Review of Literature

Review of literature

6

CHAPTER 2

REVIEW OF LITERATURE

2.1. History of ethanol as biofuel

Ethanol as a fuel has been used throughout man‟s long history. Even the

invention of ignition engines was done with bioethanol. 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). Then in 1906, the alcohol tax was lifted, which renewed the

interest in ethanol and in 1908, Henry Ford designed the automobile car „Model

T‟ to run on ethanol. By 1914, the production of ethanol had rebounded slightly

and reached 10 million gallons (Morris, 1993). But in 1919, due to the

emergence of petroleum as fuel, the use of ethanol as fuel decreased again. This

prohibition was ended in 1933 and by the early 1940s the production of ethanol

rebound again when it was used during World War II for fuel and to make

synthetic rubber. During this period, about 600 million gallons of ethanol was

produced annually in the U.S (Morris, 1993). At the end of World War II,

demand for ethanol dwindled and continued to decline for the next two decades,

mostly due to cheap petroleum imports. But again the oil embargo by Arab

countries in 1973 created petroleum shortages, resulting in significant increase

in gasoline price (Campbell and Laherrere, 1998). Since the 1970‟s, the gasoline

shortage accelerated the concerns about the rising prices for crude oil and

increasing political instability due to which the use of ethanol as biofuel is again

under consideration worldwide.

2.2. Bioethanol: First and Second generations

2.2.1. First generation bioethanol

The bioethanol produced by fermentation of sugar (sugarcane juice, molasses,

sugar beet juice, fruit juice) and starchy feedstocks (wheat, corn, potato) are

commonly known as first generation bioethanol (Antony et al., 2007). The

ethanol production methods used are enzymatic digestion (to release sugars

Review of literature

7

from stored starches), fermentation of the sugars, distillation and drying. 1st

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). However, there are a number of

concerns about the potential drawbacks of 1st generation bioethanol (IEA, 2008),

such as;

2.2.1.1. Competition between food vs fuel: It is clear that the development of

a bioenergy options, particularly food-based bioethanol may adversely affect

food demands. It is obvious fact that higher food prices will have devastating

effects on the developing world, where disposable incomes are lower. This alarm

to use food resources for alternative biofuels

2.2.1.2. Deforestation: The constant production of first generation biofuels

might lead to major deforestation and lant thus available may changed from

permanent forest cover to agriculture.

2.2.1.3. Multi-feed stock flexibility: For commercial viability, technologies

and plant designs, which are able to process a number of different feedstocks in

a flexible way is preferable. If many single food crops used for biofuels are

seasonal then to operate a plant round the year, the storage of raw material

may accelerate the cost of biofuel production.

2.2.2. Second generation bioethanol

Now, it is clearly understood that the first-generation bioethanol production is

not a sustainable approach and these increasing criticisms have raised the

attention to use non-food crops for the production of second generation

bioethanol. The second-generation bioethanol is produced from lignocellulosic

biomass comprised of the residual non-food parts of the food-crops, as well as

other crops that are not used for food purposes and also municipal, industrial

and construction waste. 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). 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, 2008). However, there is

Review of literature

8

still much work to be done in terms of improving second generation biofuel

technology pathways, to reduce costs and to improve performance and

reliability of the conversion process.

2.3. Current Status of Bioethanol

2.3.1. Current status of Bio-ethanol production worldwide

Bioethanol production worldwide has increased considerably since the oil crisis

in 1970 (Campbel and Laherrare, 1998). Its market grew from less than a billion

litres in 1975 to more than 65 billion litres in 2008 (Biofuel Platform, 2010), and

is expected to reach 100 billion litres in 2015 (Licht, 2005). According to IEA

(2008) 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). 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), but a major reason for this is sugarcane juice.

Interestingly, the innovations introduced by the automobile industry with flex-

fuel cars, which may be fueled with ethanol and/or gasoline in any proportion

increased the market for ethanol (Anfavea, 2005; Souza, 2006).

It is noteworthy that the United States (US), the largest consumer of petroleum

products (2.42 billion litres/day or 20.7 million barrels/day in 2007), meets its

demand by importing about 58% i.e., 1.4 billion litres or 12 million barrels/day

(EIA, 2008). It is predicted that the gasoline consumption will rise further along

with the rising population, as gasoline is a primary energy source that meets

non-commercial transportation demands (EIA, 2008). 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

Review of literature

9

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).

In Europe, maximum amount of ethanol is produced from wheat and sugar-beet

and France, Germany and Spain are the European countries more strongly

committed to ethanol production (Prieur-Vernat and His, 2006). The European

Union strategy for biofuels is also to decrease their dependence on oil and

reduce the negative impact caused to the environment. The share of OECD

countries in global oil demand is also expected to decrease from 57% in 2007 to

43% in 2030 (IEA, 2009).

China has also invested much in the production of ethanol, and since it is the

world‟s largest auto market, it imported about 52% of the total transportation

oil consumed in 2008 (Fang et al., 2010). The production of ethanol for biofuels

began in China in 2001, using corn as a raw material, and by 2007, four grain-

based ethanol plants with the production of about 1.75 billion litres or 1.4

million metric tons (MMT) ethanol have been developed. However, due to the

competition for ethanol and food applications, projects on fuel ethanol based on

grains were restricted and development of “non-food ethanol” (ethanol made

from non-food crops) was supported by the Chinese government (Fang et al.,

2010). Many technologies of ethanol production based on non-food crops, such

as cassava, sweet sorghum, sweet potato, Jerusalem artichoke, Kudzuvine root,

and others, are being developed (Li and Chan-Halbrendt, 2009). Till now, the

exclusive application of gasoline containing 10% ethanol to motor vehicles has

been enforced in all areas of Heilongjiang, Jilin, Liaoning, Henan, Anhui,

Guangxi and selected areas of Hebei, Shandong, Jiangsu and Hubei provinces

(Fang et al., 2010). Similarly, Thailand has also invested in the production of

ethanol. In 2007, there were 7 ethanol plants with a total capacity of 955

thousand litres/day, comprising 130 thousand litres/day cassava ethanol and

825 thousand litres/day molasses ethanol and as a result of government

promotions, 12 new plants with a total installed capacity of 1.97 Million

litres/day are being constructed (Silalertruksa and Gheewala, 2009).

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10

Other countries like Japan and Korea etc. are also in race and hence an

indigenous and affordable source of energy has become a high priority in order

to surmount the issue of energy security and sustainability. The production of

biofuels in Japan started in 2003 and by 2007 the total amount of bioethanol

production reached approximately 30 thousand litres per year (Matsumoto et

al., 2009). Similarly Korea is also very concerned about its high CO2 emission

and dependence on imported crude oil (Kim et al., 2010). The annual

consumption of gasoline in Korea is about 10 billion litres and 3 or 5 million

litres of ethanol would be needed in order to implement 3% ethanol-blending

(E3) or 5% ethanol-blended gasoline (E5) countrywide (Kim et al., 2010).

Therefore, the Korean government announced its plan to increase the supply of

transportation biofuels from 0.2 billion litres (2008) to 5 billion litres by 2030

(KMKE, 2008).

2.3.2. Status of bioethanol production in India

In the year 2003, the Planning Commission of the Government of India brought

out an extensive report on the development of biofuel (Planning Commission,

2003) and bioethanol was identified as a principal biofuel to be developed for

the nation. In India the ethanol blend in gasoline was proposed to 10 % by

2011-2012 and 5% ethanol blend in gasoline was made mandatory in 11 states

and 3 Union territories of the nation (Sukumaran et al., 2010). In 2006, the

demand for ethanol for 5% gasoline doping/blending level was 0.64 billion

Liters, while the estimated current demand for 10 % blending is projected to be

2.2 billion Liters in 2017 (Sukumaran et al., 2010). According to 2006 estimate,

the actual production of ethanol was only 0.39 billion liters which was not

sufficient to meet the fuel demand if the entire gasoline had to be doped at 5%

level.

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 bio-ethanol

technologies, which use biomass feedstock that does not have food or feed

value. The most appropriate bio-ethanol 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

(Sukukumaran et al., 2009). According to Kim and Dale (2004), the total

Review of literature

11

bioethanol production from plant biomass is estimated to be 491Giga Liters

(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 of 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. Some

major plant species along with their biomass types and biomass production has

been listed in table 2.1.

2. 4. Lignocellulose

Plant biomass is the most abundantly available and renewable natural resource

on earth. Lignocellulose is comprised of three main polymers, cellulose,

hemicellulose and lignin (Figure 2.1) and together termed as lignocellulose. The

chemical properties of the components of lignocellulosics make them a

substrate for enormous biotechnological products (Kuhad and Singh, 1993;

Kuhad et al., 1997; Sun and Cheng, 2002; Hahn-Hagerdal et al., 2007; Kuhad et

al., 2007).

2.4.1. Cellulose

Cellulose is a glucan polymer of D-glucopyranose units, which are linked

together by β-1, 4-glucosidic bonds. The wood cellulose has an average degree

of polymerization (DP) of at least 9,000–10,000 and possibly as high as 15,000.

An average DP of 10,000 would correspond to a linear chain length of

approximately 5 μm in wood. An approximate molecular weight for cellulose

ranges from about 10,000 to 150,000 Dalton (Goring and Timell, 1962).

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Table 2.1: Availability of plant biomass (lignocellulosic material) biomass

data at national level

Species Biomass

Type

Biomass

(KT/Yr)

Total Biomass

(KT/Yr) Species

Biomass

Type

Biomass

(KT/Yr)

Total Biomass

(KT/Yr)

Arhar Husk 606.2

5658.2

Mango

Bark 2.049

8.54 Stalks 5052.0 Branches 2.049

Babul

Bark 499.1

2079.0

Leaves 2.393

Branches 499.1 Twig 2.049

Leaves 581.7

Mesquite

Bark 28.58

119.08 Twig 499.1 Branches 28.58

Bajra

Cobs 1972.3

15810.71

Leaves 33.34

Husk 1793.0 Twig 28.58

Stalks 11953.0 Mustard

Husk 1553.2 8309.8

Bamboo Leaves 92.41

2466.41 Stalks 6756.6

Stalks 2374.0

Neem

Bark 2.049

8.54 Banana Residue 11936.5 11936.5 Branches 2.049

Chir pine

Bark 0.484

2.007

Leaves 2.393

Branches 0.484 Twig 2.049

Leaves 0.555 Paddy

Husk 19938.6 169478.4

Twig 0.484 Straw 149539.8

Coconut

Fronds 7253.8

11734.4 Salvadora sp.

Bark 3.069

12.801 Husk & Pith 3166.3 Branches 3.069

Shell 1314.3 Leaves 3.594

Conifers

Bark 82.14

328.56

Twig 3.069

Branches 82.14

Shisham

Bark 80.52

335.57 Leaves 82.14 Branches 80.52

Twig 82.14 Leaves 94.01

Cotton

Boll shells 6320.6

43202.0

Twig 80.52

Husk 6320.6 Sugarcane Tops & Leaves 13211.2 13211.2

stalks 30560.8 Sunflower Stalks 1391.8 1391.8

Cotton wood

Bark 2.049

8.54 Teak

Bark 1421.2

5921.7 Branches 2.049 Branches 1421.2

Leaves 2.393 Leaves 1658.1

Twig 2.049 Twig 1421.2

Eucalyptus

Bark 341.1

1463.2.4

Tobacco Stalks 316.6 316.6

Branches 341.1 Turmeric Stalks 28.7 28.7

Leaves 398.0

Umbrella

thorn Acacia

Bark 160.85

670.15 Twig 220.2 Branches 160.85

Residues 162.8 Leaves 187.6

Groundnut shell Shell 2044.6

15675.5 Twig 160.85

Stalks 13630.9 Wheat

Pods 18650.4 111902.4

Jowar

Cobs 5044.8

29827.5

Stalks 93252.0

Husk 2017.9

White mulberry

Bark 0.959

4.007 Stalks 17152.4 Branches 0.959

Maize Cobs 5612.4

28814.1 Leaves 1.13

Stalks 23201.7 Twig 0.959

Source: (http://lab.cgpl.iisc.ernet.in/Atlas/Tables/Tables.aspx); Kuhad et al. (2011a)

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13

There are several types of cellulose in wood: crystalline and noncrystalline and

accessible and nonaccessible. 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 nonaccessible 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 nonaccessible, whereas, most of the

noncrystalline cellulose is accessible but part of the noncrystalline cellulose is

so covered with both hemicelluloses and lignin that it becomes nonaccessible

(Rowell et al., 2005; Kuhad et al., 2011a). Concepts of accessible and

nonaccessible cellulose are very important in moisture sorption, pulping,

chemical modification, extractions, and interactions with microorganisms.

2.4.2. Hemicelluloses

Unlike cellulose, hemicelluloses are not chemically homogenous (Eriksson et al.,

1990; Kuhad et al., 1997; Perez et al., 2002; Kapoor, 2007; Kuhad et al., 2011a).

The hemicelluloses are comprised of both linear and branched hetero-polymers

of D-xylose, L-arabinose, D-mannose, D-glucose, D-galactose and D-glucuronic

acid (Figure 2a). In general, the hemicellulose fraction of woods consists of a

collection of polysaccharide polymers with a lower DP than cellulose (100–200)

and containing mainly the sugars D-xylopyranose, D-glucopyranose, D-

galactopyranose, L-arabinofuranose, D-mannopyranose, D-

glucopyranosyluronic acid, and D-galactopyranosyl-uronic acid with lower

amounts of other sugars. They usually contain a backbone consisting of one

repeating sugar unit linked β-(1→4) with branch points (1→2), (1→3), and/or

(1→6). 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).

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14

Figure 2.1: Structure of different components of lignocellulosic biomass

Source: Kuhad et al., 2011a

2.4.2.1. Soft wood hemicellulose

Galacto-mannans are the principal hemicelluloses in soft wood. Their backbone

is a linear chain built up by 1, 4-linked β-D-gluco-pyranose and β-D-manno-

pyranose units (Figure 2b). The mannose and glucose units in the backbone are

partially substituted at C-2 and C-3 positions by acetyl groups, approximately 1

per 3-4 hexose units as reviewed earlier (Kuhad et al., 1997). Arabino-

glucuronoxylan is another major hemicellulosic sugar and is composed of 1,4-

linked-β-D-xylopyranose units. This chain is substituted at C-2 by 4-o-Methyl-

α-D-glucuronic acid group with approximately two such units per ten xylose

units. The xylose backbone is also substituted by α-L-arabino-pyranose units,

on the average 1.3 residue per ten xylose units (Kuhad et al., 1997, 2007,

2011a, b; Perez et al., 2002). Arabino-galactan is a minor component in both

softwoods and hardwoods. The backbone of this galactan is built up by 1, 3-

linked α-D-galacto-pyranose units, and almost every galactose unit is

substituted at C-6 position.

