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.