 2007). The yeast species identified so far for the pentose
fermentation are Candida shehatae, Pichia stipitis and Pachysolen tannophilus (Abbi et al.,
1996a, b; Palmqvist and Hahn-Hagerdal, 2000a; Mosier et al., 2005; Hahn-Hagerdal et al.,
2007; Talebnia et al., 2008; Kuhad et al., 2011b). Some other microorganisms that can
ferment pentose sugars are Clostridium sp., Klebsciella sp., Lactobacillus sp., Aeromonas
hydrophila, Rhizopus oryzae, Fusarium oxysporum and Neurospora crassa (El Kanouni et
al., 1998; Chaillou et al., 1999; Sreenath et al., 1999; Dien et al., 2003; Millati et al., 2005;
Ruiz et al., 2007; Vasudevan et al., 2007; Hahn-Hagerdal et al., 2007). However, their use is
limited due to slow rates of ethanol production, strict oxygen requirement, poor inhibitor
tolerance and by-product formation (Du Preez et al., 1984; Du Preez, 1994; Hahn- Hagerdal
et al., 2006).
While no microbial strain meets all the essential traits mentioned above there have
been efforts to develop the „ideal‟ organism through metabolic engineering (Dien et al., 2003;
Hahn-Hagerdal et al., 2006). The main goals of metabolic engineering can be summarized as
follows: (a) improvement of yield, productivity and overall cellular physiology (b) extension
of the substrate range (c) deletion or reduction of by-product formation and (d) introduction
of pathways leading to new products (Kern et al., 2007). Thus, through metabolic engineering
several of the traits have been transferred to adequate hosts. As a result, a variety of
organisms displaying attractive features for fermentation of lignocellulosics have been
engineered in the last three decades with most effort concentrated on the three most
promising microbial platforms, namely Z. mobilis, E. coli and S. cerevisiae (Zaldivar et al.,
2001; Dien et al., 2003; Hahn-Hagerdal et al., 2006).
2.13 Ethanol recovery: - Distillation and Dehydration
Under ideal conditions, an ethanol and water mixture can be separated based on their
difference in volatility. Because ethanol is more volatile than water (ethanol vaporizes at
780C whereas water vaporizes at 100
0C), upon heating the ratio of ethanol-to-water in the
vapor phase will become higher than that in the liquid phase. Therefore, in an ideal
distillation column separation the overhead product will mainly be ethanol and water will be
the main bottom product. An azeotropic mixture of ethanol (95.6%) and water (4.4%) will be
reached upon completion of distillation operation, which is determined by the difference in
the boiling points between water and ethanol (Fair, 2001). Because the ethanol water mixture
from fermentation is far from being ideal, the actual ethanol recovery process is a multistage
and highly integrated process (Wankat, 1988). There are several dehydration processes to
remove water from an azeotropic ethanol/water mixture. The first process is azeotropic
distillation is addition of a solvent (e.g., Benzene, Cyclohexane or Monoethylene glycol) to
break the ethanol-water azeotrope. When the additive is more volatile than water, separation
is called azeotropic distillation and when it is less volatile than water, it is called extractive
distillation.
Recently, distillation followed by molecular sieve dehydration operations have been
used to recover a pure ethanol product of fuel-grade (>99.5%). Molecular sieves are
crystalline metal aluminosilicates (zeolites) with a 3-D porous structure of silica and
tetrahedral alumina (Kresge and Dhingra, 2004). Zeolite materials can strongly and
preferentially adsorb water from vapor mixtures, thus they are able to remove the remaining
4.4% water content in the azeotropic mixture from the rectification column. Therefore,
minimization of total energy input is a critical requirement for an economic design of an
ethanol distillation/dehydration system. An alternative to molecular sieve material is corn
grits (Ladisch and Dyck, 1979). Corn grits can selectively remove water from an azeotropic
mixture and are advantageous in that the materials are bio-renewable, of low cost and easily
disposable. However, a major drawback of the corn grits is its mechanical stability over a
long period of time (Beery and Ladisch, 2001).
The use of membranes to recover ethanol by “pervaporation” (ethanol removal by
vacuum applied at the permeate side of a membrane) is another technique which conserves
energy by abolishing energy-expensive distillation. It is possible to concentrate ethanol from
80 to 99.5% by pervaporation (Parisi, 1986). It can also reduce yeast ethanol (and inhibitor)
toxicity problems if applied during fermentation.