Review of literature

15

A

cetyl-galactoglucomannan (softwoods)

O-acetyl-4-O-methylglucuronoxylan (hardwoods) glucomannans (hardwoods)

B

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O

O

O

OO

O

O

O

O

OOH

OH

OH

OH

OH OH

OAc

OAcOAc

CH3O

COOH

OH

OH

O

O

O OO

O

O

O

O

OO

O

O

O

O

OOH

OH

OH

OH

OH OH

OAc

OAcOAc

CH3O

COOH

OH

OH

O

O

O O

O

O

OO

OO

O

OH HO OH OH HO OH OH

OH

OH

OHHO

CH2HO

OAc

CH2OH

CH2OH

CH2OH

O

CH2OH

O

O

O O

O

O

OO

OO

O

OH HO OH OH HO OH OH

OH

OH

OHHO

CH2HO

OAc

CH2OH

CH2OH

CH2OH

O

CH2OH

O

O

O OO

O

O

O

O

OO

O

O

O

O

OOH

OH

OH

OH

OH OH

OAc

OAcOAc

CH3O

COOH

OH

OH

O

O

O OO

O

O

O

O

OO

O

O

O

O

OOH

OH

OH

OH

OH OH

OAc

OAcOAc

CH3O

COOH

OH

OH

O

O

O O

O

O

OO

OO

O

OH HO OH OH HO OH OH

OH

OH

OHHO

CH2HO

OAc

CH2OH

CH2OH

CH2OH

O

CH2OH

O

O

O O

O

O

OO

OO

O

OH HO OH OH HO OH OH

OH

OH

OHHO

CH2HO

OAc

CH2OH

CH2OH

CH2OH

O

CH2OH

O

O

O OO

O

O

O

O

OO

O

O

O

O

OOH

OH

OH

OH

OH OH

OAc

OAcOAc

CH3O

COOH

OH

OH

O

O

O OO

O

O

O

O

OO

O

O

O

O

OOH

OH

OH

OH

OH OH

OAc

OAcOAc

CH3O

COOH

OH

OH

O

O

O O

O

O

OO

OO

O

OH HO OH OH HO OH OH

OH

OH

OHHO

CH2HO

OAc

CH2OH

CH2OH

CH2OH

O

CH2OH

O

O

O O

O

O

OO

OO

O

OH HO OH OH HO OH OH

OH

OH

OHHO

CH2HO

OAc

CH2OH

CH2OH

CH2OH

O

CH2OH

OHO

O

O OO

O

O

O

O

OO

O

O

O

O

OOH

OH

OH

OH

OH OH

OAc

OAcOAc

CH3O

COOH

OH

OH

O

O

O OO

O

O

O

O

OO

O

O

O

O

OOH

OH

OH

OH

OH OH

OAc

OAcOAc

CH3O

COOH

OH

OH

O

O

O O

O

O

OO

OO

O

OH HO OH OH HO OH OH

OH

OH

OHHO

CH2HO

OAc

CH2OH

CH2OH

CH2OH

O

CH2OH

O

O

O O

O

O

OO

OO

O

OH HO OH OH HO OH OH

OH

OH

OHHO

CH2HO

OAc

CH2OH

CH2OH

CH2OH

O

CH2OH

O

O

O OO

O

O

O

O

OO

O

O

O

O

OOH

OH

OH

OH

OH OH

OAc

OAcOAc

CH3O

COOH

OH

OH

O

O

O OO

O

O

O

O

OO

O

O

O

O

OOH

OH

OH

OH

OH OH

OAc

OAcOAc

CH3O

COOH

OH

OH

O

O

O O

O

O

OO

OO

O

OH HO OH OH HO OH OH

OH

OH

OHHO

CH2HO

OAc

CH2OH

CH2OH

CH2OH

O

CH2OH

O

O

O O

O

O

OO

OO

O

OH HO OH OH HO OH OH

OH

OH

OHHO

CH2HO

OAc

CH2OH

CH2OH

CH2OH

O

CH2OH

O

(i) (ii) (iii) (iv)

(v) (vi) (vii)

Figure 2.2: (A) Structure of monosaccharides commonly present in xylan

backbone (where (i) β-D-glucopyranose; (ii) α-L-rhamnopyranose; (iii) α-L-

fucopyranose; (iv) β-D-xylopyranose; (v) β-D-mannopyranose; (vi) β-D-

galactopyranose and (vii) α-L-arabinofuranose); (B) Structures of polymeric

units of softwood and hardwood xylans.

Source: Kuhad et al., 2011b

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16

2.4.2.2. Hardwood hemicellulose

The o-acetyl-4-o-Methyl-glucurono-β-xylan (commonly known as

glucuronoxylan) is the major component of hard wood hemicelluloses (Kuhad et

al., 1997; Pereira et al., 2003) (Figure 2B). The xylan content varies between 15-

30% in different hard wood species. The backbone of xylan consists of β-D-xylo-

pyranose units linked by 1, 4-bonds, while seven of ten xylose units are

substituted by acetyl group at C-2 or C-3 position and in one of ten xylose

units, the 4-o-methy1-α-D-glucuronic acid residue unit is linked at C-1, 2

positions to the hemicellulose backbone (Bastawde et al., 1992; Kuhad et al.,

1997; Perez et al., 2002; Kapoor, 2007). Gluco-mannan is another hemicellulose

in hard woods (Figure 2). This polysaccharide comprises 2% to 5% of the wood

and is composed of β-D-gluco-pyranose and β-D- manno-pyranose units by 1,

4- bonds. Depending on the wood species, the glucose: mannose ratio varies

between 1:1 and 1:2 (Kuhad et al., 1997; Perez et al., 2002; Rowel et al., 2005;

Kapoor, 2007).

2.3.3. Lignin

Lignin is amorphous and highly complex polymer of phenylpropanoid units and

is considered to be an encrusting substance in the plant biomass (Kuhad et al.,

1997). The precursors of lignin biosynthesis are p-coumaryl alcohol, coniferyl

alcohol, and sinapyl alcohol. p-coumaryl alcohol is a minor precursor of both

softwood and hardwood lignins. Whereas, the coniferyl alcohol is the

predominate precursor of softwood lignin, and coniferyl and sinapyl alcohol are

both precursors of hardwood lignin (Alder, 1977). Softwood lignin has methoxyl

content of ~15–16%, while, hardwood lignin has a methoxyl content of ~21%.

Moreover ecological factors such as age of the wood, climate, plant sustenance

and amount of sunlight also affect the chemical structure of lignin (Kuhad and

Singh, 1993; Kuhad et al., 1997) Lignin does not have a single repeating unit

like cellulose does, but instead consists of a complex arrangement of

substituted phenolic units. Lignins can be classified in several ways, but they

are usually divided according to their structural elements. All wood lignins

consist mainly of three basic building blocks of guaiacyl, syringyl, and p-

hydroxyphenyl moieties, although other aromatic units also exist in many

different types of woods. There is a wide variation of structures within different

wood species. The lignin content of softwoods is usually in the range of 18–25%,

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17

whereas the lignin content of hardwoods varies between 25 and 35%. The

phenyl propane can be substituted at the α, β, or γ positions into various

combinations linked together both by ether (C-O-C) and carbon to carbon (C-C)

linkages (Sakakibara, 1991). Lignins from softwoods are mainly a

polymerization product of coniferyl alcohol and are called guaiacyl lignin.

Hardwood lignins are mainly syringyl-guauacyl lignin, because they are a

copolymer of coniferyl and sinapyl alcohols. The ratio of these two types varies

in different lignins from about 4:1 to 1:2 (Sarkanen and Ludwig, 1971). List of

few lignocellulosic residues and their chemical composition is shown in Table

2.2.

Table 2.2: Chemical composition of various lignocellulosic residues

Substrate Hexosans Pentosans Lignin

Barley wood 40 20 15

Birch wood 40 33 21

Coastal Bermuda grass 25 35.7 6.4

Corn cobs 42 39 14

Corn stalks 35 15 19

Corn stover 38 26 19

Cotton seed hair 80-95 5-20 0-5

Flax sheaves 35 24 22

Forage sorghum 34 17 16

Grasses 25-40 35-50 10-30

Groundnut shells 38 36 16

Hardwood stem 40-55 34-40 18-25

Leaves 15-20 80-85 0-5

Miscanthus 43 24 19

Municipal solid waste 8-15 NA 24-29

News paper 40-55 25-40 18-30

Oat straw 41 16 11

Paper 85-99 0-5 0-15

Pine 41 10 27

Rice husk 36 15 19

Rice straw 32 24 13

Rye straw 31 25 7

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Salix 41.5 22-25 25

Saw dust 55 14 21

Soft wood stem 45-50 25-35 25-35

Solid cattle manure 1.6-4.7 1.4-3.3 2.7-5.7

Sorghum straw 33 18 15

Sorted refuge 60 20 20

Soya bean stalks 34 25 20

Spruce 45 26 28

Sugarcane bagasse 33 30 29

Sweet sorghum 23 14 11

Swine waste 45 30-35 12

Switch grass 37 29 19

Waste paper from chemical pulp 60-70 10-20 5-10

Wheat straw 30 24 18

2.5. Bioconversion of lignocellulosic biomass into ethanol

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 (Figure 2.3). The hydrolysis is usually

catalyzed by cellulase enzymes, and the fermentation is carried out by yeasts or

bacteria. The factors that have been identified to affect the hydrolysis of

cellulose include porosity (accessible surface area) of the waste materials,

cellulose fiber crystallinity, and lignin and hemicellulose content (Mosier et al.,

2005; Margeot et al., 2009; Alvira et al., 2010; Kuhad et al., 2011a). 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 crystallinity, 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).

2.5.1. Pretreatment of lignocellulosic biomass

The effect of pretreatment strategies of lignocellulosic materials has been well

recognized for a long time (Mosier et al., 2005; Sanchez and Cardona, 2008;

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19

Margeot et al., 2009; Kumar et al., 2009; Alvira et al., 2010; Kuhad et al.,

2011a). The pretreatment strategies must meet the following requirements: (1)

improve the formation of sugars or the ability to subsequently form sugars by

enzymatic hydrolysis; (2) avoid the degradation or loss of carbohydrate; (3) avoid

the formation of hydrolysis and fermentation inhibitory byproducts; and (4)

cost-effectiveness of the process. Various physical, physico- chemical, chemical,

and biological processes have been used for pretreatment of lignocellulosic

materials.

Figure 2.3: Schematic representation of process for bioethanol production

from lignocellulosic biomass

Source: Kuhad et al (2011a)

2.5.1.1. Physical pretreatment

2.5.1.1.1. Mechanical comminution

Lignocellulosic residues can be pretreated by comminution through a

combination of chipping, grinding and milling to reduce cellulose crystallinity

depending on the final particle size of the material (10–30 mm after chipping

and 0.2–2 mm after milling or grinding) (Sun and Cheng, 2002). The milling

process has been found to reduce the cellulose crystallinity and subsequently

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20

improving the digestibility of the lignocellulosic biomass efficiently. Different

milling processes (ball milling, two-roll milling, hammer milling, colloid milling

and vibro energy milling) can be used to improve the enzymatic hydrolysis of

lignocelullosic materials (Taherzadeh and Karimi, 2008). The power requirement

for mechanical comminution of agricultural materials depends on the final

particle size and characteristics of plant materials (Sun and Cheng, 2002;

Mosier et al., 2005; Hendriks and Zeeman, 2009).

2.5.1.1.2. Extrusion

Extrusion process is a novel and promising physical pretreatment method for

biomass conversion to ethanol. In extrusion, the materials are subjected to

heating, mixing and shearing, resulting in physical and chemical modifications

during the passage through the extruder. Screw speed and barrel temperature

are believed to disrupt the lignocellulose structure causing defibrillation and

shortening of the fibers, which in turn increases carbohydrates accessibility for

enzymatic hydrolysis (Karunanithy et al., 2008). The different reactor

parameters must be taken into account to achieve the highest efficiency in the

process. In recent studies application of enzymes during extrusion process is

being considered as a promising technology for ethanol production (Alvira et al.,

2010).

2.5.1.1.3. Pulsed-Electric-Field Pretreatment

Pulsed-electricfield (PEF) pretreatment involves application of a short burst of

high voltage to plant materials placed between two electrodes. PEF pretreatment

can have serious effects on the structure of plant tissues. When a high-

intensity, external electric field is applied, a critical electric potential is induced

across the cell membrane, which leads to rapid electrical breakdown and local

structural changes in cell membrane and the cell wall. The electric field results

in a dramatic increase in mass permeability and, in some cases, mechanical

rupture of the plant tissue.

In biomass-to-fuel conversion, pretreatment of biomass with PEFs can expose

the cellulose in the plant fibers. Using high field strengths in the range of 5-20

kV/cm, plant cells can be significantly ruptured (Kumar et al., 2009). By

applying electric pulses with high field strengths, PEF pretreatment can create

permanent pores in the cell membrane and hence facilitate the entry of acids or

enzymes used to break down the cellulose into its constituent sugars. In the

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21

case of the chemical modification of plant tissue, particularly in lignocellulose

hydrolysis, appropriate chemicals might need to be transported into the tissue

to aid in cell-wall breakdown and digestion. Two advantages of PEF

pretreatment are that it can be carried out at ambient conditions and energy

requirement is low because of very short pulse times (100 μs). Furthermore, the

actual PEF process itself does not involve moving parts, so no complex

instrument design is required. Recently Kumar et al. (2009) had used this

process for Switch grass and found considerable improvement in the enzymatic

digestibility of the substrate.

2.5.1.1.4. Pyrolysis

Pyrolysis of lignocellulosic materials was carried out at temperatures above

300°C, under which the cellulose is decomposed to produce residual char and

gaseous products. The dilute acid hydrolysis of residual char resulted in more

than 80% conversion of cellulose to reducing sugars with more than 50%

glucose (Fan et al., 1987). Moreover, the process efficiency can be enhanced

under oxygen rich conditions (Shafizadeh and Bradbury, 1979), and also with

the addition of some specific catalysts such as zinc chloride or sodium

carbonate improved process efficacy (Sun and Cheng, 2002; Zwart et al., 2006;

Kumar et al., 2009).

2.5.1.1.5. Ultrasound pretreatment

The effect of ultrasound waves on lignocellulosic biomass has also been

employed for extracting hemicelluloses, cellulose and lignin (Sun and

Tomkinson, 2002). Several researchers have reported the enhanced

saccharification of cellulose using ultrasonic pretreatment methods (Yachmenev

et al., 2009). Higher enzymatic hydrolysis yields after ultrasound pretreatment

may be attributed to the cavitation effects. The cavitation effect is caused by

introduction of ultrasound field into the enzyme-substrate suspension greatly

enhance the transport of enzyme macromolecules toward the substrate surface.

Furthermore, mechanical impacts, produced by the collapse of cavitation

bubbles, provide an important benefit of opening up the surface of solid

substrates for enzymatic action. The maximum effects of cavitation occur at 50

°C, which is the optimum temperature for many enzymes (Yachmenev et al.,

2009).

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22

2.5.1.1.6. Microwave pretreatment

Microwave-based pretreatment can be considered a physicochemical process

since both thermal and non-thermal effects are often involved. Pretreatments

were carried out by immersing the biomass in dilute chemical reagents and

exposing the slurry to microwave radiation for residence times ranging from 5 to

20 min. Zhu and coworkers (2006) have identified alkalis as suitable chemical

reagents for microwave-based pretreatment. Among different alkalis used, the

sodium hydroxide is observed as the most effective alkali reagent.

2.5.1.2. Chemical pretreatment

2.5.1.2.1. Acid hydrolysis

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

sulfuric 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). Recently the focus of

dilute acid hydrolysis processes, remained on using less severe conditions and

achieve 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). There are primarily two types of dilute acid

pretreatment processes: high temperature, continuous-flow process for low solid

load (5–10%), and low temperature, batch process for high solid load (10–40%).

Although dilute acid pretreatment can significantly improve the cellulose

hydrolysis, its cost is usually higher than some physico-chemical pretreatment

processes such as steam explosion or Ammonia Fiber Explosion/Expansion

(AFEX). Moreover, the neutralization of pH is necessary for the downstream

enzymatic hydrolysis and fermentation processes.

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23

Recently, few organic acids such as fumaric or maleic acids are also used as

alternatives to inorganic acids enhance cellulose hydrolysis for ethanol

production. These acids when compared with sulfuric acid in terms of

hydrolysis yields from wheat straw and formation of sugar degradation

compounds during pretreatment, showed higher efficiency with less amount of

furfural (Kootstra et al., 2009).

2.5.1.2.2. Alkaline hydrolysis

Alkaline hydrolysis is one of the critical method used to pretreat the plant

biomass, however the effect of alkaline pretreatment method depends on the

lignin content of the materials (Fan et al., 1987; McMillan, 1994; Sun and

Cheng, 2002; Mosier et al., 2005; Kumar et al., 2009; McIntosh and Vancov,

2010; Gupta et al., 2011a). The mechanism of alkaline hydrolysis is believed to

be saponification of intermolecular ester bonds cross-linking xylan

hemicelluloses and other components, for example, lignin and other

hemicellulose. Dilute NaOH treatment of lignocellulosic materials caused

swelling of lignocellulosic materials, leading to an increase in internal surface

area, a decrease in the degree of polymerization, and crystallinity, separation of

structural linkages between lignin and carbohydrates, and disruption of the

lignin structure is commonly reported (Sun and Cheng, 2002; Carrillo et al.,

2005). Recently, Hu and coworkers (2008) used microwave, and radio frequency

based dielectric heating in the alkali pretreatment of switchgrass to enhance its

enzymatic digestibility. In this strategy switchgrass could be treated on a large

scale at high solid loading with uniform temperature distribution (Hu et al.,

2008, Hu and Wen, 2008).

2.5.1.2.3. Ionic liquids (ILs) pretreatment

The use of ionic liquids (ILs) as solvents for pretreatment of cellulosic biomass

has recently received much attention (Kumar et al., 2009; Kuhad et al., 2011a).

ILs are salts, typically composed of large organic cations and small inorganic

anions, which exist as liquids at relatively low temperatures; often at room

temperature. Their solvent properties can be varied by adjusting the anion and

the alkyl constituents of the cation. These interesting properties include

chemical and thermal stability, non-flammability, low vapour pressures and a

tendency to remain liquid in a wide range of temperatures (Hayes, 2009). Since

no toxic or explosive gases are developed, ionic liquids are also known as

„„green” solvents. Carbohydrates and lignin can be simultaneously dissolved in

Review of literature

24

ILs with anion activity (e.g. the 1-butyl-3 methylimidazolium cation (C4mim)+)

because ILs form hydrogen bonds between the non-hydrated chloride ions of the

IL and the sugar hydroxyl protons in a 1:1 stoichiometry. As a result, the

intricate network of non-covalent interactions among biomass polymers of

cellulose, hemicellulose, and lignin is effectively disrupted while minimizing

formation of degradation products. However, most of the work showing the

effectiveness of ILs has been carried out using pure crystalline cellulose, and its

applicability to a more complex combination of constituents in lignocellulosic

biomass requires more extensive studies. The use of ILs has also been already

demonstrated on some lignocellulosic feedstocks such as straw (Li et al., 2009)

or wood (Lee S.H. et al., 2009). The ILs technology is under development,

therefore the commercial recovery methods have not been fully developed. In

addition, techniques need to be worked out to recover hemicellulose and lignin

from solutions after extraction of cellulose (Hayes, 2009).

2.5.1.2.4. Lime treatment

Lime pretreatment removes the lignin fraction from the polysaccharide fraction,

thus making the remaining polysaccharides vulnerable to enzyme digestion

(Kim and Holtzapple, 2005, O‟Dwyer et al., 2007). With regard to process

operation, different conditions are employed for different types of cellulosic

materials: 100°C for 13 h for corn stover, 150°C for 6 h with 14 atm for poplar

wood, and 100°C for 2 h for switchgrass (Mosier et al., 2005). The oxygenation of

reaction mixture can greatly improve delignification, especially when treating

lignin rich woods. The process of lime pretreatment involves slurrying the lime

with water, spraying it onto the biomass and storing thhe material in a pile for

few weeks (Mosier et al., 2005, Kim and Holtzapple, 2005). Recently, Park and

coworkers developed a novel a one-batch lime-pretreatment process called

„„calcium capturing by carbonation (CaCCO)”. The CaCCO process entraps the

formed CaCO3 in the reaction vessel throughout the conversion process and no

solid-liquid separation approach has been used (Park et al., 2010).

2.5.1.2.5. Organosolv process

In the organosolv process, an organic or aqueous organic solvent mixture with

inorganic acid catalysts (HCl or H2SO4) is used to break the internal lignin and

hemicellulose bonds (Xu et al., 2003; Zhao et al., 2009). The solvents used in

the process include methanol, ethanol, acetone, ethylene glycol, triethylene

glycol, tetrahydrofurfuryl alcohol and organic acids such as oxalic,

Review of literature

25

acetylsalicylic and salicylic acid etc. (Sun and Cheng, 2002; Itoh et al., 2003; Xu

et al., 2003; Kumar et al., 2009; Zhao et al., 2009; Kuhad et al., 2011a). A high

yield of xylose is usually obtained with the addition of acid. Solvents used in the

process are drained from the reactor, evaporated, condensed and recycled to

reduce the process cost. Removal of solvents from the system is necessary

because the solvents may be inhibitory for growth of organisms, enzymatic

hydrolysis, and subsequent fermentation (Itoh et al., 2003; Xu et al., 2003; Zhao

et al., 2009; Kuhad et al., 2011a).

2.5.1.2.6. Oxidative delignification

The pretreatment of lignocellulosic biomass with hydrogen peroxide greatly

enhanced its susceptibility to enzymatic hydrolysis. About 50% lignin and most

hemicellulose were solubilized by 2% H2O2 at 30 °C within 8 h, and 95%

efficiency of glucose production from cellulose was achieved in the subsequent

saccharification by cellulase at 45 °C for 24 h (Azzam, 1989). Besides, alkaline

peroxide (Bjerre et al., 1996; Lissens et al., 2004; Martín 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 (20 g straw/l, 170 °C, 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).

2.4.1.2.7. Ozonolysis

Ozone is used to degrade lignin and hemicellulose in many lignocellulosic

materials. The degradation was essentially limited to lignin and hemicellulose

was slightly attacked, but cellulose was un-affected which resulted into

increased in vitro digestibility of the cellulosic substrates (Kumar et al., 2009).

According to Vidal and Molinier (1988), the ozone treatment removed 60% lignin

from wheat straw which in turn enhance the enzymatic saccharification rate by

5 times. 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 (García-Cubero et al., 2009). However, requirement of large amount of

ozone makes the process expensive and commercially unfeasible.

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26

2.5.1.3. Physico-chemical pretreatment

2.5.1.3.1. Ammonia fiber explosion (AFEX) and Ammonia Recycle Percolation

Process

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). The concept of AFEX is similar to steam explosion. In

ammonia fiber/freeze expansion (AFEX) process, a 5–15% ammonia solution

flows through a column reactor that is packed with biomass at 1 mL/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).

Moreover, to reduce the cost and protect the environment, ammonia must be

recycled after the pretreatment. In an ammonia recovery process (ARP), aqueous

ammonia (10-15 wt %) passes through biomass at elevated temperatures (150-

170 °C) with a fluid velocity of 1 cm/min and a residence time of 14 min, after

which the ammonia was then withdrawn from the system by a pressure

controller for recovery. In the ARP method, the ammonia is separated and

recycled and since, the ammonia pretreatment does not produce inhibitors for

the downstream biological processes, so washing is not required (Sun and

Cheng, 2002; Galbe and Zacchi, 2007; Margeot et al., 2009). Generally, AFEX

and ARP processes are not differentiated in the literature, although AFEX is

carried out in liquid ammonia and ARP is carried out in an aqueous ammonia

solution (10-15%). It is also observed that the ammonia fiber explosion

pretreatment simultaneously reduces lignin content and removes some

hemicellulose and decrystallize cellulose. The cost of ammonia, and especially of

ammonia recovery, elevates the cost of the AFEX pretreatment (Mosier et al.,

2005; Kumar et al., 2009; Alvira et al., 2010).

2.5.1.3.2. Auto-hydrolysis (Steam explosion)

Steam explosion is commonly reported method for lignocellulosic materials. The

process causes hemicellulose degradation and lignin transformation due to high

temperature, thus increasing the potential of cellulose hydrolysis (Lee J.M. et

al., 2009, Boluda-Aguilar et al., 2010). Steam explosion pretreatment typically

Review of literature

27

subjects lignocellulose to temperatures between 160 and 260°C (corresponding

pressures of 100–700 psia; psia = pounds per square inch absolute;

atmospheric pressure is 14.5 psi) with saturated steam for a period of ten

seconds to several minutes, followed by a flashing process to explosively release

the steam. This treatment results in an explosive disruption of the lignocellulose

material, thus “opening up” the substrate to increase digestibility (Mosier et al.,

2005). The factors that affect steam explosion pretreatment are residence time,

temperature, chip size and moisture content (Duff and Murray, 1996; Lee et al.,

2009; Alvira et al., 2010). 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 (Palmqvist and Hahn-Hagerdal, 2000 a, b;

Chandel et al., 2007a, Jurado et al., 2009).

2.5.1.3.3. 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 CO2 would form carbonic acid and increase the hydrolysis

rate. 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.

2.5.1.3.4. 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 corn stover (Mosier et

al., 2005), sugarcane bagasse (Laser et al., 2002) and wheat straw (Pérez et al.,

2008). Pressured reactors are used to keep the water in the liquid state at high

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28

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

depolymerization; 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 (Weil et al., 1998;

Mosier et al., 2005; Lu and Mosier, 2008; Alvira et al., 2010).

2.4.1.3.5. Control pH liquid Pretreatment

The controlled pH liquid hot water pretreatment process will maximize the

solubilization of the hemicellulose fraction as liquid soluble oligosaccharides

with minimum formation of monomeric sugars. The minimization of complete

hydrolysis to monosaccharides minimizes the subsequent degradation of these

sugars to various aldehydes during pretreatment. By controlling the

depolymerization of hemicellulose, the major xylose containing product is

soluble xylan oligosaccharides (Weil et al., 1998). These oligosaccharides must

be subsequently hydrolyzed to fermentable sugars by enzymes or acids. In

liquid hot water pretreatment, acetic acids and other organic acids are liberated

as a result of the cleavage of O-acetyl and uronic acid substitution on

hemicellulose by the action of water. These acids help catalyze further

hemicellulose solubilization. They also may degrade the resulting monomeric

sugars to furfural, which may have negative effects on the subsequent

fermentation. With careful addition of base or buffer, controlled pH

hydrothermolysis can maintain the pH of the liquid phase between 5.5 and 7.0

during the whole process, thus minimizing the formation of degradation

products. Thus controlled pH liquid hot water is a modified version of hot water

pretreatment which provides greater control of the chemical reactions that

occur during pretreatment (Weil et al., 1998; Lu and Mosier, 2008).

2.5.1.4. Biological pretreatment

The pretreatment has become a necessity to maximize the hydrolysis of

cellulosics and eventually the production of ethanol. The advantages of

biological delignification of plant amterial over chemical and mechanical

pretreatment methods include (i) mild reaction conditions, (ii) avoids the use of

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29

toxic and corrosive chemicals, (iii) higher product yield, (iv) fewer side reactions,

less energy demand and (v) less reactor resistance to pressure and corrosion.

(Lee, 1997; Kuhar et al., 2008; Sanchez, 2009). In situ microbial delignification

appears to be a feasible strategy to achieve improved depolymerization of

hemicellulose and cellulose.

The white-rot fungi (WRF) are the most effective microorganisms for biological

pretreatment as they degrade lignin more extensively and rapidly than any other

known group of organisms (Eriksson, 1993; Kuhad et al., 1997; Keller et al.,

2003; Kuhad et al., 2007). 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). Thus, selected lignin-degrading WRF with

comparatively low cellulase and xylanase activities could be advantageous for

efficient delignification and eventually in the reduction of chemical and energy

inputs for chemical or enzymatic hydrolysis of the substrate(s).