2.14 Economic evaluation of lignocellulosic bioethanol production
The first detailed technical reports found in the literature concerning the US cases dates back
to the mid-80‟s (Chandel et al., 2007b; Gnansounou and Dauriat, 2010). In 1987, Stone &
Webster Engineering Corporation studied the economic feasibility of wood-based ethanol
plant which includes feedstock handling, acid catalyzed steam explosion pretreatment,
enzyme production and hydrolysis, concentration of glucose, fermentation, distillation and
anaerobic digestion and on the basis of constant US$ (1984) the ethanol selling price was
estimated to be $0.93/l or $3.5/gal. Similarly, another report released by Chem Systems, Inc.
in 1987 which consisted of separate hydrolysis and fermentation of hardwood, on-site
enzyme production, carbon dioxide recovery and furfural production, estimated an ethanol
selling price of $0.54/ l or $2.06/gal.
Later on, NREL reported the lignocellulose conversion to ethanol following acid
hydrolysis at a cost of ~ $0.05/l or $ 0.20/gal ethanol (Aden et al., 2002). They also reported
that though enzymatic hydrolysis has great potential for improvement but the saccharifying
enzymes are very expensive (~US$ 0.08- 0.13/l ethanol or 0.3-0.5/gal ethanol) (Aden et al.,
2002). Therefore, over the past decade, much effort was devoted to reduce the cellulases
production cost. Aden and coworkers, (2002) estimated that if the enzyme cost comes less
than 2.67 cents/l or 10 cents/gal of ethanol, the cost of ethanol production could drop as low
as $0.28/l or $1.07/gal (in 2002 dollars) and in another report NREL has aimed to achieve
this goal by 2012 (Aden, 2008). Concerning the R&D in lignocellulosic bioethanol, a “multi-
year program plan” was released and was updated every two years, including 2005 (US DOE,
2005), 2007 (US DOE, 2007) and 2009 (US DOE, 2009). The detailed updates of the
technology model are provided by Aden (2008); Aden and Foust, (2009) and Humbird and
Aden (2009).
Besides US, European research institutions have also made significant contributions
to the techno-economic evaluation of bioethanol production (Hamelinck, 2004;
Kuijvenhoven, 2006). In the REFUEL project (2006-2008) by the European Commission,
seven EU institutes evaluated the prospects for biofuels in terms of resource potential and
costs (Gnansounou and Dauriat, 2010). The economic evaluation was based on constant € of
2002 and expected a net production cost of $0.90/l or €0.62/l in 2010, $0.85/l or €0.59/l in
2020 and $0.72/l or €0.50/l in 2030 (Londo et al., 2008).
In another case study, Sassner and coworkers, (2008) compared the techno-economic
performances for the conversion of different lignocellulosics (Spruce, corn stover and salix)
to ethanol which required estimation of annual production cost including annualized capital
cost and annual operation costs. According to them, the annual production costs (US$) vary
significantly, i.e. $0.66-0.69/l ethanol (spruce), 0.67-0.86 (corn stover) and 0.72-0.87 (salix).
Similarly, Wingren et al., (2003) performed a techno-economic evaluation of simultaneous
saccharification and fermentation (SiSF) based softwood-to ethanol process. The economic
evaluation uses the same approach as by Sassner et al., (2008) and the production cost varies
between 0.546 to 0.591 US$/l. While in another study, Wright and Brown, (2007) evaluated
the economics of advanced biochemical process (pretreatment, saccharification, fermentation
and distillation) for producing bioethanol from plant fibres. Based on US$ (2007) the capital
costs and the operating costs for bioethanol production was 1.06-1.48 and 0.35-0.45 $/l
ethanol or 4.03-5.60 and 1.34-1.69 US$/gal ethanol respectively. Moreover on anticipating
further improvements in bioconversion technologies, the projected capital cost and operating
costs for future plants are estimated to be 3.33-4.44 and 0.40-0.89 US$/gal ethanol
respectively (Hamelinck et al., 2005; Wright and Brown, 2007). At the same time, Galbe et
al., (2007) also reviewed the different studies on the process economics of ethanol production
from lignocellulosic materials published during the last decade and found that the variation of
the production cost could be in the range of US$ 0.13–0.81/l of ethanol. Recently,
Gnansounou and Dauriat, (2010) proposed a six-step application of cost evaluation to the
design of lignocellulosics ethanol pathways (1) to identify desired ethanol characteristics (2)
to target selling price of lignocellulosic ethanol (3) to target cost of lignocellulosic ethanol (4)
to target cost of each step of the supply pathway (5) cost management activities and (6)
continuous improvement.