Few studies have been reported on the pretreatment of plant biomass with WRF

for its affect on cellulose hydolysis. 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

coworkers (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). Recently our group has demonstrated the

potential of insitu pretreatment of P. juliflora with Crinipellis sp. RCK-1 before

its acid or enzymatic hydrolysis in increasing sugar yield and in turn producing

ethanol as a biofuel (Kuhar et al., 2008).

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 (SSF) from beech wood chips after bio-organosolvation

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30

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 hemicelluloses 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. Recently our group has studied the SSF of P. juliflora and L.

camara followed by acid hydrolysis and observed that fungal treatment

significantly reduce the amount of inhibitors generated and eventually the

requirement of detoxiying agent was also reduced (Gupta et al., 2011b)

2.5.2. Detoxification

Thermochemical pretreatments are the most commonly used treatment for the

deconstruction of plant materials. However, under such stringent conditions,

the carbohydrate moieties of lignocellulosic biomass undergo non-selective

degradation resulted in generation of fermentation inhibitory products (Chandel

et al., 2007a; Gupta et al., 2009, 2011b; Kuhad et al., 2010b, 2011a). On the

basis of their origin these inhibitors are divided into three groups, furfurals,

hydroxymethyl furfurals (HMF) and phenolics. Furfural and hydroxymethyl

furfurals are pentose and hexose sugars degradation compounds, respectively.

While, the phenolics are the degradation compounds of acid soluble lignin

(Chandel et al., 2007a). These compounds depending on their concentration in

the hydrolysates 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

dehydrogenase (ADH) contributes to the acetaldehyde excretion, which resulted

in the prolonged lag phase in the microbes (Palmqvist and HahnHagerdal,

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

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31

matrix. Therefore in order to make these hydrolysates fermentable their

detoxification to remove the inhibitors without much sugar loss is essential.

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 hydrolysates and on the species of

microorganisms employed.

2.4.2.1. Physical detoxification

The physical methods for detoxification of hydrolysates includes vacuum

evaporation, extraction, activated charcoal adsorption and ion exchange

treatment. Vacuum evaporation of hydrolysates reduced the contents of volatile

compounds such as acetic acid, furfural and vanillin, present in the

hydrolysates and thus improves their fermentability. Converti et al. (2000)

reported that evaporation is suitable to remove the acetic acid, furfural and

other volatile compounds from hemicellulose hydrolysates improving the

fermentative process for xylitol production. However, the method also has a

drawback to moderately enhance the concentration of non-volatile toxic

compounds (extractives and lignin derivatives) which could lead to increased

fermentation inhibition. Parajo and coworkers (1998) used vacuum evaporation

method with wood hydrolyzate and observed an increase in concentration of

lignin derivatives and extractives which resulted in longer fermentation time

(from 24 to 96 h) leading to poor productivity. Similar observations were also

reported by other workers as well (Larsson et al., 1999; Silva and Roberto,

2001). As an additional report the ethyl acetate extraction has also been

reported to increase the ethanol yield in fermentation in Pichia stipitis (Pasha et

al., 2007).

In contrast, Weil et al. (2002) used two ion exchange resins XAD-4 and XAD-7

for the detoxification of hydrolysates and observed that the strain E. coli KO11

could ferment the detoxified hydrolysate nearly as rapidly as the mixed sugars

control with in 80h and produced equal amount of ethanol. Similarly, Chandel

and colleagues (2007) reported a significant decrease of furans (63.4%), total

phenolics (75.8%) and acetic acid (85.2%) when an industrial resin DIAION

(HPA 25) was used for the detoxification of acid hydrolysate of sugarcane

bagasse. Interestingly, Wickaramasinghe and Grzenia (2008) when compared

the efficiency of anion exchange membrane to that of anion exchange resin for

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32

acetic acid removal, concluded that the membrane exhibited better performance

in terms of dimensionless throughput and product loss. The membrane binds

acetic acid more efficiently than that of resins and the total volume of waste

water will be less for a membrane based system compared to resins. The use of

zeolites had also been reported to improve the ethanol yield during fermentation

by removing fermentation inhibitors with almost no loss of fermentable sugars

(Ranjan et al., 2009).

As another method, activated charcoal adsorption has gaining considerable

attention. The charcoal removed the furfural derivatives efficiently, but do not

have similar impact on acetic acid. Miyafuji et al. (2003) reported that since,

many furans and phenolics compounds are hydrophobic, thus the wood

charcoal prepared at higher temperature can remove the inhibitory compounds

more effectively compared to the wood charcoal than at lower temperature.

Gupta et al. (2009) reported the removal of HMF (38.24%), furfural (29.31%),

acetic acid (45.26%) and caffeic acid (74.92%) from the acid hydrolysate of

Prosopis juliflora with activated charcoal. Similar results have been reported in

the work of Kuhad et al. (2010b) and Gupta et al. (2011b).

2.5.2.2. Chemical detoxification

Chemical methods for detoxification include precipitation of toxic compounds

and ionization of some inhibitors under certain pH values, the latter being able

to change the degree of toxicity of the compounds (Van Zyl et al., 1998; Roberto

et al., 1991; Martinez et al., 2001; Mussatto, 2002).

Detoxification of lignocellulosic hydrolysates by alkali treatment, i.e., increasing

the pH to 9–10 with Ca(OH)2 (overliming) and readjustment to 5.5 with H2SO4,

has been described as early as 1945 by Leonard and Hajny. After an overliming

treatment (pH 10), causing the formation of a large precipitate, the ethanol

productivity was further increased. The detoxifying effect of overliming is due

both to the precipitation of toxic components and to the instability of some

inhibitors at high pH. This has been demonstrated by the fact that pre-

adjustment to pH 10 with NaOH of a strongly inhibiting dilute-acid hydrolysate

of spruce prior to fermentation resulted in twice as high ethanol yield (and

comparable to the yield in a reference fermentation containing glucose and

nutrients) as after only adjustment to fermentation pH (5.5) (Palmqvist, 1998).

Martinez et al. (2001) reported that using Ca(OH)2 to adjust the pH of sugarcane

bagasse hemicellulose hydrolyzate (overliming treatment) to 9.0, proved to be a

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33

very efficient detoxification method. Purwadi and coworkers (2004) overlimed

the detoxified hydrolysates of forest residues to various pH ranging from 9.0 to

12.0 using Ca(OH)2 followed by readjustment of the pH to 5.0. They observed

that increasing the pH, time and/or temperature resulted in more effective

degradation of furans and resulted in better fermentability for the hydrolysates.

Similar observations were reported by other researchers as well (Alriksson et al.,

2006, Chandel et al., 2007; Gupta et al., 2009; Kuhad et al., 2010a).

2.5.2.3. Biological detoxification

Biological method of detoxification involved the use of specific enzymes or

microorganisms that act on toxic compounds present in the hydrolysates and

change their composition. The enzymes used commonly for detoxification are

oxidases and dehydrogenases. The enzyme peroxidases and laccase from white

rot fungi are most commonly used for the removal of phenolics while the

furfuryl dehydrogenase was used for the furans. The detoxification mechanism

of these enzymes probably involves oxidative polymerization of low molecular

weight compounds or the enzymatic transformation to the non-toxic forms.

These enzymes showed significant improvement in enhancing the sugar

consumption and ethanol yields. Johnsson and coworkers (1998) used laccase

and peroxidase enzyme of white rot fungus Trametes versicolor to detoxify the

willow wood hydrolysates, which resulted in approximately 2-3 enhanced

ethanol productivity due to the action removal of phenolics compounds. Similar

observations were reported by other groups as well (Schneider, 1996; Palmqvist

et al., 1997; Silva and Roberto, 2001; Chandel et al., 2007). The use of

microorganisms has also been proposed to selectively remove inhibitors from

lignocellulosic hydrolysates. In similar aspects, Koopman et al. (2010)

elucidated the degradation pathway of the most hazardous inhibitors, Furfural

and HMF in the bacterium Cupriavidus basilensis HMF 14, which has the

ability to metabolize furans. They concluded that the non-specific

dehydrogenases abundant in the cell could participate in the conversion, given

that the necessary activity NAD dependant furfuryl dehydrogenase is present.

2.5.3. Enzymatic hydrolysis

Enzymatic hydrolysis of cellulose is carried out by cellulase enzymes which are

highly specific. The products of the hydrolysis are usually reducing sugars

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34

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– 50°C) 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; Kuhad et al., 2011c). Fungi that have been reported to produce

cellulases include Sclerotium rolfsii, P. chrysosporium and species of

Trichoderma, Aspergillus, Schizophyllum, Fusarium and Penicillium (Sternberg,

1976; Fan et al., 1987; Duff and Murray, 1996; Kuhad et al., 1999; Sun and

Cheng, 2002; Kuhad et al., 2011c). Of all these fungal genera, Trichoderma has

been most extensively studied for cellulase production.

The cellulase system contains of three major enzyme components:

endoglucanase (EG; EC 3.2.1.4), cellobiohydrolase (CBH; EC 3.2.1.91), and β-

glycosidase (EC 3.2.1.21). Evidence suggests that these enzymes act

synergistically (Din et al., 1994; Teeri et al., 1998; Boraston et al., 2004; Gupta

et al., 2009). The exoglucanase (CBH) act on the ends of the cellulose chain and

release β-glucosidase as the end product; endoglucanase (EG) 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) (Figure 2.4).

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). Initially, amorphous non-crystalline regions are attacked,

because they are more accessible and easier to be hydrolyzed. After the initial

hydrolysis stage, as the percentage of crystalline regions in the substrate rises,

the enzymatic hydrolysis rate falls rapidly. Such observed kinetic behavior

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35

correlates to the high-crystalline nature of the bulk of cellulose fraction, and

inaccessibility of the glycosidic bonds is the key rate-limiting factor for the low

enzymatic hydrolysis. Besides the recalcitrance of the substrate, there are a few

factors that also limit cellulase efficiency during the hydrolysis process, which

include end-product inhibition, thermal deactivation of the native protein, non-

specific binding to lignin (Yang and Wyman, 2004), and irreversible adsorption

of the enzymes to the heterogeneous substrate (Taniguchi et al.,2005).

Mechanistically, the reactions catalyzed by cellulases are suggested to involve

general acid-base catalysis by a carboxylate pair at the enzyme active site. One

residue acts as a general acid and protonates the oxygen of the O-glycosidic

bond; at the same time, the other residue acts as a nucleophile. Depending on

the distance between the two carboxylic groups, either inverting (~10 Å

distances) or retaining (~5 Å-distances) mechanisms are observed in cellulases

(Withers, 2001).

Figure 2.4: Schematic representation of enzymatic saccharification of

cellulose. (Source: Kuhad et al. 2011)

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36

Besides cellulases, hemicellulases are another group of polysaccharide

degrading enzymes that are specific to the hemicellulose substrate. As a

heterogeneous, branched polymer, hemicelluloses require enzyme activities

specific to as many as 21 different bonds (Collins et al., 2005; Polizeli et al.,

2005; Shallom and Shoham, 2003). Thus, a consortium of hemicellulases is

needed for a complete breakdown of hemicellulose. Endo-1,4-xylanases are

needed for hydrolyzing the backbone 1,4-β-linked xylose residues; acetylxylan

esterases participate in cleavage of the acetyl ester bonds; and β-D-xylosidases

are utilized for the hydrolysis of xylan oligomers through exo-type attack

(Kapoor et al., 2008). Similar to cellulases, xylanases have separate catalytic

domains and carbohydrate binding modules (CBMs) (Divne et al., 1998; Teeri,

1997; Teeri et al., 1998). Synergism and concerted action among xylanases

enhance the effectiveness in hetero-polymeric xylan hydrolysis. It has also been

reported that β-xylosidases remove the short-chain oligosaccharides,

minimizing the end product inhibition of exoxylanases; thus, overall xylan

hydrolysis efficiency is increased (Polizeli et al., 2005; Hu et al., 2008). Also, by

adding acetylxylan esterases, acetic acids will be liberated and a less acetylated

xylan is exposed for greater accessibility to endoxylanase action (Polizeli et al.,

2005).

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 reusage of the enzymes is also an attractive

methodology to reduce enzymatic hydrolysis costs (Singh et al., 1991; Ramos et

al., 1993; Gregg et al., 1998; Lee et al., 1995; 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

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37

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.

2.5.4. Fermentation

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) hydrolysates is a

complex process. There are two major streams of sugars i.e., pentose-rich sugar

syrup and hexose rich sugars coming from hemicellulose and cellulose

separately. The general requirements of an organism for efficient ethanol

production from lignocellulosic hydrolysate is that it should give a high ethanol

yield, a high productivity, high tolerance against inhibitors, able to ferment at

low pH and be able to withstand high ethanol concentrations.

2.4.4.2. Pentose fermentation

During the last three decades, a number of laboratories have demonstrated the

utilization of pentose sugars by various yeasts, fungi, and bacteria for the

production of alcohols and other fermentation products (Table 2.3).

Table 2.3: List of microorganisms that can ferment pentose sugars

Bacteria Reference Fungi and Yeasts Reference

Aeromonas hydrophila Singh and Mishra, 1993 Candida boidinii Vandeska et al., 1996

Bacillus macerans Dien et al., 2003 Candida shehatae Abbi et al., 1996a

Bacillus polymyxa Singh and Mishra, 1993 Fusarium oxysporum Jeffries and Jin, 2004

Bacteriodes Polygramatis Patel, 1984 Mucor corticolous Millati et al., 2005

Clostridium acetobutylicum El Kanouni et al., 1998 Mucor hiemalis Millati et al., 2005

Clostridium Thermosellum Herrero and Gomez, 1980 Mucor indicus Millati et al., 2005

Escherichia coli Yomano et al., 1998 Neurospora crassa Deshpande et al., 1986

Klebsciella oxytocea Ingram et al., 1999 Pachysolen tannophilus Schneider et al., 1981

Lactobacills pentosus Chaillou et al., 1999 Paecilomyces sp NF1 Mountfort et al., 1991

Lactobacillus casei Roukas and Kotzekidou,

1998

Pichia stipitis Gupta et al., 2009

Lactobacillus pentoaceticus Chaillou et al., 1999 Rhizopus orizae Millati et al., 2005

Lactobacillus plantanum Sreenath et al., 1999

Lactobacillus xylosus Sreenath et al., 1999

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38

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 orizae, Fusarium oxysporum

and Neurospora crassa (El Kanouni et al., 1998; Chaillou et al., 1999; Screenath

et al., 1999; Dien et al., 2003; Millati et al., 2005; Ruiz et al., 2007; Vasudevan

et al., 2007; Hahn-Hagerdal et al., 2007). To date several studies have been

carried out for the fermentation of xylose rich hydrolysates from various

lignocellulosic materials and a summary of recent studies is shown in table 2.4.

Moniruzzaman (1995) achieved a theoretical ethanol yield of 78% during the

fermentation of enzymatic hydrolysate of steam exploded rice straw, however, a

2 to 3 h lag due to diauxic phenomenal metabolic shift from glucose to xylose

was also observed. While, in another study, using the acid and the auto-

hydrolysate of rice straw, C. shehatae NCIM 3501 showed enhanced ethanol

production in auto- hydrolysate (23.1 g/L) than in acid hydrolysate (20.0 g/L)

because of lower inhibitors concentration (Abbi et al., 1996a). Although, the

pentose fermentation process does not require intensive aerobic fermentation

because of high cell mass synthesis, low ethanol yields and higher aeration

energy consumption. However, aeration is required for the biomass production,

which was a major problem during the fermentation of non-detoxified

hydrolysate. Interestingly, the fermentation of non-detoxified corn stover

hydrolysate at higher aeration improved the ethanol production which was due

to the higher xylose consumption translating higher biomass concentration

(Agbogbo et al., 2007). In a similar study the degree of aeration showed a

prominent effect on xylose utilization, ethanol production and xylitol

minimization during the fermentation of membrane treated sugar maple

hydrolysate using P. stipitis NRRL Y-7124 (Stoutenberg, 2008). Further to

enhance the ethanol production, different detoxification strategies were used by

various researchers (Chandel et al., 2007a). The removal of toxic inhibitors from

fermentation broth significantly improved the ethanol yield (2.4- fold) and

productivity (5.7- fold), compared to neutralized hydrolysate. Similarly, the

fermentation of sugarcane bagasse acid hydrolysate with C. shehatae NCIM

3501 showed maximum ethanol yield (0.48 g/g) from ion exchange treated

hydrolysate, followed by activated charcoal (0.42 g/g), laccase (0.37 g/g),

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39

overliming (0.30 g/g) and neutralized hydrolysate (0.22 g/g) (Abbi et al., 1996a).

While in another study, the sequential application of overliming with sodium

sulfite addition resulted in maximum ethanol yield and productivity of 0.32 g/g

and 0.065 g/L/h, respectively (Okur and Sarcaglu, 2008).

Table 2.4: Summaries of the recent work on lignocellulose hydrolysate

fermentation using non-recombinant strains of microorganisms.

Substrates Organism Sugar

(g/L)

Ethanol

(g/L)

Ethanol Yield

(g/g)

Reference

Corn cob P. stipitis 30 10.4 0.34 Saracoglu and Arslan, 2000

Cashew apple bagasse S. cerevisiae 50 20 0.4 Rocha et al., 2009

Cassava waste S. cerevisiae 38 8.73 0.23 Raman and Pothiraj, 2008

Corn stover P. stipitis 40 15.92 0.4 Zhu et al., 2009

Corn stover P. stipitis 60 25 0.42 Agbogbo et al., 2006

Corn stover P. stipitis 40 15 0.37 Agbogbo and Wenger, 2007

Newspaper S. cerevisiae 14.64 5.64 0.39 Kuhad et al., 2010b

Lantana camara S. cerevisiae 36.5 17 0.48 Kuhad et al., 2010c

Poplar P. stipitis 39 12 0.31 Fenske et al., 1998

Prosopis juliflora P. stipitis 18 7.1 0.39 Gupta et al., 2009

Prosopis juliflora S. cerevisiae 40 18 0.45 Gupta et al., 2009

Red oak spent sulfite liquor P. stipitis 49 20.2 0.41 Nigam et al., 2001b

Red oak wood chips P. stipitis 36 14.5 0.4 Nigam et al., 2001a

Rice straw P. stipitis 15 6 0.4 Moniruzzaman, 1995

Rice straw C. shehatae 20 9 0.45 Abbi et al., 1996a

Rice straw P. stipitis 33 14.9 0.45 Huang et al., 2009

Secondary fiber fines S. cerevisiae 5.4 2.12 0.39 Jeffries and Schartman, 1999

Sugar cane bagasse C. shehatae 30 8.67 0.29 Chandel et al., 2007a

Sugar cane bagasse P. tannophilus 63.5 19 0.34 Cheng et al., 2008

Sugar cane bagasse S. cerevisiae 79 30 0.38 Vasquez et al., 2007

Sugar maple P. stipitis 35 12.4 0.35 Stoutenberg et al., 2008

Sun flower seed hull P. stipitis 34 11 0.32 Okur and Saracoglu, 2008

Sun flower stalks S. cerevisiae 40 17.2 0.43 Sharma et al., 2004

Switch grass P. stipitis 39 14 0.36 Fenske et al., 1998

Wheat straw P. stipitis 52 22.3 0.43 Nigam et al., 2001c

Review of literature

40

Several strategies have been employed for the enhanced conversion of pentose

sugars into ethanol. In an attempt to ferment glucose and xylose

simultaneously, Grootjen and coworkers used a co-culture cultivation of

alginate immobilized S. cerevisiae and P. stipitis in a conventional bioreactor,

where, the P. stipitis cells inoculum were taken in comparatively higher amount,

which allowed more xylose utilization under anaerobic conditions and the

fermentation appears to be simultaneous (Grootjen et al., 1990). Furthermore,

the modified stirred tank reactor (STR) system equipped with two teflon-made

HPLC filters air diffusers with improved mixing and less shearing during co-

culture strategy improved the ethanol yield up to 80 % with calcium alginate

immobilized P. stipitis and S. cerevisiae (De Bari et al., 2004). Interestingly, an

agar sheet sandwiched between two chambered bioreactor has also been used

for the co-immobilization of S. cerevisiae and C. shehatae during the mixed

sugar (glucose and xylose) fermentation, however, the cell proliferation in the gel

clogged the microporous membrane which in turn limited the mass transfer

(Lebeau et al., 2007). In another study to overcome the problem of glucose

catabolite repression, sieve plates adjusted STR with a movable device was used

for the coculture of immobilized Zymomonas. mobilis and free P. stipitis to

improve the fermentation efficiency (Fu et al., 2009). Recently, a calcium

alginate immobilized recombinant S. cerevisiae strain ZU-10 hasve been used

for the fermentation of detoxified corn stover hemicellulosic hydrolysate, which

showed the consumption of more than 92 % xylose with an enhanced ethanol

yield and productivities with higher tolerance to fermentation inhibitors (Zhao

and Xia, 2010).

Despite various improvements of these amendments, it has been observed that

the repeat culture with the same batch of immobilized microorganism under the

same conditions resulted in decreased performance (Fu et al., 2009).

Alternatively yeast cells immobilized by self flocculation have shown many

advantages such as no requirement of support matrix, maintained biomass and

enhanced ethanol tolerance. Moreover flocculated yeast cells can be recovered

by sedimentation from fermentation broth. However, CO2 bubbles produced

during ethanol fermentation can alter the settling zone and disturb the

sedimentation of yeast floc whereas a specially designed baffle can overcome

this problem (Zhao and Bai, 2009). There are also some reports of yeast cells

immobilization for biocatalysts development for simultaneous saccharification

and fermentation (SSF). Fujita and coworkers constructed a yeast-based whole-

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41

cell biocatalyst displaying T. reesei xylanase II on the cell-surface and showed

xylan degradation by recombinant cells (Fujita, 2002). Whereas, intensive

attempts are have also been made for the development of yeast cells displaying

cellulase activities on the cell surface so to decrease the usage of exogeneously-

added cellulase use (Tsai et al., 2009; Jeon et al., 2009).

A high -productivity system that involved a membrane bioreactor with cell

recycling of Z. mobilis ZM4 capable of converting both glucose and xylose to

ethanol had been developed (Joachmisthal et al., 2000). Similar strategy was

also applied during the continuous cultivation of a recombinant xylose

fermenting S. cerevisiae TMB 3001 on a xylose-glucose mixture (Roca and

Olsson, 2003). Interestingly, in case of lignocellulose hydrolysate containing

mixed sugars, it was observed that the recycled cells get adapted to the

fermentation inhibitors present in the hydrolysate and showed better results

(Sreenath and Jeffries, 2000). The same was further confirmed by Pruwadi and

coworkers, where continuous cultivation of high cell density flocculating yeast

in toxic dilute acid hydrolysate of spruce residues in a singly and serial

bioreactor adapted to the medium and reduced the requirement of any

detoxification (Purwadi et al., 2007). Recently, a fuzzy optimization of

continuous fermentation with cell recycling for ethanol production was carried

out (Wang and Lin, 2010). From the computational results, the overall

productivity of the continuous fermentation process with cell recycling allowed a

higher dilution rate with 7.3 fold higher productivity (Wang and Lin, 2010).

2.5.4.2. Hexose sugar fermentation

A variety of microorganisms ranging from bacteria, fungi and yeasts are known

to ferment hexose sugars (Table 2.5), however, the most common and efficient

microbes used for hexose fermentation are Saccharomyces cerevisiae and

Zymomonas mobilis (Hahn-Hagerdal et al., 2007). 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

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42

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). Recently our group has 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 (Gupta et al.,

2009; Kuhad et al., 2010a).

Table 2.5: Microorganisms fermenting hexose sugars to ethanol

Organisms Reference Organisms Reference

Candida shehatae Abbi et al., 1996a Rhizopus miehei Millati et al., 2005

Fusaruium sporium Mamma et al., 1995 Rhizomucor pusillis Millati et al., 2005

Kloeckera apiculata Aguilera et al., 2006 Saccharomyces cerevisiae Kuhad et al., 2010b

Kluyeromyces marxianus Ballesteros et al., 2004 S. bayarus Belloch et al., 2008

Mucor indicus Abdenifar et al., 2009 S. paradoxus Belloch et al., 2008

M. hiemalis Millati et al., 2005 S. kudriazevii Belloch et al., 2008

M. corticolous Millati et al., 2005 S. cariocanus Belloch et al., 2008

Neurospora crassa Mamma et al., 1995 S. mikatae Belloch et al., 2008

Pachysolen tannophillus Abbi et al., 1996a S. pastorianus Belloch et al., 2008

Pichia stipitis Gupta et al., 2009 Schizosaccharomyces pombe Hu et al., 2005

Pichia membranifaciens Aguilera et al., 2006 Terulospora delbruecki Aguilera et al., 2006

Rhizopus oryzae Abdenifar et al., 2009 Zymomonas mobilis

Besides the enzymatic hydrolysates, several studies have been carried out to

use the S. cerevisiae to utilize the hexose sugars present in the acid hydrolysate

of lignocellulosic substrates. Brandberg and coworkers (2005) fermented a non-

detoxified dilute acid wood hydrolysate using S. cerevisiae ATCC 96581 under

continuous mode (with or without cell recycling) and observed that, 99% of the

sugars were converted at a cell concentration of 6 g/L leading to an ethanol

concentration of 17 g/L and a productivity of 1.6 g/L/h. To improve the

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43

productivity, three different types of cell retention process i.e., cross-flow

filtration with recirculation, sedimentation, and immobilization in calcium

alginate were evaluated He observed that when 75% of the cells were retained

by filtration, the hexose conversion increased to 94%, and the ethanol

production rate was 2.3 g/L/h. Comparable results were achieved with cell

recirculation by a settler and immobilization. When the dilution rate was

increased to 0.2/h, neither filtration nor sedimentation could prevent washout

of the cells. On the other hand, culture immobilization increased the ethanol

productivity to 3.5 g/L/h, although hexose conversion dropped (Brandberg et

al., 2007).

Taherzadeh and his group (2001) used a dilute acid hydrolysate supplemented

with defined mineral media for the fermentation with S. cerevisiae CBS 8066

immobilized in Ca-alginate beads and observed that the 79- 86% of glucose was

consumed with 0.45 and 0.47 g/g ethanol yield. Further to improve the ethanol

productivity, the same group immobilized S. cerevisiae cells in different bio-

processes such as CSTR, fluidized bed bioreactor (FBBR) and combined CSTR

and FBBR (Purwadi and Taherzadeh 2008). In CSTR operated at dilution rates

between 0.22 and 0.86/h, conversion of an initial glucose concentration

decreased from 100% to 77%, but productivities increased from 2.1 to

6.6 g/L/h. If fluidized bed bioreactor (FBBR) was connected to the CSTR,

glucose conversion was higher than 99%. In contrast, a single FBBR gave a

higher glucose conversion of 92% at a dilution rate of 0.86/h and a higher

ethanol productivity of 7.4 g/L/h (Purwadi and Taherzadeh, 2008).

Besides entrapment, encapsulated S. cerevisiae cells were also used for

fermentation, as encapsulation has the advantages of checking the diffusion of

nutrients as well as the cell leakage. Talebnia and Taherzadeh (2006)

encapsulated S. cerevisiae CBS 8066 cells and observed that ethanol

productivity increased with increasing dilution rate from 1.1 to 4.2 g/L/h, and

cell viability was high under all conditions ranging between 77% and 90%. The

flocculating S. cerevisiae strain has also been used for the fermentation of acid

hydrolysates (Tang et al., 2006; Purwadi et al., 2007).