2.15 Commercialization of bioethanol
As a consequence of the mandatory targets of blending ethanol, the demand for bioethanol is
increasing rapidly in industrialized countries worldwide and it is expected that the market for
cellulosic ethanol will become mature in the next 5-10 years (Gnansounou and Dauriat,
2010). Moreover, the international ethanol market has been stimulated by governmental
policies of incentive for the use of renewable fuels. In expansion the international market is
very regional with the largest producers also being the largest consumers (Almeida and Silva,
2006). Currently, there are 448 bioethanol production units installed in Brazil (Udop, 2009),
but the country still needs expansion of ethanol production (Soccol et al., 2010). In the US,
ethanol is used in two forms: mixed with gasoline in the maximum proportion of 10%, or in
mixtures containing 85% ethanol and 15% gasoline, as an alternative fuel (EIA, 2008). In
2011, the US produced 13.9 billion gallons ethanol from its 209 ethanol refineries located in
29 states which is an increased production from 2010 (13.2 billion gallons) and 2000 (1.63
billion gallons) (RFA, 2012). While, in EU most of the members states seem to be on track to
meet or even exceed the first interim target in 2012 and will have to increase their RES shares
more rapidly in the future to meet the 2020 target. In India as well, the addition of 5% ethanol
to gasoline is mandatory in 10 states and 3 territories and in the next step, the supply of
ethanol mixtures with gasoline will be expanded to the whole country. Some efforts will also
be directed to increase the ethanol percentage in the mixture to 10% (Prasad et al., 2007).
Sweden also uses mixtures containing 5% ethanol in gasoline, while in Canada and some
regions of China mixtures containing up to 10% ethanol in gasoline may be found (Souza,
2006). In Japan, the replacement of 3% of gasoline by ethanol is authorized (Orellana and
Bonalume Neto, 2006), but efforts will be made to increase this value to 10% (Souza, 2006).
In Thailand, renewable energy policy promotes the use of a 10% blend of bioethanol with
90% gasoline (Silalertruksa and Gheewala, 2009).
Many countries worldwide such as Brazil, United States, China, India, Russia, Japan,
Malaysia, Canada, Europe, Korea, Taiwan, etc. are developing their own bioethanol
commercialization plans and strategies. Additionally, in order to accelerate the uptake of
bioethanol towards commercialization, exemption from both federal and provincial fuel
excise taxes has been provided (Mabee and Saddler, 2010; Mussatto et al., 2010), which acts
as a rebate for the bioethanol producer. Besides a few other important strategies or policies
such as government and private grants funding in R&D, subsidy to bioethanol producers ,
production of ethanol fueled vehicles, etc. are also recommended in order to promote
cellulosic ethanol production as a substitute for conventional transportation fuel (GBEP,
2008; Mabee and Saddler, 2010; Mussatto et al., 2010). However the major recommendations
in most of the policies are as described by Tan et al., (2008) (a) government and private funds
should be made available for R&D to reduce the cost of bioethanol production (b) incentives
and tax rebates should be provided to bioethanol producing companies and (c) the production
of bioethanol should be promoted by the introduction. Besides policies to promote
bioethanol, there are direct investments in R&D, pilot and demonstration plants. As a result,
several R&D projects as well as pilot plants and demonstration projects on second generation
bioethanol are being implemented worldwide
2.16 Environmental aspects
Ethanol represents closed carbon dioxide cycle because after burning of ethanol, the released
carbon dioxide is recycled back into plant material because plants use CO2 to synthesize
cellulose during photosynthesis cycle (Wyman, 1999). Ethanol production process only uses
energy from renewable energy sources; no net carbon dioxide is added to the atmosphere,
making ethanol an environmentally beneficial energy source. In addition, the toxicity of the
exhaust emissions from ethanol is lower than that of petroleum sources (Wyman and Hinman,
1990). Ethanol derived from biomass is the only liquid transportation fuel that does not
contribute to the green house gas effect.