As an additional approach, simultaneous saccharification and fermentation

(SSF) has also been employed for improved ethanol production. In the SSF

process, the stages are virtually the same as in separate hydrolysis and

fermentation (SHF) systems, except that both are performed in the same

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44

reactor. It has been shown that SSF reduces the processing time, which in turn

leads to increase in the production of ethanol (Ballesteros et al., 1991; Wyman,

1994; Alfani et al., 2000; Soderstrom et al., 2005). During the SSF process, the

liberated glucose is quickly converted to ethanol by fermenting microorganism,

the inhibition of cellulase by the reaction end-products is reduced and this

single fermenter process eliminates a portion of the investment cost by reducing

the number of fermenters required for start-up. In addition, the SSF process

reduces the contamination risk due to the presence of ethanol in the medium

(Banat et al., 1998; McMillan, et al., 1999; Soderstrom et al., 2005; Ohgren et

al., 2007). However, the performance of the SSF process is limited by the

different optimum temperatures for enzymatic saccharification and microbial

fermentation. The optimum temperature for cellulases is usually in the range of

40 to 50°C, while the optimum temperature of ethanol production for the most

common ethanologenic yeast, S. cerevisiae, is between 30 and 37°C (Suryawati

et al., 2008). Hence, the coexistence of two process conditions for microbe and

enzyme hinders the efficacy of process. In order to overcome this notable

problem, thermotolerant yeast strains have been used in high temperature SSF

processes (Suryawati et al., 2008). In such cases, the performance of SSF with

thermotolerant yeast at temperatures closer to the optimum temperatures for

cellulase activity could enhance saccharification, reduce the operation time and

reduce cellulase dosage (Suryawati et al., 2008). Szczdodrak and Targonski

(1988) tested a total of 58 yeast strains belonging to 12 different genera and

capable of growing and fermenting sugars at temperatures of 40–46 ◦C. They

selected several strains belonging to the genera Saccharomyces, Kluyveromyces

and Fabospora in view of their capacity to ferment glucose, galactose and

mannose at 40, 43 and 46 ◦C, respectively. Ballesteros and colleagues (1991)

studied different treatments to improve the thermotolerance of some species

belonging to the genera Saccharomyces, and Kluyveromyces and the best results

were obtained with Kluyveromyces marxianus LG. Bollók and coworkers (2000)

also used Kluveromyces strain in SSF experiments of softwood and almost 70 %

ethanol conversion was obtained. Harikrishna et al. (2001) have reported final

ethanol concentrations of 2–2.5% (w/v) in 72 h SSF of lignocellulosic wastes

with thermotolerant yeast at 10% (w/v) initial substrate concentration. While,

Ballesteros and coworkers (2004) used the mutated K. marxianus strain CECT

10875 and achieved a SSF yield of 50 to 72 % in 72-82 h using various

lignocellulosic feedstocks (poplar and eucalyptus, Sorghum sp. bagasse, wheat

Review of literature

45

straw and Brassica carinata residue). Further, Ohgren and colleagues (2007)

and coworkers compared the SHF and SSF in steam exploded corn stover and

observed that SSF gave a 13% higher overall ethanol yield than SHF. Recently, a

respiratory-deficient mutant of the thermotolerant yeast Candida glabrata

(Cgrd1), was subjected to ethanol production by high-temperature SSF under

aerobic conditions to achieve maximum ethanol (17.0 g/L) within 48 h at 66.6%

of its theoretical yield and with 0.35 g/L h productivity (Watanabe et al., 2010).

2.4.5. 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 78°C whereas water vaporizes at 100°C), 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 adding

benzene or cyclohezane to the mixture. When these components are added to

the mixture, it forms a heterogeneous azeotropic mixture in vapour-liquid-liquid

equilibrium, which when distilled produces anhydrous ethanol in the column

bottom, and a vapour mixture of water and cyclohexane/benzene. While in

extractive distillation, consists of adding a ternary component, which will

increase the ethanol‟s relative volatility. When the ternary mixture is distilled, it

will produce anhydrous ethanol on the top stream of the column. The third

method uses molecular sieves to remove water from ethanol. In this process,

ethanol vapour pressure pass through a bed of molecular sieve beads. The

bead‟s pores are sized to allow absorption of water while excluding ethanol.

Recently, distillation followed by molecular sieve dehydration operations have

been used to recover a pure ethanol product of fuel-grade (>99.5%). Molecular

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46

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. In the past 40 years, installation of multistage,

pressure distillation systems have reduced energy consumption by 50%,

compared to the earlier all-atmospheric pressure systems (Madson, 2003). In

modern molecular sieve dehydration systems, “pressure swing adsorption” is

employed to remove the water content, where a relatively high pressure is

applied at the water removal stage and a relatively low pressure is applied at the

desiccant regeneration stage. Therefore, operation temperature can be kept

almost constant, and the heat of adsorption can be effectively stored and further

supplied to the regeneration stage (Swain, 2003). 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 biorenewable, 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).

2.5. Strain Improvement for ethanol fermentation

2.5.1. Mutation

There are several reports where mutagenised recombinant strains showed

enhanced ethanol production over their parent strains (Wahlbom et al., 2003;

Watanabe et al., 2005; Kim et al., 2007; Liu and Hu, 2010). In an early report, a

recombinant S. cerevisiae strain TJ1 mutagenised with ethyl methane sulfonate

(EMS) was found to have lower xylose reductase (XR) activity but high Xylitol

dehydrogenase (XD) and xylulokinase (XKS) activities than the parent strain,

which in turn resulted in 1.6 fold increase in ethanol production (Tantirungkij et

al., 1993). Further, a mutant of S. cerevisiae TMB 3001 capable of utilizing

xylose under anaerobic condition was developed by sequential EMS

mutagenesis and adaptation of the mutant strain under microaerobic and

anaerobic conditions (Sonderegger and Sauer, 2003). Similar strategy was used

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47

to develop two EMS mutagenised S. cerevisiae strains 3399 and 3400 with

improved growth on xylose (Wahlbom et al., 2003). Since the most efficient

glucose utilizing microbes are not able to metabolize the pentoses anaerobically,

therefore strategies like natural selection and random mutations were also

tested (Matushika et al., 2009a). Besides EMS, several other mutagens had also

been used to obtain mutants derepressed for pentose metabolism. Sreenath and

Jeffries (2000) used 2-deoxyglucose (2-DOG) mutated strain, which showed

considerable improvement in xylose utilization. While in another report, a UV

mutagenised P. stipitis NRRL Y-7124 strain was found to produce higher ethanol

than the wild strain (Bajwa et al., 2009).

Site-directed mutagenesis is another efficient approach used to obtain the

mutants for better xylose fermentation. Watanabe and coworkers (2005) used

multiple site- directed mutagenesis of the NAD+-dependent XDH from P. stipitis

and introduce a structural zinc atom for the complete reversal of the coenzyme

specificity. The selected mutants were found to exhibit significant

thermostability and enhanced catalytic activity with NADP+. Similarly, several

PsXDH mutants were generated with complete reversal of coenzyme specificity

toward NADP+ by multiple site-directed mutagenesis within the coenzyme-

binding domain and with increased thermostability by refining the structural

zinc-binding loop without affecting their activities (Annaluru et al., 2007). In

addition one of these S. cerevisiae mutant (MA-R5) under the control of a strong

constitutive promoter showed particularly high ethanol production from xylose

and low xylitol yield by fermentation of not only xylose as the sole carbon

source, but also a mixture of glucose and xylose (Watanabe et al., 2007;

Matushika et al., 2008; 2009b). Additionally, an ethanologenic E. coli mutant

that is, devoid of foreign genes, has also been developed by combining the

activities of pyruvate dehydrogenase and the fermentative alcohol

dehydrogenase and the mutant was found able to ferments glucose or xylose to

ethanol with 82% ethanol yield under anaerobic conditions (Kim et al., 2007).

2.5.2. Protoplast Fusion

Protoplast fusion provides characteristic advantage such as promotion of high

frequencies of genetic information between organisms for which poor or no

genetic exchange has been demonstrated or which are genetically

uncharacterized (Heluane et al., 1993; Lin et al., 2005; Pasha et al., 2007). In

the presence of a fusogenic agent such as polyethylene glycol (PEG), protoplasts

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48

are induced to fuse and form transient hybrids or diploids. Several reports on

protoplast fusion between pentose and hexose utilizing yeasts showed efficient

utilization of both sugars with higher biomass yield. Heluane and coworkers

(1993) transferred the genes of xylose utilization from P. tannophilus to S.

cerevisiae. The hybrids reassembled the S. cerevisiae parent morphologically but

displayed the ability to use the pentose sugars (xylose) similar to P. tannophilus.

The same has been supported by other workers, where a fusant of

Schizosaccharomyces pombe and Lentinus edodes were found to utilize xylan as

carbon source (Lin et al., 2005). In an another study, the protoplasts of

thermotolerant S. cerevisiae and mesophilic xylose-utilizing C. shehatae were

fused by electrofusion and the fusant yeast gave an ethanol yield of

approximately 0.459 g/g with productivity of 0.67 g/L/h and fermentation

efficiency of 90% and showed higher temperature tolerance up to 40°C as well

(Pasha et al., 2007). Moreover, using a combinatorial approach, a xylose

fermenting fusant (F6) of C. shehatae and S. cerevisiae was developed, which

showed improved ethanol production (28 %) than its parental strain (Li et al.,

2008). In this strategy the C. shehatae was first adapted for ethanol tolerance

and then mutagenised the adapted strain by UV irradiation and thus selected a

respiration deficient mutant RD-5 was selected. Further, the protoplasts of RD-

5 and S. cerevisiae were fused and the resultant fusant strain F6 showed 28%

higher ethanol production than the parent Candida shehatae stain, with the

production level of 18.75 g/L from 50 g/L xylose. Recently a strategy of genome

shuffling was also used, in which the genomes of 6 UV mutagenised P. stipitis

strains (WT, Ps302, GS301, GS302, GS401 and GS402) were shuffled and after

the third and fourth rounds of genome shuffling, putative improved mutant

colonies were pooled, re-grown and spread on hardwood spent sulphite liquor

(HW SSL) gradient plate (Bajwa et al., 2010). Of these, 2 mutants (GS401 and

GS402) from the fourth round could grew in 80% (v/v) HW SSL, and another 2

mutants (GS301 and GS302) from the third round could grow in 85% (v/v) HW

SSL. While, P. stipitis WT and PS 302 could not grow in any of the HWSSL

concentrations. Thus the mutated strains showed improved inhibitors tolerance

against HWSSL hardwood spent sulphite liquor (Bajwa et al., 2010).

2.5.3. Adaptation

Fermentation of wood-derived hydrolysates is problematic because of the toxic

inhibitors released during thermo-chemical hydrolysis, however, the adaptation

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49

approach can be an alternative means to improve the microbial strains (Yomano

et al., 1998; Agbogbo et al., 2008; Zhu et al., 2009). There are several reports on

enhancement of ethanol yield and productivity using adapted strains of P.

stipitis and C. shehatae for the fermentation of undetoxified or partially

detoxified hydrolysates (Liu et al., 2004; Attfield and Bell, 2006; Zhu et al.,

2009). For instance, an ethanologenic yeast when adapted against inhibitors by

repeated sub-culturing in a medium with furfural and HMF up to a

concentration of 10-20 mM was found to grow more efficiently than its parent

strain in the presence of inhibitors (Liu et al., 2004). While, another strategy of

natural selection and breeding was used to develop non-recombinant strains of

S. cerevisiae that could grow efficiently on xylose (Attfield and Bell, 2006). By

breeding and natural selection over 23 mating cycles and 1463 selection days, a

non-genetically modified S. cerevisiae (MBG-2303) was obtained, which grew

aerobically on xylose and demonstrated 57 fold higher biomass production than

the control strain (Attfield and Bell, 2006). Later on, it was demonstrated that

adaptation of P. stipitis CBS 6054 on solid agar produced more ethanol (19.4

g/L) than liquid adapted (18.4 g/L) and unadapted strains (16.3 g/L) (Agbogbo

et al., 2008). Recently, studies were carried out on adaptation of P. stipitis CBS

5776 strain which on fermentation of steam exploded prehydrolyzate of corn

stover showed improved ethanol yield of 15.92 g/L with 80.34% theoretical yield

(Zhu et al., 2009).

Moreover, the evolutionary adaptation approaches have also been applied to

recombinant strains to improve their fermentation capability. Lawford and his

group improved the xylose-fermenting recombinant strains of Z. mobilis 39767

to tolerate higher concentration of acetic acid by subculturing in a medium

containing 10-50 % of hydrolysate and the adapted isolates demonstrated a

significant improvement in ethanol productivity compared to un-adapted strains

(Lawford and Rousseau, 1999). Similarly, an engineered E. coli KO11 was

developed to tolerate high ethanol concentration using a long term adaptation

strategy of alternative serial selections for liquid and solid medium. The mutants

(LY01, LY02 and LY03) demonstrated more than 50 % survival rate in 10 %

ethanol (0.5 min exposure) and also decrease in fermentation time (Yomano et

al., 1998). In almost all previous efforts of evolutionary adaptation the organism

was first subjected to genetic engineering, which was followed by adaptive

selection (Sonderegger and Sauer, 2003; Kuyper et al., 2005; Wisselink et al.,

2009). However, recently a new strategy consisting genetic engineering,

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50

mutation with EMS followed by two-step evolutionary adaptation (under

sequential aerobic and oxygen limited conditions) has also been attempted (Liu

and Hu, 2010). The strain thus developed showed four fold increase in its

specific growth rate compared to the parental strain. Interestingly the activity of

critical enzymes of xylose metabolism (XR, XDH and XK) remain unchanged

suggesting that chemical mutagenesis and evolutionary adaptation might have

created a new genetic traits making the mutants capable of xylose metabolism

in the favor of xylose metabolism (Liu and Hu, 2010).

2.5.4 Genetic Manipulation for improved ethanol fermentation

Substantial progress in the genetic engineering of different microbes for the

conversion of xylose or pentose sugars to ethanol has been achieved (Karhumaa

et al., 2005; Matushika et al., 2008; Liu and Hu, 2010; Kuhad et al., 2011b).

Although the genetically engineered host strains of bacteria and yeast showed

tremendous improvement in final ethanol yields and efficient utilization of

pentose sugars, but the information about the usage of genetically modified

organisms for large scale pentose fermentation is scarcely available. The potent

recombinant microbes are listed in table 2.6.

2.5.4.1. Genetic engineered Saccharomyces cerevisiae

S. cerevisiae produces ethanol from hexose sugars but cannot ferment xylose or

arabinose, however, the yeast is able to metabolize the xylose isomer, xylulose

and recombinant DNA technologists have taken advantage of this in creating

xylose-fermenting strains.

Ho and coworkers (1998) were the first to produce successfully a recombinant S.

cerevisiae strain capable of effective xylose fermentation and xylose and glucose

co-fermentation, where plasmids with XR and XDH genes from P. stipitis and

XKS gene from S. cerevisiae were transformed into S. cerevisiae for the co-

fermentation of glucose and xylose. Similar strategy for improved xylose

utilization and ethanol production from have also been reported by other groups

(Karhumaa et al., 2007; Matushika et al., 2008; Matushika and Swayama, 2008;

Vleet and Jeffries, 2009).

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51

Table 2.6: List of pentose utilizing recombinant Yeasts and Bacterial strains

Strain Sugar used

(g/L)

Ethanol

production

(g/L)

Ethanol yield

(g/g)

Ethanol

productivity

(g/L/h)

Reference

E. coli KO11 80 X 41.6 102 0.87 Ohta et al., 1991

E. coli KO11 90 X 41 89 0.85 Yomano et al., 1998

E. coli KO11 140 X 59.5 Yomano et al., 1998

E. coli FBR5 95 X 41.5 90 0.59

E. coli FBR5 A:X:G

15:30:30 34.0 90 0.92 Dien et al., 2000

E. coli LY01 140 X 63.2 88 0.66 Yomano et al., 1998

K. oxytoca M5A1 100 X 46 95 0.96 Ohta et al., 1991

K. oxytoca P2 A:X:G

20:40:20 34.2 84 0.35 Bothast et al., 1994

Z. mobilis AX101 A:G:X

20:40:40 42 84 0.61 Mohagheghi et al., 2002

Z. mobilis CP4 25 X 11.0 86 0.57 Zhang et al., 1995

Z. mobilis CP4 G:X

65:65 24.2 95 0.81 Zhang et al., 1995

Z. mobilis CP4 60 X 23 94 0.32 Lawford and Rousseau, 1999

Z. mobilis ZM4 G:X

65:65 62 90 1.29 Joachimsthal et al., 1999

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52

Z. mobilis ATCC 39767 G:X:A

30:30:20 33.5 82-84 0.82-0.65 Chou et al., 1997

S. cerevisiae F12 G:X

50:50 26 52 NA Sonderegger et al., 2004

S. cerevisiae BH42 G:X

50:50 28 56 NA Sonderegger et al., 2004

S. cerevisiae A4 G:X

50:50 19 38 NA Zaldivar et al., 2002

S. cerevisiae 1400 G:X:A:Gal

31 :15 :10 :2 22 90 0.92 Moniruzzaman et al., 1997

S. cerevisiae 1400 80 X 27 66 1.12 Moniruzzaman et al., 1997

S. cerevisiae 1400 50 X 1.5 6 Ho et al., 1998

S. cerevisiae PRD1 21.7 1.6 14 0.07 Kotter and Ciriacy, 1993

S. cerevisiae TJ1 50 X 2.7 10.6 0.02 Tantirungkij et al., 1993

S. cerevisiae RBW 202-AFX 20 X 8.4 84 NA Kuyper et al., 2004

S. cerevisiae RBW 202-AFX 50 X 19.5 78 NA Madhavan et al., 2008

S. cerevisiae RWB 217 20 X 8.6 86 NA Kuyper et al., 2005

S. cerevisiae H158 80 X 17.6 44 NA Johansson et al., 2001

S. cerevisiae H158 80 X 21.6 54 NA Johansson et al., 2001

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S. cerevisiae H 2673 50 X 11.5 46 NA Verho et al., 2003

S. cerevisiae H 2723 50 X 12 48 NA Verho et al., 2003

S. cerevisiae H 2684 50 X 20.5 82 NA Verho et al., 2003

S. cerevisiae ZU-10 80 X 30.2 75.6 0.50 Zhao and Xia, 2009

S. cerevisiae LEK 122 20 X 2.5 24.5 0.025 Liu and Hu, 2010

S. cerevisiae LEK 122 50 X 6.35 25.4 0.064 Liu and Hu, 2010

S. cerevisiae LEK 513 50 X 8.13 32.5 0.113 Liu and Hu, 2010

S. cerevisiae TMB 3001 10 X 2.4 48 NA Sonderegger and Sauer, 2003

S. cerevisiae TMB 3001 G :X

5 :15 2.5 25 0.15 Eliasson et al., 2000

S. cerevisiae TMB 3001 50 X 15.5 62 0.22 Jeppsson et al., 2002

S. cerevisiae TMB 3001 10 X 4.4 88 0.061 Traff-Bjerre et al., 2004

S. cerevisiae TMB 3008 50 X 19 76 0.27 Jeppsson et al., 2002

S. cerevisiae TMB 3030 50 X 14 56 NA Jeppsson et al., 2003

S. cerevisiae TMB 3037 50 X 17 68 NA Jeppsson et al., 2003

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S. cerevisiae TMB 3050 50 X 14.5 58 NA Karhumaa et al., 2005

S. cerevisiae TMB 3057 50 X 16.5 66 0.13 Karhumaa et al., 2007

S. cerevisiae TMB 3066 50 X 21.5 86 0.073 Karhumaa et al., 2007

S. cerevisiae TMB 3120 10 X 4.6 92 0.064 Traff-Bjerre et al., 2004

S. cerevisiae TMB 3251 50 X 17 68 0.24 Jeppsson et al., 2002

S. cerevisiae TMB 3253 50 X 14 56 NA Jeppsson et al., 2003

S. cerevisiae TMB 3254 50 X 14 56 NA Jeppsson et al., 2003

S. cerevisiae TMB 3255 50 X 20.5 82 0.29 Jeppsson et al., 2002

S. cerevisiae TMB 3256 50 X 18 72 NA Jeppsson et al., 2003

S. cerevisiae TMB 3261 50 X 17 68 NA Jeppsson et al., 2003

S. cerevisiae TMB 3266 10 X 4.6 92 0.064 Traff-Bjerre et al., 2004

S. cerevisiae TMB 3267 10 X 4.1 82 0.057 Traff-Bjerre et al., 2004

S. cerevisiae TMB 3270 50 X 18 72 0.32 Jeppsson et al., 2006

S. cerevisiae TMB 3271 50 X 15.5 62 0.28 Jeppsson et al., 2006

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S. cerevisiae TMB 3400 8 X 1.44 36 NA Wahlbom et al., 2003

S. cerevisiae TMB 3400 20 X 5 50 NA Wahlbom et al., 2003

S. cerevisiae TMB 3400 50 X 17 68 0.12 Karhumaa et al., 2007

P. stipitis FPL UC7 80 X 30.4 76 0.41 Shi et al., 1999

P. stipitis FPL-shi 21 80 X 38.4 96 0.43 Shi et al., 1999

P. stipitis FPL-shi 31 80 X 24.8 62 0.15 Shi et al., 2002

Where,

A= arabinose,

G = glucose,

X= xylose,

Gal = galactose,

M = mannose.

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56

Thereafter to achieve reduction in xylitol formation during xylose fermentation,

recombinant S. cerevisiae strains expressing PsXR and PsXDH and

overexpressing of ScXKS were constructed that lowered the oxidative PPP

activity through the GND1 (6-phosphogluconate dehydrogenase) and ZWF1

genes (glucose- 6-phosphate dehydrogenase) (Jeppsson et al., 2002). These

mutants showed increase in ethanol yield and xylose consumption rate

compared to the parent strain. Alternatively the shift of metabolic flux towards

ethanol formation appeared a significant strategy to improve the intracellular

cofactor concentrations in S. cerevisiae (Bro et al., 2006). The impact of over-

expression of NADH kinase (encoded by the POS5 gene) on glucose and xylose

metabolism in recombinant xylose-utilizing S. cerevisiae has also been studied

(Huo et al., 2009). The expression of NADH kinase in cytosol instead of

mitochondria redirected the carbon flow from CO2 to ethanol during aerobic

growth on glucose, whereas under anaerobic growth the flux directed toward

ethanol and acetate fermentation (Huo et al., 2009).

The approach of protein engineering has also been investigated to reduce xylitol

excretion and enhancing ethanol yield using recombinant S. cerevisiae. Using

this approach an improved ethanol production accompanied by decreased

xylitol formation was achieved in recombinant S. cerevisiae expressing mutated

PsXR (having reduced affinity for NADPH), PsXDH and ScXKS (Jeppsson et al.,

2006). Besides, the heterologous expression of xylose specific transporters in

recombinant S. cerevisiae for improved ethanol production has also been tested.

The SUT1 gene (Weierstall et al., 1999) coding a sugar transporter in P. stipitis,

has been successfully expressed in S. cerevisiae (Katahira et al., 2006).

Moreover, the glucose/xylose-facilitated diffusion transporter and

glucose/xylose symporter from C. intermedia, encoded by Gxf1 and GXxf2 genes

(Leandro et al., 2008) have been expressed in S. cerevisiae (146), where the

recombinant xylose-fermenting S. cerevisiae strain harboring Gxf1 showed

faster xylose uptake and ethanol production (Runquist et al., 2009). Recently, a

combinatorial approach of genetic engineering, chemical mutagenesis and

evolutionary adaptation has been used to improve the xylose utilization. The S.

cerevisiae strain W303-La was introduced with XI and XKS gene from P. stipitis

NRRL7124 to make S. cerevisiae LEK 122. Thereafter, the selected strain was

chemically mutagenised with EMS followed by their evolutionary adaptation for

xylose utilization and growth under oxygen-limited conditions (Liu and Hu,

2010).

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57

2.5.4.2. Genetically engineered Zymomonas mobilis

The xylose utilization in Z. mobilis was developed by integrating XI and XKS from E.

coli, Xanthomonas campestris and K. penumoniae in its genome (Feldman et al., 1992;

Zhang et al., 1995). In another work, the operons encoding xylose assimilation and

pentose phosphate pathway enzymes were transformed into Z. mobilis to generate pZB5

strain for the effective fermentation of xylose to ethanol (Zhang et al., 1995; Ruiz et al.,

2007). Further to construct improved strains with higher ethanol productivities and

yields, pZB5 was transformed into Z. mobilis ethanol producing strain ZM4; ATCC

31821, which showed the capability of converting a mixture of 65g/L of glucose and 65

g/L of xylose to 62 g/L ethanol in 48h with an overall yield of 0.46 g/g (Joachmisthal et

al., 2000). Following the similar approach, another group incorporated five genes of

arabinose utilization from E. coli ara A (coding for L-arabinose isomerase), ara B (coding

for L-ribulokinase), ara D (coding for L-ribulose-5 phosphate-4-epimerase), tal and tkt in

Z. mobilis ATCC 39767 (Deanda et al., 1996). A number of other improvements have

also been made into Z. mobilis strains and the newest strain Z. mobilis AX101 fermentsed

both arabinose, xylose and glucose and carriesd seven necessary recombinant genes as

part of chromosomal DNA (Chou et al., 1997; Lawford et al., 2002; Mohagheghi et al.,

2002). However, these strains showed acetic acid sensitivity (Mohagheghi et al., 2002).

To address the problem of sensitivity to toxic fermentation inhibitors, a new strain of Z.

mobilis ZM4/AcR (pZB5) was developed with increased acetate resistance that has

enhanced performance in the presence of 12 g sodium acetate per L at pH 5 (Jeon et al.,

2009). Recently, in a recent effort, transcriptomic and metabolomic profiles for Z. mobilis

ZM4 under aerobic and anaerobic fermentations have been elucidated using microarray,

high-performance liquid chromatography and gas chromatography-mass spectrometry

(GC-MS) analysis (Yang et al., 2009).

2.5.4.3. Genetically engineered Escherichia coli

The construction of E. coli strains to selectively produce ethanol was one of the

first successful applications of metabolic engineering. Ingram‟s group has done

extensive work on the development of efficient recombinant E. coli strains for

ethanol production. They eliminated the dependence of host on alcohol

dehydrogenase (ADH) activity by combining adh B and pdc (coding for pyruvate

decarboxylase, PDC) genes of Z. mobilis to form pet operon (Ingram et al., 1987;

Beall et al., 1991). Considering a number of factors that influence ethanol

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58

production such as substrate range and growth conditions, E. coli strains ATCC

11303 was chosen as the host for the pet plasmid (Alterthum and Ingram,

1989). Further, to improve the genetic stability, the pet operon was integrated

into the chromosome of ATCC 11303. This strain was further modified by

deletion of the succinate production gene (frd) to prevent the formation of

succinate, a major byproduct of E. coli metabolism. The final strain so developed

strain (KO11) was able to convert glucose and xylose to ethanol at theoretical

yields of 100% in rich media containing ample yeast extract (Ohta et al., 1991).

To further improve pentose fermentation by KO11, a number of spontaneous

mutants defective in glucose transport were selected and two such strains SL28

and SL40 when fermented using individual or mixture of xylose and glucose

produced ethanol more efficiently (20 %) than the parent strain KO11 (Lindsay

et al., 1995). The recombinant E. coli strain was further improved to achieve

better xylose fermentation, the glycolytic flux and the growth rate of

recombinant strain (Gonzalez et al., 2002). Later on to address this problem, a

lactate producing recombinant of KO11 was reengineered for ethanol production

by deleting genes encoding for fermentative routes for NADH and randomly

inserting a promoter-less cassette containing the complete Z. mobilis ethanol

pathway into KO11 chromosome (Attfield and Bell, 2006).

In contrast, Dien and coworkers (2000) developed new ethanologenic strains of

E. coli such as FBR3, FBR4 and FBR5 with plasmid pLOI297 having Z. mobilis

pyruvate to ethanol converting enzymes. Alternatively a homo-ethanologenic

strain of E. coli SE2738 from wild type E. coli K-12 W3110 was also developed;

where the mutant strain exhibited 82 % theoretical ethanol yield when grown on

xylose under anaerobic conditions (Kim et al., 2007). In a recent study a

minimal E. coli cell for efficient ethanol production from hexoses and pentoses

was developed using elementary mode analysis to dissect the metabolic network

into its basic building blocks (Trinh et al., 2008). Later on, using the similar

approach, a glycerol to ethanol converting E. coli strain was designed by

reducing the functional space of central carbon metabolism to a total of 28

glycerol utilizing pathways (Trinh et al., 2009). More recently an attempt has bee

made, E. coli has also been attempted to engineer E. coli for the production of

ethanol from fatty acid feedstocks, resulting in ethanol yield much higher than

the theoretical maximum obtained from sugars (Clomburg and Gonzalez, 2010).

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2.6. Economic evaluation of cellulosic ethanol production

The first detailed technical reports found in the litreature 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 pre-treatment, 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/litre or $3.5/gal. Similarly, another report released by

Chem Systems, Inc. (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/ litre or

$2.06/gal.

Later on, NREL reported the lignocellulose conversion to ethanol following acid

hydrolysis at a cost of ~ $0.05 per litre or $ 0.20 per gallon 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 per litre ethanol or 0.3-0.5 per gallon 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/litres or 10 cents per gallon of ethanol,

the cost of ethanol production could drop as low as $0.28/litre or $1.07 / gallon

(in 2002 dollars) and in another report NREL has aimed to achieve this goal by

2012 (Aden, 2008). Concerning the RD&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

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60

a net production cost of $0.90/litre or €0.62/litre in 2010, $0.85/litre or

€0.59/litre in 2020 and $0.72/litre or €0.50/litre in 2030 (Londo et al., 2008).