Regarding sustainability issues with bioethanol, it is apparent that fossil fuel
combustion is contributing to an elevation of greenhouse gas (GHG) emissions (especially
CO2) and consequentially is causing changes to the earth‟s climate (Stern, 2007). Road
transport fuel combustion is currently responsible for around 20% of GHG emissions. The
reduction of GHG pollution is the main advantage of utilizing biomass conversion into
ethanol (Demirbas, 2007). Ethanol contains 35% oxygen that helps complete combustion of
fuel and thus reduces particulate emission that pose health hazard to living beings. A study
conducted by He et al., (2003) on the ethanol blended diesel (E10 and E30) combustion at
different loads found that addition of ethanol to diesel fuel simultaneously decreases cetane
number, high heating value, aromatics fractions and kinematic viscosity of ethanol blended
diesel fuels and changes distillation temperatures. These factors lead to the complete burning
of ethanol and less emissions. With its ability to reduce ozone precursors by 20-30%,
bioethanol can play a significant role in reducing the harmful gases in metro cities
worldwide. Ethanol blended diesel (E-15) causes the 41% reduction in particulate matter and
5% NOx emission (Subramanian et al., 2005). One of the disadvantage in using ethanol as
fuel is that aldehyde predominantly acetaldehydes emissions are higher than those of
gasoline. However acetaldehydes emissions generate less adverse health effects in
comparison to formaldehydes emitted from gasoline engines (Gonsalves, 2006).
2.17 Future prospects of bioethanol
Bioethanol offers great benefits for safeguarding the environment, boosting the rural
economy and ensuring fuel security. Interestingly, the world‟s focus is switching over from
corn and sugarcane to cellulosic or plant biomass as renewable raw material for production of
bioethanol (Campbell and Laherrere, 1998; Saha et al., 2005; Himmel et al., 2007; Kuhad et
al., 2011a). Nevertheless there are significant scientific, technological, sociological and
political challenges facing future bioethanol production. Besides these there are some ethical
challenges raised by increasing future bioethanol production:
Economics (affordability)
Food to fuel (changes in agricultural land use)
Genetic engineering (employment of GM-feedstock)
Local environment (localization/building of new bio-refineries: demands on fresh
water)
Bio-buisness (potential monopolization of bioresources or patents)
The major factor affecting the efficiency of the conversion of lignocellulosic materials
into energy products is the hydrolysis/saccharification of lignocellulose. The key to a
successful cellulosic ethanol production is to develop effective pretreatment technology
leading to rapid and high yield hydrolysis of lignocellulose; converting it to fermentable
sugars for subsequent fermentative production of ethanol. An efficient pretreatment strategy
should be developed that can harness maximum sugars and can fractionate lignin in a
recoverable form. Moreover for the efficient saccharification of cellulosics, approach of bio-
prospecting for novel cellulase and saccharifying enzymes should be carried out. In addition,
high throughput screening techniques and better expression systems for efficient production
of membrane proteins and enzyme complexes such as cellulosomes are in need of
development. Since the higher sugar concentration will lead to higher ethanol in the
fermentation, therefore strategies such as continuous feeding of substrate or fed-batch
enzymatic saccharification should be adopted to improve the sugar concentration in the
enzymatic hydrolysate. Moreover, to reduce the enzyme cost research is needed in the
direction to recover and reuse the enzymes.
Another major concern is the generation of microbial inhibitors during the
pretreatment process, which represent a significant carbon loss and consequently
lignocellulosic ethanol economy is largely affected due to lower ethanol yield. Although,
various detoxification strategies have been applied to remove these inhibitors for improved
hemicellulosic hydrolysate‟s fermentability (Palmqvist and Hahn-Hagerdal, 2000a,b; Mosier
et al., 2005; Chandel et al., 2007a; Gupta et al., 2009; Kuhad et al., 2010b), however, the
process of detoxification also increase the processing cost. Therefore, there is an imperative
need for bio-prospecting of new microbes capable of converting pentose sugars present in the
hydrolysate efficiently even in the presence of the toxic inhibitors (Zhang et al., 2010)
Further efforts are required to improve the fermentation efficiency of both the sugar
hydrolysates i.e. enzymatic hydrolysate (C-6 sugars) and the acid hydrolysate (C-5 sugars).
To further make the bioethanol production process successful at industrial scale with
reduction in capital and operation cost, some integrated unit operations using robust
microorganisms for better product yields should be adopted (Zhang, 2008). An ideal up-
scaling strategy needs to be fully integrated to evaluate the complete system (e.g., enzymes,
nutrients, product yields and titres and yeasts) with sufficient flexibility to investigate
alternative process configurations. From a process scale-up perspective, the challenges lie not
only with finding the most efficient organism for hemicellulose conversion but also to make
an intelligent use of the entire feedstock during process integration.