In another case study, Sassners 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. (2008) performed a techno-economic evaluation of

simultaneous saccharification and fermentation (SSF) based softwood-to-

ethanol process. The economic evaluation uses the same approach as by

Sassners 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 $/litre ethanol or 4.03-5.60 and 1.34 - 1.69 US$/gallon 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$/gallon 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 per litre of ethanol. Recently, Gnansounou and Dauriat (2010)

proposed a six-step application of cost evaluation to the design of

lignocellulosics ethanol pathways; (i) 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.

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2.7. 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 Dauriats, 2010). Moreover, the international

ethanol market has been stimulated by governmental policies of incentive for

the use of renewable fuels (Table 2.7). 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 bio-ethanol with 90% gasoline (Silalertruksa

and Gheewala, 2009).

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Table 2.7: List of major policies and plans for bioethanol worldwide

Policies/Plans/Act Country

Brazilian alcohol Program (Proalcool) Brazil

Ministry of Mines and Energy, Brazil Brazil

Brazilian Sugarcane Industry Association

(UNICA) Brazil

Brazilian Renewable Energy and Incentive

Program (PROINFA) Brazil

Agriculture and Agri-food Canada (AAFC) Canada

National Science and Engineering Research

Council (NEERC, Canada) Canada

National Development and Reform Commission

(NDRC) China

National High-Tech Program (China) China

Organisation for economic cooperation and

development (OECD) Europe

European Union Strategy (EU) EU Countries

International Energy Agency (IEA) IEA countries (28)

Ministry of Petrolium and Natural Gases (MPNG) India

Ministry of New and Renewable Energy (MNRE) India

Planning Commission, Govt. of India India

National Energy Policy (Malayasia) Malayasia

Ministry of Agriculture (Russia) Russia

Bureau of Energy (BOE, Taiwan) Taiwan

United National Development Programme

(UNDP)

United Nation's member

countries (166)

Energy Information Administration (EIA) United States of America

Environment Protection Agency (EPA) United States of America

United States Department of Agriculture (USDA) United States of America

National Renewable Energy Laboratory (NREL) United States of America

Department of Energy (DOE, US) United States of America

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Many countries world wide such as Brazil, United States, China, India, Russia,

Japan, Malaysia, Canada, Europe, Korea and 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 and 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 rebatement 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 (Table 2.8). The demonstration

plants are at different levels of maturity, few of them are in advanced stage

(Abengoa, Inbicon, BioGasol), while others are in design phase.

A. SEKAB

The pilot plant of SEKAB, located in Örnsköldsvik, Sweden, was started ethanol

production in 2005. The plant has a capacity to produce 300–400 litres

ethanol/day from 2 tons biomass (Gnansounou, 2010). The main feedstock for

ethanol production is wood chips from pine trees. While other substrates such

as sugarcane bagasse, wheat and corn stover as well as energy grass can be

also used. The plant can operate with either two stage dilute acid hydrolysis or

dilute acid pre-treatment followed by simultaneous saccharification and

fermentation (Galbe et al., 2005). The technology is already been in action in a

pilot plant operated on a 24/7 basis since 2004, and the accumulated running

time now exceeds 28,000 hours (www.sekab.com/cellulosic-ethanol/demo-

plant). Moreover, the technology is ready to a scale of commercial ethanol plant

integrated with heat and power generation plant (Biofuel digest, 2009).

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Table 2.8: List of few advanced biofuels project including annual capacity

(million gallons) by company for 2009-2013.

Companies 2009 2010 2011 2012 2013 Technology

Abengoa 0.00 0.00 15.00 15.00 15.00 Enzymatic hydrolysis

ADM 0.00 0.00 0.00 1.00 1.00 Enzymatic hydrolysis

AE Biofuels 0.01 10.01 10.01 10.01 10.01 Enzymatic hydrolysis

American Process 0.00 0.00 0.89 0.89 0.89 Enzymatic hydrolysis

BlueFire Ethanol 0.01 0.01 3.91 3.91 22.91 Acid hydrolysis

Coskata 0.05 0.05 0.05 100.0 100.05 Gasification

DDCE 0.00 0.25 0.25 0.25 0.25 Enzymatic hydrolysis

Enerkem 0.30 0.30 10.30 10.30 20.30 Gasification

Fulcrum 0.01 0.01 0.01 10.51 10.51 Gasification

Haldor Topsoe 0.00 0.00 0.00 0.80 0.80 Enzymatic hydrolysis

Inbicon 1.40 1.40 1.40 19.40 19.40 Enzymatic hydrolysis

IneosBIO 0.00 0.00 8.00 8.00 8.00 Enzymatic hydrolysis

Iogen 0.48 0.48 23.48 23.48 23.48 Enzymatic hydrolysis

KL Energy 1.30 1.30 1.30 1.30 1.30 Enzymatic hydrolysis

LanzaTech 0.01 0.51 0.51 0.51 0.51 Gasification

Lignol 0.01 0.01 0.01 0.01 0.01 Enzymatic hydrolysis

Logos

Technologies 0.00 0.00 0.00 0.80 0.80 Enzymatic hydrolysis

Mascoma 0.20 0.20 0.20 20.20 20.20

Consolidated

bioprocessing

POET 0.02 0.02 25.02 25.02 25.02 Enzymatic hydrolysis

Range Fuels 0.00 20.00 20.00 20.00 20.00 Gasification

Scottish Bioenergy 0.01 0.01 0.01 0.01 0.01 Enzymatic hydrolysis

SEKAB 0.01 0.01 0.01 0.01 0.01 Enzymatic hydrolysis

St1 Biofuels Oy 0.00 0.00 0.01 0.01 0.01 Enzymatic hydrolysis

Terrabon 0.00 0.10 0.10 0.10 0.10 Hydrogenation

TMO Renewables 0.01 0.01 0.01 0.01 0.01 Enzymatic hydrolysis

UPM-Kymmene 0.00 0.00 0.00 0.68 0.68 Enzymatic hydrolysis

Verenium 1.40 1.40 1.40 37.40 37.40 Enzymatic hydrolysis

Weyland / Statoil

Hydro 0.00 0.01 0.01 0.01 0.01 Enzymatic hydrolysis

ZeaChem 0.00 0.25 0.25 0.25 0.25 Gasification/fermentation

http://www.biofuelsdigest.com/blog2/2010/02/08/advanced-biofuels-planned-capacity-to-reach-1-38-billion-gallons-by-2013/

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65

B. Abengoa Bioenergy

Abengoa Bioenergy has built their demonstration plant in Babilafuente,

Salamanca (Spain). The plant can produce 5 Million Liters ethanol per year and

can process 70 tones agricultural residues per day (Gnansounou, 2010). The

feedstocks are wheat straw and corncob and the conversion technology used is

the separate hydrolysis and fermentation (SHF). The construction of biomass

plant was completed in 2008, and in 2009, the plan betan its operation

reaching 5000 operating hours, with a yield of 200L/tonne of straw. The plant

has target to achieve a yield higher than 300L/ tonne, as well to ferment C-5

sugars. Abengoa is now developing a commercial scale in Hugoton, Kansas

(USA), to validate this technology, which can produce 100 million Liters per

annum (www.abengoa.com/corp/web/en/noticias_y_publicaciones/noticias/

historico/2011/02_ febrero/bio_20110217.html).

C. BioGasol

The company BioGasol has its pilot plant in the Technical University of

Denmark, which has a capacity of 10 tonnes ethanol per year. The company is

also manufacturing one of the first Danish demonstration- ethanol plants (5.2

million litres) of second generation feedstock (Ganansounou, 2010). The process

technology consists of pretreatment, SSF, fermentation of xylose using a

proprietary thermophilic anaerobic bacterium and production of biogas from

processed water (Ganansounou, 2010). Since the beginning of 2011, Biogasol

has been working in phase 2 of the “”BornBiofuel” Demonstration plant project,

after developing cost effective and scaleable solutions wit in pretreatment and

C-5 fermentation during phase-1 (2008-2010). The integrated plant will

demonstrate Biogasol core technologies and will be located a Aakirke by on the

island of Bornholm, Denmark (www.biogasol.com/BornBioFuel-177.aspx)

D. Inbicon and DONG Energy

Inbicon, a subsidiary of DONG Energy, has installed a cellulosic ethanol

demonstration plant in 2009 in Kalundborg (Gnansounou, 2010). The

Kalundborg will plant produce 5.4 million litres of ethanol per year from wheat

straw and will consist of SSF with a hydrothermal pre-treatment (Gnansounou,

2010; Larsen et al., 2008). The lignin will be vaporized according to the

„„Integrated Biomass Utilization System (IBUS)” concept by a cogeneration plant

to provide heat. The electricity will be sold to the electrical network (Larsen et

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66

al., 2008). The next scale up from the Inbicon Biomass Refinery will be a

commercial size that will produce 5.4 million liters a year from 30,000 Metric

tones of biomass probably wheat straw or corcob and stover

(www.inbicon.com/Biomass%20Refinery/Pages/Inbicon_Biomass_Reninery_

at_Kalundborg.aspx).

E. Shell Oil

In Canada, a Shell‟s fuel station in Ottawa (jointly run with Iogen), became the

first fuel station in the country to serve cellulosic ethanol. The station offers a

10 % blend of gasoline and wheat straw ethanol manufactured at a

demonstration-scale cellulosic ethanol of Iogen Energy Corporation (Biofuel

digest, 2009; gas2.org/2009/06/10/shell-announces=Ce10-cellulosic-ethanol-

available-now-at-ottawa-station/). The company first partnered this in 2002 at a

lower share but subsequently increased its ownership stake in Iogen‟s

technology to 50 % in 2007 (Biofuel digest, 2009; www.shell.ca/home/

content/can-en/aboutshell/media_centrre/news_and_media_releases /archieve

/2008/July12_biofuel.html).

F. Novozymes

Novozymes launched the Cellic product family, the first commercial enzymes for

cellulosic bioethanol, which have the best cost performance from amongst any

cellulosic ethanol enzymes commercially available (Biofuel digest, 2009).

Besides, Novozymes has also launched Spirizyme Ultra and Liquozyme SC 4X

for saccharification and liquification of starch, respectively (Biofuel digest,

2009). These enzymes yielded better saccharification with eventually increase

cellulosic ethanol production. Novozymes has developed a research partnership

with several international companies. It has collaborated with a Latin American

environmental firm Cetrel, which will convert sugarcane bagasse to biogas,

using enzymes from Novozymes. Similarly, they collaborated with an Indian

fermentation based company PRAJ during the Climate Change Summit in

Copenhagen in 2009. Recently Novozyme is launching Cellic CTec3-which is 1.5

times better than the previous generation Cellic Ctec-2and 5 times better than

the standard biomass degrading enzyme sin the market

(www.novozymes.com/en/solution/bioenergy/cellulosic-ethanol/cellulosic-

extraction/pages/default.aspx).

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67

G. Verenium Corporation and BP Biofuels

Verenium operated an integrated cellulosic ethanol pilot plant and a 5.29

million liters per year demonstration scale facilities in Jennings, Louisiana.

This plant is used for strengthening the company for advanced fermentation

and to use a vast range of feedstocks for cellulosic ethanol production (Biofuel

digest, 2009). The plant also serves as a R&D facility to develop new enzymes

for optimizing the cellulosic ethanol production. However, on July 15, 2010,

these plants were acquired by BP Biofuels, North America under an MOU

agreement (http://ir.verenium.com/releasedetail.cfm?ReleaseID=488510;

www.renewableenergyworld.com/rea/news/article/2010/07/bp-to-acquire-

vereniums-cellulosic-biofuel-platform).

H. POET

The first pilot plant of cellulosic ethanol was began in Scotland, SD. This plant

has an annual product capacity of 20,000 gallons of cellulosic ethanol, but

more importantly,it allows new technogies and process to validate. This help in

reducing the ethanol production cost for $4.13/gallon to $3.0/gallon

(www.poet.com/innovation/cellulosic/pilot.asp). Poet has committed to produce

3.5 billion gallons of cellulosic ethanol by 2022

(www.poet.com/innovation/cellulosic/plan.asp). Now they have announced to

launch its first commercial scale demonstration plant under the proect-

LIBERTY in IOWA in late 2013. The plant is expected to produce 25 million

gallon per annum cellulosic ethanol (www.projectliberty.com).

2.8. Future Prospects

Ethanol has always been considered a better short- and mid-term liquid biofuel,

as it reduces the dependence on reserves of crude oil and promises cleaner

combustion leading to a healthier environment. 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). However, to

reduce the final production cost, technological bottlenecks in commercial

production of lignocellulosic ethanol from pentose sugars need to be addressed.

In recent years, a significant development in ethanol conversion has been

made, however the current industrial activity of bioethanol production is limited

mainly because of the raw material processing, cost-effective sugar release from

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biomass, and unavailability of efficient fermentation strategies. The process for

conversion of structural carbohydrates to sugars requires adequate

deconstruction of lignocellulosic biomass leading to maximum sugar yield. 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 bioprospecting 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

(Mosier et al., 2005; Chandel et al., 2007a; Gupta et al., 2009; Kuhad et al.,

2010b; Palmqvist and Hahn-Hagerdal, 2000a, b), however, the process of

detoxification also increase the processing cost. Therefore, there is an

imperative need for bioprospecting of new microbes capable of converting

pentose sugars present in the hydrolysate efficiently even in the presence of the

toxic inhibitors or use of robust hosts, such as Bacillus (Geobacillus) strains

(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). An approach of simultaneous saccharification and

fermentation (SSF) should be employed to exploit both the sugars in a more

economic way. Various fermentation strategy such as CSTR and fed-batch

strategy should be used to enhance the ethanol concentration. The approach of

genetic engineering should be used to improve the microorganisms in terms of

higher ethanol tolerance, inhibitor tolerance, osmotolerance and co-

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fermentation of C-5 and C-6 sugars. Additionally, the better elucidation of

pentose sugars transport at the molecular level and characterization of kinetic

and regulatory properties, including quorum-sensing mechanisms, should be

given high priority because it may provide the basis for the simultaneous use of

pentose as well as hexose sugars released during biomass hydrolysis (Galazka

et al., 2010). Moreover, the approach of flux analysis to divert or increase the

activity of certain crucial enzymes for ethanol production with efficient xylose

utilization must be a priority (Matushika et al., 2009b).

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

titers, 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.

Before we freeze in the dark, we must prepare to make the transition from nonrenewable carbon

resources to renewable bioresources.

Ragauskas and coworkers (2006)